Abstract
The coccidian parasite Cyclospora cayetanensis is the causative agent for foodborne outbreaks of cyclosporiasis and multiple fresh produce recalls annually. In recent years, this organism has been reported in the water near produce growing operations during outbreak investigations, prompting a call for more research on its environmental prevalence in the United States. Currently, there is a lack of performance data available on methods for conducting this research, including the performance of DNA extraction methods for molecular testing. Extraction methods for environmental samples must be efficient due to the often-limited amount of target nucleic acid and the potential for molecular inhibitors present in an environmental sample. This study assessed the performance of C. cayetanensis nucleic acid extraction seeded into surface water, produce wash water, and tap water by two methods designed for use with environmental samples: the PowerViral and UNEX methods. The PowerSoil extraction method (2 g) was assessed for C. cayetanensis extraction from seeded sewage sludge – an environmental sample type used to evaluate parasite carriage within communities. Extraction performance of the PowerViral and UNEX methods were also assessed for the detection of the foodborne bacterial pathogen Salmonella and a surrogate for foodborne viruses, murine norovirus (MNV) seeded into surface water, produce wash water, and tap water. The PowerViral method resulted in consistent detection (83–100 %) of C. cayetanensis, S. enterica, and MNV across all water types. Detection rates for the UNEX method ranged from 56 to 100 % prevalence for tap water and wash water, but there were no detections for any microbe from surface water. The PowerSoil method resulted in poor recovery of C. cayetanensis from sludge (≤1 % recovery), while both the PowerViral and UNEX methods effectively recovered C. cayetanensis from sludge (4–36 % recovery).
Keywords: Cyclospora cayetanensis, Salmonella enterica, Murine norovirus, DNA extraction, RNA extraction, Molecular methods
1. Introduction
Cyclospora cayetanensis is a coccidian parasite that causes the foodborne illness cyclosporiasis. C. cayetanensis is excreted as an environmentally resistant oocyst from the infected human host (Centers for Disease Control and Prevention, 2021; Quintero-Betancourt et al., 2002). United States outbreaks of cyclosporiasis were typically linked to fresh produce grown in endemic countries outside of the U.S. until outbreaks in 2018 and 2020 were associated with domestically grown produce (Abanyie et al., 2015; Hadjilouka and Tsaltas, 2020; U.S Food and Drug Administration, 2020, 2021). While fresh produce is often identified as the source of cyclosporiasis outbreaks, oocysts of C. cayetanensis require, on average, seven to fourteen days to sporulate into their infectious state after shedding from the host (Smith et al., 1997). This makes food contamination from harvest workers or person-to-person transmission unlikely, but instead agricultural waters could serve as an effective vehicle for contaminating growing produce and allow for sporulation to occur prior to consumption. To effectively evaluate U.S. produce irrigation waters for C. cayetanensis, the performance of testing methods for C. cayetanensis in environmental samples must be understood.
The most sensitive methods available for the detection of C. cayetanensis in environmental samples employ real-time, quantitative polymerase chain reaction (qPCR); therefore, efficient nucleic acid extraction and purification is an important first step in detection. Extraction is particularly important for effective C. cayetanensis detection because rupturing the chemically resistant oocyst wall is necessary for nucleic acid release (Erickson and Ortega, 2006; Qvarnstrom et al., 2018a; Shields et al., 2013). Nucleic acid extraction method performance can greatly affect the subsequent interpretation of C. cayetanensis qPCR data and different commercial kits have been shown to yield varying DNA output quantities, qualities, and subsequent target amplification efficiencies (Almeria et al., 2018; Assurian et al., 2020b; Iker et al., 2013; O’Brien et al., 2021; Qvarnstrom et al., 2018a; Shields et al., 2013; Zielińska et al., 2017). Factors to consider when choosing an extraction method include: sample matrix, sample volume to be extracted, target organism type (e.g., parasite, bacteria, and/or virus), target nucleic acid type (e.g., RNA and/or DNA), and predicted target concentration in tested samples (Thatcher, 2015). Moreover, environmental water concentrates contain high amounts of naturally occurring substances that could act to inhibit PCR. These include humic substances, which are the product of microbial breakdown of plant material and are co-concentrated with the molecular targets (Abbaszadegan et al., 1993; Gentry-Shields et al., 2013). Humic substances interfere with PCR by chelating the ions needed for the polymerase and have been shown to interfere with the molecular reaction (Tsai and Olson, 1992). For this reason, an inhibitor removal step is beneficial to an environmental nucleic acid extraction technique.
Previous research assessing the performance of extraction methods for C. cayetanensis detection have largely focused on extraction from food or clinical samples (Qvarnstrom et al., 2018a; Shields et al., 2013; Temesgen et al., 2020). However, the utility of these methods for environmental water and sewage sludge samples is unknown, as previous methodological studies using environmental samples have emphasized the performance of sample concentration, oocyst isolation or molecular detection methods, neglecting the assessment of DNA extraction efficacy (Borchardt et al., 2009; Kahler et al., 2021; Shields and Olson, 2003; Totton et al., 2021).
Our study assessed the efficacy of two extraction methods for detecting C. cayetanensis in three water types with varying levels of organic matter and inhibitors – surface, produce wash, and tap – as well as sewage sludge, which was previously tested to evaluate parasite carriage within communities (Kahler et al., 2024). To evaluate the performance for co-extraction of other food safety related microbial targets, all water samples were also seeded with the gram-negative bacteria, Salmonella enterica, and an RNA, enteric virus surrogate, murine norovirus-1 (MNV). Two extraction methods were evaluated for water samples: the Qiagen AllPrep® PowerViral® DNA/RNA kit (Qiagen, Hilden, Germany) and the Universal Nucleic Acid Extraction (UNEX) method (Hill et al., 2015). Three extraction methods were evaluated for sewage sludge samples: the two evaluated for water samples and the Qiagen RNeasy PowerSoil Total RNA kit with the RNeasy PowerSoil DNA Elution kit. These three extraction methods were selected due to their ability to co-extract DNA and RNA, incorporate bead beating steps to facilitate cell lysis, and utilize inhibitor removal mechanisms. The three extraction methods evaluated were designed for different input volumes, which can impact the sensitivity of pathogen detection. The PowerViral and UNEX methods require smaller input volumes, 200 μL and up to 750 μL, respectively, while the PowerSoil method accepts larger amounts (2 g). Two method variations were evaluated for the PowerViral and UNEX methods. Two input volumes were assessed to evaluate whether increasing sample volume negatively impacted extraction efficiency. Two types of beads for bead beating were assessed to evaluate the impact of bead composition on extraction efficiency.
2. Materials and methods
2.1. Environmental sample collection and concentration
Surface and tap water samples were collected from September to November 2020 from streams, ponds, and buildings in DeKalb County, Georgia, U.S. Produce wash water and municipal sewage sludge samples were collected between January 2020 through December 2021 in Tift County, GA. Sample processing and nucleic acid extraction were conducted at the Centers for Disease Control and Prevention (CDC) in Atlanta, GA.
Tap water (100L) was collected at two different time points (two samples) directly from a municipal-fed tap by dead-end ultrafiltration (DEUF) following previously described methods (Kahler et al., 2015). Prior to ultrafiltration, tap water samples were treated with 0.01 % sodium polyphosphate (NaPP) (Sigma-Aldrich No. 305553, St. Louis, MO) to neutralize free chlorine residual. Surface water samples (50–75 L) were collected on different days from a small tributary to the South Fork Peachtree Creek (N = 3) and from Murphey Candler Lake in Atlanta, GA (N = 1). Surface waters were ultrafiltered similar to tap water, and all ultrafilters were backflushed as previously described (Kahler et al., 2021). Surface and tap water sample backflush was then further concentrated by centrifugation at 4000 xg for 15 min. All but 10 mL of the supernatant was removed, which was then used to resuspend the pelleted sample. The resulting concentrates were stored at −20 °C until microbe seeding and nucleic acid extraction.
Spent produce wash water was collected at two separate time points (two samples) by pumping from the produce dump tank into a 20-L cubitainer at a packing house facility. Water was concentrated by continuous flow centrifugation (CFC) as previously described (Kahler et al., 2021). CFC eluates were further concentrated by centrifugation at 4000 xg for 15 min. The resulting concentrates (15 mL) were stored at −20 °C until microbe seeding and nucleic acid extraction.
Municipal sludge samples (1 L) were collected from two locations in the wastewater treatment plant processing system at three separate time points for each sampling location. The two locations include the thickener recovery system (REC) used to generate sludge for land application (N = 3) and the return activated sludge (RAS) from the aeration basin (N = 3). Samples were frozen at −20 °C upon arrival at CDC until further processing (3–19 months). Thawed sludge samples were heat inactivated in a 60 °C water bath for one hour. After cooling, approximately 500 mL was concentrated by centrifugation at 4000 xg for 15 min. The supernatant was removed, and sludge concentrates were stored at −20 °C until microbe seeding and nucleic acid extraction.
2.2. Microbe preparation and sample seeding
Flow-sorted C. cayetanensis oocysts were purified from human stool pooled specimens collected in Indonesia, and oocysts were quantified into aliquots of one million oocysts following previously published methods (Qvarnstrom et al., 2018b). The sporulation status of the oocysts was observed on a Zeiss Primostar microscope at 40× magnification (Zeiss, Oberkochen, Germany). The prepared C. cayetanensis stock was stored at 4 °C. Salmonella enterica (ATCC 19585) was grown in nutrient broth and aliquots were stored in 20 % glycerol at −80 °C. Murine norovirus 1 (MNV) was cultured in RAW 264.7 cells (Wobus et al., 2004). MNV was harvested from the lysates of infected cells after one freeze-thaw cycle, aliquoted and stored at −80 °C. Salmonella stocks were enumerated by spread plating on nutrient agar and MNV stocks were enumerated by plaque assay (Cromeans et al., 2010).
Thawed water and sludge concentrates were seeded immediately prior to extraction. For each microbe stock, 10-fold serial dilutions were made to achieve concentrations close to the desired seed quantity. Dilutions were made using PBS containing 0.01 % Tween 80 and 0.001 % Antifoam Y-30 emulsion to facilitate cell dispersion. Water concentrates were seeded with 1,000C. cayetanensis oocysts, 1,000 CFU of S. enterica, and 1,000 PFU of MNV, regardless of input volume. Sludge concentrates were only seeded with C. cayetanensis oocysts (no S. enterica or MNV) at seed levels of either 1,000 total oocysts, regardless of extraction method input volume, or with 500 oocysts per gram input volume.
2.3. Nucleic acid extraction methods
The PowerViral and UNEX extraction methods were evaluated for tap, surface, produce wash water, and sludge samples (Table 1). The PowerSoil method was evaluated for sludge sample concentrates, which allowed for extraction of a larger sample volume (2 g vs. 0.2 g wet sludge for PowerViral and UNEX) (Table 2). Extractions were performed in triplicate for each sample type and seed level to account for potential heterogeneity within the sample matrix and for variation in the microbe seed quantity across the three replicates. For all extraction methods and sample types, an extraction blank consisting of nuclease-free water was extracted alongside samples. For a subset of extracts, nucleic acid concentration (ng/μL) and purity (A260/280 ratio) was measured using a NanoDrop One spectrophotometer (Thermo Fisher Scientific, Waltham, MA) according to manufacturer’s instructions.
Table 1.
Number of replicate water sample extractions using the PowerViral and UNEX methods using different input volumes, bead beating bead composition (bead tube), and UNEX lysis buffer incubation conditions.
| Extraction method | PowerViral | UNEX | ||||||||
|---|---|---|---|---|---|---|---|---|---|---|
| Input volume (μL) | 200 | 375 | 200 | 375 | ||||||
| Bead tubea | PV | ME | PV | UNEX | UNEX | ME | ME | UNEX | ||
| Heated incubation (Y/N)b | N | N | N | Y | N | Y | N | Y | N | |
| Water type | Sample | No. of replicate extractions | ||||||||
| Surface | A | 3 | 3 | 3 | -c | 3 | - | 3 | - | 3 |
| B | 3 | 3 | 3 | - | 3 | - | 3 | - | 3 | |
| C | 3 | - | - | 3 | - | - | - | - | - | |
| D | 3 | - | - | 3 | - | - | - | - | - | |
| Tap | A | 3 | 3 | 3 | - | 3 | - | 3 | - | 3 |
| B | 3 | 3 | 3 | 3 | - | 3 | - | 3 | - | |
| Wash | A | 3 | 3 | 3 | 3 | - | 3 | - | 3 | - |
| B | 3 | - | - | 3 | - | - | - | - | - | |
PV = bead tube provided in PowerViral kit; ME = Matrix E tube; UNEX = bead tube indicated for UNEX method.
Heated incubation of lysis buffer prior to bead beating. Y: Yes; N: No.
-: no extractions conducted for these conditions.
Table 2.
Number of replicate sludge sample extractions using the PowerViral, UNEX, and PowerSoil methods with different C. cayetanensis seed quantities.
| Extraction method | PowerViral | UNEX | PowerSoil | |||
|---|---|---|---|---|---|---|
| Input amount | 0.2mL | 0.2mL | 2g | |||
| Oocyst seed quantity | 100 | 1000 | 100 | 1000 | 1000 | |
| Sludge type | Sample | No. of replicate extractions | ||||
| RASa | A | 3 | 3 | 3 | 3 | 6 |
| B | 3 | 3 | 3 | 3 | 6 | |
| C | 3 | 3 | 3 | 3 | 6 | |
| RECb | A | 3 | 3 | 3 | 3 | 3 |
| B | 3 | 3 | 3 | 3 | 6 | |
| C | 3 | 3 | 3 | 3 | 3 | |
RAS: return activated sludge from aeration basin.
REC: thickener recovery system sludge.
2.3.1. PowerViral extraction
Two hundred microliters of water concentrates and 0.2 g sludge concentrates were extracted using the AllPrep® PowerViral® DNA/RNA kit (Qiagen, Hilden, Germany) according to the manufacturers protocol with the following modifications. Dithiothreitol (DTT) (Fisher, Fair Lawn, NJ) was added to the lysing buffer at a concentration of 40 mM to reduce RNAse activity instead of β-mercaptoethanol, per manufacturer’s recommendation (see Supplementary Information Table S1 for comparison data). Bead beating was performed on a FastPrep-24™ Classic bead beating grinder and lysis system (MP biomedicals, Irvine, CA) at a speed of 6 m/s for 60 s, and the final elution was completed with 100 μL of TE buffer.
Method variations were assessed for two sets of surface and tap water samples and one set of wash water samples (Table 1). During these experiments, an additional set of samples were extracted using input volumes of 375 μL by proportionally scaling up reagents. A separate set of samples were extracted using Lysing Matrix E (ME) 2-mL bead tubes (MP Biomedicals, Irvine, CA). The ME bead tubes contain 1.4 mm ceramic spheres, 0.1 mm silica spheres and one 4 mm glass bead, while the PowerViral bead tubes contain 0.1 mm glass beads.
2.3.2. UNEX extraction
Two hundred microliters of water concentrates and 0.2 g sludge concentrates were extracted using the UNEX method (Hill et al., 2015). Sample concentrates were added to FastPrep® 2-mL Lysing Matrix tubes (hereafter referred to as UNEX bead tubes) (MP Biomedicals, Irvine, CA) containing ~200 mg each of 0.2 μm and 0.5 μm zirconium oxide beads (Union Process, Akron, OH). Two hundred microliters of UNEX lysis buffer (Microbiologics, St. Cloud, MN) and 20 μL Proteinase K (Qiagen, Hilden, Germany) were added to the bead tubes and the contents were mixed by briefly vortexing. Samples were incubated before bead beating to allow for cell lysing and Proteinase K digestion. Initially, incubation was performed for 15 min at room temperature. However, data from the first two experiments (Surface A and B; Tap A) showed that the UNEX method yielded lower than expected target organism recovery based on the seed level. Thereafter, the incubation conditions were adjusted to 56 °C for 1 h to attempt to improve protein digestion (Qvarnstrom et al., 2018a).
Bead beating was performed on a FastPrep-24™ Classic at a speed of 6 m/s for 60 s, followed by centrifugation at 10,000 xg for 30 s to pellet cell debris. Bead beating supernatants were added to HiBind® RNA Spin Columns (Omega BioTek, Norcross, GA) and then centrifuged at 10,000 xg for 1 min to capture DNA and RNA. The columns were washed first with 500 μL of 100 % ethanol, then 500 μL of 70 % ethanol, followed by centrifugation after each at 10,000 xg for 1 min. A final dry spin was performed at 10,000 x g for 1 min to remove any residual ethanol from the silica columns. Nucleic acid was eluted from the columns with 100 μL of TE buffer by centrifuging at 9000 x g for 1 min. The eluates were passed through the OneStep PCR Inhibitor Removal Kit, following manufacturer’s instructions (Zymo Research, Tustin, CA).
Method variations were assessed for two sets of surface and tap water samples and one set of wash water samples in the same manner as for PowerViral extractions (Table 1).
2.3.3. PowerSoil extraction
Two grams of sludge concentrates were extracted following the manufacturers protocol using the RNeasy PowerSoil Total RNA kit (Table 2). After the initial RNA elution, the RNeasy PowerSoil DNA Elution kit (Qiagen, Hilden, Germany) was used to extract the DNA from each sample. The final DNA and RNA pellets were each resuspended with 100 μL TE buffer.
2.4. Molecular analysis
All nucleic acid extracts were stored at 4 °C and subjected to qPCR within 24 h of DNA extraction. qPCR was performed for the detection of C. cayetanensis targeting the 18S rRNA gene (Qvarnstrom et al., 2018a), for S. enterica targeting the ttrRSBCA locus (Malorny et al., 2004), and for MNV targeting a region of the RNA-dependent RNA polymerase gene (Hill et al., 2007). Water extracts were analyzed in duplicate for each target with 5 μL template in 50 μL reaction volumes. TaqMan™ Environmental Master Mix 2.0 (Applied Biosystems, Waltham, MA) was used for the C. cayetanensis and S. enterica assays, and TaqMan™ Fast Virus 1-Step Master Mix was used for MNV assays. Bovine serum albumin (0.5 mg/mL) and T4 Gene 32 protein (gp32, New England Biolabs, Ipswich, MA) (25 μg/mL) were added to the reaction mixture for all three targets to improve amplification efficiency and reduce inhibition effects.
qPCR was performed in a 7500 Real-Time PCR System with software version 2.3 (Applied Biosystems). Cycling conditions for C. cayetanensis and S. enterica each started with denaturation at 95 °C for 10 min, followed by 45 cycles of denaturation at 95 °C for 15 s, then annealing and fluorescence acquisition at 67 °C for one minute for the C. cayetanensis assay and at 60 °C for the S. enterica assay. qPCR cycling conditions for MNV were one cycle of reverse transcription at 50 °C for five minutes, denaturation at 95 °C for 20 s, followed by 45 cycles of denaturation at 95 °C for 20 s, and annealing and fluorescence acquisition at 56 °C for one minute. The amplification threshold was set to 0.02 delta normalized reporter value (ΔRn) for C. cayetanensis, 0.03 ΔRn for S. enterica, and 0.05 for MNV. Positive controls consisted of genomic DNA or RNA of each organism, and no-template controls consisted of nuclease-free water. Extraction blanks from each extraction method, positive controls, and no-template controls were analyzed in duplicate with each instrument run.
Sludge extracts were analyzed for C. cayetanensis following the same qPCR assay specifications as for water samples with the following modifications. All samples and controls were analyzed in triplicate. A 1:4 dilution was analyzed in addition to each undiluted sample for PowerSoil extracts. Five hundred copies of a commercially synthesized DNA oligo was added to each qPCR as a non-competitive internal amplification control (IAC) to monitor for reaction inhibition as previously described (U.S. Food and Drug Administration, 2017). A standard calibration curve consisting of a synthetic gBlock gene fragment (IDT gBlock, Coralville, IA) was generated with dilutions ranging from 106 through 100 copies per reaction (U.S. Food and Drug Administration, 2017) and included in three instrument runs to generate a pooled standard curve equation for quantification of the C. cayetanensis target.
Gene targets were considered detected if amplification occurred at a quantification cycle threshold (Cq) less than 40 for both replicates for water extracts and for at least two out of three replicates for sludge extracts. Sludge extracts were scored as detected if either set of technical replicates (undilute or 1:4 dilution) met these criteria. A mean Cq of 40 was assigned to all non-detect water extracts for statistical analyses. For sludge extracts with 2 of 3 positive replicates, a Cq of 40 was assigned to the non-detect replicate and then averaged with the other two Cq vales for statistical analyses. Detected C. cayetanensis 18S rRNA gene target copy number was calculated by inputting the mean Cq of the detected technical replicates into the pooled standard curve equation. Sludge extracts in which the IAC had a ≥ 3 Cq value difference from the no-template control were classified as inhibited but were included in analyses using data from the technical replicates (undilute or 1:4) that produced the higher number of detected gene copies.
2.5. Statistical analysis
Statistical analyses for nucleic acid concentration and purity comparisons were performed using Excel’s data analysis tool. Comparisons between the PowerViral and UNEX extraction methods for water samples were made using paired two-tailed t-tests. One way ANOVA was performed to compare nucleic acid purity for sludge samples.
Statistical analyses for extraction method performance were performed in R version 4.2.2 (R Core Team, 2021). Multiple linear regression analysis was used to assess the impact of method variations (e.g., input volume, bead tube type, heated incubation) on microbe recovery from water samples, as measured by Cq values. Separate models were built for each combination of extraction method (PowerViral or UNEX), sample type (tap or surface water), and microbe (C. cayetanensis, S. enterica, MNV).The mean Cq of the qPCR replicates for each extract was used as the response variable and indicator variables for input volume (200 μL vs. 375 μL) and bead tube (Matrix E vs. PowerViral or UNEX bead tubes, respectively) were used as the predictor variables. Use of a heated incubation step was also included as a predictor in regression models for the UNEX method. Models for wash water were not constructed for either extraction method because all but one PowerViral wash water extractions used the PowerViral bead tube and all UNEX wash water extractions utilized a heated incubation step. Accordingly, a negative regression coefficient that corresponded to an estimated mean difference in Cq that was significantly different from zero at the 5 % significance level was interpreted as evidence that the corresponding methodological variation was associated with superior microbial recovery.
Comparison of PowerViral and UNEX recovery from water samples was assessed using non-parametric Wilcoxon signed-rank tests to compare the distributions of Cq values for each microbe and water type. Method variations that were not associated with significantly different Cq values in the regression analyses were considered to perform equivalently and all data from such variations were included in the performance analysis comparison to the other extraction method. Recovery was compared within a water sample (e.g., A, B, C) when data from all equivalent method modifications were available (N = 6 extractions per method), and when 67 % of all extracts (N ≥ 8/12) were classified as detects. This was done to prevent bias from non-detects or unpooled data.
Multiple linear regression was performed to determine if extraction method (PowerSoil, PowerViral, or UNEX), sludge type (REC or RAS), or the sludge sample extracted (e.g. A, B, C) impacted C. cayetanensis percent recovery from sludge. Comparison of the performance of the extraction methods for sludge samples was compared using percent recovery instead by comparing of Cq value because the seed level was not the same for each extraction method for sludge. Percent recovery was calculated by dividing the detected number of C. cayetanensis 18S rRNA gene copies by the estimated seeded gene copy number. The estimated seeded gene copy number was obtained by multiplying the number of seeded oocysts by the number of gene copies of the 18S rRNA gene per genome. Microscopy observations of the C. cayetanensis oocyst stock revealed exclusively unsporulated oocysts (Assurian et al., 2020a). An unsporulated C. cayetanensis oocyst contains two haploid genomes, containing an average of 17.5 copies of the 18S rRNA gene. Therefore, the 1000-oocyst seed was estimated to contain 35,000 gene copies and the 100-oocyst seed was estimated to contain 3500 gene copies.
3. Results
3.1. Effect of input volume and bead beating method on PowerViral performance for water samples
The PowerViral extraction method resulted in consistent detection of C. cayetanensis, S. enterica, and MNV from all water sample types and method variations (Table 3, Table S2). Neither the input volume nor the bead type affected the Cq values for S. enterica or MNV in any water type.
Table 3.
Number of replicate extract detections from water samples seeded with C. cayetanensis, S. enterica, and MNV using the PowerViral and UNEX methods using different input volumes, bead beating bead composition (bead tube), and UNEX lysis buffer incubation conditions. Gray shading indicates the method variations that performed equivalently for each extraction method and only these data were used for analyses between methods.
| Extraction method | Power Viral | UNEX | ||||||||
|---|---|---|---|---|---|---|---|---|---|---|
| Input volume (μL) Bead tubea Heated incubation (Y/N)b |
200 PV ME N N |
375 PV N |
200 UNEX UNEX ME ME Y N Y N |
375 UNEX Y N |
||||||
| Water type | Sample | C. cayetanensis detections (N=3) | ||||||||
| Surface | A B C D |
3 3 3 3 |
3 3 - - |
3 3 - - |
- c - 0 0 |
0 0 - - |
- - - - |
0 0 - - |
- - - - |
0 1 - - |
| Tap | A B |
3 3 |
3 3 |
2 3 |
- 3 |
3 - |
- 3 |
1 - |
- 3 |
3 - |
| Wash | A B |
3 3 |
3 - |
3 - |
3 3 |
- - |
3 - |
- - |
3 - |
- - |
| Water type | Sample | S. enterica detections (N=3) | ||||||||
| Surface | A B C D |
3 3 3 3 |
3 2 - - |
3 2 - - |
- - 0 0 |
0 0 - - |
- - - - |
0 0 - - |
- - - - |
0 1 - - |
| Tap | A B |
3 3 |
3 3 |
3 3 |
- 2 |
2 - |
- 0 |
1 - |
- 3 |
2 - |
| Wash | A B |
NDd 3 |
2 - |
3 - |
ND 3 |
- - |
2 - |
- - |
3 - |
- - |
| Water type | Sample | MNV detections (N=3) | ||||||||
| Surface | A B C D |
- 3 3 3 |
- 3 - - |
- 3 - - |
- - 0 0 |
- 0 - - |
- - - - |
- 0 - - |
- - - - |
- 0 - - |
| Tap | A B |
3 3 |
3 3 |
3 3 |
- 3 |
2 - |
- 3 |
2 - |
- 3 |
3 - |
| Wash | A B |
3 3 |
3 - |
3 - |
1 3 |
- - |
2 - |
- - |
1 - |
- - |
PV = bead tube provided in PowerViral kit; ME = Matrix E tube; UNEX = bead tube indicated for UNEX method.
Heated incubation of lysis buffer prior to bead beating. Y: Yes; N: No.
-: no extractions conducted for these conditions.
ND: no data obtained due to a seeding error for these replicates.
C. cayetanensis recovery from tap water using the PowerViral method was impacted by input volume and bead type. Mean Cq was 3.66 cycles lower when input volume was 200 μL versus 375 μL (i.e., approximately 1-log10 more target copies recovered with 200 μL input) [(95 % confidence interval (95 %CI): 2.21–5.11 cycles; p < 0.001)]. Mean Cq was 2.06 cycles lower when bead beating was done with the Matrix E bead tube versus the PowerViral bead tube (95 %CI: 0.61–3.51 cycles; p = 0.01).
C. cayetanensis recovery from surface water using the PowerViral method was also impacted by input volume, but not bead type. Mean Cq was 1.6 cycles lower when input volume was 200 μL versus 375 μL (95 % CI: 0.32–2.88 cycles; p < 0.05). However, mean Cq values for the PowerViral and Matrix E bead tubes were not significantly different (p = 0.93). All positive controls, no-template controls, and extraction blanks performed as expected.
3.2. Effect of input volume and bead beating method on UNEX performance for water samples
The UNEX extraction method resulted in inconsistent detection across water types. Detection rates for all microbes were higher for tap and wash water samples than for surface water, which had few detections (Table 3, Table S3). Analyses for the UNEX method could only be performed for tap water, as there were too few detections from surface water. Mean S. enterica Cq values were not impacted by input volume (p = 0.57), bead tube (p = 0.06), or heated incubation (p = 0.96).
C. cayetanensis recovery using the UNEX method was not impacted by input volume, as the mean Cq values for the 200 μL and 375 μL input volumes were not significantly different (p = 0.98). However, recovery was greater for both the UNEX bead tube and the heated incubation. Mean Cq was 4.18 cycles lower when bead beating was done with the UNEX bead tube versus the Matrix E tube (95 %CI: 2.67–5.69 cycles); p < 0.001) and was 2.57 cycles lower when extracted after a heated incubation (95 %CI: 1.33–3.80 cycles; p < 0.001).
MNV recovery using the UNEX method was not impacted by input volume (p = 0.22) or bead type (p = 0.48). However, a heated incubation significantly improved MNV recovery from tap water, resulting in mean Cq values that were 7.56 cycles lower than samples without heated incubation (i.e., over 2-log10 more target copies recovered using heated incubation) (95 %CI: 6.37–8.75 cycles; p < 0.001). All positive controls, no-template controls, and extraction blanks performed as expected.
3.3. Comparison of extraction method detection rates for water samples
PowerViral and UNEX method performance were compared using data from the best performing extractions for each method (Sections 3.1 and 3.2). Data from the method variations that performed equivalently were pooled (Table 3, gray shading). The PowerViral data set comprised extractions with a 200 μL input volume and both bead tubes. The UNEX data set comprised extractions with 200 μL and 375 μL input volumes, a heated incubation step, and the UNEX bead tube. From these pooled data sets, C. cayetanensis, S. enterica, and MNV were detected in nearly 100 % of surface water extracts using the PowerViral method (17/18 S. enterica detections), but there were no detections for any microbe using the UNEX method. Almost all microbes were detected in all tap water extracts for both the PowerViral and UNEX methods except for S. enterica in one 200 μL UNEX extraction. C. cayetanensis was detected in 100 % of wash water samples using both the UNEX and PowerViral methods. S. enterica was detected in 83 % of PowerViral extracts and 100 % of UNEX extracts for wash water. MNV was detected in 100 % of wash water samples using the PowerViral method, but only 56 % of samples using the UNEX method.
3.4. Comparison of extraction method recovery for water samples
Recovery performance between the PowerViral and UNEX methods were compared using the median Cq achieved by each method (Table 4). Extracts from samples Tap B and Wash A were the only samples to meet the inclusion criteria for comparison of recovery between the PowerViral and UNEX methods. Recovery of C. cayetanensis was not significantly different between the methods for tap water (W = 8.5, p = 0.15) or wash water (W = 7, p = 0.09). MNV recovery was greater using the PowerViral method, as median MNV Cq values were significantly lower for tap water (W = 0, p = 0.002) and wash water (W = 0, p = 0.005) samples extracted with the PowerViral method than those extracted with the UNEX method. Analyses for S. enterica could only be completed for tap water samples, for which recovery was greater using the PowerViral method, which had significantly lower median Cq value than UNEX extractions (W = 0, p = 0.002).
Table 4.
Median and range of the mean Cq values for C. cayetanensis, S. enterica and MNV across extraction replicates for each water sample (Tap B and Wash A) using PowerViral and UNEX methods.
| Organism | Sample | Methoda | Median Cq Value (range) |
|---|---|---|---|
| C. cayetanensis | Tap B | PowerViral | 31.23 (30.54–31.73) |
| UNEX | 31.66 (30.77–32.79) | ||
| Wash A | PowerViral | 32.63 (31.74–34.42) | |
| UNEX | 33.95 (33.01–34.39) | ||
| S. enterica | Tap B | PowerViral | 32.06 (31.67–32.47) |
| UNEX | 33.27 (32.61–37.68) | ||
| MNV | Tap B | PowerViral | 26.86 (26.20–27.09) |
| UNEX | 29.90 (29.30–30.24) | ||
| Wash A | PowerViral | 27.96 (27.81–28.12) | |
| UNEX | 35.44 (34.65–40) |
The PowerViral category included all samples with a 200 μL input volume that were extracted with either the PowerViral or Matrix E bead tube. The UNEX category included all samples with either a 200 μL or 375 μL input volume extracted with the UNEX bead tube and a heated incubation.
3.5. Comparison of extraction method nucleic acid concentration and purity
Water nucleic acid concentrations (ng/μL dsDNA) and purity ratios (260/280 nm absorbance ratio) from 200 μL extractions using the method-specific bead tubes are presented in Table S4. UNEX extract DNA concentrations were 12 times higher than PowerViral extracts for surface water (p = 0.04), and 19 times higher for tap water (p = 0.01). PowerViral extract purity ratios were closer to pure DNA than UNEX extracts, as measured by the deviation from the 1.8, which is considered a pure nucleic acid purity ratio. Mean purity ratio difference from 1.8 was 2.4 times higher for UNEX extracts (mean ratio difference = 1.0, SD = 0.05) than for PowerViral extracts (mean ratio difference = 0.43, SD = 0.11) for surface water (p = 0.009), and 2.2 times higher for tap water (UNEX: mean ratio difference = 0.97, SD = 0.04; PowerViral mean ratio difference = 0.45, SD = 0.06); p = 0.001).
3.6. Comparison of extraction method detection rates and inhibition for sewage sludge
C. cayetanensis was detected in 100 % of sludge extracts using the UNEX and PowerViral methods, regardless of seed quantity (Table 5). Using the PowerSoil method, in which samples were seeded with 1000 oocysts, C. cayetanensis was detected in all REC sludge extracts and 89 % of RAS extracts. Mean C. cayetanensis percent recoveries from sludge samples were 17–36 % using the PowerViral method and 4–12 % using the UNEX method (Table 5). Mean C. cayetanensis recoveries for the PowerSoil method were ≤ 1 %. Inhibition from RAS sludge was observed in 11–44 % of PowerViral extracts, 22–33 % of UNEX extracts, and 84 % of PowerSoil extracts (Table 5). While the percentage of inhibited extracts from REC sludge was 0–1 % for the PowerViral and UNEX methods, 58 % of PowerSoil extracts exhibited inhibition. The mean IAC Cq differences for inhibited extracts are also presented in Table 5.
Table 5.
Number of replicate extract C. cayetanensis detections from sludge samples, mean percent recovery and standard deviation, number of inhibited extracts, and mean difference in internal amplification control Cq values for inhibited samples.
| Sludge type | Extraction method | Seed quantity | N | Detects | Mean % recovery (SDV) | Inhibited extracts (%) | Mean IAC Cq difference for inhibited extracts |
|---|---|---|---|---|---|---|---|
| RAS | PowerSoil | 1000 | 18 | 16 | 0.17 (0.13) | 15 (83) | 6.65 |
| PowerViral | 1000 | 9 | 9 | 28 (5) | 1 (11) | 3.16 | |
| PowerViral | 100 | 9 | 9 | 17 (13) | 4 (44) | 6.42 | |
| UNEX | 1000 | 9 | 9 | 10 (2) | 2 (22) | 3.32 | |
| UNEX | 100 | 9 | 9 | 4 (3) | 3 (33) | 5.85 | |
| REC | PowerSoil | 1000 | 12 | 12 | 1.1 (0.6) | 7 (58) | 5.07 |
| PowerViral | 1000 | 9 | 9 | 32 (7) | 0 | NAa | |
| PowerViral | 100 | 9 | 9 | 36 (9) | 0 | NA | |
| UNEX | 1000 | 9 | 9 | 11 (3) | 0 | NA | |
| UNEX | 100 | 9 | 9 | 12 (4) | 1 (11) | 3.12 |
NA: not applicable; no inhibition observed in these extracts.
3.7. Comparison of extraction method recovery for sewage sludge
Detected copies of the 18S rRNA gene were 2.43 log10 copies per oocyst seeded greater when extracting with the PowerViral method versus the PowerSoil method (p < 0.001) and were 2.02 log10 copies per oocyst seeded greater when extracting with the UNEX method versus the PowerSoil method (p < 0.001). Neither the type of sludge (REC or RAS) nor the sample extracted (e.g., A, B, C) were significant predictors of detected gene copies per oocyst seeded (p > 0.05). All no-template controls and extraction blanks performed as expected. The C. cayetanensis pooled standard curve slope was −3.50, intercept was 38.0, and R2 was 0.998.
3.8. Comparison of extraction method nucleic acid concentration and purity
Sludge extract nucleic acid concentrations and purity ratios are presented in Table S5. For REC sludge, one-way ANOVA demonstrated that DNA purity was significantly different for at least one of the extraction methods (p = 0.0001). For RAS sludge, there was no significant difference in the deviation from pure DNA by extraction method (p = 0.09). Statistical comparisons of nucleic acid concentration were not performed, as the input amounts for the extraction methods were not equivalent for all three extraction methods.
4. Discussion
The PowerViral and UNEX extraction methods resulted in similar recovery of C. cayetanensis nucleic acid in tap and wash water, while the PowerViral method resulted in greater recovery from surface water. However, the PowerViral extraction method resulted in greater recovery of S. enterica and MNV than the UNEX extraction method across all water types. For sludge samples, both the PowerViral and UNEX resulted in better recovery of C. cayetanensis than the PowerSoil method. Several factors could have contributed to the performance differences between methods, including microbe characteristics, sample type, and efficacy of cell lysis and inhibitor removal.
Extraction recovery was highly variable based on sample type, with surface water proving the greatest challenge. We evaluated method robustness for water samples in two ways. The first was simply evaluating different types of water samples with different type and levels of organic matter and potential inhibitory substances. The second was adding organic matter by increasing the input volume from 200 μL to 375 μL. While PowerViral provided consistent detection of all microbes, C. cayetanensis recovery from surface and tap water was lower with the 375 μL input volume. This suggests that the PowerViral bead beating process may have been hindered by the additional organic matter or that the additional volume adversely affected the bead to sample ratio. It is important to note that the manufacturer-specified input volume is 200 μL. The PowerViral volumes and processes were modified accordingly for the 375 μL input volume, but additional beads were not added.
UNEX method recovery was greatly affected by the sample matrix, with consistent detection of all microbes in tap water, inconsistent detection of MNV in wash water, and no detections of any of the microbes in surface water. Although the larger 375 μL input volume had no effect on UNEX recovery for tap water, we did not study the effect of scaling up the UNEX to the designed maximum input volume of 750 μL. The overall performance of the UNEX method suggests it may be most effective for use with low inhibitory substance water matrices such as potable water.
Varying recovery between different cell lysis methods could also explain some of the performance differences between the methods. Cell lysis was facilitated by lysis buffer, heat (for UNEX), and bead beating. Bead beating for the UNEX and PowerViral extractions was performed on the FastPrep instrument, ensuring the same bead beating force for these extractions. However, bead beating for the PowerSoil method was performed on a vortexer instead of a bead beater. The poor recovery of C. cayetanensis using the PowerSoil method compared to the UNEX and PowerViral methods highlight the importance of bead beating for these organisms. The lower shearing force of the vortexer compared to a bead beater could have resulted in incomplete lysis of the C. cayetanensis oocysts. For bacteria and viruses, lysis buffer alone may be sufficient to lyse cells and virions, but physical disruption by bead beating is critical for chemically tolerant organisms such as C. cayetanensis (Cama and Ortega, 2024; Qvarnstrom et al., 2018a).
Extraction performance for sludge samples varied greatly by extraction method and sample type. While detection rates were nearly 100 % for all extraction methods, the percent recovery of the PowerSoil method was markedly lower than either the PowerViral or UNEX methods. The RAS sample matrix, with a higher proportion of organic matter, presented a greater challenge to the PowerSoil kit than the other extraction methods. Mean PowerSoil percent recovery from RAS sludge was 6-fold lower than for REC sludge.
The poor performance of the PowerSoil method may be due, in part, to ineffective removal of qPCR inhibitors. The PowerSoil method produced a higher proportion of inhibited extracts than the PowerViral and UNEX methods. The PowerViral and PowerSoil methods utilize the same flocculation procedure for inhibitor removal (Braid et al., 2003). In contrast the UNEX inhibitor removal process utilizes polyvinylpolypyrrolidone (PVPP) to bind humic acids as nucleic acid is passed through the PVPP columns. Despite the differences between the PowerViral and UNEX methods, our results demonstrate that each of these methods adequately remove inhibiting substances from sludge samples. This study was subject to several limitations. The microbial seed quantities were not directly quantified. Instead, seed quantities were calculated mathematically based on the dilution factors of the microbial stock concentrations. Therefore, differences in performance between methods could have been due, in part, to variations in seed quantities between the extracts. Extractions were performed in triplicate to help overcome this potential limitation, and our data show low Cq standard deviation values, which suggests that differences in method performance are not likely due to seeding variation. This was a small study and the ability to do statistical analyses was limited in some cases. The impact of inhibition on the qPCR results for water sample extracts could not be assessed, as IACs were not included in water extract qPCRs. While IACs provided data on how inhibitory the sludge extracts were to qPCR, it was not possible to assess how the individual aspects of the extraction processes facilitated or hindered nucleic acid recovery and inhibitor removal (e.g. filter clogging limiting DNA binding, insufficient elution from filter, inhibitors overwhelmed removal system, etc.).
5. Conclusions
Data from this study can help inform selection of extraction methods for use in the detection of food safety related microbes from environmental samples including tap water, irrigation water, and sewage sludge. The following conclusions from this study can be used to inform method selection:
The PowerViral extraction method resulted in consistent detection across water sample types.
The UNEX extraction method resulted in inconsistent detection across water types.
Surface water presented the greatest challenge to UNEX performance.
Bead beating is critical for recovery of C. cayetanensis, and vortex-beating may not be sufficient to lyse oocysts.
A heated lysis buffer incubation may improve UNEX recovery for some organisms.
The method-specified bead beating beads performed as well or better than alternative bead types.
The PowerViral and UNEX methods outperformed the PowerSoil method for sludge samples.
Nucleic acid purity was a better indicator of method performance than yield for water samples, but there was no correlation between purity or yield and method performance for sludge samples.
Supplementary Material
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://doi.org/10.1016/j.mimet.2025.107195.
Acknowledgements
We thank David Holcomb for assistance with statistical analyses and manuscript review. The use of trade names and names of commercial sources is for identification only and does not imply endorsement by the Centers for Disease Control and Prevention or the U.S. Department of Health and Human Services. The findings and conclusions are those of the authors and do not necessarily represent those of the Centers for Disease Control and Prevention.
Funding
This research received no external funding.
Footnotes
CRediT authorship contribution statement
Amy Kahler: Writing – original draft, Methodology, Investigation, Conceptualization. Jessica Hofstetter: Writing – original draft, Methodology, Investigation, Formal analysis, Conceptualization. Camila Rodrigues: Writing – review & editing. Mia Mattioli: Writing – review & editing, Supervision, Formal analysis, Conceptualization.
Declaration of competing interest
The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
Data availability
Data will be made available on request.
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Supplementary Materials
Data Availability Statement
Data will be made available on request.