Environmental DNA (eDNA) once shed can exist in numerous states with varying behaviors including degradation rates and transport potential. In this study we consider three states of eDNA: 1) a membrane-bound state referring to DNA enveloped in a cellular or organellar membrane, 2) a dissolved state defined as the extracellular DNA molecule in the environment without any interaction with other particles, and 3) an adsorbed state defined as extracellular DNA adsorbed to a particle surface in the environment. Capturing, isolating, and analyzing a target state of eDNA provides utility for better interpretation of eDNA degradation rates and transport potential. While methods for separating different states of DNA have been developed, they remain poorly evaluated due to the lack of state-controlled experimentation. We evaluated the methods for separating states of eDNA from a single sample by spiking DNA from three different species to represent the three states of eDNA as state-specific controls. We used chicken DNA to represent the dissolved state, cultured mouse cells for the membrane-bound state, and salmon DNA adsorbed to clay particles as the adsorbed state. We performed the separation in three water matrices, two environmental and one synthetic, spiked with the three eDNA states. The membrane-bound state was the only state that was isolated with minimal contamination from non-target states. The membrane-bound state also had the highest recovery (54.11 ± 19.24 %), followed by the adsorbed state (5.08 ± 2.28 %), and the dissolved state had the lowest total recovery (2.21 ± 2.36 %). This study highlights the potential to sort the states of eDNA from a single sample and independently analyze them for more informed biodiversity assessments. However, further method development is needed to improve recovery and reduce cross-contamination.

      Adsorbed DNA State: Sheared salmon sperm DNA (Invitrogen, Waltham, MA) was diluted to 100 ng/µL in 6 mL nuclease-free molecular grade water (Sigma-Aldrich, St. Louis, MO) in a 15 mL tube with 300 g (50 mg/mL) montmorillonite clay K10 (Fluka, Buchs, CH). One no-adsorbent control tube was created by diluting salmon DNA to 100 ng/µL in 1 mL nuclease-free molecular grade water, but with no clay. The tubes were shaken at 600 rpm for 48 hours. Previous work demonstrated that most of the DNA adsorption in similar concentrations reached equilibrium well within this timeframe (Kirtane et al., 2020). At 48 hours, the tube with the salmon DNA and clay was centrifuged at 4500 xg for five minutes and the supernatant was separated from the pelleted clay with a pipette. The pelleted clay was then washed using 6 mL nuclease-free molecular grade water by vortexing followed by centrifugation at 4500 xg for five minutes and the supernatant was separated to remove any non-adsorbed salmon DNA. This wash process was repeated one more time to remove any non-adsorbed salmon DNA. Finally, 4.5 mL of nuclease-free molecular grade water was added to suspend the clay pellet to create the adsorbed eDNA state spike. The control tube and all the supernatants from each washing step were stored in independent tubes at -20 ˚C.

Dissolved DNA State: DNA from ten (~0.25 g each) pieces of store-bought chicken breast was extracted using the DNeasy Blood and Tissue Kit (Qiagen, Hilden, Germany) according to the manufacturer’s protocol. Each extraction was eluted in 200 µL Buffer AE. The ten extractions were then combined and vortexed to create the dissolved DNA spike.

Membrane-bound DNA state: Mouse skin cells from cell line B16-F10 derived from mouse C57BL/6J (Jackson Laboratories, ME, USA) were resuspended in Dulbecco's Modified Eagle's Medium (ThermoFisher Scientific, Waltham, MA) containing 10 % Fetal Bovine Serum (ThermoFisher Scientific, MA) and 1 % Penicillin-Streptomycin (10,000 U/mL) (ThermoFisher Scientific, MA) in a 15 mL tube. Cells were spun at 125 x g for 5 minutes and the cell pellet was resuspended in 10 mL growth media and seeded into a tissue culture dish (TPP, Horsforth, UK). Cells were incubated for 10 days at 37 °C, 5 % CO2, and 95 % humidity to let them attach and recover to a concentration of 1x10⁶ cells/m to 20 million cells counted using an automated Cell Counter System (Countess TC20, Biorad, Hercules, CA) which assessed cell viability via trypan blue exclusion. These cells were spun down at 125 x g for 5 minutes in a 50 mL tube and resuspended in 20 mL (1x10⁶ cells/mL) of fresh growth media and used as a spike within 6 hours. The tube was centrifuged at 125 x g for five minutes to pellet the cells. Before spiking the cells, the supernatant was removed using a pipette and discarded. The pellet was then washed to remove any dissolved DNA by resuspending the pellet in 30 mL of Phosphate Buffer Saline (PBS) solution (0.137 M sodium chloride, 0.0027 M potassium chloride, 0.01 M sodium phosphate dibasic, 0.0018 M potassium phosphate monobasic, pH = 7.4). This was followed by centrifuging at 125 x g for five minutes, and the PBS supernatant was discarded as before. The cells were then resuspended in 50 mL of PBS to create the membrane-bound DNA spike.

Experimental procedure for state spiking 

A total of three water matrices were used in the experiment: Milli-Q tap water (from Independent Q-POD® ultrapure water dispensing unit (Merck, Darmstadt, Germany), water from Lake Zurich, and water from Sihl River. Ten liters of water near Lake Zurich outlet (47°21'59.2"N 8°32'39.7"E ) and Sihl river (47°22'35.8"N 8°32'07.4"E) were collected on October 7, 2021, in the morning of the experiment and transported to the lab within 1 hour. At the lab the pH, turbidity (absorbance), and temperature (˚C) of the water matrices were tested using a HI-98194 multiparameter probe (Hanna Instruments, Woonsocket, RI) (Table 1). Three replicates for each water type were created for five treatments (Figure 2). The treatments consisted of spiked DNA from one of each state (i.e., membrane-bound DNA, adsorbed DNA, and dissolved DNA), one where all three states combined were combined, and a control with no spiked DNA (Figure 2). For each treatment, the desired state/s were spiked into 50 mL of water matrix. The volume of spiked states was 500 µL for membrane-bound DNA, 100 µL for adsorbed DNA-bound clay solution, and 50 µL for dissolved DNA (Figure 2). These spike volumes were chosen for experiment logistical reasons. Specifically, to reduce the chance of accidental double-spiking and contamination as a different set of pipettes and tips were used to spike each state/species. This difference in spike volume is not expected to impact the results of the experiment because the concentration of each individual eDNA state was constant throughout the experiment. The water was then filtered through a 0.22 µm Isopore polycarbonate filter (GTTP02500, Millipore, Burlington, MA) in 25 mm Swinnex filter holders (Millipore, Burlington, MA) using a 50 mL syringe (Figure 1). 15 mL of the filtrate was transferred to a 50 mL falcon tube for dissolved DNA extraction. After filtration, air was passed through to remove any residual water and the filter was immediately removed from the housing and placed in a 1.5 mL tube with 600 µL of phosphate buffer (0.12 M Na₂HPO₄, 0.12 M NaH₂PO₄, pH = 9) and shaken at 400 rpm for 20 min to desorb adsorbed DNA (Figure 1). The tube was then centrifuged at 13,000 rpm for 2 min. The supernatant was aspirated with a pipette and stored in a separate 1.5 mL tube. The tube with the supernatant was used to extract the adsorbed DNA, while the tube with the filter and pellet was used to extract the membrane-bound DNA. All three fractions (filtrate, supernate, and filter with pellet) were immediately frozen at -20 ˚C until DNA extraction.

Table 1: Characteristics of different source water matrices used in this study

 DNA extraction methodologies for state separation

The three fractions of the DNA were extracted using three methods chosen specifically to isolate the desired state of eDNA (Figure 1). The dissolved DNA in the filtrate was concentrated using ethanol precipitation, the membrane-bound DNA on the filter and in the pellet was extracted following the lysis step using phenol-chloroform-isoamyl purification and concentrated using ethanol precipitation, and the adsorbed DNA in the phosphate buffer was extracted using a magnetic bead extraction protocol. One negative control was included in every batch of extractions for each method (N = 9).

Ethanol precipitation

15 mL of filtrate was used for the isolation of dissolved DNA using ethanol precipitation. Samples were thawed and 1.2 mL of 5M sodium chloride and 33 mL of absolute ethanol (200-proof) were added to the tube. The tube was vortexed and incubated overnight at -20 °C. The tubes were then centrifuged at 10,000 xg at 4 ˚C for one hour. The supernatant was discarded. 5 mL of 75 % ethanol was added, inverted by hand ten times, and centrifuged at 10,000 xg for 30 mins. The supernatant was discarded, and the pellet was air-dried for 30 minutes. The pellet was then dissolved in 100 µL TE buffer which was then passed through the ZYMO Onestep PCR inhibitor removal kit and stored in 1.5 mL tubes at -20 °C until molecular analysis. This inhibitor removal step was used only for dissolved DNA samples extracted with the ethanol precipitation method.

Phenol-chloroform-isoamyl extraction 

The membrane-bound DNA from the filters was extracted using a phenol-chloroform-isoamyl (PCI) protocol (Deiner et al., 2015). We added 700 µL of Longmire Lysis Solution (100 mM Tris pH 8.0, 0.5 mM EDTA, 0.2% SDS, 200 mM NaCl) and 12 µL Proteinase K (40 mg/mL) to each of the 2 mL tubes containing the filters. The tubes were gently vortexed prior to overnight incubation at 56 °C to facilitate cell membrane lysis. After the incubation, the lysate was transferred to a new sterile 2 mL tube with a pea-sized volume of grease (high vacuum, Dow Corning®). We then added 550 µL of PCI (25:24:1, Sigma, buffered pH8.0) to all tubes followed by shaking at 20 °C at 1,000 rpm. The tubes were then centrifuged at 10,000 xg for five minutes. The supernatant was transferred to another new sterile 2 mL tube with a pea-sized volume of grease to which we added 550 µL of CI (24:1, Sigma). This tube was also shaken for 5 min at 1000 rpm followed by centrifugation at 10,000x for 5 min. The supernatant was transferred to new 2 mL tubes (without grease) containing 44 µL of 5M NaCl and 1,100 µL of 200-proof ethanol and incubated at -20 °C overnight. The incubated tubes were centrifuged for 30 min at 10,000 x at 4 °C. The supernatant was carefully pipetted out and the pellet was washed twice with 75 % ethanol. The pellet was then allowed to air dry and eluted in 100 µL of TE buffer until molecular analysis.

Magnetic bead extraction 

A magnetic bead extraction was used to extract and purify formerly adsorbed DNA in phosphate buffer using a version of Powersoil^® DNA isolation protocol (Qiagen, Hilden, Germany) using homemade reagents (Sepulveda et al., 2019). The 600 µL of supernatant phosphate buffer containing the desorbed DNA was pipetted into a new 2 mL tube, ensuring the filter or the pellet at the bottom of the tube was not disturbed in the process. We then added 100 µL of protein precipitation solution and inhibitor flocculation solution and vortexed for ten seconds. The tubes were then placed in the freezer at 20 °C for 20 minutes. The tubes were removed and vortexed for ten seconds before centrifugation at 10,000 xg for five minutes. The supernatant was transferred to a new tube with 100 µL of 20% Sera-Mag SpeedBead Carboxylate modified magnetic beads (GE Healthcare Life Sciences, Pittsburgh, PA) in hybridization buffer. The tube was gently mixed by inversion and another 100 µL of hybridization buffer was added. The tube was gently mixed by inversion (10 x) and incubated at room temperature for ten minutes. The tube was then placed on a magnetic rack on a shaker (400 rpm) and shaken until all the beads migrated to the magnet (~ 20 minutes). The supernatant was then pipetted out without disturbing the magnetic beads. Two wash steps were performed where 1 mL of 75 % ethanol was added to the tube. The tube was then removed from the magnetic rack and vortexed for ten seconds, placed back onto the magnetic rack, and shaken until all the beads migrated to the magnet (~ 5 min). The ethanol was then pipetted out without disturbing the magnetic beads. The tubes were removed from the magnetic rack and air-dried for 20 minutes. The beads were then suspended in 100 µL TE buffer and pipette mixed until in solution and incubated at room temperature for ten minutes. The tube was then placed back onto the magnetic rack for 5 minutes and the TE buffer eluate was pipetted out and passed through a 2 mL EconoSpin® Mini Spin column (Epoch, Fremont, CA) by centrifuging at 10,000 xg for one min to remove any residual magnetic beads in the solution and stored at -20 °C until molecular analysis.

Development of target-specific primers and TaqMan hybridization probes

We designed a multiplex quantitative PCR (qPCR) with four parallel assays to be run on the Roche 480 light cycler (Roche, Basel, Switzerland). Compatible fluorescent dyes (FAM, VIC, TexasRed, and CY5) were selected as recommended by the PrimeTime Multiplex Dye Selection tool (web tool available from IDT DNA). Reference sequences for primer design were obtained from GenBank (Clark, Karsch-Mizrachi, Lipman, Ostell, & Sayers, 2016) (Table S1). To detect and quantify mitochondrial eDNA from mice (Mus musculus) and chicken (Gallus gallus) we designed TaqMan® qPCR assays targeting the mitochondrial NADH dehydrogenase subunit 2 (ND2) gene, a well-established phylogenetic marker in vertebrates. As a nuclear marker, the single copy gene TGFb1 coding for the transformation growth factor 1 in mice was selected. Previously designed chum salmon (Oncorhynchus keta) primers for the cytochrome oxidase I gene (COI) were used with a modified TaqMan® probe that did not have the minor grove binder (Homel et al., 2021) (Table 2) to decrease the cost of the probe. The TaqMan assays for detection and quantification of the nuclear Tgfb1 gene in mice and mitochondrial ND2 genes of chicken and mice were designed using the Primer Express Software version 3.0 (Applied Biosystems, Waltham, MA) using default parameters.

Table 2: qPCR assays with their corresponding target genes, represented eDNA state and fluorophores (in bold).

Specificity testing 

All qPCR assays were tested for specificity in-silico and experimentally. The in-silico testing was conducted with NCBI Primer-BLAST tool (Ye et al., 2012) and OligoAnalyzer tool (IDT, Coralville, IA). In Primer-BLAST, the specificity parameters were set to ensure a minimum of three mismatches and at least two mismatches within the last five base pairs of the 3’ end on each primer and probe between the target and non-target organisms used in this study. OligoAnalyzer was used to test the likelihood of dimer formation between the various primers and probes. Using the default “qPCR” parameters we checked that ΔG > -9 kcal/mole ensured a low likelihood of self- or hetero-dimers formation in between any primer and probe combinations. To experimentally test the specificity of the multiplex qPCR we amplified standard curve of a single target in the multiplex reaction setup described below. This was repeated by using each of the four target amplicons independently as template in the multiplex qPCR setup. The resulting data was analyzed for cross-amplification or cross-reporting of targets as only one target should be reported from the multiplex qPCR regardless of all assays being available. The efficiencies of the single-species standard curves were compared to the efficiency of the multiplexed standard curves to ensure reliable quantification. The multiplex qPCR negative controls used throughout the experiment ensured no false positives due to dimer formation.

 qPCR preparation and cycling conditions

The qPCR reactions were performed in 10 µL reactions in 384 well plates on a Roche Light Cycler 480. Each reaction included 5 µL Taqman^TM Multiplex Master Mix (Applied Biosystems^TM), 0.03 µL of each primer at 100 nM, 0.025 µL of each Taqman probe at 100 nM, 1 µL DNA extract, and 3.92 µL of molecular grade water to bring the volume up to 10 µL. For simplex qPCR, the same reaction mixture was used but only one set of primers and probes were added and the volume of molecular grade water was adjusted to keep the reaction volume to 10 µL. After an initial incubation for ten minutes at 95 °C, we performed 40 cycles with a denaturation step for 15 seconds at 95 °C and an annealing/extension step for 30 seconds at 60 °C. For the preparation of all qPCR plates, we used the mosquito® LV pipetting robot (SPT Labtech Ltd, England) for the efficient and accurate preparation of qPCR plates. 

 qPCR quality control and data interpretation

The Light Cycler was calibrated for multiple emission spectra for the multiplex qPCR using a color compensation protocol utilizing the four fluorophores used in this study. We incorporated six replicates of the six-point standard curve on each qPCR ranging from 10^7 copies/reaction to 100 copies/reaction. These standards were made by combining four individual gBlock gene fragments (Integrated DNA Technologies) that represent the target sequences from the four qPCR assays used in this study (Table S2). The qPCR efficiency was calculated using [E = -1+10^(-1/slope)] where E is the qPCR efficiency and the slope is calculated with pooled six-point standard curves from all plates for enumerating copy numbers of the target amplicons. This efficiency and intercept were then used in the quantification of our experimental replicates by converting Cp values to copy numbers. We also used this pooled standard curve to determine the Limit of Detection (LOD) and Limit of Quantification (LOQ) using previously described statistical criteria (Klymus et al., 2020). The LOD is described as the lower standard dilution concentration where 95 % of the replicates demonstrate amplification and the LOQ is described as the lowest standard concentration with a coefficient of variation (CV) value below 35 %. Each qPCR plate also included six qPCR negative control wells with molecular grade water instead of template DNA to identify any contamination in the reagents or during qPCR setup. A random subset of samples (N = 18) using the two environmental water matrices were diluted 1:100 to test for qPCR inhibition.

 eDNA state recovery

Quantification of total eDNA yield

The total eDNA yield of all treatment samples (N = 45), each spike (N = 3), and extraction controls (N = 9) were measured in 384 well plates using reagents from Qubit dsDNA HS Assay Kit (Qubit Digital, London, UK), and analyzed by Spark® Multimode Microplate Reader (Tecan, Männedorf, CH). We used a seven-point standard curve at concentrations of 0, 0.1, 0.2 0.5, 1, 5, and 10 ng/µL by diluting the 10 ng/µL standard provided by the manufacturer. Each reaction well consisted of 48 µL of Qubit HS 1x reaction mixture and 2 µL of DNA standard, sample, spike, or control. Accurate pipetting was facilitated by a mosquito® LV pipetting robot (SPT Labtech Ltd, England). All standards and samples were analyzed in triplicate.

State-specific isolation efficiency and percent recovery      

To calculate the extraction recovery of the DNA added to each replicate of the experiment, first, the spiked DNA for each of the three states was quantified. The concentration of target DNA recovered from the experimental samples was then quantified and compared with the spike to calculate the percent recovery of each DNA state.

For quantification of the dissolved state spike, 1 µL of the dissolved DNA spike was directly analyzed in triplicate using simplex qPCR to enumerate the dissolved DNA target copies per uL. This value was multiplied by the spike volume (50 µL) to calculate the total copy number of the spiked DNA. Using Equation 1, percent recovery was calculated where C_s is the number of DNA copies spiked in the dissolved state (per 15 mL), C_i is the initial spike concentration, V_tot is the total volume of the filtrate (50 mL), and V_s is the volume of the filtrate analyzed (15 mL). This volume correction is necessary as only 15 mL of filtrate could be used in the extraction of dissolved DNA using the ethanol precipitation method. This volume adjustment was needed as ethanol precipitation in 50 mL tubes can utilize a maximum of 15 mL of filtrate to maintain a 1:2 ratio of filtrate to ethanol.

For quantification of the membrane-bound DNA spike, 2 mL of mouse cell spike solution was frozen at -20 °C. Three replicates of the spike were created by extracting 50 µL of the saved spike using the same phenol-chloroform-isoamyl protocol described above and analyzed with qPCR to enumerate the membrane-bound DNA target copies spiked into the experiments.

To calculate the percent recovery of the membrane-bound DNA from experimental samples we used equation 2, where C_s is the concentration of mouse DNA (mitochondrial or nuclear) detected in experimental samples (per 50 mL) and C_i is the initial spike concentration of mouse DNA.

 The concentration of adsorbed DNA spike was calculated using equation 3, where C_sp is the concentration of spiked adsorbed DNA (copies DNA/mg clay), C_i (copies/µL) is the initial concentration of salmon DNA solution, V_i (µL) is the initial volume of salmon DNA solution, C_sup (copies/µL) is the concentration of salmon DNA in the supernatant after 48 h of adsorption, C_w1 (copies/µL) and C_w2 (copies/µL) are concentrations of salmon DNA in the supernatant of the first and second wash step, V_w (µL) is the volume of water used in the wash steps and M_c (mg) is the mass of montmorillonite clay added to the adsorption reaction. Finally, the adsorbed DNA was suspended in 3 mL of nuclease-free molecular-grade water to achieve a final adsorbed DNA spike concentration of 100 mg/mL. Concentrations of target DNA copies in C_i, C_sup, C_w1, and C_w2 were quantified in triplicate using simplex qPCR.

To calculate the extraction recovery of the adsorbed DNA in each sample, equation 4 was used where Cs is the concentration of salmon DNA detected from experimental samples (per 50 mL) and C_sp is the theoretical adsorbed DNA spike concentration calculated in equation 3.

Statistical analyses 

We conducted an analysis of variance (One-way ANOVA) using the extraction methods, water matrix type, and the state of spiked DNA as dependent variables, and percent recovery as the independent variable. We used a one-way ANOVA to test whether a state isolation protocol was able to enrich the target state. This test was repeated for each of the three state isolation protocols used. A one-way ANOVA test was also used to test if any state isolation protocol was able to outperform others with respect to the percent recovery of the target state. Finally, another one-way ANOVA was also used to determine whether a given water matrix had a significant impact in determining the success of eDNA state isolation based on the increased recovery of a target state. We used a student's t-test to evaluate the effect of spiking a given state individually in a sample or multiple states spiked together. The t-test was also used for testing the variation in the recovery of mitochondrial and nuclear DNA recovery from spiked mouse cells. We conducted the Shapiro-Wilk test of normality, and Levene’s test to check the homogeneity of variances to ensure our data met the assumptions of parametric t-tests and ANOVA. All ANOVA tests that rejected the null (α = 0.01) were followed up with Tukey’s post hoc test to identify what dependent variables caused a significant difference. All data analysis was conducted in R version 4.1.3 using the package tidyverse package (Wickham et al., 2019).