Original stock suspensions of purified and viable C. parvum oocysts (Iowa isolate–Cat# P102C@1 × 10/9) were obtained from Waterborne Inc. (New Orleans, LA). The concentration of the stock oocyst suspension (10⁹ oocysts in 50 mL) was determined using a Neubauer hemocytometer at a magnification of 200×. The stock of C. parvum oocysts was suspended in 50 mL de-ionized (DI) water with penicillin, streptomycin, gentamicin and amphotericin B. The C. parvum oocysts were stored in the dark at 4°C for 18 months. Prior to each experiment, the stock oocyst suspension was mixed in a vortexer at 3,000 rpm for 15 min to disperse the C. parvum oocysts that had settled to the bottom of the 50 mL centrifuge tube. This mixing process was designed to optimize the uniformity and consistency of the inoculum used to prepare the experimental oocysts concentration for use in the soil column experiments.

The four different soils used in this study were collected in bulk form from fallow and cultivated pasture fields in the U.S. states of Illinois and Utah. The difference in the textural characteristics of these soils resulted in their classification as loamy sand and sandy loam based on their particle size analyses. One of the loamy sand soils collected in Kankakee County, IL is classified as a coarse-loamy, mixed, superactive, mesic Typic Endoaquoll belonging to the Gilford series which is a very deep, poorly drained or very poorly drained soil, and that formed in loamy over sandy sediments on outwash plains, near-shore zones (relict), and flood-plain steps. The Gilford soil series has a negligible potential for surface runoff, with a saturated hydraulic conductivity that is high in the upper part and very rapid in the lower part. Its soil permeability is moderately rapid in the upper part and rapid in the lower part. The other loamy sand soil, from Kankakee County, belongs to the Sparta series. The Sparta series is classified as a coarse-loamy, mixed, superactive, mesic Typic Endoaquoll, which is a very deep, and excessively drained soil. Formed in sandy outwash that has been reworked by wind these soils have a saturated hydraulic conductivity ranging from 10.00 to 100.00 micrometers per second. The Greenson sandy loam soils collected in Cache County (Utah) are classified as fine-silty, mixed, superactive, mesic Oxyaquic Calcixerolls; they are very deep, somewhat poorly drained, or moderately well-drained soils that formed from lacustrine deposits. Found on low lake terraces, the Greenson soil series is either somewhat poorly drained or moderately well-drained with a low-to-medium surface runoff, a slow-to-moderate permeability (moderately low to high saturated hydraulic conductivity), and a generally high organic matter content, from 3 to 9%. Collected in Cache County, UT, the Lewiston sandy loam soil is classified as coarse-loamy, mixed, superactive, mesic Aquic Calcixeroll that is a very deep, somewhat poorly drained soil formed in lacustrine sediments. Also found on lake terraces, the soil is somewhat poorly drained, has slow or very slow runoff and moderate permeability. The soil organic matter content ranges generally from 1 to 3%.

The physicochemical characteristics of these soils, presented in Table 1, were determined by the Utah State University Analytical Laboratories (USUAL). Soil texture and particle size analyses were performed using the hydrometer method (Gee and Bauder, 1986). Total carbon analyses were performed using a Primac^SLC Analyzer (Skalar Inc., Buford, GA, USA). Saturated soil paste extracts were prepared for the electrical conductivity and pH measurements (Rhoades, 1996). The cation exchange capacities of the soils were measured using the sodium acetate/ammonium acetate replacement method. Soil elements and ions were determined by ion chromatography using a Dionex ICS-3000 Ion Chromatography (IC) system. The soils were air-dried at 37°C, and sieved (2 mm), which were then stored in lidded buckets at room temperature until used.

All solutions were prepared with deionized water (18 MΩ·cm⁻¹ resistivity; Milli-Q, Millipore Corp, Bedford, MA). For all flow-through column experiments, the artificial rainfall solutions contained 1 mM KCl as a background electrolyte. Potassium bromide (KBr) was used as a conservative tracer and was measured using an Orion™ Bromide Electrode (ThermoFisher Scientific, Waltham, Massachusetts, USA). The preparation of the C. parvum oocysts inoculum consisted of adding about 1.5 × 10⁶ C. parvum oocysts (quantified and enumerated using the qPCR method) from the stock oocysts suspension to 53 mL of artificial rainfall/background solution (1 mM KCl) that was spiked with 10 mM KBr tracer. Fifty of the 55 mL that constituted the inoculum were used in the flow-through column experiments and 5 mL were kept for further quantification of the initial input concentration (C₀) of C. parvum oocysts applied to each column. The inoculum applied to the surface of each column contained a measured concentration of C. parvum oocysts of about 2.85 × 10⁴ mL⁻¹, corresponding to a total of 1.56 × 10⁶ ± 575,000 oocysts.

The columns used in these experiments were composed of plastic rings with dimensions of 20 cm in length and 9.5 cm in internal diameter. A ring 5 cm in height was placed at the bottom of the column, followed by the placement of 2-cm rings to achieve a total column height of 30 cm, of which 20 cm were used for the experiments. Rings were compressed together with top and bottom column holders, which were held with four rods positioned parallel to the length of the column on the outside perimeter of the rings, and bolts to prevent sliding. The rods were bolted through a screen section of a funnel that also had an additional mesh to prevent soil particles from moving through. The column was then clamped to an individual stand. A separate clamp allowed connection of the bottom part of the funnel below the column and an additional rod built in the stand allowed connection of a nozzle above the column. The soil was packed into each column. To achieve a uniform and homogeneous soil packing and a soil bulk density of 1.5 g·cm⁻³ of soil, the total amount of soil (2,125 g) used to form the soil column was poured in three equal amounts and compacted with a rod. Four experimental columns were run simultaneously. One pump equipped with four cartridges dispensed the artificial rainfall solutions to the four nozzles, and a single air tube with brass couplings was used to pump air into these four nozzles. Rainfall was applied to the surface of each column using an individual nozzle (XA nozzle system ¼, 303 from BETE Fog Nozzle Inc., Greenfield, MA). The nozzle was connected to the water on one side through a peristaltic pump (Cole-Parmer, Vernon Hills, Illinois, USA) and to the air on the other side. The flow rate and the air pressure at the nozzle were adjusted at the pump and through a Parker Watts miniature precision regulator gauge (1/4 in; 60 psi; Parker Hannifin Corp., Cleveland, OH) to ensure that the nozzle spray covered the entire soil surface. The height of the nozzle above the soil surface was 10 cm.

Each soil column experiment was performed in duplicate. Once the experimental columns were loaded with soils and placed in their stands, rainfall simulation was initiated with the artificial rainfall. Subsequent to achieving steady state and equilibrium conditions and a determination of constant outflow, C. parvum oocysts were released at the soil surface by pouring the 50 mL inoculum into the column. No rainfall was applied during the duration of the inoculum release at the soil surface until the inoculum had infiltrated. Rainfall was then resumed. In soil column experiments characterized by an occurrence of water ponding at the soil surface, the rainfall was discontinued until the ponded water had infiltrated and was then resumed. The soil flow velocities ranged from 0.09 to 0.16 cm·h⁻¹ with all of the column outflows collected in 50 ml centrifuge tubes. After about 6 pore volumes (PV) had passed through the columns, the rainfall was stopped, the column set-up was dismantled, and the columns were sliced with a thin metal sheet. A total of 10 column layers were sliced: 1 cm for the top layer, 2 cm for each of the following seven rings, with the remaining 5 cm ring divided in two, to obtain two layers of approximately 2.5 cm each.

Each soil layer was sampled to determine the water content and the concentration of C. parvum oocysts in the soil profile. To establish the water content in the soil layers, three wet soil samples of 5 g each were collected randomly in each soil layer. These wet soil samples were then placed in aluminum foil cup holders, weighed, and oven dried at 105°C for 24 h. The weight of the dried samples was then recorded and the water content was established gravimetrically.

All of the soil remaining in each layer after sampling for water content was collected for purposes of measuring the concentration of C. parvum oocysts in the soil. 25 g of wet soil was placed in each of between 9 and 11 50 mL plastic centrifuge tubes, depending on the layer. The method developed by Koken et al. (2013) was used to isolate C. parvum oocysts from the soil. The C. parvum oocysts attached to soil particles were released by the addition of 20 mL of Tween 80 at two critical micelle concentrations (CMC) to each of the 50 mL centrifuge tubes, which were then fixed in a in a rotational shaker, perpendicular to the axis of rotation, for 24 h. Two tubes from each soil layer were selected to undergo the C. parvum oocyst isolation procedure, while the remaining tubes were stored. In each of the selected tubes, the soil was underlaid by 10 mL of cold sucrose solution with a density of 1.18 g·mL⁻¹. These tubes were then centrifuged at 2,500 × g for 15 min to construct a sucrose concentration gradient and retain the C. parvum oocysts. The resulting supernatant was then transferred into a new 50 mL conical-bottomed centrifuge tube and subjected to centrifugation at 2,500 × g for 15 min. Thereafter, the supernatant was removed, leaving 4 mL in the tube. A first wash was performed by vortexing the tube, re-suspending the pellet by adding 35 mL of deionized water, followed by centrifugation at 2,500 × g for 15 min. A second wash was conducted following the procedure described in the first wash. For the third wash, supernatants were removed and the remaining 4 mL of the two tubes from each layer were vortexed and transferred into one 15 mL centrifuge tube. Deionized water was added to reduce the final volume to 12 mL. These 15 mL tubes were centrifuged at 2,500 × g for 15 min. In each case, the supernatant was removed to leave 3 mL of pellet in the 15 mL tube, and the pellet was re-suspended by vortex. All centrifugations involving 50 and 15 mL tubes were performed using a refrigerated centrifuge (Model 5810R, Eppendorf AG, Hamburg, Germany). A 300 μL aliquot was taken from the 3 mL, and placed in a microcentrifuge tube to which 900 μl of TE buffer was added. After vortexing briefly, the microcentrifuge tube was centrifuged at 14,000 × g in a microcentrifuge (Model 5424, Eppendorf AG) for 5 min. The supernatant was then removed, leaving a 0.5 mL sample for DNA extraction and PCR analyses.

Soil effluent samples were collected in 50 mL centrifuge tubes and the method developed by Koken et al. (2013) was again used to concentrate C. parvum oocysts from the soil. Selected tubes were centrifuged at 2,500 × g for 15 min to concentrate the C. parvum oocysts in the pellet. The supernatants were discarded and the remaining 5 mL containing the pellets were transferred into 15 mL centrifuge tubes. Deionized water was added to each 15 mL tube to bring the final volume to 8 mL. The tubes were then spun at 2,500 × g for 15 min. The supernatant was removed and the remaining 1.7 mL were transferred to microcentrifuge tubes. These tubes were centrifuged at 14,000 × g for 5 min. The supernatants were removed, leaving 0.5 mL, to which 900 μL of TE buffer was added. After being vortexed, the microcentrifuge tubes were centrifuged at 14,000 × g for 5 min, and the supernatant was removed to leave 0.25 mL of pellet to which 0.25 mL of TE buffer was added, yielding a 0.5 mL of sample for DNA extraction and PCR analyses. All centrifugation processes involving 50 mL and 15 mL tubes were performed using an Eppendorf 5810 R refrigerated centrifuge. All centrifugations involving microcentrifuge tubes were performed in an Eppendorf 5424 microfuge.

The molecular biological methods used for the detection and enumeration of C. parvum oocysts, which include DNA extraction and qPCR analyses, using the procedures described in detail by Koken et al. (2013). Briefly, 7-point standard curves were prepared with DNA from 10,000; 1,000; 100; 10; 1; 0.1; and 0.01 oocysts (per 5 μL volume), which were obtained by pooling two samples from the C. parvum oocyst stock suspension. Each standard curve was used during the analysis of six microtiter plates containing samples obtained from soil and leachates. Six plates were run, with each plate containing its own set of standards for the standard curve. After DNA isolation, each sample was resuspended in 30 μL of DNA elution buffer (Cat# D3004-4; Zymo Research, Irvine, CA). A 5 μL sample DNA was used for each qPCR reaction, and each sample was tested in triplicate. Samples were analyzed in a 20 μL reaction volume with 50 cycles per run using a QuantStudio 12K Flex Real Time PCR system (Life Technologies, Carlsbad, CA, USA). Taqman® chemistry was used, and the primers and probe indicated in Koken et al. (2013) were synthesized as PrimeTime® qPCR assays by Integrated DNA Technlogies (Coralville, Iowa, USA) on the basis of GenBank accession number AF190627 by Jothikumar et al. (2008) as follows: 5′-ACTTTTTGTTTGTTTTACGCCG-3′ (forward primer), 5′-AATGTGGTAGTT GCGGTTGAA-3′ (reverse primer) and 5′-FAM-ATTTATCTCTTCGTAGCGGCG-BHQ-3′ (probe). The results were compiled and analyzed using the QuantStudio software (Life Technologies). After analysis, the data from each plate were exported to an Excel spreadsheet. Mean data for each sample (averaging of replicates) were compiled as aliquot data. These values reflected the number of oocyst genomes in 5 μL of each sample. These data were used to calculate the number of oocysts in the entire 30 μL sample. The standard deviation for each sample was also computed.

Results of the transport and retention of C. parvum oocysts in soils were expressed as the percentage of C. parvum oocysts retrieved in the soil leachates and matrices, which reflect the behavior of C. parvum oocysts in soils of different physicochemical properties. All statistical analyses were performed using SAS®Studio software. For the statistical analysis of the transport and retention results of C. parvum in soils, a t-test was applied to determine the least significant difference (LSD) for the percentage of C. parvum oocysts retrieved in the soil leachates and matrices of Sparta, Lewiston, Gilford, and Greenson series soils. Statistical significance was accepted at p < 0.05. Results are expressed in graphs as mean ± standard deviation.

Physicochemical analyses of the soils used in this study indicate that soils in the Greenson series and Lewiston series are sandy loam soils, while the Sparta and Gilford series were characterized as loamy sand soils (Table 1). The clay contents were highest in the Greenson series soil (19.5%), with Lewiston (12.5%) and Sparta (9.5%) series soils having less clay and the Gilford series soil (7.9%) having the least clay. The organic matter content ranged from 1.28% in Lewiston series to 2.91% in Gilford series soils. The lowest pH values were observed in the Gilford (5.2) and Sparta (6.9) series soils, while the highest pH values were reported in Greenson (7.4) and Lewiston (7.5) series soils. The total calcium concentrations ranged from a low of 9.36 mg·kg⁻¹ for the Sparta series, to a high of 42.09 mg·kg⁻¹ for the Greenson series soil. With the addition of artificial rainfall, the highest flow velocity (0.16 cm·h⁻¹) was observed in the Lewiston series soil, while the lowest flow velocity (0.10 cm·h⁻¹) was reported in the Greenson series soil (Table 2).

Tracer tests were used to evaluate the hydraulic properties of the soils. The concentration of bromide ions entering the column, C₀, and in the effluent, C, were used to calculate BTCs as C/C₀ as a function of PV (Figure 1). The non-reactive chemical tracer added to the C. parvum oocysts mixture was measured in every leachate sample as mg·L⁻¹ of leachate. No correlation was observed between the number of C. parvum oocysts in the leachate and the tracer concentration. Additionally, the BTCs of the tracer and the C. parvum oocysts did not display the same behavior. The tracer BTCs in the Sparta, Lewiston, and Gilford experiments presented a monotonic increase, with a C/C₀ value reaching a peak ranging from about 0.13 to 0.20 that occurred at 0.56 to 1 PV, and then a monotonic decrease. Regarding the Greenson series, the tracer BTCs contained two peaks, a first peak at the 0.12 PV and a second peak at 1.5–3.5 PV, possibly indicating a fast and slow convective transport; however this possible dual flow had no effect on the mobility and transport of C. parvum oocysts.

Even though the experiments were conducted under unsaturated conditions, with free drainage at the bottom of the columns, the degree of saturation for the replicates of soil columns of all four soil series is high, always above 70% (Figure 2). The effect of such high saturations on C. parvum oocysts was not determined, but given the size of these oocysts, it is more than likely that their transport in the soils would have been even slower, and their retention more intense had the degree of saturation been significantly lower.

The leachates from all four soil series were analyzed to assess the prevalence and concentrations of C. parvum oocysts. Leachate samples were collected over about 6 PV of each soil column. The numbers of C. parvum oocysts in the soil leachates from two replicates for each of the four soils are represented in the breakthrough curves (BTCs) as C/C₀, where C is concentration of C. parvum oocysts detected in leachates and C₀ is the concentration of C. parvum oocysts detected in the input solution (Figure 1). A visual inspection of the soil columns did not reveal any cracks or holes.

In the Sparta series soil, C. parvum oocysts were not detected in the leachates of either of the replicates (Figure 1). The BTCs of the tracer, however, displayed a fast increase and a tailing decrease. The heights of the tracer BTCs were measured as C/C₀ ranging from 0.17 to 0.20. The peaks of the BTC of the tracer occurred at 0.70 and 0.80 PV.

In the Lewiston series soil, C. parvum oocysts were detected in the soil leachates of both replicates (Figure 1). The peaks of the C. parvum oocysts BTC were observed at 2.35 PV for a C/C₀ value of 0.006 and at 1.86 PV for a C/C₀ value of 0.005, for the first and second replicates (R1 and R2), respectively. The peak of the tracer BTCs occurred at about 1.00 PV with C/C₀ values of approximately 0.16 to 0.18. These results indicate the transport of C. parvum oocysts within the soil matrix and their prevalence in the soil leachates in considerable concentration.

In the Gilford series soil, C. parvum oocysts were present in only a few samples (Figure 1), with C/C₀ values of 0.004 at 0.53 PV and 0.008 at 2.03 PV for R1, and 0.011 at 5.2 PV for R2. The peak of the tracer BTCs reached a C/C₀ value of approximately 0.13 at 0.56 PV and 0.69 PV, respectively for replicates 1 and 2.

In the Greenson series soil, C. parvum oocysts were present in the leachates of both soil columns (Figure 1). Their peaks reached values of C/C₀ of 0.004 at 0.56 PV and 0.007 at 2.68 PV, respectively, for R1 and R2. Two peaks were observed in each tracer BTC; an early and a late peak occurred in each of the replicates. The BTCs of the tracer displayed early peaks at approximately 0.12 for R1, and 0.20 PV for R2. Late and smaller peaks in the BTCs of the tracer were observed between 1.5 and 2 PV in R2, and at approximately 3.5 PV in R1. The occurrence of two peaks demonstrated the presence of fast and slow infiltration processes in this soil series for the tracer; however this dual-flow phenomenon did not induce two different transport behaviors for C. parvum oocysts.

Following the flow and transport experiments in soil, the soil columns were sliced in layers and soil samples were analyzed to establish the spatial distribution and concentration of C. parvum oocysts in the soil profile. The numbers of C. parvum oocysts in the soil matrices from the two replicates for each of the four soils are presented as number of C. parvum oocysts per gram of dry soils (Figure 2).

In the Sparta series soil (Figure 2), C. parvum oocysts were detected in the top layers of the soil profiles of each of the replicates and in the deepest layer of the second replicate (R2). The highest concentration of C. parvum oocysts was detected in the top layer of the first replicate (R1). Although C. parvum oocysts were not detected in the soil leachates, their transport through the soil matrices of Sparta series was established, as they were detected and enumerated in the topmost soil layers of R1 and R2, and the deepest soil layer of R2.

In the Lewiston series soil (Figure 2), C. parvum oocysts were detected in the soil matrices of both of the R1 and R2 soil column replicates. The highest number of C. parvum oocysts in the soil column of replicate 1 was detected in the top layer, with about 2 C. parvum oocysts per gram of dry soil. In contrast, the highest number of C. parvum oocysts in the soil column of R2 was found in the deepest layer, with about 215 oocysts per gram of dry soil. Our results demonstrated the transport of C. parvum oocysts throughout the soil matrix of the Lewiston series soil.

In the Gilford series soil (Figure 2), C. parvum oocysts were found only in the top layers of the soil column in both replicates, whereas in the Greenson series soil (Figure 2), no C. parvum oocysts were detected in the soil column from R1, while only a few oocysts detected in the top layer the soil column from R2.

Throughout the transport experiments, higher numbers of C. parvum oocysts were detected in the soil matrices than in the soil leachates of all four soil series. Although the relative proportions of C. parvum oocysts in the soil matrices and soil leachates from the different soil series varied, the trends of total numbers of C. parvum oocysts recovered from each of the replicates of each soil series differed (Table 3). The highest recovery of the number of C. parvum oocysts in soil matrices and leachates was 2.718% in the soil column of replicate 2 of the Lewiston series soil. The majority of C. parvum oocysts recovered were from the soil matrices in all of our experiments, except in the first replicate of the Greenson series soil, where C. parvum oocysts were not detected in the soil matrices. The distribution of percentage recovery between soil matrices and leachates in this case (Greenson first replicate) was attributed to soil leachate only. Similarly, no C. parvum oocysts were detected in the leachates of either replicate of the Sparta series soil, which resulted in C. parvum oocysts being recovered only from the soil matrices.

The recovery of C. parvum oocysts in the leachates was very low, ranging from 0 to 0.006% (Table 3). C. parvum oocysts were detected in the leachates of three out of four soil types. Overall, the recovery of C. parvum oocysts in leachates was impacted by the soil type. Significance differences (p < 0.05) were detected among the percentages of C. parvum oocysts retrieved in the soil leachates of the Sparta, Lewiston, Gilford, and Greenson series (Figure 3). In sandy loam soils, the highest C. parvum oocyst recovery was observed in leachates from the Lewiston series soil (0.004–0.006%). In loamy sand soils, the highest recovery of C. parvum oocysts was detected in the leachates from the Gilford series soil (0.002–0.004%).

Cryptosporidium parvum oocysts were detected in the soil of all replicates of all four soils, except for the soil from the first replicate of the Greenson series soil. Overall, the C. parvum oocyst recoveries in the four different soils were not significantly different. No significant differences (p < 0.05) were observed among the percentages of C. parvum oocysts retrieved in the soil matrices for the Sparta, Lewiston, Gilford, and Greenson series (Figure 4). C. parvum oocysts recovery in soils ranged from 0 to 2.718%. The highest C. parvum oocyst recovery from sandy loam soils was reported for the Lewiston series soil (0.060–2.718%). For the loamy sand soils, the Sparta series and Gilford series, soils showed C. parvum oocyst recoveries ranging from 0.092 to 0.165% and 0.345 to 0.919%, respectively.

Although numerous studies have been undertaken to investigate the effects of soil type on the mobility and transport of microorganisms (i.e., bacteria and viruses) in soil, research on the fate and transport of protozoan cysts is scarce (Gerba et al., 1975; Bitton and Harvey, 1992). In soil, the mobility of bacteria is controlled by adsorption, straining, and sedimentation processes (Reddy et al., 1981; Gannon et al., 1991a,b; Tan et al., 1992), whereas in the case of viruses, mobility is governed mainly by adsorption as a results of their small size (Tan et al., 1992). For protozoa and their cysts, such as C. parvum oocysts, which have a diameter of 4–6 μm, adsorption, straining, and sedimentation are expected to impact their mobility in a manner similar to that of bacteria (Mawdsley et al., 1996b).

Soil type is a critical parameter that affects the mobility of microorganisms (Bitton et al., 1974; Bashan and Levanony, 1988; Tan et al., 1991; Huysman and Verstraete, 1993; Mawdsley et al., 1996a). The presence of clay and organic matter, in particular, affects the adsorption processes of bacteria and viruses as a result of their large surface area and negative charge (Reddy et al., 1981).

This study has demonstrated that leaching of C. parvum downward through the soil profile and the vadose zone does occur, although the extent of their transport is affected by the soil type. The soil type was a determinant in the transport of C. parvum oocysts through soil and into leachate, as larger quantities with concentrations of C. parvum oocysts found in sandy loam soils (i.e., Lewiston and Greenson series) as opposed to loamy sand soils (i.e., Sparta and Gilford series).

Potential interaction of the C. parvum oocysts with surfaces and interfaces they encounter, particularly the surfaces of soil particles, during the soil infiltration process is possible. Among the soil components, clay particles have the largest surface areas for potential interactions. Consequently, soil with high clay content provides the highest probability and potential for C. parvum oocysts to interact with clay surfaces, which may result in greater adsorption of C. parvum oocysts to soil clay particles (Balthazard-Accou et al., 2014). In soil, most of the organic matter (OM) is generally combined with minerals that consist of clay and silt particles (Cheshire et al., 2000). Therefore, since most of the OM in soil is bound to the clay and silt surfaces, we can postulate that clay-OM and silt-OM complexes may also facilitate the interplay between C. parvum oocysts and OM, as C. parvum oocysts collide with clay surfaces that are coated with OM. Soil that contains high clay and silt contents may enhance and promote a higher degree of interaction between OM present at the surface of minerals and C. parvum oocysts that encounter these surfaces during their transport through soil. Interactions of OM and surfaces of C. parvum oocysts is a complex phenomenon, and previous studies have suggested the influence of OM on the fate and transport of C. parvum oocysts in porous media. In particular, an increase in OM concentration has been found to induce an increase in the breakthrough of C. parvum oocysts in sand and a decrease in the collision efficiency (Abudalo et al., 2010). The adsorption of OM on the surface of C. parvum oocysts has been postulated to be mediated by the presence of the polyvalent cations Ca²⁺ (Dai and Hozalski, 2003).

As observed in transport experiments in the Greenson series soil, the highest combination of clay, organic matter, and Ca²⁺ resulted in the highest transport through the soil matrix and the highest release of C. parvum oocysts in leachates (Tables 1, 3 and Figure 1). These results demonstrate the role that clay particles, organic matter, and Ca²⁺ play in the transport of C. parvum oocysts. These results and proposed description of an enhanced transport process of C. parvum oocysts provided by adsorption of organic matter at the oocysts surface mediated by Ca²⁺ and promoted by the presence of clay are in agreement with similar proposed enhanced-transport phenomena in other contexts. In our study, sandy loam soils have a higher content of clay (12.5 and 19.5% for Lewiston and Greenson series, respectively) than loamy sand soils (9.5 and 7.5% for Sparta and Gilford series, respectively). Consequently, a soil that contains higher concentrations of clay and silt particles offers larger surface areas for C. parvum oocysts to potentially encounter and a higher probability of C. parvum oocysts to interact with these surfaces as they are transported through the soil. The largest CEC values were observed for sandy loam soils Lewiston (115 mmol_c·kg⁻¹) and Greenson (175 mmol_c·kg⁻¹) compared to loamy sand soils Sparta (86 mmol_c·kg⁻¹) and Gilford (840 mmol_c·kg⁻¹). In our experiments, a larger number of C. parvum oocysts were observed in leachates from soils with high clay content (0.004–0.006% oocysts recovered in water in Lewiston and Greenson series, with clay content of 12.5 and 19.5%, respectively) than in leachates from soils with low clay content (0–0.003% oocysts recovered in water in Sparta and Gilford series, with clay content of 9.5 and 7.5%, respectively). Indeed, direct and indirect effects of clay particles, as well as interplay effects of other soil parameters have been identified to significantly govern the fate and transport of C. parvum oocysts. Since clay and silt surfaces are the sites where most of the OM resides and the probability of interaction of C. parvum oocysts with OM increases as the percentage of clay and silt particles increases. As previously reported, when C. parvum oocysts are placed in the presence of elements from natural OM, OM may adsorb onto their surface and increase their negative charges and hydrophobicity which enhance the electrostatic repulsion between the OM-coated C. parvum oocysts and the negatively charged soil particle surfaces that they encountered, and therefore increase their mobility and impact their transport (Mawdsley et al., 1996a; Ongerth and Pecoraro, 1996; Considine et al., 2002; Dai and Hozalski, 2002, 2003; Mohanram et al., 2010, 2012). These phenomena may result in enhanced transport, as observed in sandy loam soils. Adsorption of organic matter on the surface of C. parvum oocysts induces an increase in zeta potential that counteracts the negative charge.

In addition, the presence of Ca²⁺ has been postulated as possible mediator in the coating of C. parvum oocysts by OM (Dai and Hozalski, 2003). Soils that had the highest concentration of Ca²⁺, Greenson series (42.09 mg·kg⁻¹ and Lewiston series (37.10 mg·kg⁻¹), and the highest clay content (12.5 and 19.5% for Lewiston and Greenson series, respectively) had the greatest breakthrough of C. parvum oocysts, as corroborated by the C. parvum oocysts BTC results and the analyses of the physicochemical properties of the soils. Our findings support and demonstrate the critical role of Ca²⁺ in mediating the transfer of OM from the surface of minerals to the surface of C. parvum oocysts. In particular, the Lewiston and Greenson soil series that had the highest soil Ca²⁺ concentration, the highest clay content, and the highest silt content resulted in the highest breakthrough of C. parvum oocysts among all soils, although the OM concentrations of all soils were similar (3.4, 0.8, 4.20, and 3.80% OM for Sparta, Lewiston, Gilford, and Greenson, respectively) (Tables 1, 3 and Figure 1). We can then postulate that the interplay of clay (and silt to some extent), OM, and Ca²⁺ facilitated and mediated the transfer of OM to C. parvum oocysts surface, which resulted in the breakthrough of C. parvum of oocysts through the soil. Based on our findings, this interplay between these biogeochemical compounds and processes—clay and silt content, presence of OM, Ca²⁺ content, and Ca²⁺ mediated transfer of OM from mineral surfaces to C. parvum oocysts—is critical in controlling the mobility and transport of C. parvum oocysts in soils. Our observations are also in agreement with previous research findings by Ferguson et al. (2003) and Peng et al. (2008) that also highlighted the role of soil texture and soil physico-chemical properties on the fate and transport of C. parvum oocysts in soil.

Compared with the total number of C. parvum oocysts applied to the soil columns surface, a highest oocyst recovery rate reported for soil samples (2.718%) was considerably greater than the highest oocyst recovery rate reported for leachate samples (0.006%). These recovery results are in agreement with those from Petersen et al. (2012), with the highest recovery rate of oocysts at 9.8% for soil samples and 0.03% for leachate samples. The median of the efficiency of C. parvum oocysts recovery from the different soil matrices in our experiment was calculated as 0.128%. This is similar to the previously reported efficiency of C. parvum oocyst recovery from soils in other studies (Mawdsley et al., 1996a; Petersen et al., 2012). Mawdsley et al. (1996a) investigated the effect of incubation time on the number of C. parvum oocysts extracted from soil initially inoculated with C. parvum oocysts. Mawdsley et al. (1996a) found that the efficiency of C. parvum occyst recovery from soil decreases rapidly as time passes, from 61.6% for an incubation time of 15 min to 0.3% for an incubation time of 7 days. A substantially higher number of oocysts were recovered from the sandy loam soils (Lewiston and Greenson series) than the loamy sand soils (Sparta and Gilford series), as shown in Table 3. However, the distributions of C. parvum oocysts within the soil profiles were similar. Most of the C. parvum oocysts were encountered in the soil top layers (0–5 cm) and in the bottom layers (17.5–20 cm). Our C. parvum oocyst recoveries are in agreement with previously published results (Mawdsley et al., 1996a; Petersen et al., 2012). Also, our time of inoculation of C. parvum oocysts in soils and the volume of soil samples treated for C. parvum extraction are higher than in previously reported studies (Mawdsley et al., 1996a; Petersen et al., 2012).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.