Does each bead count? A reduced-cost approach for recovering waterborne protozoa from challenge water using immunomagnetic separation

Giardia duodenalis and Cryptosporidium spp. are two of the most prominent aetiological agents of waterborne diseases. Therefore, ef ﬁ cient and affordable methodologies for identifying and quantifying these parasites in water are increasingly necessary. USEPA Method 1623.1 is a widely used and validated protocol for detecting these parasites in water samples. It consists of a concentration step, followed by parasite puri ﬁ cation and visualization by immuno ﬂ uorescence microscopy. Although ef ﬁ cient, this method has a high cost particularly due to the immunomagnetic separation (IMS) step, which is most needed with complex and highly contaminated samples. Based on this, the present study aimed to determine whether it is possible to maintain the ef ﬁ ciency of Method 1623.1 while reducing the amount of beads per reaction, using as a matrix the challenge water recommended by the World Health Organization. As for Giardia cysts, a satisfactory recovery ef ﬁ ciency (RE) was obtained using 50% less IMS beads. This was evaluated both with a commercial cyst suspension (56.1% recovery) and an analytical quality assessment (47.5% recovery). Although RE rates obtained for Cryptosporidium parvum did not meet Method 1623.1 criteria in any of the experimental conditions tested, results presented in this paper indicated the relevance of the described adaptations, even in challenge water. adhering to beads after dissociation may impact recovery levels.

Almost 40% of these deaths are caused by parasitic protozoa, especially Giardia duodenalis and Cryptosporidium parvum, which are zoonotic aetiological agents responsible for more than 2.8 million cases per year of gastrointestinal infections worldwide (Squire & Ryan ). These infections are the second most common cause of death in early childhood (Checkley et al. ; Platts-Mills et al. ).
The repeated prevalence of these protozoa in surface water denotes significant risks to human health, especially due to their low ID50 (the number of cysts and/or oocysts needed to infect 50% of exposed people), which has been estimated to fall between 10 and 2.000 for C. parvum (Robert-Gangneux & Dardé ) and between 10 and 100 for G. duodenalis (Rendtorff ). In this context, assessing the microbiological quality of drinking water is mandatory to ensure its safety for consumption (WHO ).
Despite the growing trend in pathogen epidemiological investigations in developing countries (Squire & Ryan ), the vast majority of studies are still carried out in developed countries, where laboratories and general health infrastructure are much more accessible than those in developing countries (Snelling et al. ). Cryptosporidium is associated with moderate to severe diarrhoea and increased mortality in low-income regions (Sunnotel et al. ; Snelling et al. ), and both parasites negatively affect child growth and development (Squire & Ryan ).
Malnutrition and HIV status are also important contributors to the increased prevalence of Cryptosporidium spp. and G. duodenalis in developing countries. Climate change and population growth are also predicted to increase both malnutrition and the recurrent prevalence of these parasites in water sources (Squire & Ryan ).
Over time, various methodologies have been developed to detect these organisms in water samples. The limitations in early-stage methodologies for protozoan recovery may result in a slight prevalence in surface water, for instance, leading to the incorrect assumption of low contamination (Efstratiou et al. a). Also, the efficiency of the critical step, that of oocyst recovery, in these methods is mostly low and extremely variable, ranging from 0 to 140% ( Regardless of the detection method employed, large volumes of water are usually required in order to increase the likelihood of detecting cysts and oocysts in the sample. However, the concentration process often leads to an accumulation of debris, such as large organic particles and algal cells, making it necessary to include a sample clarification step, which aims to separate the target organisms from this debris (McCuin et al. ). In this scenario, IMS is a well-established technique that employs magnetic beads coated with an antibody specific to protein targets on the surface of microorganisms such as Giardia spp. and Cryptosporidium spp., to allow their recovery from different matrices (Di Giovanni et al. ; Yakub & Stadterman-Knauer ).
Although this technique has operational advantages and presents better results than other methods (Hsu & Huang ), the high cost of the immunomagnetic beads severely limits its use in limited-resource situations (Feng et al. ). Reducing the cost of IMS methodology is, therefore, crucial to ensure that, even in low-and middle-income countries, effective detection of pathogens in water becomes practically feasible. Such a development would lead to standardization of the methodologies across all laboratories and more consistent and reliable results worldwide. As immunomagnetic beads are a primary cost of the method, we, therefore, investigated the efficiency of the IMS method when the amount of beads per sample is serially reduced as a step towards achieving this specific goal.

METHODOLOGY Sampling
Test water consisted in an increase of turbidity and true colour to a natural water source. In short, a 5 L groundwater sample was mixed with humic acid (20 mg L À1 ) and kaolinite (60 mg L À1 ) in order to reach about 40 NTU of turbidity, 250 HU of true colour and 10 mg L À1 of dissolved organic carbon (DOC). These characteristics represent the so-called challenge water proposed by the World Health Organization (WHO ) for water testing.
In our study, 5 L batches were used for each test, and these 5 L batches were divided into five samples of 1 L each.
The groundwater used in this study came from an artesian well which is fed by the waters of the Guarani Aquifer System. The well is located on the campus of the São Carlos School of Engineering, São Carlos, São Paulo, Brazil.
Specifically, for this work, prior to the beginning of the experiments, the well water was submitted to Method 1623.1 (MF þ IMS þ IFA) for the detection of (oo)cysts of Giardia spp. and Cryptosporidium spp. The aforementioned method was used for the analysis of all samples included in this study and is, therefore, detailed in the subsection 'Sample processing'.
Viable cysts and oocysts were shipped and stored in phosphate-buffered saline containing antibiotics at 2-8 C and were utilized within a maximum of 60 days after receipt.
Approximately 697 ± 8 cysts and 700 ± 10 oocysts were spiked together into each of four of the 1 L samples, with the remaining 1-L sample being used as a blank control (i.e., without protozoa).
Prior to the tests, the suspensions were analysed to quantify the inoculum. For that, 5 μL of each suspension was spiked on a glass slide, in triplicate, and left at room temperature for 4 h for drying. Next, the commercial kit Merifluor™ (Meridian Diagnosis) and DAPI (4 0 ,6-diamidine-2 0 -phenylindole dihydrochloride) dye (USEPA ) were applied to the sample. Visualization was performed by immunofluorescence microscopy (Olympus ® BX51).
The final concentration (microorganisms/μL) was given by the average of the results observed in the three slides. The resulting liquid was then subjected to a double centrifugation process (1,500 × g; 10 min; room temperature) to form a pellet containing the target parasites. At the end of the process, samples were resuspended in 5 mL of appropriate kit buffer, and then subjected to IMS in order to purify the protozoa.

Sample processing
It is worth mentioning that throughout the filtration technique, the filter membranes may need to be replaced if they clog and interrupt the flow of the sample. The amount of membranes used depends directly on the characteristics of the study water.

Sample purification and protozoa isolation
The Dynabeads™ GC-Combo (Applied Biosystem) kit was applied in this step following the manufacturer's recommendations; this kit was also used for the dissociation step, which was carried out three times using 100 μL of 10% hydrochloric acid, in each time. As part of our aim to obtain an effective but more affordable methodology, assays were performed under four different conditions with a serial reduction in the amount of beads in each.
The first assay was performed according to the standard protocol of Method 1623.1, in which 100 μL of each bead type was added to the sample. For the second assay, the bead volume was reduced by 50%, and, in the third, the final amount of bead added to the sample was 25% of the standard protocol. The 4th assay was performed without any beads. Apart from these reductions, all other conditions were kept the same for each assay.
Considering the results obtained during the tests, an extra test was included in order to investigate whether the addition of double the volume of 10% hydrochloric acid (200 μL) in the standard amount of beads (100 μL) would positively influence RE.

Microorganism visualization
At the end of each of the three rounds of acid dissociation, 50 μL of the sample (non-adhered material) was recovered and added directly to one of the wells of the glass slide supplied with the Merifluor™ Kit, which was previously sensitized with 5 μL of sodium hydroxide 1 M.
After the drying period of the samples on slides (4 h), (oo)cysts were stained using the commercial Merifluor™ (Meridian Diagnosis) kit and visualized by immunofluorescence light microscopy (Olympus ® BX51). As a confirmatory test, DAPI dye was added to all the samples (USEPA ).

Recovery rate
Recovery efficiency (RE) of the method is calculated according to Equation (1), where RE is the recovery rate after the complete protocol (%); C 1 is the (oo)cysts enumerated in the first acid dissociation; C 2 is the (oo)cysts enumerated in the second acid dissociation; C 3 is the (oo)cysts enumerated in the third acid dissociation; and NP is the number of inoculated protozoa.

Recovery efficiency
The cysts of G. duodenalis and oocysts of C. parvum were clearly observed against the background in all samples ( Figure 1), regardless of the condition of the test, following the first and second acid dissociations. After the third round of acid dissociation, no cysts or oocysts were visualized. RE data obtained from different methodologies carried out in this study are compiled in Table 1.
As for operational purposes, it is worth mentioning that five membranes were used in each batch filtration.

Statistical analysis
The Shapiro-Wilk test indicated that data for percent C. parvum recovery without adding beads did not follow a normal distribution. Although the boxplot shown in Figure 2 perhaps visually suggests that the data is normally distributed, this hypothesis was not confirmed. Therefore, C. parvum recoveries were analysed by non-parametric statistics. Although the Kruskal-Wallis test suggested significant   Figure 2(b), G. duodenalis recoveries were prioritized for the analytical quality assess-

ment. A comparison among all experimental conditions is
shown in Table 2 which shows that the 50 μL-bead dosing led to significant differences in the sample means against all of the other conditions.

Analytical quality assessment
Colorseed™ was used to validate the lowest IMS bead concentration that still provided an acceptable RE value. This was determined to be 50 μL of each bead suspension dissociated with two rounds of 100 μL of 10% hydrochloric acid.
Under these conditions, RE reached similar values to those of a test with commercial protozoan suspensions (Table 1).
Comparing the values obtained herein with those standardized by Method 1623.1, our data were satisfactory for Giardia spp. regarding both RE (47.5%) and CV (4%). Concerning Cryptosporidium spp., RE was 17% below the value recommended by Method 1623.1, while the CV for Cryptosporidium spp. met the USEPA criteria (7.1%).

Cysts and oocysts attached to the beads
In order to verify the efficiency of the acid dissociation procedure, 50 μL of the bead suspensions obtained at the end of the IMS step were stained with Merifluor™ and imaged using fluorescence microscopy. Figure 3 displays the image captures of the best experimental condition obtained in   (2), where PA is the total amount of protozoan adhered to the beads after three acid dissociation procedures (%); P 1 is the (oo)cysts attached to the beads after the first acid dissociation; P 2 is the (oo)cysts attached to the beads after the second acid dissociation; P 3 is the (oo) cysts attached to the beads after the third acid dissociation; and NP is the number of inoculated protozoa.

DISCUSSION
The quality of water resources is a fundamental aspect of the Some cysts and oocysts were detected in the absence of IMS, but the RE was insignificant (Table 1).
The fact, pointed out by the results of this study, that the IMS methodology regardless of the number of beads was efficient only for the recovery of G. duodenalis cysts was Our protozoa recovery from challenge water also suggests that the third dissociation may be dismissed since no organisms were visualized after it, further reducing the cost of the protocol, as no labelling of an extra microscopy slide should be required for immunofluorescence.
Our revised methodology represents a significant improvement compared with those previously carried out.
Particularities inherent to the matrix and the methodology itself may influence results. The WHO challenge water, which was used in this study, presents a much higher turbidity than filtered or treated water from water treatment plants  countries and a reduced-cost approach might assist in reaching this goal.
As previously mentioned, the high cost of the methodology does not fall exclusively onto the IMS, but it is, in fact, the main expense. The Merifluor™ kit, additional use of DAPI, the epifluorescence microscope and all the necessary infrastructure to carry out the method are direct contributors to its enhancement. However, none of the aforementioned items/reagents can be removed from the global protocol without causing its mischaracterization and most likely loss of results. In this sense, we opted for the careful optimization of one of the methodological steps as an attempt to generate financial savings. The cost reduction in the IMS procedure is reflected by the increased durability of the kit, which, according to our results, can be used in 100 samples instead of 50, as recommended by the manufacturer. Therefore, the alternative offered by our study (50 μL of beads), which complies with the USEPA criteria at least for G. duodenalis allows doubling the capacity of the Dynabeads™ kit leading to a significant reduction in costs. In addition, the inference that the third acid dissociation step is not necessary for the success of the methodology also impacts its cost, as less IFA reagents, DAPI and hydrochloric acid will be required per sample.

CONCLUSIONS
Based on the results obtained from this study, we suggest an adaptation to the purification step described in Method 1623.1 in order to provide a methodology with a better cost-benefit that still provides the recovery rate necessary for (oo)cysts, even from complex matrices.
Although none of the conditions explored here was satisfactory for C. parvum oocyst recovery, the results point to a significant cost reduction of G. duodenalis cyst detection, since half of the volume of immunomagnetic beads (50 μL) used in our study complied with the USEPA recovery efficiencies.
The development of cost-effective protocols to detect and monitor waterborne parasites in water (e.g., Cryptosporidium spp. and G. duodenalis) is crucial to more effectively evaluate the water quality in developing countries having a direct impact on public health. However, this will continue to be extremely challenging, not least because scientists in developing countries face lower absolute levels of funding and must often pay far too expensive and unsustainable costs for consumables and equipment.
Further studies are recommended to improve the organism-bead dissociation process, seeking to increase the protozoa detection protocol performance in the sample purification phase.