Detection and quantification of soil-transmitted helminths in environmental samples: A review of current state-of-the-art and future perspectives.

It is estimated that over a billion people are infected with soil-transmitted helminths (STHs) globally with majority occurring in tropical and subtropical regions of the world. The roundworm (Ascaris lumbricoides), whipworm (Trichuris trichiura), and hookworms (Ancylostoma duodenale and Necator americanus) are the main species infecting people. These infections are mostly gained through exposure to faecally contaminated water, soil or contaminated food and with an increase in the risk of infections due to wastewater and sludge reuse in agriculture. Different methods have been developed for the detection and quantification of STHs eggs in environmental samples. However, there is a lack of a universally accepted technique which creates a challenge for comparative assessments of helminths egg concentrations both in different samples matrices as well as between locations. This review presents a comparison of reported methodologies for the detection of STHs eggs, an assessment of the relative performance of available detection methods and a discussion of new emerging techniques that could be applied for detection and quantification. It is based on a literature search using PubMed and Science Direct considering all geographical locations. Original research articles were selected based on their methodology and results sections. Methods reported in these articles were grouped into conventional, molecular and emerging techniques, the main steps in each method were then compared and discussed. The inclusion of a dissociation step aimed at detaching helminth eggs from particulate matter was found to improve the recovery of eggs. Additionally the selection and application of flotation solutions that take into account the relative densities of the eggs of different species of STHs also results in higher egg recovery. Generally the use of conventional methods was shown to be laborious and time consuming and prone to human error. The alternate use of nucleic acid-based techniques has improved the sensitivity of detection and made species specific identification possible. However, these nucleic acid based methods are expensive and less suitable in regions with limited resources and skill. The loop mediated isothermal amplification method shows promise for application in these settings due to its simplicity and use of basic equipment. In addition, the development of imaging soft-ware for the detection and quantification of STHs shows promise to further reduce human error associated with the analysis of environmental samples. It may be concluded that there is a need to comparatively assess the performance of different methods to determine their applicability in different settings as well as for use with different sample matrices (wastewater, sludge, compost, soil, vegetables etc.).


a b s t r a c t
It is estimated that over a billion people are infected with soil-transmitted helminths (STHs) globally with majority occurring in tropical and subtropical regions of the world. The roundworm (Ascaris lumbricoides), whipworm (Trichuris trichiura), and hookworms (Ancylostoma duodenale and Necator americanus) are the main species infecting people. These infections are mostly gained through exposure to faecally contaminated water, soil or contaminated food and with an increase in the risk of infections due to wastewater and sludge reuse in agriculture. Different methods have been developed for the detection and quantification of STHs eggs in environmental samples. However, there is a lack of a universally accepted technique which creates a challenge for comparative assessments of helminths egg concentrations both in different samples matrices as well as between locations. This review presents a comparison of reported methodologies for the detection of STHs eggs, an assessment of the relative performance of available detection methods and a discussion of new emerging techniques that could be applied for detection and quantification. It is based on a literature search using PubMed and Science Direct considering all geographical locations. Original research articles were selected based on their methodology and results sections. Methods reported in these articles were grouped into conventional, molecular and emerging techniques, the main steps in each method were then compared and discussed. The inclusion of a dissociation step aimed at detaching helminth eggs from particulate matter was found to improve the recovery of eggs. Additionally the selection and application of flotation solutions that take into account the relative densities of the eggs of different species of STHs also results in higher egg recovery. Generally the use of conventional methods was shown to be laborious and time consuming and prone to human error. The alternate use of nucleic acid-based techniques has improved the sensitivity of detection and made species specific identification possible. However, these nucleic acid based methods are expensive and less suitable in regions with limited resources and skill. The loop mediated isothermal amplification method shows promise for application in these settings due to its simplicity and use of basic equipment. In addition, the development of imaging soft-ware for the detection and quantification of STHs shows promise to further reduce human error associated with the analysis of environmental samples. It may be concluded that there is a need to comparatively assess the performance of different methods to determine their applicability in different settings as well as for use with different sample matrices (wastewater, sludge, compost, soil, vegetables etc.

Introduction
It is estimated that over 1.5 billion people are infected with at least one species of soil-transmitted helminths (STHs) worldwide (WHO, 2015), with the majority of these infections caused by the roundworms (Ascaris lumbricoides and Strongyloides stercoralis), whipworms (Trichuris trichiura) and hookworms (Necator americanus or Ancylostoma duodenale) (Strunz et al., 2014). Ascariasis is reported in 771.7-891.6 million people, while 429.6-508.0 million people have trichuriasis and 406.3-480.2 million are infected with hookworm (Pullan et al., 2014). Most of these infections occur in tropical and subtropical regions of the world where poverty results in poor sanitary conditions (Stolk et al., 2016). STHs infections are mostly caused by exposure to faecally contaminated water, soil or contaminated food (Keraita and Amoah, 2011). Wastewater and sludge reuse is reported to contribute significantly to the high risk of infections. In endemic areas, wastewater is estimated to contain up to ∼3000 eggs/L (Kamizoulis, 2008;Mara and Sleigh, 2010).
The association between ascariasis and wastewater use among farmers has been reported by several studies (Seidu et al., 2008;Blumenthal et al., 2001;Pham-Duc et al., 2013;Rutkowski et al., 2007;Trang et al., 2006;Habbari et al., 1999), where consumers of the farm produce are also at risk of infection. The highest health risks for consumers of wastewater irrigated produce are with crops which are eaten raw, for example, salad crops and some root crops or crops grown close to the soil surface (e.g. lettuce) (WHO, 2006). Wastewater for unrestricted reuse in agriculture should contain ≤1 helminth egg per liter to reduce the risk of STHs infections to below the WHO guidelines target level (WHO, 2006). This requires sensitive detection and a consistent quantification of STHs eggs in wastewater, sludge or other sample matrices. Accurate detection and quantification of STHs eggs in environmental samples is challenging. The heterogeneity of the occurrence of STHs in environmental samples is problematic for laboratory testing due to varying amounts of moisture, solids, quantity of samples and soil particles (Collender et al., 2015). Another challenge with environmental samples is the need to recover small numbers of STHs eggs from large volumes of samples (Maya et al., 2006;Mes, 2003).
Over the last few years, different techniques for detecting and quantifying the total number and the viable and non-viable fractions of STHs eggs in environmental samples has been developed and applied. The choice of technique used is largely influenced by the different types of samples (Maya et al., 2006). One reason for this may be the lack of published quality assurance/quality control (QA/QC) data on the various methods (Bowman et al., 2003). In addition to the more traditional methods based on sedimentation and/or flotation, which mainly involves the separation and concentration of eggs and the microscopical identification and quantification of these eggs, several new techniques have been developed. These new techniques make the identification and quantification of helminth eggs more efficient and sensitive. The advent of genomic sequencing and the wealth of data generated have markedly increased the feasibility of developing polymerase chain reaction (PCR)-based methods as diagnostic tools for helminth parasites (Gyawali et al., 2015). Defined gene sequences of STHs eggs can be detected with PCR, quantitative PCR (qPCR) and other nucleic acid based methods from small quantities of samples (Gordon et al., 2011). These techniques can also identify STHs eggs to species level (Gordon et al., 2011). Furthermore advanced nucleic acid based techniques (like multiplex PCR) makes it possible to target more than one species at a time (Pontes et al., 2002). The reliable quantification of viable eggs is essential both related to risk based target values and for the validation of sanitation system performance.
The lack of a globally accepted method for the detection and quantification of STHs eggs in environmental samples poses a challenge for comparative assessments of egg concentrations both in different sample matrices as well as between locations. This article presents a review of commonly used methods for the detection and quantification of STHs eggs in environmental samples, with a comparison of the advantages and disadvantages of each technique. We also review new and emerging techniques that may be applied for the detection and quantification of STHs eggs in the environment.

Search strategy
The review is based on a literature search using PubMed and Science Direct, with the following keywords and strings (soiltransmitted helminths OR intestinal parasites OR Ascaris spp OR hookworm OR Trichuris spp OR Toxocara spp OR Taenia) AND (wastewater OR water). The organism search-string were repeated with AND (sludge OR compost); AND (vegetables OR crops OR plants); as well as with AND (soil OR urine diversion (UD) toilet waste OR biosolids). Original research articles were selected and the methodology section of each article was assessed to determine the method used in the detection and quantification of the STHs. All articles were considered irrespective of the year of publication.
Methodological studies on the concentration of STHs in environmental samples were included but articles and publications related to clinical STHs infections and drug efficiency/administration reports were excluded. All geographical areas were considered for the review, however only articles in English were included. A total of 195 articles were reviewed and information from 164 used, based on a clear method for detection of the STHs. Publications reporting the same method were assessed in relation to methodological variations and the type of sample matrices. When variations were not evident, one major reference to the methodology was considered and the other was captured as an additional method.

Conventional methods
Most methods used for the detection and quantification of STHs in environmental samples involve the recovery of STHs eggs from the sample matrix and quantification of STHs eggs or larvae using microscopy. These methods are collectively referred to as 'conventional methods' due to the application of basic techniques that includes sedimentation, followed by an extraction phase and then flotation with zinc sulphate (Ayres and Mara, 1996) before viewing under a microscope. The methods reviewed are categorized based on the major steps involved in the analysis. These steps mostly are the source of method variability, relating to sample types and quantity, pre-treatment, separation of STHs eggs, microscopy and viability determination.

Sample types and quantity
This section is focused primarily on the sample volumes studied and reported in the reviewed literature, with the subsequent sections below dealing with the various method variations and the influence of the different steps on the recovery of STHs eggs from the various sample matrices.

Wastewater/water
Helminth egg concentrations in wastewater (especially treated wastewater and water) are normally very low (Mara and Silva, 1987;Schwartzbrod et al., 1987;Ayres et al., 1991), mainly due to the dilution of egg concentration in the environment. Therefore, most methods use large sample volumes for detection. The concentration of the eggs is largely dependent on the prevalence of infection in a study area (see Table 1 below). When the prevalence of STH infections in a study area is low larger sampling volumes applies, where volumes up to 200 L has been reported (Levantesi et al., 2010). Most methods used for the detection of STHs eggs in environmental samples are a modification of the United States Environmental Protection Agency (USEPA) (Schwartzbrod, 1998) and the World Health Organization (WHO) methods (Ayres and Mara, 1996). These methods recommend the use of one liter samples ( Molleda et al., 2008). Furthermore, the volume of water analyzed is directly related to the level of detection achieved (Bowman et al., 2003). For instance, the WHO guidelines for wastewater reuse in agriculture recommends <1 viable helminth egg/Liter for unrestricted irrigation (WHO, 2006). Therefore methods that aim to determine the quality of wastewater for irrigation must consider volume sufficient to reach the limit of detection. High solid content in the sample may result in lower rates of egg recovery due to interference of the solids (Rocha et al., 2016). Data presented in Table 1 below shows that even when sample volumes are the same, the amount of helminth egg in the wastewater differs greatly from one region to the other, which is dependent on prevalence of infection in the study area.

Sludge/Compost/Biosolids
Sludge, biosolids and compost may contain high concentration of STHs eggs since these will attach and concentrate together with particulate material and accumulate in the sludge (Shamma and Al-Adawi, 2002) during wastewater treatment. Eggs of STHs settle much easier into sludge during primary treatment than bacteria and viruses due to their larger sizes (Yeager and O'Brien, 1983). The concentration of STHs eggs in sludge samples varies greatly depending on the prevalence of infection in the relevant population. Table 2 shows concentration of eggs in sludge from different locations, with a higher concentration in sludge samples from developing countries irrespective of the sample size. The amount of sludge sampled is often related to the dry weights, which are in the interval from 2 g (Scaglia et al., 2014;Pecson et al., 2007;Nelson and Darby, 2001) to 5 g (Bowman et al., 2003). This contrasts with the USEPA method that recommends 300 g as a dry weight sample size (Cappizzi-Banas et al., 2004;Maya et al., 2012;Gantzer et al., 2001). The variation in the weight of solid sample, as well as is the case for liquid samples, is mainly influenced by prevalence of the STHs in the study area based either on literature or past experiences. Large samples of sludge have been found to lower the rate of egg recovery, mainly due to the interference of solids. Bean and Brabants (2001) observed that the recovery of Ascaris spp eggs from a 50 g sludge sample was twice as high as from a 300 g of sludge and biosolids. It is generally recommended that samples should contain between 5 g (Reimers et al., 1989) to 50 g (USEPA, 2003) of total solids. Table 2 contains information of the sample weight and the study area which may play a critical role in the amount of eggs recovered. Studies performed in developed countries (France, Italy etc.) reported lower concentration of STHs eggs (Scaglia et al., 2014;Gantzer et al., 2001), as compared to studies in developing countries (Ghana, Mexico, Burkina Faso) (Buitrón and Galván, 1998;Koné et al., 2007;Bastos et al., 2013). With larger sample sizes the concentration of reported STHs egg is high irrespective of the location of study (Reinoso and Becares, 2008;Engohang-Ndong et al., 2015;Bowman et al., 2003).

Soil
The concentration of STHs eggs in soil can be very high depending on the sampling site especially in unsanitary environments or where open defecation is practiced. Soil sampling for other helminths in addition to STHs eggs is important due to the possibility of zoonotic infections from infected animals, particularly dogs and cats (e.g. Toxocara canis, Toxocara cati and Ancylostoma caninum). Direct contact with dogs or cats that harbor adult Toxocara worms is unlikely to cause an infection in humans because once shed, the eggs must undergo a period of development in the environment before they can become infective (Paquet-Durand et al., 2007). The sample size for soil is measured in dry weights, with sample sizes between 10 and 50 g of dry soil ( Mizgajska, 1997;Fajutag and Paller, 2013). Samples mostly represent the top 0-5 cm layer of the soil (Paquet-Durand et al., 2007;Mizgajska, 1997;Fajutag and Paller, 2013;Nooraldeen, 2015). Important factors that influence the recovery of helminth eggs from soil include soil type and texture (Nunes et al., 1994). Sand and sandy soils are reported to result in higher recoveries of helminths than from clay, silt and silty clay-soils (Oge and Oge, 2000). Sandy soils also result in less variation in the number of eggs recovered, which might be due to the homogeneity of sandy soils based on the greater particle sizes held loosely together compared to other soil types (Oge and Oge, 2000). In addition to the investigated soil type, the study location also plays a critical role in the determination of soil weight to sample, as shown in Table 3. Since the prevalence of STHs eggs in soil samples vary from one location to the other, sampling should account for the local prevalence of the STH where possible.

Plants
With an increase in the use of wastewater for irrigation of crops, as well as the use of sludge, biosolids and compost as fertilizers or soil conditioners in agriculture, there is an enhanced likelihood of plants becoming contaminated with pathogens with the further transmission of STHs to consumers (WHO, 2006). Different types of plants irrigated with wastewater or fertilized with sludge have different likelihoods for transmission (e.g. differences between salad crops and foliage plants). This may be reflected in the sampling strategy as well as in the contamination levels. The amount/weight of vegetable sampled or analyzed varies between methods. Preferred weights of plant samples normally falls between 100 and 250 g (Adamu et al., 2012;Maikai et al., 2012;Adanir and Tasci, 2013;Fallah et al., 2016). These weights are mostly of vegetables, with lower weights for grass (Paquet-Durand et al., 2007;Robertson et al., 2002). Several reported studies fail to report on the weight of plant material analyzed (Su et al., 2012;Hassan et al., 2012;Said, 2012;Rostami et al., 2016). The specific weight is essential for risk calculation for quantitative microbial risk assessment (QMRA) and risk of infections based on consumption. However, the concentration of STHs eggs reported from different studies is largely influenced by the quality of wastewater used for irrigation, as well as the STHs egg concentration in sludge or compost used as fertilizers. Table 4 presents the prevalence of positive STHs in vegetable or plant samples from different countries.

Egg recovery
The variation in recovery is influenced by methodological approaches in each of the analytical stages as well as the sample matrix, type of STHs eggs, available reagents/materials and the discretion of the researcher. Table 5 summarizes some of these variations reported in literature. The sampling strategy for STHs eggs should reflect the uneven distribution in the environment. Their recovery will be enhanced through homogenization. Homogenization is most relevant for solid samples such as sludge, biosolids, compost and the solid waste fraction from source separation toilets (Bowman et al., 2003). It is usually achieved through blending or mixing (Bowman et al., 2003;Cappizzi-Banas et al., 2004;Pecson et al., 2007). Blending also breaks down coarse materials into finer particles which improves the recovery of the eggs (Koné et al., 2007).

Separation of eggs from particles
Separation of STHs eggs from solids in the sample is of major concern especially when considering sludge, compost and biosolids, due to their heterogeneity. Samples with a high concentration of particles (including soil particles) could easily trap the eggs, or become attached to the eggs which will result in lower recovery. Different solutions have been used to recover eggs from particulate matter. These solutions are mainly anionic detergents that break the bonds between the STHs eggs and particles in the samples. The most commonly used detergent solutions are Tween 80, Triton X-100 (Molleda et al., 2008;Forslund et al., 2010), 7X (Sengupta et al., 2011;Saddoud et al., 2007;Konaté et al., 2013) and ammonium bicarbonate (Moodley et al., 2008;Trönnberg et al., 2010). Although most researchers favor the use of detergents for dissociation, some have reported a lower recovery for treated versus untreated samples (Rocha et al., 2016). Earlier reports showed that the use of detergents reduced egg viability due to damage of egg integrity (Jaskoski, 1954).
Although there is a lack of comparative data to determine the best detergent for dissociation of helminth eggs from environmental samples, the use of 7X has been reported to give better recoveries (Bowman et al., 2003). This detergent is water soluble at all concentrations and comprises of anionic surface active agent and special solvents which may account for the improved recovery. In addition 7X does not form precipitates when in contact with salt solutions used during the flotation step (Rocha et al., 2016). The first report of the recovery of STHs eggs from soil was in 1928 by Caldwell and Caldwell (David, 1977), where antiformin resulted in a good recovery (especially in clay soils) (David, 1977). Saturated ammonium bicarbonate has been used to improve recovery of eggs from samples with a high content of soil (Moodley et al., 2008;Collender et al., 2015). A variety of solutions have been used in the analysis of vegetable or plant samples. Table 5 includes the different separation solutions used in the analysis of wastewater or sludge. Solutions such as physiological saline (Adanir and Tasci, 2013;Maikai et al., 2012;Hassan et al., 2012;Adamu et al., 2012), phosphate buffered saline (PBS) (Said, 2012), Tween 80 (Kozan et al., 2005;Pacquet-Durand et al., 2007) and water (Su et al., 2012;Kłapeć, 2009) are commonly used in contrast to Tween 80/20 or 1% 7X that is more common in other sample matrices. Again, due to a lack of comparative data on the best solutions to use for washing of plant samples, the choice is mostly reliant on the discretion of the researcher.

Filtration of samples
After dissociation of eggs from larger particles within samples, there is a need to separate the eggs from these particles. One step used to achieve this separation is filtration or the use of sieves to retain larger particles whiles allowing the eggs of interest to pass into the filtrate for further analysis (Bowman et al., 2003;Katakam et al., 2014;Engohang-Ndong et al., 2015). The choice of pore size for this step is the most important factor to consider. Most parasite eggs have dimensions between 25 m-150 m (Yeh et al., 2015;Bouchet et al., 2003). Therefore, to allow eggs to pass through, pore sizes from 4 m to125 m (sizes varying based on the helminth egg of interest), are commonly used (Gaspard et al., 1995;Gantzer et al., 2001;Mizgajska, 1997;Blaszkowska et al., 2013;Horiuchi et al., 2013;Fajutag and Paller, 2013). In some instances the interest is to retain the eggs on the sieves instead and therefore smaller pore sizes such as 20 m (Maya et al., 2010;Landa-Cansigno et al., 2013) or even as low as 8 m are chosen (Buitrón and Galván, 1998). In instances where the STHs of interest are restricted to nematodes, sieves of pore sizes between 32 and 36 m are used (Katakam et al., 2014;Maya et al., 2010;Gaasenbeek and Borgsteede, 1998). Table 5 gives examples of pore sizes of sieves that have been used to analyze different samples. It is evident that the variation in the pore size is mostly influenced by the helminth species of interest, as well as the sample matrix. For instance, most researchers working with vegetable or plant samples do not include this step due to decreased amounts of particulate matter. The influence of pore sizes of sieves on the recovery of eggs from different sample matrices has not been reported but filtration may reduce particulate material in the sample which would make the microscopy step easier, while also enhancing the accuracy in identification and quantification of the eggs. Efficiency in microscopy stage is improved due to a reduction in the particles that may obstruct the eggs on the slides. However, filtration may also result in a lower recovery of eggs due to the trapping of particle associated eggs or clumped eggs.

Sedimentation
The eggs in the filtrate need to be separated from the liquid phase. Separation of the solid particles (which includes the eggs) in samples from the liquid phase is mostly achieved through sedimentation. Sedimentation could be passive or with the use of centrifuge where different speeds are applied. This difference in the centrifugation speeds may introduce variation to the concentration of eggs recovered, however there is no data to ascertain the best speed. The time used for passive sedimentation varies between methods applied with setup time ranging from 1 h (Molleda et al., 2008;Yen-Phi et al., 2010) to overnight (Riahi et al., 2009;Saddoud et al., 2007;de Victorica and Galvan, 2003). The effect of passive sedimentation on the recovery of eggs could be influenced by several factors. These include the sample matrix and volume, the dimension of the container used for the sedimentation and the duration of the sedimentation. A high solid content in the sample may interfere with the settling of the eggs with a potential loss of some eggs (Rocha et al., 2016). The sedimentation rate is also greatly influenced by the dimension of the container used (Shuval et al., 1986). The WHO method (Ayres and Mara, 1996) recommends the use of an open-topped, straight-sided container of at least 10 L volume, aimed at making the removal of the supernatant easier and to permit thorough rinsing. In addition, the duration of the sedimentation is crucial and the sedimentation rates differ from one helminth species to the other. Adequate time is necessary for the settling of the eggs, accounting for the viscosity of the sample as well as the dimension of the container. For instance, Ascaris spp eggs (relative density of 1.13) have a settling velocity of 0.65 m/h, Trichuris spp eggs (relative density of 1.15) of 1.53 m/h while Taenia spp eggs (relative density of 1.23) will have a settling velocity of almost 2 m/h (David and Lindquist, 1982;Dryden et al., 2005). Inadequate time for sedimentation would result in the loss of some eggs, whereby overnight sedimentation as with the USEPA and Tulane methods would be an advantage (Mara and Sleigh, 2010).

Flotation
A critical step that results in method variation is the flotation step, where the main aim is to separate eggs from other materials in the sample that were not removed during the filtration or sedimentation steps. Flotation achieve separation by creating a gravity gradient that allows particles of interest (in this case the eggs) to float while heavier particles (like soil or other heavier particles) settle and are discarded. In this step the loss of STHs eggs relates to the specific gravity of the solution versus the density range for eggs, ranging from 1.05 to 1.23 (David and Lindquist, 1982). A variety of flotation solutions used include zinc sulphate (Trönnberg et al., 2010;Zamudio-Pérez et al., 2013;Amahmid et al., 2002), magnesium sulphate (Karkashan et al., 2014), sodium nitrate (Blaszkowska et al., 2013,) and sucrose solutions (Kouraa et al., 2002). For the recovery of all STH eggs, independent of species, the flotation solutions used should be heavier than 1.25 specific gravity (s.g) (David, and Lindquist, 1982). Since some methods use flotation solutions of specific gravity lower than 1.3 only some of the STHs species eggs will be recovered in consistent numbers. For example the USEPA and the Tulane method both use magnesium sulphate of specific gravity of 1.2 (Bowman et al., 2003;Saddoud et al., 2007;Konaté et al., 2013;Sengupta et al., 2011). Flotation solutions (e.g. sodium chloride and zinc sulphate) with lower specific gravity (1.18) are used in other methods (Yaya-Beas et al., 2016;Molleda et al., 2008;García et al., 2013). In addition to the STHs, Taenia spp eggs, with a specific density of 1.23 (David and Lindquist, 1982) is of main interest in environmental sampling and may not be recovered with low specific gravity solutions. Other adverse effects with flotation solutions are found with the use of sucrose, where increased viscosity has been shown to hinder the recovery of eggs. This interferes with the movement of the eggs as well as other particles (Bowman et al., 2003). Saturated sucrose has been shown to deform STHs eggs (Collender et al., 2015), and sodium nitrate forms crystals that would interfere with the quantification (Santarém et al., 2009). Nunes et al. (1994) found that sodium dichromate gave the best recovery ratio, which most certainly is due to the high specific gravity (s. g = 1.34) of the sodium dichromate used. Quinn et al. (1980) found magnesium sulphate (s. g = 1.27) plus 5% potassium iodide (KI) to be the most efficient solutions in the recovery of helminth eggs. Dada and Lindquist, (1979), found that zinc sulphate gives a better recovery of STHs eggs than sodium dichromate of the same specific gravity (1.20). This was later confirmed by Oge and Oge (2000). Table 5 shows some of the flotation solutions used and the variation in specific gravity and the corresponding STHs eggs that were recovered.

Phase extraction
Some recovery methods include a phase extraction step after the flotation. This aims at removing lipid-soluble and ether-soluble material from the sample. It results in the partitioning of the sample into acidic aqueous and lipophilic phases (Beaver et al., 1984), where a plug of waste (remaining particles) material is formed at the interphase between these. Pellets containing the eggs are deposited at the bottom of the centrifuge tubes. The commonly used reagents for the lipophilic extraction include ethyl acetate and diethyl ether (Bornay-Llinares et al., 2006;Horiuchi et al., 2013;Verbyla et al., 2016;de Victorica and Galvan, 2003;Ayres and Mara, 1996;Gaspard et al., 1996;Rude et al., 1987;USEPA, 1999). Reagents for the hydrophilic extraction include a mixture of sulfuric acid and acetoacetic buffer (Verbyla et al., 2016;de Victorica and Galván, 2003;Ayres and Mara, 1996;Fuhrimann et al., 2015). Satchwell (1986) found that the inclusion of a phase extraction step removes about 40% of contaminants (such as proteins and lipids) but results in the loss of 95% of the eggs. This may be due to distortion of the eggs or toxic effects. Several studies have documented the detrimental effects of some of these chemicals, such as ethyl acetate and acetoacetic buffer, on egg integrity and subsequently viability (Nelson and Darby, 2001;Rocha et al., 2016). Amoah et al. (submitted manuscript) found acetoacetic acid to result in the loss of egg viability. Due to the effect of this step on egg integrity it has been recommended to replace it with a sieving step, to remove proteins, lipids and other contaminant molecules (USEPA, 2003). However if the step is still used, then the exposure of the eggs to the chemicals must be within the shortest time possible in order to reduce their adverse impact (Nelson and Darby, 2001).

Viability determination
For the purposes of risk estimation and in accordance with some international and national standards and guidelines (USEPA, 2003;WHO, 2006), it is important to determine the number of viable STH eggs per amount of sample. This is, however, excluded in some studies which did not differentiate between viable and non-viable eggs (Fugazzola and Stancampiano, 2012;Bornay-Llinares et al., 2006;Pacquet-Durand et al., 2007;Hassan et al., 2012;Maikai et al., 2012). Viability assessment was excluded from these studies due to difficulties in the assessment process and time constraints. The most widely used viability assessment method is the time consuming incubation to achieve the development of the larvae. Other authors report viability based on morphological integrity of the eggs and their response to staining with vital dyes. The viability assessment method is influenced by factors such as materials and available equipment, experience of the researcher and personal preferences. The morphological determinations include size, shape and the presence of visible larvae and are used as a criterion for viability of eggs during microscopy. Viability determination based on the presence of visible motile larvae could be subjective based on the experience of the microscopists.
The limitation with the incubation to determine viability is due to its time consuming nature. When applied as part of verification monitoring, for example, in instances such as the reuse of wastewater or sludge in agriculture, the time lag for data may be a deterrent. This warrants the use of stains differentiating viable and non-viable eggs based on the permeability of the egg shells. Commonly used stains include Lugol's iodine (Benti and Gemechu, 2014;Ajeagah, 2013;Adanir and Tasci, 2013;Su et al., 2012), safranin O (Koné et al., 2007Konaté et al., 2013) trypan blue (Maya et al., 2012;Buitrón andGalván, 1998) andeosin Y (de Victorica andGalvan, 2003). Karkashan et al. (2015) compared different stains against conventional incubation and found that conventional incubation detects 86% of viable eggs which is lower than the viable eggs determine through safranin stain (97%), crystal violet stain (92%) and methylene blue (87%). The use of BacLight stain gave between 78 and 85% viability and the lowest viability (39%) was reported with trypan blue staining (Karkashan et al., 2015). The use of vital stains therefore presents an opportunity to determine viability of eggs without the prolonged incubation time required. Despite their potential advantages, some stains are toxic to embryos and require sample examination within a few minutes of application (Karkashan et al., 2015). Karkashan et al. (2015) found that only the BacLight stain is not toxic to helminth eggs.

BacLight Dead/Live method
This method is based on the BacLight LIVE/DEAD Bacterial Viability Kit (Molecular Probes, Invitrogen, Eugene, USA) and was initially developed for the enumeration of viable bacteria. It is based on detecting the difference in the membrane integrity of viable and non-viable cells. These have a differential intake of two specialized membrane permeable DNA-labelling dyes, Syto 9 which fluoresces green with a maximum emission of 498 nm and propidium iodide (PI) which fluoresces red with a maximum emission of 617 nm. The detection of viable eggs using this technique is based on the differential staining of viable eggs with Syto 9 whiles the PI stains the non-viable eggs (Lisle et al., 1998). Dabrowaska et al. (2014) found 58% live, 38% dead and 3.7% as insufficiently stained eggs of Ascaris spp, Toxocara spp and Trichuris spp in sludge samples. The authors concluded that the method was suitable for viability determination. These results were obtained from seeded samples and its performance was not compared with other methods. However Karkashan et al. (2015) reported between 78 and 85% viability of Ascaris spp eggs using the BacLight LIVE/DEAD staining technique. They also concluded that staining with this kit was the only procedure that does not affect the viability of eggs after analysis. Therefore, the use of the BacLight LIVE/DEAD staining is emerging as a rapid way of determining of the viability of STHs eggs without the need for incubation.

Nucleic acid based techniques
Advances in molecular technology provide an opportunity for an accurate and fast detection and quantification of STHs eggs in different sample matrices. PCR techniques have emerged as very specific, sensitive and rapid methods for the detection of different pathogens in a variety of matrices, from wastewater to soil and food items (Gomez-Couso et al., 2004;Ishiwata et al., 2004;Le Cann et al., 2004;Deisingh and Thompson, 2004). Several different PCR based methods are available for detection of pathogens, for example quantitative polymerase chain reaction (qPCR), multiplex polymerase chain reaction (mPCR), droplet/digital polymerase chain reaction (ddPCR). Despite the advancements in the area of molecular diagnostics only a few studies have considered their use for the detection of STHs eggs in environmental samples, even though they have found widespread use in clinical settings (Rocha et al., 2016;Pecson et al., 2006;Raynal et al., 2012).

Nucleic acid extraction
One of the main hindrances for the effective use of molecular methods in the detection of STHs eggs is the extraction of nucleic material of good quantity and quality. The extraction is hindered by the tough egg shell (van Frankenhuyzen et al., 2011). Furthermore the presence of high amounts of suspended solids in most of the sample matrices also impedes nucleic acid extraction (Bass et al., 2015) and may inhibit PCR reactions. Separation of eggs from these solid particles is mostly done before the extraction of DNA (Bass et al., 2015) using flotation and/or sedimentation steps. The method of choice for DNA extraction varies between studies (Raynal et al., 2012;Khouja et al., 2010;Mugambi et al., 2015;Loreille et al., 2001). Table 6 presents commonly used nucleic acid extraction methods, including commercial nucleic acid extraction kits and the molecular application of the product. The main component in these commercial kits is a proteinase enzyme which helps in the lysis of the egg shell and finally an elution in different solutions depending on the manufacturer. Most of these kits are optimized for extraction of bacterial genome but the egg shells of STHs eggs are much tougher than the cell walls of bacteria. Therefore the use of such kits may result in a lower nucleic acid yield. Destruction of the egg shells to release the nucleic material could result in a higher nucleic acid yield, which has been reported with methods using sonication steps (Loreille et al., 2001) and beads (Raynal et al., 2012;Gyawali et al., 2015;Gatcombe et al., 2010).

Polymerase chain reactions
The development of PCR has transformed the detection of microorganisms in different sample matrices. However, the major limitation with conventional PCR is its inability to quantify the microorganisms detected (Zhang and Fang, 2004). There are other interferences with PCR analysis, some of which include the quality of the samples (purity, quantity and integrity of DNA); the inadequate homogenization of reagents used; the inaccurate setting of the baseline and calibration curves; and pipetting and dilution of standards (Rocha et al., 2016). Despite the limitations of PCR methods, they have proven to be more sensitive than microscopy based methods in detecting low numbers of eggs (Oliveira et al., 2010;Guy et al., 2003). Since PCR methods are based on species or group specific gene sequences, the outcome is more specific than morphological methods. The latter sometimes limit species differentiation, for example of hookworm or Ascaris eggs (Valero et al., 2009;Ai et al., 2010). Advances in PCR technology has also made it possible to detect multiple parasites using multiplex PCR by amplifying more than one target of interest using multiple primer pairs (Henegariu et al., 1997). The design and selection of the multiple primer pairs can make the reaction specific for the target organism (Gordon et al., 2011). qPCR has the ability to amplify and simultaneously detect and quantify the DNA amplification in real time (Bass et al., 2015) which is a major advancement over conventional PCR. This real-time detection, important for the absolute quantification of targeted DNA sequences, depends on the flores-cence generated as the reaction proceeds. The level of fluorescence is directly proportional to the quantity of target amplicons accumulated (van Frankenhuyzen et al., 2011). There are several reagents or chemicals available such as intercalating dyes, hydrolysis and hybridization probes (Gyawali et al., 2015), that could be added to facilitate the detection of the end product and aid in the quantification of the parasites. The main advantages associated with qPCR are its sensitivity, specificity, reproducibility and wide quantification range (Gyawali et al., 2015).
Although qPCR emerged as a suitable and powerful method for quantification, its use in environmental sample analysis has limitations. The quantification of low concentration of STH eggs from environmental samples is a challenge and the potential presence of PCR inhibitors in the samples may interfere with the PCR reactions (Toze, 1999;Shannon et al., 2007). Specificity of group-specific primers, florescent probes and nucleic acid extraction efficiency are also critical points (Durant et al., 2012). In addition, the quality of template nucleic acid and amplification of DNA of non-viable STHs eggs are also important factors that affect the analysis (Zeehaida et al., 2011).
Despite the advantages, there are limited environmental studies that have utilized qPCR for detection of STHs. A qPCR method was developed by Pecson et al. (2006) to determine the levels of total and viable Ascaris eggs using the first internally transcribed spacer (ITS-1) region of ribosomal DNA (rDNA) and rRNA. The transcription product of the ITS-1 region is short lived within the cell due to rapid enzymatic degradation and absent in non-viable eggs resulting in an estimate of viability (Pecson et al., 2006). The detection limit of the rDNA-based method was approximately one larvated egg or 90 single-celled eggs whereas the detection limit for the rRNA-based method was 968 single-celled eggs. This is a limitation in areas where the prevalence of infection is very low. This study promotes the applicability of the qPCR method in the detection of viable STHs eggs due to its ability to detect a single larvated egg. Gyawali et al. (2015) also reported a qPCR method for the rapid concentration, sensitive and specific detection of hookworm (Ancylostoma caninum) eggs from wastewater matrices. The morphological similarity of A. caninum to the human hookworms (Ancylostoma duodenale and Necator americanus) makes it a preferred choice to determine the specificity of qPCR in quantification of closely related STHs eggs. The detection limit of this developed qPCR was found to be less than one A. caninum egg per 1 L of secondary treated wastewater, four A. caninum eggs per 1 L of raw wastewater and per ∼4 g of treated sludge. A duplex qPCR assay targeting the ribosomal RNA gene internal transcribed spacer (ITS2) for the detection and differentiation of the eggs of Toxocara canis and Toxocara cati (Nematoda, Ascaridoidea) in soil and fecal samples was reported by Durant et al. (2012). The detection limit of the assay in spiked samples was 2 eggs/g of soil. This study reported that the duplex qPCR could be used for the detection of T. canis and/or T. cati eggs in fecal samples as well as in soil samples. Several other studies have shown that qPCR is much more sensitive than the conventional microscopy based methods in the detection of helminths from different environmental samples (Zeehaida et al., 2011;Sultana et al., 2013;Saugar et al., 2015;Kramme et al., 2011;Schär et al., 2013).

Loop-mediated isothermal amplification
LAMP is a nucleic acid amplification method with extremely high sensitivity and specificity, able to discriminate between single nucleotide differences (Parida et al., 2008). It is characterized by the use of a DNA polymerase that has low sensitivity to inhibitors and a set of four primers specially designed to recognize six different sequences on the target gene (Paris et al., 2007). In this technique amplification occurs only when all primers bind, thus forming a product. It can amplify a few copies of genetic material to 10 9 within an hour (Notomi et al., 2000) thereby reducing the time required for analysis. LAMP is based on an isothermal reaction and only a heating block or hot water bath is required for the reaction to progress as outlined in Fig. 1. There is no need for a thermocycler. Additionally, using white magnesium pyrophosphate results in precipitation of the DNA fragments (Mori et al., 2001) and the turbidity caused by this reaction is proportional to the amount of DNA synthesized. As a result, it is possible to evaluate the reaction in real time by measuring the turbidity or, more importantly, by visualization with the naked eye, making it a suitable candidate technique for field applications (Nkouawa et al., 2009). As illustrated in Fig. 1, LAMP is a simple method that does not require sophisticated equipment and is therefore a cheaper alternative than other nucleic acid based methods (especially the PCRs). It could have potential in develop-ing regions of the world where laboratory resources are scarce. The LAMP method has been used in the detection of T. saginata, T. solium and T. asiatica DNAs in human faecal samples with higher sensitivity (88.4%) than multiplex PCR (37.2%) (Gordon et al., 2011). Liang et al. (2009) also reported a success rate of 92% in using LAMP method to detect Entamoeba histolytica in faecal samples as compared to nested PCR. The method has been applied in other applications in the detection of different parasites in a variety of sample matrices, such as Necator americanus in faecal samples (Mugambi et al., 2015) and detection of Echinococcus granulosus (Salant et al., 2012).

Digital PCR
Droplet digital PCR (ddPCR), a third generation PCR technology, was introduced to provide absolute quantification of targeted genes applicable for pathogens. Digital droplet PCR uses microwells or microfluidic chambers, which are simply referred to as wells, that split samples into several nanoliter partitions (Hindson et al., 2013). The advantages of ddPCR over qPCR-based assays are that ddPCR is based on endpoint PCR (efficiency of primer/probe annealing is minimized) and does not require the use of standards for accurate quantification. Most importantly, ddPCR is a high throughput assay with an approximate 15,000-20,000 PCR reactions per well (Baker and Ensink, 2012). Baker and Ensink (2012) described commercially available digital PCRs based on either a chip or on droplets. The application of droplet digital PCR in the detection of zoonotic pathogens in poultry processing water samples was reported by Rothrock et al. (2013). The results showed that ddPCR out-performed qPCR and culture based methods used for processing poultry zoonotic pathogens. The frequent application of digital PCR (chip-based or droplets) for detection of STHs in environmental samples has not been explored yet. Further research is needed to optimize this method in terms of sensitivity and precision for detection of STHs in environmental samples.

Parasite identification using image analysis software
The final step involved in most conventional methods is microscopy which may be time consuming and dependent on the technician/microscopists' experience. The identification of STHs eggs including all the processing steps and microscopy takes   (Jiménez et al., 2016).
Recently, an automated microscopy step has been developed, where the use of software to identify and count the number of STHs eggs is included (Jiménez et al., 2016;Gomes et al., 2015). This automation technique still requires the separation and concentration of the eggs from the environmental samples although they eliminate human involvement in the final detection and quantification of eggs. In addition, the images of these eggs must be captured with cameras with good resolution for easy analysis with the software's. Jiménez et al. (2016) reports sensitivities between 80 and 100% and close to 100% specificity, while in an earlier publication using software called Parasitology DSS, Gomes et al. (2015) reported that false results may occur. The study by Gomes et al. (2015) reported that 81.86% of parasites were correctly identified by the software and 18.13% could not be identified. The image analysis technique needs further validation with different sample matrices as the presence of debris and poor picture qualities were found to be the main issues leading to false reports (Gomes et al., 2015). An alternative approach that uses fluidic geometries to concentrate eggs into a single field of view (FOV) and combining it with a mobile phone (Nokia Lumia 1020) with digital photo-microscopy has been reported by Sowerby et al. (2016). Combined with an additional objective lens, this photo-microscopy system provided sufficient resolution for single FOV images of nematode eggs due to the extended field of depth which enhances the quality of the images. However, these software identification systems still relies on effective sample processing just as is the case in the more conventional methods. In addition, improved optics are needed to be able to provide good images that would make the use of mobile phones a feasible option in the detection of STHs eggs, which naturally would be very beneficial for low income areas.

Flow cytometry
Flow cytometry simultaneously measures and analyzes multiple physical properties of a single particle, such as cells and potentially eggs/cysts, as they flow in a fluid stream through a beam of light. Properties such as relative size, relative granularity or internal complexity and relative fluorescence intensity are used in the differentiation of one cell from the other. Any suspended particle or cell from 0.2-150 micrometers in size is suitable for analysis (Vesey et al., 1997). STHs eggs could be analyzed using flow cytometry, but there is no report of its use in the detection of STHs eggs either from clinical or environmental samples. However, flow cytometry has been used in the detection of Cryptosporidium parvum oocysts in water samples (Vesey et al., 1994;Hoffman et al., 1997;Power et al., 2003) to determine experimental parasite loads in mice (Arrowood et al., 1995) and to enumerate viable oocysts (Vesey et al., 1997;Campbell et al., 1993).
Since STHs eggs fall within the cell sizes that could, in theory, be analyzed with flow cytometry, it may be possible to use this technique in the detection and quantification of these eggs. Flow cytometry is able to analyze cells based on size whereas the different STHs species could be differentiated based on their sizes. In addition, it may be possible to determine the viability of eggs. This technique is able to differentiate between cells based on complexity which is dependent on the internal structures of the cell. As STHs eggs mature there is an increase in cell numbers in each egg. This increases the complexity and it may be possible to estimate the stage of development of the egg and therefore predict its viability. Further analysis could be achieved through staining with fluorescent dyes (El-Kowrany et al., 2015;Beers et al., 2015), which could make it possible to differentiate viable eggs from non-viable eggs through differential staining. The BacLight Live/Dead staining procedure has been coupled with flow cytometry to determine viability of bacterial cells (Schumann et al., 2003;Kramer et al., 2009;Berney et al., 2007), and since this technique has been used in the determination of viable STHs eggs, (Dabrowaska et al., 2014;Karkashan et al., 2015) it could be coupled with flow cytometry to adequately determine the viability of STHs eggs. The main limitation to the widespread use of flow cytometry is the cost involved in the purchase and use of the equipment which hinders its routine use especially in developing countries.

Conclusion
Environmental samples pose a challenge for the detection and quantification of STHs eggs. The development of methods/techniques which are applicable to the variety of sample matrices is imperative for uniformity. Many of the methods in use have analytical steps that are similar from method to method. It can be concluded from this review that some of the major steps that influence egg recovery are, but not limited to, filtration/sieving, egg recovery from particles, sedimentation, flotation and microscopy. Microscopy which is the last step in most conventional methods could also introduce an element of variability due to human subjectivity and experience. Other main challenges of conventional methods as presented in this review is the time needed to process and analyze samples, the additional identification of STHs eggs of similar species or differentiation of eggs of the same genus into different species. The development of molecular techniques has shown the potential to solve many of the challenges of the conventional techniques, but these techniques have shortfalls as well. The high cost involved in sample analysis is the main challenge with the use of molecular techniques for routine use. However the LAMP method promises to be a fast and cost-effective technique for the molecular detection of STHs eggs in environmental samples. In addition, there are new and emerging techniques that potentially could be used for the efficient and sensitive detection and quantification of STHs eggs in environmental samples. Some of these new techniques are flow cytometry, LAMP, digital droplet PCR and the use of image analysis softwares. There is the need to validate and compare all techniques to determine their applicability in environmental samples. For instance, the use of conventional methods is cheap and more suited to laboratories found in lowincome countries. However, the conventional methods as shown in Table 7 are time consuming, laborious and may result in systematic errors, which could be reduced with molecular methods. These can be rapid, sensitive and identify the STHs down to species level; a feature that is difficult with conventional methods. Despite the aforementioned advantages of the molecular methods, these are expensive due to the high cost of equipment and consumables and the need for constant supply of electricity, which is a challenge in developing countries. The improvements in the molecular techniques such as the LAMP method could considerably reduce the cost involved in analysis and provides an opportunity for more rapid, sensitive and cheap means of analysis.
The validation and subsequent comparison of techniques would help determine the most cost-effective and efficient technique for uniform detection and quantification of STHs eggs in environmental samples.