Page 5: Guidelines for Canadian Drinking Water Quality: Guideline Technical Document - Enteric Protozoa: Giardia and Cryptosporidium
Partie II. Science et considérations techniques
The most widely recognized and used method for the detection of Giardia and Cryptosporidium in water is the U.S. Environmental Protection Agency's (EPA) Method 1623, as this method allows for the simultaneous detection of these protozoa and has been validatedin surface water (U.S. EPA, 2005, 2006a). Although other methods for the detection of Giardia and Cryptosporidium in waterexist, they have demonstrated lower recoveries and increased variance compared with EPA Method 1623 (Quintero-Betancourt et al., 2002). Like most methods used for the detection of Cryptosporidium and Giardia in water, EPA Method 1623 consists of four steps: 1) sample collection, 2) sample filtration and elution, 3) sample concentration and separation (purification) and 4) (oo)cyst detection. These steps are described in the following sections. Some emerging detection methods are also discussed, along with methods used for assessing (oo)cyst viability and infectivity.
Water samples can be collected as bulk samples or filtered in the field and then shipped on ice to a laboratory for processing as quickly as possible (ideally, within 24 hours). The volume of water collected depends on the expected level of (oo)cysts in the water (i.e., site specific); the lower the expected density of (oo)cysts, the greater the sample volume needed. In most cases, between 10 and 1000 L of water are collected. In the case of raw water, samples are typically collected near and at the depth of the drinking water intake point, in an effort to sample the source water used for supplying drinking water.
(Oo)cysts are generally present in small numbers in faecally contaminated water; as such, bulk water samples must be filtered to concentrate the pathogens to a detectable level. Typically, water is pumped through a filter, and (oo)cysts, along with extraneous particulate materials, are retained on the filter. Filtration can be achieved using a variety of filter types, including wound filters, membrane filters, hollow fibre filters and compressed foam filters. These filters vary in terms of the volume of water that they can process, their filtration rates, their practicality, their compatibility with subsequent processing steps, their cost and their retention ability. These differences account for the wide range of recovery efficiencies reported in the literature (Sartory et al., 1998; DiGiorgio et al., 2002; Quintero-Betancourt et al., 2003; Ferguson et al., 2004). A number of filters have been validated by EPA Method 1623 (U.S. EPA, 2005). Once filtration is complete, entrapped (oo)cysts on the filter are released through the addition of eluting solutions, producing a filter eluate.
(Oo)cysts in the filter eluate are further concentrated through centrifugation and separated from other particulates through immunomagnetic separation (IMS)/immunocapture. Alternatively, flotation (i.e., density gradient centrifugation) can be used for (oo)cyst separation; however, this approach has been associated with significant (oo)cyst losses and does not effectively remove other biological materials (e.g., yeast and algal cells) (Nieminski et al., 1995), which may affect subsequent (oo)cyst detection.
The partially concentrated (oo)cysts are then centrifuged, resulting in the formation of a pellet. This pellet is resuspended in a small volume of buffer. The concentrate is mixed with (oo)cyst-specific monoclonal antibodies attached to magnetized beads, also referred to as immunomagnetic beads. These beads will selectively bind to (oo)cysts. A magnetic field is then applied, resulting in the separation of (oo)cyst-bead complexes from extraneous materials. These materials are removed, the (oo)cyst-bead complex is dissociated and the beads are extracted, resulting in a concentrated suspension of (oo)cysts. Several studies have assessed the recovery potential of the IMS step alone. Fricker and Clancy (1998) reported that (oo)cysts added to (i.e., seeded into) low-turbidity waters can be recovered with efficiencies above 90%. In comparison, mean oocyst and cyst recoveries for turbid waters ranged from 55.9% to 83.1% and from 61.1% to 89.6%, respectively, for turbid waters (McCuin et al., 2001). Others have reported similar recoveries (Moss and Arrowood, 2001; Rimhanen-Finne et al., 2001, 2002; Sturbaum et al., 2002; Ward et al., 2002; Chesnot and Schwartzbrod, 2004; Greinert et al., 2004; Hu et al., 2004; Ochiai et al., 2005; Ryan et al., 2005b). Although IMS aids in reducing false positives by reducing the level of debris on slide preparations for microscopic analysis, it is a relatively expensive procedure, with few manufacturers supplying the immunomagnetic beads. Moreover, it has been reported that high levels of turbidity and/or iron (Yakub and Stadterman-Knauer, 2000), along with changes in pH (i.e., optimum pH of 7) (Kuhn et al., 2002), may inhibit IMS.
Once samples have been concentrated and (oo)cysts have been separated from extraneous materials, a number of detection techniques can be applied. The most commonly used detection approach is the immunofluorescence assay (IFA). Alternative detection methods, such as the polymerase chain reaction (PCR), flow cytometry and other molecular approaches, are increasingly being used. Molecular detection methods are generally more rapid and sensitive and have the potential of being paired with a variety of other methods to provide species/genotype information. However, only small volumes can be processed using these methods, and some methods (e.g., PCR) are susceptible to environmental inhibitors.
Following sample concentration and separation, a portion of the (oo)cyst suspension is transferred to a microscope slide. Fluorescently labelled antibodies directed at specific antigens on the (oo)cyst surface are then applied to the slide and allowed to incubate. Direct immunofluorescence microscopy is then used to locate fluorescing bodies, which are potential (oo)cysts. This process, referred to as an IFA, requires specialized equipment and a high level of technical skill. It can be highly sensitive, however, because some autofluorescent algae are very close in size and staining characteristics to (oo)cysts; final identification of (oo)cysts often requires additional staining and microscopy. In most cases, a DAPI stain is applied. Because DAPI binds to deoxyribonucleic acid (DNA), it will highlight (oo)cyst nuclei and facilitate their identification.
Flow cytometry can be used as an alternative technique for detecting (oo)cysts following concentration. Flow cytometry allows the sorting, enumeration and examination of microscopic particles suspended in fluid, based on light scattering. Fluorescently activated cell sorting (FACS) is the flow cytometric technique that is used to enumerate and separate Cryptosporidium and Giardia from background particles. Typically, immunofluorescent antibodies are introduced into the (oo)cyst suspension, and the suspension is passed through a beam of light (within the flow cytometer). As particles pass through the stream of light, their fluorescence is measured, and they are then sorted into two or more vials.
FACS has proven to be highly sensitive and specific and is being used more and more as an alternative (oo)cyst detection technique (Vesey et al., 1997; Bennett et al., 1999; Reynolds et al., 1999; Delaunay et al., 2000; Lindquist et al., 2001; Kato and Bowman, 2002; Lepesteur et al., 2003; Hsu et al., 2005). This approach has the advantage of being rapid, allowing for high throughput. However, flow cytometers are expensive, and their operation requires significant user training. In addition, like IFA, this procedure can be adversely influenced by the presence of autofluorescent algae and antibody cross-reactivity with other organisms and particles. FACS also requires confirmation of (oo)cysts by microscopy, which is why it is often coupled with EPA Method 1623. Although FACS shows promise, it is still in the development stage and is not used for routine analysis.
A number of molecular approaches have also been used in the detection of Giardia and Cryptosporidium (oo)cysts. A brief description of some of these methods is provided below. It is important to note that although molecular methods have many advantages, they also possess significant disadvantages that make them unsuitable for routine analysis of water. There are currently no validated molecular methods for the detection of Giardia and Cryptosporidium in water.
PCR is the most commonly used molecular method for detection of (oo)cysts. This method involves lysing (oo)cysts to release DNA and then introducing primers that are targeted at specific Giardia or Cryptosporidium coding regions (e.g., 18S ribosomal ribonucleic acid [rRNA]) and amplification of these regions. PCR can be highly sensitive (i.e., level of a single (oo)cyst) and specific (Deng et al., 1997, 2000; Bukhari et al., 1998; Di Giovanni et al., 1999; Kostrzynska et al., 1999; Rochelle et al., 1999; Hallier-Soulier and Guillot, 2000; Hsu and Huang, 2001; McCuin et al., 2001; Moss and Arrowood, 2001; Rimhanen-Finne et al., 2001, 2002; Sturbaum et al., 2002; Ward et al., 2002). It can be combined with other molecular techniques, such as restriction fragment length polymorphism (RFLP), to discriminate between species and genotypes of Cryptosporidium and Giardia (Morgan et al., 1997; Widmer, 1998; Lowery et al., 2000, 2001a,b), although this approach can be problematic, in that it can produce similar banding patterns for different species and genotypes. PCR is also amenable to automation, and reverse transcriptase (RT) PCR may permit discrimination of viable and non-viable (oo)cysts. However, PCR inhibition by divalent cations and humic and fulvic acids is a significant problem (Sluter et al., 1997). In an effort to remove these inhibitors, samples must go through several purification steps. In addition to inhibition, inefficient (oo)cyst lysis is often an issue. Despite these problems, many PCR assays have been developed for detection of waterborne (oo)cysts (Stinear et al., 1996; Kaucner and Stinear, 1998; Griffin et al., 1999; Lowery et al., 2000; Gobet and Toze, 2001; Karasudani et al., 2001; Ong et al., 2002; Sturbaum et al., 2002; Ward et al., 2002).
Other emerging molecular methods for detection of (oo)cysts include fluorescence in situ hybridization (FISH), real-time PCR and microarrays. FISH involves hybridizing a fluorescently labelled oligonucleotide probe that is targeted at the 18S rRNA region of Giardia and Cryptosporidium. This technique has shown some success, but it is limited by relatively weak signals (i.e., (oo)cysts do not fluoresce sufficiently) and related difficulties in microscopic interpretation (Deere et al., 1998; Vesey et al., 1998; Dorsch and Veal, 2001). Real-time PCR is a modified PCR that involves oligonucleotide probes that fluoresce. As the target region within (oo)cysts is amplified, the emitted fluorescence is measured, thereby allowing quantification of the PCR products. This method has several advantages, including the lack of post-PCR analysis, increased throughput, decreased likelihood of contamination (i.e., closed vessel system), ability to quantify (oo)cysts (MacDonald et al., 2002; Fontaine and Guillot, 2003; Bertrand et al., 2004) and ability to assess (oo)cyst viability (when paired with cell culture) (Keegan et al., 2003; LeChevallier et al., 2003). This approach has other unique advantages, including its ability to differentiate between species of Cryptosporidium and Giardia (using melting curve analysis) (Limor et al., 2002; Ramirez and Sreevatsan, 2006) and simultaneously detect different microorganisms (Guy et al., 2003). Although this assay has several advantages over traditional PCR and IFA and has proven useful in identification and enumeration of (oo)cysts, it requires a real-time PCR analyser, which is very costly and may limit its widespread use. Microarrays represent a very novel approach to (oo)cyst detection. A microarray is a collection of microscopic DNA spots, usually on a glass slide, against which pathogen DNA is hybridized. This approach has proven useful in the detection and genotyping of Giardia and Cryptosporidium (Straub et al., 2002; Grow et al., 2003; Wang et al., 2004), although more research is required to determine its specificity and sensitivity.
An integral part of the Giardia and Cryptosporidium detection process involves determining recovery efficiencies. As mentioned previously, there can be significant losses of (oo)cysts during the concentration and separation processes. In addition, the characteristics of the water (e.g., presence of suspended solids, algae) can have a significant impact on recovery efficiency. As a result, the true concentration of (oo)cysts in a water sample is almost always higher than the measured concentration. Thus, recovery efficiencies are determined to better approximate the actual concentration of (oo)cysts. The recovery efficiency is generally measured by introducing a known number of (oo)cysts into the water sample (i.e., seeding) before the sample is analysed. Ideally, the recovery efficiency should be determined for each sample; however, because this is expensive, recovery efficiency data are usually collected for a subset of samples. With the introduction of commercial preparations containing a certified number of (oo)cysts, this process has become more cost-effective and routine.
Several studies have evaluated the recovery efficiencies achieved using EPA's Method 1623 with different types of filters (McCuin and Clancy, 2003; Ferguson et al., 2004; Hu et al., 2004; Wohlsen et al., 2004; Karim et al. 2010). Recoveries ranged significantly and correlated with variations in raw water quality, highlighting the importance of an internal control with each water sample.
A major drawback of existing methods for the detection of Giardia and Cryptosporidium is that they provide very limited information on the viability or human infectivity of (oo)cysts, which is essential in determining their public health significance. Whereas viability can be assessed relatively easily and rapidly, assessment of infectivity is much more complex. Methods used to evaluate viability and infectivity are very costly because of the need for maintaining cell lines, animals and qualified staff; as a result, they are not typically applied to the assessment of (oo)cysts.
A variety of in vitro and in vivo methods have been developed to assess viability and infectivity. In vitro methods include excystation, fluorogenic dye inclusion/exclusion (i.e., staining), reverse transcriptase-polymerase chain reaction (RT-PCR) and fluorescence in situ hybridization (FISH). In vivo methods include animal infectivity assays and cell culture. A brief discussion of these methods is provided in the following sections.
Viability (but not infectivity) can be estimated by subjecting (oo)cysts to conditions similar to those in the gut, in an effort to stimulate excystation (i.e., release of trophozoites/sporozoites). Excystation "cocktails" and conditions vary considerably and may result in conflicting observations. If (oo)cysts are capable of excystation, they are considered viable. Giardia can be excysted using acid and enzymes such as trypsin and grown in TYI-S-33 medium (Diamond et al., 1978; Rice and Schaefer, 1981), but the excystation rate for Giardia is often low. Cryptosporidium parvum oocysts can also be excysted as a measure of viability (Black et al., 1996). However, excystation methods have been shown to be relatively poor indicators of Cryptosporidium oocyst viability. Neumann et al. (2000b) observed that non-excysted oocysts recovered after commonly used excystation procedures are still infectious to neonatal mice.
Various staining methods have been developed to assess (oo)cyst viability, based on the inclusion or exclusion of two fluorogenic dyes, DAPI and PI (Robertson et al., 1998; Freire-Santos et al., 2000; Neumann et al., 2000b; Gold et al., 2001; Iturriaga et al., 2001). Three classes of (oo)cysts can be identified: 1) viable (inclusion of DAPI, exclusion of PI), 2) non-viable (inclusion of both DAPI and PI) and 3) quiescent or dormant (exclusion of both DAPI and PI, but potentially viable). In general, DAPI and PI give good correlation with in vitro excystation (Campbell et al., 1992). Neumann et al. (2000a) demonstrated a strong correlation between DAPI/PI staining intensity and animal infectivity of freshly isolated C. parvum oocysts. These stains have also been successfully used in conjunction with fluorescently labelled antibodies (used in FACS) to determine the viability and infectivity of (oo)cysts in water samples, because their fluorescence spectra do not overlap with those of the antibodies (Belosevic et al., 1997; Bukhari et al., 2000; Neumann et al., 2000b). In spite of these positive correlations, dye inclusion/exclusion, like excystation procedures, overestimates the viability and potential infectivity of (oo)cysts (Black et al., 1996; Jenkins et al., 1997).
RT-PCR can also be applied to the direct detection of viable (oo)cysts in water concentrates (Kaucner and Stinear, 1998). RT-PCR amplifies a messenger ribonucleic acid (mRNA) target molecule. As only viable organisms can produce mRNA, this experimental method may prove useful in assessing (oo)cyst viability. For example, when compared with the IFA DAPI/PI method, the frequency of detection of viable Giardia increased from 24% with IFA to 69% with RT-PCR. An advantage of this approach is that it can be combined with IMS, allowing for simultaneous detection and viability testing (Hallier-Soulier and Guillot, 2000, 2003); it can also be quantitative. RT-PCR, like other PCR-based methods, is highly susceptible to environmental inhibition and suffers from inefficient extraction of nucleic acids from (oo)cysts.
FISH has shown modest success in differentiating between living and dead (oo)cysts (Davies et al., 2005; Lemos et al., 2005; Taguchi et al., 2006); however, false positives are common (Smith et al., 2004). As 18S rRNA is present in high copy numbers in viable (oo)cysts but in low numbers in non-viable (oo)cysts, it is a useful target for assessing viability. Further research is required to validate this assay for use in assessing (oo)cyst viability. Like DAPI/PI staining, FISH is limited by its inability to assess (oo)cyst infectivity. Further research is required to validate this assay for use in assessing (oo)cyst viability.
The most direct method for assessing (oo)cyst viability and infectivity is to inoculate a susceptible animal and monitor for (oo)cyst shedding and any histological evidence of disease development. Giardia and Cryptosporidium are used to infect experimental animals such as the gerbil (for Giardia) (Belosevic et al., 1983) and the neonatal CD-1 mouse (for Cryptosporidium) (Finch et al., 1993). This approach has shown moderate success (Delaunay et al., 2000; Korich et al., 2000; Matsue et al., 2001; Noordeen et al., 2002; Okhuysen et al., 2002; Rochelle et al., 2002), but it is not practical, as most analytical laboratories do not maintain animal colonies, and animal infectivity assays are expensive to perform. In addition, there is limited knowledge on the diversity of species and genotypes of Giardia and Cryptosporidium that can infect animal models (i.e., some species/genotypes may not be infectious for a particular animal host). Even with this information, this approach is not sensitive enough for environmental monitoring (i.e., high median infective dose [ID50]). These assays are typically reserved for research purposes, such as assessing disinfection effectiveness, rather than for routine assessment of (oo)cyst viability/infectivity.
Unlike Giardia, Cryptosporidium is an intracellular parasite that relies on host cells for replication. Thus, oocysts cannot be grown in cell-free culture media. In vitro cell culture assays for Cryptosporidium infectivity assessment overcome several of the limitations associated with the use of animal models. These assays involve exposing oocysts to excystation stimuli followed by their inoculation into a cultured mammalian cell line, such as human ileocaecal adenocarcinoma (HCT-8) cells, which support the parasite's growth and development. Oocysts are typically inoculated on HCT-8 cell monolayers. After a 24- to 48-hour incubation, the cell monolayer is examined for the presence of Cryptosporidium reproductive stages using either an indirect IFA (Slifko et al., 1997) or PCR (Rochelle et al., 1997).
This approach has been used to estimate the infectivity of oocysts in water (Di Giovanni et al., 1999; Hijjawi et al., 2001; Weir et al., 2001; Rochelle et al., 2002; Johnson et al., 2005; Schets et al., 2005; Coulliette at al., 2006) and has been shown to yield results comparable to those of the mouse infectivity model (Hijjawi et al., 2001; Rochelle et al., 2002; Slifko et al., 2002). In other comparison studies, average percent viabilities were comparable for cell culture, excystation and DAPI/PI assays (Slifko et al., 1997).
There are several advantages to the cell culture assay, including its high sensitivity (i.e., detection of a single viable oocyst), applicability to analysis of raw and treated water samples, ease of performance and rapid turnaround time for results. Another advantage of this approach is that C. parvum and C. hominis can be maintained in vitro for long periods of time, facilitating viability and immunotherapy studies. In addition, cell culture can be combined with other methods, including PCR, to more accurately assess viability/infectivity. Cell culture PCR (CC-PCR) has proven useful in assessing watershed contamination and in estimating risk (Joachim et al., 2003; LeChevallier et al., 2003; Masago et al., 2004). Although cell culture infectivity assays have several advantages, they also possess a number of disadvantages, including the need to maintain a cell line and poor reproducibility among similar samples for quantitative assessments. Moreover, existing cell culture methods detect only C. parvum and C. hominis; very little is known about how other Cryptosporidium species and genotypes of human health concern infect culture systems. The development of C. parvum in a host cell-free culture was recently reported (Hijjawi et al., 2004), but could not be reproduced (Girouard et al., 2006).
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