Practical Guidance for Clinical Microbiology Laboratories: Laboratory Diagnosis of Parasites from the Gastrointestinal Tract

There are many different clinical manifestations of parasitic infections of the gastrointestinal tract. The presentations of these parasitoses vary depending on the infecting parasite, as well as on a variety of host factors that are incompletely understood. However, it is clear that the patients with severely compromised immune responses usually have more-severe disease. These patients are also at risk for infections by parasites that do not commonly cause disease in immunocompetent individuals (11). The duration of parasitosis and the load of parasites also affect the clinical manifestations of disease.

The prevalences of the different types of gastrointestinal parasitic infection that are encountered vary by locale. This variation is impacted largely by the mechanism of transmission, the number of parasitized individuals in the area, the adequacy of public health measures to handle human and animal waste, and the ability of public health measures to provide clean drinking water for inhabitants of the area (27, 28). The prevalence of many gastrointestinal parasitic infections is therefore great in resource-poor countries that have a high burden of disease and inadequate public health facilities to handle waste and provide clean drinking water (28).

The mechanism of transmission is important for predicting which types of parasites are likely to be encountered. For example, one expects to encounter patients infected with Enterobius vermicularis in both resource-rich and resource-poor countries given that the eggs are infectious soon after passage and child-to-child transmission is possible either directly or through fomites (i.e., it does not matter if the children sharing toys are in Manhattan or in sub-Saharan Africa). In contrast, one is far less likely to encounter hookworm infections in locales where shoes are common than in areas where the citizens are often barefooted.

The number of parasitized individuals in the community affects the likelihood of infection or reinfection due to the increased number of opportunities for infection. For example, a child with pica in an area of low endemicity is less likely to acquire an infection by a geohelminth, such as Trichuris or Ascaris, than a similar child in an area where parasites are highly endemic because they are more likely to encounter parasite eggs in the dirt. Additionally, the areas that have large numbers of infected individuals are often resource poor and unable to appropriately handle human waste, so contamination of the environment and subsequent infections become the norm.

Resource-rich countries that adequately handle human waste significantly diminish the likelihood that parasitic cysts and eggs that originate from a human source will contaminate the environment, the food supply, or the drinking water. These countries forbid the use of human waste as fertilizer. There are nonhuman sources of some gastrointestinal pathogens (e.g., Cryptosporidium oocysts derived from animals) that may contaminate the water supply (29). Therefore, it is imperative that in addition to wastewater treatment being employed, modern drinking water treatment be employed. Failures in the system responsible for clean drinking water, which we often take for granted, demonstrate how important these systems are in public health. An often-cited example is that of the drinking water sources in Milwaukee, WI, in 1993 that were contaminated with Cryptosporidium hominis from human sewage effluent that impacted the drinking water facility intake, rather than water runoff from bovine feces (30). This contamination overwhelmed the clean drinking water facility's ability to inactivate the oocysts of Cryptosporidium and caused the largest waterborne outbreak by this parasite in U.S. history.

When the factors described above are considered, the list of gastrointestinal pathogens that one may expect to find in a citizen of the United States or another resource-rich country is very different from those encountered in an infected individual from a resource-poor region of the world where parasites are highly endemic. Trends in immigration may bring patients from countries where parasites are endemic who present with otherwise-rare or -infrequent pathogens. Additionally, travel is easier than ever, and adventurous excursions can place individuals at risk for infections that are uncommon in their home locale. Thus, taking a thorough clinical history is key (see below).

The expansion of medical knowledge in the past decade is incredible. The medical profession has responded through increased specialization and subspecialization. In the past, a surgeon might specialize as an orthopedic surgeon, whereas now, it is common to find practices with individuals who specialize in only knee or hip disease. Therefore, it is unreasonable to expect individuals who are not subspecialty trained in microbiology or infectious diseases to keep abreast of changes in clinical microbiology, one of the fastest-paced fields.

A clinical parasitologist or clinical microbiologist with expertise in parasitology is perfectly positioned to help educate physicians and provide guidance in test selection. These laboratorians, whenever possible, should participate in educating the next generation of physicians, not just to teach them at that point in their career but also to inform them that highly trained laboratorians remain available to assist them as needed throughout their professional careers. Additionally, this group should participate, whenever possible, in medical technology training programs. This training should go beyond the basic training of specimen processing, testing, and results reporting and should include preparing the technologist for his/her role as an integral member of the health care delivery team. Practicing medical technologists are the “front lines” and can notify and work with the laboratory director when unsuspected findings are discovered or untoward events occur. This type of engagement translates into improved patient care.

A clinical microbiologist should work with the medical staff to formulate the test requisition forms, which are largely becoming solely electronic. They should work to aid clinicians in finding and using the most appropriate test for the clinical scenario encountered (see below). They should play an active role in monitoring test utilization and use instances of inappropriate utilization as opportunities for education.

The importance of location in determining the type of parasite that the patient may have acquired has been noted above and should be disclosed as part of taking a thorough history. It remains remarkable after many years of practice and attendance in infectious diseases/microbiology teaching conferences how often the clues to the definitive diagnosis were present in the clinical history or, unfortunately, should have been present had a thorough clinical history been taken.

The clinical history is designed to discover epidemiologic risk factors that are important for guiding testing. In addition to general aspects of a history assessment, specific questions concerning past medical history, countries of previous residence, travel, outdoor activities, family, food, and drinking water should be addressed. Specific examples of the importance of each of these follow.

Where a patient lives or has lived is important for an assessment of the risk of having acquired parasites in the patient's native country that are not endemic in their current country (Cyclospora cases have been reported from the United States, Canada, and the United Kingdom). It is very common to find evidence of multiple gastrointestinal parasites in the stools of children who have been adopted from a resource-poor country where parasites are highly endemic into a low-prevalence, resource-rich country (31). If the history did not include the location of prior citizenship, then an ova and parasite examination (O&P) of the stool may not have been performed, as the individual may have been asymptomatic. Note that, technically, parasite eggs rather than ova are examined; however, the abbreviation O&P remains conventionally accepted. Although there are no accepted general guidelines in these cases, routine parasitology examinations (ova and parasites) may be an appropriate option. Travel history similarly discloses potential risks to the traveler (Cyclospora, Mexico), as does questioning about specific outdoor activities (Cryptosporidium, swimming in late summer, contact with calves). A history of backpacking and drinking stream water is classically associated with giardiasis, for example. Family history and past medical history are important to disclose inherited genetic diseases or other conditions that may put patients at increased risk for certain parasitic diseases. For example, individuals with common variable immunodeficiency are at increased risk for Giardia lamblia (G. duodenalis, G. intestinalis) infections, which tend to be severe (32). Food and drink histories are among the most important, since many parasitic infections are acquired through ingestion. For example, the discovery that a patient is a bear hunter would make one consider trichinellosis, when in the absence of a classical presentation it might otherwise not be considered (33).

Tests in the clinical parasitology section, with the exception of rapid immunoassays, are manual, are time-consuming, and require personnel expertise. Therefore, to preserve limited resources, these tests should be ordered judiciously. As noted above and in Table 2, designing a user-friendly test menu that guides the clinician to the appropriate test is important but has become challenging since in some electronic medical record systems tests may simply be listed alphabetically, requiring the providers to “hunt and peck” to find the right test. Additionally, some tests may sound alike, without differences clearly delineated. Therefore, it is not surprising that providers may select an inappropriate test. Test menu design is an area that is often not given due consideration and is therefore responsible for many unnecessary orders. Some laboratories may elect to use a case history form to guide appropriate testing (Fig. 1).

It is also useful to periodically monitor who in the practice is ordering which tests. Most of the electronic order entry systems have the ability to create “order sets” for providers. Order sets decrease the time needed to search for individual tests. Unfortunately, these sets are often wasteful and filled with more tests than are needed. Order sets are not frequently curated and kept current. By way of example, if a primary care physician who sees predominantly individuals from areas where parasites are not endemic had a standard O&P in their “infectious diarrhea” order set, then the more-labor-intensive O&Ps would be regularly ordered when they would likely have been satisfied with the Giardia lamblia (G. duodenalis, G. intestinalis) immunoassay (IA) screening option.

There are a number of interventions that can be used to decrease unnecessary orders. Tailoring the order set for the pathogens most likely to be encountered is one approach, while educating clinicians is another option. For example, a key take-home message is that a patient from an area of endemicity or an immunocompromised patient may warrant additional tests.

A number of interventions can be useful in averting unnecessary testing, which includes testing for parasites. One of these is to electronically block same-day duplicate orders, should they occur (35). Another is to electronically block orders for O&Ps and stool cultures for patients who have been hospitalized longer than 3 days. In both instances, the clinician can override the electronic blockade by contacting the laboratory (2–6, 10). These interventions have diminished the number of orders that have been placed without thoughtful consideration, while still allowing clinicians to order the test if they really believe that it is clinically necessary.

The number of patients with compromised immune systems in our health care system continues to increase, due to the greater longevity of transplant recipients, better virologic control in human immunodeficiency virus (HIV)-infected individuals, and the use of newer “biologic” immunomodulating agents for diseases such as Crohn's disease, rheumatoid arthritis, and psoriasis. These patients are at risk for more-severe disease caused by commonly encountered agents, as well as disease caused by pathogens less commonly encountered in an immunocompetent host. Therefore, when investigating the cause of a likely infectious diarrheal syndrome in an immunocompromised host, both common and less common agents should be considered (5, 36). Although a patient is immunocompromised, an epidemiologic exposure history is still important to obtain, as this may disclose the most likely pathogen. Providers may choose to begin their investigation with the Giardia/Cryptosporidium immunoassay, given the excellent sensitivity of this assay and the inclusion of both a common pathogen (i.e., Giardia) and an organism known to infect immunocompromised hosts (i.e., Cryptosporidium). If negative, a clinical history will suggest the likelihood of a positive finding in the standard O&P. Both the modified acid-fast stain and a standard O&P will disclose the presence of Cystoisospora, an important pathogen in immunocompromised hosts, as well as Cyclospora, a foodborne pathogen associated with travel and the consumption of imported produce. Finally, if the cause remains undetermined, one should consider an assessment of Microsporidia. Although Microsporidia are taxonomically now classified as fungi, most testing remains in the parasitology sections of the laboratory (5). If the morphological assessment for Microsporidia is negative and clinical suspicion remains high, then a PCR analysis should be considered because of its superior sensitivity.

Clinical laboratories should follow the requirements related to standard precautions, which state that all patients and all laboratory specimens are potentially infectious and should be handled accordingly (37–40). “Standard precautions” replaces earlier terms such as “blood and body fluid precautions,” “universal precautions,” and “body substance precautions” found in Occupational Safety and Health Administration (OSHA) documents. While the OSHA documents place the emphasis on blood-borne pathogens, such as HIV and hepatitis B and C viruses, standard precautions recognize that all infectious agents and all other potentially infectious material, except sweat, pose a risk to the health care worker (37).

Methods include those used to minimize exposure to infectious agents, to shield the laboratory worker from infectious material through a set of engineering and work practice controls, and to use personal protective equipment (PPE). In situations where differentiation between body fluid types is difficult or impossible, all body fluids shall be considered potentially infectious.

Also, the OSHA regulations require that employers provide hepatitis B vaccination and postexposure evaluation and follow-up, communicate the hazards to employees, and maintain appropriate records (40). Employees who decline immunization against hepatitis B virus are required to sign a hepatitis B vaccine declination form.

Unfortunately, recruitment and retention of qualified individuals are major problems for most clinical laboratories throughout the United States and many other countries (42–45). The closure of training programs for medical laboratory scientists (MLS) and medical laboratory technicians (MLT) has contributed to this shortage; many of these closures were related to financial constraints. Between 1983 and 2009, approximately 64% of MLS programs were closed (42, 46). The number of MLS candidates passing the American Society of Clinical Pathology (ASCP) certification exam decreased from 6,000 in 1983 to a low of 1,892 in 2005 (http://www.ascp.org/certification). In spite of extensive recruitment from high school through college and the opening/reopening of training programs, there remains a large shortage of trained laboratory personnel. Most training programs are designed with two different segments, the first of which includes didactic training in various diagnostic disciplines and the second of which includes hands-on clinical training. Often, due to existing personnel shortages, the clinical laboratory training sites can no longer support the extensive teaching required for the clinical rotation portion of the training program.

Diagnostic decision making in clinical medicine is very dependent on clinical laboratory testing. The number, type, complexity, and cost of parasitology tests are growing, possibly creating more confusion regarding which tests are most appropriate at a time when emphasis is on reducing health care costs. This has led to scrutinizing the overordering of tests and ordering tests for the wrong purpose. Thus, it is important that physicians and laboratory staff communicate with each other to understand which laboratory tests are necessary and appropriate for diagnosis and treatment. In order for clinicians to maximize the expertise and resources of the laboratory staff, clinicians must provide accurate and relevant information, including the following. Additional information can be found in Fig. 1.

Clinicians may not have the in-depth knowledge that is necessary to choose the appropriate parasitology test. There are several approaches that a laboratory can provide to facilitate selecting the appropriate laboratory test to establish a diagnosis. One very useful method includes medical algorithms. They are logical and sequential and can be automated and incorporated into software programs, as designed by medical microbiologists. Educating clinicians can be accomplished in a number of ways.

Computer and test result comments can serve as excellent teaching aids for clinicians, many of whom may have little-to-no training in medical parasitology. When using these comments, it is recommended that clinicians use “canned” consistent comments (a template) rather than entering free-text comments. Specific examples can be seen in Appendix 2 (see Tables A5 and A6).

As travel has become more accessible, the rate of tropical infections across the world is expected to increase; more health care professionals both at home and abroad are going to encounter these diseases more often. Disorders of parasitic etiology will play an important and expanding role in all aspects of medicine. Often, pathology personnel may be involved in training various groups of physicians, residents, and/or fellows regarding clinical aspects of medical parasitology diagnosis. In many cases, these students have had very limited exposure to these gastrointestinal protozoan and helminth infections. Tables for training can be found in Appendix 3, while a list of general medical parasitology references can be found in Appendix 4.

The test that is to be requested depends on which infectious agent is suspected. It is very important that the ordering physician be aware of what procedures are performed with the O&P and that this test may not recover all parasites. Additional stains for Microsporidia and Cyclospora/Cystoisospora/Cryptosporidium may need to be ordered separately, and immunoassays for selected organisms may also not be included in the O&P.

Collection of stool for parasite examination should always be performed before barium is administered to a patient for radiologic exams. Stool specimens containing the opaque, chalky suspension are unacceptable, and intestinal protozoa may be masked for 5 to 10 days after ingestion of barium. Other substances, such as castor oil or mineral oil, bismuth, antibiotics, including antimalarial medication, and nonabsorbable antidiarrheal preparations, may interfere with parasite recovery, and collection should be postponed for 5 to 10 days after administration to allow clearance of these substances.

There are many stool collection methods available for specimens suspected of containing parasites. When collection methods are selected, a thorough understanding of the advantages and disadvantages of each must be reviewed. Unless the stool specimens are properly collected and processed, these infections may not be detected. Therefore, specimen rejection criteria have become very important for the best results.

Fecal specimens should be collected in clean, wide-mouth containers with tight-fitting lids. The specimen should not be contaminated with water or urine, which may contain elements that can be mistaken for fecal parasites. Stool specimens should be placed in leak-proof bags when being transported to the laboratory for analysis. If postal delivery services are used, any diagnostic specimen must be packed according to national or international regulations (e.g., labeling with UN code 3373, the three-container approach) for packaging and shipping of biological specimens. Specimens need to be labeled with the proper patient identifiers, including patient's name and identification number along with the time and date of specimen collection. The specimen must also be accompanied by a request form indicating which laboratory procedures should be performed. Any additional relevant information should be included with the specimen submission.

It is recommended that multiple stool samples be examined prior to ruling out a parasitic infection. Historically, three specimens collected on alternate days within a 10-day period should be examined; however, some may argue that one or two stool exams are adequate. Many organisms, particularly intestinal protozoa, do not appear in the stool in consistent numbers; concentrations of trophozoites and cysts may vary on a daily basis. Physicians should be aware that the probability of detecting clinically relevant parasites in a single specimen may be as low as 50 to 60% but is >95% if three samples are examined by O&P (3, 51, 52).

All fresh specimens should be handled carefully, since each specimen represents a potential source of infectious material. Standard safety precautions, including the use of personal protective equipment, the use of a biological safety cabinet when working with infectious materials, and proper handling of chemicals, should be followed (7, 53–56). Material safety data sheets (MSDS) should be reviewed for all reagents used in the laboratory. It is also mandatory that appropriate policies to prevent eating, drinking, or smoking within the laboratory are in place.

Formaldehyde vapor monitoring must be performed to ensure that the exposure does not pose a risk to laboratory personnel. Initial monitoring should be performed and repeated any time that there is a change in the use of formalin, which may result in an increase in exposure.

Quality control should be performed on a regular basis and documented. The frequency of quality control assessments will depend on the test and the expertise of personnel. Microscope calibration should be performed on all instruments used to examine specimens by the O&P.

Fresh stool specimens are mandatory for the recovery of motile protozoan trophozoites (amoebae, flagellates, ciliates). The trophozoite stage is normally found in cases of diarrhea. Once trophozoites have passed out of the body, they do not encyst but disintegrate if not examined promptly or put into preservative. Most helminth eggs and larvae, Cyclospora/Cystoisospora/Cryptosporidium oocysts, and microsporidial spores survive for extended periods of time. In general, liquid stools should be examined within 30 min of passage (not 30 min after arriving in the laboratory). If this is not possible, the specimen should be placed in a preservative and then transported to the laboratory. Once in preservative, motility will be lost. Semiformed or soft stools should be examined within 1 h of passage and usually contain both cysts and trophozoites. Formed stools should be examined within 24 h after passage and contain mainly cysts. If these specimens cannot be examined within the suggested time frames, again, the specimen should be placed in preservatives.

If there are delays from the time of passage until examination in the laboratory, the use of fecal preservatives should be considered. To preserve protozoan morphology and to prevent continued development of various helminth eggs and larvae, the stool can be placed in preservative immediately after passage (by the patient or hospital staff). Once placed in the preservative, adequate mixing of the specimen is mandatory. To ensure the proper ratio of preservative to stool, commercial vials are marked with a “fill-to” line on the collection container.

There are many preservative options provided by commercial vendors. When selecting an appropriate preservative, make sure that it is compatible with all stains and test kits used in your parasitology laboratory. Formalin, sodium acetate-acetic acid-formalin (SAF), mercuric chloride polyvinyl alcohol (PVA), modified (nonmercury) PVA, and nonformalin, nonmercury, non-PVA preservatives are commercially available (5, 57–59) (Table 4). Disposal regulations for compounds containing formalin and mercury are becoming stricter, and disposal is a factor.

The most commonly performed test in the parasitology laboratory is the complete O&P. It consists of a direct wet mount, concentration, and permanent-stain smear (2–5).

The Cyclospora/Cystoisospora/Cryptosporidium parasites are apicomplexan protozoa, and they are intracellular, oocyst-forming parasites. The three genera recognized as causing human diarrhea and thought to be spread via contaminated food and water are Cyclospora, Cystoisospora, and Cryptosporidium. Cyclospora infection has been identified as common in tropical and subtropical regions, usually with a wet season peak. It is considered endemic in Haiti, Peru, Guatemala, Venezuela, Southeast Asia, Nepal, and India (60–62). There have been a series of large outbreaks of foodborne infection in the United States and Canada traced to imported berries and vegetables, including raspberries, mesclun, and basil. Symptoms include an acute watery diarrhea that may extend for several months if untreated (60–62). In AIDS patients, Cyclospora infection can also cause biliary tract involvement manifested by acalculous cholecystitis and cholangitis (62). Cystoisospora has a similar distribution: Latin America, the Middle East, Southeast Asia, and tropical areas in Africa. As with the other Cyclospora/Cystoisospora/Cryptosporidium infections, illness is more pronounced in the very young, travelers, and AIDS patients. In contrast to infection with Cyclospora and Cryptosporidium, Cystoisospora infection in AIDS patients may include systemic spread to lymph nodes, liver, and spleen. Despite appropriate treatment, infections may become chronic in these patients (63).

Why is it necessary to resort to staining or methods other than direct microscopy to identify these organisms? The spherical oocysts of Cryptosporidium are only 4 to 6 μm in diameter. At this size, they are too small to reliably identify in wet preparations by light microscopy. The unsporulated 8- to 10-μm oocysts of Cyclospora are perfectly round in outline and contain greenish inclusions, or morulas. However, despite these properties being quite identifiable, their comparatively small size means that they can be confused with smaller amoebae or pus cells. Although of larger size at 10 to 20 by 20 to 30 μm, the oocysts of Cystoisospora are unsporulated or contain a contracted sporont when passed. These oocysts are hyaline in appearance and can be poorly visualized by direct light microscopy. For all three species, the use of alternative methods other than direct microscopy is required to enhance detection.

Although these parasites are often considered together, there are features of Cryptosporidium that set it apart from other species. It is considered either as basal to all other Apicomplexa or more closely related to the gregarine parasites, with which Cryptosporidium shares many life cycle features. When describing the genus in 1907, Tyzzer (280) chose the name Cryptosporidium to emphasize the difference from other parasites, notably the absence of a sporocyst and the presence of naked sporozoa within the oocyst. These oocysts differ in having no need for sporocyst maturation, they are immediately infectious, and importantly, they are capable of autoinfection (64, 276). The immense significance of Tyzzer's discovery was not appreciated for many decades.

Further impetus for study of Cryptosporidium came from the veterinary world with the description of Cryptosporidium meleagridis in turkey poults in Scotland in 1955 (65). 1978 saw the recognition of Cryptosporidium associated with scouring in neonatal calves in Iowa (66). As the importance of cryptosporidial illness in calves was becoming recognized, Henriksen and Pohlenz in Denmark made the discovery that oocysts stain acid fast in a Ziehl-Neelsen (ZN) stain; this was a largely empirical finding based on testing a variety of stains from a neighboring bacteriology laboratory (67). The first report of human illness came in 1976 with the identification of Cryptosporidium in rectal biopsy specimens of a 3-year-old child (68). A subsequent review of more cases in humans suggested that infection seemed limited to immunosuppressed patients (69). In 1983, first reports describing Cryptosporidium in intestinal biopsy specimens of AIDS patients were seemingly in agreement with this theory. Initial efforts at staining oocysts from fecal specimens included the use of Giemsa and Kinyoun stains (69). Later that year, at a time when there were only an estimated 1,000 AIDS cases worldwide, a comparison study of specimens from eight patients using 15 different staining options confirmed that modified acid-fast staining performed best (70). Prior to the introduction of antiretroviral therapy, the diagnosis of Cryptosporidium was recognized as one of the AIDS-defining illnesses. The often-unrelenting symptoms included chronic diarrhea or severe cholera-like illness and could also have involved biliary tract colonization, cholecystitis, and cholangitis (71). The prediction that infection would be confined to the immunocompromised proved to be erroneous. Cryptosporidium was soon identified in immunocompetent patients as a cause of self-limiting, watery diarrhea of several weeks' duration; infection was also more common among young children than among adults (72). In subsequent years, Cryptosporidium has also been identified as a significant cause of illness in patients with malignancy and other types of immunodeficiency (73).

In the intervening years, many Cryptosporidium species and possible subspecies have been identified, with many exhibiting a very limited host range. The two species commonly detected in humans are C. parvum and C. hominis. The latter species does not typically infect animals, but C. parvum is commonly infects cattle. This species is recognized as an environmental problem in water catchments in rural communities. In total, there are approximately 20 species or genotypes of Cryptosporidium that have been identified as causing illness in humans, but most of these generally have a more restricted host range (74). The waterborne outbreak in Milwaukee, WI, that affected 400,000 people has been attributed to human feces (C. hominis from the sewage treatment plant) entering the drinking water system after a failure of the water treatment plant filtration system (30, 75).

With the advent of PCR assays, the dimension of illness caused by Cryptosporidium has been dramatically reestimated (76). In an analysis of the global burden of Cryptosporidium, estimates quoted show that 15 to 25% of diarrhea in childhood is attributable to this parasite (77). Furthermore, those most at risk are the immunocompromised, the malnourished, and infants of low birth weight. Review articles have stressed that any form of infection, even asymptomatic or postsymptomatic, can result in significantly reduced growth rates by the age of 6 to 9 years (78, 79). The importance of this infection has been confirmed in the Global Enteric Multicenter Study (GEMS) of the etiology of diarrhea in children (to 59 months) in sub-Saharan Africa and south Asia. Based on detection by immunoassay, results showed that Cryptosporidium infection was the second-most-common infection in infants and was associated with the highest risk of death in toddlers (80). PCR testing in Uganda has also shown concomitant respiratory tract involvement among children with diagnosed intestinal infection (80).

These recent figures illustrate that infection rates in developing countries have been significantly underestimated. In poorly resourced regions, where access to molecular techniques may not be available, it is important that attention is focused on improving existing technology, especially microscopy and staining techniques. The introduction of LED-sourced fluorescence has reduced the ongoing cost and maintenance of fluorescence microscopy, which should enhance the test options for the diagnosis of Cyclospora/Cystoisospora/Cryptosporidium and microsporidian parasites.

In considering alternative approaches for concentrating and staining oocysts of Cryptosporidium, it is essential to have a gold standard method for comparison. It is generally accepted that a fluorescent antibody test meets that requirement. First developed by Sterling and Arrowood (81), the assay was based on a monoclonal antibody to oocyst cell wall antigen. Similar versions of this assay are now marketed commercially by a number of manufacturers (71). Apart from cost, a limitation of this test is the requirement that the fecal smear must be processed or cleared in some way to ensure that fluorescence can easily be detected. This test will be described in more detail below.

The presence of acid-fast lipids in the cell wall and the property of autofluorescence are all common to Cryptosporidium, Cyclospora, and Cystoisospora (88, 90) (Fig. 3). This shared property means that the modified acid-fast stain still has relevance for the combined screening for all three parasites, especially in immunocompromised patients and for those with a history of overseas travel. At this stage, no other single test offers this flexibility. Henriksen and Pohlenz were the first (67) to recognize that lipids in the wall of Cryptosporidium oocysts could be stained with an acid-fast stain. Their method involved flooding a smear with strong carbol fuchsin, decolorizing with 5% sulfuric acid in alcohol, and counterstaining with methylene blue. This stain was referred to as a modified Ziehl-Neelsen (ZN) stain, as there was no heating step. An alternative approach adopted by Garcia et al. (70) was also referred to as a modified acid-fast stain. This method was more similar to that using the ZN stain in that heating of the carbol-fuchsin step was required. The atypical step was decolorizing with 5% H₂SO₄, without alcohol (acid-alcohol is used for acid-fast staining of mycobacteria). Garcia et al. also suggested the pretreatment of mucoid smears with 10% KOH to break up the mucus and improve adherence of the sample to the slide (70). In 1991, Casemore (91) described another method based on the standard solutions used for a ZN stain. Smears were stained for 20 min with either strong carbol-fuchsin or Kinyoun stain without heating and decolorized with 1% HCl in methanol, followed by counterstaining in malachite green or methylene blue.

There are many minor variations in performance of modified acid-fast stains, but the consensus now is to use 1 to 3% H₂SO₄ as a decolorizer and to avoid the use of alcohol (1% H₂SO₄ has been favored if the stain is for dual-purpose detection of Cyclospora). There is no heating step involved with the Kinyoun stain. It contains high concentrations of both phenol and basic fuchsin, and this is thought to give better penetration of the dye. It is postulated that phenol may change surface tension, allowing dye to enter hydrophobic surfaces. Once the dye is bound, it cannot be removed by acid treatment. Another variation in acid-fast staining is the addition of dimethyl sulfoxide (DMSO) to the phenol-basic fuchsin and the incorporation of acetic acid with malachite green to act as a combined decolorizer-counterstain. It was reported that by this method, staining of internal details of oocysts was more detailed than seen with conventional ZN stains (92). The action of DMSO is thought to allow better penetration of the fuchsin stain into the lipid layer.

Although Cryptosporidium, Cyclospora, and Cystoisospora are all identified as staining acid fast in modified acid-fast stains, shortcomings have been identified. One of the principal criticisms is that ghost cells or poorly stained oocysts occur, suggesting varied levels of uptake of the dye and ease of destaining (83). In particular, Cyclospora does not stain uniformly; there are usually many unstained ghost cells present (62). It has been suggested that to counteract this variability, the strength of decolorizer in the Kinyoun stain should be reduced to 1% H₂SO₄ (90). In contrast, the staining of Cystoisospora in the modified acid-fast stain is regarded as a suitable diagnostic test (88). Oocysts stain a mottled-pink color, but sporonts or sporocysts stain bright crimson (63).

To perform the modified acid-fast stain in our laboratory (NR), smears are prepared directly from fecal suspensions in SAF with no prior concentration step. Smears are briefly heat fixed on a hot plate at 60°C for 5 min and then fixed in methanol for 5 min. The prior, brief heat fixing is recommended to increase adherence and may improve staining of Cyclospora. The staining method used is that of Casemore (91), as described above. This method provides good contrast in staining and little obvious occurrence of ghost cells in control smears. The stained smear is first coated with a light layer of oil and examined at a ×100 magnification using strong lighting. Viewing at low magnification allows screening of a large area of the slide. Any pink objects are easily detected and are then inspected at ×1,000 magnification to identify the typical morphology of Cryptosporidium or other oocysts. Oocysts of Cryptosporidium are commonly found in thicker areas of the slide. Staining artifacts may be encountered; these generally take the form of variously sized, undifferentiated particles that are possibly coagulated protein. Bacterial and fungal spores may also stain acid fast and are seen quite commonly but can readily be distinguished by their morphology and intensity of staining. There is enormous variation in the choices of recipe used for modified acid-fast stains, ranging from ZN-strength carbol-fuchsin to the stronger Kinyoun reagent, applied with or without heating. For decolorizer, the options range from 1 to 10% H₂SO₄ with or without alcohol to the use of 1 to 3% HCl in alcohol. The choice of counterstain is usually either methylene blue or malachite green. The enormous variability underscores the necessity for adequate control material and for some estimation of the quantitative capability, perhaps in identifying standardized controls. In general, despite the variety of approaches used, the modified acid-fast stains can be used for the screening and identification of these species.

There has been a long history of study of Microsporidia, with early focus being on insect Infections, but they have been recognized subsequently as parasites of all animal groups (109, 110). Microsporidia are eukaryotic species formerly considered protozoa but now thought to be more closely related to fungi. They exist by harnessing host cell metabolism to support a proliferative stage followed by spore formation. The identifying feature of microsporidian spores is that they contain a coiled tubule which is capable of evagination and penetration of the host cell membrane to transfer infective sporoplasm into the cell. Other features that are shared with fungi include the presence of chitin and trehalose, similarities in cell cycles, and certain gene organizations. Microsporidia are now considered to be highly derived fungi that underwent genetic and functional losses, thus resulting in one of the smallest eukaryotic genomes known. However, at this point, clinical and diagnostic issues and responsibilities may remain with the parasitologists.

Typical spore sizes range from 1.5 to 5 μm wide and 2 to 7 μm long; however, the organisms found in humans tend to be quite small, ranging from 1.5 to 2 μm long. Until recently, awareness and understanding of human infections have been limited; with increased understanding of AIDS and more recently other immunosuppressive illnesses, more attention has been focused on these organisms. Limited availability of electron microscopy (EM) capability has also played a role in the inability to recognize and diagnose these infections. However, with the introduction of newer diagnostic methods, the ability to identify these parasites has definitely improved, particularly in solid-organ and bone marrow transplant recipients.

Recent studies have highlighted the fact that Microsporidia are more widespread than first thought (281). Asymptomatic human and animal carriage illustrate that the environmental spread of spores is likely, especially through water (282). Those most at risk for severe infection include many categories of immunocompromised patients, not just HIV patients. Microsporidial infections have been detected in patients who have undergone bone marrow or solid-organ transplants and in patients with severe chronic illness requiring immunosuppressive therapies (119–123, 139, 140). The use of combination staining methods will reduce the laboratory requirement for single-focus assays. In this respect, combined stains that target Microsporidia and Cryptosporidium are of great practicality, as both constitute significant infections of immunocompromised and immunocompetent patients.

For most parasitic diseases, the antigen(s) is generally the assay component which has the most influence on test sensitivity and specificity. Parasites generally have more than one life cycle stage that may have both mutually shared antigens and stage-specific antigens. The matrix to which antigens are bound for use in a specific procedure also influences which antigen subset will be available for antibody binding.

The ova and parasite examination (O&P) is traditionally for detection of fecal parasites. Since the O&P is a labor-intensive, multistep test that requires a skilled microscopist, other test options have been developed. Immunoassays are available for a limited number of pathogenic protozoa, including Giardia lamblia (G. duodenalis, G. intestinalis), Cryptosporidium spp., the Entamoeba histolytica/Entamoeba dispar group, and Entamoeba histolytica. Commercially available antigen test formats include direct fluorescent antibody (DFA), enzyme immunoassay (EIA), and immunochromatographic lateral flow assays (LFAs) and are more sensitive and specific than the O&P (7, 95, 100, 101, 141–150). Immunoassays for antigen detection, however, are usually limited to one or two organisms only. When choosing which method is best for your laboratory, parameters such as test performance characteristics, skill level of technologists, availability of equipment, volume of requests, specimen collection requirements, and kit cost should be considered.

Fresh or preserved fecal samples are acceptable specimens for immunoassay testing. It is best to refer to the recommended collection procedures for each specific kit. All current EIA and LFA kits that include testing for the Entamoeba histolytica/E. dispar group and Entamoeba histolytica require fresh, unpreserved stool specimens only (149). Fresh, unpreserved specimens can be stored at 2 to 8°C and tested within 48 h or frozen. Most test kits for Giardia lamblia (G. duodenalis, G. intestinalis) and Cryptosporidium spp. accept stools preserved in a number of fecal fixatives. Ten percent formalin, merthiolate-iodine formalin (MIF), sodium acetate-acetic acid-formalin, non-polyvinyl alcohol (PVA) preservatives, and the single-vial universal fixatives are acceptable and should be tested within 2 months of collection. Stool specimens collected in Cary-Blair transport medium (or a similar medium) should be refrigerated or frozen and tested within 1 week. Specimens collected in mercury or PVA are not acceptable.

Immunodetection of antigens in stool specimens using the monoclonal-antibody-based DFA, EIA, and LFA formats are available through various manufacturers. Several kits combine tests to detect Cryptosporidium spp., Giardia lamblia (G. duodenalis, G. intestinalis), and the E. histolytica/E. dispar group (141–149). Only one EIA kit and one rapid test are specific for Entamoeba histolytica; additional kits are due to be released in the near future. All kits have high specificities (90 to 100%) and sensitivities (63 to 100%). DFA kits have sensitivities and specificities of >95%. Enhanced sensitivity for the DFA is attained by using concentrated fecal sediment. EIAs are also very sensitive; however, false-positive results have been known to occur (151).

Special larval-nematode detection methods are used mainly for diagnosing Strongyloides infection when the direct fecal smear or the FEA concentration technique, both of which are relatively insensitive tests, yields negative results. Because Strongyloides are highly intermittent with their shedding patterns, which makes the routine ova and parasite examination unreliable, the consequences of a missed infection might have serious implications for patient management in certain population groups. It is therefore necessary to exclude this organism using a more sensitive testing process (152).

Various migration techniques can be used for the detection of Strongyloides (and other nematode larvae, such as hookworm and Trichostrongylus species) in human stools and are generally categorized into water emergence and agar plate culture methods, with the former becoming increasingly less popular. It is important to remember that all fecal specimens for any larval-detection method must be freshly passed and not refrigerated.

Although it is possible to correlate the egg production of Ascaris lumbricoides, Trichuris trichiura, the hookworms, and Schistosoma mansoni with worm burden, the quantitation of helminth eggs is rarely required in routine diagnostic laboratories. The number of eggs detected in a stool specimen does not significantly influence anthelmintic drug therapy, nor is it needed for posttreatment follow-up.

Most laboratories do not offer worm burden counts; however, when teaching students parasitology, a semiquantitative density chart (Table 5) can aid as a scoring guide of parasite recognition. This semiquantitation method is also used for proficiency testing reporting of helminths.

Helminth egg counting is an inexact science, with many factors contributing to variations and errors in eggs per gram (EPG). These factors include the fecundity of worms and variability in daily egg production, stool consistency influenced by diet and fecal transit times, and the experience and accuracy of the testing technician. It should therefore be taken into consideration when applying the results of helminth egg counts that they do not necessarily correlate with or accurately predict the true worm burden of the host.

The most common egg counting techniques have traditionally been the direct smear (Beaver) (5–7, 47), the Stoll method (5–7, 47), and the Kato-Katz method (5–7, 47, 159–168); however, the FLOTAC (159–164) and McMaster (161, 166) methods are becoming increasingly more popular. Further studies are needed to validate the mini-FLOTAC with other quantitative techniques (McMaster and Kato-Katz) and in different settings where other soil-transmitted helminths are also endemic (169).

Schistosome eggs are normally detected in stool or urine using either direct microscopy or concentration methods. Determining their viability is also a significant factor for the clinician to consider with patient management, as viable eggs represent an active infection. If they are nonviable, it may indicate a “burnt-out” infection or that treatment has been successful even though eggs can still be passed in stool or urine for months afterwards. If stool or urine has been preserved with a fixative or been refrigerated, the ability to determine the viability of eggs seen is nullified; therefore, the hatching of the eggs to release the miracidia in fresh, nonfixed stool is a procedure which not only confirms egg viability but may also increase the sensitivity of detection of a schistosomal infection (5). Certainly if schistosomiasis is suspected as a potential diagnosis, fresh, nonfixed specimens are recommended.

The standard method described for hatching eggs, particularly from those species whose eggs are found in stools, entails the use of an elbowed sidearm flask (few laboratories have these available), but similar vessels, such as Erlenmeyer flasks, are quite satisfactory (5–7).

Fresh, nonrefrigerated feces are homogenized in saline and coarse filtered through gauze. This process is twice repeated with the fecal sediment obtained. The saline suspension is then decanted, replaced with chlorine-free water, and added to a 500-ml sidearm flask so that the water rises 20 to 30 mm vertically in the side arm (the eggs will not hatch in saline). The flask is tightly covered in foil, leaving only the sidearm exposed. Place a bright light near the exposed sidearm and then stand the flask at room temperature, ensuring that the light does not heat the sidearm fluid. With a hand lens, observe for minute white, moving organisms in the sidearm and pipette a drop or two off for microscopic confirmation. An enhancement of this method applied to hatching the miracidia of S. mansoni was shown to be more sensitive than the Kato-Katz method (170). S. mansoni organisms are phototrophic and actively seek a light source; however, S. japonicum is reported to be not attracted to light (171), so this property should be taken into account when attempting to hatch eggs from this species.

The diagnosis of Schistosoma haematobium infection is confirmed by detecting the eggs in urine sediment or using a filtration technique, so there is seldom any point in using egg-hatching techniques for this species. Provided the specimen has not been preserved or refrigerated, viability can frequently be determined by observing either an active miracidium within the egg or flame cell movement (wafting) within the larva. There are four flame cells (sometimes termed solenocytes), one near each corner of the embryo (172). Trypan blue staining is another reliable way of determining viability (173). If viability is still undetermined, urine (preferably that collected between 10:00 a.m. and 2:00 p.m., when there is a daily peak excretion of eggs) is diluted approximately 1 in 10 with chlorine-free tap water in a large flask, in which miracidial hatching will occur in 10 to 15 min. Against a black background, they can be seen with the naked eye as actively motile minute white bodies. S. haematobium miracidia are relatively nonresponsive to light stimulation and will not rise to the source of an illuminated flask (174).

Note that since adult schistosomes may occur in unexpected sites (S. mansoni from urine), both urine and stool specimens must be collected without preservatives and should not be refrigerated before being processed. Regardless of the species suspected, both urine and stool should be examined for every patient with potential schistosomiasis.

Sigmoidoscopy is a clinical procedure that can be used to obtain specimens for the diagnosis of Entamoeba histolytica or schistosome infection, although the advent of sensitive immunoassays and DNA-based technologies for detecting E. histolytica (179) has probably reduced the need for this procedure. Sigmoidoscopy may still be indicated when permanent-stain films or immunoassays of a series of stool specimens are negative or equivocal for E. histolytica yet clinical suspicion remains. It is essential that there be a close liaison between the clinician and the laboratory scientist to ensure that specimens are correctly collected and processed.

Sigmoidoscopy may reveal the characteristic ulcers of intestinal amebiasis, especially in more severe disease, and allow the clinician to target lesions on the mucosa for specimen collection. Both direct wet preparations and slides for stained films should be made from material from the mucosal surface, which must be scraped or aspirated rather than wiped with absorbent tipped swabs. Specimens should also be taken from multiple sites, with each processed individually. If biopsy specimens for histological examination are taken, they should be preserved in 10% buffered formalin.

If a slide made for direct wet microscopy is to be of any value, a coverslip needs to be placed over the slide (suffused with a drop of saline) at the bedside, and the edges of the slide should be sealed with an agent such as nail polish to prevent drying before the slide is transported stat to the laboratory. At the laboratory, examine for trophozoite motility, initially under a 10× objective magnification. If trophic forms are detected, confirm any red blood cell (RBC) ingestion under a 40× objective magnification. An immediate diagnosis of E. histolytica can be made only if hematophagous trophozoites are seen in the wet preparation. Active directional motility is sometimes described as a diagnostic feature of E. histolytica, but this can be a subjective observation and is only suggestive at best. Nuclear detail cannot be seen in unstained preparations. The use of these direct mounts is currently uncommon and has been supplanted by the use of preserved material.

Fixatives such as those containing PVA or the newer single vials (universal fixatives) should be supplied at the bedside after sigmoidoscopy for making permanent slides for staining. Mucoid sigmoid material needs to be dabbed or scraped onto a glass slide and fixed immediately or mixed thoroughly with a drop or two of fixative containing PVA and then air dried. The making of permanent-stain slides should take precedence over the direct wet preparation, especially if the specimen amount is limited.

A permanent-stain film, such as one stained with trichrome, will confirm ingested RBCs for a definitive diagnosis of E. histolytica; however, the absence of such does not necessarily invalidate a diagnosis, and other features, such as nuclear details of trophozoites, need to be considered (5). The cyst stage of Entamoeba sp. is unlikely to be found from sigmoidoscopy specimens.

It should also be noted that although the WHO has recommended that intestinal E. histolytica infection be diagnosed with a specific test, such as E. histolytica-specific immunoassays (180), these methods have been validated only for fecal specimens and, as such, should not be used without additional validation for mucosal specimens collected after a sigmoidoscopy. DNA-based diagnostics have become increasingly more affordable and are about 100 times more sensitive than ELISA antigen tests (179).

The examination of sigmoid colon biopsy specimens is also considered a sensitive test for the rapid diagnosis of S. mansoni and S. japonicum infections, particularly if fecal testing is negative yet clinical suspicion remains high (181). A number of rectal biopsy specimens taken at sigmoidoscopy or proctoscopy from inflamed or granulomatous lesions or even random sites are gently crushed flat between two glass slides to near transparency and then examined microscopically (initially 40× objective). Some methods suggest soaking the tissue in isotonic saline for 30 to 60 min before examining the tissue (and, if need be, by hydrolysis of fresh tissue in 4% KOH for 18 h at 37°C or at 56°C for fixed tissue, although this procedure will invalidate observation of viable organisms). Schistosome eggs will appear as refractile bodies, often in clusters. Determining egg size and shape and the size and position of the spine definitively identifies the species, and observing flame cell movement within the miracidia (100× objective) will determine viability. Calcified eggs appear as black bodies. Occasionally, S. haematobium eggs are found in the rectal region and need to be differentiated from superficially similar Schistosoma intercalatum eggs. The latter are usually acid fast, so Ziehl-Neelsen staining can assist identification; other schistosome eggs, such as those of S. mansoni and Schistosoma mekongi, also exhibit an acid-fast shell outline (173). S. haematobium can also be diagnosed by demonstrating its characteristic eggs from biopsy specimens taken at cystoscopy using the same technique. Some biopsy specimens should also be preserved in 10% buffered formalin for histological sectioning and staining.

At an endoscopy procedure, villus biopsies of the duodenal-jejunal region are usually taken as part of the investigation of upper intestinal disorders, including parasitic infections (5–7). Although biopsy specimens may be variously taken for histological, immunological, or biochemical studies, they are also valuable in detecting Giardia trophozoites in situ (and, more rarely, Strongyloides, Clonorchis [Opisthorchis], and microsporidial infection), especially if repeated stool examinations are negative but clinical suspicion remains. Aspirated duodenal fluid may also be collected for parasite examination.

Specimens collected by a physician at endoscopy for Giardia require immediate transportation at room temperature to the laboratory and need to be examined without delay. Biopsy specimens must be in isotonic saline to prevent desiccation.

Duodenal aspiration fluid, also collected at endoscopy, may sometimes be received for (primarily) Giardia examination (5–7). As with biopsy specimens, the fluid needs to be collected without any preservatives and sent immediately to the laboratory for stat examination. Depending upon the volume received, the specimen may need to be centrifuged (10 min at 500 × g). Examine the deposit for trophic forms, which, as for biopsy specimens, may exhibit only a barely noticeable fluttering movement. If there is mucus in the fluid, remove it with a Pasteur pipette for microscopic examination, as trophozoites tend to adhere to slough. There will be no movement if the specimen has become too cold, so identification will then need to be made by shape, binucleated appearance, and sucking disc on the organism's ventral side. Slides fixed in either Schaudinn's or PVA fixative should also be made for trichrome or iron-hematoxylin staining. If PVA fixative is to be used, mix 2 to 3 drops of the fixative onto a slide containing a drop or two of the aspiration deposit and mix well. Spread the specimen with an applicator stick to ensure that the preparation is thin and thoroughly dry before staining.

Strongyloides larvae in a wet preparation move and may demonstrate thrashing motility under a 10× objective magnification. Because the bile-stained eggs of Clonorchis (Opisthorchis), which have a size of around 30 μm, can easily be missed under ×10 magnification, scanning with a 40× objective is recommended. Examination for Microsporidia will require some specimens to be fixed in 10% buffered formalin and then centrifuged before slides are made for staining.

The string test, also known as the gelatin capsule method and marketed commercially as the Entero-Test, was devised in 1970 as a procedure to diagnose upper-small-bowel pathogens, especially Strongyloides (182). Although patient cooperation is required, it is a novel, simple, and rapid procedure used to obtain secretions from the duodenum-jejunum region and was also found to detect other parasites, including Giardia, Cryptosporidium, and Cystoisospora, without the need for endoscopy. More rarely, other parasites may also be found using this method, including Clonorchis (Opisthorchis), Fasciola, Trichostrongylus, and hookworm eggs. This testing procedure has also been applied to nonparasitological investigations, such as measuring mucosal inflammation in eosinophilic esophagitis and as a device to sample the esophageal microbiome. The use of the string test for diagnosing giardiasis has, however, probably declined over the years, being surpassed by sensitive fecal antigen detection tests that have since become available.

The string test is a gelatin capsule containing a coiled 90-cm line for children or a 140-cm line for adults and a 1-gram weight (5–7). The line consists of 20 cm of silicone rubber-covered thread and 70 cm of soft nylon yarn, one end of which is taped to the patient's cheek or ear. The capsule is swallowed and the line plays out, the gelatin dissolves in the stomach, and the weighted line passes through to the upper small intestine, where the weight becomes disengaged with the line and eventually is expelled in the stool. The line is left in position for around 4 h and then retrieved. Four to five drops of bile-stained mucus can be “milked” with thumb and forefinger off the terminal end of the string and expressed into a small tube, or the whole string can be placed into a petri dish, sealed with tape, and sent immediately to a laboratory. Disposable gloves must be worn when this procedure is performed.

A wet preparation is made from a drop of the fluid, and the complete coverslip area is examined for parasites. Giardia trophozoites adhere to any mucus portions and may exhibit fluttering motility (40× to 100× objectives), but a permanent-stain film should also be prepared. Mix a drop of the fluid on a slide and fix it in Schaudinn's fixative, or prepare a thin slide of the mucus mixed with a drop of PVA fixative and thoroughly dry the slide before staining. Other fecal fixatives appropriate for permanent staining are also acceptable. Strongyloides larvae are actively motile in the wet preparation (10× objective), but care must be taken to distinguish embryonating Strongyloides eggs from those of hookworm. Other helminth eggs are visible in the wet preparation (10× to 40× objectives). Although Cryptosporidium oocysts may appear as small refractile bodies and Cystoisospora oocysts are characteristically large and contain one or two sporonts, they need to be confirmed with a modified Ziehl-Neelsen stain or by demonstration of epifluorescence. Record also the color of the string, as a yellow-bile stain indicates that the line reached the duodenal-jejunal region.

Increasingly, culture of parasites has been deemphasized because of cutting-edge research in molecular biology that facilitates pathogen identification and diagnosis even when small numbers of organisms are present in a sample. Culture options are limited to large, more-complex laboratories and are not routinely available. It should be noted, however, that culturing parasites is essential for diverse areas of research where large numbers of parasites free of bacteria, fungi, and host materials are needed. An important advantage of in vitro culture, especially for axenic cultures, is its provision of a continuous supply of pure organisms without any interfering bacterial, fungal, or host tissue contamination. If practical, culture of the parasite should be attempted, even when identification of the organism has already been made, so that a bank of isolates can be established for further antigenic, molecular, and biochemical research, as well as epidemiologic investigations. In vitro culture is also invaluable for screening potential therapeutic agents that can interfere with the development of the parasite so that the infection can be terminated. Further, in vitro culture can be useful in elucidating isolate or strain differences that may provide helpful insights into the treatment of patients, as well as in developing and assessing the efficacies of vaccines. Additionally, cultured parasites are essential for developing animal models wherein experimental animals can be infected with the parasites to understand the pathological process and to develop effective candidate therapeutics. Although in vitro cultivation of the various parasites discussed here constitutes a continuing challenge, it nevertheless is a fruitful area of research (183).

Amoebae grown in association with unknown bacterial associates are called xenic cultures. If the amoebae are grown in association with a single known bacterium, the culture is called monoxenic; if amoebae are grown with several identified bacterial flora, then the culture is called polyxenic. If amoebae are grown as pure culture without any bacterial or other associated organisms, then the culture is called axenic.

The history of cultivation of the intestinal parasites, especially Entamoeba histolytica, is long and dates back to 1925, when Boeck and Drbohlav (184) isolated and cultured E. histolytica in a diphasic egg slant medium that they had developed for culturing intestinal flagellates. Dobell and Laidlaw (185) modified the egg slant medium by replacing Locke's solution, which contains dextrose, with Ringer's solution and by replacing the liquid overlay with egg albumin or serum diluted with Ringer's solution and adding particulate rice starch at the interphase of the liquid overlay and solid slant. In 1946, Balamuth (186) reported on an improved egg yolk monophasic medium for the cultivation of intestinal amoebae. Robinson (187) developed a complex biphasic medium containing agar slants, Bacto peptone, erythromycin, a defined “R” medium with Escherichia coli, and phthalate solution. In 1982, Diamond (188) developed a new monophasic medium (TYSGM-9) for the cultivation of E. histolytica and several lumen-dwelling protists. This medium consisted of buffered salt solution, casein digest peptone, yeast extract, gastric mucin, heat-inactivated bovine serum, and rice starch in screw-cap tubes. The amoebae could easily be examined directly within the culture tube, and the density and movement of the amoebae could be monitored without removing an aliquot. Three media, modified Boeck and Drbohlav's egg slant medium, Robinson's medium, and Diamond's TYSGM-9, are used most often for the isolation and subsequent cultivation of E. histolytica (189). Culture of E. histolytica serves only as a supplemental procedure and never replaces the primary diagnosis by routine fecal examinations, PCR, or an E. histolytica-specific antigen assay. Even when the culture system is within quality control guidelines, a negative culture is still not definitive in ruling out the presence of E. histolytica.

Previously called nonpathogenic E. histolytica, E. dispar can be grown in several culture media (modified Boeck and Drbohlav's egg slant, Robinson's, and Diamond's TYSGM-9 media) that are normally used for cultivating E. histolytica. However, E. dispar resists growing in the axenic medium designed for E. histolytica. Clark reported growing E. dispar initially in the axenic medium supplemented with irradiated bacteria and later on with Escherichia coli fixed with either glutaraldehyde or formalin (190). After a few years of growing with the killed bacteria, the amoebae finally adapted to growth in the axenic medium without any bacterial supplementation (190). Kobayashi et al. (191) designed a medium (YIGADHA-S) containing yeast-iron-gluconic acid-dihydroxyacetone-serum and a vitamin mixture to cultivate E. dispar axenically. This medium is based on Diamond's casein-free yeast extract-iron-serum (YI-S) medium (192, 193). According to those authors, the main differences from YI-S medium are replacement of glucose by gluconic acid, addition of dihydroxyacetone and d-galacturonic acid monohydrate, and sterilization by filtration. This medium promoted the axenic growth of 5 strains of E. dispar (2 strains of nonhuman primate isolates and 3 strains of human isolates).

Blastocystis is an obligate anaerobic parasite with worldwide distribution. It is a common inhabitant of the gastrointestinal tracts of humans as well as a wide variety of vertebrates, including pigs, cattle, rats, mice, chickens, and reptiles. Although identified as a possible agent of gastrointestinal illness, its pathogenic status has not been clearly established (194). After being tossed around in various taxonomic groups, it has finally been classified as a stramenopile (along with golden brown algae, brown algae, chrysophytes, diatoms, water molds/oomycetes, slime nets, bicosoecids) based on the phylogenetic analysis of the 5 rRNAs (195). A commonly used culture medium for Blastocystis spp. is modified whole-egg slant medium with a Locke solution overlay used for the isolation of Entamoeba spp. (196); it contains 30% horse serum or 20% human serum (197). Blastocystis spp. can also be grown in Diamond's TP-S-1 or Balamuth's medium, although growth is not as good as that in the egg slant medium. If the tubes containing fecal material are positive for Blastocystis spp. after 48 h of incubation, then confirm the identification using a permanent-stain smear and/or a digital image. If negative, an additional 48 h of incubation can be performed after subculture.

Karapetyan (198) was the first to culture Giardia lamblia (G. duodenalis, G. intestinalis) in 1960 in a mixed-culture medium containing the yeast Candida guilliermondii and chick fibroblasts. In 1962 (199), he reported monoxenic cultivation of Giardia duodenalis (a rabbit isolate) with Saccharomyces cerevisiae. In 1970, axenic cultivation of Giardia from the rabbit, chinchilla, and cat (200) was achieved by Meyer after migrating the trophozoites in a U-shaped glass tube and separating the protozoa from the yeast. Giardia lamblia from the duodenal aspirate of a woman with an 8-year history of giardiasis was finally isolated and cultured with the accompanying yeast in two different liquid media (HSP-1 and HSP-2) (201). The HSP-1 medium consisted of Phytone Peptone, glucose, l-cysteine hydrochloride, Hanks' balance salt solution, and 15% human serum and antibiotics (penicillin and streptomycin). HSP-2 medium consisted of 100 ml HSP-1 medium plus 7.5 ml NCTC-135 (Gibco) and 2.5 ml glutathione-cysteine reducing solution. Since some of the human sera inhibited the growth of Giardia in the HSP-2 medium, Visvesvara (202) experimented with Diamond's TPS-1 medium used for the cultivation of E. histolytica. He found that G. lamblia (G. duodenalis, G. intestinalis) did not grow in autoclaved TP-S-1 medium but grew well in TPS-1 medium that was filter sterilized. Gordts et al. (203) cultured Giardia in TPS-1 medium by directly inoculating the parasites from the human intestines. Since Panmede (a liver digest and a component of the TPS-1 medium) was no longer available and liver digest from other sources did not support good growth of Giardia, efforts were made to come up with a medium that supported the growth of not only Giardia but other protists, including E. histolytica. Finally, Diamond's TYI-S-33 medium, devised to grow E. histolytica, was used by Keister (204), with some modification to support the growth of Giardia axenically. Because of the new manufacturing methods of some of the components (casein digest peptone), the growth of these parasites was somewhat erratic; a new medium (YI-S) consisting of yeast extract, iron, and serum, which supported the growth of both Giardia and E. histolytica, was designed (193). Bingham and Meyer (205) described a method of treating mature Giardia cysts with hydrochloric acid (pH 2), which allowed the excystation of the parasite and thus facilitated its axenic cultivation. In most cases, acid treatment kills off all bacteria, and the resulting culture is axenic. Therefore, this method is preferable because invasive procedures like collection of duodenal aspirate and biopsy specimen are negated.

Although Dientamoeba fragilis was considered nonpathogenic and thought to lack a cyst stage, recent studies indicate otherwise (206, 207). Dobell (208) was probably the first to establish thriving cultures of D. fragilis in a diphasic medium that consisted of a slant of inspissated horse serum overlaid with egg whites in Ringer's solution, with rice starch added at the confluence of the solid and liquid media. It has been cultured by others in Locke egg (LE) medium, TYSGM medium, Balamuth's egg yolk infusion medium, Robinson's medium, and other media with mixed bacterial flora.

Recently, Barratt et al. (209) reviewed the previous work on the culture media used and compared different culture media for their ability to support the growth of several isolates of D. fragilis, their optimum temperatures for growth, the optimum method for the cryopreservation of the isolates, and important bacterial associates in D. fragilis cultures. Based on their exhaustive research, Barratt et al. have shown that the best culture medium to grow D. fragilis is Loffler's medium, although the LE medium and Robinson's medium support fairly good growth of the organisms. Development of either a monoxenic or an axenic culture of D. fragilis has not yet been developed.

Balantidium coli is a large, ciliated protozoon found in the gastrointestinal tracts of humans, nonhuman primates, pigs, and cattle, although its normal host is pigs. Humans can acquire the infection via the fecal-oral route, and human-to-human transmission may also occur. Humans may remain asymptomatic, as does the pig, or may develop dysentery similar to that caused by Entamoeba histolytica. B. coli is often referred to as an opportunistic pathogen (210, 211). Recently, the nomenclature of the mammal-infecting Balantidium genus has been changed to Neobalantidium based on the internal transcribed spacer (ITS) region of the small-subunit (SSU) ribosomal-DNA (rDNA) tree and morphological differences. Since Balantidium causes no pathology in poikilothermic hosts, but N. coli is pathogenic to warm-blooded animals, the term balantidiasis is retained to describe the infection in humans. Currently, the term Balantidium coli is still in use for human infections; however, the name change may occur in the future.

Although, Barret and Yarbrough (212) used a simple saline-serum medium for the cultivation of Balantidium, it can be isolated and cultivated in a wide variety of media, including LE, Jones', Robinson's, Balamuth's, Dobell's HSre with starch, and TYSGM-9 media, which have been used for the cultivation of E. histolytica (210). It can also be maintained monoxenically with Escherichia coli or other intestinal bacteria without much difficulty (210). It can be grown at a broad temperature range, from 25°C to 40°C (213).

The obligate, intracellular, protozoan parasites of the genus Cryptosporidium infect epithelial cells lining the digestive and respiratory tracts of a wide range of animal hosts. Typical of related parasites, Cryptosporidium features a complex life cycle, including asexual and sexual stages, culminating in the production of small, environmentally hardy oocysts. This monoxenous parasite can multiply to tremendous numbers in the host by “recycling” asexual stages and producing thin-walled oocysts in the gut that mature and release infectious sporozoites in the same host (autoinfection). Most human infections are attributed to two species: C. parvum (zoonotic transmission) and C. hominis (anthroponotic transmission) (214–216, 277), and the majority of in vitro studies have focused on isolates of these species.

Basic in vitro culture methods for Cryptosporidium species have been developed since the first report of success in 1983 (217). Indeed, cryptosporidial growth has been evaluated in a wide range of cell lines, but the majority of published in vitro studies use the following cell lines: human colonic tumor cells (HCT-8, CCl-224), Madin Darby canine kidney cells (MDCK, CCL-34), and human colonic adenocarcinoma cells (Caco-2, HTB-37). In vitro development of C. parvum through asexual and sexual stages is routinely accomplished in cell lines, and continuous culture has recently been reported (218, 219, 283).

Although more than 1,200 species belonging to 143 genera of the Microsporidia have been described, only parasites belonging to 9 genera and 13 species are known to cause human disease. They are Anncaliia, Encephalitozoon, Enterocytozoon, Microsporidium, Nosema, Tubulinosema, Pleistophora, Trachipleistophora, and Vittaforma. However, Enterocytozoon bieneusi, the three species of Encephalitozoon (E. cuniculi, E. hellem, and E. intestinalis), and Anncaliia algerae are the most frequently found to infect the humans. Only short-term culture of E. bieneusi is possible so far. E. cuniculi, E. hellem, E. intestinalis, and Anncaliia algerae have been cultured from urine, sputum, and corneal smears and cryopreserved indefinitely (220). However, none of these Microsporidia have been cultured directly from the stool.

Molecular tools have been used for a variety of purposes in the study of parasitology. Many of these applications involve a further understanding of the biology of parasites and, although important, are not immediately relevant to this manuscript. Similarly, these tools have been used by veterinary parasitologists and environmental microbiologists. References to these studies will be made only when there is pertinence to human infections, such as when zoonotic transmission or food- or waterborne infections are studied. There have been no FDA-approved/cleared molecular diagnostic assays in the recent past. Numerous laboratory-developed assays that target a variety of parasites have been developed. Although numerous papers have disclosed excellent sensitivity and overall performance for many of these assays, there has been limited adoption in clinical microbiology laboratories for a number of reasons.

Methods of extraction and/or more-robust PCR assays needed to be developed to overcome the various inhibitors that may be found in stool (221). Additionally, and to this day, a single assay that affords detection of the full range of the parasites that may be found via an ova and parasite microscopic examination has not been developed. Furthermore, the medical community has been quite satisfied with the sensitivity of the Giardia and/or Cryptosporidium DFA microscopy and EIAs that are commercially available and moderately priced. The added cost of PCR for select pathogens, in conjunction with the need to retain the O&P, has been a deterrent to the incorporation of molecular diagnostics into clinical parasitology.

A variety of molecular-laboratory-developed tests (LDTs) that target parasites have been developed. This section will predominantly review LDTs that target common intestinal parasites, such as Giardia and Entamoeba histolytica, and parasites that infect immunocompromised hosts, such as Cryptosporidium and Microsporidia species. These results can be particularly helpful in outbreak investigations and epidemiological studies. It is also important to remember that some state public health laboratories require that stools positive for certain organisms (Cryptosporidium) be submitted for confirmatory testing and typing. The LDTs have been developed as both monoplex and multiplex assays, with each developer including targets of interest for particular reasons. Commercial vendors have responded to the needs of the clinical laboratory through the development of syndromic test panels. These syndromic panels target the most common causes of viral respiratory tract infections, the most frequent causes of meningitis and sepsis, and (pertinent to this review) the common causes of infectious enterocolitis. These panels may replace the stool culture bench, while the Giardia/Cryptosporidium EIAs will likely significantly decrease the number of O&Ps that are performed. A number of these assays have received FDA clearance at the time of this writing, and several contain parasite targets. These assays are prepackaged and easy to use and target the bacteria, viruses, and parasites most likely to cause infectious diarrhea. The compositions of panels vary slightly from vendor to vendor, but each represents a new diagnostic tool worthy of thorough consideration.

Whether a molecular assay for a particular parasite is clinically useful (i.e., it represents a practical assay for implementation in a clinical laboratory and may improve turnaround time) is a question that will ultimately be determined in laboratories across the country. Although an assay may be technically feasible to perform, it may not be practical to implement for a variety of reasons. Potential problems include (i) the inability to obtain sufficient material for an adequate validation, a process that is under increased scrutiny by the FDA; (ii) clinical volumes being insufficient to support implementing the assay; and (c) the fact that traditional methods are competitive and possibly detect other pathogens. For example, one may initially consider a PCR assay for Ascaris lumbricoides to represent an advance in testing due to improved sensitivity over the traditional O&P. Practically, however, Ascaris is not commonly encountered in resource-rich countries, which are likely able to afford such an assay. More importantly, this assay detects only Ascaris, whereas the traditional O&P, albeit less sensitive in this scenario, detects a variety of gastrointestinal parasites, as well as commensal protozoa. The introduction of molecular tests for gastrointestinal parasites is, therefore, complicated. The patient population served, the medical question(s) being asked of the laboratory, alternative approaches, and technical, economic, and practical feasibility must all be considered and largely aligned to successfully implement these assays.

There are a number of challenges in the construction and application of multiplex assays. In addition to all the challenges encountered with monoplex assays (e.g., inhibitors, primer and probe design, etc.), the opportunity for intermolecular interactions increases significantly with each additional primer and probe set added to the mixture. Therefore, sophisticated primer and probe design programs and molecular biologists who understand these interactions are needed to design functional, complex assays. Regardless of the expertise of the technician, these multiplex assays, by the sheer nature of the increased intermolecular interactions, have a lower analytical sensitivity than corresponding monoplex assays. The most important question for clinical applications, even if the analytical sensitivity is reduced, is “Is the analytical sensitivity sufficient to produce a clinical sensitivity to appropriately categorize all patients with disease?” When this is the case, then a multiplex assay that is clinically useful has been produced. Fortunately, advances in specimen preparation (i.e., extraction methods), more-robust PCR (i.e., with DNA polymerases that are less susceptible to inhibition), and advances in PCR design software have afforded the creation of clinically useful multiplex PCR assays for enteric pathogens.

A large number of laboratory-developed PCR assays that target a variety of pathogens, depending on the interest of the investigators and the population served, have been developed. A full review of these is beyond the scope of this text, but select assays are reviewed. These studies have been important to demonstrate the feasibility of this approach to the detection of parasites in a complex matrix.

Some researchers have developed multiplex assays that target only select protozoal enteric pathogens. Stark and colleagues undertook the development and assessment of a multiplex PCR for the detection of four enteric protozoal parasites, Giardia, Cryptosporidium, Dientamoeba fragilis, and Entamoeba histolytica (266). They evaluated the newly designed multiplex PCR both against monoplex PCRs directed against the same targets and against traditional microscopy using modified iron-hematoxylin staining. They evaluated 427 fecal specimens that were routinely submitted to a clinical microbiology laboratory. The multiplex PCR detected Giardia in 28 specimens, D. fragilis in 26 specimens, E. histolytica in 11 specimens, and Cryptosporidium in 9 specimens; these results were uniformly corroborated by the four monoplex assays that targeted these organisms. The sensitivities and specificities of the morphological assessment for the four pathogens were, respectively, as follows: 50% and 100% for Giardia, 38% and 99% for D. fragilis, 47% and 97% for E. histolytica, and 56% and 100% for Cryptosporidium.

Taniuchi et al. took a different approach to the screening of stool specimens for intestinal parasites, selecting to screen for both select protozoal and select helminthic pathogens (267). This assay combined the high sensitivity of PCR with the differentiating capability afforded by the Luminex bead technology. In short, Taniuchi and colleagues developed two multiplex PCR assays, one of which contains primer mixes that target protozoal parasites and another that targets helminthic parasites. The PCR products from these reactions were then hybridized with Luminex beads that contain species-specific probes for enteric parasites. They assessed 319 fecal specimens and compared this approach with another previously described multiplex assay. They reported an 83% sensitivity and 100% specificity for this approach and recommend it as a possible screen for enteric parasites. Interestingly, in another assay, Taniuchi and colleagues developed a multiplex assay that detected Cyclospora, Cryptosporidium, and microsporidial targets, which would be particularly useful for immunocompromised patients (268).

Basuni et al. also targeted both protozoa and helminths, but with a slightly different approach (269). They designed two multiplex PCR assays, one that targeted select protozoa and another that targeted select helminths. The intestinal protozoal assay targeted E. histolytica, Giardia, and Cryptosporidium, whereas the intestinal helminth assay targeted Ancylostoma duodenale, Ascaris lumbricoides, S. stercoralis, and Necator americanus. Both assays contained internal amplification controls. The results of these multiplex assays were compared with microscopic examinations performed on direct smears and following zinc-sulfate concentration and Kato-Katz thick-smear techniques performed on 225 fecal specimens from patients suspected of having infections. Microscopy detected the presence of eight specimens positive for helminths, whereas the multiplex assay detected 46 (P < 0.001). Similarly, only 4 specimens were found to contain protozoa by microscopy, whereas 18 were detected by PCR (P < 0.001). Although the enhanced sensitivity of the molecular approach is evident, the importance of a broadly inclusive panel was also evident, as three instances of T. trichiura infections were detected by microscopy, but this organism was not present in the helminth multiplex PCR, so it was missed by that method.

There are several FDA-approved multiplex assays that include parasite targets, including the xTAG gastrointestinal pathogen panel (Luminex, Austin, TX) and the BioFire FilmArray gastrointestinal panel (bioMérieux, Durham, NC). The xTAG gastrointestinal pathogen panel was the first multiplex assay to receive FDA approval. This assay, which was originally formulated for the Luminex 100/200 system, has now been modified for the MAGPIX system. This modification retains the excellent performance of the assay on a new platform that is more user-friendly (i.e., it eliminates wash steps) and is a closed system. In addition to detecting 8 bacteria (9 in the international version) and 3 viruses, this assay detects Giardia, Cryptosporidium, and Entamoeba histolytica. An internal amplification control is also included in this assay.

Coste et al. compared this assay to conventional methods to study the etiologic agents of severe diarrhea in renal transplant recipients (270). Diarrheal stools from 49 patients were studied. Thirteen (23%) of the stool specimens were shown to contain an enteric pathogen by conventional methods, whereas 39 specimens (72%) were shown to contain an enteric pathogen using the xTAG gastrointestinal pathogen panel. Among these infected patients, one was found to have Giardia by conventional methods. When the specimens were studied with the xTAG gastrointestinal pathogen panel, the patient with Giardia was found to also have an infection with Campylobacter; additionally, another patient was found to be infected with Cryptosporidium, which was not discovered with conventional studies.

Perry et al. similarly disclosed advantages of a multiplex molecular platform when they compared two commercially available multiplex assays (i.e., the Luminex xTAG gastrointestinal pathogen panel and the Savyon Diagnostics gastrointestinal panel [available outside the United States]) with conventional diagnostics. They studied 1,000 clinical diarrheal stool specimens for the variety of pathogens included in these assays. Regarding intestinal parasites, Giardia was detected in 10 stools by the Luminex xTAG gastrointestinal pathogen panel, in 8 stools by the Savyon Diagnostics gastrointestinal panel, and in 6 stools by conventional diagnostics. The Savyon Diagnostics gastrointestinal panel contains D. fragilis, whereas the Luminex xTAG gastrointestinal pathogen panel does not. The Savyon assay detected D. fragilis in 45 stools, but no organisms of this species were detected by conventional diagnostics, and none, of course, were detected by the Luminex assay because D. fragilis was not included in the panel. The authors were able to corroborate the presence of D. fragilis in 44/45 of these stools by an alternative method (271). This study, again, confirms the need for expanded panels directed against a wider variety of parasites.

The FilmArray gastrointestinal panel is another commercially available product that has received FDA approval. In addition to detecting 12 enteric bacterial pathogens and five groups of viruses, this assay detects Cryptosporidium, Cyclospora, E. histolytica, and Giardia. The FilmArray reactions and analysis occur in a pouch that contains a number of chambers, wherein different reactions occur. Following a simple loading procedure, the pouch is placed into an instrument that controls the reactions and analysis. This simple-to-use approach brings complex molecular diagnostic capabilities to even small hospital laboratories. Buss and colleagues undertook a multicenter evaluation of the BioFire FilmArray gastrointestinal panel (272). This group prospectively collected and studied 1,556 clinical stool specimens and, in addition to the BioFire assay, tested these with conventional and other molecular assays. They determined the sensitivities of the FilmArray assay to be 100% for 12 of the 22 targets and >97.1% for 7 of the 22 targets, and the sensitivity could not be determined for the remaining analytes due to a low prevalence of the organism targets. The patients with parasitic infections detected in this study consisted of 27 patients with Giardia, 24 with Cryptosporidium, and a surprising 19 with Cyclospora, all of which were unsuspected and part of an outbreak. There were no infections with E. histolytica detected. A combination of traditional and molecular assays was used to determine the sensitivities and specificities of the individual parasites targeted within the BioFire FilmArray gastrointestinal panel. This study demonstrated the following sensitivities and specificities, respectively, for this assay: 100% and 99.5% for Giardia, 100% and 100% for Cyclospora, and 100% and 99.6% for Cryptosporidium.

There are always challenges that arise whenever new technologies are introduced and procedures are changed. One of the challenges with many organism-specific molecular diagnostic tests is that they detect only the organism for which they were designed. For example, a Giardia-specific PCR would be appropriately negative for a patient whose stool O&P disclosed Cystoisospora as the causative agent of infection. Broad-range and multiplex assays have addressed some of these issues, but gaps remain. The approach to building multiplex assays that address syndromes is generally good, since clinical findings alone are insufficient to differentiate the classes of the infectious agents causing disease (273). The current FDA-approved assays are excellent initial assays, but a second tier of assays is needed, particularly in parasitology. The helminths, including Strongyloides, and other parasites, such as D. fragilis, Blastocystis spp., Cystoisospora, and Microsporidia, among others, are not addressed. However, many of these inclusive panels are currently under development. Perhaps the most concerning challenge is the likely associated loss of microscopic morphological expertise when the majority of testing is converted from traditional to molecular assays. Next-generation sequencing, which consists of a variety of techniques to accomplish massive parallel sequencing, holds promise in several ways. Foremost, the usefulness of this type of technology to fully characterize the microbiomes of individuals with a particular disease, such as gastroenteritis, provides the opportunity to fully characterize all the microorganisms present in an attempt to determine the presence and type of pathogens, regardless of their taxonomic position. Furthermore, analysis of the transcriptome affords the ability to determine which microorganisms are likely involved in disease-producing processes and which are simply commensal or beneficial microbiota. Although this work and our understanding are nascent, this type of work will afford a more thorough understanding of the pathogens present, the pathophysiology of disease, and, finally, the construction of even more thorough syndrome-based assays.

Human parasitic infections caused by intestinal helminths and protozoans are the most prevalent infections in developing countries (Tables A7 to A11 and Fig. A1 and A2). There are several different species of intestinal protozoans, pathogenic and nonpathogenic, which have similar characteristics, so accurate identification can be difficult because of the tiny differences. The protozoans are grouped according to their locomotor organelles. The largest group contains the amoebae, which move with pseudopodia in the trophozoite form. There are specific criteria that are used to identify the trophozoite and cyst forms of the amoebae. Because of the minute details in structure required to identify these organisms, a good-quality microscope and good staining procedures are essential.

Tables A7 and A8 illustrate the details of the morphology of the trophozoite and cyst forms of the different species. Although Dientamoeba fragilis is considered to be a flagellate, the flagella are internal and not visible by light microscopy. Since they look more like amoebae, D. fragilis is grouped with the amoebae. The cyst form has recently been confirmed; however, the number of cysts in a human clinical specimen is extremely limited. Therefore, they are not included in Table A8.

Tables A9 and A10 illustrate the details of the trophozoite and cyst forms of the different species within flagellates. Some of the flagellates do not have a cyst form; therefore, no entry in Table A10 will be found for these organisms.

Table A11 illustrates the morphological details of the ciliates, the coccidians, the Apicomplexa, and Blastocystis spp. The most common pathogenic protozoan parasites are Giardia lamblia, Entamoeba histolytica, Dientamoeba fragilis, Cyclospora cayetanensis, and Cryptosporidium spp. If Blastocystis spp. are grouped with the pathogenic protozoa, they represent the most common parasites throughout the world and have been classified with the stramenopiles (brown algae) (5).

“Microsporidia” is the general term for the obligate intracellular parasites belonging to the phylum Microsporidia. They produce resistant spores of various sizes. Because they are extremely small, measuring from 1 to 4 μm in size, and have unique features, special stains are necessary to detect and identify them. www.cdc.goc/dpdx is an excellent website for laboratory diagnosis, images, and links (5).

Helminth infections are diagnosed by finding the characteristic eggs, which may vary in size from 25 to 180 μm, in the stool specimen, microscopic larvae, or macroscopic proglottids. The helminths are divided in two phyla: the Nematoda, or round worms, and the Platyhelminthes, which consist of the trematodes (flukes) and cestodes (tapeworms). Figure A1 contains images of nematode and cestode eggs, while Figure A2 contains images of trematode eggs.

The eggs have some biological variation but are uniform in size, shape, and coloration within each species. Size charts, identification keys, etc. are available (5). Excellent references are available for assistance and can be found in Appendix 4.