INTRODUCTION

Parasites play a central role in ecosystems, affecting the ecology and evolution of specific interactions (Esch and Fernandez, 1993), host population growth and regulation (Hochachka and Dhondt, 2000; Hudson et al., 1998), and community biodiversity (Hudson et al., 2002). Parasites can impact host survival and reproduction directly through pathologic effects and indirectly by reducing host condition (Boyce, 1990; Chandra and Newberne, 1977; Coop and Holmes, 1996; Dobson and Hudson, 1992; Hudson et al., 1992). Severe parasitosis can lead to blood loss, tissue damage, spontaneous abortion, congenital malformations, and death (Chandra and Newberne, 1977; Despommier et al., 1995). However, less severe infections are more common and may impair nutrition, travel, feeding, predator escape, and competition for resources or mates, or increase energy expenditure (Coop and Holmes, 1996; Dobson and Hudson, 1992; Hudson et al., 1992; Packer et al., 2003).

The close phylogenetic relationship between humans and nonhuman primates results in high potential for pathogen exchange (Ott-Joslin, 1993; Wolfe et al., 1998). Recent studies have highlighted emerging human diseases, e.g., HIV/AIDS, Ebola, with origins or likely transmission to humans that involves nonhuman primates (Gao et al., 1999; Leroy et al., 2004). Likewise, evidence from well studied ape populations suggests that epidemics of polio, respiratory diseases, and scabies originated from humans (Hill et al., 2001; Kalema-Zikusoka et al., 2002). As human population density continues to increase exponentially, speeding the reduction and fragmentation of primate habitat, greater human-primate contact is inevitable and higher rates of pathogen transmission are likely. Baseline data on patterns of parasitic infections in wild primate populations are critical to provide an index of population health and to begin to assess and manage disease risks. In addition, considering the evolutionary and ecological linkages between primates and their parasites (Stuart and Strier, 1995), one can view parasites as indicator species, potentially alerting us to imminent threats to primate conservation.

Though many studies have documented the gastrointestinal parasites of wild populations of African apes (Ashford et al., 1990, 2000; Lilly et al., 2002; McGrew et al., 1989), baboons (Appleton et al., 1986; Eley et al., 1989; Hahn et al., 2003; Müller-Graf et al., 1997), and howlers (Stoner, 1996; Stuart et al., 1990, 1998), the gastrointestinal parasites of other primate taxa remain poorly known (cf. Gillespie et al., 2004, 2005a; Stuart et al., 1993). In addition, though researchers have accomplished much admirable work, primate parasite studies have often made use of divergent methodologies, compromising the potential for longitudinal or comparative work. Here, I provide practical guidelines and standardized methodologies for the noninvasive assessment of gastrointestinal parasite infections in free-living primate populations.

PRACTICAL GUIDELINES FOR THE STUDY OF PRIMATE PARASITES

Use Standard Terminology

The American Society of Parasitologists has standardized parasitological terminology (Margolis et al., 1982), which Stuart and Strier (1995) and Bush et al. (1997) have reviewed. However, studies of primate parasites frequently perpetuate the use of inappropriate terms. Using vague language such as parasite load or alternatives to standardized terms may lead to misinterpretation of results or compromise the value of findings to the broader scientific and conservation communities. Noninvasive studies of primate parasites can readily provide data on presence or absence, richness, and prevalence of parasitic infections.

Avoid Use of Egg Counts to Provide a Measure of Infection Intensity

The aforementioned invalid assumption is pervasive in the primate parasite literature. Intensity is the number of adult individuals of a particular taxon infecting an individual host. Many factors affect the number of parasite eggs contained in host fecal material and they cannot reliably provide an index of adult worm burden, i.e., intensity. Studies of livestock; laboratory animals, including primates; and humans demonstrate that host immunity, density-dependent factors, and environmental cues can depress worm ovulation (Christensen et al., 1995; Roepstorff et al., 1996; Stear et al., 1995) and inherent differences in parasite fecundity, size, age, and sex ratio also affect egg output (Coadwell and Ward, 1982; Dineen et al., 1965; Stear et al., 1995). Even in parasite taxa in which a linear relationship exists between egg production and adult worm burden—the case for a limited number of taxa—variation in fecal condition, e.g., moisture content and consistency, and inherent temporal and spatial sampling heterogeneity, compromise intersample comparisons (Anderson and Schad, 1985; Eberhard et al., 2001). Studies of domesticated animals suggest that egg counts may provide a qualitative measure of intensity, i.e., low, moderate, high (Anderson and Schad, 1985; Tarazona, 1986); however, the generality of the relationship requires testing. Hence, noninvasive studies of primate parasites should not report intensity.

Avoid Collecting Anonymous Samples

Knowing the identity or even the age and sex of individual primates sampled improves data quality exponentially. Unless samples are from known individuals without duplication, one cannot treat them as independent data points (OIE, 2004), which limits conclusions that one can draw regarding prevalence and specific interactions for both the primates and the parasites. In addition, samples not collected immediately after defecation may be contaminated, potentially leading to misinterpretations regarding host specificity for a given parasite taxa (MAFF, 1979). Anonymous samples can be of value for providing basic presence or absence information, and researchers should seek them for long-term monitoring of unhabituated populations.

Ensure Adequate Sample Size

Though the number of independent fecal samples required for a study will depend on the questions asked, samples required to determine whether a parasite is present in a primate population are the minimum. The minimum sample size n required to detect at least 1 infection is calculated via the following formula, where α is the significance level and p is the prevalence in the population:

$$n = \ln \,(a)/\ln \,(1 - p)$$

Accepting a 0.05 level of significance and recognizing the broad range of values for gastrointestinal parasite prevalence in free-ranging primates, minimum required sample size depends greatly on expected prevalence of the parasite of interest (Fig. 1). For general surveys of primate parasites, one should apply an assumed prevalence of 5% (Leech and Sellers, 1979; OIE, 2004; Putt et al., 1988), resulting in a minimum requirement of 60 independent fecal samples (Fig. 1). Studies incorporating comparisons over space and time, among age and sex categories, or among species should aim for substantially more samples.

Fig. 1.
figure 1

Minimum sample size required to be 95% confident of finding a parasite in a population of free-ranging primates in relation to prevalence.

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Present Details of Parasite Identification

Gastrointestinal parasite classification by fecal analyses is weak by its very nature. Consequently, it is critical that researchers present details of the characteristics used to identify a given parasite taxa, e.g., color, size, defining features, and provide conservative identifications as warranted by data collected. Researchers should measure eggs, larvae, and cysts using a calibrated micrometer (n=10/species/sample) and photograph representatives (MAFF, 1979). Trichostrongyloid, strongyloid, and rhabditoid nematode eggs are similar in size and appearance, making differentiation extremely difficult (OIE, 2004), and one should culture larvae to contribute information not available from egg traits alone. Opportunistic necropsies of primates dying of natural causes allow for critical validation of egg identifications. Presentation of details of parasite identification will facilitate longitudinal and comparative studies.

Provide Supportive Data

Details regarding the environment, density, behavior, and health status of primates sampled can provide critical information that may inform patterns of parasitism observed. The following is not intended as a comprehensive overview of the topic, but instead a synopsis to stimulate future discussion of the issues.

Habitat attributes can strongly influence patterns of parasitism in free-ranging primates. For example, studies have demonstrated infection prevalence to be higher in primates ranging in humid compared to more arid habitats (Hausfater and Meade, 1982; Stuart et al., 1990, 1993). Evidence of general patterns of seasonal infection in primates is equivocal, with clear seasonal patterns of infection for some primate species (Freeland, 1977; Huffman, 1997), and no clear pattern for others (Gillespie et al., 2004, 2005a). Lastly, recent studies demonstrate that patterns of habitat disturbance including selective logging and forest fragmentation can affect primate-parasite dynamics in dramatic ways (Gillespie and Chapman, 2005; Gillespie et al., 2005b). Consequently, researchers should collect data, or cite published data, on climate, habitat, patterns of disturbance, and history of the environment in which they sample primates for parasites to improve our understanding of the interplay.

Density estimates for the primate of interest, as well as sympatric primates and other potential hosts of generalist parasites, also provide critical information. Host density is of central importance to infection rates in directly transmitted parasites (Poulin, 1998), and intraspecific studies have demonstrated that host density positively correlates with parasite prevalence and diversity (Morand and Poulin, 1998; Packer et al., 1999). However, studies that have concurrently examined primate density and habitat characteristics in relation to primate parasite prevalence reveal that habitat characteristics may be a better predictor of parasitic infection than primate density is (Gillespie, 2004; Gillespie and Chapman, 2005; Stuart et al., 1993). Future studies are needed to distinguish more systematically the effects of host density and habitat characteristics.

A wide variety of primate behaviors including patterns of ranging, grooming, interindividual and intergroup associations, and foraging may influence patterns of parasitic infection. For example, yellow baboons (Papio cynocephalus) appear to avoid potential infection by regularly rotating their sleeping sites (Hausfater and Meade, 1982). Likewise, mangabeys (Lophocebus albigena) presumably reduce the risk of contact with fecal contamination and consequent infection by traveling further on days of heavy rainfall and avoiding foraging in the same areas on consecutive days (Freeland, 1980). Similarly, Gilbert (1997) suggests that red howlers (Alouatta seniculus) reduce contact with parasites by consistently defecating above gaps in the forest vegetation.

Grooming likely affects patterns of parasitism in complex ways. Though grooming appears to be an important mechanism for removing potentially damaging ectoparasites (Freeland, 1981; Gilbert, 1997), there is no published study on how grooming behavior may alter the risk of gastrointestinal parasite infection. Grooming brings individuals into close contact, increasing the risk of transmission of parasites with direct life cycles. In addition, ectoparasites often act as intermediate hosts for gastrointestinal parasites (Despommier et al., 1995; Muller and Baker, 1990; Poulin, 1998). Consequently, primates may unintentionally infect themselves with parasites with intermediate hosts by ingesting ectoparasites while grooming.

Group size may also affect primate parasitic infections. Freeland (1779) found a significant correlation between the size of groups of mangabeys and blue monkeys (Cercopithecus mitis) and the number of protozoan infections each group maintained, and Freeland (1980) demonstrated that the prevalence of protozoan infections increased with group size for mangabeys. A meta-analysis of a wide variety of host-parasite relationships showed a positive correlation between intensity of infection by parasites and host group size (Côté and Poulin, 1995). It is expected that for generalist parasites, the frequency and duration of mixed-species associations affect patterns of parasitism in similar ways. Thus, by limiting group size and the frequency of multispecies associations, primates may be able to limit their risk of parasitic infection.

Patterns of primate foraging may also affect patterns of parasitism. Researchers have systematically documented dietary self-medication of parasitic infections in the great apes (Huffman, 1997; Huffman and Wrangham, 1994), and anecdotal evidence supports the potential generality of this behavior among primates and other taxa (Garber and Kitron, 1997; Glander, 1994; Janzen, 1978; Phillips-Conroy, 1986). Consequently, primatologists should be aware of the potential role self-medication may play in the patterns of parasitism they observe.

Though parasites are a normal component of a functioning ecosystem and low-intensity infections are often asymptomatic (Anderson and May, 1979), anthropogenic change may result in altered transmission rates, parasite host range, and parasite virulence (Daszak et al., 2000; Patz et al., 2000). Resultant changes in host susceptibility may result in elevated morbidity and mortality, and ultimately, population declines. By evaluating fecal samples for symptoms of illness, i.e., diarrhea, blood, etc., before collection, researchers provide data that one can analyze in relation to patterns of infection to examine potential relationships between parasitic infections and pathology or disease in free-ranging primates.

STANDARDIZED METHODOLOGIES

The following protocols are intended to provide primatologists with simple yet effective methods to evaluate primate gastrointestinal parasitic infections based on the collection and analysis of fecal samples.

Collection of Fecal Samples for Gastrointestinal Parasite Analysis

  1. 1.

    Prepare collection tubes containing 10% buffered formalin (preferred for helminths) or polyvinyl alcohol (preferred for protozoa).

  2. 2.

    Before collecting feces, examine macroscopically for, and note, consistency, presence of blood, mucus, tapeworm proglottids, and adult or larval nematodes.

  3. 3.

    With gloved hands, use a wooden applicator or spatula to scoop a ca. 2-g sample from within the fecal mass into the collection tube. By taking the sample from within the fecal mass, you reduce risks of contamination by free-living nematodes in the immediate environment.

  4. 4.

    Close tube and label with identification no., date, time, initials of collector, primate species, site (ideally Global Positioning System [GPS] coordinates), and age/sex/identity of individual sampled if possible.

  5. 5.

    Shake the tubes vigorously to maximize contact between sample and storage solution.

  6. 6.

    Collect replicate sample in similar fashion using RNAlater (Ambion) as preservative if planning molecular confirmation of parasite identities.

Techniques for Recovery and Examination of Gastrointestinal Parasites

Direct Smear

This method involves examining a thin smear of fecal material with normal saline on a microscope slide. Though direct smear can demonstrate the presence of helminths and protozoa, it is not ideal because it is effective only when egg, larvae, or cyst, or all, concentrations are high. In addition, large amounts of detritus in feces can interfere with identifications and quantitative assessment of egg production is not possible (MAFF, 1979). Consequently, I recommend the method only as a supplemental procedure. A combined recovery using fecal flotation and sedimentation techniques provides the best results.

Fecal Flotation

The method is optimal for separating many helminth eggs and protozoan oocysts and cysts from fecal debris. Occasionally, parasitic mites groomed from the skin will float as well. Solutions of saturated NaCl or sugar can be effective, but NaNO3 is optimal. ZnSO4 and MgSO4 are unsuitable for general analyses because they will not isolate many of the nematodes commonly infecting wild primates. The following protocol is appropriate for the analysis of primate fecal samples preserved in formalin or polyvinyl alcohol.

  1. 1.

    Add 1 g of feces to centrifuge tube.

  2. 2.

    Fill centrifuge tube 2/3 with distilled water and homogenize fecal pellet with a wooden applicator.

  3. 3.

    Centrifuge samples at 1800 rpm for 10 min.

  4. 4.

    Pour off supernatant.

  5. 5.

    Resuspend fecal material in NaNO3 solution.

  6. 6.

    Fill tube to meniscus with NaNO3 solution, and place microscope cover slip on lip of tube.

  7. 7.

    Centrifuge samples at 1800 rpm for 10 min.

  8. 8.

    Remove cover slip from centrifuge tube and place on a slide labeled with the sample number.

  9. 9.

    Scan slide using the ×10 objective lens of a compound microscope and identify and count all parasite eggs, larvae, and cysts. Use the ×40 objective lens for measurement and confirmation of identifications.

  10. 10.

    Scan slide thoroughly under ×40 objective lens to confirm presence or absence of protozoan cysts (add a drop of iodine to facilitate identification).

  11. 11.

    Measure the length and width of individual eggs, cysts, and larvae using a calibrated ocular micrometer.

  12. 12.

    Photograph representatives.

If examining undiluted fresh samples, omit steps 2–4.

Fecal Sedimentation

The method allows for the isolation and identification of trematodes (flukes) which, unlike other helminths, are too heavy to float up in NaNO3 solution. The following protocol is appropriate for the analysis of primate fecal samples preserved in formalin or polyvinyl alcohol and can use the fecal pellet remaining after the previously described flotation methodology.

  1. 1.

    Suspend fecal pellet in 40 ml of sedimentation solution (dilute soapy water) in a 50-ml beaker.

  2. 2.

    Filter the suspension through cheesecloth held over the lip of the beaker into a 50-ml centrifuge tube. Rinse cheesecloth with sedimentation solution and refilter through cheesecloth. Dispose of cheesecloth and remaining fecal pellet.

  3. 3.

    Allow filtered suspension to settle until sediment is apparent (5 min).Footnote 1

  4. 4.

    Remove supernatant by pipette and rinse remaining material into disposable beaker with sedimentation solution.

  5. 5.

    Repeat until supernatant is clear.

  6. 6.

    Transfer 5 drops of sediment to a slide labeled with the sample number and cover with 2 cover slips placed side by side.

  7. 7.

    Scan slide under ×10 objective lens and identify and count all parasite eggs, larvae, and cysts. Use the ×40 objective lens for measurement and confirmation of identifications.

  8. 8.

    Scan slide thoroughly under the ×40 objective lens to confirm presence or absence of protozoan cysts (add a drop of iodine to facilitate identification).

  9. 9.

    Measure the length and width of individual eggs, cysts, and larvae using a calibrated ocular micrometer.

  10. 10.

    Photograph representatives.

Fecal Cultures

The similarities in size and appearance of the eggs of different species of gastrointestinal nematodes are such that their differentiation is extremely difficult. Their third-stage larvae, however, are sufficiently different and it is possible to distinguish between different genera, and species in some cases. The method below is suitable for the culture of trichostrongyloid, strongyloid, and rhabditoid larvae.

  1. 1.

    Transfer 5 g of fresh feces to a beaker and thoroughly mix with an equivalent volume of vermiculite, using distilled water as needed (ca. 10 ml).

  2. 2.

    Tie off mixture inside 20×20 cm cheesecloth using a ca. 50-cm fishing line or string.

  3. 3.

    Suspend mixture in a 100-ml container with lid.

  4. 4.

    Add distilled water to container until bottom of cheesecloth makes contact with surface of water.

  5. 5.

    Allow culture to incubate at room temperature (20–21°C) for 2 wk, checking culture daily for fungal growth and water level.

  6. 6.

    Spray mixture with a fine mist of water every few days to deter fungal growth and add water as needed.

  7. 7.

    After incubation, collect larvae by pipette and transfer onto a microscope slide for examination and identification. A drop of Lugol’s iodine solution (1 g of iodine, 2 g of potassium chloride, 300 ml of distilled water) can facilitate identification.

Opportunistic Necropsy

The following protocol will allow for the isolation of adult and immature gastrointestinal parasites from primates found dead.

  1. 1.

    Procure a fecal sample from the individual for later analysis.

  2. 2.

    Ligate and remove stomach and intestines from the individual.

  3. 3.

    Tie off stomach, small intestine, large intestine, and colon with fishing line or strong string.

  4. 4.

    Wash each section of gastrointestinal tract separately as described below.

  5. 5.

    Cut open stomach and collect contents in a container.

  6. 6.

    Above same container, wash stomach wall thoroughly with water, carefully rubbing mucous membrane to remove any worms.

  7. 7.

    Slowly pour small amounts of wash onto wire-mesh stackable sieves (top screen aperture of 0.15 mm [for recovering adult worms] and bottom screen aperture of 0.038 mm [for recovering immature worms].

  8. 8.

    Wash material on screen until clear water passes through.

  9. 9.

    After washing all material in this way, invert each screen and wash adhering material into separate containers.

  10. 10.

    Carefully examine the surface of the stomach with dissecting microscope for parasites that remain attached.

  11. 11.

    Treat each section of intestine as stomach.

Preservation of Recovered Helminths

Helminths preserved in the following manner can be delivered to a qualified parasitologist for identification.

Nematodes

Because of the thick cuticle of the worms, optimal fixation requires use of a hot solution. Thoroughly wash nematodes in 2 or 3 changes of water or 1% saline, then transfer them to hot (70–80°C) 70% alcohol or 5% formalin. After cooling, store in clean fluid of the same kind.

Cestodes

Carefully wash tapeworms in 1% saline. Fix in 5–10% formalin between 2 pieces of glass or by dipping repeatedly while suspending posterior with forceps. The techniques will facilitate subsequent identification by minimizing contraction by the parasite.

Trematodes

Vigorously shake flukes in 1% saline, replace saline with 5–10% formalin while continuing to shake. Vigorous shaking throughout this process prevents contraction by the parasite.

CONCLUSIONS

Primatologists are situated to provide high-quality and needed data on gastrointestinal parasite infections in wild primate populations. The guidelines and methodologies presented here will help to achieve the goal.