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Am J Primatol. 2019;e23063. wileyonlinelibrary.com/journal/ajp © 2019 Wiley Periodicals, Inc.
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https://doi.org/10.1002/ajp.23063
Received: 3 May 2019
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Revised: 10 September 2019
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Accepted: 20 September 2019
DOI: 10.1002/ajp.23063
RESEARCH ARTICLE
A multiyear survey of helminths from wild saddleback
(Leontocebus weddelli) and emperor (Saguinus imperator)
tamarins
Gideon A. Erkenswick
1,2
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Mrinalini Watsa
1,2
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Alfonso S. Gozalo
3
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Shay Dudaie
1
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Lindsey Bailey
1
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Kudakwashe S. Muranda
1
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Alaa Kuziez
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Patricia G. Parker
1,4
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Department of Biology, University of
Missouri‐St. Louis, Saint Louis, Missouri
2
Field Projects International, Saint Louis,
Missouri
3
Comparative Medicine Branch, National
Institute of Allergy and Infectious Diseases,
National Institutes of Health, Bethesda,
Maryland
4
WildCare Institute, Saint Louis Zoo, Saint
Louis, Missouri
Correspondence
Gideon A. Erkenswick, Department of Biology,
University of Missouri‐St. Louis, One
University Blvd., Saint Louis, MO 63121.
Email: gaet4b@umsl.edu
Funding information
Sigma Xi; American Society of Mammalogists;
Division of Intramural Research, National
Institute of Allergy and Infectious Diseases;
Whitney R. Harris World Ecology Center,
University of Missouri‐St. Louis; IdeaWild;
University of Missouri‐St. Louis, Grant/Award
Number: Trans World Airlines Scholarship;
Field Projects International
Abstract
The establishment of baseline data on parasites from wild primates is essential to
understand how changes in habitat or climatic disturbances will impact parasite–host
relationships. In nature, multiparasitic infections of primates usually fluctuate temporally
and seasonally, implying that the acquisition of reliable data must occur over time.
Individual parasite infection data from two wild populations of New World primates, the
saddleback (Leontocebus weddelli)andemperor(Saguinus imperator) tamarin, were collected
over 3 years to establish baseline levels of helminth prevalence and parasite species
richness (PSR). Secondarily, we explored variation in parasite prevalence across age and
sex classes, test nonrandom associations of parasite co‐occurrence, and assess the
relationship between group size and PSR. From 288 fecal samples across 105 individuals
(71 saddleback and 34 emperor tamarins), 10 parasite taxa were identified by light
microscopy following centrifugation and ethyl‐acetate sedimentation. Of these taxa, none
were host‐specific, Dicrocoeliidae and Cestoda prevalences differed between host species,
Prosthenorchis and Strongylida were the most prevalent. Host age was positively associated
with Prosthenorchis ova and filariform larva, but negatively with cestode and the
Rhabditoidea ova. We detected no differences between expected and observed levels of
co‐infection, nor between group size and parasite species richness over 30 group‐years.
Logistic models of individual infection status did not identify a sex bias; however, age and
speciespredictedthepresenceoffourandthreeparasitetaxa,respectively,with
saddlebacktamarinsexhibitinghigherPSR.Nowthatwehavereliablebaselinedatafor
future monitoring of these populations, next steps involve the molecular characterization
of these parasites, and exploration of linkages with health parameters.
KEYWORDS
baseline data, Callitrichidae, free‐ranging, Neotropics, parasite infections
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INTRODUCTION
Parasitism has a fundamental role to play in the persistence of animal
populations in nature, and the richness of parasite communities may
serve as effective population‐and ecosystem‐level measures of health
(Hudson, 1998; Hudson, Dobson, & Lafferty, 2006). In particular, helminth
parasites, which can be detected through noninvasive sampling, may be
ideally suited for long‐term health monitoring of a primate community
(Gillespie, 2006; Howells, Pruetz, & Gillespie, 2011). They are relatively
easy to evaluate from fecal samples collected from habituated primate
groups, but can sometimes be acquired in the absence of habituation by
scat detection dogs or by searching beneath known feeding or resting
locations (Arandjelovic et al., 2015; Orkin, Yang, Yang, Yu, & Jiang, 2016).
Several long‐term research programs have successfully used temporal
parasite data to examine ecological perturbations of threatened primate
populations (Bakuza & Nkwengulila, 2009; Chapman, Gillespie, & Speirs,
2005; Gillespie & Chapman, 2008; Gillespie, Chapman, & Greiner, 2005).
In contrast, in the absence of temporal data, comparative studies
between isolated and moreurbanprimatepopulationsareeffectiveat
evaluating impacts of increased contact with humans (Salzer, Deutsch,
Raño, Kuhlenschmidt, & Gillespie, 2010; Wenz, Heymann, Petney, &
Taraschewski, 2009). Despite the utility of such studies, parts of the
world with the highest primate diversity, such as the Neotropics, remain
inadequately sampled for naturally occurring helminth parasites (re-
viewed in Hopkins & Nunn, 2007, but also see Solórzano‐García & de
León, 2018), and reliable baseline data that are required to detect
changes over time and space are not commonly available.
Since the range of factors that can explain parasite–host patterns
in nature is large (Clough, Heistermann, & Kappeler, 2010; Gillespie,
Barelli, & Heistermann, 2013; Gillespie et al., 2010; MacIntosh,
Hernandez, & Huffman, 2010; MacIntosh et al., 2012; Monteiro,
Dietz, Raboy, et al., 2007; Muehlenbein & Watts, 2010; Nunn,
Brezine, Jolles, & Ezenwa, 2014; Telfer et al., 2008) and often
dependent on the environment and time (Clough et al., 2010;
Gillespie et al., 2010), it may be best approached through longitudinal
monitoring of individuals in host communities (Clutton‐Brock &
Sheldon, 2010; Erkenswick, Watsa, Gozalo, Dmytryk, & Parker, 2017;
Stuart et al., 1998). A primary challenge has been that research on
wild primates usually requires habituation to observers, which often
constrains sample sizes, making it difficult to adequately analyze
many of these factors (Williamson & Feistner, 2011). Thus far, a
multitude of studies have offered snapshots of parasite prevalences,
focused on just one or two parasites of known interest, the sampling
of a single primate host, or on data from health inspections, or
necropsies, after animal extraction from the wild. Collectively they
have created a broad foundation of primate parasite data (see Nunn
& Altizer, 2005 for a detailed compilation; Solórzano‐García & de
León, 2018).
The next challenge will be to determine baseline levels of
prevalence and species richness, which will enable more detailed
studies that incorporate host demography and development, mode of
transmission, and change over time at the level of a population. As an
example, for almost a half‐century it has been well known that New
World monkeys are broadly infected by Plasmodium brasilianum,a
quartan malarial parasite, that may in fact be the same as the human
parasite Plasmodium malariae (Collins & Jeffery, 2007; Lalremruata
et al., 2015). However, only last year do we have the first evidence
that it may persist in a highly aggregated manner among a small
number of chronically infected nonhuman primate hosts (Erkenswick,
Watsa, Pacheco, Escalante, & Parker, 2017). In addition, long‐term
studies that incorporate more than one primate host are essential to
examine several longstanding hypotheses of how sociality influences
parasite prevalence, intensity, and diversity (Altizer et al., 2003;
Freeland, 1976, 1979), as are long‐term studies of multiple sympatric
species to examine species‐specificity of infection dynamics.
The Callitrichidae (comprised of tamarins and marmosets) are
small arboreal primates that are widely distributed throughout the
forests of South America (Sussman & Kinzey, 1984). They are
frequently found in sympatry with other New World monkeys and in
some cases have proven relatively resilient and flexible in the face of
encroachment by human populations (Gordo, Calleia, Vasconcelos,
Leite, & Ferrari, 2013; G. C. Leite, Duarte, & Young, 2011; Soto‐
Calderón, Acevedo‐Garcés, Álvarez‐Cardona, Hernández‐Castro, &
García‐Montoya, 2016). Part of their ecological flexibility may be due
to their generalist diets that include fruits, insects, tree exudates, and
fungi (Sussman & Kinzey, 1984), a characteristic that also could
expose them to a wide array of parasites that are dispersed by
intermediate arthropod hosts. Studies of endoparasites of callitri-
chids have documented overlap with other primate families including
the Ateledae, Cebidae, and Aotidae (Michaud, Tantalean, Ique,
Montoya, & Gozalo, 2003; Phillips, Haas, Grafton, & Yrivarren,
2004; Tantalean, Gozalo, & Montoya, 1990; Wolff, 1990). Consider-
ing the approximately 61 species and subspecies of Callitrichidae
(Rylands and Mittermeier, 2009), there have been only a handful of
comprehensive evaluations of helminth parasites from free‐ranging
populations (Monteiro, Dietz, Beck, et al., 2007; Müller, 2007; Wenz
et al., 2009), and only two species in which parasites have been
monitored routinely over time—the golden lion and golden‐headed
lion tamarins (Leontopithecus rosalia and L. chrysomelas, respectively;
Monteiro, Dietz, Raboy, et al., 2007).
The principle aim of this study was to characterize the helminth
assemblages from two populations of sympatric, individually identifi-
able, free‐ranging callitrichids—the saddleback tamarin (Leontocebus
weddelli, formerly Saguinus fuscicollis weddelli; Buckner, Lynch Alfaro,
Rylands, & Alfaro, 2015; Matauschek, Roos, & Heymann, 2011) and
emperor tamarin (Saguinus imperator)—from fecal samples collected
noninvasively and via an annual mark‐recapture program. By
sampling these hosts across 3 years, we estimate the prevalence of
helminth parasites, species richness, and the extent of parasite
overlap between the two host species. We also summarize changes in
infection status from the subset of animals that were screened for
helminths in two or more consecutive years. In doing so, we establish
baseline data for future comparative studies following perturbations
such as changing weather patterns due to climate change, habitat
loss/modification, or greater human encroachment.
Secondarily, as sampling came from an individually identifiable
population, we analyzed how parasite prevalence varied by host
demographic variables. As a result of greater social burdens placed
on females to compete for dominant breeding opportunities, we
predicted that an age–sex interaction would influence prevalence
and parasite species richness. We also analyzed infection data for
nonrandom associations between all pairwise parasite combinations,
on the basis that blood parasite associations were detected
previously in the same populations (Erkenswick, Watsa, Gozalo,
et al., 2017). Lastly, prior studies have detected a potential pattern
between primate group size and parasite species richness (Cote &
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Poulin, 1995; Nunn, Altizer, Jones, & Sechrest, 2003; Rifkin, Nunn, &
Garamszegi, 2012; Vitone, Altizer, & Nunn, 2004), but this associa-
tion has not been considered within the Callitrichidae, at small
taxonomic scales. We tested the hypothesis that large groups will
harbor higher parasite species richness.
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METHODS
All sampling protocols adhered to guidelines outlined by the
American Society of Mammalogists (Sikes & Gannon, 2011) and
complied with the American Society of Primatologists Principles for
the Ethical Treatment of Non‐Human Primates. Permissions for this
study were obtained from the Institutional Animal Care and Use
Committee at the University of Missouri‐St. Louis and the Directo-
rate of Forest and Wildlife Management (DGFFS) of Perú annually.
The DGFFS also granted export permits for the samples, while the
CDC and U.S. Fish and Wildlife Services approved the import of these
samples into the United States.
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Field site and study subjects
Sample collection took place annually from 2012 to 2014 in the
Madre de Dios Department of Southeastern Perú at the Estación
Biológica Rio Los Amigos (EBLA; 12°34′07″S, 70°05′57″W), which is
managed by the Asociación para la Conservación de La Cuenca
Amazonica. This site was previously known as the Centro de
Investigación y Capacitación Río Los Amigos (CICRA), see Watsa,
Erkenswick, Rehg, and Pitman (2012) for further description of the
site, including a detailed map of the location. The field station was
created in 2000, and aside from selective logging, the forest remains
intact. All sampling took place within a forest trail system that covers
approximately 900 ha of tropical rainforest that is adjacent to the Los
Amigos Conservation Concession inside the buffer zone of Manu
National Park. The collection area consists of five, nonmutually
exclusive, forest types including terra firme, primary forest, bamboo,
palm swamp, floodplain, and successional/disturbed forests (Pitman,
2008). There are two distinct seasons each year at this site—the wet
season from October to March (average monthly precipitation > 250
mm), and the dry season from April to September (136 mm ± SD
19 mm; Watsa, 2013). All sampling took place during the dry season,
from May–July each year, precluding the study of the effects of
seasonality on the parasite community in these primates.
Three callitrichines at this site, the saddleback tamarin, emperor
tamarins, and the more cryptic Goeldi’s monkey (Callimico goeldii;
Watsa et al., 2012), share forest habitat with eight other primate
species including three species of Cebidae, and two species each of
Atelidae and Pithecidae, as well as owl monkeys (Aotus nigrifrons;
Watsa, 2013). At EBLA, both S. imperator and L. weddelli have average
group sizes of 5 (range of 3–8) individuals and group compositions
are similar (Watsa, Erkenswick, & Robakis, 2017; Watsa et al., 2015).
The primary differences between S. imperator and L. weddelli are adult
weight, 515 ± 66 and 386 ± 86g, respectively, and nuances in feeding
behavior including greater amounts of fungi consumption in S.
imperator (pers. obs.; Terborgh, 1985).
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Sample collection and storage
Since 2009, an annual mark‐recapture program has been implemen-
ted on ~70 saddleback and emperor tamarins by Field Projects
International (Watsa et al., 2015). During capture, each individual is
permanently tagged with a microchip (Microchip ID Systems,
Covington, LA), and was made visually identifiable by unique patterns
of bleached rings around the tail, as well as a tricolor beaded
necklace that signified group, sex and individual identity (for the full
capture protocol see Watsa et al., 2015). In addition to collecting
fecal samples at the time of capture, we used radio telemetry to track
tamarins in 14 groups each year via a radio collar placed on the
breeding female in each group (Wildlife Materials, Murphysboro IL).
We also used both full (sleep‐site to sleep‐site, spanning ~11 hr) and
half‐day (minimum 5 hr) follows to opportunistically collect fecal
samples from all group members as they were produced.
Each animal in the study was classified into one of three age
classes based on dental eruption patterns (Watsa, 2013). Juveniles
were defined as individuals whose adult teeth were absent or not
fully erupted (<11 months old). Subadults were animals with adult
teeth, but that were juveniles in the preceding year. All remaining
individuals were assigned to the adult age class. Due to small sample
sizes from the subadult class, the juveniles and subadult classes were
combined to analyze the effects of age on parasite prevalence.
Upon sample collection during mark‐recapture and follows, all
fecal samples were transferred using sterile technique into numbered
plastic bags and stored in a chilled thermos. Upon return to
basecamp, each sample was fixed in 10% neutral buffered formalin
(1:2, feces to preservative ratio). For each sample, we recorded
species, individual ID, group, date, time of day, and type of collection
(follow or trapping event). Only samples produced by identified
individuals were included in this study. All samples were exported to
the University of Missouri‐St. Louis for analyses.
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Laboratory analysis
Isolation of parasite cysts, eggs, and larvae from fecal samples
followed a two‐step process based on sedimentation procedures as
per MacIntosh et al. (2010) and Zajac and Conboy (2012). In Step 1,
we used a fecal straining procedure in which fecal samples were (a)
diluted in 10% neutral buffered formalin, (b) strained of large debris
through cheese cloth into a plastic cup, (c) transferred to a 15‐ml
falcon tube with an empty weight already recorded, (d) centrifuged at
800gfor 5 min to form a fecal pellet, (e) removed of the supernatant
and weighed, (f) resuspended and homogenized in 5 ml of 10%
formalin. In Step 2, we followed the centrifugal sedimentation test
outlined by Zajac and Conboy (2012) with 1ml of the homogenized
suspension from Step 1. Sedimentations from Step 2 were resus-
pended in exactly 1 ml of preservative, and 80 µl aliquots were placed
onto clean slides with 22× 22 mm coverslips for full evaluation with an
ERKENSWICK ET AL.
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Olympus CX31 light microscope (Center Valley, PA) using ×200
magnification (micrographs were taken at higher power). Evaluations
of parasites were timed and tabulated using a free online data counter,
COUNT (http://erktime.github.io/count/), and each unique infection/
sample was documented with multiple micrographs taken with a Leica
ICC50 HD camera (Allendale, NJ). Three separate aliquots per sample
were evaluated with each evaluation taking an average of 10 min.
Unless infections were too rare, standard length and width
measurements from 10 representative micrographs per parasite per
species were recorded with a calibrated ruler in Image J (https://
imagej.nih.gov/ij/) to the nearest 1 µm. Sizes and measurements of all
parasite forms were compared with known references values in the
literature and identified to the lowest taxonomic scale possible.
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Statistical analysis
For each animal and for each year of the study, if a parasite was
detected in one or more fecal samples it was considered a positive
parasite infection. Average prevalence, as well as the proportion of
individuals that acquired infection, lost infection, or showed no change
in infection status, was calculated for each helminth identified by
microscopy across the 3‐year study period. Average change in
infection status was determined by selecting all instances where an
individual was sampled across a 2‐year period, either 2012–2013 or
2013–2014, and computing the mean number of individuals that
acquired, lost, or did not change infection status. Differences in annual
helminth prevalence between host species were tested with a two‐
tailed Fisher’s Exact Test, followed by p‐value adjustment for multiple
comparisons using the Holm–Bonferroni method, αlevel of .05 (Holm,
1979). To test for variation in the presence of parasitic infections
across host variables we used mixed‐effect logistic regression models
with a binary response variable and binomial errors. Fixed effects
included “sex,”“age class,”and “species”and random effects included
“animal identity”and “year”to accommodate individual resampling and
possible interannual variation. We also incorporated the number of
samples collected per animal per year as an offset to account for
temporal sampling bias (Walther, Cotgreave, Price, & Gregory, 1995).
Parasite species richness, which was a discrete numerical response
variable, was analyzed with an identical model formula but using
Poisson errors. Model selection for all models was carried out with
step‐wise term deletion by removing nonsignificant factors and
comparing nested models with a likelihood ratio test.
To test for significant correlations between group size and parasite
species richness we calculated rarified parasite community richness
estimates per group. The use of species accumulation curve estimates
are advocated by Walther et al. (1995), because raw values of parasite
community richness are easily biased by uneven sampling. We used
Spearman’s rank correlations to test if parasite community richness
estimates with similar sampling effort were associated with group size.
To identify any nonrandom parasite co‐occurrences, we compared
the prevalence of all observed pairwise co‐infections with expected
estimates of co‐infection (calculated as prevalence of A × prevalence of
B). We then plotted expected against observed values to identify
discordant levels of co‐infection, and if applicable, used a two‐sample
z‐test to compare proportions. All statistical analyses were performed
in R (R Development Core Team, 2015).
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RESULTS
In total, we collected 288 individually identified fecal samples from
105 unique tamarins (71 L. weddelli,34S. imperator) distributed across
13 groups of L. weddelli and 7 groups of S. imperator. The number of
samples collected per individual per year ranged from 1 to 7, with a
mean of 1.6± 0.98. The average fecal sample weight, following Step 1
in sample processing (see Methods), was 0.41 ± 0.22 g. Across the
study period, we were able to assess individual infection status from
two or more fecal samples 40% (n= 71) of the time. Within this 40%,
redetection of a parasite infection is significantly correlated with
parasite prevalence for both species (for L. weddelli r (8) = .95, p<0.05,
for S. imperator r (7) = .69, p< 0.05), meaning that common infections
were relatively easy to redetect but rare parasites were difficult. This
pattern held for all resampling within a year, whether obtained from a
trapping event or collected during a primate follow. Considering all sex
and age classes, our sampling included slightly more males than
females across years, and subadults of both host species were the least
sampled age group (Table 1).
We were able to differentiate 10 helminth parasites by
morphology (Figure 1). The parasite ova were grouped based on
morphology, length, width, color, shell, and internal characteristics,
and compared with New World nonhuman primate helminth ova
descriptions reported in the literature. Suspect genus and/or species
for each morpho‐group was suggested based on ova characteristics
and their similarity to those of parasites known to affect Callitrichi-
dae in the wild (Table 2). All but one rare parasitic infection, an
unidentified ova (Group A, Table 2), were found in both host species,
although prevalence profiles varied (Tables 3 and S2 for post‐hoc
adjusted p‐values). Prevalence for the Dicrocoeliidae was signifi-
cantly higher in S. imperator (Fisher’s test mean‐adjusted p< 0.05),
and Cestoda was significantly higher in L. weddelli (Fisher’s test mean‐
adjusted p< 0.05; Table 3). Our rarest parasite infections for both
host species were the unidentified ova and large larvated ova (Group
H, likely Primasubulura spp.), and also Cestoda for S. imperator. Out of
TABLE 1 Numbers of individuals sampled by species, sex, age
class, and year
L. weddelli S. imperator
Year 2012 2013 2014 2012 2013 2014
Sex 36 46 34 18 23 19
Male 19 28 17 7 14 10
Female 17 18 17 11 9 9
Age class
Juvenile 893633
Subadult 461231
Adult 24 31 30 10 17 15
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176 parasite screening events (animal‐years), at least one parasite
was detected all but five times.
With caution due to the error associated with redetecting rare
parasites, we report that individual infections status remained
primarily unchanged (Figure 2). Among S. imperator, this pattern
differed slightly for strongylid ova and Dicrocoeliidae (Figure 2 and
Table S1). Among L. weddelli, we only detected slightly more frequent
changes in individual infection status for Cestoda.
Considering each year separately, we did not detect any
significant deviations between expected and observed prevalence
of co‐infection (Figure 3); the largest absolute difference in
prevalence across all parasite combinations throughout the study
period was 0.07. We also found no evidence of a relationship
between group size (ranged 3–8) and estimated parasite species
richness within groups after controlling for sampling effort (Spear-
man’s rank correlation = −0.08, p= .665, n= 30).
Out of the 10 helminths identified, 6 were common enough to
evaluate their distributions across host species, sex and age class
variables; the unidentified ova, Gongylonematidae, and the large
larvated ova (Primasubulura sp.) were too rare to analyze prevalence
patterns using statistical models. No parasitic infection exhibited a
significant sex bias; however, age and species did predict the
presence of 4 and 3 parasites, respectively (Table 4). Relative to L.
weddelli,S. imperator was positively associated with Dicrocoeliidae
but negatively associated with cestode and strongylid ova. Relative
to adults, juveniles and subadults were negatively associated with
Prosthenorchis and filariform larva, but positively associated with
cestode and the Rhabditoidea ova. Our models of parasite species
richness identified host species as the only significant predictor
considered in this study (Table 4), which had a significantly negative
estimate for S. imperator.
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DISCUSSION
The results provided here, in combination with recent works on
hemoparasites at the same site (Erkenswick, Watsa, Gozalo, et al.,
2017), represent a benchmark against which future parasitological
surveys can be compared. Included in the parasite assemblages found
here were parasites that could be transmitted directly/indirectly or
FIGURE 1 Micrographs and measurements for each parasite
ERKENSWICK ET AL.
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trophically, which may be differentially impacted by anthropogenic
disturbance and climate change. Worth noting, we also detected
differences from previous studies of parasites in congeneric
callitrichids. In northern Peru, both Müller (2007) and Wenz et al.
(2009) conducted snapshot surveys on sympatric callitrichids, S.
fuscicollis and S. mystax, and reported a parasite assemblage that
overlaps with our findings (including Prosthenorchis,Hymenolepis,
large and small spirurids, Primasubulura, and strongylid larvae).
Phillips et al. (2004) screened a group of S. fuscicollis in the nearby
Tambopata National Reserve and identified four parasites (Trichuris,
Iodamoeba,Entamoeba, and an unidentified strongyle), none of which
could be confirmed in our study. In this study, all samples were fixed
in formalin, which is not always preferred for screening of protozoan
parasites (Zajac & Conboy, 2012). Müller (2007) and Wenz et al.
(2009) also recorded higher prevalence than our study for every
helminth except Prosthenorchis, which was considerably less common.
These differences could be explained to some extent by, for instance,
the small sample size of four individuals in the study by Phillips et al.
TABLE 2 Parasite ova and larva characterization and classification
Group Name Characterization
A Unidentified ova Eggs are subglobose with a slightly flattened side and a smooth and thick transparent shell. Eggs
morphologically resemble Spirura guianensis ova but are unembryonated (Cosgrove, Nelson, & Jones, 1963;
Thatcher and Porter, 1968). A new species, S. delicata, was reported in a tamarin but the ova not described
(Vicente, Pinto, & Faria, 1992).
B Gongylonematidae Eggs are ovoid with a thick smooth transparent shell and larvated. Suspect genus Gongylonema spp. (Orihel &
Seibold, 1972; Strait et al., 2012).
C Unidentified cestoda ova Eggs are spherical with a clear mucus thick layer and embryonated, morphologically similar, but smaller, than
Hymenolepis spp. and Paratriotaenia spp. (Guerrero, Serrano‐Martínez, Tantaleán, Quispe, & Casas, 2012;
Müller, 2007).
D Strongylid ova Eggs are ellipsoidal, slightly tapered at one end, with a smooth thin shell and are unembryonated. Suspect
genera Molineus spp. Possibly M. vexillarius (Cogswell, 2007; Cosgrove et al., 1968; Dunn, 1961; Stone, Conga,
& Santos, 2016).
E Small embryonated ova Eggs are ellipsoidal, symmetrical, with a smooth, relatively thick shell and larvated. Two common nematodes in
tamarins with similar egg morphology are Trypanoxyuris spp. (Carrasco, Tantaleán, Gibson, & Williams, 2008;
Conga et al., 2014; Guerrero et al., 2012; Stone et al., 2016; Thatcher & Porter, 1968) and Trichospirura
leptostoma (Orihel & Seibold, 1971; Vicente et al., 1992; Vicente, de Oliviera Rodrigues, Corrêa Gomes, &
Pinto, 1997). Other parasites with similar ova occasionally reported in New World nonhuman primate species
are Physaloptera dilatata and Longistriata dubia. However, Ortlepp (1922) describes P. dilatata eggs as oval,
thick‐shelled, and measuring on average 39 × 27 µ, and Vicente et al. (1997) and Gibbons and Kumar (1980)
describe L. dubia as having thin‐shelled ova measuring 62–79 × 31–41 µ. P. dilatata eggs are not ellipsoidal and
are smaller than the ova found in the current study and L. dubia eggs are larger and with a thinner shell
compared with the ova found in the current study. Hence, the ova found are most likely Trypanoxyuris spp.
and/or Trichospirura leptostoma.
F Dicrocoeliidae ova Eggs are ovoid, golden‐brown with a thick shell and operculum. Based on ova morphology and size the suspect
species is Athesmia foxi =heterolecithoides (Cogswell, 2007; Thatcher & Porter, 1968).
G Rhabditoidea ova Eggs are ellipsoidal, with a smooth thin shell, and larvated. Suspect genus Strongyloides, possibly S. cebus
(Cogswell, 2007; Conga et al., 2014; Guerrero et al., 2012; Mati, Junior, Pinto, & de Melo, 2013; Parr, Fedigan,
& Kutz, 2013; Stone et al., 2016).
H Large larvated ova
(Primasubulura)
Eggs are large, globular, with a thick irregular transparent shell and embryonated. Ova morphology is
characteristic for Primasubulura spp. Possibly P. jacchi or P. distans (Rocha, 2014; Tavela et al., 2013; Thatcher
and Porter, 1968; Vicente et al., 1997).
I Acanthocephala ova
(Prosthenorchis sp.)
Characteristic eggs are ovoid, brown, thick walled with three layers, the outer shell has fine reticular sculpting,
contains an acanthor. Suspect species Prosthenorchis elegans. Other species occasionally reported in
Callitrichidae is P. lenti, however, the ova for this species are much larger than the ones found in the current
study (Cogswell, 2007; Müller, 2007; Orihel & Seibold, 1972; Thatcher & Porter, 1968).
J Filariform larva Translucent larvae with a very long, sometimes coiled, thin tail. Tiny notch on tip of tail, characteristic of
Strongyloides larvae, was not observed. In addition, larvae were larger than reported Strongyloides rhabditiform
larvae. Little (1966) describes all stages of S. cebus larvae which are morphologically very different from the
larvae found in the tamarins.S. cebus larvae have a short and very muscular esophagus, something we did not
observe in the larvae found in the tamarins, and a shorter tail. Moreover, S. cebus eggs do not hatch until they
leave the body of the host and since the feces were collected immediately post defecation, chilled and within
6 hr placed in fixative, it is unlikely these larvae are Strongyloides spp. The morphology of the larvae found in
the tamarins agree with previous descriptions of Filariopsis barretoi larvae in South American monkeys.
Suspect genus Filariopsis spp, possibly F. barretoi (Lee, Boyce, & Orr, 1996; Orihel & Seibold, 1972; Parr et al.,
2013; Stone et al., 2016).
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(2004), but broader differences are possibly explained by ecologically
distinct sites, including a higher helminth diversity due to increased
primate species diversity (11 species compared with 4). However, an
additional factor could be a consequence of prior human activities
(such as hunting), which increased the densities of small primate
species at EBLA (Pitman, 2008; Rosin & Swamy, 2013).
Our study underscores that wild animals generally maintain multiple
parasitic infections during their lives (Cox, 2001; Petney & Andrews,
1998), and that the combination of parasites may be particularly
important. Of the 10 helminths documented in this study, 6 are of
unknown pathogenicity, 2 are probably non‐pathogenic, and 2 are known
to be pathogenic (Table 2). In this study population, Prosthenorchis sp.
(Phylum Acanthocephala) is among the most prevalent, and has been
hosted by some individuals for upwards of 12 years (Erkenswick,
unpublished data) despite numerous reports of high pathogenicity (King,
1993; Strait, Else, & Eberhard, 2012): the thorny‐headed worms attach
themselves to the intestinal mucosa of a primate host and cause
inflammatory responses, obstruction of the lumen, and lesions and ulcers
that lead to secondary infections or even peritonitis in the worst cases.
Infections with Acanthocephala have been frequently reported in wild
callitrichids (Müller, 2007; Tantalean, Gozalo, & Montoya, 1990; Wenz
et al., 2009) and occasionally in other New World primates such as the
Cebidae and Atelidae (King, 1993; Phillips et al., 2004; Wenz et al., 2009).
This suggests that observations of particularly pathogenic parasites
should be taken in the context of the broader parasite community and
changes in the environment, which may be best achieved with
longitudinal data collection (Haukisalmi, Henttonen, & Tenora, 1988).
Although our analysis did not identify nonrandom associations
between co‐infecting parasites, it is still possible that within‐host parasite
interactions are at play. Presence‐absence infection data is less sensitive
TABLE 3 Average annual prevalence by host species and parasite
Leontocebus weddelli Saguinus imperator
Class Parasite % SD %SD Diff Dispersal Pathogenic
Acanthocephala Prosthenorchis sp. 85 0.04 78 0.08 0.07 Trophic Yes
Cestoda Hymenolepis or Paratriotaenia 44 0.02 7 0.02 0.37
a
Trophic Unknown
Nematoda Unidentified ova 6 0.06 0 0 0.06 Trophic Possibly
b
Primasubulura 4 0.05 7 0.08 0.03 Trophic Unknown
Rhabditoidea ova 16 0.08 21 0.07 0.05 Trophic Unknown
Gongylonematidae 15 0.09 13 0.06 0.02 Trophic No
Sml embryonated ova 19 0.1 39 0.04 0.2 Trophic Unknown
Filariform larva 43 0.11 4 0.16 0.03 Direct Unknown
Strongylid ova 83 0.07 76 0.22 0.07 Direct No
Trematoda Dicrocoeliidae 8 0.1 4 0.12 0.32
a
Trophic Unknown
Note: Pathogenic classification reviewed in Müller (2007) with hosts appearing to tolerate well helminthic infections in the wild.
Abbreviations: % , average prevalence across the study period; Diff, difference in average prevalence between the host species; SD, standard deviation.
a
Significant differences (Fisher’s exact p< .05).
b
Spirura guianensis has been documented as pathogenic (See Group A, Table 2).
FIGURE 2 Averaged number of changes in individual infection status by host species across the study period
ERKENSWICK ET AL.
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at detecting relationships than quantitative measures of parasite intensity
or burden (Knowles et al., 2013; Lello, Boag, Fenton, Stevenson, &
Hudson, 2004). Estimates of parasite intensity were omitted from the
present study as it remains uncertain how well eggs per quantity of feces
actually represents intensity of infection (Gillespie, 2006). Our hypothesis
that larger groups would show higher parasite species richness was not
supported. This finding is consistent with observations of blood parasites
in these primates (Erkenswick, Watsa, Gozalo, et al., 2017), and may be a
consequence of low group size variation (3–8 inds.). Some callitrichids can
occur in larger groups, for example, Callithrix at 15 members (Pontes & da
Cruz, 1995; Watsa et al., 2017), but it is also possible that group size and
parasite diversity cannot be linked within the Callitrichidae. Alternatively,
the majority of parasite taxa in this study are trophically transmitted,
rather than directly transmitted, which may not be influenced by group
size to the same extent.
Interpreting these findings in light of parasite mode of transmission,
direct or trophic, is challenging because although the majority of the
parasites detected were trophically transmitted via an intermediate host
(usually an arthropod), these intermediates are unknown in most cases.
While a comprehensive study of feeding ecology is a good next step in
our host populations, two studies on sympatric S. fuscicollis and S. mystax
(closely related to S. imperator)inNorthernPerúagreethatS. fuscicollis
spends significantly more time foraging in the lower strata and on the
ground, while the opposite was true of S. mystax (Heymann, Knogge, &
Tirado Herrera, 2000; Smith, 2000). Smith (2000) documented distinct
feeding preferences based on color and size of prey, a niche
specialization, if present at our site, that might account for observed
host differences in Dicrocoeliidae and Hymenolepis infections. This could
also explain variation in parasite species richness among these hosts via
differences in intermediate host encounter rates or the persistence of
parasite free‐living phases on the ground or in certain forest strata.
Consistent with the pattern of prevalence of blood parasites in this
population (Erkenswick, Watsa, Gozalo, et al., 2017), age class predicted
the presence of two trophically transmitted parasites, though we
obtained both positive and negative relationship estimates from our
models. We suspect that differences in diet and foraging efficiency
between younger and older individuals underlie different parasite
encounter rates, but differences in immune status, and cumulative
parasite exposure as animals age are also possibilities.
By tracking prevalence of parasitic infections over time in wild
populations it is possible to infer the stability of natural parasite
communities; however, longitudinal data at the level of the individual
provides insights into the source, or lack thereof, of population stability
FIGURE 3 Observed versus expected prevalence of parasite co‐
infection. Each dot represents a unique pairwise combination of
parasites
TABLE 4 Generalized linear mixed model outcomes for each parasite and parasite species richness
Parasite Fixed effect BSEWald (χ
2
)DF p‐Value
Prosthenorchis Intercept 0.7772 0.4963
Age: Subadult −1.4417 0.5459 6.9742 1 <0.05
Cestoda Intercept −2.4263 0.3764
Species: Simp −4.1473 0.8134 25.998 1 <0.05
Age: Subadult 1.9867 0.6172 10.36 1 <0.05
Rhabditoidea ova Intercept −4.2509 0.5863
Age: Subadult 1.3999 0.5733 5.9619 1 <0.05
Filariform larva Intercept −1.7792 0.3472
Age: Subadult −1.5110 0.6875 4.8303 1 <0.05
Strongylid ova Intercept 0.4347 0.4500
Species: Simp −0.8282 0.4322 3.6726 1 0.055
Dicrocoeliidae Intercept −4.4469 0.7193
Species: Simp 2.0681 0.4903 17.794 1 <0.05
PSR Intercept −0.4953 0.1528
Species: Simp −0.6648 0.1874 12.577 1 <0.05
Sml. embryonated ova Could not reject null models
Note: Minimal, best‐fit models for the presence of each parasite and parasite species richness (PSR). Model selection began with fixed factors host “sex,”
“age class,”and “species,”while “individual identity”and “year”were incorporated as random effects, and the number of fecal samples collected for each
individual/year was included as a model offset. Infection data were insufficient to analyze the distribution of Unidentified ova, Gongylonematidae, and
Primasubulura.Simp denotes S. imperator.
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(Knowles et al., 2013). In some cases, it could even aid in identification of
parasites that have negative health consequences. For example, if
parasite prevalence is consistently low relative to the incidence of new
infections across years, and there is little evidence that individuals clear
infections, then previously infected hosts must be disappearing regularly.
In this study, we report few disparities in the rate of acquisition or loss of
parasites, and considered in concert with observed prevalence, we see no
obvious signs of negative health consequences for these parasites.
However, consideration of individual infection status should be mindful of
the high correlation between redetection and parasite prevalence when
using data derived from microscopic evaluations.
Molecular parasite identifications will be critical to assure data
comparisons over time, geographic distance, across multiple laboratories,
and to differentiate between species whose eggs or other reproductive
stages are identical morphologically (Solórzano‐García & de León, 2018).
Microscopy‐based detection of parasites can lack sensitivity (Garamszegi,
2009) and be prone to bias. For example, we observed a strong positive
correlation between redetection of parasite infections across multiple
samples from the same individuals of the same year and overall parasite
prevalence. Moreover, it is possible that the same parasite population
across two closely related hosts can exhibit higher or lower levels of
subpopulation structure (Levin & Parker, 2013; McCoy, Boulinier, &
Tirard, 2005), reflecting more or less cross species sharing of parasites.
On the other hand, complete reliance on parasite molecular markers
ignores variation in parasite life stages and the discovery of unexpected
parasite infections, although the latter may be addressed if reliable
universal eukaryotic markers are developed (Hadziavdic et al., 2014;
Hugerth et al., 2014). Also, accessing the tools necessary to employ
molecular methods is still much more difficult than those for light
microscopy, though new efforts are underway to address the technology
disparity in locations were primate biodiversity is highest (Watsa,
Erkenswick, Pomerantz, & Prost, 2019). Hence, a dual approach of
utilizing molecular and microscopy methods to screen for parasite
infections is the current ideal (Valkiūnas et al., 2008).
This study provides the first description of the helminth parasite
assemblage of free‐ranging S. imperator, alongside a new comparative
data set from L. weddelli (formerly S. fuscicollis weddelli; Buckner et al.,
2015; Matauschek et al., 2011). Although the IUCN currently classifies
both species as “least concern”(Rylands & Mittermeier, 2008), S.
imperator has a vastly smaller species range than L. weddelli,andthecore
of its distribution in Peru overlaps with a rapidly expanding illegal gold
mining hub (Asner, Llactayo, Tupayachi, & Luna, 2013), threatened by
forest fragmentation and mercury contamination. S. imperator is currently
one of the most valuable Peruvian monkeys in the illegal wildlife trade
(Watsa, 2015). These factors emphasize the importance of this data set
for this region and its comparative nature allows us to dig deeper into
factors that could affect parasite distributions in wild primates.
In conclusion, we maintain that reliance on baseline parasite
infection data may become increasingly important to understand how
a fast changing physical and climatic environment is impacting
natural wildlife populations. The Callitrichidae are nearly ubiquitous
across South American rainforests, have a propensity to be found in
sympatry with other New World primates, and some species readily
exist in and around human‐altered landscapes. They also evidence
flexible, omnivorous diets that include autotrophs, fungi, insects, and
small vertebrates. As such, regular study of the parasites from this
family could serve as a potential flagship for the regional detection of
ecological changes, or even environmental threats.
ACKNOWLEDGMENTS
We wish to thank research assistants from Field Projects International
who aided in the collection of tamarin fecal samples. We want to thank
Cindee Rettke for her assistance coordinating wet laboratory analyses,
and Aaron Erkenswick for creating a web application that was used to
record infection data. We express our gratitude to the following agencies
and organizations that provided non‐monetary support to this investiga-
tion: Centro de Ecologia y Biodiversidad, Amazon Conservation Associa-
tion, el Estación Biológica Río Los Amigos. This study was supported by
Field Projects International, the American Society of Mammalogists,
University of Missouri Trans World Airlines Scholarship Program,
Whitney R. Harris World Ecology Center, Sigma Xi Society, IdeaWild,
and the Intramural Research Program of the National Institutes of
Health, National Institute of Allergy and Infectious Diseases, Comparative
Medicine Branch.
DATA AVAILABILITY STATEMENT
The data that have been used in the production of this article are
available upon request. For a copy of the raw data, send an inquiry
email to the corresponding author.
ORCID
Gideon A. Erkenswick http://orcid.org/0000-0001-6040-1170
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SUPPORTING INFORMATION
Additional supporting information may be found online in the
Supporting Information section.
How to cite this article: Erkenswick GA, Watsa M, Gozalo AS,
et al. A multiyear survey of helminths from wild saddleback
(Leontocebus weddelli) and emperor (Saguinus imperator)
tamarins. Am J Primatol. 2019;e23063.
https://doi.org/10.1002/ajp.23063
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