DNA of Dientamoeba fragilis detected within surface-sterilized eggs
of Enterobius vermicularis
Dennis Rösera,⇑, Peter Nejsumb, Anne Josefine Carlsgarta, Henrik Vedel Nielsena,
Christen Rune Stensvolda
aLaboratory of Parasitology, Department of Microbiology and Infection Control, Statens Serum Institut, Artillerivej 5, DK-2300 Copenhagen S, Denmark
bDanish Centre for Experimental Parasitology, Department of Veterinary Disease Biology, Faculty of Health and Medical Sciences, University of Copenhagen,
Dyrlægevej 100, 2, DK-1870 Frederiksberg, Denmark
h i g h l i g h t s
" Eggs of Enterobius vermicularis were
" DNA was extracted from individual
" Dientamoeba fragilis and E.
vermicularis PCR was performed on
" Sequencing of PCR amplicons
showed evidence of D. fragilis-
" Results support hypothesis of D.
fragilis transmission by E.
g r a p h i c a l a b s t r a c t
a r t i c l ei n f o
Received 10 April 2012
Received in revised form 13 September 2012
Accepted 19 October 2012
Available online 29 October 2012
a b s t r a c t
With no evidence of a cyst stage, the mode of transmission of Dientamoeba fragilis, an intestinal protozoon
of common occurrence and suggested pathogenicity, is incompletely known. Numerous studies have sug-
gested that eggs of intestinal nematodes, primarily Enterobius vermicularis (pinworm), can serve as vec-
tors for D. fragilis, although attempts to culture D. fragilis from pinworm eggs have been unsuccessful and
data from epidemiological studies on D. fragilis/pinworm co-infection have been conflicting.
The aim of this study was to investigate whether we could detect D. fragilis DNA from pinworm eggs
collected from routine diagnostic samples (cellophane tape) and surface-sterilised by hypochlorite.
DNA was extracted from individual eggs and tested by PCR using D. fragilis- and E. vermicularis-specific
primers; amplicons were sequenced for confirmation.
In cellophane tape samples from 64 patients with unknown D. fragilis status we detected D. fragilis DNA
in 12/238 (5%) eggs, and in a patient known to harbour D. fragilis we detected D. fragilis DNA in 39/99
The finding of D. fragilis DNA within eggs of E. vermicularis strongly supports the hypothesis of D.
fragilis-transmission by pinworm and has implications for antimicrobial intervention as well as control
and public health measures.
? 2012 Elsevier Inc. All rights reserved.
Dientamoeba fragilis is a common yet little studied intestinal
protozoon suspected of causing gastrointestinal illness, particularly
in children (Windsor and Johnson, 1999; Johnson et al., 2004; Stark
et al., 2010a; de Wit et al., 2001). Despite being discovered nearly a
0014-4894/$ - see front matter ? 2012 Elsevier Inc. All rights reserved.
⇑Corresponding author. Fax: +45 32683033.
E-mail addresses: firstname.lastname@example.org (D. Röser), email@example.com (P. Nejsum), carlsgart@
yahoo.dk (A.J. Carlsgart), firstname.lastname@example.org (H.V. Nielsen), email@example.com (C.R. Stensvold).
Experimental Parasitology 133 (2013) 57–61
Contents lists available at SciVerse ScienceDirect
journal homepage: www.elsevier.com/locate/yexpr
century ago (Jepps and Dobell, 1918), research on D. fragilis has
been scarce, in part due to diagnostic difficulties (Calderaro et al.,
icance (Gijsbers et al., 2011). While some recent publications advo-
cate for the inclusion of this flagellate in the list of enteric
pathogens to be tested for in patients with gastrointestinal illness
(Stark et al., 2010a; Barratt et al., 2011a), there is still a lack of con-
clusive evidence of pathogenicity and the mechanisms hereof.
Since D. fragilis has no known cyst stage and trophozoites de-
grade rapidly in stool samples (<48 h) (Hakansson, 1936; Barratt
et al., 2011b), traditional microscopy of faecal concentrates is unli-
kely to detect D. fragilis (Johnson et al., 2004), and molecular diag-
nostics are preferred because of higher sensitivity and specificity
(Stensvold and Nielsen, 2012; Stark et al., 2010b). A recent review
on reported prevalences of D. fragilis worldwide showed a preva-
lence of 0.3–52%, depending on cohort investigated and diagnostic
method used (Barratt et al., 2011a). In Danish patients suspected of
enteroparasitic disease, Stensvold et al. (2007a,b) found a D. fragilis
prevalence of 11.7% using a permanent staining technique (Stensvold
et al., 2007a), however, a recent Danish study using real-time PCR
found a D. fragilis prevalence of 10–70%, depending on age group
The mode of transmission for D. fragilis is still unknown, and
there is neither evidence of a (pseudo-)cyst stage for D. fragilis,
nor has faecal–oral transmission by trophozoites been shown to
occur (Johnson et al., 2004; Barratt et al., 2011b).
For decades it has been suggested that eggs of nematodes, par-
ticularly Enterobius vermicularis (pinworm), could serve as vectors
for D. fragilis (Burrows and Swerdlow, 1956; Ockert, 1972; Girgin-
kardesler et al., 2008) (Fig. 1).
Transmission of protozoa via helminth eggs has previously been
shown for the bird helminth Heterakis gallinae (s. gallinarum) and
the protozoon Histomonas meleagridis (Smith and Graybill, 1920;
Ruff et al., 1970), an organism phylogenetically closely related to
D. fragilis (Gerbod et al., 2001). Like D. fragilis, pinworm has a global
distribution and a high prevalence among children (Burkhart and
Burkhart, 2005; Lacroix and Sorensen, 2000; Barratt et al.,
2011a), and previous studies have used epidemiological data to
test for association between D. fragilis and E. vermicularis. However
such studies can neither confirm nor reject vector transmission of
D. fragilis by pinworm, as they are cross-sectional and cannot take
into account differences in infection duration, making determina-
tion of cause and effect difficult.
The lifecycle and transmission of D. fragilis was recently re-
viewed by Barratt et al. (2011a), who found the pinworm vector
hypothesis controversial but plausible, and called for studies
employing PCR, DNA sequencing and electron microscopy to sub-
stantiate findings (Barratt et al., 2011b).
In a previous attempt to test this hypothesis using molecular
diagnostics, Menghi et al. (2005) successfully amplified pinworm
and D. fragilis DNA from a solution of pinworm eggs, but were un-
able to amplify D. fragilis DNA once the eggs had been treated with
DNase, indicating that the amplified D. fragilis DNA had been lo-
cated on the surface of the eggs only, and not within (Menghi
et al., 2005). In the present study we present a method that enables
reliable detection of D. fragilis DNA within the eggs of helminths.
Moreover, we report for the first time the successful detection of
D. fragilis DNA inside eggs of E. vermicularis, endorsing the hypoth-
esis that pinworm may indeed serve as a vector for D. fragilis.
2. Materials and methods
2.1. Method development
Since our aim was to be able to detect DNA of D. fragilis poten-
tially lodged inside individual pinworm eggs, a method was devel-
oped for surface-sterilizing and collecting individual eggs, while
preserving sufficient DNA within the egg to be detectable by PCR.
Hypochlorite (HClO) is a bactericidal agent known to induce
damage to DNA, RNA, and polynucleotides (Hawkins and Davies,
2002), and has previously been used to remove the uterine, but
not the chitin layer, of nematode eggs (Oksanen et al., 1990).
In order to determine the HClO concentrations sufficient to de-
grade DNA, we prepared eight solutions of 1 mL HClO (10?2–10?9)
(Sodium Hypochlorite 1%, BDH PROLABO), to which we added 1 lL
(undiluted) DNA extracted from an adult female pinworm using
QIAamp DNA Mini Kit (QIAGEN, Hilden, Germany) with a 100 lL
eluate. From these initial eight HClO-DNA solutions we then
prepared 10-, 100- and 1,000-fold dilutions, using DNase/RNase
Fig. 1. The presumed life cycle of D. fragilis. Transmission of D. fragilis by faecal–oral route and via nematode eggs (e.g., Ascaris, Enterobius spp.). There is currently no evidence
of an infective cyst or a pseudocyst stage for D. fragilis, nor has faecal–oral transmission by trophozoites, the only known stage of D. fragilis, been shown to occur. Life cycle
image courtesy of CDC–DPDx; shown with modifications.
D. Röser et al./Experimental Parasitology 133 (2013) 57–61
free water, in order to show if subsequent lowering of the HClO
concentration would result in PCR amplification (Table 1).
We then incubated all solutions for 10 min at room tempera-
ture, performing PCR with a 5 lL template and standard PCR con-
ditions and using primers targeting E. vermicularis (Table 2).
Two methods were employed for washing eggs after HClO incu-
bation and for DNA isolation/membrane rupture. The first method
ing) by centrifugation, aspirating the supernatant and washing the
pellet with DNase/RNase-free water to lower the HClO concentra-
tion below 10?4(=0.01%), the threshold concentration as deter-
mined in our hypochlorite trials. Single eggs were then subjected
to proteinase K treatment and subsequently PCR was performed
(see below). The second method employed washing single eggs
(single-egg washing) by transferring a single egg (within 1 lL 10?2
HClO) into a solution of lysis buffer and proteinase K (both provided
with the QIAamp DNA Mini kit, QIAGEN, Hilden, Germany), per-
turer, which also lowered the HClO conc. below 10?4before PCR.
Both methods proved feasible and enabled successful amplifica-
tion and sequencing of E. vermicularis DNA, though multiple-egg
washing resulted in a greater loss of eggs (through aspiration),
and single-egg washing proved more laborious. We also noted that
proteinase K was unable to rupture eggs not treated with HClO,
and that washing with DNase/RNase-free water was not sufficient
to remove DNA contaminants from eggs (data not shown).
Between Dec. 1st 2010 and Dec. 1st 2011 we collected clinical
samples from Danish patients (cellophane tape) at Statens Serum
Institut. We analysed each sample for egg density using light
microscopy, discarding samples with <10 eggs per visual field, as
it proved difficult to harvest a sufficient number of eggs from such
samples. We collected a total of 64 individual samples from 64 pa-
tients with unknown D. fragilis-status, and 1 sample with a high
egg load (>1,000 eggs per visual field) from a patient known to har-
bour D. fragilis (confirmed by real-time PCR as previously described
(Stensvold et al., 2007b; Stensvold and Nielsen, 2012)). The cello-
phane tape samples were processed averagely 36 days after
2.3. Egg recovery and surface sterilization
Eggs were scraped off cellophane tape samples using a scalpel
and transferred to a small glass jar containing 1 mL 10?2HClO
and incubated for 10 min at room temperature.
2.4. Egg washing and DNA isolation
In samples from patients with unknown D. fragilis-status, we
used single-egg washing, transferring individual eggs suspended
in 1 lL 10?2HClO into a 1.5 mL Eppendorf tube, using a micropi-
pette and a light microscope (Nikon SMZ6454, Nikon Nordic,
Copenhagen, Denmark), thus visually confirming the presence of
one egg. Between 1 and 16 individual eggs from each sample were
collected, depending on the number of eggs harvested into the
glass jar. To each tube, 180 lL of ATL buffer and 20 lL of proteinase
K (QIAamp DNA Mini kit) were added, and the remaining steps in
the DNA extraction were carried out according to the recommen-
dations of the manufacturer; however, only 20 lL AE buffer was
used to elute the DNA in the final step.
In the sample from the patient known to harbour D. fragilis, we
used multiple-egg washing (Section 2.1), transferring the entire
Hypochlorite trials; determination of hypochlorite (HClO) concentrations sufficient to degrade DNA of E. vermicularis. Row A1 shows initial concentrations of HClO (1% = 10?2),
row A2 shows initial concentration of pinworm DNA (1 lL undiluted DNA added to 1 ml 1% HClO i.e. 0.1% = 10?3) and row A3 shows obtained PCR products/presence of gel band.
Rows B1–B3, C1–C3 and D1–D3 show 10-, 100- and 1000-fold dilutions (using DNase/RNase free water) of the HClO/pinworm DNA solutions from row A1–A2 and obtained PCR
products. Rows E1–E2 show obtained PCR products from the pinworm DNA concentrations used, without added HClO.
Pinworm DNA conc.
Pinworm DNA conc.
Pinworm DNA conc.
Pinworm DNA conc.
Pinworm DNA conc.
Pinworm DNA conc.
Primers used for amplification of DNA from D. fragilis and E. vermicularis.
OrganismPrimer namePrimer sequence GenePCR product size
(in base pairs (bp))
DFpn_1f50-GCC AAG GAA GCA CAC TAT GG-30
50-Terminal of the SSU rRNA gene
(GenBank accession No. AY730405)
DFpn_364r50-GTA AGT TTC GCG CCT GCT-30
50-CAA CAC TTG CAC GTC TCT TCA-30
50-ATT GCT CGT TTG CCG ATT AT-30
50-Terminal end of the 5S region of rRNA gene195 bp
D. Röser et al./Experimental Parasitology 133 (2013) 57–61
content of the glass jar to a 1.5 mL Eppendorf tube, and wash-
ing ? 3 with 1 mL DNase/RNase free water (centrifugation at
1,500g for 5 min). After the final wash, we aspirated 10 lL of the
washing water from the supernatant, and used this as control for
DNA contamination of the water, potentially caused by hatched
or crushed eggs. Individual eggs were transferred in 1 lL of fluid
into 0.2 mL PCR tubes using a micropipette and a light microscope,
visually confirming the presence of one egg, and preparing a total
of 100 single eggs. To each PCR tube we added 1 lL Proteinase K
(QIAGEN, Hilden, Germany) and 8 lL water, spun the mixture (con-
taining one egg in 10 lL) briefly (5 s), and incubated it at 50 ?C for
30 min, then 100 ?C for 15 min.
2.5. PCR and sequencing
For conventional PCR, we used the following conditions: initial
denaturation at 95 ?C for 15 min, followed by 35 cycles of 94 ?C for
30 s, 54 ?C for 30 s and 72 ?C for 30 s; and a final elongation step at
72 ?C for 5 min (Primus HT, Biotech, Ebersberg, Germany). From
DNA extracted by use of the Qiagen protocol, we used a template
of 4 lL (of 20 lL eluate from one pinworm egg) and from the pro-
teinase K extractions, we used 5 lL (of 10 lL solution from one
pinworm egg). PCR components were as follows: 2.5 lL of 10?
PCR Rxn Buffer (Invitrogen?, Tåstrup, Denmark), 1 lL of dNTPs
(1.25 mM/base, Roche?, Hvidovre, Denmark), 1.75 lL of MgCl2
(50 mM, SIGMA?, Brøndby, Denmark), 1 lL of each primer
(10 pmol/lL,TAG Copenhagen
0.2 lL of Platinum?Taq DNA Polymerase (5 U/lL, Invitrogen?,
Tåstrup, Denmark), in a total volume of 25 lL. Positive (DNA)
and negative (H2O and water from the washing step) controls were
included in all runs. For electrophoresis we used 5 lL of the 25 lL
PCR volume, and the remaining 20 lL were used for sequencing,
which was done at least uni-directionally (Eurofins MWG Operon,
3.1. Hypochlorite trials
We found that a solution of 1% HClO was sufficient to prevent
PCR amplification of E. vermicularis DNA, and that agarose gel
bands indicating positive PCR reactions re-appeared at a threshold
concentration of 10?4HClO. Also, subsequent lowering of the HClO
concentration did not result in PCR amplification, showing that
that failing amplification was due to DNA degradation and not
PCR inhibition by HClO (Table 2).
3.2. Samples from patients with unknown D. fragilis status
A total of 238 single eggs were collected from 64 patients, and
14/238 (6%) eggs tested positive for D. fragilis (Table 2). Of these,
11 specific amplicons were successfully sequenced and shown to
be identical to GenBank acc. no. AY730405 (D. fragilis genotype
1) (Johnson and Clark, 2000); one was identical to GenBank
U37461 (D. fragilis genotype 2) and in the remaining 2/14, the
genotype could not be called because of insufficient sequence qual-
ity. In total, 8/64 (13%) patients were found to harbour D. fragilis
DNA within pinworm eggs collected from their sample.
3.3. Sample from patient known to harbour D. fragilis
A total of 100 single eggs were collected and 39/100 (39%)
tested positive for D. fragilis by PCR. Of these, 39/39 (100%) had
specific amplicons successfully sequenced and shown to be identi-
cal to GenBank AY730405 (D. fragilis genotype 1). No E. vermicularis
or D. fragilis-specific products were produced by PCR on DNA ex-
tracted from washing water, and 99/100 of the single eggs tested
positive for E. vermicularis by PCR. We also used a D. fragilis real-
time PCR on a selection of eggs, showing a mean Ct of 35.6 (30,
3–40, 9) for D. fragilis positive eggs (data not shown), indicating
only few gene copies present within individual eggs.
The present study is, to our knowledge, the first to demonstrate
the presence of D. fragilis DNA inside eggs of E. vermicularis, allow-
ing us to presume that D. fragilis can be transmitted by a vector, in
this case via eggs of E. vermicularis.
The rationale behind this presumption is the repeated detection
of D. fragilis DNA from pinworm eggs, which had been surface-ster-
ilized prior to DNA extraction using a solution of hypochlorite in
concentrations shown to render DNA non-amplifiable due to
DNA damage. In addition, since water from the final washing steps
was PCR-negative for both E. vermicularis and D. fragilis, we con-
clude that D. fragilis DNA amplified from DNA extracts from the
surface-sterilised eggs must have been present inside the pinworm
eggs and not on the surface. It is possible that intact D. fragilis tro-
phozoites (or an undetected cyst stage) can stick to the surface of
an egg, as suggested by Menghi et al. (2005), who noted amoe-
boid-like structures on the surface of the eggs. However, we con-
sider this unlikely, given that D. fragilis normally degrades
rapidly (<48 h) once passed from the intestine, and since the mean
time from sampling to processing of the cellophane tape samples
was 36 days. Also, any organism adherent to the surface of the
egg would still be subjected to the damaging effect of the
Menghi et al. (2005) collected eggs by emptying the uteri of five
female pinworms, four of which had been collected from patients
with D. fragilis co-infection, and one from a patient without D. fra-
gilis co-infection. The authors divided the egg solutions, treating
one part with DNase to degrade DNA present on the surface and
left another solution untreated. The DNase treated eggs were rup-
tured using a combination of NaOH/2-mercaptoethanol incubation
and high-velocity centrifugation (10,000 rpm), and PCR was
performed on both solutions using E. vermicularis- and D. fragilis-
specific primers, showing amplification of both pinworm and D.
fragilis DNA from eggs untreated with DNAse, but only amplifica-
tion of pinworm DNA, once the eggs had been treated with DNase,
indicating that amplified D. fragilis DNA did not stem from within
the eggs (Menghi et al., 2005).
The fact that the authors’ detected D. fragilis DNA prior to the
DNase treatment of the eggs does not necessarily imply that the
worms investigated were responsible for transmission of D. fragilis,
as the detected DNA could have originated from D. fragilis-positive
faeces. The authors commented on the risk of contamination, but
dismissed it, as all worms had successively been washed with
water and stored in RPMI 1640 (a cell culture medium). However,
in the present study we found that washing with DNase/RNase free
water instead of HClO was not sufficient to remove DNA contami-
nants from the eggs, and have no reason to assume this would not
be the case for an adult worm. Moreover, in the present study we
detected D. fragilis DNA in only a fraction of the eggs (39%) from a
co-infected patient, and it could be hypothesised that the fraction
of eggs harbouring D. fragilis could vary, depending on the pin-
worm and/or D. fragilis genotype.
The findings of the present study also warrant a consideration
of possible DNA contamination. Faecal content could contaminate
the cellophane tape samples used; however, since we harvested
eggs directly into HClO in concentrations shown to damage DNA,
and since we subsequently transferred eggs individually in as little
D. Röser et al./Experimental Parasitology 133 (2013) 57–61
as 1 lL of fluid, we consider this mode of contamination unlikely. Download full-text
Contamination by ruptured/hatched egg contents is also a possibil-
ity, as our multiple-egg washing protocol employed several succes-
sive centrifugation steps of the egg solution. It could be speculated
that such a treatment could damage a portion of the eggs, allowing
DNA to contaminate the egg-solution. To address this, we per-
formed PCR on the water from the last washing step, which tested
negative for both pinworm and D. fragilis DNA. However, as was
the case for the high density sample, the wash water was removed
first, and the time-consuming process of picking up the eggs indi-
vidually was done afterwards, with the possibility of an untimely
hatching of an egg causing contamination of the egg-solution.
However, any such false-positive eggs would still represent DNA
originally found within a pinworm egg, while the reported fraction
of pinworm eggs harbouring D. fragilis in a co-infected patient
(39%) might be inflated.
Epidemiological studies reporting either higher than expected
co-infection of pinworm and D. fragilis, or no association at all,
should be interpreted with caution. Since such studies rarely adjust
for age, and given that both parasites occur more commonly in
children than adults, the observed co-infection can represent a sig-
nificant source of bias when not age-adjusted. Also, without a lon-
gitudinal setup, a difference in duration of infection becomes
important, as a shorter time to spontaneous remission of one of
the parasites could potentially mask an association. Not all Entero-
bius may harbour D. fragilis and even if they do, the present data
suggest that eggs from such worms may not all harbour D. fragilis.
While our findings strongly support the hypothesis of D. fragilis
transmission by a pinworm vector, many issues remain to be re-
solved regarding the lifecycle of D. fragilis. First, does the presence
of D. fragilis DNA represent viable D. fragilis that can be cultured
from the pinworm eggs?
If so, is Enterobius a ‘dead end’ or a paratenic (transport) host?
How does D. fragilis exit the eggs? Does D. fragilis require transmis-
sion by a vector to complete its life cycle and does D. fragilis under-
go development and multiplication in pinworms? Do other
nematodes harbour D. fragilis, and if so, could they serve as
Using microscopy, Sukanahaketu et al. (1977) reported the
presence D. fragilis trophozoites within eggs of Ascaris lumbricoides
(Sukanahaketu, 1977), but confirmation using molecular diagnos-
tics or electron microscopy remains to be demonstrated.
As stated by Barratt et al. (2011a), protozoa closely related to D.
fragilis have been reported to display both pseudocysts and cyst-
like structures (trichomonads) and direct transmission of tropho-
zoites remain at least a theoretical possibility (Barratt et al.,
2011b), though more research is needed.
Transmission of D. fragilis by pinworm would provide an entic-
ing explanation for the high prevalence of D. fragilis, particularly
among children, who also have the highest prevalence of pinworm.
While the present findings might warrant consideration on the use
of antimicrobial intervention and control measures related to D.
fragilis and E. vermicularis, the overall uncertainty of the proposed
pathogenicity in D. fragilis should give clinicians pause before new
strategies are implemented.
Future studies should aim to generate information on the viabil-
ity, longevity, potential development and multiplication of D. fragi-
lis in pinworm eggs, with an essential first step being the successful
culture of live D. fragilis trophozoites from a pinworm egg or suc-
cessful transmission of D. fragilis in an animal model using pin-
We thank Lis Lykke Wassmann and Mario Rodríguez Ferrero for
excellent laboratory assistance.
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