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Am. J. Trop. Med. Hyg., 00(0), 2019, pp. 1–9
doi:10.4269/ajtmh.18-0242
Copyright © 2019 by The American Society of Tropical Medicine and Hygiene

Estimating Human Exposure to Rat Lungworm (Angiostrongylus cantonensis) on Hawai’i Island:
A Pilot Study
Susan I. Jarvi,1* Praphathip Eamsobhana,2 Stefano Quarta,1 Kathleen Howe,1 Steven Jacquier,1 Alexandra Hanlon,3
Kirsten Snook,1 Robert McHugh,1 Zachariah Tman,4 Jill Miyamura,5 Kuilei Kramer,1 and McKayla Meyers1
1
Department of Pharmaceutical Sciences, Daniel K. Inouye College of Pharmacy, University of Hawai’i at Hilo, Hilo, Hawaii; 2Department of
Parasitology, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand; 3Department of Statistics, Center for Biostatistics and
Health Data Science, Virginia Tech, Roanoke, Virginia; 4School of Public Health, University of California, Berkeley, California; 5Hawaii Health
Information Corporation, Kailua, Hawaii

Abstract. Angiostrongylus cantonensis is a zoonotic, parasitic nematode causing angiostrongyliasis or rat lungworm
disease. Clinical diagnosis in humans is currently conﬁrmed by detection of parasite DNA in cerebrospinal ﬂuid. This study
estimated human exposure to A. cantonensis in volunteer participants solicitated via public venues on east Hawai’i Island
using blood-based tests. Antibodies were screened in sera by crude antigen ELISA, followed by a 31-kDa dot-blot test
developed and validated in Thailand. Human participants (n = 435) donated blood samples and completed a questionnaire
to self-report relevant symptomology or clinical diagnosis. Among symptoms reported by participants diagnosed by
licensed clinicians, headaches, high eosinophil counts, stiff neck, fatigue, and joint pain were most severe during the initial
3 months of infection. ELISA results revealed 22% of the serum samples as positive, 46% as equivocal, and 32% as
negative. A subset of 186 samples was tested by dot blot, with 30% testing positive and 70% testing negative. A
signiﬁcantly higher mean ELISA value was found among recently (2014–2015) clinically diagnosed participants as than
among those with a diagnosis before 2010 (P = 0.027). All dot-blot positives were also ELISA positive and were signiﬁcantly associated with higher ELISA values compared with dot-blot negatives (P = 0.0001). These results suggest that an
ELISA using crude antigen isolated from adult A. cantonensis from Hawai’i may be an effective initial screening method for
estimating exposure to A. cantonensis in Hawai’i and likewise suggest that dot-blot tests using the 31-kDa antigen exhibit
efﬁcacy as a diagnostic for exposure.

INTRODUCTION

reports of adult worms found in the hearts and lungs of humans on autopsy, demonstrating that larvae are able to exit
the CNS in humans.7–11
Deﬁnitive diagnosis of angiostrongyliasis is made by identifying A. cantonensis nematodes in the cerebrospinal ﬂuid
(CSF) of patients or by detection of parasitic DNA in the CSF12;
both require lumbar puncture. However, diagnosis can also be
made by serological detection of A. cantonensis antigens or
speciﬁc antibodies. Glycoproteins of approximately 31 kDa
isolated from adult A. cantonensis worms have been shown to
be diagnostic for human infection in Thailand with high sensitivity and speciﬁcity.13
Eamsobhana et al.14 found an ELISA using a 31-kDa antigen
to be 100% sensitive and 100% speciﬁc in sera from human
A. cantonensis infections. This ﬁnding was supported by a
separate study,15 where nine diagnostic laboratories in Thailand were each provided with an identical 31-kDa ELISA kit to
screen sera from a range of human parasitic infections, including angiostrongyliasis, gnathostomiasis, and ﬁlariasis. In
all nine independent laboratories, the 31-kDa kit was shown to
have 100% sensitivity and 100% speciﬁcity.
A lateral ﬂow assay using the 31-kDa antigen was used to
test 97 serum samples from clinically diagnosed subjects with
parasitic infections and from 35 negative controls.16 Although
one of four of the serum samples from hookworm-infected
subjects was positive, this subject was later shown to have
had a concomitant or previous infection with A. cantonensis.16
This assay for A. cantonensis showed 100% sensitivity,
98.72% speciﬁcity, 95% positive predictive value, and 100%
negative predictive value.16 With crude antigen, crossreactivity was observed in sera from cases of gnathostomiasis, toxocariasis, ﬁlariasis, and paragonimiasis, but results
could be useful for initial screening in regions where nontarget
parasite prevalence is low.17

The nematode Angiostrongylus cantonensis is a rat lungworm, a zoonotic pathogen that in Hawai’i, as in many other
tropical locations, is the main cause of human eosinophilic
meningitis and ocular angiostrongyliasis,1 known colloquially
as rat lungworm disease. Initially discovered in China in 1935,2
the nematode is found in at least 30 countries, including many
Paciﬁc islands and the continental United States (at the time
of this writing, Louisiana, Texas, Florida, Tennessee, and
Oklahoma).3,4 The deﬁnitive host is the rat, which acquires the
third-stage (L3) larvae by eating an infected mollusk host; after
being ingested by the rat, the L3 larvae migrate to the central
nervous system (CNS), where they become subadults (L5).
From the CNS, the nematodes move to the pulmonary arteries
where they mature sexually, mate, and lay eggs. First-stage
larvae (L1) hatch from eggs and migrate up the bronchial tree,
are swallowed, and pass through the host’s digestive system
and feces. The cycle in the rat, from initial infection with L3 to
excretion of L1, takes approximately 6 to 8 weeks.5 Slugs or
snails acquire L1 larvae by ingesting rat feces, and the parasite
develops from the L1 to the L3 larval stage in these intermediate mollusk hosts. Humans are accidental hosts and
become infected by ingesting L3 larvae via intermediate or
paratenic hosts on contaminated food or in water or other
liquid. The L3 larvae, once ingested, have historically been
presumed to then migrate to the CNS or to the eye chamber (in
the case of ocular angiostrongyliasis), where the parasite remains until its death.6 There have, however, been a number of

* Address correspondence to Susan I. Jarvi, Department of Pharmaceutical Sciences, Daniel K. Inouye College of Pharmacy, University of
Hawai’i at Hilo, 34 Rainbow Dr., Hilo, HI 96720. E-mail: jarvi@
hawaii.edu

1

 2

JARVI AND OTHERS

The 31-kDa–based test was developed and validated in
Thailand.14–17 Genetic studies show little difference between
Thai and Hawaiian strains of A. cantonensis (Nei’s genetic
distance estimated at 0.03).18 We therefore expect the Thailand 31-kDa–based test to effectively detect antibodies speciﬁc for A. cantonensis in our Hawai’i study population,
thereby serving as an indicator of human exposure to this
parasite. To test these hypotheses, we initially screened serum samples with an ELISA using crude antigen from
A. cantonensis collected in Hawai’i. We then performed dot
blots on a selective subset of samples using the 31-kDa proteins isolated from Thailand A. cantonensis as antigen and
tested a further subset by real-time polymerase chain reaction
(PCR) for parasite DNA in the blood. Participants in this study
completed a questionnaire to correlate symptomology with
test results.
MATERIALS AND METHODS
Study design. This study was conducted using convenience sampling coupled with self-reported responses to a
questionnaire. Volunteer participants were solicited via radio
ads and notices posted in local newspapers and public areas
on the eastside of Hawai’i Island. Participants completed
consent forms and questionnaires with assistance from study
personnel and licensed staff of the Puna Community Medical
Center, Pahoa, Hawai’i. No preference for participants of
known A. cantonensis infection status was given.
Ethical considerations. This study was reviewed and approved by the University of Hawai’i Institutional Review Board.
Written, informed consent was obtained from adult participants and guardians of participants aged 17 years and
younger by study staff.
Sample collection. Blood samples were collected as per
standard protocol by the licensed staff of Clinical Laboratories
of Hawai’i from July to December 2015. A total of 435 blood
samples were collected from human volunteer participants
between June 11 and September 16, 2015. Serum samples
were collected in serum separator tubes (SSTs) and blood was
collected in heparinized collection vials for plasma isolation.
Serum separator tubes were centrifuged at 1,889 × g for 10
minutes and stored at 4°C until transport to the research laboratory. Whole blood in heparinized tubes was stored at 4°C
until transport. All samples were transported in an insulated
cooler with ice packs. All samples were de-identiﬁed and labeled with numerical coding sans personal data.
Sample processing. A sample of 100 μL heparinized whole
blood was transferred to a tube containing 100 μL DNA lysis
buffer (0.1 M Tris–HCl, 0.1 M EDTA, 2% SDS), gently mixed by
manual inversion, and stored at −80°C until DNA extraction
was performed. Heparinized blood samples were centrifuged
for 5 minutes at 3,000 × g. Plasma was transferred to sterile
screw-capped tubes and stored at −80°C until testing. Sera
from samples collected in SSTs were transferred to individual
sterile screw-capped aliquot tubes and stored at −80°C until
testing.
Controls. Among the participants were 15 individuals who
reported recent clinical diagnosis of angiostrongyliasis by
PCR of CSF. Sera from these individuals were used for positive controls. Sera from individuals newly arrived in Hawai’i
from the U.S. mainland who reported no exposure to
A. cantonensis were used for negative controls.

Preparation of crude antigen. The Hawai’i strain of
A. cantonensis was used to prepare crude antigen. Adult worms
were collected in the course of previous animal studies. At the
end of the studies, rats were humanely euthanized and adult
A. cantonensis were dissected from the hearts and lungs. The
live nematodes were rinsed three times in 0.01 M phosphate
buffer saline (PBS) (Life Technologies, Grand Island, NY) containing 1 × protease inhibitor (Biochem Cocktail Set V EDTAfree; Thermo Scientiﬁc). Rinsed, clean nematodes were then
placed in 200 μL PBS/protease inhibitor and stored at −80°C.
Crude A. cantonensis antigen preparation was completed with
some modiﬁcation as described in Eamsobhana et al.14 Brieﬂy,
10 male and 10 female adult A. cantonensis worms in PBS/
protease inhibitor were exposed to three freeze/thaw cycles
and homogenized with 400 μL PBS/protease inhibitor. This
preparation was sonicated (Microson Ultrasonic Disrupter,
Misonix, Inc., Farmingdale, NY) on ice four times using 3second pulses and stored overnight at 4°C. Crude antigen was
centrifuged at 2,400 × g for 15 minutes, and the supernatant
was collected. A Bradford assay kit (Bio-Rad Labs) was used to
quantify proteins recovered.
Indirect ELISA. ELISA was based on the methods described in Eamsobhana et al.19 with the following modiﬁcations: optimal concentration of antigen and serum (positive
control) was determined by checkerboard titration. A concentration of 1:200 for primary antibody was derived by titration of positive and negative serum controls with a range of 1:
100 to 1:1,000. Serum and plasma were tested, and no differences in reactivity were noted; serum was used for the remainder of the study. Each plate included two positive
controls, two negative controls, and multiple carbonate buffer
controls (without antigen). Crude A. cantonensis antigen was
diluted with 0.05 M carbonate buffer pH 9.6 to a ﬁnal concentration of 0.5 μg/mL. All washing steps were conducted on
a 405 Select TS microplate washer (BioTek). After sensitization, plates were washed four times with 300 μL PBS–0.05%
Tween 20 (PBS-T) (pH 7.4) with a 2-minute pause after every
other wash. Plates were then blocked with 125 μL 5% bovine
serum albumin PBS-Tween (PBS-T) for at least 2 hours at
room temperature with gentle shaking. Blocking solution was
removed, and plates were washed twice with PBS-T with no
pause in between. Serum (100 μL) diluted to 1:200 with PBS
was added to appropriate wells and incubated for 2 hours at
37°C with shaking. All samples were run in triplicate. After
incubation with primary antibody, plates were washed six
times with 300 μL PBS-T, with a 2-minute pause after every
other wash. A 1:1,000 dilution of horseradish peroxidase
(HRP)-conjugated goat antihuman IgG Fc (Immunology Consultant Laboratories) (100 μL) was added, and plates were
incubated for 1 hour at 37°C with gentle shaking. Plates were
then washed six times with PBS-T, pausing every other wash.
TMB-solubilized substrate solution (TMBOne®; Promega) for
HRP was added according to the manufacturer’s protocols.
Reaction was stopped with 1 N HCl after 15 seconds. Quantiﬁcation was determined on an iMark microplate absorbance
reader (BioRad) at 450 nm.
Dot-blot ELISA. Nitrocellulose membrane blots with 0.2 μg
of 31 kDa isolate of A. cantonensis were provided by P. E.,
Mahidol University, Bangkok, Thailand. Procedures were followed as previously described.17 Brieﬂy, each membrane was
placed in a sterile 12-well cell culture plate well with lid
(CytoOne) and immersed in 1 mL blocking solution (5% nonfat

 HUMAN EXPOSURE TO ANGIOSTRONGYLUS CANTONENSIS IN HAWAI’I

dry milk in PBS-T) with gentle shaking for 1 hour at room
temperature. Membranes were washed three times with PBST and shaken gently for 5 minutes. Membranes were then
incubated for 1 hour at 37°C with serum samples diluted 1:200
in 1 mL 1% nonfat dry milk in PBS and washed as described
previously. Washed membranes were immersed in HRPconjugated goat antihuman IgG Fc diluted 1:1,000 with 1%
nonfat dry milk in PBS and incubated for 1 hour at 37°C.
Membranes were again washed as earlier and blotted dry in
between washes. Blots were then transferred to new trays and
incubated at room temperature with 500 μL chloronapthol
substrate (ThermoScientiﬁc) with gentle shaking until color
appeared (∼30 minutes). Reactions were stopped by washing
twice with dH2O. Blots were dried and scanned (V500 Photo;
Epson) at 1,200 dpi.
DNA extraction. DNA extractions were performed on 58
dot-blot–positive samples using the Qiagen DNeasy Animal
Blood and Tissue Kit for nonnucleated blood with slight
modiﬁcations from the manufacturer’s suggested protocol.
Whole blood in lysis buffer (0.1 M Tris-HCl pH 8.0, 0.1 M sodium EDTA, 2% SDS) was thawed completely, and 200 μL was
combined with 20 μL proteinase K followed by overnight digestion at 55°C with gentle shaking. Extractions were otherwise completed according to the standard protocol.
Polymerase chain reaction. Samples were subjected to
PCR on a StepOnePlus RealTime PCR system (Life Technologies) using a Custom TaqMan Gene Expression Assay (Life
Technologies, assay ID A139RIC) as described.20 Cycling
conditions, primers (AcanITS1F1 and AcanITS1R1), and
probe (AcanITS1P1) were those described by Qvarnstrom
et al.21 Polymerase chain reactions were carried out in 20 μL
total volume and included 0.25 μM probe, 0.9 μM forward and
reverse primers, and 1X TaqMan Environmental Master Mix
2.0 (Life Technologies). A positive plasmid control and a
negative control containing no DNA were included in all reactions. All PCR reactions were replicated for veriﬁcation.
Indirect ELISA data analysis. Positive controls were chosen by selecting the samples with the highest mean absorbance
from recently clinically diagnosed participants. Negative control
samples originated from two newly arrived volunteers from
Nevada who had never been to the Hawaiian Islands or elsewhere in the tropics and whose sera showed consistently low
absorbance values. ELISA value (%EV) was created to compare
absorbance values as a percentage of positive and negative
controls according to the following formula: %EV = (Sample
Mean Absorbance − Negative Control Mean Absorbance)/
(Positive Control Mean Absorbance − Negative Control Mean
Absorbance) × 100. A two-sample t-test was performed in
Minitab 18 to compare mean %EV between more recent clinically diagnosed participants (2014–2015) and those clinically
diagnosed earlier (before 2010).
The consistency of ELISA absorbance data was evaluated
by measuring coefﬁcient of variability (%CV).22 An inter-assay
%CV of the mean absorbance of each sample in triplicate was
set to 15% and an intra-assay %CV of the mean absorbance
of all samples on the microplate was set to 10%. A %CV value
greater than the respective threshold was not considered
consistent and that data were discarded. Inter-assay %CV
was calculated as follows: (SD of plate means/Mean of Plate
Means) × 100. Intra-assay %CV was calculated as follows:
(SD of triplicate absorbance values/Mean of absorbance values in triplicate) × 100.

3

Dot-blot data analysis. A positive result was determined by
dot blot if a deﬁned round circle could be seen by the naked
eye in the photos taken immediately after enzymatic development. To be considered positive, circles had to appear at
the vicinity of the small prick created during the absorption of
the 31-kDa antigen and be of a size and density consistent
with positive control results. If a dot was not seen on a blot, if
an irregular shaped splotch was seen on the blot, or if the
deﬁned dot was only seen under altered contrast of the photo,
the sample was considered negative for rat lungworm exposure. Negative results were numerically assigned a value of 0.
If the dot was not visible by eye but appeared faint by enhancement, then it was assigned a value of 1. Weak positives
were assigned a value of 2; these were seen by the naked eye
without enhancement but were clearly not as dark as strong
positives. Strong positives were assigned a value of three and
were easily observed as a dark circle. For data analysis, we
conservatively considered zero and one to be negative,
whereas two and three were considered positive. To examine
differences in ELISA values between participants with positive
versus negative dot-blot test results, a nonparametric Wilcoxon rank-sum test was performed in R.23
Questionnaire data analysis. Using results from the survey
questionnaire (Supplemental File 1), ELISAs, and dot blots,
two multivariable logistic regression models were generated
to determine whether demographic (residence, type of water
supply, gender, age, education level, and ethnicity) and food
preparation (see Question 3 A-I in Supplemental File 1) variables were signiﬁcantly associated with positive versus negative ELISA and dot-blot test results. Separate models were
performed for ELISA and dot-blot test results.
Repeated measures of self-reported symptoms (nausea,
headache, meningitis, high eosinophil counts, stiff neck, skin
sensitivity, muscle soreness, fatigue, joint pain, fever, mental
clarity, memory, speech, sexual dysfunction, vision, balance,
bladder dysfunction, bowel dysfunction, and others) and their
level of severity (mildest to most severe, with higher scores
indicating more severity) were collected for each participant reporting clinical diagnosis early (< 3 months) and late
(> 3 months) in infection. Three months was selected as a reasonable time post-symptom onset to distinguish early versus
late, as Hawai’i Department of Health (HDOH) states that
“symptoms usually last between 2 and 8 weeks; symptoms
have been reported to last for longer periods of time” https://
health.hawaii.gov/docd/disease_listing/rat-lungwormangiostrongyliasis/#signs_symptoms. Normality of the severity score for each symptom was evaluated using Shapiro-Wilk
tests performed in SPSS (version 25.0).24 Because symptom
severity scores collected early and late in infection were not
normally distributed, nonparametric Wilcoxon signed-rank
tests were conducted to examine whether median change in
each symptom’s severity was signiﬁcantly different from zero.
In addition, boxplots were generated to visually assess
changes in symptom severity within and after 3 months of
infection. Statistical signiﬁcance was taken at the P < 0.05
level.
RESULTS
Demographics. A total of 435 volunteers participated in
this study, providing blood samples and responding to a
questionnaire. Study location and participants’ reported

 4

JARVI AND OTHERS

places of residence are provided in Figure 1. The majority (272/
435, 62.5%) are residents of Pahoa, HI, where sample collection took place. Eastern Hawai’i Island is rural, and most
participants (83%) reported living in a rural environment versus city or suburban environment. There is unregulated,
widespread use of rainwater catchment in east Hawai’i Island;
nearly 78% of participants reported the use of rainwater
catchment as a source of household water, as compared with
21% reporting a municipal water source. Most participants
were female (56%), Caucasian (79%), and reported some level
of college education (62%), with a mean age of 56 years
(range: 6–93 years).
Self-reported diagnosis. The participants were asked
whether they believed they had ever had angiostrongyliasis
and, if so, whether diagnosis was made by a licensed clinician. A total of 105 (24%) reported angiostrongyliasis (selfdiagnosed), 315 (72%) reported no known exposure to
A. cantonensis, and 15 (3.4%) reported clinical diagnosis. Of
the 105 who reported having had angiostrongyliasis, 81 reported the date of onset of symptoms (Figure 2). More participants reported symptoms acquired after 2013; this is
similar to incidence data reported by the HDOH and Hawaii
Health Information Corporation (HHIC) (Figure 2). Selfreported demographics as well as ELISA (%EV) and dot-blot
results of the 15 clinically diagnosed participants are provided
in Table 1. Reported dates of clinical diagnosis ranged from
1981 to 2015.
Self-reported symptoms. Frequency and severity of
common symptoms and presentations reported by the 15
clinically diagnosed patients are summarized, and earlier
stages of infection (< 3 months post-symptom onset) are
compared with those reported during later infection (> 3
months post-symptom onset) (Figure 3). Severity range was
ranked from 1 to 5, with 5 being the most severe. The

FIGURE 1. Study location on East Hawaii Island and participants’
reported place of residence. This ﬁgure appears in color at
www.ajtmh.org.

frequency and severity of symptoms reported earlier versus
later in infection appeared to decrease over time among the 15
clinically diagnosed participants; however, some participants
reported severe symptoms persisting after 3 months, for example, skin sensitivity. Figure 3 displays boxplots of symptoms that demonstrated statistically signiﬁcantly different
levels of severity earlier versus later in infection. Speciﬁcally,
nonparametric Wilcoxon signed-rank tests showed that median symptom severity was signiﬁcantly greater for headaches
(P = 0.0232), high eosinophil counts (P < 0.05), stiff neck (P <
0.05), fatigue (P < 0.05), and joint pain (P < 0.05) during the
initial 3 months of infection than reported severity of these
symptoms later than 3 months. No statistically signiﬁcant
differences were observed for the other symptoms.
ELISA. Crude antigen was isolated from 10 male and 10
female adult A. cantonensis for use in ELISAs, yielding a
protein concentration of 1.343 μg/μL. All serum samples
were tested by ELISA for the presence of antibodies to
A. cantonensis. ELISA tests were run in triplicate and data
included where the SD among replicates was < 0.025. ELISA
cutoff values were determined by the %EV of the samples
from clinically diagnosed participants. The lowest %EV seen
in samples from participants reporting recent clinical diagnoses (2014–2015) by spinal tap at Hilo Medical Center
consistently showed a %EV at or greater than 70%, and this
was chosen as the lowest threshold for classifying the sample
“positive” by ELISA for exposure to A. cantonensis. The lowest
%EV of the samples from participants reporting clinical diagnosis before 2014 by spinal tap at Hilo Medical Center
(2009) was 48, with the exception of one outlier, clinically diagnosed in 1981 whose sample showed a %EV of 17. Thus, 48
%EV served as a cutoff range for testing by dot blot, and
samples at or greater than this threshold were considered
“likely positive.” The highest %EV within negative control
samples was 16; therefore, 16 was established as the upper
end of a negative result. A %EV of 16.1–48 was considered
equivocal. For simpliﬁcation, negative samples (%EV < 16)
were numerically assigned a value of 0, equivocal (16.1–48 %
EV) assigned 1, likely positive (48.1–70 %EV) assigned 2, and
positive (> 70 %EV) assigned 3 in our datasets.
ELISA results are shown in Table 2. Overall, 28 samples
(6%) were positive, 69 (16%) were likely positive, 198 (46%)
were equivocal, and 140 (32%) were negative. Combined, 97
(22%) samples tested positive or likely positive. Of the samples from the 15 individuals who had reported clinical diagnosis of angiostrongyliasis, four (27%) tested positive, six
(40%) tested likely positive by ELISA, and ﬁve tested equivocal. None of the samples from clinically diagnosed participants returned results within our negative range. Of the 15
participants reporting clinical diagnosis, seven reported diagnoses by spinal tap. Six of these tested positive or likely
positive and one tested equivocal. Of the 105 volunteers who
reported infection with A. cantonensis by self-diagnosis, nine
(9%) tested positive, 15 (14%) likely positive, 46 (44%)
equivocal, and 35 (33%) tested negative. Of the 315 participants reporting no exposure to A. cantonensis, 15 (5%) tested
positive, 48 (15%) likely positive, 147 (47%) equivocal, and
105 (33%) negative.
Dot blot. A subset of 186 samples was tested with the 31kDa dot blot, including 28 samples from the ELISA-positive
pool of samples, 64 samples from the likely positive group, 71
from the equivocal group, and 23 from the group testing

 5

HUMAN EXPOSURE TO ANGIOSTRONGYLUS CANTONENSIS IN HAWAI’I

FIGURE 2. Self-reported numbers and year of infection with A. cantonensis as compared with numbers reported by the Hawai’i Department of
Health and Hawaii Health Information Corporation.

negative. Results are presented in Table 2. Overall, 56 (30%)
dot blots were positive and 130 (70%) were negative. Within
the 15 samples from clinically diagnosed participants, seven
(47%) were positive and eight (53%) were negative by dot blot.
Of the seven participants diagnosed by spinal tap, four samples (57%) were positive and three (43%) were negative. Of the
105 participants who self-diagnosed angiostrongyliasis, 45
were tested by dot blot, with 13 (29%) testing positive and 32
(71%) negative. Of the 315 volunteers reporting no exposure
or disease, 126 were tested by dot blot, with 36 (29%) testing
positive and 90 (71%) negative.
Dot-blot and ELISA results are compared in Table 3. Of
samples testing “positive” by ELISA, 19 (68%) tested positive

and nine (32%) tested negative by dot blot. Of those that were
“likely positive” by ELISA, 24 (38%) tested positive and 40
(62%) tested negative by dot blot. Of those “equivocal” by
ELISA, 13 (18%) tested positive and 58 (82%) tested negative
by dot blot. Of those “negative” by ELISA, zero were positive
and 23 (100%) were negative by dot blot (Table 3).
A nonparametric Wilcoxon rank-sum test demonstrated
statistically signiﬁcant differences in ELISA values between
participants with positive versus negative dot-blot test results.
Speciﬁcally, participants testing positive by the 31-kDa dot
blot had higher median ELISA values than those testing negative (P < 0.0001) (Table 3). Of the 186 participants whose
samples were tested by dot blot, 152 (82%) reported using

TABLE 1
Self-reported demographics of clinically diagnosed participants
Residence

Pahoa
Pahoa
Pahoa
Pahoa
Hilo
Pahoa
Pahoa
Pahoa
Keaau
Hilo
Hilo
Hilo
Pahoa
Pahoa
Pahoa

Gender

Age

When infected

Where infected

F
M
F
M
M
M
F
M
F
F
M
F
M
M
F

68
52
63
67
30
70
12
73
62
39
39
55
23
73
59

1981
2004
2005
2007
2008
2009
2009
2009
2010
2014
2014
2014
2015
2015
2015

Honokaa or Hilo
Kapoho (Puna)
Kapoho (Puna)
Pahoa (Puna)
Kapoho (Puna)
Papaya Farms Rd (Puna)
Puna, HI
Pahoa (Puna)
Hawaii
Hilo
Hilo
Keaau (Puna)
Puna
Puna
Puna

Diagnosis procedure

Spinal tap
Spinal tap
Antibody test
Eosinophil test
Spinal tap
Other
Spinal Tap
Symptoms
Don’t Know
Spinal Tap/CT
Spinal tap
Ab test
Spinal tap
Antibody test
Symptoms and response to
treatment

Hospital
admission

%EV

Dot blot

HMC
QMC
NR
NR
HMC
Thailand
HMC
NR
NR
HMC
HMC
NR
HMC
HMC
NR

18
58
27
60
53
69
48
16
38
100
70
32
111
87
69

P
N
N
P
N
P
N
N
N
P
P
N
P
P
N

HMC = Hilo Medical Center (Hilo); QMC = Queen’s Medical Center (Honolulu); all were diagnosed by MD or ND. NR = no response. The %EV is the ELISA value, and dot blot is listed as positive (P) or
negative (N).

 6

JARVI AND OTHERS

FIGURE 3. A boxplot analysis displaying mean severity of symptoms in 15 clinically diagnosed participants found to be signiﬁcantly different
during and after 3 months of being infected with A. cantonensis. These clinically diagnosed symptoms had higher severity during 3 months of
infection than 3 months post-infection. The lines are the severity spectrum, the bars represent most of the patients’ responses, and the cross
between bars represents the medians. *Extreme outliers. This ﬁgure appears in color at www.ajtmh.org.

rainwater catchment as a source of household water and 33
(18%) reported having a municipal water source. Because of
the large difference in sample sizes, the water source data
were not further analyzed.
ELISA and dot-blot results from clinically and self-diagnosed
positive participants from 2001 to 2015 were summarized (data
not shown). Among participants reporting clinical diagnosis, the
earliest reported year of angiostrongyliasis onset our ELISA
detected was 2004. By dot blot, the earliest reported year of
onset detected was 2007, again with the exception of one
participant reporting diagnosis in 1981 (Table 1). A two-sample
t-test demonstrated that participants with recent (2014–2015)
infections had a signiﬁcantly higher mean %EV than those
with onset of disease before 2010 (mean %EV: 78.2 versus 43,
P = 0.027).
Regression analyses. Separate multivariate logistic regression models were estimated to examine whether demographic and food preparation habit variables were
associated with A. cantonensis exposure status as measured
by ELISA and dot-blot tests. None of these variables was
found to be signiﬁcantly associated with a “positive” ELISA
or dot-blot test at the P < 0.05 level.
Polymerase chain reaction. We extracted DNA from 58
samples, including 55 positives by dot blot, and tested for the

presence of A. cantonensis DNA in the blood by real-time
PCR. Reliable signal was not detected in any samples except
the positive controls. Negative controls remained negative
throughout (data not shown).
DISCUSSION
This study represents the ﬁrst attempt in the United States
to estimate human exposure to A. cantonensis and to describe early symptoms (< 3 months onset) and later symptoms
(> 3 months onset) of angiostrongyliasis. Initial screening of
435 serum samples from participants by Hawai’i-based crude
antigen ELISA revealed a “positive” rate of 97/435 (22%).
Using a previously validated 31-kDa dot-blot test developed in
Thailand on samples that were previously screened by ELISA,
including all positive samples and subsets of the equivocal
and negative samples, we found 56/186 tested positive (30%).
These results suggest the number of humans exposed to
A. cantonensis may be greater than the number of cases
previously reported by the HDOH.
Thailand has reported approximately 47% of all angiostrongyliasis cases worldwide.1 However, the number of cases
dropped from 1,386 cases of eosinophilic meningitis in 2000
to 176 cases in 2009.18 Many of the cases in Thailand, as in

TABLE 2
Results of ELISA testing on 435 serum samples and dot-blot data on 186 serum samples
ELISA
Positive
Likely positive
Equivocal
Negative
Dot blot
Positive
Negative

n = 435
6% (28)
16% (69)
46% (198)
32% (140)
n = 186
30% (56)
70% (130)

Clinically diagnosed

Self-diagnosed as positive

Self-diagnosed as negative

n = 15
27% (4)
40% (6)
33% (5)
0
n = 15
47% (7)
53% (8)

n = 105
9% (9)
14% (15)
44% (46)
33% (35)
n = 45
29% (13)
71% (32)

n = 315
5% (15)
15% (48)
47% (147)
33% (105)
n = 126
29% (36)
71% (90)

Percent rounded to the nearest signiﬁcant number. Diagnosis category is based on self-reported questionnaire responses.

 7

HUMAN EXPOSURE TO ANGIOSTRONGYLUS CANTONENSIS IN HAWAI’I

TABLE 3
Comparison of dot-blot results with ELISA results categorized by %EV (ELISA value)
ELISA results

Dot-blot positive
Dot-blot negative
Total tested by dot blot

Positive, ³ 70 %EV

Likely positive, 48.1–70 %EV

Equivocal, 16.1–48 EV

Negative, £ 16 %EV

19 (68%)
9 (32%)
28

24 (38%)
40 (62%)
64

13 (18%)
58 (82%)
71

0 (0%)
23 (100%)
23

Percent rounded to the nearest signiﬁcant number.

China, are due to intentional ingestion of undercooked
mollusks.3,18 Studies from China showed seropositivity to
A. cantonensis in 20% of 459 subjects on Hainan Island,25
21% of 393 in another study,26 and 14% of 300 in Guangdong
Province.27 Intentional ingestion of mollusks is rare in
Hawai’i28,29; most of our subjects could not identify their infection source.
The semi-slug Parmarion martensi is a recent immigrant to
the Hawaiian Islands30 and is likely a major contributor to the
increase in angiostrongyliasis cases acquired on the island of
Hawai’i in the past 15 years.31 Many cases have originated in
the Puna district of Hawai’i Island, where P. martensi are numerous and heavily infected (> 75%) with A. cantonensis larvae.31 In a survey of wild rats in the Puna district, 54% (20/37)
had adult rat lungworms visible on necropsy and 100% of rats
lungs tested positive for A. cantonensis by real-time PCR.32
Rats captured in the Hilo district of Hawai’i Island were found
to be 94% (512/545) positive for A. cantonensis based on
ﬁndings of adult worms, encysted adult worms, L3 larvae, and/
or real-time PCR analysis of brain tissue.33 The introduction of
P. martensi combined with the high infection rate in rats may
explain the increase in human cases of angiostrongyliasis and
the high rates of human exposure in East Hawai’i Island.
Notably, East Hawai’i also has widespread use of unregulated rainwater catchment for residential as well as agricultural purposes. Our questionnaire demonstrated that some
people, including ﬁve of the participants reporting clinical diagnosis in this study, believe they were infected via contaminated rainwater in catchment systems. Water transmission of
A. cantonensis has been previously demonstrated in rats,34–36
and it has been suggested that infections in man can be acquired from the consumption of A. cantonensis L3 in water.37
Parmarion martensi is a robust climber and has routinely been
found, drowned, inside catchment tanks. Infective larvae
leave drowned hosts and can survive in water for several
weeks.38 The potential exists for water transmission if effective ﬁltration and treatment are not implemented. For example,
contaminated water may be consumed directly from the tap,
used for rinsing food, or during the course of performing
personal hygiene functions such as showering and brushing
teeth. The widespread use of rainwater catchment for
household water supply may further contribute to the high
exposure rate of humans to A. cantonensis on East Hawai’i
Island and other regions with rainwater catchment use.
Severity of presentation in human cases of angiostrongyliasis ranges from mild and ﬂu-like illness to coma, longterm disability, and death.1 In the United States, symptoms
such as headache, paresthesia, hyperesthesia, and numbness have been reported to last up to 10 months or longer.29
Among our participants, 24% believed that they had been
infected with A. cantonensis and 72% believed that they
had never been infected. Because these are self-reported

data, there is some question as to the reliability of responses
relating to self-diagnosis. However, self-reported data can be
valuable,28,39 especially when information is otherwise unavailable. The prevalence of angiostrongyliasis in participants
who self-diagnosed (Figure 2) shows a recent upward trend
similar to the number of clinical cases reported by the HDOH
and HHIC40; this correlation provides support for the reliability
of the self-reported data.
Participants self-reporting no disease had ELISA and dotblot results comparable to those reporting having contracted
the disease. This suggests that subclinical infections may be
common with mild symptoms that went undiagnosed, were
absent, or were misattributed to the ﬂu or other ailments. A
low-level exposure to A. cantonensis (e.g., potentially through
environmental exposure such as to contaminated water from
catchment)38 over time may produce antibodies as seen in
participants reporting no disease. It has been shown in rats
that low exposure levels do provide immunological protection
from lethal challenge with A. cantonensis.41
Polymerase chain reaction was used to attempt to detect
A. cantonensis DNA in the blood of the 55 dot-blot–positive
blood samples.20,21 No ampliﬁcation was detected in any of the
test samples, except for the positive controls. We have previously detected A. cantonensis DNA in peripheral rat blood
under controlled infection at several time points using the same
methods,42 and we now routinely detect A. cantonensis DNA in
canine blood samples provided by local veterinarians (Unpublished data). In our human samples, the parasite had likely
cleared from the peripheral blood; sample collection for most
participants was at least 1 year after reported symptoms or
diagnosis. It would be useful to study blood (and CSF) PCR
changes over time in human angiostrongyliasis.
Of our 435 participants, 15 (3.4%) reported clinical diagnoses
of angiostrongyliasis. Seven of these individuals reported diagnoses by spinal tap and provided contact information of the
medical center that conﬁrmed the diagnosis. Data from clinically diagnosed infections may provide further information on
the duration of the antibody response as well as the impact of
longer term symptoms. The frequency and severity of many of
the symptoms in participants reporting clinical diagnoses decreased after 3 months. Some symptoms were reported as
severe earlier in infection (< 3 months.) but were signiﬁcantly
reduced later (> 3 months). These included headache, high
eosinophil count, stiff neck, fatigue, and joint pain. However,
some participants reported several symptoms persisted at
severe levels (e.g., skin sensitivity and memory issues). A better
understanding is required of the relationships among detection,
treatment, and long-term sequelae of this disease.
In comparisons of ELISA and dot-blot results from the
participants reporting clinical diagnoses, higher ELISA values
generally correlated with positive dot-blot results, with the
exception of the person who was ﬁrst infected in 1981 who

 8

JARVI AND OTHERS

had a low %EV and a positive dot blot. The %EV was signiﬁcantly higher in participants more recently diagnosed, which
was expected given that antibody titer generally decreases as
infection senesces.43 We detected antibody by dot blot in
participants reporting onset of disease and clinical diagnosis
in 1981, 2007, and 2009. It is not clear whether antibodies
detected were to the initial exposure in these time periods or to
a more recent reexposure. All (100% or 23/23) of the samples
from dot-blot negative participants were also ELISA negative,
whereas 53% (49/92) of the samples from ELISA-positive
participants were dot-blot negative. This suggests that, as
seen with crude antigen from Thailand,17 crude antigen from
Hawai’i A. cantonensis in ELISA exhibits some level of crossreactivity. There is no true gold standard for comparison within
this dataset, but if we were to consider our dot-blot results with
the validated 31-kDa Thai proteins as the “gold standard,” our
ELISA would show 100% sensitivity and 32% speciﬁcity. All
dot-blot positives were also ELISA positive, which suggests
an ELISA test using crude antigen isolated from adult
A. cantonensis from Hawai’i appears to be an effective initial
screening method and the 31-kDa dot blot appears to have
efﬁcacy as a diagnostic for exposure to A. cantonensis in
Hawai’i.
Hawai’i Island has extremely high infection rates in rats33 and
semi-slugs,31 and this study reveals human exposure levels
higher than those reported in endemic areas (China).25–27 Effective preventive measures are needed, especially in areas of
high transmission. Effective, less invasive diagnostics are also
needed in these areas to facilitate treatments. Currently,
HDOH requires PCR-based detection of parasite DNA in the
CSF to deﬁnitively diagnose angiostrongyliasis which is expensive, invasive, and can be inconclusive. Antibody-based
testing requires only a blood sample and provides a reasonable or supplemental alternative to a lumbar puncture test,
although one has to wait some time until antibodies are produced in sufﬁcient levels for detection. Detection of parasite
DNA in the blood provides an early diagnostic test and has
been demonstrated in rats,42 dogs, and horses (Unpublished
data). Availability of a reliable blood-based diagnostic test for
exposure to A. cantonensis in humans, pets, and livestock
would beneﬁt the public’s health wherever the disease exists
or emerges.
Received March 21, 2018. Accepted for publication October 14, 2019.
Note: Supplemental ﬁle appears at www.ajtmh.org.
Acknowledgments: We would like to thank Puna Community Medical
Center and staff for assistance in participant completion of the forms
and questionnaires necessary for conducting this study, and Clinical
Laboratories of Hawai’i for their assistance with blood sample collection. We would also like to thank Peter Hoffman, PhD, for his helpful
comments in reviewing this manuscript and the volunteers who
assisted with conducting this study including Patrick Edwards, Smiley
Burrows, Doran Vaughan, Patricia Macomber, Judy Hartshom, Jeremai Cann, and Shawzy Cann. We would also like to thank all the
volunteer participants who donated blood samples and took the time
to complete the questionnaire.
Financial support: This study was funded by the Hawaii Community
Foundation - Medical Research and the Daniel K. Inouye College of
Pharmacy, University of Hawaii at Hilo, Hilo, HI.
Authors’ addresses: Susan I Jarvi, Stefano Quarta, Kathleen Howe, Steven Jacquier, Kirsten Snook, Robert McHugh, Kuilei Kramer, and
McKayla Meyers, Department of Pharmaceutical Sciences, Daniel K.
Inouye College of Pharmacy, University of Hawai’i at Hilo, Hilo, Hawaii,

E-mails: jarvi@hawaii.edu, stefanoquartaemail@gmail.com, kehauhowe@
gmail.com, stevenjacquier@gmail.com, rmchugh@hawaii.edu, kui65@
hawaii.edu, mckayla@rams.colostate.edu, and ksnook@hawaii.edu.
Praphathip Eamsobhana, Department of Parasitology, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand, E-mail:
praphathip.eam@mahidol.ac.th. Alexandra Hanlon, Department of Statistics, Center for Biostatistics and Health Data Science, Roanoke, VA,
E-mail: alhanlon@vt.edu. Zachariah Tman, School of Public Health, University of California, Berkeley, CA, E-mail: tmanz@berkeley.edu. Jill
Miyamura, Hawaii Health Information Corporation, Kailua, HI, E-mail:
miyamura.jill@gmail.com.

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 "A pilot study to determine the prevalence of human rat lungworm infection in East Hawaii Island.”
Rat Lungworm Disease (RLWD) Questionnaire
Please return completed questionnaires in the enclosed, pre-paid envelope to:
Dr. Susan Jarvi,
Daniel K Inouye College of Pharmacy, University of Hawaii at Hilo,
34 Rainbow Drive,
Hilo HI 96720.
Tel 808 932-7701,
email jarvi@hawaii.edu)
Signatures for Consent:
I agree to participate in the survey entitled Rat Lungworm Disease Questionnaire. I understand that I can
change my mind about participating in the project, at any time, by notifying the researcher. I agree to allow
information collected on this survey to be shared anonymously at appropriate, scientific conferences and
publications, to further research and education outreach on RLWD by the University of Hawaii at Hilo, Daniel
K Inouye College of Pharmacy and the Department of Public Health Services, University of Hawaii at Manoa.
Name (Print): ________________________________ Date: ____________________

Signature: _____________________________________________________________

City and state of residence:________________________________________________
Environment of residency: City   Suburb  

Rural  

Household water supply: Catchment   Municipal  
Gender: M   F  
Age: __________

1. What is the highest level of education you have completed?
❏
❏
❏
❏
❏
❏

Less than High School
High School/GED
Some College
College/University Undergraduate Degree
University Graduate Degree
University Professional Degree
1

 2. What ethnicity do you MOST identify with? Please check ONE only.
❏
❏
❏
❏
❏
❏
❏
❏
❏
❏
❏
❏
❏
❏
❏
❏

3.

Chinese
Filipino
Japanese
Korean
Vietnamese
Portuguese
Native Hawaiian
Samoan
Micronesian
Marshallese
Other Pacific Islander
Alaska Native/American Indian
Black or African American
Hispanic or Latino
White or Caucasian
Other (specify): ____________________

Approximately how often do you do the following?
a. Eat salad at local restaurants?
___daily ___weekly ___monthly ___annually ___never
b. Eat other uncooked vegetables at local restaurants?
___daily ___weekly ___monthly ___annually ___never
c. Eat raw fruit at local restaurants?
___daily ___weekly ___monthly ___annually ___never
d. Eat unwashed salad at home?
___daily ___weekly ___monthly ___annually ___never
e. Eat other uncooked vegetables at home?
___daily ___weekly ___monthly ___annually ___never
f. Eat raw fruit at home?
___daily ___weekly ___monthly ___annually ___never
g. Eat washed, uncooked salad or other fruit or vegetables at home?
___daily ___weekly ___monthly ___annually ___never

2

 h. Buy fresh fruit or vegetables at the grocery store?
___daily ___weekly ___monthly ___annually ___never
i. Buy fresh fruit or vegetables at the farmer’s market?
___daily ___weekly ___monthly ___annually ___never

4. Do you believe that you are, were, or might have been infected by rat lungworm?
  Yes
  No
a. If no, discontinue this survey
b. If yes:
i. When do you think you were infected (month/year)?___________
ii. How do you think you were infected?
1. ____Eating contaminated unwashed raw fruits
2. ____Eating contaminated vegetables
3. ____Eating infected slug/snail
4. ____ Exposure to contaminated catchment water
5. ____Other (describe): ____________________________
iii. Where do you think you were infected?
1. State: ____Hawaii ____Other (specify): ____________
2. Island / county___________________________
3. District _________________________________
4. City / town _________________________________
5. Have you ever been diagnosed with rat lungworm disease?  Yes
If no, skip to the next question. If yes:
Who made the diagnosis?
____Doctor’s office
____Emergency room
____Hospital
____Naturopath
____Urgent Care
____Other (please describe)
How was the diagnosis confirmed?
____Spinal tap
____Antibody test
____Don’t know
____Other (please describe)
3

 No

 6. If you were hospitalized, what hospital were you admitted to and what were the dates of your stay?
Hospital:______________________________________

From:___________ To:____________

7. What medications were prescribed? How many days were you taking these medications?
Number of days
____Steroids
____________
____Anthelmintics (anti-parasitic drugs)
____________
____Others (please describe)
____________
____Don’t know
8. What were your symptoms within 3 months after being infected? Please check applicable symptoms and
circle level of discomfort with 1 being mildest and 5 being most severe. (if none, skip to the next
question)
Mildest

Most Severe

___Nausea

1

2

3

4

5

___Headache

1

2

3

4

5

___Meningitis

1

2

3

4

5

___High eosinophil counts

1

2

3

4

5

___Stiff neck

1

2

3

4

5

___Skin sensitivity

1

2

3

4

5

___Muscle soreness

1

2

3

4

5

___Fatigue

1

2

3

4

5

___Joint pain

1

2

3

4

5

___Fever

1

2

3

4

5

___Mental clarity

1

2

3

4

5

___Memory

1

2

3

4

5

___Speech

1

2

3

4

5

___Sexual dysfunction

1

2

3

4

5

___Vision

1

2

3

4

5

___Balance

1

2

3

4

5

___Bladder dysfunction

1

2

3

4

5

___Bowel dysfunction

1

2

3

4

5

___ Other (please describe below)

1

2

3

4

5

4

 9. What were/are your symptoms more than 3 months after being infected? Please check applicable
symptoms and circle level of discomfort with 1 being mildest and 5 most severe. (if none, skip to the
next question)
Mildest
1

2

3

Most Severe
4
5

___Headache

1

2

3

4

5

___Meningitis

1

2

3

4

5

___High eosinophil counts

1

2

3

4

5

___Stiff neck

1

2

3

4

5

___Skin sensitivity

1

2

3

4

5

___Muscle soreness

1

2

3

4

5

___Fatigue

1

2

3

4

5

___Joint pain

1

2

3

4

5

___Fever

1

2

3

4

5

___Mental clarity

1

2

3

4

5

___Memory

1

2

3

4

5

___Speech

1

2

3

4

5

___Sexual dysfunction

1

2

3

4

5

___Vision

1

2

3

4

5

___Balance

1

2

3

4

5

___Bladder dysfunction

1

2

3

4

5

___Bowel dysfunction

1

2

3

4

5

___ Other (please describe below)

1

2

3

4

5

___Nausea

May we contact you if we have further questions? If so, please provide your contact information.
Address: _________________________________
_________________________________
Phone:

_________________________________

Email:

_________________________________

Thank you for taking the time to complete our survey, we appreciate this very much.
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