The rat lungworm (Angiostrongylus cantonensis) is a parasitic nematode that causes rat lungworm disease. It is the leading cause of eosinophilic meningitis and is a zoonotic health risk. We confirmed the presence of A. cantonensis using species-specific, quantitative PCR in 18 of 50 (36%) giant African land snails (Lissachatina fulica) collected from Miami, Florida, US in May 2013. These snails were collected from seven of 21 core areas that the Florida Department of Agriculture and Consumer Services monitor weekly. Rat lungworms have not previously been identified in these areas. Duplicate DNA extractions of foot muscle tissue from each snail were tested. Of the seven core areas we examined, six were positive for A. cantonensis and prevalence of infection ranged from 27% to 100%. Of the 18 positive snails, only five were positive in both extractions. Our results confirm an increase in the range and prevalence of rat lungworm infection in Miami. We also emphasize the importance of extracting sufficient host tissue to minimize false negatives.
The rat lungworm (Nematoda; Angiostrongylus cantonensis) is a common parasite of rats and snails globally, and the cause of the emerging infectious neurologic rat lungworm disease (Jarvi et al. 2012). Rat lungworms can cause eosinophilic meningitis in humans and animals (Reece et al. 2013). The complex life cycle of the rat lungworm has been described in detail with both definitive mammalian hosts (multiple species of rats; Rattus spp.) and intermediate gastropod hosts (land and freshwater snails and slugs) (Thiengo et al. 2013). Mature adult female nematodes are found in the pulmonary arteries of the definitive host, where they lay eggs. These eggs develop into first-stage larvae and move to the interior of the alveoli. Over time, the larvae migrate to the pharynx, are swallowed, pass through the gastrointestinal tract, and are excreted into the environment in the feces (Thiengo et al. 2013). Feces are ingested by the intermediate host and the immature nematodes develop into infective third-stage larvae (Reece et al. 2013). After ingestion of the intermediate host by a suitable mammalian host, the infective third-stage larvae penetrate the gut wall and are carried via the bloodstream to the brain (Reece et al. 2013). In the brain of the definitive host, they develop to subadults and re-enter the bloodstream where they transit to the pulmonary arteries and mature to adults (Reece et al. 2013). Rats become infected by ingesting infected intermediate hosts (snails or slugs) or paratenic hosts (Jarvi et al. 2012).
Besides the definitive and intermediate hosts, there are numerous paratenic and accidental hosts for rat lungworms. Paratenic hosts include flatworms (Pseudoceros spp.), frogs (Rana spp.), toads (family Bufonidae), freshwater prawns (Macobrachium spp.), land crabs (family Gecarcinidae), and monitor lizards (Varanus indicus) (Wang et al. 2008; Jarvi et al. 2012). Although the parasites do not develop in paratenic hosts, these hosts may be a source of infection for definitive or accidental hosts by acting as a carrier of the parasite that can be digested by definitive hosts (Morassutti et al. 2014). Accidental hosts, including humans, are primarily infected by consumption of raw or undercooked intermediate or paratenic hosts (Kim et al. 2014). Once ingested by the accidental host, the third stage larvae die when they reach the brain or spinal cord, which can lead to eosinophilic meningitis (Kim et al. 2014). In the US, the rat lungworm has been detected in accidental host animals in Louisiana, Mississippi, and Florida (Teem et al. 2013). Globally, rat lungworms have been found in many accidental host species such as grey-headed fruit bats (Pteropus poliocephalus) (Reddacliff et al. 1999), horses (Equus ferus), dogs (Canis lupus familiaris), multiple species of marsupials (subclass, Marsupialia), gang-gang cockatoos (Callocephalon fimbriatum), and tawny frogmouths (Podargus strigoides) (Reece et al. 2013).
One of the most noted intermediate hosts for the rat lungworm is the giant African land snail; Lissachatina fulica, also known as Achatina fulica (Lv et al. 2008; Kim et al. 2014). Lissachatina fulica is an invasive species that was first introduced to Florida in 1966 and was successfully eradicated by 1975 (Mead 1979). During 2011, giant African land snails were again detected in Miami, Florida, and rat lungworms identified within the snail population a year later (Florida Department of Agriculture and Consumer Services, Division of Plant Industry 2012). However, rat lungworm infection in a white-handed gibbon (Hylobates lar) and an orangutan (Pongo pygmaeus) had been recorded earlier from this area (Emerson et al. 2013). Here we report a change of protocol that resulted in detecting an increase in frequency of rat lungworm infection in giant African land snails in Miami.
We sampled 50 giant African land snails during 16–18 May 2013. These snails were collected from seven of the 21 core areas established by the Florida Department of Agriculture and Consumer Services, Division of Plant Industry, where the snails have been found around Miami–Dade County and that are monitored weekly (Fig. 1). Core areas are defined as an initial detection site surrounded by a 1.6-km-diameter buffer zone. Snails were returned to the laboratory alive, individually bagged, labeled, weighed, and measured, then killed by decapitation within 5 h of collection. The majority of the foot muscle was removed, cut into small pieces, and stored in 96% ethanol until processed. Genomic DNA (gDNA) was extracted from two 25-mg muscle samples using the DNeasy tissue kit (Qiagen, Valencia, California, USA) as per manufacturer-suggested protocols. These duplicate extractions were labeled A and B. The DNA was eluted with 100 μL of elution buffer and stored at 4 C until quantitative PCR (qPCR) amplification was performed following the methods of Qvarnstrom et al. (2010). Primers and probe are specific for the Angiostrongylus cantonensis inner transcribed spacer region 1 (ITS1). Positive controls (gDNA extracted from A. cantonensis) were included in all assays to verify qPCR performance. All snail DNA samples, the positive control, and negative control were run in triplicate. The DNA of a plasmid (pCR2.1 vector containing a single A. cantonensis ITS1 insert) was quantified using the Qubit® double-stranded DNA broad range assay (Life Technologies, Grand Island, New York, USA). The molecular mass of the plasmid was determined by multiplying the length of the plasmid (with insert) in base pairs by the average molecular mass (650) of a single base pair. The inverse of the molecular mass was then taken to calculate the number of moles/g of plasmid. Multiplication by Avogadro's number (6.022×1023) yielded the number of molecules (copies)/g of plasmid DNA. This value was used to convert the concentration of the plasmid standard to copies/μL. Dilutions of plasmid DNA were used to determine the number of copies of A. cantonensis ITS1 in snail DNA samples (Qvarnstrom et al. 2010) by comparison of the qPCR response of unknown snail DNA samples with those of a standard dilution series (102 to 108 copies/μL). Standard curves were generated by linear regression of logarithm of the dilution series concentrations versus the cycle number required for the qPCR response to cross the assay threshold. Calculations of standard measures of real-time analysis quality were calculated using ABI viiA™7 version1.2.1 (Life Technologies) and included determination of the linear regression coefficient (r2) for standard dilution series and reaction efficiency calculations. Samples containing ≥100 copies/μL, as estimated by the plasmid-based standard curve, were scored as positive. Infectivity was calculated by dividing the number of true positive outcomes by the total number of samples.
The coefficient of determination for the assay was r2 = 0.98. Of the 50 snails sampled, 18 from six cores were identified as positive for A. cantonensis via qPCR (Table 1). Only core 10 was negative, but only one snail was tested from that area and the negative result may simply reflect low sample size. Of the 18 snails found positive, only five were positive in both extractions A and B. In cases in which both samples were positive, the quantification estimates were highly variable (Table 2), indicating that rat lungworm larvae are not uniformly distributed in muscle tissue. There was no correlation between snail weight and infectivity (copies/μL).
Rat lungworms were first detected in giant African land snails sampled in Miami during 2011–12. In September 2012, 140 snails were collected from 12 of the 19 core areas that had been established at the time of their collection (Teem et al. 2013). Teem et al. (2013) followed the same qPCR protocol as used in our study and found that individuals found positive for rat lungworms were from core four only. The prevalence of infection at this site was 7.4%. We collected one snail from core four during this study and it was positive for rat lungworm. The range of L. fulica infected with A. cantonensis has since expanded within Miami. The rat lungworm does not have a specific intermediate host. It is estimated that species from over 50 nonnative gastropod families have been introduced into the US inadvertently or intentionally (Robinson 1999), any of which could be potential intermediate hosts. In addition, climate change is believed to influence both disease range (pathogen spread) and indirect (hosts) expansions by allowing parasites such as the rat lungworm or molluscan hosts to occur in areas they previously could not as their range is limited by temperature (York et al. 2014). The rat lungworm is an emerging threat to humans and wildlife that should be routinely monitored. The island applesnail (Pomacea maculate), the giant ramshorn snail (Marisa cornuarietis), and the giant African land snail are the only snail species that have been tested for rat lungworm infection in Florida (Teem et al. 2013). As giant African land snails and island applesnails are larger than most other species of snails that host rat lungworms and are widespread in Florida, multiple samples must be extracted from muscle tissue to decrease the number of false negatives. Additional research should be conducted to determine an optimal sampling protocol to adequately test for rat lungworms.
This investigation was made possible by the US Geological Survey, Sharon Gross, and Laurie Allen. We also thank the Florida Department of Agriculture and Consumer Services (including the Golf Team and Mary Yong Cong) and Ryan Braham. Any use of trade, product, or firm names is for descriptive purposes only and does not imply endorsement by the US government.