INTRODUCTION
The superfamily Metastrongyloidea comprises a number of roundworms, which infest the cardiopulmonary system of vertebrate animals, and are primarily transmitted by pulmonate molluscs (Anderson, Reference Anderson2000). Some of these parasites may also have a zoonotic potential, as in the case of Angiostrongylus cantonensis and Angiostrongylus costaricensis (Strongylida, Angiostrongylidae), the causative agents of eosinophilic meningitis and ileitis, respectively (Wu et al. Reference Wu, French and Turner1997; Wang et al. Reference Wang, Lai, Zhu, Chen and Lun2008). Among feline lungworms, Aelurostrongylus abstrusus (Strongylida, Angiostrongylidae) is regarded as the most prevalent species, whereas Troglostrongylus brevior (Strongylida, Crenosomatidae) has been reported in felids from Palestine (Gerichter, Reference Gerichter1949), Italy (Brianti et al. Reference Brianti, Gaglio, Giannetto, Annoscia, Latrofa, Dantas-Torres, Traversa and Otranto2012, Reference Brianti, Gaglio, Napoli, Falsone, Giannetto, Latrofa, Giannelli, Dantas-Torres and Otranto2013), and Spain (Jefferies et al. Reference Jefferies, Vrhovec, Wallner and Catalan2010). Clinically, feline bronchopulmonary strongyloses may range from subclinical to life-threatening conditions, featured by dyspnoea, mucoid-purulent nasal discharge, sneezing, depression and anorexia, which can lead to exitus, especially during concomitant infections or immunosuppressive conditions (Brianti et al. Reference Brianti, Gaglio, Giannetto, Annoscia, Latrofa, Dantas-Torres, Traversa and Otranto2012). These lungworms share similar life cycles (Gerichter, Reference Gerichter1949) in that, after mating, females lay eggs, from which first-stage larvae (L1) hatch, ascend the respiratory tract up to the pharynx, where they are swallowed, eventually leaving the definitive host throughout feces (Gerichter, Reference Gerichter1949; Anderson, Reference Anderson2000). L1 of A. abstrusus and T. brevior may survive in the environment for up to 60 and 142 days, respectively, until they infest land or freshwater snails, and slugs (Gerichter, Reference Gerichter1949; Ash, Reference Ash1970; Gökpinar and Yildiz, Reference Gökpinar and Yildiz2010; Ramos et al. Reference Ramos, Giannelli, Dantas-Torres, Brianti and Otranto2013). In the intermediate host, development to the second-(L2) and third-(L3) larval stage occurs in about 2 weeks, depending on the gastropod species involved, as well as environmental factors (Gerichter, Reference Gerichter1949). The life cycle completes when a feline host ingests infested gastropod molluscs or paratenic hosts, harbouring infective L3 (Anderson, Reference Anderson2000). In addition, a direct route of transmission, from the queen to her kittens, has been speculated for T. brevior, via lactation or throughout the placenta (Brianti et al. Reference Brianti, Gaglio, Napoli, Falsone, Giannetto, Latrofa, Giannelli, Dantas-Torres and Otranto2013).
The global impact of feline bronchopulmonary strongyloses, coupled with the worldwide spread of several mollusc-borne diseases (e.g. human angiostrongyliasis, schistosomiasis, bilharziasis) (Wang et al. Reference Wang, Lai, Zhu, Chen and Lun2008; Majoros et al. Reference Majoros, Dán and Erdélyi2010; Soldánová et al. Reference Soldánová, Selbach, Kalbe, Kostadinova and Sures2013), spurred the interest of the scientific community on the biology of A. abstrusus and T. brevior (Di Cesare et al. Reference Di Cesare, Castagna, Meloni, Milillo, Latrofa, Otranto and Traversa2011; Barutzki and Schaper, Reference Barutzki and Schaper2013; Brianti et al. Reference Brianti, Gaglio, Giannetto, Annoscia, Latrofa, Dantas-Torres, Traversa and Otranto2012). From this perspective, the knowledge on the mollusc species involved in the epidemiology of feline lungworms is crucial to their prevention and control. The morphology of A. abstrusus and T. brevior larvae in molluscs has been poorly investigated (Gerichter, Reference Gerichter1949) and mostly restricted to the former lungworm species (López et al. Reference López, Panadero, Paz, Sánchez-Andrade, Díaz, Díez-Baños and Morrondo2005; Di Cesare et al. Reference Di Cesare, Crisi, Di Giulio, Veronesi, Frangipane di Regalbono, Talone and Traversa2013). Accordingly, the limited data available on T. brevior is due to the fact that L1s cannot be easily differentiated from those of A. abstrusus (Brianti et al. Reference Brianti, Gaglio, Giannetto, Annoscia, Latrofa, Dantas-Torres, Traversa and Otranto2012, Reference Brianti, Gaglio, Napoli, Falsone, Giannetto, Latrofa, Giannelli, Dantas-Torres and Otranto2013; Otranto et al. Reference Otranto, Brianti and Dantas-Torres2013). Interestingly, the occurrence of co-infestation (Jefferies et al. Reference Jefferies, Vrhovec, Wallner and Catalan2010) indicates that both species occupy the same ecological niches, thus potentially challenging the proper diagnosis and therapeutic schedule of feline infestations (Brianti et al. Reference Brianti, Gaglio, Giannetto, Annoscia, Latrofa, Dantas-Torres, Traversa and Otranto2012, Reference Brianti, Gaglio, Napoli, Falsone, Giannetto, Latrofa, Giannelli, Dantas-Torres and Otranto2013).
Hence, the present research aimed to provide insights into the development of A. abstrusus and T. brevior in Helix aspersa snails, and morphological information of larval stages.
MATERIALS AND METHODS
Maintenance of snails
In December 2012, 50 farmed specimens of H. aspersa were purchased from a snail farming centre from Teramo (Abruzzo region, central Italy), where this species is bred for human consumption. The absence of natural infections by any metastrongyloid larvae was assessed by microscopic examination of 10 snails at 5 days before the infestation, and of all snail specimens that died naturally during the observation period. The molluscs were put in plastic boxes, filled with fresh soil, which was changed and humidified every 3 days. Feeding of snails occurred every 2 days with potatoes, lettuce and water ad libitum. In addition, the upper part of the vivarium was covered with a net, which was wetted every 8 h with a water sprayer, in order to maintain proper ventilation and humidity in the box. The vivarium was kept in a temperature-controlled room (20±1 °C), where all procedures below were carried out.
Larval collection and infestation procedures
L1 of A. abstrusus and T. brevior were isolated from the feces of a 50-day-old naturally infested female kitten. The cat was referred to the Department of Veterinary Medicine of the University of Bari (southern Italy), due to a persistent cough and dyspnoea. Larvae were retrieved by Baermann technique, as previously described (MAFF, 1986). The solution containing fecal sedimentation and larvae was firstly centrifuged at 600 g for 5 min, the supernatant removed, and the sediment observed under light microscopy (Leica®, DM LB2). Nematode species were differentiated on the basis of the size and tail morphology and then molecularly identified (see below). Single infective doses of 100 L1s each were kept in small tubes, until used for snail infestation.
Helix aspersa snails (n = 30) were deprived of food 24 h before the infestation. Then, they were placed individually in a plastic infestation chamber, composed of six circular wells, containing a potato slice (1 cm wide) (Fig. 1A), which was contaminated with the infective dose (Fig. 1B). The infestation chamber was covered with a wet gauze cloth and secured with rubber bands, after ascertaining that snails had their foot on the tuber slice, to maximize contact with larvae (Fig. 1C). Specimens were left in the infestation chamber for 48 h and then removed into their vivarium.
Fig. 1. Helix aspersa infestation with feline lungworms larvae. Snails were placed individually in a plastic infestation chamber (A), containing a potato slice, which was contaminated with the infective dose (B). The infestation chamber was covered with a wet gauze cloth and closed with several rubber bands (C).
Larval isolation and morphological identification
Larval development of A. abstrusus and T. brevior was assessed using 5 snail specimens for each of the following time points: 2 (T1), 6 (T2) and 11 (T3) days post infestation at 20±1 °C. In addition, 15 of the infested H. aspersa snails were exposed to outdoor conditions from December to April 2013 (mean temperature: 10·6 °C; min–max: 6·6–15·0 °C), and finally examined 120 days post infestation (T4), when snails recovered from natural hibernation, as described in the following.
Snails were pooled and digested in a solution of 1% HCl (150 mL) and 1·2 g powdered pepsin (A/S N Foss Electric, Hillerod, Denmark), after their foot was separated from the body and cut into small pieces (1–2 mm). The suspension was stirred on a magnetic, heated plate at 37 °C for 35 min, strained through a 180 μm sieve to remove undigested material, collected in plastic tubes, centrifuged at 600 g for 5 min, before re-suspending the pellet in 5 mL tap water. Three replicates (100 μL each) of the suspension were microscopically examined and larvae were morphologically and morphometrically identified according to species and developmental stage (Gerichter, Reference Gerichter1949; Euzeby, Reference Euzeby1981), using an analyser program (Leica LAS® AF 4.1). Drawings were made with an optical microscope (Olympus® BX51) equipped with a camera lucida. Microscopic images and measures were taken using a digital image processing system (AxioVision rel. 4.8, Carl Zeiss®, Germany).
Molecular procedures
Single larval specimens of A. abstrusus and T. brevior (n = 10 per each species), at different developmental stages, were isolated from the medium using a 10 μL micropipette and stored in plastic vials containing phosphate buffer saline (PBS) at −20 °C, until analysed. Briefly, the DNA was extracted using a commercial kit (DNeasy Blood & Tissue Kit, Qiagen, GmbH, Hilden, Germany), in accordance with the manufacturer's instructions, and a partial 18S ribosomal RNA gene sequence (1708 bp) was amplified from individual larvae using primers NC18SF1 (5′-AAAGATTAAGCCATGCA-3′) and NC5BR (5′-GCAGGTTCACCTACAGAT-3′), as previously described (Patterson-Kane et al. Reference Patterson-Kane, Gibbons, Jefferies, Morgan, Wenzlow and Redrobe2009). Sequences were determined from both strands, using the same primers individually as for the PCR, and the electropherograms verified by eye. In order to ensure open reading frames, the nucleotide sequence was conceptually translated into amino acid sequence, using the invertebrate mitochondrial code MEGA5 software. Sequences were compared with those available in the GenBank™ database by Basic Local Alignment Search Tool (BLAST) (Altschul et al. Reference Altschul, Madden, Schäffer, Zhang, Zhang, Miller and Lipman1997).
RESULTS
All specimens of H. aspersa (n = 10) digested 5 days before the infestation, as well as those that died naturally during the pre-infestation period (n = 10), were negative for nematode larvae. Conversely, all pooled snails were positive for A. abstrusus and T. brevior. In particular, of 91 larvae retrieved from the H. aspersa examined, 48 (52·7%) were identified as A. abstrusus and 43 (47·3%) as T. brevior. Up to 27 larvae were detected in a single sample with a mean concentration of 22·75 larvae/sample (Table 1). The morphological identification was confirmed as the 18S rRNA sequences from larvae showed a 100% overall nucleotide BLAST homology with those of A. abstrusus and T. brevior (accession numbers AJ920366 and JX290562, respectively), previously deposited in GenBank™ (Table 2). At each time point, larvae of both species were at the same developmental stage with L1 found at T1, L2 at T2 and L3 at T3. After the overwintering period (T4) only A. abstrusus and T. brevior L3 were detected (Table 1).
Table 1. Aelurostrongylus abstrusus and Troglostrongylus brevior developmental larval stages detected in Helix aspersa foot, following the pepsin-HCl digestion (300 μL), at different time points post infestation
Table 2. Measurements (mean length and width±s.d.; expressed in μm) of Aelurostrongylus abstrusus and Troglostrongylus brevior larvae
Aelurostrongylus abstrusus larvae
L1 (Fig. 2A) measured 384·5±33·4 μm in length and 17·7±2·6 μm in width, and were featured by a narrowed anterior extremity, with several granules on the middle part of the body. The tail, which appeared forked-like, was bent in the form of S, with a small short appendage split on its dorsal side. The shape of the tip of the tail was constant in all specimens examined.
Fig. 2. Aelurostrongylus abstrusus. (A) First-stage larva (L1) and detail of the tail; (B) Second-stage larva (L2); (C) Third-stage larva and details of the anterior extremity and tail, ending into a rounded projection.
L2 (Fig. 2B) were 479·4±53·6 long and 27·6±4·5 wide, being incorporated into an external cuticle, which appeared empty at both end portions. The anterior extremity was rounded and the metacorpus filled by numerous refractive food granules. The tail was conoid, resembling that of the previous stage.
L3 (Figs 2C and 4A) were recovered enclosed into the two cuticles or without the external sheaths, and showed a slender body 538·9±51·8 μm long, with a maximum body width of 26·7±1·9 μm, at about mid-body. The anterior end was blunt and beard lateral alae, extending from 40–65 μm (mean 53 μm) to mid-tail, interrupted by the excretory pore at 87–96 μm (mean 92 μm) from the anterior end. The stoma, 19 μm long, was characterized by slightly cuticularized cheilorhabdion (4–5 μm long) followed by stylet-like organ with prominent barb-like points, extending into insertion in upper oesophagus (Fig. 2C). The muscular oesophagus, 183–210 μm long (mean 202 μm) and 7–10 μm wide (mean 8 μm) was almost cylindrical to the level of excretory pore and, subsequently increased in diameter up to 13–16 μm (mean 14 μm). The relative length of oesophagus to body length was 0·37–0·44 (mean 0·41) and the nerve ring was indistinctly observed at 75–80 μm from the anterior end. The tail was conical with rounded projection, measuring 34–40 μm in length (mean 36 μm) and 15–18 μm in width at anus level, with relative tail length to body length being 0·067–0·080 (mean 0·073) (Figs 2C and 4A).
Troglostrongylus brevior larvae
L1 (Fig. 3A) were 347·3±12·4 μm long and 16·1±1·5 μm wide, with a pointed anterior extremity and with granules confined to the second half of the body. The tail gradually tapered to the extremity, being featured by a deep incision, which divided the appendage into a shallower ventral spine and a conspicuous dorsal appendix, further divided near its tip. Individual variations of the shape of the caudal extremity were observed, with the tail more or less squared (Fig. 3A).
Fig. 3. Troglostrongylus brevior. (A) First-stage larva (L1) and detail of the tail; (B) Second-stage larva (L2); (C) Third-stage larva (L3) and details of the anterior extremity and tail, ending in four terminal appendages.
Fig. 4. Morphology of third stage larvae (L3) of Aelurostrongylus abstrusus (A) and Troglostrongylus brevior (B).
L2 (Fig. 3B) measured 380·7±18·6 μm in length and 24·9±2·7 μm in width. Similarly to the L2 of A. abstrusus, they were enclosed into an external cuticle, although specimens without the sheath were also observed. The anterior extremity was narrowed, widening progressively to the metacorpus, which intestinal cells were packed with many food granules. The tail tapered into an appendage similar to that of the previous stage.
L3 (Figs 3C and 4B) measured 432·1±15·3 μm in length, with the maximum body width of 20·9±1·5 μm, observed at about mid-body. Lateral alae were absent and the rounded anterior end was featured by a slightly cuticularized stoma, 3 μm long, which opened in a 121–145 μm long (mean 135 μm) muscular oesophagus (Fig. 4B). This was characterized by a 5–6 μm wide procorpus, a slightly swollen metacorpus, a long isthmus and basal bulb, 10–12 μm wide (mean 11 μm). The relative length of oesophagus to body length was 0·32–0·42 (mean 0·37). The excretory pore and the nerve ring were detected at 50–79 μm (mean 70 μm) and 60–67 μm (mean 65 μm) from anterior end, respectively. The tail, 30–38 μm long (mean 33 μm) and 13–14 μm wide at the anus level, ended in four terminal appendages (Figs 3C and 4B), with a relative length to body length of 0·081–0·103 (mean 0·090).
DISCUSSION
Results of the present study indicate that A. abstrusus and T. brevior may develop simultaneously in H. aspersa, reaching the third-larval stage in about 11 days post infestation. In addition, larvae of both metastrongyloids may overwinter in this snail species, remaining viable for at least 120 days. Data on the biology of these nematodes in the intermediate hosts are of relevance, considering that the information available is limited to the development of A. abstrusus in land (e.g. Helicella spp., Achatina fulica, Monacha syriaca and Theba pisana) and freshwater snails (i.e. Biomphalaria glabrata) or slugs (e.g. Agrolimax spp. and Limax flavus) (Gerichter, Reference Gerichter1949; Ash, Reference Ash1970; Ohlweiler et al. Reference Ohlweiler, Guimarães, Takahashi and Eduardo2010). Helix aspersa has been herein confirmed as intermediate host for A. abstrusus (Hobmaier and Hobmaier, Reference Hobmaier and Hobmaier1935; Hamilton, Reference Hamilton1969; Di Cesare et al. Reference Di Cesare, Crisi, Di Giulio, Veronesi, Frangipane di Regalbono, Talone and Traversa2013), and, for the first time, for T. brevior. Indeed, the known host range for the latter lungworm species was restricted to a small number of molluscs (i.e. Helicella barbesiana, Helicella vestalis joppensis, Chondrula septemdentata, M. syriaca, Retinella nitellina, T. pisana and L. flavus), but no information was available on their larval developmental times (Gerichter, Reference Gerichter1949).
From an epidemiological viewpoint, the environmental dissemination of this snail might impact on the distribution of feline lungworm infestations. Indeed, H. aspersa has been already reported from several areas previously considered non-endemic for this snail (e.g. Oceania and South Africa), as well as in northern European regions (Guiller and Madec, Reference Guiller and Madec2010). In some of these areas (e.g. Germany), the infestation rate by feline lungworm was shown to range from 0·5 to 15·6% (Barutzki and Schaper, Reference Barutzki and Schaper2013). Considering that larvae of A. abstrusus do not develop in snails at 4–8 °C (Gerichter, Reference Gerichter1949; Di Cesare et al. Reference Di Cesare, Crisi, Di Giulio, Veronesi, Frangipane di Regalbono, Talone and Traversa2013) but, at the same low temperature, T. brevior can reach the infective stage (Gerichter, Reference Gerichter1949), lungworm infestations in central and northern Europe could be due predominantly to T. brevior. In contrast, at mild temperatures, such as those of the Mediterranean area, both species may occur in sympatry. Based on these findings, H. aspersa may play an important role in the dissemination of both nematodes, maintaining infective larvae throughout the winter in hibernating molluscs.
The larval development time reported in the present study for A. abstrusus is shorter than that reported in H. barbesiana (11 and 18 days, at 30 °C) and Cernuella virgata (12 and 18 days, at 20 °C) (Gerichter, Reference Gerichter1949; López et al. Reference López, Panadero, Paz, Sánchez-Andrade, Díaz, Díez-Baños and Morrondo2005), whereas the development of T. brevior in H. aspersa is similar to that recorded in H. barbesiana and H. vestalis joppensis, with L2 and L3 detected 5 and 8 days post infestation, respectively (Gerichter, Reference Gerichter1949). Overall, data suggest that the occurrence of T. brevior and A. abstrusus in their intermediate hosts might display seasonal variations in relationship with the seasonal dynamic of their intermediate hosts. Indeed, snails are most prevalent in middle spring/early autumn when most kittens are born (Little, Reference Little2011). Importantly, the interactions between snails and paratenic hosts (e.g. birds, rodents and reptiles) should also be taken into account when considering the ecology of these parasites (Brianti et al. Reference Brianti, Gaglio, Giannetto, Annoscia, Latrofa, Dantas-Torres, Traversa and Otranto2012, Reference Brianti, Gaglio, Napoli, Falsone, Giannetto, Latrofa, Giannelli, Dantas-Torres and Otranto2013).
Data on the morphology of A. abstrusus and T. brevior herein presented do not concur with those previously reported in a case of co-infestation in a kitten from Spain (Jefferies et al. Reference Jefferies, Vrhovec, Wallner and Catalan2010), as larvae of both species were clearly inverted in their identification. Molecular tools may assist and expedite the proper identification of these nematodes, both in vertebrate and mollusc hosts. Nonetheless, further studies are needed to simultaneously diagnose A. abstrusus and T. brevior, instrumentally to proper therapeutics (Annoscia et al. Reference Annoscia, Latrofa, Campbell, Giannelli, Ramos, Brianti, Dantas-Torres and Otranto2013). Indeed, while active compounds have been licensed for the prevention and treatment of A. abstrusus, it is not the case for T. brevior. Overall these data may improve the current understanding on the risk period for cats to acquire the infestation from snails, as well the population dynamics of these nematodes in receptive snails. Indeed, A. abstrusus and T. brevior co-evolve in H. aspersa, potentially co-infesting the definitive hosts. Practitioners should be aware of the occurrence of this crenosomatid and perform proper diagnostic techniques when a bronchopulmonary strongyloses in a feline patient is suspected.
ACKNOWLEDGEMENTS
The authors thank Bronwyn Campbell (University of Bari) for comments on the manuscript.
FINANCIAL SUPPORT
This research received no specific grant from any funding agency, commercial or not-for-profit sectors.
