EcoHealth

, 2:307

Evidence of Batrachochytrium dendrobatidis Infection in Water Frogs of the Rana esculenta Complex in Central Italy

Authors

  • Francesca Simoncelli
    • Dipartimento di Biologia Cellulare e AmbientaleUniversità di Perugia
  • Anna Fagotti
    • Dipartimento di Biologia Cellulare e AmbientaleUniversità di Perugia
  • Raffaele Dall’Olio
    • Istituto Nazionale di Apicoltura
  • Daniela Vagnetti
    • Dipartimento di Biologia Cellulare e AmbientaleUniversità di Perugia
    • Dipartimento di Biologia Cellulare e AmbientaleUniversità di Perugia
  • Ines Di Rosa
    • Dipartimento di Biologia Cellulare e AmbientaleUniversità di Perugia
Original Contributions

DOI: 10.1007/s10393-005-8337-8

Cite this article as:
Simoncelli, F., Fagotti, A., Dall’Olio, R. et al. EcoHealth (2005) 2: 307. doi:10.1007/s10393-005-8337-8

Abstract

Batrachochytrium dendrobatidis (phylum Chytridiomycota, order Chytridiales) is the causative organism of chytridiomycosis in amphibians, a disease associated with their population decline worldwide. In this work, we report a cutaneous infection in water frogs of the Rana esculenta complex in agricultural areas of Umbria, central Italy. Histological, immunohistochemical, ultrastructural, and molecular analyses demonstrated for the first time the presence of the Batrachochytrium dendrobatidis in this complex; to date, no association between the presence of chytrid fungal infection and mortality has been found, to our knowledge. However, the presence of Batrachochytrium dendrobatidis infection in the water frogs of the Rana esculenta complex is of concern because the frogs could act as a reservoir species and contribute to the decline of less resistant species.

Key words

amphibiansBatrachochytrium dendrobatidisRana esculentaagricultural areas

INTRODUCTION

Chytridiomycosis is an emerging infectious disease involved in mass mortality events associated with both wild and captive amphibian population decline in the Americas, the Caribbean, Australia, and Europe (Berger et al., 1998; Daszak et al., 1999; Pessier et al., 1999; Bosch et al., 2001; Bradley et al., 2002; Daszak et al., 2003; Morehouse et al., 2003; Muths et al., 2003). It has been demonstrated that both wild populations of Xenopus laevis in South Africa (Weldon, 2002) and Rana catesbeiana introduced into Uruguay and Venezuela (Mazzoni et al., 2003; Hanselmann et al., 2004) carry chytrid infections without mortality. In addition, experimentally-induced infections in Tiger salamanders (Davidson et al., 2003) and bullfrogs (Daszak et al., 2004) did not comprise mortality imputable to chytridiomycosis. The infective organism of chytridiomycosis is a nonhyphal fungus, Batrachochytrium dendrobatidis, a member of the fungal phylum Chytridiomycota in the order of Chytridiales (Longcore et al., 1999; Pessier et al., 1999). Clinical signs of chytridiomycosis include abnormal posture, loss of righting reflex, lethargy, and rapid progression to death. Gross skin lesions result in excessive sloughing of the epidermis, occasional ulceration, and hyperemia (Berger et al., 1999). Amphibian chytrid fungus infects and replicates within the keratinized cell layer of epidermis and is associated with hyperkeratosis and hyperplasia (Berger et al., 1999). The life cycle of the fungus involves a spherical monocentric or colonial zoosporangium, that is the growing phase, and motile, waterborne, uniflagellated zoospores, that are the infective stage. The zoospores are released from the zoosporangium through a discharge papilla that is usually extended towards the skin surface and gives a characteristic flask-like appearance to the zoosporangium (Berger et al., 1998; Longcore et al., 1999; Pessier et al., 1999).

The composition and health status of the Rana esculenta complex of the Trasimeno Lake district (Umbria) in central Italy have been monitored in a mark-recapture study since 1997 (Bucci et al., 2000; Pascolini et al., 2003; Fagotti et al., 2005; Mosconi et al., 2005). The water frogs of the Rana esculenta complex are widely distributed throughout peninsular Italy and constitute mixed populations of a nonhybrid taxon, Rana lessonae, and hemiclonally reproducing hybrids, Ranaesculenta, that are directly analogous to the well-studied central European lessonae/esculenta system. They display varying sensitivities to environmental stressors, whereas the parental species is much more sensitive to chemical contamination and climate changes (Bucci et al., 2000; Pascolini et al. 2003; Pereira et al., 2005; Mosconi et al., 2005). In an area (Solomeo) of the Trasimeno Lake district, the frequency of the parental species of the water frog population had reduced during the period 1999–2000 with respect to 1997–1998. In addition, the presence of an Amphibiocystidium ranae infection, low tissue levels of organochlorine compounds, and anomalies in the frog development such as oversized tadpoles were observed (Pascolini et al., 2003; Fagotti et al., 2005). The Solomeo area is characterized by increasing anthropogenic pressure; extensive and intensive agricultural land use, as well as chemical industries and urbanization, have caused habitat loss and pollution.

After exceptional meteorological events, including a drought in 2003, a further decline in the parental species was observed and the Amphibiocystidium ranae infection spread over a more extended area of Trasimeno Lake (Pereira et al., 2005). During the same period, some adult frog samples (5%) from Solomeo and from another agricultural area, Anguillara, had gross skin abnormalities. Here we report for the first time the presence of Batrachochytrium dendrobatidis in the water frogs of the Rana esculenta complex in Umbria, central Italy; to date, no episode of death related to chytridiomycosis has been observed, to our knowledge. However, the presence of Batrachochytrium dendrobatidis infection in the water frogs of the Rana esculenta complex is of concern because they could contribute to the decline of less resistant amphibian species.

METHODS

Sample Collection

Eighty adult water frogs of similar size were caught at various breeding sites in two agricultural areas (40 frogs from Solomeo and 40 from Anguillara) of the Trasimeno Lake district during the reproductive season. The frogs were captured during the night and kept in separate aquaria according to capture site. The following morning, the animals were transported to the laboratory and kept under conditions of natural photoperiod at 18°–20°C. The frogs were immediately subjected to experimental procedures. For each individual, snout-vent length, mass, and sex were determined. For the subsequent taxon determination and morphological analyses, the frogs were anesthetized with 0.05% solution of 3-amino benzoic acid ethyl ester (MS-222, Sigma-Aldrich, St. Louis, MO). To avoid duplicate recordings, captured frogs were marked by toe clipping using the numbering system devised by Hero (1989). This procedure consists in the amputation of the terminal phalanx of a toe other than the longest toe that is used during ecdysis. The frogs were released into their habitat after phalanx removal.

Taxon Determination

The clipped phalangs obtained from the marking method were used to determine the taxon by Southern blotting of genomic DNA, using the centromeric satellite DNA RrS1, a molecular marker, to reliably distinguish between hybrids and nonhybrids (Ragghianti et al., 1995; Bucci et al., 2000).

Morphological Analyses

The collected frogs were examined to determine the presence of gross lesions particularly in the abdominal, pelvic, and femoral regions, and the feet. When lesions were observed, skin biopsies were taken for light and electron transmission microscopy analyses. Sloughed skin and the surrounding intact skin were fixed in 10% buffered neutral formalin and processed routinely for embedding in paraffin. Sections (5 μm) were stained with hematoxylin and eosin and observed for histological diagnosis. For electron microscopy, samples were fixed in 2.5% glutaraldehyde in cacodylate buffer, postfixed in 1% OsO4, dehydrated and embedded in Epon-Araldite resin (Sigma-Aldrich, St. Louis, MO). Thin sections were stained with uranyl acetate and lead citrate and examined using a Philips 400 TEM (Eindhoven, The Netherlands) at 60 Kv.

Immunohistochemical Assay

Paraffin blocks from histological diagnosis were used for immunohistochemistry by indirect immunoperoxidase test. Sections were processed according to a modified version of the protocol described by Berger et al. (2002). Deparaffinized sections were treated with 3% H2O2 in methanol to block endogenous peroxidase activity. After rinsing in 10 mM sodium phosphate saline buffer (PBS), sections were preincubated with normal goat serum for 30 minutes and then incubated with Rabbit 667 anti-chytrid antiserum diluted 1:1000 in PBS containing 0.1% bovine serum albumin (BSA) for 2 hours. After washing in PBS, slides were incubated with biotinylated goat anti-rabbit IgG (Vector Laboratories Inc., Burlingame, CA) and developed with the avidin-biotinyl-peroxidase complex (Vector). The antigenic complex was visualized using a 3-3′’ diaminobenzidine (DBA) substrate kit (Vector). Slides were then counterstained with hematoxylin. All procedures were carried out at room temperature.

DNA Extraction, Polymerase Chain Reaction (PCR) Protocol, and Sequencing

Chytrid genomic DNA was extracted directly from the same infected skin samples that had been embedded in paraffin for histological analysis. Twenty-micrometer sections were deparaffinized in xylene, centrifuged at high speed, and the pellet was washed with 95% and 70% ethanol. Tissues were then dried and the genomic DNA was extracted following the protocol of Wizard Genomic DNA Purification Kit (Promega, Madison, WI). The extracted DNA was amplified by PCR using two specific primers, CHY18Ssh-F:5′GGCCTACCATGGTGATAACG and CHY18Ssh-R:3′CTGGCTACCATGTCCCAACT, designed on the basis of the Batrachochytrium nuclear gene encoding small-subunit ribosomal RNA (ssu-rDNA). The PCR reaction was performed in RoboCycler 40 (Stratagene, La Jolla, CA), by using GoTaq polymerase (Promega) under the following conditions: an initial denaturation step at 94°C for 2 minutes followed by 40 cycles of denaturing at 94°C for 1 minute, annealing at 50°C for 30 seconds, extension at 72°C for 2 minutes, and a 7-minute final extension at 72°C. The product obtained was purified with Concert Rapid Gel Extraction System (Invitrogen, Carlsbad, CA) according to the manufacturer’s protocol and sequenced on an automated CEQ 8000 Genetic Analysis System sequencer (Beckman Coulter, Fullerton, CA).

RESULTS

In the field-collected water frog samples, gross skin lesions, such as excessive sloughing and roughening of the superficial epidermis, were found in four Rana lessonae specimens, corresponding to 5% of the water frogs examined, mainly in the abdominal, pelvic, and femoral regions, and in the feet (Fig. 1). However, abnormal posture, reflex loss and lethargy, typical of chytridiomycosis disease were not observed. Histological examination of fixed and paraffin-embedded skin samples showed evidence of fungal infection. Spherical and ovoidal zoosporangia, flask-shaped mature zoosporangia each with a discharge papilla and containing zoospores, and empty zoosporangia were present in the keratinized cell layer of the epidermis (Fig. 2A,B). The immunohistochemical technique, performed using the rabbit 667 anti-chytrid antiserum by indirect immunoperoxidase test (Berger et al., 2002), demonstrated the presence of Batrachochytriumdendrobatidis. The antibody adhered to the wall, cytoplasm, and septa of the zoosporangia and the internal zoospores (Fig. 2C,D). Ultrastructural analysis of chytrid thalli revealed septate zoosporangia (Fig. 3A) and mature zoosporangia filled with zoospores (Fig. 3B,C) with morphology consistent with that described by Berger et al. (1998) and Longcore et al. (1999). Each zoospore had a single flagellum (Fig. 3B), cytoplasmic extensions, and a core of aggregated ribosomes, around which one or more lipid globules, mitochondria, and nucleus were distributed (Fig. 3C). A zoospore apparently being released through the discharge papilla of a flask-shaped zoosporangium is shown in Figure 3D. The infective organism was also confirmed by molecular analysis. The resulting partial sequence is made up of 366 bp. Blast analysis of the fragment revealed the highest match with the small subunit ribosomal RNA gene sequence of Batrachochytriumdendrobatidis. Accession number for the deposited sequence is AY842255.
https://static-content.springer.com/image/art%3A10.1007%2Fs10393-005-8337-8/MediaObjects/10393_2005_8337_f1.jpg
Figure 1

Gross skin abnormalities of Rana lessonae sample infected by Batrachochytrium dendrobatidis. Rough skin (arrows) is visible in the abdominal (A) and pelvic and femoral (B) regions.

https://static-content.springer.com/image/art%3A10.1007%2Fs10393-005-8337-8/MediaObjects/10393_2005_8337_f2.jpg
Figure 2

A,B: Hematoxylin and eosin staining of Rana lessonae infected skin by Batrachochytrium dendrobatidis. Spherical and ovoidal zoosporangia (arrows), flask-shaped mature zoosporangia (arrowheads) with discharge papilla and containing zoospores, and empty zoosporangia (double arrows) are detectable in the sloughed keratinized cell layer of epidermis (E). C,D:Rana lessonae infected skin stained with B. dendrobatidis-specific indirect immunoperoxidase stain. Sporangium wall and zoospores (C) and sporangium septa (D) are stained positively for B. dendrobatidis. Scale bars are indicated.

https://static-content.springer.com/image/art%3A10.1007%2Fs10393-005-8337-8/MediaObjects/10393_2005_8337_f3.jpg
Figure 3

Transmission electron micrographs of chytrid thalli in the keratinized cell layer of epidermis of Rana lessonae. A: Septate zoosporangium (arrows). B: Mature zoosporangium containing zoospores with a flagellum (arrowhead, F). C: Morphological details of a zoospore. The ribosome core (R) is bounded by lipid globules (L), mitochondria (M), and nucleus (N). D: Flask-shaped zoosporangia, each with a discharge papilla extended towards the skin surface through which zoospores are released (arrowhead). Scale bars are indicated.

DISCUSSION

We report a cutaneous fungal infection in field-collected water frogs of the Rana esculenta complex in central Italy. The histological, immunohistochemical, ultrastructural, and molecular findings are consistent with the presence of the chytrid fungus Batrachochytrium dendrobatidis. The diagnosis was made by identifying the characteristic intracellular flask-shaped zoosporangia and septate thalli within the superficial layer of the epidermis and by observing zoospore ultrastructure (Daszak et al., 1999; Longcore et al., 1999). The visualization of the fungus was supported by polyclonal antibody to Batrachochytrium dendrobatidis in an immunohistochemical assay (Berger et al., 2002). In addition, the isolation of a partial sequence of the small subunit ribosomal RNA gene, showing the highest match (99%) with Batrachochytriumdendrobatidis, confirmed the presence of this fungal infection. To our knowledge, this is the first reported case of chytrid infection in water frogs of the Rana esculenta complex in central Italy (Umbria). In the region of Umbria, these frogs are present in a variety of aquatic and terrestrial systems and are involved in both aquatic and terrestrial food webs, and constitute an important component of the biomass of many aquatic systems. Rana lessonae live in wet habitats where the air humidity is high; Rana esculenta are found in more aquatic areas. Our field data indicate that the presence of chytrid fungal infection is present in the parental species but is not associated with mortality. The individuals infected by Batrachochytriumdendrobatidis were from areas that have intensive anthropogenic pressure. In one of these areas, low levels of organochlorine compounds were detected in both frog tissues and water of breeding sites (Fagotti et al., 2005). In addition, during the year 2003, characterized by severe drought, significant changes in plasma androgens and estradiol-17β and thyroid hormones were recorded in adult male water frogs sampled from the same areas (Mosconi et al., 2005) and the spread of Amphibiocystidium ranae infection was notable with respect to previous monitoring (Pascolini et al., 2003; Pereira et al., 2005). These results suggest that complex interactions of different stress factors, such as habitat loss, climate changes, and environmental contaminants strongly impact the water frog system examined.

To date, the water frogs examined seem resistant to the more severe aspects of chytridiomicosis. Other authors have demonstrated that several amphibian species could have differential resistance to this disease in that they do not show all the characteristic pathological signs or evidence of mortality (Weldon, 2002; Davidson et al., 2003; Mazzoni et al., 2003; Daszak et al., 2004; Hanselmann et al., 2004; Rollins-Smith and Conlon, 2005). Further studies on the possible evolution of the chytrid infection, particularly in the neo-metamorphs, are in progress. However, the presence of Batrachochytriumdendrobatidis infection in the water frogs of the Rana esculenta complex is of concern because the frogs could act as a reservoir species and contribute to the decline of less resistant species, such as the yellow-bellied toad Bombina pachypus in the northern Apennines, in which chytridiomycosis has already had lethal effects (Stagni et al., 2004). Further investigations are needed to determine if other amphibian declines, such as that suspected of the green toad Bufo viridis in Umbria [B. Ragni, personal communication], are related to the presence of the fungus.

Acknowledgments

We thank Dr. A. Hyatt for providing the rabbit 667 antiserum and Dr. S. Bucci and Dr. M. Ragghianti for supplying the molecular marker RrS1. We are also grateful to Dr. R. Paracucchi for excellent technical assistance and to Dr. Nancy Hutchinson for critical reading of the manuscript. The work was supported by a grant from MIUR (Programmi di Ricerca Scientifica di Rilevante Interesse Nazionale) and by Agenzia Regionale Umbra per lo Sviluppo e l’Innovazione in Agricoltura.

Copyright information

© EcoHealth Journal Consortium 2005