Magnetic capture hybridization of Batrachochytrium dendrobatidis genomic DNA

Journal of Microbiological Methods 90 (2012) 156–159

Contents lists available at SciVerse ScienceDirect

Journal of Microbiological Methods journal homepage: www.elsevier.com/locate/jmicmeth

Note

Magnetic capture hybridization of Batrachochytrium dendrobatidis genomic DNA David Rodriguez ⁎, Ana V. Longo, Kelly R. Zamudio Department of Ecology and Evolutionary Biology, Cornell University, Ithaca, NY 14853, USA

a r t i c l e

i n f o

Article history: Received 10 January 2012 Received in revised form 26 April 2012 Accepted 26 April 2012 Available online 2 May 2012

a b s t r a c t We hybridized biotinylated probes that anneal at multiple locations throughout the Batrachochytrium dendrobatidis (Bd) genome to selectively capture Bd genomic DNA (gDNA) by binding the probe-gDNA complex to streptavidin coated magnetic beads. We then whole genome amplified the captured gDNA. This method extends the usefulness of field-collected swabs for downstream PCR-based genomic applications. © 2012 Elsevier B.V. All rights reserved.

Keywords: Batrachochytrium dendrobatidis Chytrid Magnetic capture hybridization Whole genome amplification

Batrachochytrium dendrobatidis (hereafter Bd) is a recently discovered chytrid fungal pathogen that occurs globally and causes the disease chytridiomycosis in amphibians (Berger et al., 1998; Longcore et al., 1999). This emergent pathogen has been implicated as a major factor causing population declines and extinctions of amphibians, even in pristine habitats (Kilpatrick et al., 2010; Stuart et al., 2004). The life cycle of Bd consists of a sessile zoosporangium phase in amphibian skin within which flagellated zoospores grow and are released into water, where they in turn infect other hosts (Berger et al., 1998; Longcore et al., 1999; Piotrowski et al., 2004). Clinical signs of chytridiomycosis include hyperkeratosis, skin sloughing, and host death (Berger et al., 1998; Voyles et al., 2007). Diagnosis of Bd in frogs is typically performed by histological inspections of skin (Berger et al., 1999; Pessier et al., 1999), immunochemical assay (Berger et al., 2002), or PCR-based detection (Annis et al., 2004). In addition, real-time PCR (RT-PCR) allows quantification of Bd infection intensity from field collected swabs of amphibian skin, which is measured as number of zoospore equivalents or genomic equivalents (Boyle et al., 2004). Increasingly, studies are using swabbing and PCR-based methods to characterize the presence and infection intensity of Bd in amphibian populations (Hyatt et al., 2007) leading to a better understanding of the population dynamics of this pathogen (http://www.Bd-maps. net/). However, recent detailed studies of the evolutionary genetic relationships among Bd strains have necessarily relied on culturing to obtain genetic material (Farrer et al., 2011; Fisher et al., 2009; James et al., 2009; Morehouse et al., 2003; Morgan et al., 2007). Isolating and culturing Bd from amphibian skin is difficult, time consuming, and ⁎ Corresponding author at: E221 Corson Hall, Department of Ecology and Evolutionary Biology, Cornell University, Ithaca, NY 14853, USA. Tel.: +1 806 470 0855; fax: +1 607 255 8088. E-mail address: [email protected] (D. Rodriguez). 0167-7012/$ – see front matter © 2012 Elsevier B.V. All rights reserved. doi:10.1016/j.mimet.2012.04.021

requires sacrificing amphibian hosts (Longcore et al., 1999); thus, reliance on fungal cultures for population genetic analyses has limited fine-scale studies of this pathogen due to low sample sizes. An ideal method would permit us to obtain high-yield Bd DNA from field collected, nondestructive samples. Unfortunately, swabs from amphibian skin usually contain low quantities of Bd genomic DNA (gDNA). As is typical of environmental samples, the target DNA is mixed with large quantities of host DNA, DNA from other microorganisms, and PCR inhibitors (Rådström et al., 2004). In population genetic studies of Bd, this can lead to unreliable amplification rates and limit sample sizes across sampling sites (Velo-Antón et al., 2012). Magnetic beads are routinely used to isolate and purify cells and nucleic acids (Lund et al., 1988; Olsvik et al., 1994). In pathogen detection assays, magnetic beads in conjunction with biotinylated probes are used to target‐specific genomic regions from samples containing host DNA or other biological contaminants [e.g., clinical specimens (Mangiapan et al., 1996), food products (Chen et al., 1998), and soil (Jacobsen, 1995)]. Magnetic beads and target-specific oligonucleotide capture probes are also used to capture bacterial gDNA for pathogen detection using RT-PCR (Parham et al., 2007). In cases where gene or genome copy is low, whole genome amplification (WGA) methods can be used to increase the amount of starting genetic material for PCRbased applications from as little as one cell (Sato et al., 2005; Spits et al., 2006). The ϕ29 polymerase and random hexamers can polymerize up to 1–2 μg of gDNA from less than 1 ng of gDNA input with high sequence fidelity and genome coverage (Spits et al., 2006). In this study, we combine magnetic capture hybridization (MCH) and WGA to selectively extract and amplify Bd gDNA from swabs of amphibian skin (Fig. 1). Our method extends the usefulness of field-collected swabs for downstream genome-scale PCR-based applications and circumvents the difficulties associated with low gDNA concentrations and PCR inhibitors.

D. Rodriguez et al. / Journal of Microbiological Methods 90 (2012) 156–159

PrepMan extract

*

157

*

Magnetic capture

*

*

*

*

*

*

Bead decoupling

Elution

*

MCH

*

*

*

Probe hybridization

Bead coupling

*

Host DNA Bd DNA Inhibitors Probe Beads

*

*

*

WGA

Fig. 1. Diagram of MCH-WGA extraction method starting with PrepMan extract from amphibian skin swabs. Final WGA product can be used in downstream PCR reactions.

We used a hemocytometer to count 1 × 10 7 zoospores harvested from cultures JEL 427 (Puerto Rico) and MX01 (Mexico). We then extracted the zoospores and made serial dilutions of the DNA to use as the template for our bead optimization experiments. We tested control swabs loaded with 1 × 10 5 zoospores of strain MX01 and field-collected swabs that were previously quantified using RT-PCR (Boyle et al., 2004) to evaluate the efficiency of the selective extraction of Bd gDNA in the presence of other genomic material and impurities. We carried out all extractions using 50 μl of PrepMan Ultra (Applied Biosystems) and a 10 min incubation at 100 °C, the standard protocol for field-collected samples (Boyle et al., 2004; Hyatt et al., 2007). We used the program progressiveMauve (Darling et al., 2010) to identify two sets of repeated regions among the 17 largest supercontigs (97%) of the Bd genome (strain JEL423; http://www.broadinstitute. org). We designed three probes within these repeat regions, Bd1BioTEG (5'-TAACGTCACACTTTGACTTGATGA-3'), Bd2BioTEG (5'-GCATCATACTG TCCATTCTTGAAC-3'), and Bd2BioTEG54 (5'-GCAGGGTTCAAGAATG GACAGTATGATGCTGTATAGTGTAGTTCCAAACCTCGC-3'). We BLAST searched the probe sequences to insure they were specific to Bd based on the current database of accessioned sequences. The capture probes were modified by the addition of a triethylene glycol spacer and biotin at the 5' end. Integrated DNA Technologies synthesized and HPLC-purified the probes. We tested 1 μm (New England Biolabs) and 2.8 μm (Dynabeads M-280, Invitrogen) magnetic beads covalently coupled to streptavidin, which has a high affinity for biotin. We used 2.8 μm beads, which were designed to immobilize larger fragments (>2 kb), with the washing and binding buffer supplied in the Dynabeads kilobase BINDER kit following the manufacturer's instructions. To prepare the 1 μm magnetic beads, we added 10 μl of beads (4 mg/ml) in preservative into 0.2 ml strip tubes and then added 40 μl of binding and washing solution (BWS; 0.5 M NaCl, 0.02 M Tris, 0.001 M EDTA). The tubes were placed on a 96-well plate magnet (Agencourt Bioscience) for 30 s and the supernatant was removed leaving the beads behind. Two more washes using 50 μl of BWS solution were performed in the same manner before resuspending the beads in 10 μl of BWS. We hybridized the probe with target Bd gDNA in 30 μl reactions comprised of 10 μl of PrepMan extract, 15 μl of 12X saline-sodium citrate (SSC), 1.5 μl of double-deionized H2O, 1.5 μl of 1% sodium dodecyl sulfate

solution, 1 μl of bovine serum albumin (BSA) (400 ng/μl), and 1 μl of a single biotinylated probe (10 μM) or 0.5 μl of each biotinylated probe (10 μM), if two were used. We then denatured the gDNA at 98 °C for 15 min, carried out the probe hybridization at 40–50 °C for 1–3 h, and cooled the reaction to room temperature on a rotator for 10 min. To optimize the probe hybridization reaction, we varied number of probes (one or two), probe length (24 bp or 54 bp), and hybridization temperatures. To test for the effect of SSC concentration on the hybridization reaction, we tested higher (5X and 10X) and lower (15X and 20X) stringency conditions. To couple the gDNA-biotinylated probe complex to the magnetic beads, we added the hybridization reaction to 10 μl of washed beads and incubated the solution at 45.0–67.7 °C for 30 min in a thermocycler. At 15 min, we briefly vortexed the reaction to resuspend the beads and continued the incubation. At 30 min, the reaction was cooled to room temperature and placed on a magnet for 30 sec. We removed all of the supernatant, added 50 μl of elution buffer (5 mM Tris–Cl, 0.25 mM EDTA; pH 9.0), and vortexed the tube to resuspend the beads. We decoupled the biotin-streptavidin bond by incubating at 95 °C for 10 min. After the reactions cooled to room temperature, we placed them on a magnet for 30 s and pipetted the supernatant into a new sterile tube. The eluted product was used as the template for downstream PCR, RT-PCR, and WGA reactions using the Illustra GenomiPhi V2 DNA Amplification Kit following the manufacturer's instructions (GE Healthcare). We carried out RT-PCR on products from the hybridization optimization experiments in replicates of three to assess whether changing reaction conditions affected percent recovery. We performed RT-PCR on a ViiA7 (Applied Biosystems) following protocols outlined in Boyle et al. (2004). In addition to our RT-PCR tests of loaded control swabs and field-collected frog skin swabs, we also PCR amplified six previously characterized loci for Bd (James et al., 2009) to determine the gDNA concentration threshold for successful endpoint PCR amplification. Based on JEL423 (Broad Institute) as the reference genome, two of these markers are located on separate ends of the largest supercontig (SC1). The remaining four loci are located on supercontigs SC5, SC9, SC10, and SC15. We did this PCR verification using the following templates: (i) pre-MCH extraction products (gDNA), (ii) post-MCH products (MCH-DNA), (iii) post-WGA of pre-MCH products (WGA gDNA), and (iv) post-WGA of MCH (WGA MCH-DNA) products. For

158

D. Rodriguez et al. / Journal of Microbiological Methods 90 (2012) 156–159

each marker, we performed endpoint PCR amplifications in 25.0 μl final volumes using 1 μl of template, 17.8 μl of ddH2O, 2.5 μl of 10X PCR reaction buffer (100 mM Tris–HCl, 15 mM MgCl2, and 500 mM KCl), 0.5 μl of dNTP mix (10 mM each), 1 μl (10 μM) of upstream primer, 1 μl (10 μM) of downstream primer, 1 μl of BSA (400 ng/μl), and 0.2 μl (1 U) of Taq polymerase (Roche). We used a touchdown thermal profile beginning with denaturation for 2 min at 95 °C; followed by 20 cycles of 30 s at 95 °C, 50 s starting at 65 °C and decreasing 0.5 °C every cycle, and 30 s at 72 °C; then 20 cycles of 30 s at 95 °C, 50 s at 50 °C, and 30 s at 72 °C; with a final extension for 5 min at 72 °C. Successful amplification was verified by electrophoresis of PCR products on a 2% agarose gel stained with ethidium bromide. Capture probes Bd1BioTEG, Bd2BioTEG54, and Bd2BioTEG anneal 12, 22, and 27 times on the JEL423 reference genome (Broad Institute), respectively. We found no exact matches for these probes in other taxa in a BLAST search of GenBank; thus, we can safely assume that these probes selectively hybridize to Bd only. However, if our probes anneal on another chytrid genome, this would not affect downstream PCRbased tests for Bd, which rely on Bd‐specific markers. When comparing bead types, we found that 1 μm beads performed more efficiently than 2.8 μm beads (Fig. 2A, Kruskal–Wallis P b 0.001);

therefore, we used 1 μm beads in all subsequent MCH reactions. Percent recovery was higher when using two probes instead of one (Fig. 2B, Kruskal–Wallis P =0.04), and lower hybridization temperatures resulted in generally higher recovery, but the outcome of different temperatures was not significant (Fig. 2C, Kruskal–Wallis P =0.88). Different SSC concentrations did not significantly increase recovery, but concentrations above 5X performed better (Fig. 2D, Kruskal–Wallis P =0.30). The optimal bead coupling temperature was 45.0 °C (Fig. 2E, Kruskal–Wallis P =0.03), and increasing coupling time did not significantly increase recovery (Fig. 2F, Kruskal–Wallis P = 0.73). For field-collected and loaded swab extractions, we used the optimal conditions inferred from our previous tests (Fig. 2) and performed hybridizations at 40 °C with probes BdBioTEG1 and BdBioTEG2. We set the bead coupling temperature to 45 °C for 1 h and used 12X SSC. WGA of MCH products revealed that four out of six Bd loci successfully amplify if PrepMan extractions contain at least 9.3 genomic equivalents per microliter (g.e./μl), whereas amplifications typically failed for pre‐ and post-WGA PrepMan extractions below that threshold (Table 1). We attributed this to WGA biased towards host genetic material. This result indicates that WGA alone will not resolve the difficulties with low quantities of Bd genomic material on amphibian skin swabs; rather,

Bead Type 100

Probe 100

A

80

80

60

60

40

40

20

20

0

0

B

2 probes

1 probe

Dynabeads (2.8 µm)

New England Biolabs (1.0 µm)

Bd1BioTEG

Bd2BioTEG

24 bp 12 sites

24 bp 27 sites

Hybridization Temperature (°C)

Percent Recovery

100 80

80

60

60

40

40

20

20

Bd2BioTEG

Bd2BioTEG Bd2BioTEG54

D

0

0 40.0 40.9 42.9 45.8 49.5 52.4 54.1

56.0

5X

Bead Coupling Temperature (°C) 100

54 bp 22 sites

SSC Concentration 100

C

Bd2BioTEG54 Bd1BioTEG

80

80

60

60

40

40

20

20

0

12X

15X

20X

Bead Coupling Time (hrs) 100

E

10X

F

0 45.0

46.5

49.6

54.2

60.0 64.8

67.6

1

2

3

Fig. 2. Box plots showing percent recovery of the MCH method, measured by RT-PCR. In every case, 1 × 105 genomic equivalents of Batrachochytrium dendrobatidis were used as the genomic input for each optimization experiment.

D. Rodriguez et al. / Journal of Microbiological Methods 90 (2012) 156–159

159

Table 1 Results of Bd loci amplification tests on genetic material extracted from swabs using the MCH-WGA method. Successful amplifications at each of the six loci are denoted by (+). Reactions that were not successful or detectable by electrophoreses are denoted by (−). Sample ID

Source

Concentration (g.e./ul)a

gDNAb, c

ADK65 ADK71 ADK87 JEL427 ADK99 ADK91 ADK70 ADK68 JEL427 ADK55 JEL427

Frog skin Frog skin Frog skin Cultured Frog skin Frog skin Frog skin Frog skin Cultured Frog skin Cultured

0.8 0.9 1.4 2.1 2.5 3.5 9.3 12.2 20.8 36.8 208

−−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− ++++++

a b c d

MCH-DNAc − − − − − − − − − −

−−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− −−−−− ++++++

− − − − − − − − − −

WGA gDNAc

WGA MCH-DNAc

−−−−−− −−−−−− −−−−−− −−−−−− −−−−−− −−−−−− −−−−−− −−−−−− ++++++ −−−−−− ++++++

−−−−+− −−−−−− − − − ++ − ++++++ −−−+−− − − − ++ − ++++ − − +++++ − ++++++ ++++++ ++++++

Genomic equivalents per microliter, determined by RT-PCR. Extracted from a swab using PrepMan Ultra. Amplification determined by PCR for loci 8329x9 (SC1), 9893x2 (SC1), r6046x2 (SC5), 8702x2 (SC9), mb-b13(SC10), and CSTYN1 (SC15). Swabs loaded with cultured Bd zoospores.

it must be used after a genomic enrichment method, such as the one we describe here. Our method can be integrated into the workflow of current protocols for the detection and quantification of Bd loads from field-collected swabs. This method permits the recovery, purification, and amplification of Bd genomic material for use in downstream PCR-based applications including sequencing and genotyping. Acknowledgements We thank Steven M. Bogdanowicz and Karen Kiemnec-Tyburczy for technical assistance, and C. Guilherme Becker for field-collected swabs. This study was supported by the National Science Foundation (DBI‐0905810, DEB‐0815315, DEB‐1120249). References Annis, S.L., Dastoor, F.P., Ziel, H., Daszak, P., Longcore, J.E., 2004. A DNA-based assay identifies Batrachochytrium dendrobatidis in amphibians. J. Wildl. Dis. 40, 420–428. Berger, L., Speare, R., Daszak, P., Green, D.E., Cunningham, A.A., Goggin, C.L., Slocombe, R., Ragan, M.A., Hyatt, A.D., McDonald, K.R., Hines, H.B., Lips, K.R., Marantelli, G., Parkes, H., 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc. Natl. Acad. Sci. U. S. A. 95, 9031–9036. Berger, L., Speare, R., Kent, D.A., 1999. Diagnosis of chytridiomycosis of amphibians by histologic examination. Zoos' Print J. 15, 184–190. Berger, L., Hyatt, A.D., Olsen, V., Hengstberger, S.G., Boyle, D., Marantelli, G., Humphreys, K., Longcore, J.E., 2002. Production of polyclonal antibodies to Batrachochytrium dendrobatidis and their use in an immunoperoxidase test for chytridiomycosis in amphibians. Dis. Aquat. Organ. 48, 213–220. Boyle, D.G., Boyle, D.B., Olsen, V., Morgan, J.A.T., Hyatt, A.D., 2004. Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman PCR assay. Dis. Aquat. Organ. 60, 141–148. Chen, J.R., Johnson, R., Griffiths, M., 1998. Detection of verotoxigenic Escherichia coli by magnetic capture hybridization PCR. Appl. Environ. Microbiol. 64, 147–152. Darling, A.E., Mau, B., Perna, N.T., 2010. progressiveMauve: Multiple Genome Alignment with Gene Gain, Loss and Rearrangement. PLoS One 5. Farrer, R.A., Weinert, L.A., Bielby, J., Garner, T.W.J., Balloux, F., Clare, F., Bosch, J., Cunningham, A.A., Weldon, C., Louis, H., Anderson, L., Kosakovsky, S.L., Shahar-golan, R., Henk, D.A., Fisher, M.C., 2011. Multiple emergences of genetically diverse amphibian- infecting chytrids include a globalized hypervirulent recombinant lineage. PNAS 108, 18732–18736. Fisher, M.C., Garner, T.W.J., Walker, S.F., 2009. Global emergence of Batrachochytrium dendrobatidis and amphibian chytridiomycosis in space, time, and host. Annu. Rev. Microbiol. 63. Hyatt, A.D., Boyle, D.G., Olsen, V., Boyle, D.B., Berger, L., Obendorf, D., Dalton, A., Kriger, K., Hero, J.M., Hines, H., Phillott, R., Campbell, R., Marantelli, G., Gleason, F., Colling, A., 2007. Diagnostic assays and sampling protocols for the detection of Batrachochytrium dendrobatidis. Dis. Aquat. Organ. 73, 175–192.

Jacobsen, C.S., 1995. Microscale detection of specific bacterial-dna in soil with a magnetic capture-hybridization and pcr amplification assay. Appl. Environ. Microbiol. 61, 3347–3352. James, T.Y., Litvintseva, A.P., Vilgalys, R., Morgan, J.A.T., Taylor, J.W., Fisher, M.C., Berger, L., Weldon, C., du Preez, L.H., Longcore, J.E., 2009. Rapid global expansionof the fungal disease chytridiomycosis into declining and healthy amphibian populations. PLoS Pathol. 5, e1000458. Kilpatrick, A.M., Briggs, C.J., Daszak, P., 2010. The ecology and impact of chytridiomycosis: an emerging infectious disease of amphibians. Trends Ecol. Evol. 25, 109–118. Longcore, J.E., Pessier, A.P., Nichols, D.K., 1999. Batrachochytrium dendrobatidis gen et sp nov, a chytrid pathogenic to amphibians. Mycologia 91, 219–227. Lund, V., Schmid, R., Rickwood, D., Hornes, E., 1988. Assessment of methods for covalent binding of nucleic-acids to magnetic beads, dynabeads, and the characteristics of the bound nucleic-acids in hybridization reactions. Nucleic Acids Res. 16, 10861–10880. Mangiapan, G., Vokurka, M., Shouls, L., Cadranel, J., Lecossier, D., vanEmbden, J., Hance, A.J., 1996. Sequence capture-PCR improves detection of mycobacterial DNA in clinical specimens. J. Clin. Microbiol. 34, 1209–1215. Morehouse, E.A., James, T.Y., Ganley, A.R.D., Vilgalys, R., Berger, L., Murphy, P.J., Longcore, J.E., 2003. Multilocus sequence typing suggests the chytrid pathogen of amphibians is a recently emerged clone. Mol. Ecol. 12, 395–403. Morgan, J.A.T., Vredenburg, V.T., Rachowicz, L.J., Knapp, R.A., Stice, M.J., Tunstall, T., Bingham, R.E., Parker, J.M., Longcore, J.E., Moritz, C., Briggs, C.J., Taylor, J.W., 2007. Population genetics of the frog-killing fungus Batrachochytrium dendrobatidis. Proc. Natl. Acad. Sci. U. S. A. 104, 13845–13850. Olsvik, O., Popovic, T., Skjerve, E., Cudjoe, K.S., Hornes, E., Ugelstad, J., Uhlen, M., 1994. Magnetic separation techniques in diagnostic microbiology. Clin. Microbiol. Rev. 7, 43–54. Parham, N.J., Picard, F.J., Peytavi, R., Gagnon, M., Seyrig, G., Gagne, P.A., Boissinot, M., Bergeron, M.G., 2007. Specific magnetic bead-based capture of genomic DNA from clinical samples: Application to the detection of group B streptococci in vaginal/anal swabs. Clin. Chem. 53, 1570–1576. Pessier, A.P., Nichols, D.K., Longcore, J.E., Fuller, M.S., 1999. Cutaneous chytridiomycosis in poison dart frogs (Dendrobates spp.) and White's tree frogs (Litoria caerulea). J. Vet. Diagn. Investig. 11, 194–199. Piotrowski, J.S., Annis, S.L., Longcore, J.E., 2004. Physiology of Batrachochytrium dendrobatidis, a chytrid pathogen of amphibians. Mycologia 96, 9–15. Rådström, P., Knutsson, R., Wolffs, P., Lövenklev, M., Löfström, C., 2004. Pre-PCR processing: strategies to generate PCR-compatible samples. Mol. Biotechnol. 26, 133–146. Sato, M., Ohtsuka, M., Ohmi, Y., 2005. Usefulness of repeated GenomiPhi, a phi29 DNA polymerase-based rolling circle amplification kit, for generation of large amounts of plasmid DNA. Biomol. Eng. 22, 129–132. Spits, C., Le Caignec, C., De Rycke, M., Van Haute, L., Van Steirteghem, A., Liebaers, I., Sermon, K., 2006. Whole-genome multiple displacement amplification from single cells. Nat. Protoc. 1, 1965–1970. Stuart, S.N., Chanson, J.S., Cox, N.A., Young, B.E., Rodrigues, A.S.L., Fischman, D.L., Waller, R.W., 2004. Status and trends of amphibian declines and extinctions worldwide. Science 306, 1783–1786. Velo-Antón, G., Rodriguez, D., Savage, A., Parra-Olea, G., Lips, K., Zamudio, K., 2012. Amphibian-killing fungus loses genetic diversity as it spreads across the New World. Biol. Conserv. 146, 213–218. Voyles, J., Berger, L., Young, S., Speare, R., Webb, R., Warner, J., Rudd, D., Campbell, R., Skerratt, L.F., 2007. Electrolyte depletion and osmotic imbalance in amphibians with chytridiomycosis. Dis. Aquat. Organ. 77, 113–118.