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Keywords:

  • sponge orange band;
  • sponge disease;
  • bleaching;
  • Xestospongia muta;
  • cyanobacteria;
  • microbial consortia

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

The aim of this study was to examine sponge orange band (SOB) disease affecting the prominent Caribbean sponge Xestospongia muta. Scanning and transmission electron microscopy revealed that SOB is accompanied by the massive destruction of the pinacoderm. Chlorophyll a content and the main secondary metabolites, tetrahydrofurans, characteristic of X. muta, were significantly lower in bleached than in healthy tissues. Denaturing gradient gel electrophoresis using cyanobacteria-specific 16S rRNA gene primers revealed a distinct shift from the Synechococcus/Prochlorococcus clade of sponge symbionts towards several clades of unspecific cyanobacteria, including lineages associated with coral disease (i.e. Leptolyngbya sp.). Underwater infection experiments were conducted by transplanting bleached cores into healthy individuals, but revealed no signs of SOB development. This study provided no evidence for the involvement of a specific microbial pathogen as an etiologic agent of disease; hence, the cause of SOB disease in X. muta remains unidentified.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

To date, coral diseases have received enormous attention owing to the important role of coral reefs in global biodiversity as well as the alarming rate at which reefs have been decimated in the context of global warming (Rosenberg et al., 2007; Mao-Jones et al., 2010). In comparison, much less effort has been undertaken to investigate diseases of marine sponges. Disease epidemics can in fact cause drastic reductions of sponge populations, with negative effects on the overall reef ecology (Webster, 2007). Sponge diseases have been documented in all major ocean bodies, with the Caribbean representing a particular hot spot (Harvell et al., 1999). They typically start with the appearance of discolored patches, followed by tissue disintegration, leaving behind the exposed skeleton. The size and consistency of the sponge seem to affect the pattern of disease progression, which may spread over the entire body within weeks to months. It is noteworthy that the disease typically does not switch host species for reasons that are not understood. Research on sponge diseases is hampered by the fact that the death of any given sponge often goes unnoticed.

The factors that cause disease in sponges are still largely unknown. A clear correlation has been documented between high (Cerrano et al., 2000; Perez et al., 2000; Lopez-Legentil et al., 2008; Webster et al., 2008a; Maldonado et al., 2010) and low temperatures (Perez et al., 2006) and sponge disease. The role of temperature is important in the context of global warming and climate change, which impact the composition of entire ecosystems (Hughes et al., 2003; Hoegh-Guldberg et al., 2007; Whiteman, 2010). Among other abiotic factors, hurricanes, physical damage and water turbidity do not seem to be major causes of sponge death (Wulff, 2006). Exposure of the sponge Rhopaloeides odorabile to the heavy metal copper has been shown to cause severe necrosis as well as shifts in its microbial community composition (Webster et al., 2001). Even though it has been difficult to pinpoint the exact causes, environmental stress is likely to decrease the fitness of sponges, which may, in turn, render them more susceptible to disease.

Diverse microorganisms such as Alphaproteobacteria (Webster et al., 2002), representatives of the genera Bacillus and Pseudomonas (Cervino et al., 2006), cyanobacteria (Rützler, 1988; Olson et al., 2006), fungi (Galstoff, 1942) and viruses (Vacelet & Gallissian, 1978) have been implicated as disease-causing agents among sponges. However, in only one case have Koch's postulates been upheld, linking a causative microorganism to a specific disease (Webster et al., 2002). A number of additional studies implicated microbial pathogens as etiological agents of sponge disease (Webster, 2007); however, a bona fide sponge pathogen has rarely been identified. Recently, there has been a paradigm shift in that environmental factors rather than infectious agents are considered as the main causes of sponge disease (Luter et al., 2010).

Our study focuses on the barrel sponge Xestospongia muta (Demospongiae, Haplosclerida), one of the largest, oldest and most common members of Caribbean coral reef communities (Armstrong et al., 2006; McMurray et al., 2008) and a species whose populations are increasing as coral cover has decreased (McMurray et al., 2010). Unsaturated polyacetylenic brominated acids (Ashok et al., 1992), xestosterol and mutasterol have been isolated from X. muta (Li et al., 1981). Recently, a family of chiral tetrahydrofurans, mutafurans A–G, with antifungal activities has been obtained from this sponge species (Morinaka et al., 2007). Xestospongia muta harbors dense and phylogenetically distinct microbial communities extracellularly within its mesohyl and is therefore a representative of the high microbial abundance (HMA) group of sponges (Hentschel et al., 2006). The surface layers (pinacoderm) are populated by the Synechococcus/Prochlorococcus clade of cyanobacterial symbionts, which are also responsible for the distinctive, reddish-brown surface coloration (Steindler et al., 2005). Vertical transmission of its microbial symbionts through the reproductive elements has been demonstrated (Schmitt et al., 2008).

Massive bleaching events of Caribbean X. muta specimens were observed in Puerto Rico (Vicente, 1990), Belize and the Florida Keys (Gammill & Fenner, 2005), Curaçao (Nagelkerken et al., 2000) as well as offshore Cuba and the reefs of Cozumel (Mexico) (Gammill & Fenner, 2005). The most recent outbreak took place in June of 2009 in the US Virgin Islands, where approximately 20% of the resident X. muta population was affected (T. Smith, pers. commun.). Two types of bleaching have been described: cyclic bleaching, from which sponges recover, and fatal bleaching, synonymous with sponge orange band (SOB) disease, which usually results in sponge death (Cowart et al., 2006; Lopez-Legentil et al., 2008). The disease starts with lesions on the outer surface and spreads over the sponge body within weeks, leaving exposed skeleton behind. SOB is frequently accompanied by an orange transition band, giving rise to the name (Cowart et al., 2006). In the present study, we aimed to provide an in-depth characterization of the pathology of fatal SOB disease as well as changes in the chemical and microbial community profiles during disease progression. Furthermore, attempts to prove Koch's postulates were undertaken by conducting underwater infection experiments.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Sponge collection

Samples of healthy and diseased X. muta sponges (class Demospongiae, order Haplosclerida, family Petrosiidae) were collected by SCUBA at a depth of 4–30 m within the Florida Keys National Marine Sanctuary in November and December 2007 and September 2009. Additional sampling was conducted in June 2007 and 2008 during research expeditions throughout the Bahamas onboard the RV Seward Johnson II. The samples were transferred to the surface in seawater-containing Ziploc bags and kept cool until further processing within 1–2 h. Altogether, seven healthy and 12 diseased individuals were obtained. The term sponge ‘tissue’ is used hereafter in a colloquial sense, as sponges are made up of a confederation of cell types and do not have true tissues or organs (Brusca & Brusca, 1990).

Electron microscopy

Scanning electron microscopy (SEM) was performed on pieces of tissue of 0.5 cm3 size from selected X. muta specimens following established protocols by Scheuermayer et al. (2006). Briefly, this involved the excision of sponge tissue, storage in 6.25% glutaraldehyde phosphate-buffered solution, washing procedures with Soerensen phosphate buffer (50 mM, pH 7.4) and dehydration in an increasing acetone series. The samples were critical point dried, sputtered with gold-palladium and stored in the dehydrator until examination using the scanning electron microscope (Zeiss DSM 962, Oberkochen, Germany). Transmission electron microscopy (TEM) was performed on pieces of tissue of about 1.0 mm3 size from selected X. muta individuals following the procedure of Schmitt et al. (2008). The prepared samples were sectioned using an MT-7000 ultramicrotome (RMC, Tuscon, AZ) for examination with the transmission electron microscope (Zeiss EM10).

Chlorophyll a analysis

Chlorophyll a content was measured via spectrophotometry (Parsons et al., 1984) on three 0.5 g tissue cubes per individual. The samples were maintained in a 90% acetone : water mixture at 4 °C in the dark until analysis. After centrifugation, the absorbance of 1 mL supernatant was measured at 750, 664, 647 and 630 nm using an optic spectrophotometer (Amersham Biosciences, Ultrospec 3100 pro). The chlorophyll a content was calculated using the equations of Parsons et al. (1984) and standardized to the volume of sponge tissue.

HPLC

Lyophilized sponge samples were ground with a mortar and pestle and extracted overnight by stirring at room temperature with twice the volume of 100% methanol. After vacuum filtration, the supernatants were dried on a rotary evaporator at 40 °C. For each extract, 5.0 mg were dissolved in 1.0 mL of methanol, from which 5.0 μL was analyzed via analytical HPLC using a Jasco system (pump PU1580, gradient unit LG-980-025, degasser DG-2080-53 and UV detector MD-2010Plus) on a Chromolith RP-18e column (4.6 × 100 mm; Merck). The separation was achieved with a solvent mixture of acetonitrile and water complemented with 0.05% trifluoroacetic acid, starting with 10% acetonitrile : H2O to 100% acetonitrile over a time span of 15 min and at a flow rate of 3 mL min−1.

Preparative isolation was carried out using an HPLC Jasco system (pump PU1580, gradient unit LG-980-025, degasser DG-2080-53 and UV detector MD-2010Plus) on a Chromolith SemiPrep RP-18e column (10 × 100 mm; Merck). A solvent gradient of 10% acetonitrile : H2O to 100% acetonitrile supplemented with 0.05% trifluoroacetic acid was used over a time span of 15 min at a flow rate of 10 mL min−1. HPLC–ESI-MS/MS analysis was performed on 5.0 μL of healthy X. muta extract utilizing a triple-stage quadrupole 7000 tandem mass spectrometer system equipped with an ESI interface (Finnigan MAT, Bremen, Germany). Data acquisition and evaluation were conducted on DEC 5000/33 digital equipment (Unterfoehring, Germany) using ICIS 8.1 software (Finnigan MAT). Positive ions were detected by scanning from 170 to 900 u with a 1-s scan duration for a single spectrum.

Denaturing gradient gel electrophoresis (DGGE)

For DNA extraction, sponge samples preserved in 70% ethanol and 0.5 cm3 in size were air-dried and homogenized with a mortar and pestle in liquid nitrogen. The DNA extraction and 16S rRNA gene amplification for DGGE were performed as described in Schmitt et al. (2008), with the following modifications: the primer pair 106f with a GC-clamp and 781r was used for PCR amplification of the cyanobacterial 16S rRNA gene (Nübel et al., 1997) and the primer pair 341f with a GC-clamp and 907r (Muyzer et al., 1998) was used for the bacterial 16S rRNA gene. The PCR was conducted using a T3 Thermocycler (Biometra, Germany) using the following conditions: initial denaturation at 94 °C for 2 min, 34 cycles of denaturation at 94 °C for 1 min, primer annealing at 60 °C (cyanobacterial primers) or 57 °C (bacterial primers) for 1 min following elongation at 72 °C for 75 s. The final extension step was at 72 °C for 10 min. The size and quality of the PCR products obtained were examined on 2% agarose gels following staining with ethidium bromide. DGGE was conducted as described by Schmitt et al. (2008) on 8% w/v polyacrylamide gels. Selected DGGE bands were excised under UV-light using an ethanol-sterilized scalpel and extracted overnight with 25 μL H2O at 4 °C. Cluster analyses of the DGGE banding pattern were conducted using the software program quantity one (Bio-Rad, Munich, Germany). Dendrograms were constructed using the UPGAMA clustering method as defined by quantity one to compare the banding pattern similarities in between different samples within one gel.

Cloning and sequencing

The excised DGGE bands were reamplified with the primers 106f and 781r (Nübel et al., 1997), and the obtained PCR products were purified, ligated into the vector pGEM-T-Easy (Promega) and transformed into Escherichia coli NovaBlue cells. The plasmid DNA was isolated using standard miniprep procedures (Sambrook et al., 1989) and digested with EcoRI to confirm the correct insert size by agarose gel electrophoresis. Sequencing was conducted for up to four clones per DGGE band as described in Schmitt et al. (2008). Chimeras were identified using the program pintail (Ashelford et al., 2005) by comparison against the five most closely related 16S rRNA gene sequences from culturable bacteria and removed from the dataset.

Phylogenetic analysis

The arb program package (Ludwig et al., 2004) was used for sequence alignment and phylogenetic tree construction. The derived X. muta sequences and their closest basic local alignment search tool (blast) hits were imported into the SILVA 16S rRNA gene database (version 93) (Pruesse et al., 2007) for automatic alignment and manual refinement using the arb integrated alignment tool. Only sequences≥1200 bp were used for the calculation of neighbor joining, maximum likelihood and maximum parsimony trees. Shorter sequences were added using the parsimony interactive tool in arb without changing the tree topology. All trees were constructed using conservation filters for cyanobacteria and bootstrap analysis (1000 resamplings). The resulting trees were compared and the maximum likelihood tree was chosen for publication. The highlighted group was supported by all three treeing methods. All 16S rRNA gene sequences were deposited in GenBank (accession numbers GU590802–GU590859).

Infection experiments

The experiments were conducted at a water depth of ∼20 m on Conch Reef (24°56′863″N; 80°27′230″W). For this purpose, tissue samples of 1.5 cm diameter and ∼5 cm depth were removed with a corkborer from diseased sponges and placed in healthy individuals where identical holes had been created. Nine tissue cores were removed per diseased sponge and three cores were placed into each of three healthy individuals. This experiment was performed for a total of three diseased sponges and, in the same fashion, for one healthy individual as a control, resulting in a total of 36 transplanted cores. The overall health condition of the transplanted sponges was monitored regularly by SCUBA over 11 days.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Underwater photography

Healthy specimens of X. muta are barrel-shaped and have a brown, irregular surface, often with protrusions (Fig. 1a). Diseased individuals show a distinctive change from brown to a bleached white color. Bleaching typically starts as isolated patches penetrating the first few millimeters of the surface tissue (Supporting Information, Fig. S1). Eventually, the entire sponge body may be affected (Fig. 1b and c). This transition can be accompanied by an orange band, giving rise to the name ‘sponge orange band’ of this disease (Fig. 1b–d; Cowart et al., 2006). However, an orange band was observed only in about half of the sponges investigated in this study. In addition to color loss, massive tissue destruction and erosion was observed (Fig. 1d), which usually leads to the collapse of the entire sponge. Only sponges in the early stage of disease were actively pumping, as judged visually by the application of fluorescent dye, while the water flow had ceased in sponges in the advanced stages of disease.

image

Figure 1.  Underwater photographs of representative Xestospongia muta individuals: healthy individual #9 (a) and individuals #4, #3 and #5 in advanced stages of disease (b, c, d). Underwater photography by Hilde Angermeier.

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Electronmicroscopical observations

Tissues from five healthy and five diseased X. muta were inspected by SEM. Representative images of one healthy and one diseased individual (Fig. 1d) are shown. SEM on the surfaces of healthy X. muta revealed characteristic features. The ostia are visible as opening canals and the spicules are readily identified as spines embedded within the extracellular matrix (Fig. 2a). The flattened pinacocyte cells form an incoherent layer covering large areas of the mesohyl matrix (Fig. 2b). In contrast, the outer layers of the white tissues of a diseased sponge are made up of spicule material and only very little biomass is left (Fig. 2c). In the enlargement, filaments resembling bacteria are found to cover the spicules (Fig. 2d).

image

Figure 2.  Scanning electron micrographs of the surfaces of healthy (a, b) and diseased (c, d) Xestospongia muta individuals. The scale bars represent 200 μm (a, c) and 20 μm (b, d). O, osculi; S, spicules; P, pinacocytes; Ac, aquiferous canals; F, filaments.

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Particular care was taken to visualize the cellular processes in the orange band transition zone. For this purpose, samples were compared from sponges collected in the Bahamas (one healthy and two diseased individuals investigated), which appeared to be in an early stage of disease, and from Florida (two healthy and three diseased individuals investigated), which showed more severe disease symptoms. In the Bahamas sponges, flattened pinacocytes were still visible on the surfaces of the orange band transition zone (Fig. 3a). The cyanobacteria were embedded close to the surface within the collagen matrix. TEM revealed large amounts of intracellular cyanobacteria that were apparently subject to digestion by sponge archaeocytes (Fig. 3b and c). By comparison, in the sponges from Florida, pinacocytes were entirely missing on the surface and the cyanobacteria were fully exposed (Fig. 3d). TEM revealed cellular destruction of nearly all cells inspected (Fig. 3e and f).

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Figure 3.  Scanning electron micrographs of the orange band transition zone of Xestospongia muta sponges collected from the Bahamas (a–c) and from Florida (d–f). P, pinacocytes; Cy, cyanobacteria; N, nucleus; S, spicules; C, collagen; and B, bacteria. The scale bars represent 10 μm (a), 5 μm (d) and 1 μm (b, c and e, f).

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Chlorophyll a analysis

The chlorophyll a value of healthy X. muta individuals was 32.1 μg g−1±4 (n=9) (Fig. 4). The chlorophyll a concentrations in diseased sponges decreased from brown tissues (32.9 μg g−1±11.2, n=2) to orange tissues (29.9 μg g−1±10, n=3) to white tissues (7.5±4.2 μg g−1, n=3).

image

Figure 4.  Chlorophyll a concentrations in tissues of healthy (n=9±SE) and diseased (n=3±SE) Xestospongia muta individuals.

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HPLC analysis

HPLC analysis was performed on two healthy and five diseased X. muta specimens. Representative chromatograms of one healthy (Fig. 5a) and one diseased sponge are shown (Fig. 5b–d). HPLC analysis of the healthy individual revealed five major peaks (A–E) (Fig. 5a). Five partially pure compounds were identified from these peaks based on UV/MS data and comparison with published X. muta metabolites. Peak A represented mutasterol (Rt 6.3 min) (Li et al., 1981), peak B mutafuran A, B, E or F (all of which have equal molecular masses) (Rt 6.9 min) (Morinaka et al., 2007), peak C mutafuran G (Rt 7.0 min) (Morinaka et al., 2007), peak D 18-bromo-octadecadiene-diyonic acid (Rt 7.3 min) (Patil et al., 1992) and peak E 5,28-stigmastadien-3β,24-diol (Rt 7.9 min) (Duque et al., 1985). The HPLC chromatogram of the brown tissue of the diseased sponge showed that the major peaks, including peaks A–E, were still present, albeit at lower concentrations (Fig. 5b). The peak profile was significantly reduced in orange tissues, leaving only a single peak visible (Fig. 5c), and peaks were entirely absent in white tissues of the diseased sponge (Fig. 5d).

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Figure 5.  HPLC chromatograms (at 220 nm) of methanolic extracts obtained from a healthy individual (a) as well as brown (b), orange (c) and white (d) tissues of a diseased Xestospongia muta individual (Fig. 1d). The letters A–E depict the identified compounds from the healthy sponge.

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DGGE

Inspection of the DGGE gel following amplification with cyanobacteria-specific primers revealed consistently few bands in the healthy sponge and in the brown and orange tissues of the five diseased sponges (Fig. 6a, Table S1). The DGGE banding patterns of the corresponding white tissues were much more heterogeneous. While the conspicuous DGGE bands from the healthy tissues were largely missing, a large number of new bands appeared that formed no consistent pattern. Altogether, 61 bands were excised from the cyanobacterial DGGE gel. Seven bands could not be reamplified with the primers 106f and 781r and were thus discarded. Up to four clones per DGGE band were sequenced. Four clones belonging to the phylum Verrucomicrobia as well as two chimeric sequences were removed from the dataset. Altogether, 58 cyanobacterial 16S rRNA gene sequences were obtained, of which one sequence was derived from healthy X. muta, while 17 sequences were derived from brown, 19 sequences from orange and 21 sequences from white tissues of diseased X. muta specimens.

image

Figure 6.  DGGE of cyanobacterial 16S rRNA genes in one healthy and five diseased Xestospongia muta individuals. Arrows point to the excised bands (a). UPGAMA cluster analysis of the cyanobacteria-specific DGGE gel banding pattern (b).

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The DGGE banding pattern was analyzed using the software program quantity one and a dendrogram was constructed using the clustering method UPGAMA (Fig. 6b). The DGGE lanes from white tissues formed one coherent cluster with similarities ranging from 36 to 56%. All DGGE lanes obtained from healthy sponge or from brown and orange tissues of diseased sponges formed another cluster with 56% similarity. Within this, two major groups were evident: one comprised of four DGGE lanes from brown tissues (71% similarity) and another one of DGGE lanes from orange tissues (82% similarity). The DGGE lane from the healthy sponge was placed most closely to the DGGE lanes from orange tissues.

Phylogenetic tree construction revealed that all 37 cyanobacterial sequences, which were derived from the healthy sponge as well as from the brown and orange tissues of the diseased sponges, form one coherent, monophyletic sequence cluster together with the previously known Synechococcus/Prochlorococcus clade of sponge symbionts (Fig. 7, Table S1). This cluster contains representatives of clades B and L Synechococcus spongiarum (Erwin & Thacker, 2008) and also of the vertically transmitted cyanobacterial 16S rRNA gene phylotypes described by Schmitt et al. (2008). From the white tissues, only four cyanobacterial DGGE sequences belonged to the cyanobacterial symbiont cluster (DGGE 28, DGGE 34, DGGE 51 and DGGE 61). The remaining 17 sequences from white tissues were affiliated with diverse cyanobacterial genera such as Limnothrix, Plectonema, Acaryochloris, Leptolyngbya and Myxosarcina. Their closest reference sequences were obtained from freshwater (DGGE 30), seawater (DGGE 57), hypolithic slime (DGGE 43), microbial mat (DGGE 36), rock oyster (DGGE 31), coral reef (DGGE 48) as well as coral sediment (DGGE 29, 33-3, 40), healthy corals (DGGE 33-1, 38, 44, 41), mucus of black band diseased (BBD) corals (DGGE 37) and BBD corals (DGGE 39) as well as from white syndrome affected dead coral skeleton (DGGE 33-2, 42).

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Figure 7.  Phylogenetic maximum likelihood tree with cyanobacterial 16S rRNA gene DGGE sequences and references. Sequences of this study are shown in bold. Full circles indicate bootstrap support≥90% and open circles bootstrap support≥75%. The arrow points towards the outgroup, Rhodothermus marinusAF217494. The scale bar indicates 10% sequence divergence. The gray box shows the monophyletic, sponge-specific, cyanobacterial symbiont cluster.

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Infection experiments

All of the sponges, into which tissue cores had been transplanted, responded in the same manner (Fig. 8). Within 11 days the tissues around the cores appeared to heal up. A rejection of the foreign material was not observed. Donor material had been taken from sponges in the early and late stages of disease but no difference in recipient sponge response was observed. Specifically, there was no difference in recipient sponge response to either diseased or healthy cores.

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Figure 8.  Underwater photography of the transplanted cores from a healthy individual (a) and from diseased sponges (b–d) into healthy Xestospongia muta recipients after 11 days of transplantation.

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Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Despite the careful characterization of SOB disease of the Caribbean barrel sponge X. muta, this study provided no evidence for the involvement of a specific microbial pathogen as an etiologic agent of disease, and attempts at infection with transplanted SOB tissue were not successful. SEM of SOB tissue revealed massive destruction of the pinacoderm accompanied by the loss of both chlorophyll a and characteristic secondary metabolites. Electronmicroscopical observations of SOB tissues in the early stage of the disease (Bahamas collection), as judged by the presence of disease symptoms, revealed a notably increased phagocytosis of cyanobacteria that indicates that the host was actively reacting to SOB progression. Tissue from SOB at later stages (Florida collection) exhibited largely degenerated cells. Active phagocytosis of symbiotic cyanobacteria was observed previously in diseased tissues of the mangrove sponge Geodia papyracea (Rützler, 1988) and in diseased pustules of Ircinia sp. (Maldonado et al., 2010). For cnidarians with algal symbionts, the intriguing hypothesis has been advanced that tissue bleaching is due to an overly aggressive innate immune response, with loss of control over apoptosis and necrosis being the cause of uncontrolled bleaching and ultimately death (Dunn et al., 2004; Weis, 2008). Unraveling the mechanisms of interaction between host phagocytes and microbial symbionts (i.e. recognition, differentiation between self and nonself) will clearly be instrumental to understanding sponge health and disease.

Microbial community profiling by DGGE revealed a distinct shift from the Synechococcus/Prochlorococcus clade of cyanobacterial sponge symbionts in normal sponge tissue towards a heterogeneous mix of cyanobacteria as SOB advances (Figs 6 and 7). Interestingly, several DGGE band sequences of bleached tissues were most closely related to sequences derived from healthy corals (DGGE bands 41, 44), coral mucus (DGGE band 37) or coral disease (DGGE bands 39, 42) (Table S1). The appearance of filament-forming cyanobacterial genera, Limnothrix, Plectonema, Leptolyngbya, in white tissues is consistent with the appearance of filaments in scanning electron microscopy of white X. muta tissues (Fig. 2d). Similarly, a shift from a stable bacterial DGGE banding pattern in healthy individuals towards a heterogeneous mixture of DGGE bands in brown, orange and white tissues of SOB sponges was observed (Fig. S2). Moreover, cultivation efforts from the orange band transition zone yielded bacterial isolates most closely related to Gammaproteobacteria from BBD-affected corals (data not shown). These data are consistent with previous observations of diseased tissue from the sponge Aplysina aerophoba in which shifts from the symbiotic microbiota towards a more diverse microbial consortium were documented (Webster et al., 2008b). Several of the pathogenic phylotypes identified in A. aerophoba were also most closely related to known BBD coral pathogens. Likewise, bleaching in X. muta leads to a disruption of the symbiotic, archaeal ammonia-oxidizing community that was replaced with sediment-derived phylotypes in the late stage of SOB (Lopez-Legentil et al., 2010).

Microbial population shifts away from the stable consortium are well known in coral diseases where the change even predates the bleaching event (Cooney et al., 2002; Frias-Lopez et al., 2002; Pantos et al., 2003; Gil-Agudelo et al., 2006; Bourne et al., 2008). Remarkably, the dominant phylotype during pre- and postbleaching of the coral Acropora millepora was a typically sponge-affiliated gammaproteobacterial clade (Spongiobacter sp.) (Bourne et al., 2008). It thus appears that disturbances of the natural microbiota of sponge and coral hosts are a common feature of disease and that certain opportunistic bacteria are then able to colonize the newly exposed, chemically undefended invertebrate tissues. Whether these bacteria are simply efficient colonizers or whether they actively contribute to the infection process remains to be investigated.

The absence of evidence for a specific pathogen in SOB disease of X. muta is noteworthy. Our findings are fully consistent with a recent publication by Luter et al. (2010) showing that microorganisms are not responsible for the formation of brown spot lesions in the sponge Ianthella basta. Perhaps most revealing in both studies is the unsuccessful infection of healthy individuals either at early or at late stages of disease (Fig. 8). A previous study also demonstrated that X. muta is able to heal wounds very rapidly (Walters & Pawlik, 2005), and this appears to be true whether the wounds are open or filled with exogenous sponge tissue. The infection experiments described in the present study are only the latest attempts at infection by tissue transplantation. Previously, wedges of SOB tissues at various stages of the disease were transplanted into several healthy sponges without inducing SOB in the recipient sponges (data not shown). Consequently, SOB of X. muta appears not to be due to the involvement of a microbial pathogen. However, an infectious agent, whether bacterial, fungal or viral, can also not be excluded. Moreover, the earliest window of an infection process may have been missed in this study and methods might be insufficient to identify a pathogen, especially if present at low abundances or if specific factors necessary for the expression of virulence genes were required.

An imbalance of the natural microbial consortia appears to be a hallmark of marine sponge and coral diseases. In this context, it is interesting to note that mostly species belonging to the HMA group of sponges including X. muta (Cowart et al., 2006), R. odorabile (Webster et al., 2002), I. basta (Cervino et al., 2006; Luter et al., 2010), Ircinia sp. (Maldonado et al., 2010), Aplysina cauliformis (Olson et al., 2006) and A. aerophoba (Webster et al., 2008b) are prone to disease. As sponge diseases are nearly always reported in the context of environmental stress, the hypothesis put forward by Lesser et al. (2007) that coral diseases are, with rare exceptions, opportunistic infections secondary to physiological stress (e.g. elevated seawater temperature) appears to be most plausible also in the context of X. muta SOB disease. Indeed, temperature has been identified as a major stress factor in sponges, causing increased heat shock protein gene hsp70 expression in X. muta at temperatures>30 °C (Lopez-Legentil et al., 2008) as well as causing tissue necrosis and dramatic shifts in the microbial community composition of R. odorabile>33 °C (Webster et al., 2008a). In conclusion, while microorganisms are clearly instrumental to sponge and coral diseases, we recognize their involvement in maintaining homeostasis in the natural host-associated microbiota rather than in their function as an invading pathogen in the clinical sense.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

We gratefully acknowledge the marine operations personnel of the National Undersea Research Center (University of North Carolina, Wilmington) in Key Largo, Florida, and the crew of the RV Seward Johnson II (HBOI) for excellent support during field work. We thank S.E. McMurray (NOAA/UNCW) and C. Finelli (UNCW) for professional assistance during field work as well as P. Tabares for guidance with the chemical studies. We acknowledge T. Smith (University of the Virgin Islands) and L. Siba (Misool Eco Resort, Indonesia) for sharing information on recent Xestospongia sp. disease outbreaks as well as F. Michels (UW-PSD.DE) for image editing of the underwater photographs. H.A. was supported by a grant of the German Excellence Initiative to the Graduate School of Life Sciences, University of Würzburg. Additional financial support was provided by the DFG, SFB567-TPC3, to U.H.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Fig. S1. Photography on the surface of a healthy Xestospongia muta individual (Fig. 1a, #9) and a diseased individual (Fig. 1b).

Fig. S2. DGGE of bacterial 16S rRNA genes in two healthy and five diseased Xestospongia muta individuals (a); UPGAMA cluster analysis of the eubacteria-specific DGGE gel banding pattern (b).

Table S1. 16S rRNA gene sequence analysis of the excised cyanobacterial DGGE bands from Fig. 6a.

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