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Ecological physiology of the black band disease cyanobacterium Phormidium corallyticum

Laurie L. Richardson , Kevin G. Kuta
DOI: http://dx.doi.org/10.1016/S0168-6496(03)00025-4 287-298 First published online: 1 April 2003


Laboratory studies were carried out to assess the photosynthetic and nitrogen-fixing capabilities of the gliding, filamentous cyanobacterium Phormidium corallyticum. This species is found on coral reefs, and is one of the members of a pathogenic microbial consortium called black band disease of corals, a unique horizontally migrating microbial mat with an active sulfuretum. It was determined that P. corallyticum can perform oxygenic photosynthesis in the presence or absence of sulfide, but cannot conduct (DCMU-forced) anoxygenic photosynthesis with sulfide as electron donor. Photosynthesis vs. irradiance curves revealed a very low threshold for Pmax of <30 μE m−2 s−1. Temperature optima for photosynthetic activity were at and above 30°C. Neither a laboratory culture of P. corallyticum nor freshly collected samples of the black band microbial consortium were capable of fixing N2. Results are discussed in terms of the ecology of this coral disease.

  • Cyanobacterium
  • Microenvironment
  • Black band disease
  • Microbial mat
  • Sulfide
  • Photosynthesis vs. irradiance curve
  • Photosynthesis
  • Phormidium corallyticum

1 Introduction

Black band disease of corals is caused by a pathogenic microbial consortium that exists as a horizontally migrating, laminated microbial mat [1]. The consortium is structurally directly analogous to cyanobacterial mats found in many illuminated, sulfide-rich benthic environments such as hot spring outflows and sediments of hypersaline lagoons, but is unique in that the entire mat community migrates across the surface of coral colonies completely degrading coral tissue (Fig. 1A). It is one of a number of coral diseases believed to play an important role in the observed decline of coral reefs [2].


A: In situ photomacrograph of black band disease on the coral Montastraea cavernosa. As the band (indicated as BBD) moves across the colony at rates up to 1 cm day−1, coral tissue is completely lysed, leaving behind exposed coral skeleton (indicated as ‘dead’). Healthy polyp diameter=8 mm. B: Photomicrograph of P. corallyticum taken from the culture used in laboratory experiments in this study. The characteristic pointed tip of one end of the filament, a key morphological feature used to identify this cyanobacterium, can be seen. Scale bar=50 μm.

The black band microbial consortium is dominated in terms of biomass by a population of gliding, filamentous, phycoerythrin-rich cyanobacteria originally identified as Oscillatoria submembranacaea [3]. It is the pigmentation of the cyanobacterial matrix that makes black band appear black. A subsequent study characterized and renamed the black band disease cyanobacterium as Phormidium corallyticum, a new species within a separate genus [4], and proposed that it is the primary pathogen of the disease. The designation of the new species included documentation of a distinctive morphology in which the 4.0×4.4-μm filaments exhibited one rounded tip and one pointed tip. Two recent molecular studies [5,6] of black band disease community composition were conducted in which 16S rDNA sequencing was performed and results compared to the GenBank data base. One of these studies found a single black band-associated cyanobacterium that was most closely related to Trichodesmium tenue (93% match) [6]. The other found two different cyanobacteria, Oscillatoria cf. corallinae (92%) and a planktonic freshwater strain (AF289160, 91%) [5]. None of these constitutes a generally accepted identification (98% or higher sequence homology). Neither study found a sequence associated with the genus Phormidium.

In addition to the dominant cyanobacterial population, the black band microbial mat contains numerous sulfate-reducing bacteria [58], at times dense populations of the gliding sulfide-oxidizing bacterium Beggiatoa spp. [711], and numerous heterotrophic bacteria [5,6]. While the functional roles of the sulfate-reducing and sulfide-oxidizing members are clear, the specific roles of the many heterotrophs are not known [5].

In a previous study using oxygen- and sulfide-sensitive microelectrodes [1] we found the interior of black band to be anaerobic and to contain variable levels of sulfide that depended on the light regime. We measured sulfide concentrations of up to 800 μM (produced by the population of sulfate reducers) using this approach. It was subsequently determined experimentally that exposure to anoxia with sulfide at concentrations comparable to those found in black band is lethal to corals [12], which we interpreted as a mechanism of black band-induced coral tissue death. We also demonstrated [1,13] that an interface of oxygen and sulfide migrates vertically within the band as a result of changes in oxygenic photosynthetic activity on the part of the black band cyanobacteria, a pattern directly analogous to that found in many other cyanobacterial mat communities [14].

In addition to the microelectrode studies, behavioral studies have been conducted to examine the vertical migrations of P. corallyticum and Beggiatoa within black band [10,11]. These studies demonstrated that, again similar to other microbial mats, there is a well-defined vertical migration pattern. However, the pattern is unusual in that Beggiatoa is often present on the surface of the band under aerobic conditions with moderate to high light levels [11]. We found that P. corallyticum moved to the surface of the band during low light, but was replaced by Beggiatoa when light intensity was greater than 190 μE m−2 s−1.

Together the black band microelectrode and motility studies demonstrated that members of the black band community are exposed to a range of microenvironments of varying oxygen and sulfide concentrations which fluctuate on a diel basis and range from anoxic and sulfide-rich to supersaturated with O2. This is of interest in terms of the physiological ecology of the black band microbial community because many microbial metabolic processes are flexible and can be triggered by changes in the chemical (and light) environment. Switching between metabolic modes has been shown to occur as a direct response to exposure to differing microenvironments as a result of vertical migrations of the oxygen/sulfide interface within sulfide-rich laminated cyanobacterial mats [15,16].

The basic physiology of P. corallyticum isolated from black band disease was first investigated by Taylor in 1983 [17]. He reported photoautotrophic growth in the laboratory with 20–22 h per generation, and dark (aerobic) heterotrophic growth at doubling times ranging from 3.4 to 14 days. The latter was supported by simple carbon compounds (ribose, glycerol, sucrose, mannitol, glucose and fructose). Both the photoautotrophic and heterotrophic growth rates are in the same range as those of other cyanobacterial species [18].

Determination of the range of physiologic capabilities of P. corallyticum (and other black band cyanobacteria) would be an important step in furthering our understanding of the etiology of this coral disease. In addition to enabling growth and survival within the highly variable chemical environment within the band, the potential metabolic modes of P. corallyticum may directly influence the band by producing or consuming both oxygen and sulfide. It is already known that P. corallyticum produces and consumes O2 during the metabolic processes of oxygenic photosynthesis and (dark) aerobic respiration. Sulfide use (as a possible photosynthetic electron donor) and/or tolerance by P. corallyticum might also influence the dramatic oxygen and sulfide dynamics within the band.

A second functional role of black band disease cyanobacteria is that of potential nitrogen fixation. It is well known that coral reefs are nitrogen-limited, and also well known that nitrogen fixation can be carried out by many non-heterocystous cyanobacteria [18,19]. The black band environment is, during the daytime, anaerobic at the base of the band. The entire band is fully anoxic at night when, at times, the oxygen/sulfide interface extends above the band surface into the water column [1]. These dynamics suggest the distinct possibility that nitrogen fixation by black band cyanobacteria can occur during periods of exposure to anoxia as seen with other non-heterocystous cyanobacteria [19]. The potential for this ecologically significant metabolic activity has been pointed out in earlier studies of the microbial ecology of black band disease [4]. The potential is also addressed by Frias-Lopez et al. [6] in terms of their finding of a cyanobacterial sequence from black band disease samples with 93% homology to the known marine N2-fixing cyanobacterium Trichodesmium. Thus in addition to roles in the sulfur and oxygen cycles within the black band microbial community, P. corallyticum and other black band cyanobacteria may have an important role in nitrogen cycling.

The objective of this research was to investigate the physiology of P. corallyticum and to assess how the metabolism of this species may contribute to the functioning of the pathogenic black band disease microbial consortium. Photosynthetic capabilities of P. corallyticum in response to variations in temperature, light intensity, and exposure to sulfide were investigated, as was the potential for use of sulfide to support anoxygenic photosynthesis and the potential for nitrogen fixation.

2 Materials and methods

2.1 Laboratory strain

The cultured strain used in all laboratory experiments was isolated as a single filament of P. corallyticum from a fresh sample of black band disease collected on Algae Reef in the northern Florida Keys (Fig. 2). Microscopic examination of this (and all other) black band samples revealed a predominance of cyanobacteria that matched the morphological description of P. corallyticum [4]. The isolate was obtained from a sample collected between 17.30 and 18.30 h on July 31, 1991 at a water depth of 7 m. Water temperature was 30.1°C. The isolation was performed using the gliding method of R. Castenholz [20] in which individual filaments are observed (using a dissecting microscope) to glide out from a point inoculum on an agar plate and then excised using sterile watchmaker's forceps. For our isolation we used agar plates made with ASNIII, and inoculated individual filaments into 80 ml of ASNIII (buffered with EPPS with the pH adjusted to 8.1) in 125-ml Erlenmeyer flasks. Successful growth of inoculated filaments occurred only when the flasks were wrapped in two layers of paper towels placed approximately 25 cm from a 30 μE m−2 s−1 light source. Unwrapped flasks never exhibited growth, which we interpret as due to photosensitivity. The culture was not axenic although repeated efforts to obtain a pure culture were implemented. The contaminants were small, Gram-negative bacteria that apparently live in (on on) the copious polysaccharide secreted by the cyanobacterium. When the culture had grown to form dense clumps, stock cultures (transferred on a weekly basis) were maintained at 30°C, on a 12:12-h light:dark cycle at a light intensity of 53 μE m−2 s−1. Culture medium (used both during experiments and for culture maintenance) consisted of ASNIII buffered to pH 8.1 with EPPS. The cultured P. corallyticum is shown in Fig. 1B.


Sites where black band was collected and used as the source of the laboratory culture of P. corallyticum (closed star), and where freshly collected black band mat was used in photosynthesis and nitrogen fixation experiments (closed and open stars). The closed star indicates Algae Reef, and the open star Grecian Rocks, both offshore of Key Largo, Florida.

2.2 Freshly collected black band for comparative laboratory experiments

In addition to laboratory experiments that used the clonal culture, experiments were conducted using freshly collected black band. For these experiments black band was collected from two sites, Algae Reef (maximum depth 8 m) and Grecian Rocks (maximum depth 2.5 m), both offshore of Key Largo, Florida. These sites are indicated in Fig. 2. Freshly collected material was used the next day for experiments on photosynthetic capability and/or nitrogen fixation. Collection was by sterile 60-ml syringes in which a mass of black band was removed. The sampling syringe was maintained in ambient sea water (in a cooler) in low light (the lid was open slightly), during which time samples formed a dense clump within the sample syringe due to the contraction of the matrix of cyanobacterial filaments. Collections for photosynthesis and nitrogen fixation experiments occurred throughout the months of July and August, 1996, between 10.00 and 14.00 h. Water temperature ranged from 29 to 30°C. For these experiments no effort was made to reisolate P. corallyticum and a dispersion of the entire black band community, which was present as a matrix of P. corallyticum with associated bacteria, was used.

2.3 Photosynthetic capabilities of P. corallyticum

Photosynthetic growth rates of laboratory cultures of P. corallyticum were measured under different experimental conditions by measuring the photo-incorporation of [14C]NaHCO3. Experiments were conducted to determine photosynthetic rates using the following three potential photosynthetic modes: (1) oxygenic photosynthesis (aerobic conditions); (2) photosynthesis with exposure to sulfide; and (3) ‘DCMU-forced’ anoxygenic photosynthesis. DCMU-forced anoxygenic photosynthesis occurs when DCMU (3-(3′,4′-dichlorophenyl)-1,1-dimethylurea), an inhibitor of electron transport in photosystem (PS) II [21], is used to block electron transport, and an alternative electron donor is provided for PSI. In our experiments sulfide was provided as a potential electron donor to enable anoxygenic photosynthesis.

Photosynthesis experiments were carried out using a photosynthetron, originally developed [22] for use in phytoplankton photophysiology experiments. A photosynthetron enables one to generate a P vs. I (photosynthesis vs. irradiance) curve in a single experiment by incubating multiple replicate samples at a range of light intensities. For our experiments we used triplicate experimental vials at each of 15 light intensities (n=45 simultaneous incubations). Neutral density (and black) filters provided a light regime that ranged from complete darkness to a maximum of 79.7 μE m−2 s−1 in 7.1% intervals, with triplicate incubation wells at each irradiance level. Light was provided by three cool white fluorescent (F15T12) bulbs with reflective fixtures (aluminum foil). All light measurements were made using a Biospherical Instruments QSL100 averaging scalar irradiance meter, which measures photosynthetically active radiation.

P vs. I curves were generated for a range of incubation temperatures. Each experiment measuring oxygenic photosynthesis was repeated three times at each of the following temperatures: 18, 20, 25, 28, 30, 35, 37, and 40°C. An additional experiment was carried out at 16°C (only one triplicate run). These temperatures extend beyond the high and low temperature extremes of coastal waters at the study reefs and were selected to examine the full temperature range for growth. Experiments aimed at investigating photosynthesis in the presence of sulfide and anoxygenic photosynthesis (DCMU and sulfide) were carried out (in triplicate) at fewer temperatures (20, 25, 30, and 35°C), selected from the data provided by the experiments that measured oxygenic photosynthesis. In all cases experimental temperatures were maintained by immersion of the photosynthetron in a flow-through water bath that consisted of a large aquarium. The light source shown through the glass walls of the aquarium.

To quantify oxygenic photosynthesis subsamples of P. corallyticum, from a culture that was freshly inoculated and incubated overnight, were added as 0.5 ml of a uniform suspension to replicate 20-ml scintillation vials (n=45). Each vial contained ASNIII to which NaCO3 and [14C]NaCO3 had been added to a specific activity of 0.05 μCi ml−1. Total volume was 6.0 ml. At the same time that experimental vials were inoculated, three additional vials that contained only ASNIII (no added label) were inoculated to determine the dry weight of the inoculum. Immediately after the experiment was initiated (see below) the contents of the three unlabeled (non-radioactive) vials were filtered onto preweighed filters, dried for 48 h at 60°C, weighed with pre-weight values subtracted, and averaged. After inoculation (under low light conditions) samples were incubated for 2 h in the photosynthetron. At the end of the incubation period, each sample was killed by adding formalin to a final concentration of 1.5%. Samples were then filtered onto 934-AH glass fiber filters and were rinsed with sterile ASNIII medium followed by a 2.0% HCl solution. Filters were placed in scintillation cocktail and 14C was measured using a Beckman LS 3801 scintillation counter (Beckman, USA). The amount of label incorporated was then expressed based on the dry weight of the inoculum. These methods were adapted from Castenholz and Utkilen [23].

Measurements of anoxygenic photosynthesis (DCMU-forced) and oxygenic photosynthesis during exposure to sulfide were performed using the same experimental protocol described above with the following changes. Anaerobic media for these experiments were prepared by bubbling ASNIII medium with reagent grade N2 gas for 30 min to strip oxygen from the liquid. The media were then stored in an anaerobic (Coy) hood until used. All solutions and materials were added to the vials under anaerobic conditions immediately prior to each experiment. Seven-milliliter scintillation vials with Hungate caps and septa were used in order to maintain anaerobic conditions. Sulfide was introduced through the septum to produce a final concentration of 0.5 mM (added from a stock solution of Na2S·9H2O). This concentration was chosen based on our direct measurements of sulfide in black band disease. Sulfide concentration was monitored in parallel (non-radioactive) incubations of P. corallyticum using the colorimetric Pachmayr assay calibrated with freshly prepared sulfide solutions of known sulfide concentration [24]. It was not necessary to add additional sulfide in order to maintain a sulfide concentration of 0.5 mM during the 2-h incubation period.

To investigate the potential for DCMU-forced anoxygenic photosynthesis, DCMU was added to a concentration of 10 μM (along with 0.5 mM sulfide). Cultures of P. corallyticum prepared for these experiments were exposed to 0.5 mM sulfide for 6 h prior to the experimental treatments to allow for sulfide adaptation [25]. As a control, in one experiment at each temperature DCMU was added (10 μM) but no alternative electron acceptor was provided.

In order to determine if the ability to carry out anoxygenic photosynthesis had been lost by the cultured P. corallyticum, additional experiments were performed using freshly collected black band samples under the following conditions: light, aerobic; light and sulfide (anaerobic); light, sulfide, and DCMU (anaerobic); light and DCMU (aerobic control); and dark, aerobic (additional control). In these experiments the concentrations of DCMU and sulfide were the same as detailed above. All experimental conditions were incubated simultaneously at a temperature of 30°C and at a light intensity of 79.7 μE m−2 s−1. This temperature and light intensity were selected based on the results of the oxygenic photosynthesis experiments, detailed below. The experiment using freshly collected black band was repeated three times, with three replicates of each condition in each experiment.

2.4 Nitrogen fixation

The ability to fix N2 was determined using the acetylene reduction technique [26], and was carried out with both cultured P. corallyticum and freshly collected black band disease samples. Three replicates were used for each of the following experimental conditions (all anaerobic): light; dark; light with sulfide; dark with sulfide; and light with DCMU. Anaerobic conditions were attained as described above, and DCMU and sulfide were used at the concentrations described above.

Prior to N2 fixation experiments, cultures of P. corallyticum were prepared that were nitrogen-limited. A series of 125-ml Erlenmeyer flasks containing 80 ml nitrogen-free ASNIII medium (prepared by omitting all nitrogen nutrient sources from the medium recipe) were inoculated sequentially each day (with overnight cultures) and observed for pigment loss. When pigments were lost, it was assumed that the cultures that were inoculated into the nitrogen-free medium 1 day later in the sequence were nitrogen-limited but not visibly stressed. These cultures were used for the N2 fixation assays. Viable but nitrogen-limited P. corallyticum (or freshly sampled black band not subjected to nitrogen limitation) was transferred into O2-free 60-ml serum vials, and acetylene was injected to a final concentration of 15% using a gas-tight syringe. Controls for each experimental condition (listed above) were run without the addition of P. corallyticum or freshly collected black band.

During incubation, fluorescent lighting (two F15T12 CW fluorescent tubes) provided a light intensity of 53.1 μE m−2 s−1. Dark incubations were double wrapped in aluminum foil. Temperature was maintained in a temperature-controlled incubation chamber at 30°C. Parallel incubations were monitored (using the Pachmayr assay, by sampling through the septum) to ensure that sulfide concentrations of 0.5 mM were maintained during the experiment. Sulfide additions were found to be unnecessary in order to maintain sulfide concentrations.

After an incubation period of 12 h, the gas phase was sampled by syringe through the septa using evacuated vacutainers. The gas phase was then analyzed using an HP 5890 gas chromatograph (Hewlett Packard, USA) equipped with a flame ionization detector and a GS Al (30 m×0.53 mm) column. The carrier gas was helium (6 ml min−1) with an oven temperature of 70°C for 2 min, then 70–100°C at 10°C min−1 (no hold), 100–200°C at 25°C min−1, and 200°C for 3 min. A splitless injector was used (injector and detector at 250°C).

3 Results

3.1 Photosynthetic capabilities of P. corallyticum

The maximum rate of oxygenic photosynthesis (Pmax) obtained by cultured P. corallyticum occurred under aerobic conditions at temperatures of 30°C, 35°C, and 37°C (Fig. 3A). No significant differences were detected between photosynthetic rates at these three temperatures (Table 1). The average Pmax was 6.96×10−3 mg C mg dry wt−1 h−1. At temperatures below 30°C (16, 18, 20, 25, and 28°C), Pmax values decreased along with each decrease in temperature (Fig. 3A). Pmax was also lower at 40°C, and was comparable to that at 18°C.


P vs. I curves under aerobic (A) and anaerobic, sulfide-rich (B) conditions at nine different temperatures. Anaerobic incubations were sparged with 100% N2 and contained 0.5 mM sulfide. Each curve represents the average of three experiments with three replicates within each experiment (each data point represents nine measurements). 100% light=79.7 μE m−2 s−1; light levels are in 7.1% (of 100%) increments. Error bars indicate SEM.

View this table:

Maximum photosynthetic rate (Pmax), measured as mg C mg dry wt−1 h−1 for photosynthesis under aerobic (top) and anaerobic, sulfide-rich (bottom) conditions at optimal growth temperatures (30°C, 35°C, and 37°C)

Temperature (°C)Pmax (10−3 mg C mg dry wt−1 h−1)Min (10−3 mg C mg dry wt−1 h−1)Max (10−3 mg C mg dry wt−1 h−1)S.E.M.
Anaerobic, 0.5 mM sulfide
  • Minimum and maximum rates, showing the S.E.M., are included.

Photosynthesis continued to occur under anaerobic conditions with exposure to 0.5 mM sulfide (including preadaptation). These experiments (Fig. 3B) were conducted at four temperatures (20, 25, 30, and 35°C). Under these conditions, Pmax occurred at 30°C and 35°C. There was no significant difference between the photosynthetic rates at these two temperatures (Table 1). The average Pmax was 6.62×10−3 mg C mg dry wt−1 h−1, slightly lower than that in the absence of sulfide. At the lower temperatures, Pmax values decreased similar to the oxygenic photosynthesis experiments.

To determine if the measured photosynthesis in the presence of sulfide (Fig. 3B) was due to oxygenic or anoxygenic photosynthesis (or both), experiments were performed to assess the capability of (DCMU-forced) anoxygenic photosynthesis with sulfide as electron donor. There was no detectable photosynthetic activity (14C incorporation) either in the presence of DCMU plus 0.5 mM sulfide (Fig. 4A), or in the control (DCMU exposure with no electron donor provided, Fig. 4B). These experiments were conducted at four temperatures (20, 25, 30, and 35°C), the same as those of the experiment shown in Fig. 3B.


Anoxygenic (DCMU-forced) photosynthesis experiments. Incubations were carried out similar to those of Fig. 3, except that 10 μM DCMU was added. A: DCMU plus 0.5 mM sulfide (N2-sparged). B: DCMU only (control).

3.2 Light requirements

Photosaturation (Pmax) occurred at a minimum light intensity of 27.9 μE m−2 s−1 (35% of the maximum light provided). This value was consistent for all experimental conditions in which photosynthesis was carried out (with and without sulfide present) and was independent of temperature. The compensation intensity (calculated) remained similarly constant at 16.8 μE m−2 s−1. Pmax was observed from the threshold through 100% of the light provided (79.7 μE m−2 s−1). Although the maximum light intensity provided is low, light saturation was reached by P. corallyticum under each experimental condition. No higher light intensities were investigated (see Section 4 concerning cyanobacterial clumping behavior and self-shading).

3.3 Comparison of cultured and freshly collected P. corallyticum

The photosynthetic capabilities of freshly collected black band disease, present as a matrix of P. corallyticum along with other members of the consortium, was measured for comparison with the culture (Fig. 5 and Table 2). When incubated at optimal temperature (30°C) and light intensity (79.7 μE m−2 s−1) photosynthetic activity in the absence of sulfide was slightly higher for the freshly collected black band samples than for the culture, however the difference was not statistically significant (P>0.05). When exposed to sulfide, however, the rate of photosynthetic activity of freshly collected black band was significantly higher than the photosynthetic rate of cultured material (P<0.05). Similar to the laboratory culture, freshly collected black band samples did not exhibit DCMU-forced anoxygenic photosynthesis with sulfide as electron donor. (Note: in Fig. 5 the data presented for the culture experiments were extracted from the experiments shown in Figs. 3 and 4 in which incubations were conducted at the same temperature of 30°C and light regime of 79.7 μE m−2 s−1 used for the experiments with freshly collected black band).


Comparison of the maximum photosynthetic rates of cultured P. corallyticum and freshly collected black band disease for the following conditions: light (aerobic); light, 0.5 mM sulfide (anaerobic); light, 0.5 mM sulfide, 10 μM DCMU (anaerobic); light, 10 μM DCMU (aerobic, control); and dark (aerobic, control). Each experiment was repeated three times with triplicate incubations. All experiments were conducted at 30°C at a light level of 79.7 μE m−2 s−1.

View this table:

Comparison of photosynthetic rates of cultured P. corallyticum and freshly collected black band mat at 30°C

TreatmentPhotosynthetic rate (10−3 mg C mg dry wt−1 h−1)Min (10−3 mg C mg dry wt−1 h−1)Max (10−3 mg C mg dry wt−1 h−1)S.E.M.
Freshly collected black band mat
Sulfide, DCMU0.120.0680.180.062
Cultured P. corallyticum
TreatmentPmaxa (10−3 mg C mg dry wt−1 h−1)Min (10−3 mg C mg dry wt−1 h−1)Max (10−3 mg C mg dry wt−1 h−1)S.E.M.
Sulfide, DCMU0.050.0460.250.015
  • Incubations were conducted under the following conditions: light (aerobic); light, 0.5 mM sulfide (anaerobic); light, 0.5 mM sulfide, 10 μM DCMU (anaerobic); light, 10 μM DCMU (aerobic, control); and dark (aerobic, control). Freshly collected black band mat (dominated by P. corallyticum) was obtained from active black band disease on the reef.

  • aThe Pmax values of the culture are expressed as the average of Pmax over all light intensities at 30°C.

3.4 Nitrogen fixation

Neither the laboratory culture of P. corallyticum nor freshly collected black band were found to fix N2 (i.e. there was no acetylene reduction) under the suite of environmentally relevant conditions tested (light, anaerobic; dark, anaerobic; light, anaerobic with sulfide; dark, anaerobic with sulfide; and light, anaerobic with DCMU). (See SEC2 for details of these experiments.)

4 Discussion

4.1 Photosynthetic capabilities of P. corallyticum

P. corallyticum can perform oxygenic photosynthesis in the presence and absence of sulfide, but cannot perform (DCMU-forced) anoxygenic photosynthesis with sulfide as electron donor. The maximum photosynthetic rates (measured at different temperatures) of cultured P. corallyticum occurred during oxygenic photosynthesis in the absence of sulfide (Fig. 3A). The average rate (6.96×10−3 mg C mg dry wt−1 h−1) was attained at temperatures at and above 30°C and was only slightly higher than the oxygenic photosynthetic rate (6.62×10−3 mg C mg dry wt−1 h−1) attained during exposure (including preadaptation) to 0.5 mM sulfide. The difference was not statistically significant (P>0.05). Pmax during exposure to sulfide similarly occurred at temperatures at and above 30°C (Fig. 3B).

The same photosynthetic capabilities were present using black band samples freshly collected on the reef. The only difference was an elevated Pmax (for the freshly collected sample) under both sulfide-exposed and non-exposed conditions (Fig. 5 and Table 2). Under these conditions Pmax values were increased to >8×10−3 mg C mg dry wt−1 h−1. The enhanced rate may be due to the fact that the cultured P. corallyticum had been maintained in a sulfide-free environment in the laboratory for a period of 5 years before these experiments were conducted.

No detectable photosynthesis occurred when either cultured P. corallyticum or freshly collected black band was exposed to 0.5 mM sulfide along with 10 μM DCMU. This indicates, first, that the cultured P. corallyticum is incapable of anoxygenic photosynthesis with sulfide as an electron donor to PSI, and second, that the negative results obtained using the culture were not due to a loss of this metabolic capacity after 5 years of maintenance under aerobic laboratory conditions. The freshly collected black band sample was not manipulated to produce new unialgal cultures, therefore other cyanobacterial black band disease members may have been present in small numbers. If other phototrophs present in the freshly collected black band used in these experiments were capable of DCMU-forced anoxygenic photosynthesis, it was not detected in our experiments.

Our experiments revealed that P. corallyticum isolated from black band is capable of sulfide-resistant oxygenic photosynthesis with both PSII and PSI functioning normally, as described by Cohen et al. [27], and that this phototrophic mode was not inhibited (i.e. the rate was not decreased) by the presence of sulfide. This capability is quite uncommon. Most photosynthetic organisms, cyanobacteria included, are sulfide-intolerant (meaning that any level of sulfide permanently damages the photosynthetic capacity of the organism) [27].

There are four metabolic strategies by which cyanobacteria photosynthesize in sulfide-rich environments. These are: (1) sulfide-sensitive oxygenic photosynthesis, (2) sulfide-resistant oxygenic photosynthesis, (3) sulfide-insensitive oxygenic photosynthesis concurrent with sulfide-dependent anoxygenic photosynthesis; and (4) sulfide-sensitive oxygenic photosynthesis replaced by sulfide-dependent (anoxygenic) photosynthesis [18,25,2733]. Cyanobacteria with the first strategy are not able to tolerate sulfide at any significant level for extended periods. These cyanobacteria are unable to use sulfide in anoxygenic photosynthesis, and have impaired oxygenic photosynthesis, although some are capable of survival. This is the most common reaction to sulfide in cyanobacteria [27]. The second adaptation to sulfide allows continued oxygenic photosynthesis in the presence of sulfide in which PSII has a resistance to sulfide toxicity that enables complete or partial functioning of both photosystems [27]. This is less common than sulfide sensitivity. The third strategy consists of oxygenic photosynthesis that occurs concurrently with anoxygenic photosynthesis in that electrons are donated to both PSI and PSII. This adaptation is the most versatile mode of sulfide adaptation as it allows these cyanobacteria to function efficiently in environments with fluctuating sulfide levels [27]. It is believed that most mat-building cyanobacteria have this type of adaptation [27]. The final strategy is that of a sulfide-intolerant PSII (it is shut off) and a very efficient use of anoxygenic photosynthesis with sulfide as an electron donor to PSI. This adaptation is best suited to environments with prolonged exposure to high sulfide levels. Very few cyanobacteria have been shown to have this ability [25,27]. We have demonstrated that P. corallyticum utilizes the second form of sulfide tolerance, that of continued oxygenic photosynthesis in the presence of sulfide.

This physiological capability is most likely critical for survival within the black band disease microbial consortium. As discussed previously, the microenvironment in the black band microbial mat is routinely sulfide-rich and anoxic [1]. This characteristic is integrally important to black band disease activity as the combination of anoxia and sulfide is lethal to coral [12]. Of further benefit, this environment may protect P. corallyticum from competition from other photosynthetic organisms, including other cyanobacterial species, which are unable to tolerate sulfide.

4.2 Temperature and photosynthetic activity in black band disease

The rate of photosynthesis by P. corallyticum was strongly dependent on temperature (Fig. 3). Pmax was attained at temperatures of 30, 35, and 37°C during oxygenic photosynthesis in the absence of sulfide, and at 30 and 35°C in the presence of sulfide (the experiment was not performed at 37°C). This optimal temperature range agrees with the results of Taylor [17] who documented maximal growth rates of P. corallyticum at 30 and 32°C. (Higher temperatures were not tested.)

At temperatures between 20 and 30°C (20, 25, and 28°C) there was still substantial photosynthetic activity (Fig. 3). At 18 and 40°C photosynthesis was still measurable, but much lower (Pmax was <2×10−3 mg C mg dry wt−1 h−1). At the lowest temperature tested, 16°C, there was no detectable photosynthetic incorporation of CO2. The overall pattern of photosynthesis in response to temperature was the same during the presence or absence of sulfide.

Our laboratory results are in agreement with previous reports of field studies which demonstrated that black band disease is seasonal on reefs within higher tropical latitudes. It is commonly reported that black band disease is most active during the warm months of the year when water temperature is above 28°C. It is also reported that black band, although less prevalent, has often been observed between 25 and 28°C [3,17]. Our data are in agreement with these results in that we found maximal photosynthesis above 28°C, and moderate photosynthesis at both 25 and 28°C. Our data also support the numerous reports that black band disease is relatively inactive at temperatures less than 25°C [9]. As such our findings support the hypothesis that water temperature (as opposed to, for example, light intensity) is one of the most important factors in the seasonality of black band disease. However, temperature may not be the only factor involved in disease seasonality. We found that P. corallyticum is capable of active photosynthesis at 18 and 20°C albeit at reduced rates. Temperatures below 20°C are quite rare on reefs in the Florida Keys during the coldest months of the year, yet they do occur [34]. Thus our laboratory results are in agreement with our earlier report of active black band disease on one reef in Florida during winter months at a temperature of 19.6°C [35].

4.3 Nitrogen fixation

We found no detectable nitrogen fixation (acetylene reduction) using either the laboratory culture of P. corallyticum or freshly collected black band. This result is of importance because the presence of dense populations of cyanobacteria in anaerobic microenvironments found on nitrogen-limited coral reefs immediately raises the question of the potential for a new source of fixed nitrogen for the reef community. As mentioned above, many non-heterocystous cyanobacteria have the potential for nitrogen fixation, however this potential is not always expressed. Zehr and Capone [36] clearly point out that even when nitrogenase (nif), and in particular the even more highly conserved dinitrogenase reductase (nifH), gene sequences are detected in natural samples using molecular (sequence) techniques, only the potential for nitrogen fixation has been demonstrated. They emphasize that the demonstration of nitrogen fixation must include use of an assay for the actual expression of nitrogenase activity.

The recent molecular studies of the microbial diversity of the black band microbial consortium revealed more than 500 bacterial sequences associated with 12 bacterial divisions [5,6]. Since PCR amplification was used, however, many of these may be present in small numbers and not be of importance to the overall functioning of black band disease. Thus, some of these may very well be nitrogen fixers, however present in numbers too low to be detectable when assaying nitrogen fixation (acetylene reduction) by the entire band community. One of these studies [6] reported a single cyanobacterial sequence in two of three black band samples analyzed. This sequence was 93% homologous to that of GenBank accession number AF013029, which corresponds to the cyanobacterium T. tenue. These authors conclude that if present in the anaerobic black band, this species would actively fix nitrogen supported by photosynthesis. However, no assays were conducted to demonstrate nitrogen fixation activity.

4.4 Physiological ecology of P. corallyticum in the black band disease microbial consortium

The temperature range that supports optimum photosynthetic growth of P. corallyticum is in close agreement with the reported seasonality of black band disease on coral reefs. This species, like most cyanobacteria that live in temperate and subtropical zones, exhibits increased growth rates as water temperature increases over a range that is naturally occurring in its environment. A water temperature on Florida's reefs at and above 30°C is common during late summer and early fall, and it is exactly these months which correspond to the most active black band season. Our results are also in agreement with the limited data available, to date, concerning temperature optima of the other known and isolated microbial pathogens of coral that have been studied in the laboratory. These include Vibrio shiloi, a bacterium known to cause coral bleaching [37,38]; Aspergillus sydowii, which causes aspergillosis of gorgonians (soft corals) [39]; and Aurantimonas coralicida, the bacterial pathogen of the coral disease plague type II [40]. Each of these coral pathogens has a temperature optimum at or above 30°C.

Less straightforward, but probably very relevant, is the light requirement of P. corallyticum. We have shown that this species attains Pmax at very low light levels. This is fairly common for cyanobacteria [41], and may be an integral part of the development of the black band microbial mat. Many cyanobacteria, including our culture of P. corallyticum, perform ‘clumping behavior’, in which cells glide together to form clumps [20]. This is a self-shading behavior that occurs in response to high light conditions in the natural environment [42], and has been shown to result in the attainment of a light regime within the clumps that is ideal for the support of optimal photosynthesis by the clumping cyanobacteria [23]. In terms of black band disease, this behavior may be critical in the transition between an aerobic biofilm of filaments of P. corallyticum (the only reported natural reservoir of a black band disease cyanobacterium [43]) and the anaerobic black band microbial mat. Black band disease almost always starts at the top or on sides of coral colonies, expanding in a ring-shaped pattern. The initiation point coincides with the presence of sediment patches in depressions on top (or on the side) of colonies, on which cyanobacterial biofilms are observed to develop ([43]; H. Hudson, personal communication). Although it is not known how black band becomes active, it is possible that as the biomass of P. corallyticum and other microorganisms in the biofilm increases via growth, the physical concentration of cells may be enhanced by cyanobacterial clumping in response to light. Cellular concentration would enhance the development of anaerobic microzones within the biofilm. As these microzones develop, the continued presence of anoxic zones would lead to enrichment for sulfate reducers, which would then allow for an accumulation of sulfide leading to stable anaerobic and sulfide-rich zones. Exposure of the adjacent coral tissue to this environment would result in coral tissue death. An alternative scenario would be that as the population increases in concentration the development of an anaerobic microzone would allow nitrogen fixation by non-heterocystous black band cyanobacteria. This new source of fixed nitrogen in the nitrogen-limited reef environment would then potentially allow for the rapid development of a microbial community, including the population of sulfate reducers. The ability to fix nitrogen has been tested (this study) for only one of the black band cyanobacteria. In any event, after the transition to anoxia, and by some as yet unknown mechanism that most likely involves chemotaxis, the new microbial mat community would begin to actively migrate and lyse coral tissue, and the transition to black band disease would be complete.

Our study was focused on assessing the physiology of one member of the black band community that is widely cited as the primary pathogen and that we routinely find in black band on reefs of the Florida Keys. Our laboratory strain of P. corallyticum was, as discussed above, isolated and identified based on the morphological description of this species put forth by Rützler and Santavy [4]. This culture was sequenced and included in the study of Cooney et al. [5] and was found to be most closely matched (97% homology) to an entirely different cyanobacterial genus (Geitlerinema sp., PCC 7105; AF132780). This strain was not found in the molecular analyses of black band conducted by either Frias-Lopez et al. [6] or Cooney et al. [5]. It is intriguing to note, in this context, that in the study of Frias-Lopez et al. [6] one of the three black band samples analyzed contained no cyanobacteria, even though the paper states that each of the black band samples was dominated by filamentous cyanobacteria that were morphologically similar to P. corallyticum [6].

It is our belief that the entire highly structured black band community functions as a pathogenic consortium. As such, there may be no individual primary pathogen. To understand this highly complex and fascinating community in terms of the roles of individual members, future work will require the use of an integrated approach that includes both molecular and physiologic techniques [5,13]. In particular more studies using fluorescent in situ hybridization with metabolic-specific as well as taxonomic-specific molecular probes should be carried out to assess the relative numbers and metabolic importance of the more than 500 bacteria currently detected in this consortium. Such an approach would ideally be coupled with confocal laser microscopy to view the relative abundance of different species and metabolic potential in the different laminations of the band.


We are grateful to the staff of the Florida Keys National Marine Sanctuary for providing boat support. The comments of two anonymous reviewers greatly improved the manuscript. This research was supported by the National Oceanographic and Atmospheric Administration's National Undersea Research Program administered by the University of North Carolina at Wilmington (Grant UNCW9416) and by the Environmental Protection Agency Region 4 (Grant X-984298-97-0). This is contribution number 55 from the Tropical Biology Program at Florida International University.


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View Abstract