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Research Article
31 January 2018

A New Niche for Anoxygenic Phototrophs as Endoliths


Anoxygenic phototrophic bacteria (APBs) occur in a wide range of aquatic habitats, from hot springs to freshwater lakes and intertidal microbial mats. Here, we report the discovery of a novel niche for APBs: endoliths within marine littoral carbonates. In a study of 40 locations around Isla de Mona, Puerto Rico, and Menorca, Spain, 16S rRNA high-throughput sequencing of endolithic community DNA revealed the presence of abundant phylotypes potentially belonging to well-known APB clades. An ad hoc phylogenetic classification of these sequences enabled us to refine the assignments more stringently. Even then, all locations contained such putative APBs, often reaching a significant proportion of all phototrophic sequences. In fact, in some 20% of samples, their contribution exceeded that of oxygenic phototrophs, previously regarded as the major type of endolithic microbe in carbonates. The communities contained representatives of APBs in the Chloroflexales, various proteobacterial groups, and Chlorobi. The most abundant phylotypes varied with geography: on Isla de Mona, Roseiflexus and Chlorothrix-related phylotypes dominated, whereas those related to Erythrobacter were the most common in Menorca. The presence of active populations of APBs was corroborated through an analysis of photopigments: bacteriochlorophylls were detected in all samples, bacteriochlorophyll c and a being most abundant. We discuss the potential metabolism and geomicrobial roles of endolithic APBs. Phylogenetic inference suggests that APBs may be playing a role as photoheterotrophs, adding biogeochemical complexity to our understanding of such communities. Given the global extent of coastal carbonate platforms, they likely represent a very large and unexplored habitat for APBs.
IMPORTANCE Endolithic microbial communities from carbonates, which have been explored for over 2 centuries in predominantly naturalistic studies, were thought to be primarily composed of eukaryotic algae and cyanobacteria. Our report represents a paradigm shift in this regard, at least for the marine environment, demonstrating the presence of ubiquitous and abundant populations of APBs in this habitat. It raises questions about the role of these organisms in the geological dynamics of coastal carbonates, including coral reefs.


Over the past two hundred years, naturalists have extensively studied the endolithic habitat within intertidal carbonates. Evidence from as early as the mid-1800s from Agassiz, reported by Duerden (1) and Kölliker (2) describe the presence of vegetable parasites within mollusk shells and corals. These descriptions eventually extended to a range of substrates and settings, including marine carbonates (37), terrestrial limestones and marbles (8), corals (9, 10), and microbialites (11). Much of the work focused on the boring algae and cyanobacteria, known as euendoliths, which can actively penetrate the carbonate substrate to establish a home within the solid rock. The crypto- and chasmoendoliths, which colonize pore spaces and cracks (12), respectively, have received less attention but are undoubtedly common. Endolithic communities play significant bioerosive roles in the natural environment (13, 14), can become pests of bivalve fisheries (1517), and judging by the presence of microfossils in the rock record, have been active in their roles since the Precambrian (18, 19).
The deployment of early molecular methods for community fingerprinting (clone libraries, denaturing gradient gel electrophoresis [DGGE]) provided expanded accounts of marine and terrestrial endolithic communities of carbonates as well as other substrates (7, 2022). They revealed that the endolithic habitat can harbor complex communities of microbes, with important heterotrophic components, particularly when the substrate rock is naturally porous or when it has been made porous through excavation by euendoliths. This level of complexity was clear in the first high-throughput multisample survey of community diversity from intertidal outcrops (23), which we conducted on Isla de Mona, Puerto Rico (PR). Results of that survey made two things apparent: the diversity and complexity of these communities had been dramatically underreported in the literature, and they could host a potentially wide range of metabolic niches previously unrecognized in this environment (23). The level of diversity found in endolithic communities was comparable to that of other microbial communities such as biological soil crusts (24) and microbialites (25), containing representatives of a variety of microbial metabolisms, from fermenters to sulfur oxidizers. Among these, one could discern many phylotypes potentially allied with anoxygenic phototrophic bacteria (APBs).
Anoxygenic phototrophic bacteria are a phylogenetically widespread metabolic guild distributed among six different bacterial phyla. Canonically, APBs have been delineated into five groups: the green sulfur bacteria in the phylum Chlorobi, the green nonsulfur bacteria of the phylum Chloroflexi, the purple sulfur bacteria of the Gammaproteobacteria, the purple nonsulfur bacteria of the Alphaproteobacteria and Betaproteobacteria, and the Heliobacteria within the Firmicutes (26). Over the past 2 decades, new APBs have been discovered, including the aerobic anoxygenic phototrophic bacteria found within the Alphaproteobacteria and Betaproteobacteria (27), along with one representative each in the Acidobacteria (28) and the Gemmatimonadetes (29). Some groups of APBs are typically found within a narrow range of habitats: green and purple sulfur bacteria are limited to locations in which there is a ready supply of an electron donor, typically hydrogen sulfide, such as at the anoxic bottom of meromictic lakes, ocean sediments, hot springs (3032), microbial mats (33), stratified marine and estuarine waters (34, 35), and subglacial lakes (36). Green nonsulfur bacteria have been found in hot springs, hypersaline microbial mats, and some marine sediments (37). Purple nonsulfur bacteria occur in a wider range of habitats, including the open ocean and Antarctic lakes, but they rarely are the dominant phototroph within a community (38). Here we report that the endolithic habitat must be now added to this short list.


Incidence and composition of endolithic APBs.

Our molecular study, which included 30 discrete intertidal carbonate outcrop samples obtained around Isla de Mona, PR, and 11 obtained around Menorca, Spain (Fig. 1), showed that APBs were abundantly and universally present in this environment. Sequence reads belonging to operational taxonomic units (OTUs) that were classifiable as likely phototrophs (oxygenic or anoxygenic) accounted for a large percentage of the total, on average 21%. As expected from the literature, oxygenic phototrophs (largely cyanobacteria, but also some algae) were either dominant or very well represented (Fig. 2). A significant proportion of phototrophic sequence reads could be classified as belonging to one of several groups of APBs, according to our stringent phylogenetic placement criteria (OTU sequence was placed within a clade formed exclusively by known cultivated phototrophs with >70% confidence; see Materials and Methods). Such APB sequences were found in every sample, ranging from 0.8% to 89% of total phototrophs. On average, they accounted for 30% of the total phototroph count. In close to one-fifth of the samples, APBs were more abundant than oxygenic phototrophs.
FIG 1 (a and b) Sampling locations (yellow dots) in Isla de Mona (a) and Menorca (b), chosen primarily by accessibility and to encompass a variety of substrate mineral compositions. (c) Isla de Mona limestone/dolostone cliffs, with characteristic intertidal notch (box). (d) Limestone cliffs from Menorca showing intertidal notch (box). (e) Close up of typical Menorca sampling site, showing chipped-off area with a pale coloration. (f) A sample rock chip of Isla de Mona raised reef, showing green endolithic growth below the surface.
FIG 2 Distribution of 16S rRNA gene reads (left) among phototrophic bacterial groups in endolithic communities from Isla de Mona and Menorca, based on phylogenetic taxon assignment, and the corresponding mineralogical compositions of their substrates based on XRD analyses (right). Sample identifiers are to the very left, their initial capital letters corresponding to the sites of origin in Fig. 1.
Phylogenetic analyses at a family-level resolution of these APBs revealed that two distinct groups were both common and abundant: the Chloroflexaceae (green nonsulfur bacteria) and the Erythrobacteraceae (in the Alphaproteobacteria). Other groups that were well represented included the Chlorobiaceae, various purple nonsulfur proteobacteria, and the Chromatiaceae in the Gammaproteobacteria. Samples from Isla de Mona supported relatively larger populations of Chloroflexaceae, while the endolithic microbiome in Menorca presented a larger relative abundance of Erythrobacteraceae. While most samples were taken in the intertidal zone, for a few samples in Isla de Mona that were collected in the subtidal zone (3.5-, 4.6-, and 9.1-m deep), we detected a large relative abundance of Chlorobiaceae and Chromatiaceae, which were absent or very rare in intertidal samples. At this level of phylogenetic resolution, we could not detect any obvious influence of the mineralogical composition of the substrate (which included calcite, aragonite, dolomite, and apatite) on the APB community composition (Fig. 2). No phototrophic representatives of the Betaproteobacteria, the Heliobacteria, the Acidobacteria, or the Gemmatimonadetes were detected.

Major endolithic APB phylotypes.

Interestingly, only a few OTUs (defined as groups of sequences that are 97% self-similar) made up the vast majority of all sequences attributable to APBs (Table 1). Two of these OTUs were by far the most abundant: OTU 31154957, a representative of the genus Roseiflexus (39), and OTU 582344, a representative of Erythrobacter sp. strain NAP1 (40). Both were present in all samples. Other OTUs in these two families were also quite important. In some samples, we found that OTU 112750, attributable to Prosthecochloris in the Chlorobi, was dominant in relative abundance among the APB phylotypes.
TABLE 1 Major putative APB OTUs within intertidal carbonates
OTUClassificationa% of total phototrophic sequences
Isla de MonaMenorca
3114957Roseiflexus sp.12.490.71
NROTU1Chlorothrix sp.8.26 
582344Erythrobacter sp. strain NAP10.4729.05
NROTU6Erythrobacter sp. strain NAP10.496.48
112750bProsthecochloris sp.2.17 
Classification assigned by placement in phylogenetic tree.
Present in one sample as 57.57% of the total phototrophic sequences.

Differential distribution with geography.

The most abundant APB phylotypes throughout Isla de Mona intertidal samples were members of the genera Roseiflexus, Chlorothrix, and Chloroflexus in the phylum Chloroflexi (Fig. 3). In stark contrast, the most abundant APBs in Menorca samples were the aerobic APB Erythrobacter NAP1. Yet, the latter phylotype could also be found in Isla de Mona, albeit making up a small contribution to the APB guild, and conversely, the Chloroflexi OTUs that dominated communities in Isla de Mona also contributed to the communities of Menorca. Additionally, a similar differential incidence could be detected for the purple nonsulfur members of the Rhodovulum and Rubrimonas, the latter being more abundant in Isla de Mona and the former in Menorca.
FIG 3 Phylogenetic distribution of endolithic APB phylotypes detected in marine carbonates on Isla de Mona and Menorca. Phyla in the bacterial phylogenetic tree (upper left) known to contain phototrophs are shown in color. Detailed subtrees for each of such phyla that found APB representatives in our survey are shown as enlargements. Circles of variable area to the right of individual clades in these subtrees represent the average % of total phototrophic sequences assignable to the clade in Isla de Mona (orange) or Menorca (blue). All trees were constructed using maximum likelihood algorithms.

Endolithic morphological diversity.

Confocal microscopy using autofluorescence in the visible and near-infrared (NIR) spectra of preparations in which the carbonate substrate had been dissolved away (41) was used in an attempt to visually confirm the presence of APBs and to gauge morphological and pigmentation diversity. Communities showed a range of morphological diversity. At one end, some samples displayed apparently monospecific beds of filamentous cyanobacteria, with abundant chlorophyll (Chl) a and little NIR fluorescence (Fig. 4a). However, other samples contained heterogenous mixtures of unicells, clusters, and thin filaments, with various fluorescence profiles, including cells that were only fluorescent in the NIR range (clearly APBs) (Fig. 4b) and cells that fluoresced in both the NIR and visible ranges, which probably correspond to chlorophyll d- and f-containing cyanobacteria.
FIG 4 Confocal laser scanning microscopy images of endolith communities after dissolution of their carbonate substrates. Chips were dissolved to liberate the biomass from the surrounding mineral and then imaged with a 405-nm laser. Chl a emission (false color red channel) was measured between 660 and 690 nm and near-infrared emission (false color green channel) was captured in the 740- to 800-nm range. Images show a wide diversity of morphotypes within and between samples, ranging from virtually monotypic beds of red-fluorescing cyanobacteria (a, likely Mastigocoleus), to diverse assemblages of unicellular, colonial, and filamentous types showing both red and NIR emissions (b).

Photosynthetic pigments in endolithic communities.

To confirm beyond the molecular signatures and confocal microscopy if active anoxygenic phototrophy was occurring within the endolithic microbiome, we analyzed lipid-soluble extracts by high-performance liquid chromatography (HPLC) with online diode array spectroscopy detection to isolate and identify major photopigments. As expected, we found concentrations of Chl a within the typical range that had been determined elsewhere in these systems (42, 43). But, we also found bacteriochlorophylls (Bchls) (Table 2; see also Table S1 in the supplemental material): Bchl c (as various isomers [44]) was detected in every sample, Bchl a was detected in eight samples from Isla de Mona and in all samples from Menorca, and Bchl d was detected in two samples from Isla de Mona and four samples from Menorca. Bacteriochlorophyll concentrations were generally much higher in samples from Isla de Mona than in Menorca. The average ratio of Chl a to total Bchl on Isla de Mona was 1.9 (Table 2), while on Menorca, the ratio was one order of magnitude higher, at 11.03 (Table 2). Interestingly, chlorophylls b, d, and f were also detected in low quantities in samples from both Isla de Mona and Menorca (Table S1).
TABLE 2 Detected pigments from intertidal carbonates
PigmentIsla de Mona (n = 29)Menorca (n = 11)Total (n = 40)
No. (%) of samples detectedAvg concn (mg/m2)Concn range (mg/m2)No. (%) of samples detectedAvg concn (mg/m2)Concn range (mg/m2)No. (%) of samples detectedAvg concn (mg/m2)Concn range (mg/m2)
Chl a29 (100)8.950.11–52.5611 (100)10.140.71–21.9540 (100)9.280.11–52.56
Chl b6 (21)0.040.00–0.2711 (100)0.580.01–1.6917 (43)0.190.00–1.69
Chl d3 (10)0.010.00–0.1611 (100)0.120.01–0.5614 (35)0.040.00–0.56
Chl f3 (10)0.010.00–0.229 (82)0.190.00–0.6112 (30)0.060.00–0.61
BChl a8 (28)<0.010.00–<0.0111 (100)0.530.01–1.3319 (48)0.150.00–1.33
BChl c29 (100)8.580.14–41.6611 (100)1.930.02–8.1940 (100)6.750.02–41.66
Bchl d2 (7)<0.010.00–0.124 (36)0.030.00–0.146 (14)0.010.00–0.14


The prevalence of bacterial families associated with anoxygenic phototrophy within the endolithic communities became evident upon close examination. Admittedly, however, assigning metabolism to phylotypes on the basis of automated taxonomic assignment, with often poorly curated databases, carries some uncertainties. To make our assignments stringent, we constructed our own curated databases and phylogenetic trees (available at and counted as “phototrophs” only OTUs that would fall within clades formed exclusively by known cultivated phototrophs with >70% confidence. This strict assignment, in fact, likely led to an underestimation of the relative abundance and number of phylotypes of APBs, in that we could have excluded any APBs that were close to but not within APB clades and obviously could not have detected any APBs with no known cultured representatives. However, while likely conservative, our results give us confidence in our finding that APBs are indeed a widespread and significant component of endolithic communities. The fact that this component could have been missed during almost 2 centuries of research is perplexing. It is possible that the spectral overlap of some chlorophylls and bacteriochlorophylls in extracted forms may have disguised these biomarkers (45). Perhaps the shared morphological characteristics of small thin filamentous cyanobacteria, such as Halomicronema (46) or Plectonema terebrans (15) and Chloroflexi (47), rendered them hard to discern under the microscope. And yet, an ad hoc literature review returned some corroborating evidence for the presence of APBs in endolithic communities from coral skeletons: spectroscopy revealed absorption peaks in the IR range, attributable to the presence of bacteriochlorophylls (48), and Yang et al. (49) report directly the presence of populations of Prosthecochloris spp. (Chlorobi).
Bacteriochlorophylls are diagnostic biomarkers for APBs, as they are integral to the reaction centers and antenna complexes at the core of their phototrophic capacity (50). We first examined samples using confocal microscopy, looking for the characteristic profile of near-infrared fluorescence associated with APBs. Though many morphotypes contained various levels of NIR fluorescence, this evidence was insufficient, in that it could also be attributed to the tail of Chl d or Chl f fluorescence. Still, some cells were exclusively fluorescent in the NIR range, indicating an abundant presence of bacteriochlorophylls. However, HPLC pigment composition analysis offered a more direct way of identification, showing beyond doubt their presence in all samples. Consistent with the dominance of Chloroflexi in Isla de Mona, Bchl c, a characteristic photopigment of the Chloroflexi (37), was by far the most abundant bacteriochlorophyll present. Conversely, Bchl a, which is characteristic of the Erythrobacteraceae (51), was more abundant in Menorca, consistent with the dominance by Erythrobacter. The detection of Bchl d, a primary photopigment of Chlorobi and some Chloroflexi (52, 53), was also expected given the abundance of Chlorobi in sample K003. However, the concentrations of total Bchl did not correlate well in absolute terms with our molecular tallies, suggesting that we could have missed novel APB populations with our stringent phylogenetic litmus test. Additionally, our differential ability to detect Bchl a in each site due to differences in the storage protocols may have also played a role. In any event, these analyses confirmed the presence and breadth of APBs.
While it is clear that the geographical extent of our sampling is insufficient to establish biogeographical patterns of distribution, the switch in intertidal endolith APB dominance between Isla de Mona and Menorca, involving Roseiflexus/Chlorothrix on the one side and Erythrobacter on the other, was internally consistent and quite significant. It will be interesting to determine in future studies if the pattern holds in other locations with larger biogeographical provinces; but, in the interim, a potential ecophysiological explanation could be put forward. It is known that in terrestrial environments, temperature can in fact drive biogeographic patterns of microbial phototroph distribution (54), and our two sites experience rather different temperature regimes. Isla de Mona has a minimum yearly seawater temperature of 25°C, while in Menorca, winter temperatures can dip down to 13°C (55). A literature review shows that the minimal reported temperature for growth in marine Chloroflexi is 18°C (47), whereas it can be as low as 10°C for Erythrobacter (51), and purple nonsulfur alphaproteobacteria can grow at temperatures as low as 5°C (56). This suggests that temperature may be a significant factor in determining the composition of APBs in intertidal carbonates.
Crucial to establishing the functional impact of APBs on endolithic communities and their geochemical impact on carbonates is to determine their metabolism in situ. Because most of the major APB OTUs in our survey (i.e., in Table 1) are allied with taxa known to act as photoheterotrophs in nature (37, 40, 57) and because of the absence of an obvious source of electron donors in our samples, we hypothesize that endolithic APBs likely conduct photoheterotrophy as their predominant metabolic function as endoliths, generating ATP through photophosphorylation and consuming organic compounds, including neutral and acidic sugars produced by cyanobacteria (58) as their source of carbon. Considering that diffusion limitation is one of the most important constraints in endolithic habitats (59), photoheterotrophic APBs could add a component of endolithic element cycling, consuming excess sugars, fermentation by-products, and even molecular oxygen (26), along with the release of CO2 back into the environment. Furthermore, photoheterotrophy has a demonstrable effect on carbonate geochemistry; Rhodovulum growing photoheterotrophically on acetate and lactate raised the external pH and precipitated carbonate, but it did not do so when grown on neutral sugars (60). Similar results (61) were obtained with Rubrivivax isolates.
Even though cyanobacteria have a mineral substrate preference at the single OTU level (23), we did not detect any such preference within APBs. This apparent independence of mineral substrate would be consistent with the notion that APBs are not actively carrying out carbonate dissolution, but rather depend on the boring action of cyanobacteria for endolithic space, a hypothesis that will require direct experimentation to formally test.
In summary, we have identified APBs as important endoliths of marine carbonates, with Chloroflexi (Roseiflexus and Chlorothrix), Erythrobacter (Erythrobacter sp. NAP1), and purple nonsulfur alphaproteobacteria as the most important types. Endolithic APBs could potentially play important metabolic roles in these communities and, in turn, exert geomicrobial effects on coastal carbonates.
It is of interest to compare the relevance of this new habitat for APBs to that of existing ones. Our samples had a depth-integrated average biomass of some 7 mg Bchl · m−2, which is much less than that observed in microbial mats (860 mg Bchl · m−2 [33]) or lake blooms (some 500 mg · m−2 [62]) but much more than that found in the open ocean (0.1 mg · m−2 [63]). When these areal densities are multiplied by the global extent of the respective habitats considered (64, 65), it becomes clear that endolithic APB biomass constitutes potentially a significant reservoir, slightly upwards of 105 kg of Bchl globally if our survey is representative of most outcrops. This reservoir is much larger than that in microbial mats (some 80 kg Bchl) or in the open ocean (3 × 104 kg Bchl) and similar in magnitude to that of lake blooms (1 × 105 kg Bchl; assuming that as much as 1/10 of the surfaces of all lakes stratify and are sufficiently eutrophic to support these blooms). Considering these simple calculations, the shallow interior of carbonates must be regarded as a globally major reservoir of APB biomass.


Sampling collection.

Samples from intertidally exposed hard carbonate rock were collected from Isla de Mona (18.0867°N, 67.8894°W), a small (11 km by 7 km) carbonate island 66 km west of Puerto Rico, having obtained permits from the Departamento de Recursos Naturales y Ambientales (Commonwealth of Puerto Rico), and from Menorca (39°58′0″N, 4°5′0″E), a populated carbonate-rich island 200 km east the Iberian Peninsula (Fig. 1). The study did not involve endangered or protected species. Rock samples were broken off from large boulders or cliff walls using a geological hammer, which was washed in seawater near the respective sampling site. Most samples were collected within the intertidal notch typical of carbonate cliffs, but a few were collected by scuba diving below the tidal ranges (only at Isla de Mona, K samples). Initial samples were then aliquoted with an ethanol-sterilized chisel and hammer. Samples were divided for mineralogical analysis (kept air dried) and either preserved in 70% ethanol (Isla de Mona) or frozen at −80°C (Menorca) for eventual DNA and lipid-soluble pigment extraction. The samples were then shipped at room temperature (Isla de Mona) or in liquid nitrogen (Menorca), reaching the laboratory in less than a week, and stored frozen at −80°C until analysis.

X-ray diffraction.

Subsamples were ground into a fine powder with a small amount of 100% ethanol. X-ray diffraction (XRD) patterns were collected using a Panalytical X'Pert Pro diffractometer mounted in the Debye-Scherrer configuration with a Cu K-α monochromatic X-ray source. The continuous scan mode was used in the 10 to 90° 2θ range to collect data. The relative mineral composition was identified using the X'Pert High Score plus software with a Rietvald refinement using default parameters.

Confocal microscopy.

Selected subsamples were dissolved over a period of 48 h in 200 mM EDTA (pH 5) (for calcite or aragonite) or 250 mM cyclohexylenedinitrilotetraacetate (CDTA; pH 5) (for dolomite) (41). The resulting liquids were then filtered using a GE black polycarbonate (PC) 0.8-μm-pore-size filter. The filter was placed on a drop of immersion oil on a microscope slide and covered with a second drop, over which a coverslip was placed, and kept at −20°C until imaging. Images were collected using a Leica SP5 confocal microscope at a 1,024 by 1,024 pixel resolution, with a minimum line average of 1 and a scan rate maximum of 400 Hz. Samples were excited using a 405-nm-wavelength laser. Innate Chl a emission (the “red” channel) was collected between 660 nm and 690 nm. Near-infrared emission (the “green” channel) was collected between 740 nm and 800 nm. A maximum intensity z projection of each channel was then visualized using Fiji (66).

Endolithic community DNA extraction.

Samples for DNA extraction were brushed aggressively with sterile toothbrushes and sterilized Milli-Q water to remove any epilithic biomass, which was already very sparse. Samples were chipped to get rid of the deep rock and to obtain smaller volumes of the top several millimeters, where the endolithic phototrophic biomass was conspicuous by color. To ensure a consistent sampling effort, we measured and cut pieces of chips with surface areas of 8 cm2. The pieces were then ground using sterile mortars as described by Wade and Garcia-Pichel (41), and 0.5 g of the sample was placed into the bead tube of a MoBio PowerPlant Pro kit (Mo Bio Laboratories, Inc., Carlsbad, CA, USA) for DNA extraction. We followed the protocol provided with one exception: prior to the first lysis step, we homogenized the bead tubes horizontally on a vortex mixer at 2,200 rev/min for 10 min and added 7 freeze-thaw cycles to ensure better disruption of the endolithic cells. DNA in the extract was quantified using a Qubit 2.0 HS DNA kit (Life Technologies, Carlsbad, CA, USA).

16S rRNA gene library preparation and Illumina sequencing.

The V3 to V4 variable region of the 16S rRNA gene was targeted for amplification using PCR primers 341F (5′-CCTACGGGNGGCWGCAG [67]) and 806R (5′-GGACTACVSGGGTATCTAAT [68]) with a barcoded forward primer. PCR amplification was performed using the HotStartTaq Plus master mix kit (Qiagen, USA) with the following parameters: 94°C for 3 min, followed by 28 cycles of 94°C for 30 s, 53°C for 40 s, and 72°C for 1 min, followed by a final 5-min elongation step at 72°C. The PCR products were then further purified and pooled to generate a single DNA library using the Illumina TruSeq DNA library preparation protocol. The library was sequenced using Illumina MiSeq following the manufacturer's guidelines. The library preparation, sequencing paired-ends assembly, and first quality trimming (with a Phred score cutoff of Q = 25) were performed commercially (MrDNALab, Shallowater, TX, USA).

Bioinformatics pipeline.

Paired sequences were then processed using the QIIME 1.9 analysis pipeline (69). First, chimera sequences were detected and removed utilizing the VSEARCH de novo chimera checking algorithm (70). Next, we ran the script using default parameters (removal of barcodes, removal of sequences less than 200 bp, and removal of sequences containing homopolymer runs longer than 6) to prep the data set for OTU picking, for which we used script with modified parameters. Briefly, OTUs were clustered at 97% using SortMeRNA (71) and SUMAclust (72), and taxonomy was assigned using SortMeRNA and the reference Greengenes 13_8 release database (73). Singletons were removed after OTU picking.

Reference phylogenetic tree building and OTU placement.

To determine which OTUs were likely phototrophic, we imposed the rule that for a given OTU to be considered phototrophic, it would have to phylogenetically belong to a clade that was composed of only known phototrophs. To facilitate this task, we constructed reference trees for each of the bacterial groups present in our samples known to contain phototrophic clades. The trees contained only sequences obtained from bona fide cultured isolates of known metabolism. Representative sequences for each tree were obtained from the SILVA SSU database (74) and aligned using MAFFT (75) and Guidance2 (76). The Guidance2 alignment with low-scoring columns removed for each group was then used for all further analyses. The reference trees were constructed on the CIPRES high-performance computing cluster (77) using the RAxML-HPC2 (78) workflow with the ML+Thorough bootstrap (1,000 bootstraps) method and the GTRGAMMA model. In all, we constructed 9 trees representing the following taxa: Chloroflexaceae, Chlorobiaceae, Rhodospirillaceae, Rhodocyclaceae, Comamonadaceae, Rhodobacteraceae, Rhizobiales, Erythrobacteraceae, and Chromatiales.
Next, we filtered the OTU table to include all OTUs that had been previously (see above) automatically assigned to taxa that contained known phototrophic bacteria. OTU representative sequences were then aligned to the appropriate reference alignment using PaPaRa (79) and placed into the edges of the respective reference trees using the evolutionary placement algorithm (based on the maximum likelihood model) feature of RAxML8 (80). Placement trees were visualized using the ITOL3 website (81). An OTU was considered a likely phototroph if it was placed within a phototroph clade (node) on the respective reference phylogenetic tree with better than 70% certainty.

Pigment extraction and analysis.

Lipid-soluble pigments were extracted as follows. First, 3 g of powdered sample (same sample as used for DNA) was suspended in a 7:2 acetone-methanol mixture and sonicated twice for 30 s in an ice bath in the dark. Extracts were centrifuged at 2,100 × g for 10 min and decanted, and the supernatants were filtered through a 0.22-μm-pore-size nylon filter. These steps were repeated until the supernatants were devoid of color. The resulting filtered supernatants were then dried under a N2 stream in the dark and then resuspended in 100% HPLC-grade acetone. HPLC analysis was conducted on an Agilent 1100 with an online photodiode array detector, using the protocol of Frigaard et al. (82) on a Novapak C18 3.9 mm by 300 mm (60-Å pore size, 4-μm particles) column. The gradient was composed of solvent A (methanol-acetonitrile-water, 42:33:25 by vol) and solvent B (methanol-acetonitrile-ethyl acetate, 50:20:30 by vol), and elution was performed as follows: at the time of injection, 30% B; linear increase to 100% for 52 min; constant for 15 min; and a return to 30% for 2 min. The flow rate was 1.0 ml · min–1, and the column temperature was 30°C. Pigments were identified by a comparison of the retention times and the spectra against true standards of Chl a and Bchl a from Sigma-Aldrich. All other pigments were identified from known spectra (45) and from extracts of Chloroflexus aurantiacus grown anaerobically. The injected pigment mass was calculated from the chromatogram using the equation m = FA (em · d)−1, where m is the mass of BChl or Chl in milligrams, F is the solvent flow rate (1 ml · min−1), A is the peak area (in absorbance units times seconds), em is the extinction coefficient in liter/mg/cm, and d is the path length of the PDA detector (1 cm). Extinction coefficients were taken from Ley et al. (83). We then converted the masses to mg/m2 using a per-sample surface area-to-volume ratio.

Data availability.

Isla de Mona sequences were deposited under GenBank accession numbers KT972744 to KT981874. Menorca sequences were deposited under GenBank BioProject identification (ID) number PRJNA396581.


This work was funded by National Science Foundation grant no. EAR 1224939 to F.G.-P., and by a Marie Skłodowska-Curie IOF grant awarded to E.C.
We thank the staff in ASU's Goldwater Materials Science Facility for analytical support and A. Garrástazu for field support and medical assistance.

Supplemental Material

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cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 84Number 415 February 2018
eLocator: e02055-17
Editor: Harold L. Drake, University of Bayreuth
PubMed: 29222097


Received: 20 September 2017
Accepted: 29 November 2017
Published online: 31 January 2018


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  1. bioerosion
  2. carbonate
  3. intertidal
  4. microbiomes



School of Life Sciences, Arizona State University, Tempe, Arizona, USA
Center for Fundamental and Applied Microbiomics, Biodesign Institute, Arizona State University, Tempe, Arizona, USA
Estelle Couradeau
School of Life Sciences, Arizona State University, Tempe, Arizona, USA
Laboratoire Biogéosciences, UMR6282, Université de Bourgogne, Dijon, France
Brandon Guida
School of Life Sciences, Arizona State University, Tempe, Arizona, USA
Susanne Neuer
School of Life Sciences, Arizona State University, Tempe, Arizona, USA
Center for Fundamental and Applied Microbiomics, Biodesign Institute, Arizona State University, Tempe, Arizona, USA
Ferran Garcia-Pichel
School of Life Sciences, Arizona State University, Tempe, Arizona, USA
Center for Fundamental and Applied Microbiomics, Biodesign Institute, Arizona State University, Tempe, Arizona, USA


Harold L. Drake
University of Bayreuth


Address correspondence to Ferran Garcia-Pichel, [email protected].

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