Manganese-Cycling Microbial Communities Inside Deep-Sea

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Manganese cycling microbial communities inside deep-sea manganese nodules Marco Blöthe, Anna Wegorzewski, Cornelia Müller, Frank Simon, Thomas Kuhn, and Axel Schippers Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/es504930v • Publication Date (Web): 28 May 2015 Downloaded from http://pubs.acs.org on June 4, 2015

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Environmental Science & Technology

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Manganese cycling microbial communities inside deep-sea

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manganese nodules

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Marco Blöthe1, Anna Wegorzewski1, Cornelia Müller2, Frank Simon3, Thomas Kuhn1 and Axel

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Schippers1*

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Germany

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Leibniz Institute for Applied Geophysics, Stilleweg 2, 30655 Hannover, Germany

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Leibniz Institute of Polymer Research Dresden, Hohe Str. 6, 01069 Dresden, Germany

Federal Institute for Geosciences and Natural Resources (BGR), Stilleweg 2, 30655 Hannover,

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*Corresponding author: Stilleweg 2, D-30655 Hannover, Germany

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E-mail: [email protected]; Phone: +49(0)511-643-3103; Fax +49(0)511-643-2304

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Polymetallic nodules (manganese nodules) have been formed on deep sea sediments over

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millions of years and are currently explored for their economic potential, particularly for

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cobalt, nickel, copper and manganese. Here we explored microbial communities inside nodules

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from the northeastern equatorial Pacific. The nodules have a large connected pore space with a

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huge inner surface of 120 m2/g as analyzed by computer tomography and BET measurements.

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X-ray photoelectron spectroscopy (XPS) and electron microprobe analysis revealed a complex

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chemical fine structure. This consisted of layers with highly variable Mn/Fe ratios (< 1 - > 500)

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and mainly of turbostratic phyllomanganates such as 7 and 10 Å vernadites alternating with

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layers of Fe-bearing vernadite (δ-MnO2) epitaxially intergrown with amorphous feroxyhyte (δ-

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FeOOH). Using molecular 16S rRNA gene techniques (clone libraries, pyrosequencing, real-

ACS Paragon Plus Environment

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time PCR) we show that polymetallic nodules provide a suitable habitat for prokaryotes with

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an abundant and diverse prokaryotic community dominated by nodule-specific Mn(IV)-

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reducing and Mn(II)-oxidizing bacteria. These bacteria were not detected in the nodule-

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surrounding sediment. The high abundance and dominance of Mn-cycling bacteria in the

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manganese nodules argue for a biologically driven closed manganese cycle inside the nodules

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relevant for their formation and potentially degradation.

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Introduction

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Marine encrustations such as nodules and crusts consist of Mn-Fe oxi-hydroxide precipitations and

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are found in oceans worldwide with economically important deposits occurring in the Clarion and

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Clipperton Zone (CCZ) of the Pacific Ocean, which extends from off the west coast of Mexico to as

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far west as Hawaii1-3. Polymetallic nodules in the CCZ consist predominantly of Mn (31%), Fe (6%),

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Cu (1.2%), Ni (1.3%), and Co (0.2%), but are also enriched in Mo, Zr, Li, Y, and rare earth elements

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(REEs)4.

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Current conditions of the CCZ seafloor are oxic (~ 150 µmol/l O2) at least down to 2 to 2.5 m

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sediment depth5. The sediments consist of clay and siliceous ooze1 in water depths of 4000 to 5000 m

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which are at or below the carbonate compensation depth6. Soon after the discovery of manganese

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nodules, theories about their origin and formation were proposed and discussed7-9. All assumed that a

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primary oxidation of manganese to MnO2 is followed by agglomeration of the MnO2 into nodules. It

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was hypothesized that manganese in seawater first adsorbs to nodule substance by a rapid abiotic

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reaction (equation 1) and consecutively undergoes a bacterial oxidation to Mn(IV) (equation 2)

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becoming part of the nodule matrix (MnMnO3):

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(1)

H2MnO3 + Mn2+

→ MnMnO3 + 2 H+

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(2)

MnMnO3 + 0.5 O2 + 2 H2O → 2 H2MnO3 .

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Bacteria are major players in sediment diagenesis by releasing manganese (and other metals) to the

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pore fluids, which then take part in the process of nodule formation. Less is known about bacterial

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involvement in the nodule build-up process and the so far supposed importance of microorganisms

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for nodule formation was only inferred from the enrichment/isolation of Mn(II)-oxidizing bacteria

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from manganese nodules and their activity determination8. Energy-dispersive X-ray spectroscopy

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showed that microorganisms were associated with nodule parts rich in Mn and Fe9. Apart from

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biomineralization via Mn(II) oxidation, the presence of dissimilatory Mn(IV) reducing prokaryotes


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on the nodules may indicate ongoing nodule degradation. Microbial communities on ferromanganese

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oxides in terrestrial habitats have been reported from freshwater sediments10, caves11, or soils12; these

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examples suggest that phylogenetic diverse prokaryotes are involved in manganese oxidation. Similar

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microbial diversity investigations in marine environments have been reported for ferromanganese

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crusts13, for deep-sea surface sediments14-15 and for manganese nodules16-17 but without giving hints

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for microbial manganese cycling.

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To comprehensively explore microbial communities inside manganese nodules with respect to the

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nodule formation processes, the mineralogical composition and the internal structure of nodules from

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six different areas of the CCZ were analyzed by X-ray photoelectron spectroscopy (XPS), high-

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resolution micro computer tomography (µ-CT) and electron microprobe analyses (EMPA). The

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nodule associated microbial communities were analyzed by quantitative, real-time PCR (qPCR), 16S

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rRNA-gene clone libraries and primer-tagged pyrosequencing. We found that the nodules have a

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large connected pore space providing a suitable habitat for prokaryotes. The most abundant group of

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microorganisms in the nodules could be assigned to the Gammaproteobacteria, with Shewanella and

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Colwellia as the dominant genera, distinct from the communities present in the sediment surrounding

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the nodules. The dominance of these Mn-cycling bacteria argues for a biologically driven closed

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manganese cycle inside the nodules.

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Methods

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Sampling. Nodules and surrounding sediments were sampled with a multicorer during the scientific

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cruise SO-205 with R/V Sonne from six different working areas inside the German license area in the

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CCZ in 2010 (Supplementary Fig. S1 and Table S1). Nodules from single multicorer tubes were

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rinsed with filter sterilized seawater to remove the surrounding sediment. All samples were either

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immediately used for experiments onboard or frozen at – 20 °C for later DNA-based analyses. The


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frozen samples were transported on dry ice to the BGR laboratory in Hannover, Germany. Further

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nodules for mineralogical analyses were collected with a box corer.

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X-ray photoelectron spectroscopy (XPS). XPS analyses were carried out on individual subsamples

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of three nodules from the CCZ. The XPS spectra were recorded using a Kratos AXIS Ultra XPS

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system. A detailed description of the method is given elsewhere4. For XPS measurements, three

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nodules (27KG, 32KG, 49KG) were sampled from the surface of a box corer together with the

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sediment and the near-bottom water and were kept in their natural position for transportation. All

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nodules were smooth on the surface, exposed to the bottom water, and slightly porous on the bottom

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side (which was buried in the upper few cm of the sediment).

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High-resolution micro computer tomography (µ-CT). The distribution of pores, fractures, the pore

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network as well as pore size were determined by µ-CT analysis of two small nodules from the CCZ.

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The high-resolution scans were performed with the Phoenix nanotom s 180 CT scanner. The scans of

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the nodule were carried out with a water-cooled nanofocus X-ray tube (180kV/15W). The further

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visualization of the pore space and 3D imaging were performed with the program AVIZO Fire. For

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CT analysis two small nodules (44KG, 49KG; 2 x 2.5 x 1.5 cm) were dried for 48h at 40°C.

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Electron microprobe analyses (EMPA). The chemical composition of the pore fillings between the

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dendritic growth structures was investigated for a few nodules using electron microprobe analyses

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(CAMECA SX - 100). Thick sections of nodules were prepared. Measurements of genetically

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different layers were carried out using a 15/20 kV accelerating voltage and 40 nA beam current. The

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detailed description of the method is given elsewhere4.

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Onboard experiments for testing microbial Mn2+ removal from seawater. After sampling,

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manganese nodules were washed by flushing with filter-sterilized seawater to remove attached

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sediment and surface-sterilized by boiling for 30min in a glass beaker with filter sterilized seawater

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as described elsewhere 8. The nodules were transferred to an alcohol-flamed mortar and crushed with

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a sterile pestle to sandy grain size. Two gramms of pulverized material was transferred to a 125ml


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Erlenmeyer flask with 18ml filter sterilized seawater and 2ml of sterile 0.2M MnSO4 solution.

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Peptone was added to a finale concentration of 1%. For the glucose assay 1ml of the 0.2M MnSO4

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and 1ml of a sterile 10% glucose solution was added. Manganese concentration in the supernatant

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was determined by the formaldoxime method.

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Extraction of nucleic acids, and amplification of bacterial and archaeal 16SrRNA genes.

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Material for DNA extraction was cut off from frozen nodules with a hand drill and a circular saw

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(diameter 2.5cm). Material was taken from four different parts of nodules which are named in the

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following (I) hydrogenetic part, (II) diagenetic part, (III) core part and (IV) equatorial rim (Fig. 1).

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The hydrogenetic part is the smooth part of the nodules that was exposed to the near-bottom seawater

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and the diagenetic part is the rough surface texture part of the nodules, which sits in the sediment.

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The equatorial rim is marking the position up to which the nodules were embedded in the sediment.

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The core part is the inner part of the nodules. To prevent cross contamination from moving particles

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produced by the circular saw blade, we first cut off the material from the equatorial rim and then

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scraped off any particles from the nodule surface with a sterile spatula. In a second step surface

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material was cut off from the hydrogenetic and diagenetic parts. Material from the core part was

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removed after breaking the nodule apart. The circular saw was flame sterilized before each cutting

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step. Finally, the material was crushed to millimeter-sized or smaller fractions using sterilized pistil

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and mortar and environmental DNA was extracted from 10g of crushed material or sediment using

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the Ultraclean soil DNA kit (MoBio Laboratories) following the manufacturer’s protocol.

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The amplification of 16S rRNA genes from Bacteria was performed by PCR with the universal

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bacterial

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TACCTTGTTACGACTT- 3’)18 from Thermo Scientific. PCR mix was prepared from Thermo

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Scientific 2xMasterMix (final concentration: 75mM Tris-HCl (pH 8.8), 1.5mM MgCl2, 0.2 mM each

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of dNTP, 0.5µM each of primer, 0.652U ThermoPrime Taq DNA Polymerase, 100ng/µL BSA) and a

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2µL template of extracted DNA in a total reaction of volume 50µL. Negative controls without

primers

GM3F

(5′-AGAGTTTGATCMTGGC-3’)


and

GM4R

(5′-

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template were used as a contamination check. Reaction mixtures were held at 95°C for 2min

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followed by 35 cycles of 94°C for 30s, 52°C for 30s, and 72°C for 90s, with a final extension step of

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5 min at 72°C. PCR for Archaea was carried out with the primers 109f (5’-ACK GCT CAG TAA

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CAC GT-3’)19 and 912r (5’-CTC CCC CGC CAA TTC CTT TA-3’)20 from Thermo Scientific. The

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composition of the Master mix was the same as described for the bacterial PCR. These thermocycling

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conditions were used: one cycle at 95°C for 5min; 26–30 cycles at 95°C for 1min, 52°C for 1min,

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and 72°C for 3min; and one cycle at 72°C for 6min. PCR amplification of DNA extracts using group-

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specific primers (Microsynth, Switzerland) for Shewanella was performed as described 21.

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Clone library construction and phylogenetic analyses. Products of PCR reactions were cloned and

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sequenced by the company Microsynth (Switzerland). Overlapping sequencing from both sides of the

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16S rRNA genes was performed. Contigs were constructed with the software Geneious Pro 6.0 and

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checked for chimera with UCHIME 22. In total 620 archaeal with more than 800bp and 922 bacterial

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sequences with more than 1300 bp were obtained. These sequences were aligned with the SILVA

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Incremental Aligner (SINA

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software package (v.5.5

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operational taxonomic units (OTU, 97% similarity) and to calculate the coverage, diversity indices

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and to perform analysis of molecular variance (AMOVA). For the AMOVA analysis sequences were

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organized by clone libraries and the P value significance tests for the variance components were

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carried out using 10000 permutations. One sequence from each OTU harboring at least 5 sequences

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was picked as a representative and imported to the SILVA_111NR template tree using the ARB

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program suite. An additional 10 sequences for each OTU representative were selected based on the

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phylogenetic affiliation (minimum distance 5%, in total 300) in the SILVA_111NR. Selected

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reference sequences together with the OTU-representatives were used for tree construction using

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maximum likelihood algorithm (RAxML) with GTRGAMMA as rate distribution model, the general

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bacteria filter provided in ARB and 1000 bootstraps.

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) and the SILVA_111NR database and processed with the ARB

). This alignment was used in the Mothur v 1.28 program


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to build

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Quantitative PCR. Extracted DNA from nodules and surface sediment samples was amplified by

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qPCR using the device ABI Prism 7000 (Applied Biosystems) and master mixes from the companies

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Applied Biosystems, Eurogentec, or Invitrogen. Each DNA extract was measured in triplicate. The

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copy numbers of the 16S rRNA gene were quantified for Archaea 26 and Bacteria 27 using TaqMan

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assays and converted to cell numbers using conversion factors of 1.5 for Archaea and 4.1 for

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Bacteria, as previously done 28.

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Pyrosequencing. Amplicon pyrosequencing was performed on a 454 GS FLX Titanium system

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(Roche, Penzberg, Germany) as recently described 29. Briefly, barcoded amplicons for multiplexing

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were prepared using the primers Ba27f (5’-AGA GTT TGA TCM TGG CTC AG-3’) and Ba519r

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(5’-TAT TAC CGC GGC KGC TG-3’) extended with the respective A or B adapters, key sequence

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and multiplex identifiers (MID) as recommended by Roche. Pyrotag PCR was performed in a

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Mastercycler ep gradient (Eppendorf) with the following cycling conditions: initial denaturation

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(94°C, 5min), followed by 28 cycles of denaturation (94°C, 30s), annealing (52°C, 30s) and

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elongation (70°C, 60s). Each 50µl PCR reaction contained 1× PCR buffer, 1.5mM MgCl2, 0.1mM

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dNTPs, 1.25U recombinant Taq polymerase (Fermentas, St. Leon-Rot, Germany), 0.2µg ml-1 bovine

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serum albumin (Roche), 0.3mM of each MID-primer (Biomers, Ulm, Germany) and 1µl of template

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DNA. Processing of sequences using Mothur v2.28, including denoising and chimera removal, was

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performed according to the standard operating procedure

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allowing 0 mismatch to the barcode and to the primer and a maximum homopolymer length of 8

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bases. Sequences shorter than 250bp or longer than 500bp were removed. Chimeras were removed

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using UCHIME. Amplicon sequences were sorted according to their barcodes and primers.

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Sequences remaining after these initial steps were further processed with the online tools SILVAngs

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30

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. Briefly, sequences were screened by

.

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Results and Discussion

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Nodule porosity and chemistry. High-resolution computer tomography (µ-CT) analysis discovered

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a connected pore system inside polymetallic nodules and revealed a high internal porosity ranging

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from 26 to 61 vol.-% (Fig. 2, Table 1) with a high mean surface area of 119 m2/g and a mineral

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density of 2.83 g/cm3. Overall, the pore sizes decreased from the surface towards the inner parts of

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the nodules. The outer 11 µm of the nodules contained 38% small pores (90%), rarefaction analysis revealed that the top sediment harbor greater bacterial richness

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than the nodule (Fig. S7) and the molecular data show that the bacterial community inside the

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nodules is clearly distinct from that in the sediment (Table S7). The same trend applies to the

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archaeal community, except that the community from the hydrogenetic part of the nodule is more

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similar to the sediment than the rest of the nodule. This interpretation is supported by the

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dissimilarity measure value θYC (Fig. S8), and by the analysis of molecular variance (AMOVA)

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(Table S7 and Table S8). Comparing the bacterial community from the nodule with the bacterial

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community published for seafloor basalt from the East Pacifc Rise and Hawaii 46 the cumulated

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nodule clone libraries (all 16S rRNA gene clone libraries combined) shared fewer OTUs with these

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hard rock based habitats than with the sediment (data not shown), a result which clearly demonstrate

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that the manganese nodule harbors a distinct bacterial community.

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Overall, the data suggest that the microbial diversity in the sediment is higher than in the nodule and

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the core part of the nodule showed a slightly increased diversity compared to the nodule outer part.

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Similar observations have been reported previously for nodules from the South Pacific Gyre based on

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pyrosequencing16 and for iron-manganese concretions from the Baltic Sea based on DGGE

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analyses17. Since it is assumed that the microbial diversity is positively correlated with cell density

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linked to the availability of potential energy sources, the higher total organic carbon (TOC) content

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of the surface sediment (

Manganese-Cycling Microbial Communities Inside Deep-Sea

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