Cable bacteria (CB) are multicellular, filamentous
bacteria within the family of Desulfobulbaceae that transfer electrons longitudinally from
cell to cell to couple sulfide oxidation and oxygen reduction in surficial
aquatic sediments. In the present study, electrochemical reactors that
contain natural sediments are introduced as a tool for investigating the
growth of CB on electrodes poised at an oxidizing potential. Our experiments
utilized sediments from Yaquina Bay, Oregon, USA, and we include new
phylogenetic analyses of separated filaments to confirm that CB from this
marine location cluster with the genus “Candidatus Electrothrix”. These CB may belong to
a distinctive lineage, however, because their filaments contain smaller
cells and a lower number of longitudinal ridges compared to cables described
from other locales. The results of a 135 d bioelectrochemical reactor
experiment confirmed that these CB can migrate out of reducing sediments and
grow on oxidatively poised electrodes suspended in anaerobic seawater. CB
filaments and several other morphologies of Desulfobulbaceae cells were observed by scanning
electron microscopy and fluorescence in situ hybridization on electrode
surfaces, albeit in low densities and often obscured by mineral
precipitation. These findings provide new information to suggest what kinds of
conditions will induce CB to perform electron donation to an electrode
surface, further informing future experiments to culture CB outside of a
sediment matrix.
Introduction
Long-distance electron transfer (LDET) is a mechanism used by certain
microorganisms to generate energy through the transfer of electrons over
distances greater than a cell-length. These microorganisms may pass
electrons across dissolved redox shuttles, nanofiber-like cell appendages,
outer-membrane cytochromes, and/or mineral nanoparticles to connect
extracellular electron donors and acceptors (Li et al., 2017; Lovley, 2016).
Recently, a novel type of LDET exhibited by filamentous bacteria in the
family of Desulfobulbaceae was discovered in the uppermost centimeters of various aquatic,
but mainly marine, sediments (Malkin et al., 2014; Trojan et al., 2016).
These filamentous bacteria, also known as “cable bacteria” (CB),
electrically connect two spatially separated redox half reactions and
generate electrical current over distances that can extend to centimeters,
which is an order of magnitude longer than previously recognized LDET
distances (Meysman, 2017).
The unique ability of CB to perform LDET creates a spatial separation of
oxygen reduction in oxic surface layers of organic-rich sediment from
sulfide oxidation in subsurface layers (Meysman, 2017). The spatial
separation of these two half reactions also creates localized porewater pH
extremes in oxic and sulfidic layers, which induces a series of secondary
reactions that stimulate the geochemical cycling of elements such as iron,
manganese, calcium, phosphorus, and nitrogen (Kessler et al., 2018; Rao et
al., 2016; Seitaj et al., 2015; Sulu-Gambari et al., 2016a, b). In
addition to altering established perceptions of sedimentary biogeochemical
cycling and microbial ecology (Meysman, 2017; Nielsen and Risgaard-Petersen,
2015), CB also possess intriguing structural features that may inspire new
engineering applications in areas of bioenergy harvesting and biomaterial
design (Lovley, 2016; Meysman et al., 2019).
Much is still unknown about the basic mechanism(s) that CB use to perform
LDET. It has been suggested that when long filaments form, a chain of cells
at the sulfidic terminal catalyzes anodic half reactions (e.g., 0.5H2S+2H2O→0.5SO42-+ 4e-+5H+), while a cathodic half reaction (O2+ 4e-+4H+→2H2O) is completed by cells at the oxic terminal. Electron
transfer then occurs along the longitudinal ridges of CB filaments via
electron hopping promoted by extracellular cytochromes positioned within a
redox gradient or via conductive electronic structures such as pili (Bjerg
et al., 2018; Cornelissen et al., 2018; Kjeldsen et al., 2019; Meysman et
al., 2019; Pfeffer et al., 2012). These hypotheses await further
verification, and CB remain uncultured and difficult to grow outside of
sediment. This difficulty complicates efforts to study them using different
techniques, such as electrochemical assays and metatranscriptomics.
In a previous benthic microbial fuel cell (BMFC) experiment in a marine
estuary (Reimers et al., 2017), we serendipitously observed the attachment
of CB to carbon fibers serving as an anode in an anaerobic environment above
sediments. This finding suggested that CB possess the ability to donate
electrons to solid electron acceptors, and it indicated a range of cathodic
potentials favorable for electron transfer (Reimers et al., 2017). However
further investigations are still needed to study the conditions that allow
the attachment of CB to a poised electrode and to document electron transfer
mechanisms at their cathodic terminus. In the present study, we first
clarify the phylogenetic placement of CB found in sediments from Yaquina
Bay, Oregon, where the BMFC was previously deployed. Then, we describe the
design of a bioelectrochemical reactor configured to mimic the environment
in the anodic chamber of a BMFC and verify conditions that can induce CB
attachment on electrodes. Results assert that when oxygen is not available,
CB can glide through sediments and seawater to an electrode poised at
oxidative potentials. Thus, the present study provides new information about
the chemotaxis of CB in environments other than sediments, revealing key
conditions for their attachment to surfaces and growth in both natural and
engineered environments.
Materials and methodsStudy site and sediment collection
Several studies suggest that CB may be found widely in coastal sediments
possessing high rates of sulfide generation coupled with organic matter
mineralization (Larsen et al., 2015; Malkin et al., 2014; Pfeffer et al.,
2012). Therefore, to initiate this enquiry, sediment with these two
characteristics was collected from Yaquina Bay, Oregon, USA, using a hand
shovel at a site on an intertidal mud flat (IMF, 44∘37′30 N, 124∘00′26 W). The IMF site is located about 3 km upstream from the site where the BMFC was deployed in the abovementioned
study (Reimers et al., 2017). The top 20 cm of these sediments were sieved
through a 0.5 mm mesh size metal screen to remove macrofauna and shell
debris. Then the sieved sediments were allowed to settle and stored in
sealed buckets in a cold room at 5 ∘C.
Sediment incubation
To cultivate CB of Yaquina Bay, IMF sediment was initially incubated for 60 d. These first incubations were started 2 d after collection and
performed after homogenizing the sieved sediments under a flow of N2
and then packing the sediment into triplicate polycarbonate tubes (15 cm
height and 9.5 cm inner diameter). These cores were submerged in an aquarium
containing aerated seawater collected from Yaquina Bay and held at
15 ∘C, a temperature that is about average for the mudflats of
Yaquina Bay (Johnson, 1980). Once a distinctive suboxic layer was evident
from color changes in the top centimeters of the cores, profiles of
porewater pH, O2, and H2S were measured to 2–3 cm depth with
commercial microelectrodes (Unisense A.S., Aarhus, Denmark) to confirm
geochemical evidence of CB activity (see below). Multiple small sub-cores
(0.5 cm diameter, 3 cm in length) were then taken out from each incubated
core using cut-off syringes. Some of these sediment plugs were washed gently
to reduce the volume of fine particles, and CB biomass was further separated
out from the sediment matrix by using custom-made tiny glass hooks following
Malkin et al. (2014). Sediment plugs and separated filamentous biomass were
frozen or fixed for subsequent phylogenetic and microscopic
characterizations.
Reactor configuration and operation
To mimic the conditions where CB were found attached to electrode fibers in
a BMFC (Reimers et al., 2017), a bioelectrochemical reactor was assembled
from a polycarbonate core tube (15 cm height and 11.5 cm inner diameter,
Fig. 1) as a second phase of this research. A lid, a center rod to locate
and support the electrodes, and a perforated bottom partition were made from
polyvinyl chloride (PVC, McMaster-Carr, Elmhurst, IL). Three carbon brush
electrodes, which would serve as two anodes and a control electrode
(Mill-Rose, Mentor, OH, 2 cm in diameter and 8.9 cm total length), were
inserted through septa within holes in the core lining to meet the center
rod and were spaced radially at 120∘ angles from each other.
Schematic of the bioelectrochemical reactor design used in this
study: (a) lateral view of the reactor, (b) electrical circuit of the
reactor, and (c) bird's-eye view of the reactor cap and electrode arrangement.
Dimensions are in cm. A1 and A2 represent the current monitored in
duplicate anodes, V1 represents the potential monitored between the
duplicate anodes and cathode, and V2 and V3 represent potentials
monitored between the duplicate anodes and the reference electrode.
To initiate the experiment, the reactor was placed inside an 8 L plastic
beaker (with perforated walls) containing 3 cm of IMF sediments at the
bottom. Enough additional IMF sediment was then placed inside the reactor to
form an 8 cm thick layer after settling and compacting. In this configuration,
the sediment–water interface was approximately 1 cm away from the lower
extent of the carbon brush electrodes. The beaker was then gently lowered
inside an aquarium filled with Yaquina Bay seawater until fully submerged,
and the reactor was left uncapped. Seawater in the aquarium was maintained
at 15 ∘C and bubbled to maintain air saturation. A fuel cell
circuit was completed by placing a 10 cm long carbon-fiber brush cathode
(Hasvold et al., 1997) and a reference electrode (Ag/AgCl [3 M KCl],
MI-401F, Microelectrodes, Inc., Bedford, NH) into the seawater outside the
reactor tube (Fig. 1a).
The reactor was monitored in an open-circuit state for 31 d to allow the
development of a CB population within the top centimeters of sediment as had
been observed in the previous incubations. Microelectrode profiling was used
to characterize the vertical distribution of porewater pH and concentrations
of O2 and H2S on day 13 and 24 of reactor incubation. On day 31,
carbon fiber samples were trimmed off the unpoised anode brushes as initial
reference samples, and the reactor was sealed to create fully anoxic
conditions. Beginning on day 44, under seal, cathode versus anode potentials
were poised at 300 mV by regulating two of the three anode carbon brushes
with an individual custom-designed potentiostat circuit board (NW
Metasystems, Bainbridge Island, WA) (Fig. 1b). The third brush was kept at
open circuit as a continuing control. Electrode potentials of the anode
(versus reference) and whole cell and the current flow between anodes and
cathode were monitored and recorded every 7 min with a multichannel
data logger (Agilent Technologies, Santa Clara, CA, model 34970A fitted with
two 34901A multiplexer modules) wired to the potentiostat outputs. The
electrodes were poised for more than 3 months. On day 48 microelectrode
profiling was repeated by lowering the sensors through the ports in the
reactor lid (Fig. 1c). The pH microelectrode was broken at the start of this
profile and therefore no pH or calculated total sulfide are reported (only
H2S). On day 135, the anodes and control electrode were extracted
through the side openings in the bioreactor tube for SEM (scanning
electron microscopy) and CARD-FISH (catalyzed reporter deposition–fluorescence in situ hybridization)
analyses (described below). At the experiment's end, the final pH of the
seawater inside of the anodic chamber was measured by microelectrode (see
below).
Microelectrode measurements
The sediments incubated in open cores and in the bioelectrochemical reactor
were each profiled with O2, pH, and H2S microelectrodes to
show through geochemical signatures evidence of CB activity (Malkin et al.,
2014). Microelectrodes had tip diameters of 100 µm. The O2
microelectrodes were calibrated in air-purged seawater (as 100 % air
saturation) and in a solution of sodium ascorbate and NaOH (both to a final
concentration of 0.1 M, as 0 % O2 saturation). Vertical oxygen
microprofiles were recorded starting from 2 mm above either the
sediment–water interface or in the reactor above the carbon brush, at a
step size of 400 µm. Vertical pH and H2S microprofiles were
measured concurrently at the same spatial interval. The pH microelectrode
was calibrated by using standard pH 4, 7, and 10 buffer solutions (Ricca
Chemical, Arlington, Texas, USA). H2S microelectrodes were
calibrated by generating an 11-point calibration relationship by standard
addition, from 0 to 7.48 µM H2S at pH =1.6. A 3 mM standard
solution was made from crystal Na2S9H2O (> 98.0 %,
MilliporeSigma, Burlington, MA) in an anoxic glove box. Total sulfide
concentration at each profile depth was derived from pH and H2S
according to equilibrium relationships given in Millero et al. (1988).
SEM
To confirm the presence of CB and to examine the characteristic longitudinal
ridges and cell–cell junctions of CB, filaments extracted from the sediments
and carbon fibers from the reactor electrodes were visualized by scanning
electron microscopy (SEM). Samples were dehydrated in a graded series of
ethanol solutions from 10 % to 100 %. Specimens were then mounted on aluminum
SEM stubs with double-sided carbon tape, critical-point dried using an EMS 850 Critical Point Dryer, and sputter-coated with gold and palladium using a
Cressington 108 sputter coater. The resultant specimens were observed under
a FEI Quanta 600FEG ESEM at 5–15 kV. This instrument also provided
elemental spectra by X-Ray Energy Dispersive Spectrometry (EDS).
Representative microelectrode depth profiles of oxygen (blue), pH
(red), and ΣH2S or H2S (yellow) in (a) IMF sediment after
53 d of incubation and in the bioelectrochemical reactor at (b) day 13,
(c) day 24, and (d) day 48.
CARD-FISH
Catalyzed reporter deposition–fluorescence in situ hybridization (CARD-FISH)
was used to microscopically identify Desulfobulbaceae filaments using
a Desulfobulbaceae-specific oligonucleotide probe (DSB706; 5′-ACC CGT ATT CCT CCC
GAT-3′) labeled with horseradish peroxidase (Lücker et al.,
2007). In preparation for CARD-FISH, sediment samples were fixed with a 1:1
(vol : vol) ethanol and phosphate-buffered saline solution and stored at -20 ∘C until analysis. Extracted bacterial filaments and carbon
fibers cut from the carbon brush electrodes were treated with a fixative
solution containing 1.25 % glutaraldehyde and 1.3 % osmium tetroxide.
Fixed samples were stored at -20 ∘C until analysis. Sediment and
bacterial filament samples were first retained on polycarbonate membrane
filters and then mounted onto a glass slide by using 0.2 % agarose
(Malkin et al., 2014). Carbon fiber samples were mounted directly onto a
glass slide without first retaining on a filter. Mounted samples were
sequentially permeabilized by 10 mg mL-1 of lysosome (2 h at 37 ∘C) and achromopeptidase (1 h at 37 ∘C). After permeabilization,
glass slides were incubated in H2O2 (0.15 % in methanol) for 30 min at room temperature (∼25∘C) to inactivate the
endogenous peroxidases. The hybridization process was performed in a
standard hybridization buffer at 46 ∘C with 45 % formamide for 7 h (Wendeberg, 2010). Alexa Fluro 488 (ThermoFisher, Waltham, Massachusetts, USA) was deposited on samples in the presence of 0.15 % H2O2.
Two-color CARD-FISH was performed on some carbon fiber samples to look for a
previously observed co-occurrence of CB and other electroactive bacteria on
electrode surfaces (Reimers et al., 2017). To perform the CARD-FISH,
horseradish peroxidases on the hybridized DSB706 probes were inactivated by
0.15 % H2O2. The inactivated samples were then hybridized with
a Desulfuromonadales-specific oligonucleotide probe (DRM432; 5′-CTT CCC CTC TGA CAG
AGC-3′) modified with horseradish peroxidase in standard
hybridization buffer at 46 ∘C with 40 % formamide for 5 h and
sequentially stained with Alexa Fluro 555 (ThermoFisher, Waltham, Massachusetts, USA). A counter stain, 4′,6-diamidino-2-phenylindole (DAPI), was
applied to all samples after the deposition of fluorescent probe(s).
Hybridization samples were visualized using confocal laser scanning
microscopy (CLSM) (LSM 780, Zeiss, Jena, Germany).
Cable bacteria filaments recovered from Yaquina Bay sediments. (a) A cable bacteria filament under SEM. (b) A thin type of cable bacteria
filament under SEM. (c) Multiple filaments of cable bacteria clumped
together under SEM. The blue arrow indicates a section of cable, and the red arrow
indicates a cable bacteria filament covered with a mineral coating. (d) Identification of the filaments belonging to Desulfobulbaceae using catalyzed reporter
deposition–fluorescence in situ hybridization (DSB 706 probe + Alexa Fluor 488 in
green and DSB DAPI in blue). (e) Phylogenetic tree of Desulfobulbaceae 16s rRNA gene sequences
recovered from IMF sediment and extracted biomass samples. Color boxes
indicate previously recognized species of cable bacteria. The scale bar shows
5 % sequence divergence.
Microbial community characterizations
To investigate the phylogeny of the CB discovered in Yaquina Bay, genomic
DNA was extracted from three sediment plug samples and from two separated
filamentous biomass samples using a MoBio PowerSoil DNA Extraction Kit. To
avoid insufficient cell lysis, all samples went through five to seven freeze–thaw
cycles before the use of the extraction kit (Roose-Amsaleg et al., 2001).
Bacterial 16S rRNA genes were amplified by polymerase chain reaction (PCR) with random primers 357wF
(5′-CCTACGGGNGGCWGCAG-3′) and 785R (5′-GACTACHVGGGTATCTAATCC-3′). Amplification and sequencing of DNA
(Illumina MiSeq Reagent Kit v3, 2×300 bp) was performed by the
Center of Genome Research and Biocomputing at Oregon State University.
Sequences were processed using DADA2 (v.1.10) in R (3.5.0), as described in a
previous study (Callahan et al., 2016). Sequences were aligned to the Silva
SSU Ref NR database (v.132) and clustered into operational taxonomic units
(OTUs) at 97 % similarity. Representative sequences classified into the
family of Desulfobulbaceae were tagged and aligned to 16S rRNA gene sequences from
previously identified CB (Trojan et al., 2016). A phylogenetic tree was
constructed using RaxML with 1000 bootstraps (Stamatakis, 2014). Sequences
from this study were deposited to the Genbank's Sequence Read Archive
(MK388690-MK388723, PRJNA587126).
The current production (blue), the anodic potential (black), and
cathodic potential (orange) over time during the reactor experiment. The
reference electrode was an Ag/AgCl electrode with a saturated KCl filling
solution. This figure only shows measurements associated with one of the
duplicate electrodes.
SEM images illustrating (a, b) cable bacteria filaments with
visible ridges and cell–cell junctions incorporated into the biofilms on
carbon fiber electrode surfaces. Red pointers indicate cell–cell junctions.
(c, d, e, f) Short bacterial filaments without typical
morphological features of cable bacteria. Yellow arrows indicate the
locations of elongated cells. (g, h) Mineral-encrusted bacterial
filaments. (i) Image of control electrode surface after culture.
(a–c) Confocal microscope images illustrating cable
bacteria filaments on the carbon fibers that served as an anode. (d–f) Colonies of cells belonging to Desulfobulbaceae. Red circles indicate a possible
doublet of the long cells. Cells were visualized using catalyzed reporter
deposition–fluorescence in situ hybridization (DSB 706 probe + Alexa Fluor 488,
green; DRM 432 + Alexa Fluor 555, red; and DAPI, blue).
Results and discussionCable bacteria activity in the sediments of Yaquina Bay
During the initial open incubations of IMF sediment, the top centimeter of
each core changed from dark to light gray, and a brownish layer formed from
the sediment–water interface to ∼0.2 cm depth. Hallmarks of
the activity of CB were documented by microelectrode profiling after 53 d
of culture. These hallmarks were a sulfide-free suboxic zone and opposing pH
extremes at approximately 0.2 cm and 1–1.5 cm deep (Fig. 2a). Although a
faint smell of sulfide was detected during collection of the sediment, total
sulfide concentrations detected by microelectrode profiling were low
compared to previous studies of marine sediments hosting CB (Malkin et al.,
2014). The pH minimum within the anoxic layers of cultured sediment was 6.0,
indicating acidification coupled to sulfide or iron sulfide oxidation.
SEM revealed that cells within extracted filaments were 0.5 to 1.2 µm
wide and 2 to 3 µm long (Fig. 3a, b, c). Typical morphological
features of CB including longitudinal ridges and cell–cell junctions were
observed, though a smaller number of ridges (8–10) were usually spotted
compared to 16–58 in other characterizations (Malkin et al., 2014). Certain
filaments extracted from sediments were covered by heterogeneous coatings of
mineral particles as was observed recently by Geerlings et al. (2019) (Fig. 3c). These particles have similar elemental compositions to some authigenic
clays (Burdige, 2006), showing enrichments of silicon, aluminum, magnesium,
and iron. In our open incubation samples, some thinner filaments were also
seen that displayed no obvious longitudinal ridges, although cell–cell
junctions were still visible (Fig. 3b). Extracted filaments reacted
positively to the DSB 706 probe and DAPI (Figs. 3d and S1 in the Supplement).
When analyzing the 16S rRNA gene sequence data, we found that one of the
candidate CB genera, “Candidatus Electrothrix”, was relatively abundant in sediment plug
samples (2.9 %) and predominant (83.5 %) in separated filamentous
biomass samples. The most abundant Desulfobulbaceae OTUs within these samples were aligned
with a previously established taxonomy framework of CB (Trojan et al., 2016)
(Fig. 3e). Partial 16s rRNA sequences of CB have been discovered in sediment
samples from the US East Coast, Gulf of Mexico, and certain sites on the US
West Coast from SILVA or GenBank databases (Trojan et al., 2016). Our
studies have provided the first combined microscopic and genetic
observations of CB in sediments from the northeastern Pacific coast of the United
States, reinforcing the suggestion that CB bacteria are distributed
globally. This result also indicates that Yaquina Bay, OR, where we deployed
the BMFC, indeed harbors a rich population of CB.
Encouraging the growth of cable bacteria on poised electrodes
Geochemical hallmarks of CB developed within 2 weeks of culture within the
bioelectrochemical reactor (Fig. 2b). The pH minimum within the sulfidic
layer and the pH maximum in the oxic layer of sediment became more extreme
by day 24 (Fig. 2c), indicating that a CB population was actively mediating
electrogenic sulfide oxidation and transporting electrons to reduce oxygen.
After sealing the reactor, oxygen concentration in the overlaying seawater
dropped below detection limits and the open-circuit anode potential fell to
-104 mV (versus Ag/AgCl). Once poised with the potentiostat, the cathode and
anode potentials became stable at approximately 330 and 30 mV versus
Ag/AgCl, respectively. When microelectrode profiling was performed on day 48, the measurements indicated that the overlying seawater was anoxic
(except right below the sample port) and that free H2S was detectable
right below the sediment–water interface but not in the water column (Fig. 2d). The pH of the seawater measured inside of the reactor chamber at the
experiment's end was 6.2, consistent with sulfide oxidation under anaerobic
conditions within the anolyte seawater. Current collection started to
increase once the anodic potential became stable, indicating that the anode
brushes were being used as an electron acceptor. Current records collected
from duplicate electrodes were similar, and the current in each steadily
increased to ∼30.5±2.5µA by day 86,
stabilized, then rose again to a peak of ∼75±8µA on day 101. After this maximum, current decreased and restabilized
at ∼30±5µA. These electrochemical results are
portrayed in Fig. 4. The cause of the current rise and subsequent fall (Fig. 4) is unknown but is a common occurrence in marine BMFC experiments (Nielsen
et al., 2009; Ryckelynck et al., 2005). It is likely that such behavior is a
result of a varying supply of reductants from the underlying sediment,
changes in the anodic biofilm, and finally loss of anode surface area due to
mineral deposition induced by microbial activity and/or the applied
electrical potential. Coatings containing iron, phosphorus, sulfur, silicon,
and aluminum are often found on anode surfaces of BMFCs in marine
environments and were seen by SEM in the present study (see below). Cyclic
voltammetry (CV, Fig. S2a) performed on the anode brushes at day 52 and
100 yielded broad and poorly defined electrochemical signals. The
interpretation of such voltammograms may be complicated by a high
uncompensated resistance between working electrode and reference electrode
(Babauta and Beyenal, 2015). While an oxidation peak can be clearly
identified at potentials near where the anode was held, the peak current did
not increase with an increase in scan rate (Fig. S2b). The peak oxidation
current also did not change much between day 52 and day 100. This CV
behavior suggests that any current generated by the biomass of electroactive
bacteria, including CB, was obscured during scans by current arising from
irreversible redox reactions, such as oxidation of dissolved iron. A
reduction peak was unidentifiable throughout the scans, a common phenomenon
in sediment MFCs (Babauta and Beyenal, 2015). Taken together, these results
demonstrate that the electrode surface was altered during the course of the
bioreactor experiment by mineral and chemical precipitate deposition (Imran et
al., 2019).
Examining the attachment of cable bacteria on the anode
The hypothesis that led to the bioreactor experiments in this study was that
an electrode poised at an oxidative potential can produce redox conditions
and geochemical gradients that attract CB and that will lead to their
electron donation to an electrode. Several observations that were made on
harvested electrodes affirm this hypothesis. Firstly, under SEM, bacteria
filaments with visible longitudinal ridges and cell–cell junctions were
found integrated into biofilms on the surfaces of poised electrodes (Fig. 5a, b). As observed in the initial IMF sediment examinations, filaments
appeared to contain a smaller number of ridges (8 to 10) compared to
previously reported CB filaments and others were without pronounced ridges
along their longitudinal axes. The latter examples did show cell–cell
junctions and appeared to have wrinkled surfaces (Fig. 5c, d, e, f).
Secondly, many of the bacteria filaments observed on the electrode surfaces
were encrusted, suggesting mineral deposition similar to that observed at
the oxic terminal of CB filaments in sediments (Fig. 5g, h). EDS
indicated that these deposits contained iron, phosphorus, oxygen, and
silicon (Fig. S3b). The control electrode that was not positively poised
displayed no mineral deposition and nearly no cell growth (Fig. 5i).
Thirdly, most of the bacterial filaments on the poised electrode surfaces
reacted positively with the Desulfobulbaceae-specific probe. CARD-FISH performed in the
present study revealed that the anodic carbon fibers harbored many short
bacterial filaments, as well as colonies belonging to the family of
Desulfobulbaceae (Fig. 6a, b, c, d, e, f). Clear cell–cell junctions were observed along
many of the fluorescent filaments. However, the complexity of the carbon
fiber samples often hampered clear microscopic visualization of fluorescent
cells. Application of an additional Desulfuromonadales-specific oligonucleotide probe (DRM432)
confirmed the presence of other known electrogenic bacteria on the carbon
fibers near Desulfobulbaceae cells as well (Fig. 6b, c, d).
Though a global occurrence has recently been indicated (Malkin et al.,
2014), CB successfully evaded microbiological survey for quite a long time.
One of the reasons is likely that the phylogeny of CB is overshadowed by the
broad family of Desulfobulbaceae, which are often highly abundant in marine sediments
(Kuever, 2014). Another reason may be a resistance of the cells of CB to
routine cell lysing techniques that have been used with many DNA extraction
kits (Trojan et al., 2016). Therefore, their identification in various
studies has relied on microscopic observations of their unique filamentous
form and morphological features (ridges and cell–cell junctions), combined
with fluorescence in situ hybridization labeling (Malkin et al., 2014, 2017;
Malkin and Meysman, 2015). The electrochemical reactor in this study was
anoxic for more than 100 d. The observation of CB on the anodic carbon
fibers at the end of this experiment confirms that, although they may not
have been abundant, they can survive under such conditions and were likely
using the anode as an electron acceptor (as suggested previously by Reimers et
al., 2017). Besides the recognized forms of CB, short filaments within the
family of Desulfobulbaceae that possessed different morphologies were also observed on the
anode surface. Aller et al. (2019) suggest that redox environment may play
an important role in controlling the length of CB filaments. For example, in
the bioturbated zone associated with the tube worm Chaetopterus variopedatus, in which redox
conditions often oscillated between oxic and hypoxic, CB were present
predominately in short filaments. Assuming the CB can use an electrode as an
electron acceptor, the distance between the electron donor and acceptor
utilized by CB may be short, reducing the advantage of forming long
filaments.
The closest culturable relative to CB, Desulfobulbus propionicus, can utilize an electrode as an
electron acceptor to oxidize S0, H2, and organic acids like
pyruvate, lactate, and propionate (Holmes et al., 2004). While CB appear to
possess features like motility and an ability to form loops and bundles that
are similar to large sulfur bacteria (but distinct from the D. propionicus), our SEM and
CARD-FISH examinations suggest that CB on oxidative electrode surfaces may
produce extracellular structures to transfer electrons to an electrode
and/or to insoluble Fe(III)-oxides similar to D. propionicus (Bjerg et al., 2016; Holmes
et al., 2004; Jørgensen, 2010; Pfeffer et al., 2012). Admittedly,
indisputable proof of electron transfer from CB to electrodes still awaits
growth in purer biofilms and cultures.
Summary and implications for the electrode-associated growth of cable
bacteria
The present study introduces bioelectrochemical reactors as an approach to
investigate filamentous cable bacteria and their unique ability to transfer
electrons. Furthermore, we confirm that an active population of filamentous
CB are present in Yaquina Bay, Oregon, USA, where CB were previously found
attached to carbon-fiber electrode surfaces within a BMFC (Reimers et al.,
2017). Moreover, by incubating intertidal sediment collected from Yaquina
Bay in a reactor mimicking the anodic chamber of a BMFC, we observed that CB
can be drawn to electrodes at oxidative electrical potentials. Thus, we have
further evidence that CB can survive under anoxic conditions in the presence
of an oxidative electrode serving as an electron acceptor. The
bioelectrochemical reactor study also showed attachment of CB to an
oxidative electrode when the surrounding seawater was stripped of hydrogen
sulfide and at a pH ∼6.2. However, the observed CB density
and the overall density of recognizable cells were both relatively low on
electrode surfaces, as the respirable surface area appeared to become
limited by the deposition of mineral coatings. More work is needed to
determine conditions or experimental designs that may attract CB to an
electrode while not also leading to excessive mineral precipitation on
electrode surfaces. Developing ex situ culture techniques of CB and using these
approaches to gain insight into their electron transfer will contribute to
the overall understanding of this group of bacteria, their genomic makeup,
and their survival in both natural and engineered environments.
Data availability
Partial 16S rRNA gene and raw sequences are deposited in the Genbank's Sequence Read Archive (MK388690-MK388723, PRJNA587126).
The supplement related to this article is available online at: https://doi.org/10.5194/bg-17-597-2020-supplement.
Author contributions
CL and CER conceived the study. YA designed and assembled the
bioelectrochemical reactor. CL performed the microscopic examinations and
microprofiling and analyzed the microbial community and phylogenies. CL
also wrote the manuscript, with major rewrites and editing contributed by CER.
Competing interests
The authors declare that they have no conflict of interest.
Acknowledgements
This research was funded through grant N00014-17-1-2599 from the Office of
Naval Research to Clare E. Reimers. We thank Teresa Sawyer, the Electron Microscopy Facility
Instrument Manager at Oregon State University, for assistance with the SEM
imaging and Anne-Marie Girard for valuable advice on the confocal
microscope imaging. The authors wish to acknowledge the Confocal Microscopy
Facility of the Center for Genome Research and Biocomputing at Oregon State
University, which is supported in part by an award (no. 1337774) from the
National Science Foundation. We also thank our lab members for providing
critical comments on the manuscript.
Financial support
This research has been supported by the Office of Naval Research (grant no. N00014-17-1-2599) and the National Science Foundation (grant no. 1337774).
Review statement
This paper was edited by Jack Middelburg and reviewed by two anonymous referees.
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