Shewanella oneidensis MR-1 nanowires are outer membrane and periplasmic extensions of the extracellular electron transport components

Sahand Pirbadian, Sarah E. Barchinger, Kar Man Leung, Hye Suk Byun, Yamini Jangir, Rachida A. Bouhenni, Samantha B. Reed, Margaret F. Romine, Daad A. Saffarini, Liang Shi, Yuri A. Gorby, John H. Golbeck, and Mohamed Y. El-Naggar
PNAS September 2, 2014 111 (35) 12883-12888; first published August 20, 2014 https://doi.org/10.1073/pnas.1410551111

  1. Edited by Harry B. Gray, California Institute of Technology, Pasadena, CA, and approved July 30, 2014 (received for review June 9, 2014)

Significance

Bacterial nanowires from Shewanella oneidensis MR-1 were previously shown to be conductive under nonphysiological conditions. Intense debate still surrounds the molecular makeup, identity of the charge carriers, and cellular respiratory impact of bacterial nanowires. In this work, using in vivo fluorescence measurements, immunolabeling, and quantitative gene expression analysis, we demonstrate that S. oneidensis MR-1 nanowires are extensions of the outer membrane and periplasm, rather than pilin-based structures, as previously thought. We also demonstrate that the outer membrane multiheme cytochromes MtrC and OmcA localize to these membrane extensions, directly supporting one of the two models of electron transport through the nanowires; consistent with this, production of bacterial nanowires correlates with an increase in cellular reductase activity.

Abstract

Bacterial nanowires offer an extracellular electron transport (EET) pathway for linking the respiratory chain of bacteria to external surfaces, including oxidized metals in the environment and engineered electrodes in renewable energy devices. Despite the global, environmental, and technological consequences of this biotic–abiotic interaction, the composition, physiological relevance, and electron transport mechanisms of bacterial nanowires remain unclear. We report, to our knowledge, the first in vivo observations of the formation and respiratory impact of nanowires in the model metal-reducing microbe Shewanella oneidensis MR-1. Live fluorescence measurements, immunolabeling, and quantitative gene expression analysis point to S. oneidensis MR-1 nanowires as extensions of the outer membrane and periplasm that include the multiheme cytochromes responsible for EET, rather than pilin-based structures as previously thought. These membrane extensions are associated with outer membrane vesicles, structures ubiquitous in Gram-negative bacteria, and are consistent with bacterial nanowires that mediate long-range EET by the previously proposed multistep redox hopping mechanism. Redox-functionalized membrane and vesicular extensions may represent a general microbial strategy for electron transport and energy distribution.

Reduction–oxidation (redox) reactions and electron transport are essential to the energy conversion pathways of living cells (1). Respiratory organisms generate ATP molecules—life’s universal energy currency—by harnessing the free energy of electron transport from electron donors (fuels) to electron acceptors (oxidants) through biological redox chains. In contrast to most eukaryotes, which are limited to relatively few carbon compounds as electron donors and oxygen as the predominant electron acceptor, prokaryotes have evolved into versatile energy scavengers. Microbes can wield an astounding number of metabolic pathways to extract energy from diverse organic and inorganic electron donors and acceptors, which has significant consequences for global biogeochemical cycles (2⇓–4).

For short distances, such as between respiratory chain redox sites in mitochondrial or microbial membranes separated by <2 nm, electron tunneling is known to play a critical role in facilitating electron transfer (1). Recently, microbial electron transport across dramatically longer distances has been reported, ranging from nanometers to micrometers (cell lengths) and even centimeters (5). A few strategies have been proposed to mediate this long-distance electron transport in various microbial systems: soluble redox mediators (e.g., flavins) that diffusively shuttle electrons, conductive extracellular filaments known as bacterial nanowires, bacterial biofilms incorporating nanowires or outer membrane cytochromes, and multicellular bacterial cables that couple distant redox processes in marine sediments (6⇓⇓⇓⇓⇓⇓–13). Functionally, bacterial nanowires are thought to offer an extracellular electron transport (EET) pathway linking metal-reducing bacteria, including Shewanella and Geobacter species, to the external solid-phase iron and manganese minerals that can serve as terminal electron acceptors for respiration. In addition to the fundamental implications for respiration, EET is an especially attractive model system because it has naturally evolved to couple to inorganic systems, giving us a unique opportunity to harness biological energy conversion strategies at electrodes for electricity generation (microbial fuel cells) and production of high-value electrofuels (microbial electrosynthesis) (14).

A number of fundamental issues surrounding bacterial nanowires remain unresolved. Bacterial nanowires have never been directly observed or studied in vivo. Our direct knowledge of bacterial nanowire conductance is limited to measurements made under ex situ dry conditions using solid-state techniques optimized for inorganic nanomaterials (6, 7, 10, 11), without demonstrating the link between these conductive structures and the respiratory electron transport chains of the living cells that display them. Intense debate still surrounds the molecular makeup, identity of the charge carriers, and interfacial electron transport mechanisms responsible for the high electron mobility of bacterial nanowires. Geobacter nanowires are thought to be type IV pili, and their conductance is proposed to stem from a metallic-like band transport mechanism resulting from the stacking of aromatic amino acids along the subunit PilA (15). The latter mechanism, however, remains controversial (13, 16). In contrast, the molecular composition of bacterial nanowires from Shewanella, the best-characterized facultatively anaerobic metal reducer, has never been reported. Shewanella nanowire conductance correlates with the ability to produce outer membrane redox proteins (10), suggesting a multistep redox hopping mechanism for EET (17, 18).

The present study addresses these outstanding fundamental questions by analyzing the composition and respiratory impact of bacterial nanowires in vivo. We report an experimental system allowing real-time monitoring of individual bacterial nanowires from living Shewanella oneidensis MR-1 cells and, using fluorescent redox sensors, we demonstrate that the production of these structures correlates with cellular reductase activity. Using a combination of gene expression analysis, live fluorescence measurements, and immunofluorescence imaging, we also find that the Shewanella nanowires are membrane- rather than pilin-based, contain multiheme cytochromes, and are associated with outer membrane vesicles. Our data point to a general strategy wherein bacteria extend their outer membrane and periplasmic electron transport components, including multiheme cytochromes, micrometers away from the inner membrane.

Results and Discussion

In Vivo Imaging of Nanowire Formation.

Previous reports demonstrated increased production of bacterial nanowires and associated redox-active membrane vesicles in electron acceptor (O2)-limited Shewanella cultures (7, 10, 19). To directly observe this response in vivo, we subjected Shewanella oneidensis MR-1 to O2-limited conditions in a microliter-volume laminar perfusion flow imaging platform (Materials and Methods) and monitored the production and growth of extracellular filaments from individual cells with fluorescent microscopy. The cells and attached filamentous appendages were clearly resolved (Fig. 1 and Movies S1–S4) at the surface–solution interface using NanoOrange, a merocyanine dye that undergoes large fluorescence enhancement upon binding to proteins (20, 21). This dye was previously used to label bacterial nanowires recovered from chemostat cultures (7, 22). In all our experiments, the production of filaments coincided with the formation of separate or attached spherical membrane vesicles, another observation consistent with previous electron and atomic force microscopy measurements of Shewanella nanowires (19). In Fig. 1A, a leading membrane vesicle can be clearly seen emerging from one cell 20 min after switching to anaerobic flow conditions, followed by a trailing filament. These proteinaceous vesicle-associated filaments were widespread in all of the S. oneidensis MR-1 cultures tested; the response was displayed by 65 ± 8% of all cells (statistics obtained by monitoring 6,466 cells from multiple random fields of view in six separate biological replicates). As a representative example, Movie S5 shows an 83 × 66-μm area where the majority of cells produced the filaments. The length distribution of the filaments is plotted in SI Appendix, Fig. S1, showing an average length of 2.5 μm and reaching up to 9 μm (100 randomly selected filaments from six biological replicates).

Fig. 1.
Fig. 1.

In vivo observations of the formation and respiratory impact of bacterial nanowires in S. oneidensis MR-1. (A) A leading membrane vesicle and the subsequent growth of a bacterial nanowire observed with fluorescence from the protein stain NanoOrange. Extracellular structure formation was first observed in the t = 0 frame, captured 20 min after switching from aerobic to anaerobic perfusion. (Scale bars: 5 µm.) (B) Combined green (RedoxSensor Green) and red (FM 4-64FX) fluorescence images of a single cell before and after (35 min later) the production of a bacterial nanowire. Movie S6 shows the time-lapse movie of B. (Scale bars: 5 µm.) (C) The time-dependent RedoxSensor Green fluorescence for nanowire-producing cells, compared with neighboring cells that did not produce nanowires (n = 13, mean cellular pixel intensity ± SE). The nanowires were first observed at the t = 0 time point.

The filaments described here were the only extracellular structures observed in our experiments, and possess several features of the conductive bacterial nanowires previously reported in O2-limited chemostat cultures. Specifically, the dimensions of these filaments (10, 23, 24), their association with membrane vesicles (19), and their production during O2 limitation (7) led us to conclude that these structures are the bacterial nanowires whose conductance was previously measured ex situ under dry conditions (10). Additionally, when we labeled cells grown in O2-limited chemostat cultures with the same fluorescent dyes, we observed identical structures with the same composition as the perfusion cultures reported here (see below).

The Production of S. oneidensis MR-1 Bacterial Nanowires Is Correlated with an Increase in Cellular Reductase Activity.

To directly measure the physiological impact bacterial nanowire production has on S. oneidensis, we labeled cells with RedoxSensor Green (RSG) in the perfusion imaging platform described above. RSG is a fluorogenic dye that yields green fluorescence upon interaction with bacterial reductases in the cellular electron transport chain, and has been previously demonstrated to be an indicator of active respiration in pure cultures and environmental samples (25⇓–27). Because the redox-sensing ability of RSG was not previously characterized in Shewanella, we first confirmed that electron donor (lactate)-activated respiration increases RSG fluorescence in aerobic cultures relative to starved controls (SI Appendix, Fig. S2), and that the addition of specific electron transport inhibitors abolishes RSG fluorescence (SI Appendix, Fig. S3). S. oneidensis MR-1 cells displayed a significant increase in RSG fluorescence concomitant with nanowire production (Fig. 1 B and C and Movie S6), indicating increased respiratory activity compared with nearby control cells that did not produce nanowires in the same field of view under identical perfusion conditions. DMSO, which is respired extracellularly by Shewanella (28), was available as a terminal electron acceptor in all RSG-labeled experiments.

S. oneidensis MR-1 Nanowires Are Outer Membrane and Periplasmic Extensions.

Membrane vesicles have previously been observed to be associated with Shewanella nanowires (19) (Fig. 1A). To test the extent of membrane involvement in nanowire formation, we labeled S. oneidensis MR-1 cells with the membrane stain FM 4-64FX. This styryl dye is membrane-selective as a result of a lipophilic tail that inserts into the lipid bilayer and a positively charged head that is anchored at the membrane surface (29, 30). The amphiphilic nature of this molecule hinders it from freely crossing the membrane into the cellular interior except through the endocytic pathway, as extensively characterized in eukaryotic cells (30, 31). FM 4-64 has also been widely used in bacterial cells and shown to specifically label membranes but not extracellular protein filaments such as flagella (32, 33), except in a few bacterial species where flagella are coated in membrane sheaths (34). To our surprise, the entire length of the Shewanella nanowires was clearly stained with this reliable lipid bilayer dye (Fig. 1B, red channel, and Fig. 2), indicating that membranes are a substantial component of Shewanella nanowires, contrary to previous suggestions that these structures are pilin based. We stained cells producing both nanowires and membrane vesicles with NanoOrange and FM 4-64FX, demonstrating that proteins and lipid colocalized on these extracellular structures, consistent with being derived from the cell envelope (Fig. 2A).

Fig. 2.
Fig. 2.

Bacterial nanowires contain lipids, proteins, and periplasm. (A) Shewanella oneidensis MR-1 cells and attached bacterial nanowires, stained with both NanoOrange (Upper) and FM 4-64FX (Lower), indicating the presence of proteins and membranes, respectively. (Scale bars: 5 µm.) (B) Bacterial nanowires from S. oneidensis MR-1 strains containing GFP only in the cytoplasm (Upper) or in the periplasm as well (Lower). The green and red channels monitor GFP and FM 4-64FX fluorescence, respectively. The nanowires display green fluorescence only when GFP is present in the periplasm (Lower Left). (Scale bars: 2 µm.)

Most known bacterial vesicles are composed primarily of outer membrane and periplasm. To determine whether Shewanella nanowires contain periplasm, we expressed either GFP fused to a signal sequence that enables GFP export to the periplasm (SI Appendix, Fig. S4) after folding in the cytoplasm (35) or YFP fused to the S. oneidensis periplasmic [Fe–Fe] hydrogenase large subunit HydA (SI Appendix). We observed fluorescence along the bacterial nanowires in both constructs (Fig. 2B and SI Appendix, Fig. S5). However, no fluorescence was detected along nanowires from a strain expressing cytoplasmic-only GFP (Fig. 2B). Taken together, these results indicate that Shewanella nanowires are outer membrane extensions containing soluble periplasmic components.

It has previously been proposed, but never demonstrated, that pili are important components of Shewanella nanowires. To test whether pili play a role in S. oneidensis nanowire production, we harvested RNA (36) from chemostat cultures where O2 served as the only electron acceptor, before and at time intervals after transition from electron donor (lactate) limitation to O2 limitation. Electron acceptor limitation is known to result in increased production of bacterial nanowires, as previously demonstrated in chemostat cultures (7, 10) (and confirmed by electron microscopy). We then used qPCR to measure changes in the expression of key genes necessary for type IV pilus assembly: pilA, encoding the type IV major pilin subunit; mshA, encoding the mannose sensitive hemagglutinin (msh) pilin major subunit; and pilE and fimT, encoding type IV minor pilin proteins. Expression of all these pilin genes either remained constant or decreased after electron acceptor limitation, when nanowire production was observed (Fig. 3A). Furthermore, mutants lacking the type IV pilin major subunit (ΔpilA) or both the type IV and msh pilus biogenesis systems (ΔpilM–Q/ΔmshH–Q) (37) produced bacterial nanowires and displayed an increase in reductase activity in the perfusion imaging platform (Fig. 3B and SI Appendix, Fig. S6)—a response identical to wild-type S. oneidensis MR-1. The chemostat qPCR and perfusion pilus-deletion observations both support the conclusion that Shewanella nanowires are distinct from pili.

Fig. 3.
Fig. 3.

The genetic and molecular basis of bacterial nanowires. (A) Expression of pilin genes in S. oneidensis MR-1 measured by qPCR. Chemostat cultures were grown under aerobic conditions (dissolved oxygen tension of 20%) at 30 °C for 48 h before the dissolved oxygen tension (DOT) was reduced to 0%. Samples were harvested from cultures right before reducing the DOT (t < 0, reference sample), at t = 0, and in 15-min intervals for 1 h. The fold change in gene expression relative to the reference sample was calculated by 2−ΔΔCT from at least four reactions of three independent chemostat cultures, using recA for normalization. (B) Combined green (RedoxSensor Green) and red (FM 4-64FX) fluorescence images of the ΔpilA strain, lacking the type IV pilin major subunit PilA, before and after (95 min later) the production of a bacterial nanowire. ΔpilA is capable of producing bacterial nanowires with a similar respiratory impact as wild-type S. oneidensis MR-1, evidenced by the increase in reductase activity (green fluorescence) after nanowire production. (Scale bars: 2 µm.) (C) Expression of key genes mtrC, mtrA, and omcA encoding extracellular electron transport proteins from the same chemostat cultures as A. (D) Labeling with antibodies against MtrC (Left) or OmcA (Right) and membrane fluorescence (FM 4-64FX) images of wild-type S. oneidensis MR-1 (Upper) compared with the ΔmtrC/omcA control strain (Lower). Nanowire-localized MtrC and OmcA are observed in the wild-type strain. (Scale bars: 2 µm.)

Why were the membrane and periplasmic components of these structures overlooked in previous studies? One important factor is the difficulty of isolating the appendages and separating them from cells before downstream compositional analysis (e.g., liquid chromatography–tandem mass spectrometry). In fact, membrane and periplasmic proteins were previously identified in a study focused on developing optimal methods for removing the Shewanella filaments, although it was not possible to rule these proteins out as an artifact of the shearing procedure (38). The present in vivo microscopy work circumvents some of the previous artifact problems by fluorescently labeling specific cellular components (protein, lipid, and periplasm) on intact nanowires attached to whole cells. In light of these new results, we revisited the chemostat samples from our previous conductance study, and noted that in those samples, SEM images also revealed vesicular morphologies, nonuniform cross-sections, and diameters >5 nm (larger than pili; SI Appendix, Fig. S7A and the source-drain devices in ref. 10). These suggestive links did not become clear until the present in vivo results. We fixed samples from O2-limited chemostat cultures and labeled them with FM 4-64FX. The nanowires from chemostat cultures contained both protein and membrane (SI Appendix, Fig. S7B), further demonstrating that these structures are the same as those observed in the perfusion cultures in vivo. Though the net patterns of gene expression measured in the planktonic chemostat cultures may differ from the surface-attached perfusion cultures, it is important to stress that identical membrane extension phenotypes were observed in both these experiments where O2 served as the sole electron acceptor in limiting concentrations.

Localization of the Decaheme Cytochromes MtrC and OmcA Along Nanowires.

As a metal reducer, Shewanella has evolved an intricate EET pathway to traffic electrons from the inner membrane, through the periplasm, and across the outer membrane to external electron acceptors, including minerals and electrodes. This pathway includes the periplasmic decaheme cytochrome MtrA, as well as the outer membrane decaheme cytochromes MtrC and OmcA, which may interface to soluble redox shuttles or, via solvent-exposed hemes, directly to the insoluble terminal acceptors (39). Previous work has shown that S. oneidensis cells lacking mtrC and omcA produce nanowires that are not capable of electron transport (10). In light of the structural finding that Shewanella nanowires are outer membrane and periplasmic extensions, we examined whether expression of these periplasmic and outer membrane cytochromes increases during nanowire production. We measured the expression levels of mtrA, mtrC, omcA, and dmsE, a periplasmic decaheme cytochrome required for maximal extracellular respiration of DMSO (28), during and after the transition to O2 limitation in chemostat cultures. These EET components had significantly increased expression in response to electron acceptor limitation (Fig. 3C and SI Appendix, Fig. S8). To determine whether these cytochromes are indeed components of the membrane extensions, we performed immunofluorescence with MtrC- and OmcA-specific antibodies (40) following in vivo observation of the target nanowires in the perfusion imaging platform (Movies S7 and S8). We observed MtrC and OmcA localization at the periphery of the cell, as expected. We also observed clear localization of these cytochromes along the membrane-stained bacterial nanowires (25 of 35 nanowires labeled with anti-MtrC and 19 of 22 nanowires labeled with anti-OmcA), whereas no fluorescence was detected from ΔmtrC/omcA negative control cells or their membrane extensions (none of 22 nanowires labeled with anti-MtrC and none of 20 nanowires labeled with anti-OmcA; Fig. 3D).

Though the conductance of Shewanella nanowires was previously only demonstrated under nonphysiological conditions (10), the data reported here are consistent with membrane extensions that could function as nanowires to mediate EET from live cells. Localization of MtrC and OmcA to these membrane extensions provides the most compelling evidence to date, and directly supports the proposed multistep redox hopping mechanism (13, 17, 18), allowing long-range electron transport along a membrane network of heme cofactors that line Shewanella nanowires (Fig. 4). We have shown that S. oneidensis nanowires contain periplasm (Fig. 2B); therefore, it is also possible that periplasmic proteins and soluble redox cofactors may contribute to electron transport through these extensions.

Fig. 4.
Fig. 4.

Proposed structural model for Shewanella nanowires. S. oneidensis MR-1 nanowires are outer membrane (OM) and periplasmic (PP) extensions including the multiheme cytochromes responsible for extracellular electron transport.

Intermediate Steps in Nanowire Formation.

The extension of outer membrane filaments, and their functionalization with electron transport proteins, may represent a widespread strategy for EET. Virtually all Gram-negative bacteria produce outer membrane vesicles, and can alter the rate of production and composition of those vesicles in response to various stress conditions (19, 41). More recent electron microscopy reports describe membrane vesicle chains and related membrane tubes (also referred to as periplasmic tubules) that form cell–cell connections in the social soil bacterium Myxococcus xanthus (42) as well as the phototrophic consortium Chlorochromatium aggregatum (43). Consistent with these reports, we observed both a transition from vesicle chains to smoother filaments (Movie S9), as well as nanowires connecting separate Shewanella cells (Movie S4).

To gain a clearer picture of the role of membrane vesicles in Shewanella nanowire formation, we performed atomic force microscopy (AFM) on the same bacterial nanowires after observing their growth with fluorescent imaging under perfusion flow conditions. Such observations were not possible in steady-state chemostat cultures where the nanowire growth is not confined to the surface–solution interface. Perfusion was stopped and the samples were fixed quickly after observing the early signs of nanowire production, allowing us to examine the initial stages of nanowire formation with nanoscale resolution using AFM (Fig. 5). We measured the nanowires to be 10 nm in diameter under dry conditions, consistent with previous observations (10, 23, 24) and roughly corresponding to two lipid bilayers. In addition, we were able to resolve different morphologies corresponding to different stages of nanowire formation (Fig. 5), consistent with the chain-to-filament transition in Movie S9. The morphologies observed ranged from vesicle chains (Fig. 5A) to partially smooth filaments incorporating vesicles (Fig. 5B, also consistent with SEM imaging in SI Appendix, Fig. S7A), and finally continuous filaments (Fig. 5C). In addition to possibly mediating EET up to micrometers away from the inner membrane, the vesiculation and extension of outer membranes into the quasi one-dimensional morphologies observed here increase the surface area-to-volume ratio of cells. This shape change can present a significant advantage, increasing the likelihood cells will encounter the solid-phase minerals that serve as electron acceptors for respiration.

Fig. 5.
Fig. 5.

Correlated atomic force microscopy (AFM) and live-cell membrane fluorescence of bacterial nanowires. (A–C) Tapping AFM phase images of S. oneidensis MR-1 cells after producing bacterial nanowires in the perfusion flow system. The sample is fixed and air-dried before AFM imaging. (Scale bars: 2 µm.) (Insets) In vivo fluorescence images of the same cells/nanowires at the surface/solution interface in the perfusion platform. The cells and the nanowires are stained by the membrane stain FM 4-64FX. (Scale bars: 1 µm.) The morphologies observed range from vesicle chains (A) to partially smooth filaments incorporating vesicles (B), which is consistent with SEM imaging of chemostat samples (SI Appendix, Fig. S7A), and, finally, continuous filaments (C). See also Movie S9, which captures the transition from a vesicle chain to a continuous bacterial nanowire.

Given the ubiquity of membrane vesicles and related extensions in Gram-negative bacteria, the localization of electron transport proteins along membrane extensions in a manner consistent with bacterial nanowires that could mediate extracellular electron transport, and the finding of nanowire-based cell–cell connectivity, our results raise the intriguing possibility of redox-functionalized membrane extensions as a general microbial strategy for EET and cell–cell signaling. Our study motivates further experimental and theoretical work to build a detailed understanding of the full biomolecular makeup, electron transport physics, and physiological impact of bacterial nanowires.

Materials and Methods

Cell Growth Conditions.

All Shewanella strains were grown aerobically in LB broth from a frozen (−80 °C) stock, at 30 °C, up to an OD600 of 2.4–2.8. The cells from these precultures were pelleted by centrifugation at 4226 × g for 5 min, washed twice, and finally resuspended in a defined medium consisting of 30 mM Pipes, 60 mM sodium DL-lactate as an electron donor, 28 mM NH4Cl, 1.34 mM KCl, 4.35 mM NaH2PO4, 7.5 mM NaOH, 30 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, and 0.05 mM ferric nitrilotriacetic acid. In addition, vitamins, amino acids, and trace mineral stock solutions were used to supplement the medium as described previously (7). The medium was adjusted to an initial pH of 7.2.

Perfusion Chamber Conditions.

Washed cells were slowly injected into the perfusion chamber (d = 250 μm, w = 9,580 μm, L = 7,110 μm, described in detail in SI Appendix) and the flow was stopped to allow the cells to settle on the coverslip with a surface density of ∼100 cells in an 83 × 66-µm field of view while being imaged by the inverted microscope. After inoculation, sterile defined medium (100 mL in an inverted 135-mL sealed glass serum bottle) was connected to the perfusion chamber inlet and passed through at a flow rate of 5 ± 1 µL/s that remained constant for 3 h.

As previously detailed (7, 19), transition to electron–acceptor limitation was required to promote the production of bacterial nanowires and membrane vesicles; this was accomplished using one of two methods. In the first method, the experiment is started with fully aerobic medium flow before switching to supply bottles containing medium made anaerobic by boiling and purging with 100% N2. This method has the advantage of providing a precise time point for entering acceptor (O2) limitation (Fig. 1A and Movies S1 and S4). In the second method, O2 becomes limited close to the coverslip surface at high cell density, even with aerobic medium, as a result of the laminar flow (no mixing between adjacent layers) and no-slip condition (zero velocity at the surface); this is confirmed with the simple calculation outlined in SI Appendix. Using method 2, nanowires and membrane vesicles were consistently observed after a lag period of 90–120 min from the start of perfusion flow (Movies S2 and S3). We observed higher nanowire production rates and better consistency using this method, possibly because the slower transition exposed the surface-bound cells to a wide range of acceptor concentrations, compared with the abrupt transition in the first method. Both methods resulted in identical membrane extensions as observed using membrane fluorescence (FM 4-64FX), protein fluorescence (NanoOrange), and the accompanying increase in reductase activity (RedoxSensor Green).

Immunofluorescence with MtrC or OmcA Antibody.

S. oneidensis MR-1 and ∆omcA/mtrC (SI Appendix, Table S2) were used in the perfusion chamber experiment as described above. As soon as the nanowires were produced and observed through staining by FM 4-64FX, the media flow was stopped and the chamber was opened under sterile medium such that the coverslip remained hydrated. The sample (coverslip with attached cells) was fixed with 4% (vol/vol) formaldehyde solution in PBS for 1 h at room temperature (RT). After rinsing in PBS, the sample was incubated in 0.15% glycine at RT for 5 min to quench free aldehyde groups and reduce background fluorescence. The sample was then transferred to a blocking solution of 1% BSA in PBS for 5 min, and reacted with the diluted polyclonal rabbit-raised MtrC or OmcA-specific primary antibody (40) at RT for 30 min (MtrC Ab: 2.6 µg/mL, OmcA Ab: 1.9 µg/mL, both in 1% BSA/PBS). After rinsing four times in PBS, the sample was incubated with anti-rabbit FITC-conjugated secondary antibody (Thermo Scientific Pierce antibodies, catalog no. 31635; 7.5 µg/mL in 1% BSA/PBS) at RT for 30 min. Finally, the coverslip was rinsed twice in PBS before immunofluorescence imaging in the green channel. To perform immunofluorescence on the same cells and nanowires observed during live imaging (Movies S7 and S8 and Fig. 3D), we modified the coverslips with surface scratches that acted as fiducial markers.

Chemostat Growth and qPCR Analysis of the Transition from Electron-Donor to Electron-Acceptor Limitation.

S. oneidensis MR-1 was grown in chemostat medium (SI Appendix, Table S3) at 30 °C and 20% dissolved oxygen tension using continuous flow bioreactors (BioFlo 110; New Brunswick Scientific) with a dilution rate of 0.05 h−1 and an operating liquid volume of 1 L, as previously described (7). After 48 h of this aerobic growth, a reference sample was taken. The dissolved oxygen tension was then manually dropped to 0% by adjusting the N2/air mixture entering the reactor, and the first sample after electron acceptor limitation (t = 0) was taken at this time. O2 served as the sole terminal electron acceptor throughout the experiment. Samples were subsequently taken at 15-min intervals for 1 h. At each time point, 10 mL of cells were harvested in ice-cold 5% citrate-saturated phenol in ethanol to prevent further transcription and protect the RNA. Samples were taken from three independent biological replicates (different chemostats). Total RNA was prepared using a hot phenol extraction, as previously described (36). Five micrograms of total RNA from each of the time points described above was used as input for reverse-transcriptase reactions with the SuperScript III First-Strand Synthesis System, as per the manufacturer’s protocol (Life Technologies). Subsequent cDNA was then diluted and used as template in qPCR experiments with SYBR Select Master Mix (Life Technologies). Fold change in gene expression relative to the reference sample was calculated by 2−ΔΔCT from at least four reactions of three independent biological replicate samples, using recA for normalization. Similar results were obtained using rpoB for normalization. The sequences of the primers used for each gene are shown in SI Appendix, Table S4.

Acknowledgments

The pHGE-PtacTorAGFP plasmid was generously provided by Prof. H. Gao (Zhejiang University), and pProbeNT was kindly provided by Dr. Steven Lindow (University of California, Berkeley). Atomic Force and Electron Microscopy were performed at the University of Southern California Centers of Excellence in NanoBioPhysics and Electron Microscopy and Microanalysis. The development of the in vivo imaging platform and chemostat cultivation was funded by Air Force Office of Scientific Research Young Investigator Research Program Grant FA9550-10-1-0144 (to M.Y.E.-N.). Redox sensing measurements, compositional analysis, and the localization of multiheme cytochromes were funded by Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the US Department of Energy Grant DE-FG02-13ER16415 (to M.Y.E.-N.). RT-PCR experiments and genetic analyses were funded by National Science Foundation Grant EF-1104831 (to J.H.G.). M.F.R., S.B.R., R.A.B., and D.A.S. were supported under the Shewanella Federation consortium funded by the Genomics: Genomes to Life program of the US Department of Energy Office of Biological and Environmental Research.

Footnotes

  • ↵1To whom correspondence should be addressed. Email: mnaggar{at}usc.edu.

  • Author contributions: S.P., S.E.B., J.H.G., and M.Y.E.-N. designed research; S.P., S.E.B., K.M.L., H.S.B., Y.J., and Y.A.G. performed research; R.A.B., S.B.R., M.F.R., D.A.S., and L.S. contributed new reagents/analytic tools; S.P., S.E.B., J.H.G., and M.Y.E.-N. analyzed data; and S.P., S.E.B., J.H.G., and M.Y.E.-N. wrote the paper.

  • The authors declare no conflict of interest.

  • This article is a PNAS Direct Submission.

  • This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1410551111/-/DCSupplemental.

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S. oneidensis nanowires are membrane extensions
Sahand Pirbadian, Sarah E. Barchinger, Kar Man Leung, Hye Suk Byun, Yamini Jangir, Rachida A. Bouhenni, Samantha B. Reed, Margaret F. Romine, Daad A. Saffarini, Liang Shi, Yuri A. Gorby, John H. Golbeck, Mohamed Y. El-Naggar
Proceedings of the National Academy of Sciences Sep 2014, 111 (35) 12883-12888; DOI: 10.1073/pnas.1410551111
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