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

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).

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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).

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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.

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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.

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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.

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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.