Shewanella oneidensis MR-1 nanowires are outermembrane and periplasmic extensions of theextracellular electron transport componentsSahand Pirbadiana, Sarah E. Barchingerb, Kar Man Leunga, Hye Suk Byuna, Yamini Jangira, Rachida A. Bouhennic,Samantha B. Reedd, Margaret F. Romined, Daad A. Saffarinic, Liang Shid, Yuri A. Gorbye, John H. Golbeckb,f,and Mohamed Y. El-Naggara,g,1

aDepartment of Physics and Astronomy, University of Southern California, Los Angeles, CA 90089; bDepartment of Biochemistry and Molecular Biology,Pennsylvania State University, University Park, PA 16802; cDepartment of Biological Sciences, University of Wisconsin, Milwaukee, WI 53211; dPacificNorthwest National Laboratory, Richland, WA 99354; eDepartment of Civil and Environmental Engineering, Rensselaer Polytechnic Institute, Troy, NY 12180;fDepartment of Chemistry, Pennsylvania State University, University Park, PA 16802; and gMolecular and Computational Biology Section, Department ofBiological Sciences, University of Southern California, Los Angeles, CA 90089

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

Bacterial nanowires offer an extracellular electron transport (EET)pathway for linking the respiratory chain of bacteria to externalsurfaces, including oxidized metals in the environment andengineered electrodes in renewable energy devices. Despite theglobal, environmental, and technological consequences of thisbiotic–abiotic interaction, the composition, physiological relevance,and electron transport mechanisms of bacterial nanowires remainunclear. We report, to our knowledge, the first in vivo observationsof the formation and respiratory impact of nanowires in the modelmetal-reducing microbe Shewanella oneidensis MR-1. Live fluores-cence measurements, immunolabeling, and quantitative gene expres-sion analysis point to S. oneidensis MR-1 nanowires as extensions ofthe outer membrane and periplasm that include the multiheme cyto-chromes responsible for EET, rather than pilin-based structures as pre-viously thought. These membrane extensions are associated withouter membrane vesicles, structures ubiquitous in Gram-negativebacteria, and are consistent with bacterial nanowires that mediatelong-range EET by the previously proposed multistep redox hoppingmechanism. Redox-functionalized membrane and vesicular exten-sions may represent a general microbial strategy for electron trans-port and energy distribution.

extracellular electron transfer | bioelectronics | respiration |membrane cytochromes

Reduction–oxidation (redox) reactions and electron transportare essential to the energy conversion pathways of living cells

(1). Respiratory organisms generate ATP molecules—life’s uni-versal energy currency—by harnessing the free energy of elec-tron transport from electron donors (fuels) to electron acceptors(oxidants) through biological redox chains. In contrast to mosteukaryotes, which are limited to relatively few carbon com-pounds as electron donors and oxygen as the predominantelectron acceptor, prokaryotes have evolved into versatile energyscavengers. Microbes can wield an astounding number of meta-bolic pathways to extract energy from diverse organic and in-organic electron donors and acceptors, which has significantconsequences 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 fa-cilitating electron transfer (1). Recently, microbial electrontransport across dramatically longer distances has been reported,ranging from nanometers to micrometers (cell lengths) and evencentimeters (5). A few strategies have been proposed to mediatethis long-distance electron transport in various microbial sys-tems: soluble redox mediators (e.g., flavins) that diffusivelyshuttle electrons, conductive extracellular filaments known asbacterial nanowires, bacterial biofilms incorporating nanowires

or outer membrane cytochromes, and multicellular bacterialcables that couple distant redox processes in marine sediments(6–13). Functionally, bacterial nanowires are thought to offer anextracellular electron transport (EET) pathway linking metal-reducing bacteria, including Shewanella andGeobacter species, tothe external solid-phase iron and manganese minerals that canserve as terminal electron acceptors for respiration. In additionto the fundamental implications for respiration, EET is an es-pecially attractive model system because it has naturally evolvedto couple to inorganic systems, giving us a unique opportunity toharness biological energy conversion strategies at electrodes forelectricity generation (microbial fuel cells) and production ofhigh-value electrofuels (microbial electrosynthesis) (14).A number of fundamental issues surrounding bacterial nano-

wires remain unresolved. Bacterial nanowires have never beendirectly observed or studied in vivo. Our direct knowledge ofbacterial nanowire conductance is limited to measurementsmade under ex situ dry conditions using solid-state techniquesoptimized for inorganic nanomaterials (6, 7, 10, 11), withoutdemonstrating the link between these conductive structures andthe respiratory electron transport chains of the living cells that

Significance

Bacterial nanowires from Shewanella oneidensis MR-1 werepreviously shown to be conductive under nonphysiologicalconditions. Intense debate still surrounds the molecularmakeup, identity of the charge carriers, and cellular respiratoryimpact of bacterial nanowires. In this work, using in vivofluorescence measurements, immunolabeling, and quantitativegene expression analysis, we demonstrate that S. oneidensisMR-1 nanowires are extensions of the outer membrane andperiplasm, rather than pilin-based structures, as previouslythought. We also demonstrate that the outer membrane mul-tiheme cytochromes MtrC and OmcA localize to these mem-brane extensions, directly supporting one of the two models ofelectron transport through the nanowires; consistent with this,production of bacterial nanowires correlates with an increasein cellular reductase activity.

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. contributednew 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.1To whom correspondence should be addressed. Email: mnaggar@usc.edu.

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

www.pnas.org/cgi/doi/10.1073/pnas.1410551111 PNAS | September 2, 2014 | vol. 111 | no. 35 | 12883–12888

MICRO

BIOLO

GY

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 17

, 202

0

display them. Intense debate still surrounds the molecularmakeup, identity of the charge carriers, and interfacial electrontransport mechanisms responsible for the high electron mobilityof bacterial nanowires. Geobacter nanowires are thought to betype IV pili, and their conductance is proposed to stem froma metallic-like band transport mechanism resulting from thestacking of aromatic amino acids along the subunit PilA (15).The latter mechanism, however, remains controversial (13, 16).In contrast, the molecular composition of bacterial nanowiresfrom Shewanella, the best-characterized facultatively anaerobicmetal reducer, has never been reported. Shewanella nanowireconductance correlates with the ability to produce outer mem-brane redox proteins (10), suggesting a multistep redox hoppingmechanism for EET (17, 18).The present study addresses these outstanding fundamental

questions by analyzing the composition and respiratory impact ofbacterial nanowires in vivo. We report an experimental systemallowing real-time monitoring of individual bacterial nanowiresfrom living Shewanella oneidensis MR-1 cells and, using fluo-rescent redox sensors, we demonstrate that the production ofthese structures correlates with cellular reductase activity. Usinga combination of gene expression analysis, live fluorescencemeasurements, and immunofluorescence imaging, we also findthat the Shewanella nanowires are membrane- rather than pilin-based, contain multiheme cytochromes, and are associated withouter membrane vesicles. Our data point to a general strategywherein bacteria extend their outer membrane and periplasmicelectron transport components, including multiheme cyto-chromes, micrometers away from the inner membrane.

Results and DiscussionIn Vivo Imaging of Nanowire Formation. Previous reports demon-strated increased production of bacterial nanowires and associ-ated redox-active membrane vesicles in electron acceptor(O2)-limited Shewanella cultures (7, 10, 19). To directly observethis response in vivo, we subjected Shewanella oneidensis MR-1to O2-limited conditions in a microliter-volume laminar perfu-sion flow imaging platform (Materials and Methods) and moni-tored the production and growth of extracellular filaments fromindividual cells with fluorescent microscopy. The cells and at-tached filamentous appendages were clearly resolved (Fig. 1 andMovies S1–S4) at the surface–solution interface using Nano-Orange, a merocyanine dye that undergoes large fluorescenceenhancement upon binding to proteins (20, 21). This dye waspreviously used to label bacterial nanowires recovered from

chemostat cultures (7, 22). In all our experiments, the productionof filaments coincided with the formation of separate or attachedspherical membrane vesicles, another observation consistentwith previous electron and atomic force microscopy measure-ments of Shewanella nanowires (19). In Fig. 1A, a leadingmembrane vesicle can be clearly seen emerging from one cell 20min after switching to anaerobic flow conditions, followed bya trailing filament. These proteinaceous vesicle-associated fila-ments were widespread in all of the S. oneidensis MR-1 culturestested; the response was displayed by 65 ± 8% of all cells (sta-tistics obtained by monitoring 6,466 cells from multiple randomfields of view in six separate biological replicates). As a repre-sentative example, Movie S5 shows an 83 × 66-μm area wherethe majority of cells produced the filaments. The length distri-bution of the filaments is plotted in SI Appendix, Fig. S1, showingan average length of 2.5 μm and reaching up to 9 μm (100 ran-domly selected filaments from six biological replicates).The filaments described here were the only extracellular

structures observed in our experiments, and possess several fea-tures of the conductive bacterial nanowires previously reported inO2-limited chemostat cultures. Specifically, the dimensions ofthese filaments (10, 23, 24), their association with membranevesicles (19), and their production during O2 limitation (7) led usto conclude that these structures are the bacterial nanowireswhose conductance was previously measured ex situ under dryconditions (10). Additionally, when we labeled cells grown inO2-limited chemostat cultures with the same fluorescent dyes, weobserved identical structures with the same composition as theperfusion cultures reported here (see below).

The Production of S. oneidensis MR-1 Bacterial Nanowires Is Correlatedwith an Increase in Cellular Reductase Activity. To directly measure thephysiological impact bacterial nanowire production has on S. onei-densis, we labeled cells with RedoxSensor Green (RSG) in theperfusion imaging platform described above. RSG is a fluorogenicdye that yields green fluorescence upon interaction with bacterialreductases in the cellular electron transport chain, and has beenpreviously demonstrated to be an indicator of active respirationin pure cultures and environmental samples (25–27). Because theredox-sensing ability of RSG was not previously characterized inShewanella, we first confirmed that electron donor (lactate)-activatedrespiration increases RSG fluorescence in aerobic cultures relative tostarved controls (SI Appendix, Fig. S2), and that the addition ofspecific electron transport inhibitors abolishes RSG fluores-cence (SI Appendix, Fig. S3). S. oneidensis MR-1 cells displayed

Fig. 1. In vivo observations of the formation and re-spiratory impact of bacterial nanowires in S. oneidensisMR-1. (A) A leading membrane vesicle and the sub-sequent growth of a bacterial nanowire observed withfluorescence from the protein stain NanoOrange. Ex-tracellular structure formation was first observed in thet = 0 frame, captured 20 min after switching fromaerobic to anaerobic perfusion. (Scale bars: 5 μm.) (B)Combined green (RedoxSensor Green) and red (FM4-64FX) fluorescence images of a single cell beforeand after (35 min later) the production of a bacterialnanowire. Movie S6 shows the time-lapse movieof B. (Scale bars: 5 μm.) (C ) The time-dependentRedoxSensor Green fluorescence for nanowire-producing cells, compared with neighboring cellsthat did not produce nanowires (n = 13, mean cel-lular pixel intensity ± SE). The nanowires were firstobserved at the t = 0 time point.

12884 | www.pnas.org/cgi/doi/10.1073/pnas.1410551111 Pirbadian et al.

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 17

, 202

0

a significant increase in RSG fluorescence concomitant with nano-wire production (Fig. 1 B and C andMovie S6), indicating increasedrespiratory activity compared with nearby control cells that did notproduce nanowires in the same field of view under identical per-fusion conditions. DMSO, which is respired extracellularly by She-wanella (28), was available as a terminal electron acceptor in allRSG-labeled experiments.

S. oneidensis MR-1 Nanowires Are Outer Membrane and PeriplasmicExtensions. Membrane vesicles have previously been observed tobe associated with Shewanella nanowires (19) (Fig. 1A). To testthe extent of membrane involvement in nanowire formation, welabeled S. oneidensis MR-1 cells with the membrane stain FM4-64FX. This styryl dye is membrane-selective as a result of a li-pophilic tail that inserts into the lipid bilayer and a positivelycharged head that is anchored at the membrane surface (29, 30).The amphiphilic nature of this molecule hinders it from freelycrossing the membrane into the cellular interior except throughthe endocytic pathway, as extensively characterized in eukaryoticcells (30, 31). FM 4-64 has also been widely used in bacterial cellsand shown to specifically label membranes but not extracellularprotein filaments such as flagella (32, 33), except in a few bac-terial species where flagella are coated in membrane sheaths(34). To our surprise, the entire length of the Shewanella nano-wires was clearly stained with this reliable lipid bilayer dye (Fig.1B, red channel, and Fig. 2), indicating that membranes area substantial component of Shewanella nanowires, contrary toprevious suggestions that these structures are pilin based. Westained cells producing both nanowires and membrane vesicleswith NanoOrange and FM 4-64FX, demonstrating that proteinsand lipid colocalized on these extracellular structures, consistentwith being derived from the cell envelope (Fig. 2A).Most known bacterial vesicles are composed primarily of outer

membrane and periplasm. To determine whether Shewanellananowires contain periplasm, we expressed either GFP fused toa signal sequence that enables GFP export to the periplasm (SIAppendix, Fig. S4) after folding in the cytoplasm (35) or YFP fusedto the S. oneidensis periplasmic [Fe–Fe] hydrogenase large subunitHydA (SI Appendix). We observed fluorescence along the bacterialnanowires in both constructs (Fig. 2B and SI Appendix, Fig. S5).However, no fluorescence was detected along nanowires froma strain expressing cytoplasmic-only GFP (Fig. 2B). Taken together,

these results indicate that Shewanella nanowires are outer mem-brane extensions containing soluble periplasmic components.It has previously been proposed, but never demonstrated, that

pili are important components of Shewanella nanowires. To testwhether pili play a role in S. oneidensis nanowire production, weharvested RNA (36) from chemostat cultures where O2 served asthe only electron acceptor, before and at time intervals aftertransition from electron donor (lactate) limitation to O2 limita-tion. Electron acceptor limitation is known to result in increasedproduction of bacterial nanowires, as previously demonstrated inchemostat cultures (7, 10) (and confirmed by electron micros-copy). We then used qPCR to measure changes in the expressionof key genes necessary for type IV pilus assembly: pilA, encodingthe type IV major pilin subunit; mshA, encoding the mannosesensitive hemagglutinin (msh) pilin major subunit; and pilE andfimT, encoding type IV minor pilin proteins. Expression of allthese pilin genes either remained constant or decreased afterelectron acceptor limitation, when nanowire production was ob-served (Fig. 3A). Furthermore, mutants lacking the type IV pilinmajor subunit (ΔpilA) or both the type IV and msh pilus biogenesissystems (ΔpilM–Q/ΔmshH–Q) (37) produced bacterial nanowiresand displayed an increase in reductase activity in the perfusionimaging platform (Fig. 3B and SI Appendix, Fig. S6)—a responseidentical to wild-type S. oneidensis MR-1. The chemostat qPCRand perfusion pilus-deletion observations both support the con-clusion that Shewanella nanowires are distinct from pili.Why were the membrane and periplasmic components of these

structures overlooked in previous studies? One important factor isthe difficulty of isolating the appendages and separating them fromcells before downstream compositional analysis (e.g., liquid chro-matography–tandem mass spectrometry). In fact, membrane andperiplasmic proteins were previously identified in a study focusedon developing optimal methods for removing the Shewanella fila-ments, although it was not possible to rule these proteins out as anartifact of the shearing procedure (38). The present in vivo mi-croscopy work circumvents some of the previous artifact problemsby 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 samplesfrom our previous conductance study, and noted that in thosesamples, SEM images also revealed vesicular morphologies, non-uniform cross-sections, and diameters >5 nm (larger than pili; SIAppendix, Fig. S7A and the source-drain devices in ref. 10). Thesesuggestive links did not become clear until the present in vivoresults. We fixed samples from O2-limited chemostat cultures andlabeled them with FM 4-64FX. The nanowires from chemostatcultures contained both protein and membrane (SI Appendix, Fig.S7B), further demonstrating that these structures are the same asthose observed in the perfusion cultures in vivo. Though the netpatterns of gene expression measured in the planktonic chemostatcultures may differ from the surface-attached perfusion cultures, itis important to stress that identical membrane extension pheno-types were observed in both these experiments where O2 served asthe sole electron acceptor in limiting concentrations.

Localization of the Decaheme Cytochromes MtrC and OmcA AlongNanowires. As a metal reducer, Shewanella has evolved an in-tricate EET pathway to traffic electrons from the inner mem-brane, through the periplasm, and across the outer membrane toexternal electron acceptors, including minerals and electrodes.This pathway includes the periplasmic decaheme cytochromeMtrA, as well as the outer membrane decaheme cytochromesMtrC and OmcA, which may interface to soluble redox shuttlesor, via solvent-exposed hemes, directly to the insoluble terminalacceptors (39). Previous work has shown that S. oneidensis cellslacking mtrC and omcA produce nanowires that are not capableof electron transport (10). In light of the structural findingthat Shewanella nanowires are outer membrane and periplasmic

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

Pirbadian et al. PNAS | September 2, 2014 | vol. 111 | no. 35 | 12885

MICRO

BIOLO

GY

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 17

, 202

0

extensions, we examined whether expression of these periplasmicand outer membrane cytochromes increases during nanowireproduction. We measured the expression levels of mtrA, mtrC,omcA, and dmsE, a periplasmic decaheme cytochrome requiredfor maximal extracellular respiration of DMSO (28), during andafter the transition to O2 limitation in chemostat cultures. TheseEET components had significantly increased expression in re-sponse to electron acceptor limitation (Fig. 3C and SI Appendix,Fig. S8). To determine whether these cytochromes are indeedcomponents of the membrane extensions, we performed immu-nofluorescence with MtrC- and OmcA-specific antibodies (40)following in vivo observation of the target nanowires in theperfusion imaging platform (Movies S7 and S8). We observedMtrC and OmcA localization at the periphery of the cell, asexpected. We also observed clear localization of these cyto-chromes along the membrane-stained bacterial nanowires (25 of35 nanowires labeled with anti-MtrC and 19 of 22 nanowireslabeled with anti-OmcA), whereas no fluorescence was detectedfrom ΔmtrC/omcA negative control cells or their membraneextensions (none of 22 nanowires labeled with anti-MtrC andnone of 20 nanowires labeled with anti-OmcA; Fig. 3D).Though the conductance of Shewanella nanowires was pre-

viously only demonstrated under nonphysiological conditions(10), the data reported here are consistent with membraneextensions that could function as nanowires to mediate EETfrom live cells. Localization of MtrC and OmcA to these mem-brane extensions provides the most compelling evidence to date,and directly supports the proposed multistep redox hoppingmechanism (13, 17, 18), allowing long-range electron transportalong a membrane network of heme cofactors that line Shewa-nella nanowires (Fig. 4). We have shown that S. oneidensisnanowires contain periplasm (Fig. 2B); therefore, it is also pos-sible that periplasmic proteins and soluble redox cofactors maycontribute to electron transport through these extensions.

Intermediate Steps in Nanowire Formation. The extension of outermembrane filaments, and their functionalization with electrontransport proteins, may represent a widespread strategy for EET.Virtually all Gram-negative bacteria produce outer membranevesicles, and can alter the rate of production and composition ofthose vesicles in response to various stress conditions (19, 41).More recent electron microscopy reports describe membranevesicle chains and related membrane tubes (also referred to asperiplasmic tubules) that form cell–cell connections in thesocial soil bacterium Myxococcus xanthus (42) as well as the

Fig. 3. The genetic and molecular basis of bacterial nanowires. (A) Expression of pilin genes in S. oneidensis MR-1 measured by qPCR. Chemostat cultureswere 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 ingene expression relative to the reference sample was calculated by 2−ΔΔCT from at least four reactions of three independent chemostat cultures, using recAfor normalization. (B) Combined green (RedoxSensor Green) and red (FM 4-64FX) fluorescence images of the ΔpilA strain, lacking the type IV pilin majorsubunit PilA, before and after (95 min later) the production of a bacterial nanowire. ΔpilA is capable of producing bacterial nanowires with a similar re-spiratory 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.)

Fig. 4. Proposed structural model for Shewanella nanowires. S. oneidensisMR-1 nanowires are outer membrane (OM) and periplasmic (PP) extensionsincluding the multiheme cytochromes responsible for extracellular elec-tron transport.

12886 | www.pnas.org/cgi/doi/10.1073/pnas.1410551111 Pirbadian et al.

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 17

, 202

0

phototrophic consortium Chlorochromatium aggregatum (43).Consistent with these reports, we observed both a transition fromvesicle chains to smoother filaments (Movie S9), as well asnanowires connecting separate Shewanella cells (Movie S4).To gain a clearer picture of the role of membrane vesicles in

Shewanella nanowire formation, we performed atomic forcemicroscopy (AFM) on the same bacterial nanowires after ob-serving their growth with fluorescent imaging under perfusionflow conditions. Such observations were not possible in steady-state chemostat cultures where the nanowire growth is not con-fined to the surface–solution interface. Perfusion was stoppedand the samples were fixed quickly after observing the early signsof nanowire production, allowing us to examine the initial stagesof nanowire formation with nanoscale resolution using AFM(Fig. 5). We measured the nanowires to be 10 nm in diameterunder dry conditions, consistent with previous observations (10,23, 24) and roughly corresponding to two lipid bilayers. In ad-dition, we were able to resolve different morphologies corre-sponding to different stages of nanowire formation (Fig. 5),consistent with the chain-to-filament transition in Movie S9. Themorphologies observed ranged from vesicle chains (Fig. 5A) topartially smooth filaments incorporating vesicles (Fig. 5B, alsoconsistent with SEM imaging in SI Appendix, Fig. S7A), and fi-nally continuous filaments (Fig. 5C). In addition to possiblymediating EET up to micrometers away from the inner mem-brane, the vesiculation and extension of outer membranes intothe quasi one-dimensional morphologies observed here increasethe surface area-to-volume ratio of cells. This shape change canpresent a significant advantage, increasing the likelihood cellswill encounter the solid-phase minerals that serve as electronacceptors for respiration.Given the ubiquity of membrane vesicles and related exten-

sions in Gram-negative bacteria, the localization of electrontransport proteins along membrane extensions in a mannerconsistent with bacterial nanowires that could mediate extra-cellular electron transport, and the finding of nanowire-basedcell–cell connectivity, our results raise the intriguing possibility ofredox-functionalized membrane extensions as a general micro-bial strategy for EET and cell–cell signaling. Our study motivatesfurther experimental and theoretical work to build a detailedunderstanding of the full biomolecular makeup, electron trans-port physics, and physiological impact of bacterial nanowires.

Materials and MethodsCell Growth Conditions. All Shewanella strains were grown aerobically in LBbroth 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 for5 min, washed twice, and finally resuspended in a defined medium con-sisting of 30 mM Pipes, 60 mM sodium DL-lactate as an electron donor, 28mM NH4Cl, 1.34 mM KCl, 4.35 mM NaH2PO4, 7.5 mM NaOH, 30 mM NaCl, 1mM MgCl2, 1 mM CaCl2, and 0.05 mM ferric nitrilotriacetic acid. In addition,vitamins, amino acids, and trace mineral stock solutions were used to sup-plement the medium as described previously (7). The medium was adjustedto an initial pH of 7.2.

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

As previously detailed (7, 19), transition to electron–acceptor limitationwas required to promote the production of bacterial nanowires and mem-brane vesicles; this was accomplished using one of two methods. In the firstmethod, the experiment is started with fully aerobic medium flow beforeswitching to supply bottles containing medium made anaerobic by boilingand purging with 100% N2. This method has the advantage of providing aprecise time point for entering acceptor (O2) limitation (Fig. 1A and MoviesS1 and S4). In the second method, O2 becomes limited close to the coverslipsurface at high cell density, even with aerobic medium, as a result of thelaminar flow (no mixing between adjacent layers) and no-slip condition(zero velocity at the surface); this is confirmed with the simple calculationoutlined in SI Appendix. Using method 2, nanowires and membrane vesicleswere consistently observed after a lag period of 90–120 min from the start ofperfusion flow (Movies S2 and S3). We observed higher nanowire productionrates and better consistency using this method, possibly because the slowertransition exposed the surface-bound cells to a wide range of acceptor con-centrations, compared with the abrupt transition in the first method. Bothmethods resulted in identical membrane extensions as observed using mem-brane fluorescence (FM 4-64FX), protein fluorescence (NanoOrange), and theaccompanying increase in reductase activity (RedoxSensor Green).

Immunofluorescence with MtrC or OmcA Antibody. S. oneidensis MR-1 andΔomcA/mtrC (SI Appendix, Table S2) were used in the perfusion chamberexperiment as described above. As soon as the nanowires were producedand observed through staining by FM 4-64FX, the media flow was stoppedand the chamber was opened under sterile medium such that the coverslipremained hydrated. The sample (coverslip with attached cells) was fixed with4% (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 for5 min to quench free aldehyde groups and reduce background fluorescence.The sample was then transferred to a blocking solution of 1% BSA in PBS for5 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

Fig. 5. Correlated atomic force microscopy (AFM) and live-cell membrane fluorescence of bacterial nanowires. (A–C) Tapping AFM phase images ofS. 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 nanowiresare stained by the membrane stain FM 4-64FX. (Scale bars: 1 μm.) The morphologies observed range from vesicle chains (A) to partially smooth filamentsincorporating vesicles (B), which is consistent with SEM imaging of chemostat samples (SI Appendix, Fig. S7A), and, finally, continuous filaments (C). See alsoMovie S9, which captures the transition from a vesicle chain to a continuous bacterial nanowire.

Pirbadian et al. PNAS | September 2, 2014 | vol. 111 | no. 35 | 12887

MICRO

BIOLO

GY

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 17

, 202

0

Ab: 1.9 μg/mL, both in 1% BSA/PBS). After rinsing four times in PBS, thesample 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 beforeimmunofluorescence imaging in the green channel. To perform immuno-fluorescence on the same cells and nanowires observed during live imaging(Movies S7 and S8 and Fig. 3D), we modified the coverslips with surfacescratches that acted as fiducial markers.

Chemostat Growth and qPCR Analysis of the Transition from Electron-Donor toElectron-Acceptor Limitation. S. oneidensis MR-1 was grown in chemostatmedium (SI Appendix, Table S3) at 30 °C and 20% dissolved oxygen tensionusing 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, aspreviously described (7). After 48 h of this aerobic growth, a referencesample was taken. The dissolved oxygen tension was then manually droppedto 0% by adjusting the N2/air mixture entering the reactor, and the firstsample 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 timepoint, 10 mL of cells were harvested in ice-cold 5% citrate-saturated phenolin ethanol to prevent further transcription and protect the RNA. Sampleswere taken from three independent biological replicates (different chemo-stats). Total RNA was prepared using a hot phenol extraction, as previouslydescribed (36). Five micrograms of total RNA from each of the time points

described above was used as input for reverse-transcriptase reactions withthe SuperScript III First-Strand Synthesis System, as per the manufacturer’sprotocol (Life Technologies). Subsequent cDNA was then diluted and used astemplate in qPCR experiments with SYBR Select Master Mix (Life Technolo-gies). Fold change in gene expression relative to the reference sample wascalculated by 2−ΔΔCT from at least four reactions of three independentbiological replicate samples, using recA for normalization. Similar resultswere obtained using rpoB for normalization. The sequences of the primersused for each gene are shown in SI Appendix, Table S4.

ACKNOWLEDGMENTS. The pHGE-PtacTorAGFP plasmid was generously pro-vided by Prof. H. Gao (Zhejiang University), and pProbeNT was kindly providedby Dr. Steven Lindow (University of California, Berkeley). Atomic Force andElectron Microscopy were performed at the University of Southern CaliforniaCenters of Excellence in NanoBioPhysics and Electron Microscopy and Micro-analysis. The development of the in vivo imaging platform and chemostatcultivation was funded by Air Force Office of Scientific Research Young Inves-tigator Research Program Grant FA9550-10-1-0144 (to M.Y.E.-N.). Redox sensingmeasurements, compositional analysis, and the localization of multiheme cyto-chromes were funded by Division of Chemical Sciences, Geosciences, and Bio-sciences, Office of Basic Energy Sciences of the US Department of Energy GrantDE-FG02-13ER16415 (to M.Y.E.-N.). RT-PCR experiments and genetic analyseswere 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 Feder-ation consortium funded by the Genomics: Genomes to Life program of the USDepartment of Energy Office of Biological and Environmental Research.

1. Gray HB, Winkler JR (2003) Electron tunneling through proteins. Q Rev Biophys 36(3):341–372.

2. Myers CR, Nealson KH (1988) Bacterial manganese reduction and growth with man-ganese oxide as the sole electron acceptor. Science 240(4857):1319–1321.

3. Nealson KH, Belz A, McKee B (2002) Breathing metals as a way of life: Geobiology inaction. Antonie van Leeuwenhoek 81(1-4):215–222.

4. Kowalchuk GA, Jones SE, Blackall LL (2008) Microbes orchestrate life on Earth. ISME J2(8):795–796.

5. El-Naggar MY, Finkel SE (2013) Live wires. Scientist 27(5):38–43.6. Reguera G, et al. (2005) Extracellular electron transfer via microbial nanowires. Na-

ture 435(7045):1098–1101.7. Gorby YA, et al. (2006) Electrically conductive bacterial nanowires produced by She-

wanella oneidensis strain MR-1 and other microorganisms. Proc Natl Acad Sci USA103(30):11358–11363.

8. Marsili E, et al. (2008) Shewanella secretes flavins that mediate extracellular electrontransfer. Proc Natl Acad Sci USA 105(10):3968–3973.

9. von Canstein H, Ogawa J, Shimizu S, Lloyd JR (2008) Secretion of flavins by Shewanellaspecies and their role in extracellular electron transfer. Appl Environ Microbiol 74(3):615–623.

10. El-Naggar MY, et al. (2010) Electrical transport along bacterial nanowires from She-wanella oneidensis MR-1. Proc Natl Acad Sci USA 107(42):18127–18131.

11. Malvankar NS, et al. (2011) Tunable metallic-like conductivity in microbial nanowirenetworks. Nat Nanotechnol 6(9):573–579.

12. Pfeffer C, et al. (2012) Filamentous bacteria transport electrons over centimetre dis-tances. Nature 491(7423):218–221.

13. Snider RM, Strycharz-Glaven SM, Tsoi SD, Erickson JS, Tender LM (2012) Long-rangeelectron transport in Geobacter sulfurreducens biofilms is redox gradient-driven. ProcNatl Acad Sci USA 109(38):15467–15472.

14. Rabaey K, Rozendal RA (2010) Microbial electrosynthesis—revisiting the electricalroute for microbial production. Nat Rev Microbiol 8(10):706–716.

15. Vargas M, et al. (2013) Aromatic amino acids required for pili conductivity and long-range extracellular electron transport in Geobacter sulfurreducens. MBio 4(2):e00105–e00113.

16. Strycharz-Glaven SM, Tender LM (2012) Reply to the ’Comment on “On electricalconductivity of microbial nanowires and biofilms”’ by N. S. Malvankar, M. T. Tuominenand D. R. Lovley. Energy Environ Sci 5(3):6250–6255.

17. Pirbadian S, El-Naggar MY (2012) Multistep hopping and extracellular charge transferin microbial redox chains. Phys Chem Chem Phys 14(40):13802–13808.

18. Polizzi NF, Skourtis SS, Beratan DN (2012) Physical constraints on charge transportthrough bacterial nanowires. Faraday Discuss 155:43–62, discussion 103–114.

19. Gorby Y, et al. (2008) Redox-reactive membrane vesicles produced by Shewanella.Geobiology 6(3):232–241.

20. Grossart HP, Steward GF, Martinez J, Azam F (2000) A simple, rapid method fordemonstrating bacterial flagella. Appl Environ Microbiol 66(8):3632–3636.

21. Jones LJ, Haugland RP, Singer VL (2003) Development and characterization of theNanoOrange protein quantitation assay: A fluorescence-based assay of proteins insolution. Biotechniques 34(4):850–854, 856, 858 passim.

22. Leung KM, et al. (2011) Bacterial nanowires: Conductive as silicon, soft as polymer.Soft Matter 7(14):6617–6621.

23. El-Naggar MY, Gorby YA, Xia W, Nealson KH (2008) The molecular density of states inbacterial nanowires. Biophys J 95(1):L10–L12.

24. Leung KM, et al. (2013) Shewanella oneidensis MR-1 bacterial nanowires exhibitp-type, tunable electronic behavior. Nano Lett 13(6):2407–2411.

25. Kalyuzhnaya MG, Lidstrom ME, Chistoserdova L (2008) Real-time detection of activelymetabolizing microbes by redox sensing as applied to methylotroph populations inLake Washington. ISME J 2(7):696–706.

26. Cologgi DL, Lampa-Pastirk S, Speers AM, Kelly SD, Reguera G (2011) Extracellularreduction of uranium via Geobacter conductive pili as a protective cellular mecha-nism. Proc Natl Acad Sci USA 108(37):15248–15252.

27. Gray DR, Yue S, Chueng C-Y, Godfrey W (2005) Bacterial vitality detected by a novelfluorogenic redox dye using flow cytometry. Abstracts of the American Society ofMicrobiology Meeting (Am Soc Microbiol, Washington, DC).

28. Gralnick JA, Vali H, Lies DP, Newman DK (2006) Extracellular respiration of dimethylsulfoxide by Shewanella oneidensis strain MR-1. Proc Natl Acad Sci USA 103(12):4669–4674.

29. Betz WJ, Mao F, Smith CB (1996) Imaging exocytosis and endocytosis. Curr OpinNeurobiol 6(3):365–371.

30. Bolte S, et al. (2004) FM-dyes as experimental probes for dissecting vesicle traffickingin living plant cells. J Microsc 214(Pt 2):159–173.

31. Griffing LR (2008) FRET analysis of transmembrane flipping of FM4-64 in plant cells: IsFM4-64 a robust marker for endocytosis? J Microsc 231(2):291–298.

32. Rosu V, Hughes KT (2006) sigma28-dependent transcription in Salmonella enterica isindependent of flagellar shearing. J Bacteriol 188(14):5196–5203.

33. Blair KM, Turner L, Winkelman JT, Berg HC, Kearns DB (2008) A molecular clutchdisables flagella in the Bacillus subtilis biofilm. Science 320(5883):1636–1638.

34. Lowenthal AC, et al. (2009) Functional analysis of the Helicobacter pylori flagellarswitch proteins. J Bacteriol 191(23):7147–7156.

35. Luo Q, Dong Y, Chen H, Gao H (2013) Mislocalization of Rieske protein PetA pre-dominantly accounts for the aerobic growth defect of Tat mutants in Shewanellaoneidensis. PLoS ONE 8(4):e62064.

36. Rhodius VA, Gross CA (2011) Using DNA microarrays to assay part function. MethodsEnzymol 497:75–113.

37. Bouhenni RA, et al. (2010) The role of Shewanella oneidensis MR-1 outer surfacestructures in extracellular electron transfer. Electroanalysis 22(7-8):856–864.

38. Howard EC, Petersen ER, Fitzgerald LA, Sheehan PE, Ringeisen BR (2011) Optimalmethod for efficiently removing extracellular nanofilaments from Shewanella onei-densis MR-1. J Microbiol Methods 87(3):320–324.

39. Clarke TA, et al. (2011) Structure of a bacterial cell surface decaheme electron con-duit. Proc Natl Acad Sci USA 108(23):9384–9389.

40. Reardon CL, et al. (2010) Role of outer-membrane cytochromes MtrC and OmcA in thebiomineralization of ferrihydrite by Shewanella oneidensis MR-1. Geobiology 8(1):56–68.

41. Kulp A, Kuehn MJ (2010) Biological functions and biogenesis of secreted bacterialouter membrane vesicles. Annu Rev Microbiol 64:163–184.

42. Remis JP, et al. (2014) Bacterial social networks: Structure and composition of Myx-ococcus xanthus outer membrane vesicle chains. Environ Microbiol 16(2):598–610.

43. Wanner G, Vogl K, Overmann J (2008) Ultrastructural characterization of the pro-karyotic symbiosis in “Chlorochromatium aggregatum” J Bacteriol 190(10):3721–3730.

12888 | www.pnas.org/cgi/doi/10.1073/pnas.1410551111 Pirbadian et al.

Dow

nloa

ded

by g

uest

on

Feb

ruar

y 17

, 202

0