Diffusion-Based Recycling of Flavins Allows Shewanella oneidensis

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Environmental Processes

Diffusion-based recycling of flavins allows Shewanella oneidensis MR-1 to yield energy from metal reduction across physical separations Kyle Michelson, Reinaldo Enrique Alcalde, Robert A Sanford, Albert J Valocchi, and Charles J. Werth Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b04718 • Publication Date (Web): 06 Mar 2019 Downloaded from http://pubs.acs.org on March 6, 2019

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Environmental Science & Technology

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Category: Environmental Processes

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Title: Diffusion-based recycling of flavins allows Shewanella oneidensis MR-1

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to yield energy from metal reduction across physical

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separations

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Authors: Kyle Michelson†,‡, Reinaldo E. Alcalde‡, Robert A. Sanford§, Albert J. Valocchi†,

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Charles J. Werth‡,*

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†Department

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Champaign, 205 North Mathews Avenue, Urbana, IL 61801.

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‡Department

of Civil and Environmental Engineering, University of Illinois at Urbana-

of Civil, Architectural, and Environmental Engineering, University of Texas at

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Austin, 301 East Dean Keeton Street, Austin, TX 78712.

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§Department

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Avenue, Champaign, IL 61820.

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Keywords:

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of Geology, University of Illinois at Urbana-Champaign, 605 East Springfield

Extracellular electron transfer, metal reduction, electron shuttling, Shewanella

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ACS Paragon Plus Environment


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Abstract. We fabricated a microfluidic reactor with a nanoporous barrier to characterize electron

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transport between Shewanella oneidensis MR-1 and the metal oxide birnessite across a physical

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separation. Real-time quantification of electron flux across this barrier by strains with different

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electron transfer capabilities revealed that this bacterium exports flavins to its surroundings when

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faced with no direct physical access to an electron acceptor, allowing it to reduce metals at

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distances exceeding 60 microns. An energy balance indicates that flavins must be recycled for S.

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oneidensis MR-1 to yield energy from lactate oxidation coupled to flavin reduction. In our

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system, we find that flavins must be recycled between 24 and 60 times depending on flow

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conditions. This energy saving strategy, which until now had not been systematically tested or

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captured in environmentally relevant systems, suggests that electron shuttling microorganisms

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have the capacity to access and reduce metals in physically distant or potentially toxic

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microenvironments (i.e., pores with soluble and transiently-sorbed toxins) where direct contact is

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limited or unfavorable. Our results challenge the prediction that diffusion-based electron

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shuttling is only effective across short distances, and may lead to improved bioremediation

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strategies or advance biogeochemical models of electron transfer in anaerobic sediments.

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INTRODUCTION Bacteria store energy in the form of ATP by mediating intracellular redox reactions

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between electron donors and electron acceptors. In anaerobic sediments, however, soluble

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electron acceptors are often unavailable for intracellular respiration1. While some species of

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bacteria respond to electron acceptor limitations by entering a dormant state of decreased

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metabolic activity, others have evolved mechanisms for the transport of respiratory electrons to

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the outer membrane in a process defined as extracellular electron transport (EET)2. Dissimilatory

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metal-reducing bacteria (DMRB), most notably of the Shewanella and Geobacter genera, have

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evolved several strategies to generate ATP from the reduction of insoluble electron acceptors

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such as Fe(III) and Mn(IV) that are abundant in anaerobic sediments1. These include the

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localization of c-type cytochromes (c-Cyts) to the outer membrane3–6, secretion of extracellular

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polymeric substances (EPS) with embedded c-Cyts7,8, production of electrically conductive

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appendages (i.e. nanowires)9,10, and electron shuttling via redox mediators (e.g. flavins)11–13.

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Nanowires and electron shuttles have the additional advantage of being able to reach

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spatially distant electron acceptors in narrow pore spaces and toxic microenvironments. For

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example, Geobacter sulfurreducens reduces U(VI) outside its cell wall via pilus-based nanowires

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to avoid periplasmic accumulation of toxic U(IV)9,14. G. sulfurreducens KN400 can use

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nanowires in association with flavin cofactors to reduce metal oxides in nanopores at distances of

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15 μm15. While S. oneidensis MR-1 also produces conductive nanowires, electron shuttling of

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flavins appears to be the dominant pathway for metal oxide reduction used by this species11,12.

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Flavin electron shuttling involves the reduction of flavins by outer membrane c-Cyts OmcA and

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MtrC that serve as the terminal reductases of the metal-reducing (Mtr) pathway16,17. Once

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reduced, flavins may transfer electrons to a metal oxide as diffusion-based shuttles or cofactors

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bound to OmcA and MtrC. The latter mechanism has been validated by electrochemical

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measurements of current generation in Shewanella biofilms18, but can only be used when cells

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are in direct contact with an electron acceptor. Diffusion-based shuttling could potentially reduce

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metals over much greater distances than nanowires. However, shuttling of endogenously

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produced flavins requires extensive recycling to offset the metabolic cost of biosynthesis11. This

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can be difficult in open environments where shuttles may diffuse beyond a radius of recapture.

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Despite these limitations, there is some evidence for diffusion-based shuttling. For

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example, Fe(III) oxide in nanoporous glass and alginate beads was reduced in batch culture by

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Shewanella spp., presumably by flavins19,20. Current production by S. oneidensis MR-1 in

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membrane bioreactors sharply declined when supernatant containing flavins was replaced with

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media without flavins, which would affect diffusion-based electron shuttling11. However, little is

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known about the recycling efficiency of flavins, the length scale of diffusion-based shuttling, or

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the relative contribution of competing pathways that may also be capable of metal reduction

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across a physical separation. Regarding the latter, metal reduction by endogenously produced

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thiols (e.g. cysteine) through sulfur metabolism is an alternate pathway that has yet to be

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characterized in DMRB. This mechanism is potentially significant because S. oneidensis MR-1 is

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capable of disulfide bond reduction and electron transfer to reduced and oxidized thiols21, but the

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aforementioned studies with flavins in electrochemical reactors suggest it is not dominant13.

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Characterizing diffusion-based EET has the potential to transform how we conceptualize and

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model biogeochemical reactions, and may lead to new bioremediation strategies. Bioremediation

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could be improved by amending groundwater with flavins or thiol precursors, for example, or

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stimulating the growth of electron shuttling DMRB. Biogeochemical reactions in nanopores,

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which account for > 90% of the surface area in porous media22, could be modeled more

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accurately if the rate and extent of metal reduction across a physical separation is known.

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Our objective is to measure the rate and extent of metal reduction by S. oneidensis MR-1

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across a physical separation and characterize the energetics of long-range EET. Based on the

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aforementioned association of flavins with metal reduction, and the effect of soluble flavins on

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current generation, we hypothesize that flavins will serve as the dominant electron shuttles and

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reduce spatially distant metal oxides by diffusion. Given the high ATP cost of flavin

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biosynthesis, and evidence supporting flavin reduction at outer membrane protein sites, we

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hypothesize that flavins will be reused many times during metal reduction. Our approach to test

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these hypotheses involves the use of a silicon-etched microfluidic reactor with a nanoporous

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barrier that is capable of separating S. oneidensis MR-1 from the Mn(IV) mineral birnessite.

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Microfluidic reactors can be operated on a microscope, enabling real-time, in-situ visualization

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in space and time of biofilm growth and birnessite reduction by brightfield and fluorescent

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microscopy, and mineral analysis by Raman spectroscopy. The role of flavins in metal reduction

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is tested using wild-type and mutant strains, where the latter are deficient in flavin export or

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reduction. Electron flux is quantified by measuring flavin and thiol concentrations in reactor

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effluent and monitoring birnessite reduction by optical microscopy and Raman spectroscopy.

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MATERIALS AND METHODS

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Cell Growth Conditions. Wild-type S. oneidensis MR-1 (referred to as MR-1), a knock-out

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mutant lacking the FAD transporter (referred to as Δbfe)13, and a double-deletion mutant lacking

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two outer membrane electron transport proteins (referred to as ΔomcA/ΔmtrC)23 were used in

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this work. They were each cultured at pH 7.0 and 30 °C in anoxic, bicarbonate-buffered

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freshwater medium containing the following per liter of distilled water: 0.5 g NH4Cl, 0.14 g

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KH2PO4, 0.2 g MgCl2·6H2O, 0.15 g CaCl2·2H2O, and 2.5 g NaHCO3. This was supplemented

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with a 2% w/v solution of cysteine-free casamino acid digest, a non-chelated SL-10 trace

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elements solution, and a selenite plus tungstate solution24. Vitamins and chelating agents were

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omitted to minimize the risk of abiotic reduction and dissolution. Sulfur was provided by

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methionine at a concentration of 12.5 μM. Lactate was supplied as the electron donor at a

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concentration of 20 mM. Batch culture experiments were performed with the three MR-1

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species, in some cases with flavins added, to support interpretation of microfluidic reactor

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experiments. All batch culture experiments are listed in Table S1. Experimental details are in the

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Supporting Information, and each batch experiment was performed in triplicate.

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Birnessite Synthesis. Birnessite [(Na, K)0.6(Mn4+, Mn3+)2O4 · 1.5H2O] was synthesized by

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oxidizing manganese chloride in a basic solution of potassium permanganate25, and as described

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in our recent work15. Fresh batches of birnessite were prepared every 3 months and analyzed by

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Raman spectroscopy before each experiment to verify the purity of the birnessite. TEM images

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from our previous work showed that birnessite particles were < 100 nm in length15. However,

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particles aggregate and do not pass through the nanoporous barrier.

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Microfluidic Reactor Setup and Operation. Microfluidic reactors were etched in silicon using

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photolithography. A schematic illustrating the major steps of fabrication is shown in Figure S1,

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and detailed fabrication steps are provided in the Supporting Information. The main feature of

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our reactor is a thin, nanoporous barrier measuring 2.5 μm in width and 2 cm in length. The

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barrier bisects two parallel flow channels that are 250 μm wide and 10 μm deep. An array of slits

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(i.e. pores) measuring 3 μm (L) x 2.5 μm (W) x 0.18 μm (D) are etched into the top of the barrier

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at 3 μm intervals to allow the diffusion of solutes between each channel but not the passage of

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cells. Flow was delivered to the channels at a rate of 2.2 μL/h, corresponding to a linear velocity

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of 1.2 cm/min and a residence time of 90 seconds. Reactors were stored in the dark to prevent

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photodegradation of flavins and maintained at a temperature of 25 °C ± 1 °C. Reactors were

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disinfected for 12 hours with 1 M acetic acid26 and flushed with 3,000 pore volumes of sterile DI

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water before starting an experiment. Reactor effluent was collected every 24 hours for analysis

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of flavins, thiols, and aqueous manganese.

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Experiments consisted of four stages as summarized in Table S2. In Stage 1, birnessite

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was infused through one flow channel and immobilized at the nanoporous barrier by inducing

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cross flow. This was achieved by closing the effluent port on the birnessite side and opening the

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effluent port on the bacteria side. After fixing the birnessite, the reactor was inoculated in Stage 2

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by infusing fumarate grown cells through the adjacent channel. In Stage 3, flow was closed off to

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the bacteria side while the birnessite channel was infused with 20 mM lactate as the electron

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donor and a combination of 0.5 mM nitrate and 5 mM fumarate as soluble electron acceptors.

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Chemotaxis toward the barrier was observed within hours, and flow through the birnessite side

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was maintained for 12-24 hours until a biofilm was established at the barrier. In Stage 4, both

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channels were infused with media containing only lactate and media so that birnessite remained

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as the sole electron acceptor. Experimental conditions and results for microfluidic experiments

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are summarized in Table S3, and all experiments were performed in triplicate unless noted

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otherwise.

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Analytical Methods. Aqueous Mn(II) captured in effluent samples was analyzed on a Varian

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710-ES inductively coupled plasma optical emission spectrometer (ICP-OES) after acidification

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of samples in 10 mM oxalic acid and 2% HCl. Flavins and thiols were collected from both

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outlets and analyzed by reverse-phase high performance liquid chromatography (HPLC).

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Detailed analytical methods for the determination of flavins and thiols are in the Supporting

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Information.

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Black and white images were taken with an Andor Zyla 5.5 camera on a Nikon Eclipse

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TI-E inverted microscope as previously described15. The resolution of our objectives was 0.56

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μm for 550 nm light, which is sufficient to measure the extent of metal reduction over time.

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Autofluorescence of c-Cyts produced by MR-1 was imaged through a 400 nm dichroic mirror at

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excitation and emission wavelengths of 340-380 nm and 435-485 nm, respectively. Color images

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were taken with a Lumenera Infinity 3-1UR camera.

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Manganese reduction in the microfluidic reactor was estimated by measuring the total

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area of birnessite reduced in the reactor and using birnessite density and reactor depth to convert

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area to concentration. Birnessite and rhodochrosite (MnCO3) area was determined by optical

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thresholding using Nikon NIS software. This technique involves looking at contrast in grayscale

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and selecting pixels above a threshold value. Reactor depth was measured along the entire

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reactor by optical profilometry during the fabrication process and averaged. Density of birnessite

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at the nanoporous barrier was calculated from the average oxidation state (AOS) of Mn, the Mn

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content of birnessite from a stock solution analyzed by ICP, and the mass of Mn per unit area of

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birnessite inside triplicate reactors that were pulverized and also analyzed by ICP (SI Materials

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and Methods).

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Manganese solids inside the reactor were identified with the Horiba LabRAM HR

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Evolution confocal Raman system. Raman spectra were taken between 0 and 1200 cm-1 using a

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532 nm diode-pumped solid-state (DPSS) laser. The laser was calibrated using silicon, and

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reference spectra from the RUFF mineral database was used to identify minerals inside the

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reactor. Birnessite spectra were obtained at 10 mW and averaged over 5 scans with an

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acquisition time of 60 seconds. Rhodochrosite spectra were obtained at 2.5 mW and averaged

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over 20 scans with an acquisition time of 30 seconds. Rhodochrosite deposited as a lower density

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and less cohesive layer, and required a lower laser power to prevent movement of particles

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during analysis. A greater scan number was used to compensate for the decrease in laser power.

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RESULTS

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The nanoporous barrier physically separates S. oneidensis MR-1 from birnessite. A photo

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of the microfluidic reactor and a close-up of the nanoporous barrier with 180 nm deep slits is

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shown in Figures 1a,b. For reference, the diameter of MR-1 across its smallest axis at 22 °C is

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610 ± 110 nm27. An illustration of the experimental set-up, with partially reduced birnessite

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separated from a biofilm of MR-1, is shown in Figure 1c. Autofluorescence of c-Cyts28 in MR-1

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was used to monitor the location of cells within the reactor and check for passage across the

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barrier (Fig. S2). In earlier (faulty) reactor designs with deeper slits, cell passage was visible

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within the low-density layer of rhodochrosite, and cell motility within rhodochrosite could be

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observed using brightfield microscopy. With our present design, we did not observe any cells

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across the barrier or within the slits. In several experiments, MR-1 was grown in the absence of

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birnessite to provide a clear view of cells by brightfield and fluorescence microscopy. As shown

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in Figure S3, cells were retained in one channel of the reactor. Media containing lactate and

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fumarate was also inoculated with effluent from the abiotic channel to check for contamination,

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but no growth was observed. Together, these results demonstrate the necessary performance of

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the barrier in retaining cells.

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Birnessite reduction across a physical separation is biologically driven. Birnessite reduction

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by MR-1 in a representative experiment is shown in Figure 2, where reduction of the dark brown

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birnessite to the tan colored rhodochrosite is indicated in the bright field images. While the

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reduction front in Figure 2 appears diffuse, this focal plane was only chosen to highlight the

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location of the nanoporous barrier. At the silicon/barrier interface where rhodochrosite

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precipitates, the reduction front is sharp despite the barrier being slightly out of focus (Fig. S4).

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Images taken from this focal plane were used to track the extent of birnessite reduction over

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time. In all experiments, birnessite reduction began after a lag period of 1-3 days after the start of

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Stage 4, which we attribute to the acclimation of MR-1 to the absence of aqueous phase electron

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acceptor (i.e., fumarate and nitrate). In no cases was birnessite reduction observed in Stage 3,

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when fumarate and nitrate were amended to the reactor. Within 10 days from the start of

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reduction, the 40-60 μm layer of birnessite was completely reduced to soluble Mn(II) and a solid

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product that was identified in-situ as rhodochrosite (MnCO3) by Raman spectroscopy (Fig. S5).

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The cumulative production of Mn(II) was determined based on the average oxidation

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state (AOS), manganese content, and density of birnessite, as well as the area of birnessite that

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was reduced in the microfluidic reactor over time (Materials and Methods). The AOS,

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determined in triplicate by permanganate back titration, is 3.96 ± 0.03, similar to reported

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values25. The manganese content (i.e., Mn/birnessite), determined in triplicate by ICP-OES on a

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mass fraction basis is 0.44 ± 0.02, also similar to reported values29. The density of birnessite in

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the reactor, determined in triplicate by pulverizing reactors with known depth and birnessite area

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(SI Materials and Methods) then analyzing by ICP-OES, is 0.298 ± 0.017 g/cm3. Channel depth

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of unbonded reactors was measured by optical profilometry during the fabrication process, and

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SEM cross-sections of dried reactors showed that deposited birnessite spanned the entire depth

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of the channels (Fig. S6). Birnessite density and reactor depth was used to convert area to

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concentration and calculate the Mn(II) production rate. Based on these values, we determined

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that the Mn(II) production rate during birnessite reduction is 437 ± 43 pmol/day at a flow rate of

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2.2 μL/h, with 4009 ± 405 pmol Mn(II) produced after 10 days. Cumulative birnessite reduced

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(i.e., Mn(II) produced) is shown in Figure 3.

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Mn(II) production was also quantified in two experiments by analyzing effluent by ICP-

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OES. The Mn(II) production rate according to ICP-OES was 9-14% lower than that calculated

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by area during birnessite reduction (Materials and Methods), and we attribute the slightly lower

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production rate to the precipitation of rhodochrosite. When rhodochrosite was dissolved in oxalic

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acid at the end of these experiments, the cumulative production of Mn(II) was 104-110% of that

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calculated by area. We attribute this discrepancy to small particles of birnessite that deposit

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upstream and are not counted as part of the birnessite area. We decided to use area rather than

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effluent concentration to quantify Mn(II) production because there is no lag between in-situ and

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measured reduction, as there is with effluent measurements.

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Abiotic control experiments (i.e., same conditions except no cells) were conducted in the

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microfluidic reactor, and showed no reduction or dissolution of birnessite after one month of

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continuous flow; this demonstrates that birnessite reduction over time is not due to abiotic

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reduction or dissolution with media. Confocal images of fluorescent beads embedded in

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birnessite in a replicate reactor indicated that birnessite particles were immobilized throughout

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depth and thickness once deposited on the nanoporous wall (Fig. S7). This discounts the

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possibility that the migration of particles accounts for the disappearance of birnessite. Solid-state

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electron transport through birnessite particles was ruled out due to the low conductivity of

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manganese oxide (10-5-10-6 S/m)30.

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Birnessite reduction is consistent with flavin electron shuttling. Effluent collected from the

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original experiment containing MR-1 opposite birnessite in the microfluidic reactor was

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analyzed for electron shuttles. Riboflavin (RF) and flavin mononucleotide (FMN) were

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consistently detected by HPLC at masses of 5.12 ± 0.28 and 12.79 ± 0.63 pmol, respectively, as

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the combined mass from both channels measured over a 24-hour collection period. This is shown

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in Figure 3. Recovery of total flavins in control experiments with birnessite but without cells was

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96% in the microfluidic reactor under similar conditions during infusion of 60 nM RF and 40 nM

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FMN (Figure S8). Additionally, we did not detect any of the known degradation byproducts of

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photolysis or hydrolysis (e.g. lumichrome, lumiflavin)31.

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To test the role of flavins more thoroughly, we repeated our experiments with the mutant

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lacking bfe, the bacterial FAD transporter13. This mutant is capable of flavin biosynthesis but not

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flavin export across its outer membrane. In batch culture, we observed a 12- and 37-fold

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decrease in RF and FMN, respectively, despite wild-type rates of birnessite reduction. In the

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microfluidic reactor, however, we observed a dramatic decrease in the reduction rate by

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approximately 5 times, corresponding to a decrease in RF and FMN concentrations by 6 and 8

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times, respectively (Fig. 4). Mn(II) production in the microfluidic reactor was restored to wild-

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type rates by providing bfe cells with an exogenous supply of RF and FMN at concentrations

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similar to those produced by wild-type cells (Fig. 4). This result discounts the possibility that

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pleiotropic effects resulting from the mutation may have influenced the rate of birnessite

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reduction and reinforces the role of flavins in birnessite reduction.

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As a control, we repeated our experiments with the ΔomcA/ΔmtrC mutant, which lacks

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the two outer membrane c-Cyts of the Mtr pathway. In batch culture, only a small fraction of

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birnessite was reduced, even when supplemented with flavin. In the microfluidic reactor, no

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reduction was observed (Fig. 4). These results were expected, and are consistent with prior work

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showing that these outer membrane proteins are necessary for electron transfer to flavins32–34.

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We tested whether thiols were serving as electron shuttles, since MR-1 is capable of

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disulfide bond reduction21. Hence, a portion of the effluent was reduced using tris(2-

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carboxyethyl)phosphine (TCEP) and derivatized with 4-Chloro-3,5-dinitrobenzotrifluoride

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(CNBF) to quantify thiols (SI Materials and Methods)35. We detected three thiols including

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cysteine, homocysteine, and glutathione, but only in the channel containing MR-1. Cysteine

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production began several days after the start of Stage 4, while homocysteine and glutathione

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were detected in trace amounts at the limit of detection and could not be accurately quantified.

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Cysteine was present at concentrations between 200-800 nM (Fig. S9), but its concentration in

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the effluent was not correlated to the Mn(II) production rate (i.e. the concentration of cysteine

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was variable, while the reduction rate of birnessite was constant). Therefore, our results do not

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support thiols serving as electron shuttles for birnessite reduction across a physical separation.

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Finally, we explored the possibility of conductive EPS being secreted across the barrier.

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To date, 42 types of c-Cyts have been detected on the outer membrane of MR-136, and 20 redox

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proteins have been extracted from EPS37. Outer membrane c-Cyts OmcA and MtrC are

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associated with the EPS of MR-16,38, and may be responsible for its reported conductivity8. We

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ran two additional experiments in sacrificial reactors to investigate the potential of long-range

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EET via the secretion of EPS. The reactors were dried, broken apart, and analyzed by SEM.

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While EPS was clearly present in the biofilm, it was not observed across the barrier in either

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reactor (Fig. 5 and Fig. S10), conclusively showing that EPS was not involved in birnessite

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reduction. We also note the absence of nanowires, which should be visible by SEM given their

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diameter of 50-150 nm23. While imaging nanowires can be difficult, they are also under 10 μm in

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length10, which rules out their involvement in EET beyond this distance.

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Diffusion-based shuttling and recycling of flavins accounts for the electron transfer

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imbalance. We ran an additional pair of experiments with variations in flow rate to confirm the

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role of diffusion-based electron shuttling and recycling. After biofilm was established opposite

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birnessite, the influent and effluent ports for the birnessite-containing channel were initially

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closed to limit the loss of soluble species, then opened once half of the birnessite layer was

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reduced. After several days, the birnessite ports were closed again. If flavins were used more

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than once (i.e. recycled) for birnessite reduction, we would expect the opening of these ports to

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coincide with a decrease in Mn(II) production due to the removal of flavins from the purged flow

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channel (as a result of the steeper concentration gradient of flavins across the birnessite). In line

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with this expectation, we observed a clear decrease in Mn(II) production (based on birnessite

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reduced) immediately following the onset of flow in the birnessite channel (Fig. 6). During

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stopped flow, average Mn(II) production was 1002 ± 152 pmol/day. During continuous flow, this

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decreased to 396 ± 14 pmol/day, very close to the value of 437 ± 43 pmol/day obtained in the 10-

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day continuous flow experiment. During the final stopped flow phase, the average reduction rate

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increased to 1271 ± 67 pmol/day.

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The number of electrons available from flavins can be compared to the number of

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electrons transferred to birnessite to indicate if the former are being recycled to reduce the latter.

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As illustrated in Figure 3, 17.9 ± 0.8 pmol/day of flavins were measured in the effluent of the

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microfluidic reactor under continuous flow conditions, while 437 ± 43 pmol/d of birnessite were

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reduced in the same experiment. We note that flavin concentration in the effluent reached steady-

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state before the start of reduction, indicating that soluble flavin was in equilibrium with any

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flavin that may have been adsorbed to the birnessite. Reduction of birnessite by flavins via

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diffusion-based shuttling is a two-electron transfer process18. Therefore, for flavins to be

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responsible for birnessite reduction, they must be reused ~24 times under continuous flow

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conditions. Under stopped flow conditions, 21.6 ± 1.2 pmol/day of flavins were produced and

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1271 ± 67 pmol/day of birnessite were reduced, indicating that flavins were reused ~60 times.

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We observed Brownian motion of particles in the reduced region between birnessite and the

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nanoporous barrier, which is consistent with our assumption that diffusion is responsible for

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electron transport between the biofilm and deposited birnessite. We also determined from a

308

column breakthrough experiment (Fig. S11, SI Materials and Methods) that flavins reversibly

309

sorb to birnessite, with a calculated retardation factor of 44; this would enhance flavin recycling

310

because flavin diffusion between birnessite and the nanoporous barrier would be fast relative to

311

flavin diffusion through the birnessite and subsequent loss to the flow channel.

312

Implications. Diffusion-based recycling of endogenously produced flavins may allow bacteria to

313

access metal oxides in difficult to access microenvironments, such as within intra-aggregate soil

314

and clay pore spaces too small for cell passage, and in contaminated groundwater plumes with

315

extreme pH or elevated antibiotic concentrations. We did not expect reduction to occur across

316

the entire layer of immobilized birnessite in our microfluidic reactors since diffusion-based 15



Page 16 of 31

317

shuttling of flavins at physiological concentrations is predicted to be effective only at short

318

distances39. This assumption was based on comparisons between measured electron flux in

319

anode-grown biofilms and theoretical electron flux by diffusion-based shuttling. Our results also

320

suggest that flavins play a less important role in mediating electron transfer when the cell is in

321

direct contact with birnessite (i.e., batch experiments), possibly due to more efficient flavin

322

recycling under well-mixed conditions.

323

The recycling mechanism our results support is important because it allows MR-1 to

324

recoup energetic losses from flavin biosynthesis. To put this into perspective, we can compare

325

the energy requirements for flavin synthesis to the energy generated through the oxidation of

326

lactate and acetate (E°′ = −0.42 V) coupled to the reduction of RF (E°′ = −0.21 V) and FMN (E°′

327

= −0.24 V) reported under conditions similar to our own (20 mM lactate, 250 nM flavin, pH 7)11.

328

With RF and FMN accounting for 29% and 71% of the flavins measured in our effluent,

329

respectively, the total energy available from this redox couple is calculated at -72.8 kJ/mol-

330

lactate using the Nernst equation, where negative values correspond to a net gain in free energy.

331

The energetic cost to produce ATP at pH 7 assuming typical intracellular concentrations of ADP

332

and ATP for anaerobic bacteria (2 mM ATP, 1 mM ADP) is 43.9 kJ/mol-flavins40.

333

Approximately 25 mol-ATP/mol-flavin are required for synthesis41, resulting in a free energy

334

demand of 1098 kJ/mol-flavin. If flavins are not recycled, this would correspond to a minimum

335

daily energy requirement of 4.8 × 10-4 J/d to reduce the 437 pmol/day of birnessite measured in

336

the continuous flow microfluidic experiment, while the energy gained from coupling lactate

337

oxidation to flavin reduction (based on MnO2 reduced) is only -1.6 × 10-5 J/d. Therefore, the

338

energy gain from flavin reduction is insufficient to offset the energy requirement for flavin

339

biosynthesis. Recycling flavins 24 times, for example (as we determined experimentally),

16


Page 17 of 31


340

reduces the energetic cost of flavin synthesis to 2.0 × 10-5 J/d, which is very close to the

341

breakeven point despite representing an overall cost of 4.1 × 10-6 J/d. While the actual energy

342

gain depends on specific biochemical pathways and solute concentrations, the calculations reveal

343

that flavin recycling may allow MR-1 to yield energy from birnessite reduction at a distance.

344

These calculations suggest that biofilm growth should be slow or stagnant during diffusion-

345

based, flavin-mediated reduction of birnessite. Indeed, biofilm area in the microfluidic reactor

346

remained constant after switching from fumarate to birnessite as a terminal electron acceptor

347

(Fig. 2). However, the interpretation of these qualitative observations is limited since we did not

348

investigate the metabolic status of cells within the biofilm.

349

The results presented here, summarized in Table S3, suggest that conventional models of

350

electron transport in anaerobic sediments should be revised to account for diffusion-based

351

electron shuttling across physical separations. We predict that electron acceptors sequestered

352

deep within narrow pores and in toxic microenvironments, including iron and manganese oxides,

353

metalloids (e.g. selenium), and radionuclides (e.g. uranium), are accessible to Shewanella spp.

354

and other microorganisms capable of flavin synthesis and diffusion-based electron shuttling.

355

Injecting electron shuttles at contaminated sites or stimulating the growth of shuttle-producing

356

DMRB may improve the efficiency of bioremediation and should be explored in future work.

357

ASSOCIATED CONTENT

358

Supporting Information

359

Supplementary materials and methods, process diagram for reactor fabrication (Figure S1),

360

autofluorescence of cytochromes (Figure S2), retention of cells by the nanoporous barrier

361

(Figure S3), interface between birnessite and rhodochrosite (Figure S4), Raman spectra of

17



Page 18 of 31

362

birnessite and rhodochrosite (Figure S5), SEM cross-section of reactor with deposited birnessite

363

(Figure S6), fluorescent beads embedded in deposited birnessite (Figure S7), recovery of flavins

364

in abiotic control experiments (Figure S8), cysteine production during birnessite reduction

365

(Figure S9), close-up of debonded reactor (Figure S10), breakthrough curves to determine flavin

366

retardation in birnessite (Figure S11), summary of conditions and results for batch culture

367

experiments (Table S1), description of stages in microfluidic experiments (Table S2), and

368

summary of conditions and results for microfluidic experiments (Table S3). This information is

369

available free of charge via the Internet at http://pubs.acs.org/.

370

AUTHOR INFORMATION

371

Corresponding Author

372

*Phone: +01 (512) 232-1626; e-mail: [email protected].

373

Notes

374

The authors declare no competing financial interest.

375

ACKNOWLEDGEMENTS

376

This work was supported by the U.S. Department of Energy under Award DE-SC0006771,

377

Office of Biological and Environmental Research (BER), Office of Science, US Department of

378

Energy (DOE), and partially supported by the National Aeronautics and Space Administration

379

(NASA) through the NASA Astrobiology Institute under Cooperative Agreement No.

380

NNA13AA91A issued through the Science Mission Directorate, and the National Science

381

Foundation Graduate Research Fellowship Program (NSF GRFP) under Grant No. DGE-

382

1610403 to Reinaldo E Alcalde. We thank J. Gralnick (UMN) for providing MR-1 and

383

ΔOmcA/ΔMtrC mutants, and B. Keitz (UT) for providing the bfe mutant.

18


Page 19 of 31


384

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TOC Art

25



A Birnessite

C

Oxidized Flavin Reduced Flavin Bacteria

Page 26 of 31

Birnessite, Mn(IV) Rhodochrosite, Mn(II)

Lactate + Cells

B

1 μm

514

Figure 1. (A) Photograph of the microfluidic reactor. (B) SEM image of the silicon-etched

515

nanoporous barrier at a 45-degree incline. (C) Illustration of birnessite reduction to rhodochrosite

516

by flavin electron shuttling and a recycling mechanism.

26


Page 27 of 31


Day 1

Day 6 Birnessite

Rhodochrosite

Biofilm

Day 3

Day 10

517

Figure 2. Time progression of birnessite reduction across the nanoporous barrier over 10 days by

518

S. oneidensis MR-1.

27


5000

Page 28 of 31

200

Cumulative Mn

4500

180

Cumulative flavin

4000

160

3500

140

3000

120

2500

100

2000

80

1500

60

1000

40

500

20

0

Cumulative flavins produced (pmol)

Cumulative birnessite reduced as Mn(II) (pmol)


0 1

2

3

4

5

6

7

8

9

10

Day 519

Figure 3. Cumulative moles of flavin (as the sum of RF and FMN) produced by S. oneidensis

520

MR-1 and moles of birnessite reduced during continuous flow. Error bars indicate data from

521

triplicate experiments.

28




Page 29 of 31

4000 WT

3500

bfe bfe + flavin

3000

OmcA/MtrC

2500 2000 1500 1000 500 0 0

2

4

6

8

Day

522

Figure 4. Cumulative birnessite reduced in the microfluidic reactor by wild-type (WT), Mtr

523

mutant (ΔomcA/ΔmtrC), bfe export mutant (bfe), and bfe cells of S. oneidensis MR-1

524

supplemented with FMN and RF (bfe + flavin). Flavin supplementation consisted of 5 pmol RF

525

and 15 pmol FMN per day.

29



Page 30 of 31

10 μm 526

Figure 5. SEM image of the birnessite side of the nanoporous barrier after debonding a

527

microfluidic reactor.

30




Page 31 of 31

10000

Stopped flow

Continuous flow

slope = 1002 ± 152

slope = 396 ± 14

Stopped flow

9000 8000 7000 6000 5000 4000

slope = 1271 ± 67

3000 2000 1000 0 0

2

4

Day

6

8

10

528

Figure 6. Cumulative birnessite reduced under different flow conditions. Flow through the

529

bacteria channel was continuous. Flow through the birnessite channel was either stopped (white)

530

or continuous (blue). The slope is equal to pmol reduced per day for each stage.

31


Diffusion-Based Recycling of Flavins Allows Shewanella oneidensis

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