M.D.Y, S.M.G-S., and L.M.T. acknowledge the Office of Naval Research (Award #N0001415WX01038 and N0001415WX00195), the Naval Research Laboratory, and the Naval Research Laboratory Nanosciences Institute; M.Y.E.-N. is supported by the U.S. Department of Energy Grant DE-FG02-13ER16415.

1. Interdigitated microelectrode array (IDA) preparation
<ol>
	<li>Obtain commercially available IDA electrodes patterned on a nonconductive substrate or synthesize them using standard lithographic methods.28 
	NOTE: IDA dimensions and/or materials can be varied based on desired conditions for different experiments. IDAs used here were obtained commercially and consisted of two interdigitated gold microelectrodes patterned on a glass substrate connected to large electrode pads at opposite ends of the array. The electrodes are exposed while the bus lines connecting the electrodes to the large contact pads are coated with thin insulating material. IDAs used here are comprised of two sets of 10 &#181;m-wide and 2 mm long gold microelectrode bands (source and drain) spaced 5 &#181;m apart patterned on a flat glass surface. IDAs used here had 65 electrode pairs (130 total interdigitated bands). Alternating bands are connected to electrode pads at opposite ends of the array.</li>
	<li>Wire and insulate IDA
	<ol>
		<li>Attach a wire to each large electrode pad using conductive silver epoxy.
		<ol>
			<li>Prepare conductive epoxy according to manufacturer&#39;s instructions for the specific epoxy in use with a mixing rod or pipette tip (instructions may vary by manufacturer).</li>
			<li>Place a wire on each gold pad, secure in place using lab tape. Cover the wire and pad with silver epoxy using a mixing rod or pipette tip.</li>
			<li>Carefully move to an 80 &#176;C oven for 1 h to cure or cure based on manufacturer&#39;s recommendations for the specific conductive silver epoxy used.</li>
			<li>After the epoxy cures, use a multimeter to ensure electrical connectivity between the end of the wire and the pads. The resistance between the wire and the pad should be &#60; 5 &#937;. Also, use the multimeter to verify that no conductive epoxy is connecting multiple electrode pads, as this can short the array. If conductive epoxy is connecting multiple leads, use a scribe to isolate the leads.</li>
		</ol>
		</li>
		<li>Insulate the epoxied connection by coating the connection with a thermal, electrical, and waterproof insulating material. Flame retardant polyurethane resins are often suitable.
		<ol>
			<li>Remove the tip of a 15-mL conical centrifuge tube (at approximately the 2.5 mL mark) as a mold for the insulating material. Make two small holes in the bottom for the wires to protrude through using a 21g needle.</li>
			<li>Insert the IDA into the mold and wires through the holes in the bottom of the mold.</li>
			<li>Prepare the specific insulating material. Follow instructions provided with the specific resin obtained.</li>
			<li>Pipette the insulator into the mold so that the silver epoxy is completely covered (~2.5 mL) and allow to dry according to manufacturer&#39;s specifications.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Electrochemical reactor setup, testing, and inoculation
<ol>
	<li>Set up the electrochemical cell.
	<ol>
		<li>Insert IDA, counter electrode, and reference electrode into the electrochemical cell. 
		NOTE: The electrochemical reactor used can vary widely, as long as all of the electrodes fit inside. One consideration is that the reference electrode should be as close as possible to the working electrode given the practical limits of the reactor. Here, we use a single chamber reactor with the reference electrode ~2-3 cm from the working electrodes. Also, the counter electrode should be larger than the working electrode to ensure that it is not limiting in the system.</li>
		<li>If working with a pure culture, sterilize the reactor with the counter electrode inside. Sterilize the reference electrode by soaking in bleach for 10 min and sterile deionized water for 10 min. Sterilize the IDA by dipping in bleach for 5 s followed by sterile deionized water for 10 s before insertion into the reactor.</li>
		<li>Fill the electrochemical cell with sterile medium suitable for biofilm growth. For G. sulfurreducens17,18, use freshwater medium excluding fumarate. For biocathode MCL,9 use artificial seawater medium.9</li>
		<li>Connect the electrodes to a bipotentiostat. Connect one IDA electrode to the working lead 1, the other IDA electrode to the working lead 2, the reference electrode to the reference lead, and the counter electrode to the counter lead.</li>
	</ol>
	</li>
	<li>Electrochemical testing of IDAs. 
	NOTE: The major goal of this test is to make sure that the two electrodes are electrically isolated. All electrochemical techniques are available in the software used to control the bipotentiostat.
	<ol>
		<li>Perform control conductivity tests (before biofilm growth) to ensure proper IDA function using a bipotentiostat.
		<ol>
			<li>Measure the open circuit potential of each electrode for 1 min. 
			NOTE: On some instruments, open circuit potential must be achieved by using the galvanostatic collection program and setting the current to 0 mA.</li>
			<li>Measure the open circuit potential of electrode 2 while performing cyclic voltammetry on electrode 1 between +0.2 V to +0.6 V (vs. SHE) at 20 mV/s. 
			NOTE: Other bounds and scan rates for CV can be used if desired. However, avoid potentials that will result in hydrogen or oxygen generation.</li>
			<li>Verify that the open circuit potential measured at electrode 2 does not mirror the potential of electrode 1 during the CV using the potentiostat software. 
			NOTE: The open circuit potential of electrode 2 may change, but it should be independent of the potential of electrode 1.</li>
			<li>Repeat steps 2.2.1.1 through 2.2.1.3 except control the potential of electrode 2 and measure the open circuit potential of electrode 1.</li>
		</ol>
		</li>
		<li>Set the potential of working electrodes to the desired growth potential of the relevant electroactive biofilm using the bipotentiostat software. For example, use +0.5 V (vs. SHE) for Geobacter sulfurreducens or +0.31 V (vs. SHE) for biocathode MCL.</li>
	</ol>
	</li>
	<li>Grow relevant electroactive biofilm
	<ol>
		<li>Inoculate the electrochemical reactor from a stock culture/enrichment of the desired electrochemically active microorganisms using standard aseptic microbiological techniques. For standard tests, inoculate in a 1:20 ratio (inoculum to reactor volume) of an OD600 = 0.5 culture.</li>
		<li>Set the stirring in the reactor to the desired level (~200 rpm) and the incubator/ water bath to the desired temperature based on the growth conditions of the biofilm of interest. For G. sulfurreducens, use 30 &#176;C for optimal growth.</li>
		<li>Incubate the system based on specific requirements of the microorganism(s) of interest until the biofilm bridges the gap separating the two electrodes. For stationary G. sulfurreducens biofilm, incubate for ~7-10 days. For biocathode MCL, incubate ~7 days. In each case, control the temperature at 30 &#176;C.</li>
	</ol>
	</li>
</ol>
3. Electrochemical gating experiments
<ol>
	<li>Select the experimental parameters that will be used to determine the current-potential dependence for the gating measurements.
	<ol>
		<li>Determine the range of gate potentials that will be applied to the IDA to obtain the conducted current (ISD) versus gate potential (EG) curve for the system. 
		NOTE: The range of gate potentials examined should cover all potentials with possible redox activity. If no information about the system of interest is available, a wide potential range should be used (-0.55 to +0.6 V vs. SHE). A trial and error approach can be used to fine tune the range based on the system under study. Gate potential is defined as: 
		<img alt="Equation 1" src="/files/ftp_upload/54671/54671eq1.jpg" /> 
		where ED is the potential of the drain electrode and ES is the potential of the source electrode. The range of gate potentials used is constrained by the requirements and limitations of the specific system of interest.18 
		NOTE: Source and drain potentials that will result in oxygen and hydrogen evolution should be avoided as these processes can damage the biofilm.</li>
		<li>Determine the source-drain voltage (VSD) that will be used as the driving force for electron transport through the film from the source to the drain. Source-drain voltage is defined as: 
		<img alt="Equation 2" src="/files/ftp_upload/54671/54671eq2.jpg" /> 
		NOTE: The source-drain voltage should be sufficiently small so that the ISD scales linearly with VSD.17</li>
		<li>Choose a scan rate (v) at which EG is changed linearly with time that is independent of ISD so that the system can be approximated to be at steady state for each gate potential. 
		NOTE: A scan rate of less than 0.002 V/s is often used for biological systems.29,30</li>
		<li>Set up the bipotentiostat software to perform the gating measurements (i.e. sweep the gate potential) over the selected range, at the selected source-drain voltage, and at the desired scan rate based on the above considerations. 
		NOTE: For previous gating measurements using G. sulfurreducens,17,19 EG = -0.55 V to 0.6 V (vs. SHE), VSD = 0.01 V or 0.1 V, v = 0.001 V/s. For Biocathode MCL, EG = 0.25 V to 0.7 V (vs. SHE), VSD = 0.002 V, v = 0.0002 V/s.
		<ol>
			<li>Additionally, perform a baseline measurement with VSD = 0.000 V (i.e., a CV at each individual electrode taken simultaneously) at the same v chosen in Step 3.1.3.</li>
		</ol>
		</li>
		<li>Perform gating measurements using the conditions in 3.1.4 under both turnover (with soluble electron donor or acceptor present) and non-turnover (without soluble electron donor or acceptor) conditions. 
		NOTE: Non-turnover conditions are advantageous because measurements are not obscured by the oxidation or reduction of soluble compounds for cellular metabolism, although similar results should be obtained regardless of which condition is used after subtracting background currents (detailed in 3.1.8).
		<ol>
			<li>Achieve turnover conditions by using the same reactor medium as used for bacterial growth on the electrode. This medium contains a soluble electron donor, such as acetate for G. sulfurreducens,17 or acceptor, such as oxygen for Biocathode MCL.</li>
			<li>Achieve non-turnover conditions by making the same reactor medium used for bacterial growth on the electrode, except omit the soluble electron donor or acceptor. After ensuring that the potentiostat is off, remove the medium aseptically and add in fresh medium without the electron donor for anodic systems or acceptor for cathodic systems. Alternatively, a continuous flow system can be set up to slowly replace the medium with the desired medium for nonturnover conditions. 
			NOTE: If oxygen is the soluble electron acceptor (as for biocathode MCL), sparge the system with a mixture of ~15% CO2 and 85% N2 (or a gas mixture that will maintain the correct pH in the medium).</li>
		</ol>
		</li>
		<li>After completion of the gating measurements, use the potentiostat software to set the potential of each electrode back to the growth potential to allow the system to re-stabilize (using the same values as in 2.2.2).</li>
		<li>If all the conditions described above are met (ISD is independent of v and scales linearly with VSD), convert ISD values to conductivity using the following equations as previously described31 
		<img alt="Equation 3" src="/files/ftp_upload/54671/54671eq3.jpg" /> 
		<img alt="Equation 4" src="/files/ftp_upload/54671/54671eq4.jpg" /></li>
		<li>where G is the conductance and S is a scaling factor that is system dependent, and factors in variables such as electrode size, gap size, and biofilm height. For certain systems, S can be determined from predetermined equations.31 Alternatively, S can be calculated numerically using a modeling software, as detailed previously.17</li>
		<li>Subtract the background currents to identify the shape and magnitude of the conducted current. Either subtract the current generated at VSD = 0.00 V from the current generated at VSD = 0.01 V or subtract the drain current from the source current generated with a VSD = 0.01 V. Either method removes background currents, leaving only the conducted current.</li>
	</ol>
	</li>
	<li>Temperature dependent electrochemical gating measurements
	<ol>
		<li>Determine the range of temperatures of interest. This range is highly dependent on the system under study, however physiologically relevant temperatures should be used. 
		NOTE: Previous studies have used a temperature range of 10 &#176;C to 30 &#176;C to study electron transport under physiologically relevant conditions for mesophilic microorganisms.17</li>
		<li>Obtain a recirculating water bath or incubator to regulate reactor temperature and a control reactor to ensure that the set point and actual temperature of the medium is the same.
		<ol>
			<li>Place a thermometer or thermocouple in a control reactor where the IDA would be.</li>
		</ol>
		</li>
		<li>Make ISD measurements (see 3.1.4) over the range of temperatures selected under turnover and nonturnover (described in Step 3.1.5) conditions using the bipotentiostat following one of the two procedures described below.
		<ol>
			<li>Generate ISD vs. EG curves, as described above (see 3.1), for each temperature over the temperature range of interest. For materials that exhibit redox conductivity, such as G. sulfurreducens and Biocathode MCL, the peak current from each temperature is used to determine the ISD vs. T dependence. 
			NOTE: This method is used to generate a full ISD vs. EG curve for each temperature, but requires more time than the second option.</li>
			<li>Alternately, set and hold the IDA at the gate potential that yields maximum conducted current using the bipotentiostat (obtained from the ISD vs EG curve, Step 3.1.4) and record the maximum conducted current using the bipotentiostat while the temperature is cycled between the range of temperatures selected using the onboard controls of the water bath or incubator. 
			NOTE: With the electrode/reactor geometry used here, the ISD and T are allowed to stabilize for at least 20 min and an average stable ISD is used for each temperature. More or less stabilization time may be required based on the specific system. This method is faster than the first and causes less stress to the biofilm. However, full gating curves are not generated.</li>
			<li>Cycle the temperature from one set point to the other and back again using the onboard controls of the incubator or the water bath to determine the reversibility of the reaction to ensure that the temperature cycling does not harm the biofilm.</li>
		</ol>
		</li>
		<li>Reset the temperature back to the normal growth temperature using the onboard controls of the incubator or water bath and allow the system to stabilize. 
		NOTE: For a redox conductor, the ISD versus 1/T data can be fit with the Arrhenius rate expression, which allows the calculation of activation energy as follows: 
		<img alt="Equation 5" src="/files/ftp_upload/54671/54671eq5.jpg" /> 
		<img alt="Equation 6" src="/files/ftp_upload/54671/54671eq6.jpg" /> 
		where Ea is the activation energy for electron transfer between adjacent redox cofactors and k is the Boltzmann constant.</li>
	</ol>
	</li>
</ol>

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S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tunable+metallic-like+conductivity+in+microbial+nanowire+networks.">Tunable metallic-like conductivity in microbial nanowire networks.</a> Nat Nano. 6, 573-579 (2011).</li><li>Ing, N. L., Nusca, T. D., Hochbaum, A. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Geobacter+sulfurreducens+pili+support+ohmic+electronic+conduction+in+aqueous+solution.">Geobacter sulfurreducens pili support ohmic electronic conduction in aqueous solution.</a> Phys Chem Chem Phys. 19, 21791-21799 (2017).</li><li>Fricke, K., Harnisch, F., Schr&#246;der, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=On+the+use+of+cyclic+voltammetry+for+the+study+of+anodic+electron+transfer+in+microbial+fuel+cells.">On the use of cyclic voltammetry for the study of anodic electron transfer in microbial fuel cells.</a> Energ Environ Sci. 1, 144-147 (2008).</li><li>Marsili, E., Rollefson, J. B., Baron, D. B., Hozalski, R. M., Bond, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microbial+biofilm+voltammetry:+direct+electrochemical+characterization+of+catalytic+electrode-attached+biofilms.">Microbial biofilm voltammetry: direct electrochemical characterization of catalytic electrode-attached biofilms.</a> Appl Environ Microbiol. 74, 7329-7337 (2008).</li><li>Kankare, J., Kupila, E. -. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In-situ+conductance+measurement+during+electropolymerization.">In-situ conductance measurement during electropolymerization.</a> J Electroanal Chem. 322, 167-181 (1992).</li><li>Byun, H. S., Pirbadian, S., Nakano, A., Shi, L., El-Naggar, M. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetic+Monte+Carlo+Simulations+and+Molecular+Conductance+Measurements+of+the+Bacterial+Decaheme+Cytochrome+MtrF.">Kinetic Monte Carlo Simulations and Molecular Conductance Measurements of the Bacterial Decaheme Cytochrome MtrF.</a> Chem Electro Chem. 1, 1932-1939 (2014).</li><li>El Kasmi, A., Wallace, J. M., Bowden, E. F., Binet, S. M., Linderman, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Controlling+interfacial+electron-transfer+kinetics+of+cytochrome+c+with+mixed+self-assembled+monolayers.">Controlling interfacial electron-transfer kinetics of cytochrome c with mixed self-assembled monolayers.</a> J Am Chem Soc. 120, 225-226 (1998).</li><li>Bortolotti, C. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Reorganization+Energy+in+Cytochrome+c+is+Controlled+by+the+Accessibility+of+the+Heme+to+the+Solvent.">The Reorganization Energy in Cytochrome c is Controlled by the Accessibility of the Heme to the Solvent.</a> J Phys Chem Lett. 2, 1761-1765 (2011).</li><li>Gallaway, J. W., Calabrese Barton, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetics+of+Redox+Polymer-Mediated+Enzyme+Electrodes.">Kinetics of Redox Polymer-Mediated Enzyme Electrodes.</a> J Am Chem Soc. 130, 8527-8536 (2008).</li><li>Thackeray, J. W., White, H. S., Wrighton, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Poly(3-methylthiophene)-coated+electrodes:+optical+and+electrical+properties+as+a+function+of+redox+potential+and+amplification+of+electrical+and+chemical+signals+using+poly(3-methylthiophene)-based+microelectrochemical+transistors.">Poly(3-methylthiophene)-coated electrodes: optical and electrical properties as a function of redox potential and amplification of electrical and chemical signals using poly(3-methylthiophene)-based microelectrochemical transistors.</a> J Phys Chem-US. 89, 5133-5140 (1985).</li><li>Jugnet, Y., Tourillon, G., Duc, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+of+Intrinsic+Extended+&#960;-Bonding+Band+and+Metalliclike+Behavior+in+Undoped+and+Doped+Electropolymerized+Poly+(3-methylthiophene)+Films.">Evidence of Intrinsic Extended &#960;-Bonding Band and Metalliclike Behavior in Undoped and Doped Electropolymerized Poly (3-methylthiophene) Films.</a> Phys Rev Lett. 56, 1862-1865 (1986).</li></ol>

The authors have nothing to disclose.

During the setup of the IDA, it is critical to test that the source and the drain are not shorted together prior to electrochemical gating measurements, as this will alter the ISD vs. EG curve and could lead to erroneous results and interpretations. It is also critical to select VSD and v such that the current is linearly dependent on VSD and independent of v. If this is not the case, then the equations described above cannot be utilized to calculate conductivity.
<p class=...

Extracellular electron transport (EET) is a process that enables certain microorganisms to transport electrons between intracellular metabolic processes and insoluble electron acceptors or donors that reside outside the cell, ranging from natural minerals to electrodes. In some cases, EET enables microorganisms to form electrically conductive multi-cell thick biofilms on electrode surfaces, in which cells not in direct contact with the electrode can still utilize it as a metabolic electron acceptor or donor. There is considerable interest in such biofilms as electrode catalysts for various applications, such as microbial electrosynthesis, contaminant sensing/removal, and remote energy generation and storage,7,8,9,10,11,12,13,14 due to the diversity of metabolic processes performed by microorganisms and the durability of microbial biofilms compared to enzyme-based bioelectrodes.15,16 In addition, EET pathways may potentially be utilized to electrically control or signal changes in naturally occurring or genetically engineered microbial metabolic processes involved, for example, in production of a desired product or detection of a target analyte or stimulus. The electrical conductivity of electrocatalytic biofilms, which sets them apart from other biological materials, is a central aspect of their electrocatalytic properties, yet little is understood about the underlying EET process in the electrode environment, and that which is known is highly contested.17,18,19,20,21,22,23,24
Described here is a 2-electrode method to measure conductivity through living, electrode-grown biofilms using interdigitated electrode arrays (IDAs). IDAs consist of parallel rectangular electrodes patterned on flat glass surface such that every other band is connected at opposite sides of the array resulting in 2 electrodes (the source and drain). Careful examination of an IDA (see for example, Figure 6.12b of ref #1) reveals that that the gaps separating adjacent bands are also connected in such a way as to form a single gap that weaves back and forth across the array separating the two electrodes. The result is a long and narrow gap separating the source and drain electrodes, yielding very high source-drain currents when a conductive material is formed, cast, polymerized, or grown (in the case of the type of biofilms considered here) over the array. In addition, the small size of the electrodes results in small background current due to capacitance charging and to change in oxidation state of the conductive material with change in gate potential, since the amount of material needed to make conductivity measurements using IDAs is so small. The technique of IDA-based electrochemical gating described here, developed to characterize thin film conductive polymers,2,3,4,25 has only recently been applied to living systems.18 Another technique used to measure conductivity of living biofilms utilized a large format split source and drain electrodes and source meters to set the gate potential.26,27 However, concerns over these methods have been detailed previously.18
The protocol below encapsulates our experience with making conductivity measurements of living Geobacter sulfurreducens and biocathode MCL biofilms. G. sulfurreducens is a model electrode reducing organism able to use insoluble materials, including electrodes, as the sole metabolic electron acceptor. Additionally, it forms thick biofilms that are able to transport electrons over multiple cell lengths, making it an ideal model organism to study anodic long-distance extracellular electron transfer. We also include details for the study of biocathode MCL, an aerobic, autotrophic mixed community biofilm isolated from the cathode of a benthic microbial fuel cell. Biocathode MCL (named for the three primary constituents &#8211; Marinobacter, Chromatiaceaea and Labrenzia) is capable of oxidizing an electrode as its sole electron donor and transporting electrons over multiple cell lengths, making it an interesting cathodic system to study. Additionally, biocathode MCL has the highest reported conductivity for a living system to date using these methods. The inclusion of these diverse electroactive biofilms in this protocol is meant to highlight that this technique is applicable to measure the transport of electrons through any living biofilm able to electrically interact with electrodes.

<table><tbody><tr><td>IDAs</td><td>CH Instruments</td><td>012125</td><td>Manufactured by ALS-Japan; sold by CH Instruments</td></tr><tr><td>Wire</td><td>Digikey</td><td>W7-ND</td><td></td></tr><tr><td>Conductive silver epoxy</td><td>Electron microscopy sciences</td><td>12670-EE</td><td></td></tr><tr><td>Insulating material</td><td>3M</td><td>2131-B</td><td>Scotchast flame retardant compound</td></tr><tr><td>15 mL conical centrifuge tube</td><td>VWR</td><td>89004-368</td><td></td></tr><tr><td>21g needle</td><td>VWR</td><td>BD-305165</td><td></td></tr><tr><td>5 mL pipette tips</td><td>VWR</td><td>82018-842</td><td></td></tr><tr><td>5 mL pipettor</td><td>VWR</td><td>89079-976</td><td></td></tr><tr><td>Freshwater medium components</td><td>Sigma Aldrich</td><td>All standard laboratory chemicals</td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Ammonium chloride</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Sodium phosphate monobasic</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Sodium bicarbonate</td><td></td><td></td><td></td></tr><tr><td>Artificial seawater medium components</td><td>Sigma Aldrich</td><td>All standard laboratory chemicals</td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Sodium chloride</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Magnesium chloride hexahydrate</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Magnesium sulfate heptahydrate</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Potassium chloride</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Sodium bicarbonate</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Calcium chloride dihydrate</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Ammonium chloride</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;&amp;nbsp;&amp;nbsp; Potassium phosphate dibasic</td><td></td><td></td><td></td></tr><tr><td>Ag/AgCl reference electrode</td><td>Basi</td><td>MF-2079</td><td></td></tr><tr><td>Graphite rod counter electrode</td><td>Electron microscopy sciences</td><td>70230</td><td></td></tr><tr><td>Recirculating water bath</td><td>Thermo Scientific</td><td>152-5256</td><td></td></tr><tr><td>Bipotentiostat</td><td>Pine Instruments</td><td>WD-20</td><td>http://www.voltammetry.net/pine/aftermath/user</td></tr><tr><td>Stir bars</td><td>VWR</td><td>58947-114</td><td></td></tr><tr><td>&lt;em&gt;G. sulfurreducens&lt;/em&gt; culture</td><td>ATCC</td><td>51573</td><td></td></tr><tr><td>Jacketed reactor</td><td>Pine Instruments</td><td>RRPG085</td><td></td></tr></tbody></table>

characterizing electron transport through living biofilms

Here we demonstrate the method of electrochemical gating used to characterize electrical conductivity of electrode-grown microbial biofilms under physiologically relevant conditions.1 These measurements are performed on living biofilms in aqueous medium using source and drain electrodes patterned on a glass surface in a specialized configuration referred to as an interdigitated electrode (IDA) array. A biofilm is grown that extends across the gap connecting the source and drain. Potentials are applied to the electrodes (ES and ED) generating a source-drain current (ISD) through the biofilm between the electrodes. The dependency of electrical conductivity on gate potential (the average of the source and drain potentials, EG = [ED + ES]/2) is determined by systematically changing the gate potential and measuring the resulting source-drain current. The dependency of conductivity on gate potential provides mechanistic information about the extracellular electron transport process underlying the electrical conductivity of the specific biofilm under investigation. The electrochemical gating measurement method described here is based directly on that used by M. S. Wrighton2,3 and colleagues and R. W. Murray4,5,6 and colleagues in the 1980&#39;s to investigate thin film conductive polymers.

IDAs were wired, insulated and tested to ensure that the two electrodes were electrically isolated from each other (Figure 1). Reactors were assembled, inoculated with G. sulfurreducens, and incubated until a biofilm bridged the gap between the electrodes. The G. sulfurreducens biofilm can be visually seen to be covering the array. Other biofilms may require the researcher to do an electrochemical gating measurements to see if the two electr...

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Characterizing Electron Transport through Living Biofilms

A protocol for measuring electrical conductivity of living microbial biofilms under physiologically relevant conditions is presented.

Here we demonstrate the method of electrochemical gating used to characterize electrical conductivity of electrode-grown microbial biofilms under ...

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8.4K Views.  Naval Research Laboratory. Een protocol voor het meten van de elektrische geleiding van levende micro-organismen biofilms fysiologisch relevante voorwaarden wordt gepresenteerd.

Video: Characterizing Electron Transport through Living Biofilms

Waste Water Derived Electroactive Microbial Biofilms: Growth, Maintenance, and Basic Characterization

The growth of anodic electroactive microbial biofilms from waste water inocula in a fed-batch reactor is demonstrated using a three-electrode setup controlled by a potentiostat. Thereby the use of potentiostats allows an exact adjustment of the electrode potential and ensures reproducible microbial culturing conditions. During growth the current production is monitored using chronoamperometry (CA). Based on these data the maximum current density (jmax) and the coulombic efficiency (CE) are discussed as measures for characterization of the bioelectrocatalytic performance. Cyclic voltammetry (CV), a nondestructive, i.e. noninvasive, method, is used to study the extracellular electron transfer (EET) of electroactive bacteria. CV measurements are performed on anodic biofilm electrodes in the presence of the microbial substrate, i.e. turnover conditions, and in the absence of the substrate, i.e. nonturnover conditions, using different scan rates. Subsequently, data analysis is exemplified and fundamental thermodynamic parameters of the microbial EET are derived and explained: peak potential (Ep), peak current density (jp), formal potential (Ef) and peak separation (&#916;Ep). Additionally the limits of the method and the state-of the art data analysis are addressed. Thereby this video-article shall provide a guide for the basic experimental steps and the fundamental data analysis.

Chronoamperometric growth of anodic electroactive microbial biofilms in a fed-batch reactor using a three-electrode setup, controlled by a potentiostat, is demonstrated. The extracellular electron transfer is characterized using cyclic voltammetry in the presence of the electron donor and in the absence of the electron donor. Fundamental data analysis is demonstrated.

The growth of anodic electroactive microbial biofilms from waste water inocula in a fed-batch reactor is demonstrated using a three-electrode setup ...

Environment

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM c-Cyts) has recently emerged as a novel whole-cell analytical method to characterize the bacterial electron transport from the respiratory chain to the cell exterior, referred to as the extracellular electron transport (EET). While the pathway and kinetics of the electron flow during the EET reaction have been investigated, a whole-cell electrochemical method to examine the impact of cation transport associated with EET has not yet been established. In the present study, an example of a biochemical technique to examine the deuterium kinetic isotope effect (KIE) on EET through OM c-Cyts using a model microbe, Shewanella oneidensis MR-1, is described. The KIE on the EET process can be obtained if the EET through OM c-Cyts acts as the rate-limiting step in the microbial current production. To that end, before the addition of D2O, the supernatant solution was replaced with fresh media containing a sufficient amount of the electron donor to support the rate of upstream metabolic reactions, and to remove the planktonic cells from a uniform monolayer biofilm on the working electrode. Alternative methods to confirm the rate-limiting step in microbial current production as EET through OM c-Cyts are also described. Our technique of a whole-cell electrochemical assay for investigating proton transport kinetics can be applied to other electroactive microbial strains.

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Here we present a protocol of whole-cell electrochemical experiments to study the contribution of proton transport to the rate of extracellular electron transport via the outer-membrane cytochromes complex in Shewanella oneidensis MR-1.

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM ...

Biochemistry

Characterization of Aquatic Biofilms with Flow Cytometry

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond quickly to environmental changes and can be thus used as indicators of water quality. Currently, biofilm assessment is mostly based on integrative and functional endpoints, such as photosynthetic or respiratory activity, which do not provide information on the biofilm community structure. Flow cytometry and computational visualization offer an alternative, sensitive, and easy-to-use method for assessment of the community composition, particularly of the photoautotrophic part of freshwater biofilms. It requires only basic sample preparation, after which the entire sample is run through the flow cytometer. The single-cell optical and fluorescent information is used for computational visualization and biological interpretation. Its main advantages over other methods are the speed of analysis and the high-information-content nature. Flow cytometry provides information on several cellular and biofilm traits in a single measurement: particle size, density, pigment content, abiotic content in the biofilm, and coarse taxonomic information. However, it does not provide information on biofilm composition on the species level. We see high potential in the use of the method for environmental monitoring of aquatic ecosystems and as an initial biofilm evaluation step that informs downstream detailed investigations by complementary and more detailed methods.

Flow cytometry in combination with visual clustering offers an easy-to-use and fast method for studying aquatic biofilms. It can be used for biofilm characterization, detection of changes in biofilm community structure, and detection of abiotic particles embedded in the biofilm.

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond ...

In Situ Characterization of Shewanella oneidensis MR1 Biofilms by SALVI and ToF-SIMS

Bacterial biofilms are surface-associated communities that are vastly studied to understand their self-produced extracellular polymeric substances (EPS) and their roles in environmental microbiology. This study outlines a method to cultivate biofilm attachment to the System for Analysis at the Liquid Vacuum Interface (SALVI) and achieve in situ chemical mapping of a living biofilm by time-of-flight secondary ion mass spectrometry (ToF-SIMS). This is done through the culturing of bacteria both outside and within the SALVI channel with our specialized setup, as well as through optical imaging techniques to detect the biofilm presence and thickness before ToF-SIMS analysis. Our results show the characteristic peaks of the Shewanella biofilm in its natural hydrated state, highlighting upon its localized water cluster environment, as well as EPS fragments, which are drastically different from the same biofilm's dehydrated state. These results demonstrate the breakthrough capability of SALVI that allows for in situ biofilm imaging with a vacuum-based chemical imaging instrument.

In Situ Characterization of Shewanella oneidensis MR1 Biofilms by SALVI and ToF-SIMS

This article presents a method for growing a biofilm for in situ time-of-flight secondary ion mass spectrometry for chemical mapping in its hydrated state, enabled by a microfluidic reactor, System for Analysis at the liquid Vacuum Interface. The Shewanella oneidensis MR-1 with green fluorescence protein was used as a model.

Bacterial biofilms are surface-associated communities that are vastly studied to understand their self-produced extracellular polymeric substances (EPS) ...

Concurrent Quantification of Cellular and Extracellular Components of Biofilms

Confocal laser scanning microscopy (CLSM) is a powerful tool for investigation of biofilms. Very few investigations have successfully quantified concurrent distribution of more than two components within biofilms because: 1)&#160;selection of fluorescent dyes having minimal spectral overlap is complicated, and 2)&#160;quantification of multiple fluorochromes poses a multifactorial problem. Objectives: Report a methodology to quantify and compare concurrent 3-dimensional distributions of three cellular/extracellular components of biofilms grown on relevant substrates. Methods: The method consists of distinct, interconnected steps involving biofilm growth, staining, CLSM imaging, biofilm structural analysis and visualization, and statistical analysis of structural parameters. Biofilms of Streptococcus mutans (strain UA159) were grown for 48 hr on sterile specimens of Point 4 and TPH3 resin composites. Specimens were subsequently immersed for 60 sec in either Biot&#232;ne PBF (BIO) or Listerine Total Care (LTO) mouthwashes, or water (control group; n=5/group). Biofilms were stained with fluorochromes for extracellular polymeric substances, proteins and nucleic acids before imaging with CLSM. Biofilm structural parameters calculated using ISA3D image analysis software were biovolume and mean biofilm thickness. Mixed models statistical analyses compared structural parameters between mouthwash and control groups (SAS software; &#945;=0.05). Volocity software permitted visualization of 3D distributions of overlaid biofilm components (fluorochromes). Results: Mouthwash BIO produced biofilm structures that differed significantly from the control (p&#60;0.05) on both resin composites, whereas LTO did not produce differences (p&#62;0.05) on either product. Conclusions: This methodology efficiently and successfully quantified and compared concurrent 3D distributions of three major components within S. mutans biofilms on relevant substrates, thus overcoming two challenges to simultaneous assessment of biofilm components. This method can also be used to determine the efficacy of antibacterial/antifouling agents against multiple biofilm components, as shown using mouthwashes. Furthermore, this method has broad application because it facilitates comparison of 3D structures/architecture of biofilms in a variety of disciplines.

A protocol for the concurrent quantification and comparison of three cellular and extracellular components within biofilms is presented. The methodology involves the use of confocal laser scanning microscopy, biofilm structural analysis and visualization software, and statistical analysis software.

Confocal laser scanning microscopy (CLSM) is a powerful tool for investigation of biofilms. Very few investigations have successfully quantified ...

Immunology and Infection

Methods for Characterizing the Co-development of Biofilm and Habitat Heterogeneity

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is strongly regulated by the surrounding flow and nutritional environment. Biofilm growth also increases the heterogeneity of the local microenvironment by generating complex flow fields and solute transport patterns. To investigate the development of heterogeneity in biofilms and interactions between biofilms and their local micro-habitat, we grew mono-species biofilms of Pseudomonas aeruginosa and dual-species biofilms of P. aeruginosa and Escherichia coli under nutritional gradients in a microfluidic flow cell. We provide detailed protocols for creating nutrient gradients within the flow cell and for growing and visualizing biofilm development under these conditions. We also present protocols for a series of optical methods to quantify spatial patterns in biofilm structure, flow distributions over biofilms, and mass transport around and within biofilm colonies. These methods support comprehensive investigations of the co-development of biofilm and habitat heterogeneity.

Biofilms have complex interactions with their surrounding environment. To comprehensively investigate biofilm-environment interactions, we present here a series of methods to create heterogeneous chemical environment for biofilm development, to quantify local flow velocity, and to analyze mass transport in and around biofilm colonies.

Biofilms are surface-attached microbial communities that have complex structures and produce significant spatial heterogeneities. Biofilm development is ...

Bioengineering

Methodologies for Studying B. subtilis Biofilms as a Model for Characterizing Small Molecule Biofilm Inhibitors

This work assesses different methodologies to study the impact of small molecule biofilm inhibitors, such as D-amino acids, on the development and resilience of Bacillus subtilis biofilms. First, methods are presented that select for small molecule inhibitors with biofilm-specific targets in order to separate the effect of the small molecule inhibitors on planktonic growth from their effect on biofilm formation. Next, we focus on how inoculation conditions affect the sensitivity of multicellular, floating B. subtilis cultures to small molecule inhibitors. The results suggest that discrepancies in the reported effects of such inhibitors such as D-amino acids are due to inconsistent pre-culture conditions. Furthermore, a recently developed protocol is described for evaluating the contribution of small molecule treatments towards biofilm resistance to antibacterial substances. Lastly, scanning electron microscopy (SEM) techniques are presented to analyze the three-dimensional spatial arrangement of cells and their surrounding extracellular matrix in a B. subtilis biofilm. SEM facilitates insight into the three-dimensional biofilm architecture and the matrix texture. A combination of the methods described here can greatly assist the study of biofilm development in the presence and absence of biofilm inhibitors, and shed light on the mechanism of action of these inhibitors.

Methodologies for Studying B. subtilis Biofilms as a Model for Characterizing Small Molecule Biofilm Inhibitors

This study presents the development of reproducible methodologies to study biofilm inhibitors and their effects on Bacillus subtilis multicellularity.

This work assesses different methodologies to study the impact of small molecule biofilm inhibitors, such as D-amino acids, on the development and ...

Chronoamperometry Analysis of EET in Lactiplantibacillus plantarum with DHNA

Characterizing Mediated Extracellular Electron Transfer in Lactic Acid Bacteria with a Three-Electrode, Two-Chamber Bioelectrochemical System

Many bacteria perform extracellular electron transfer (EET), whereby electrons are transferred from the cell to an extracellular terminal electron acceptor. This electron acceptor can be an electrode and electrons can be delivered indirectly via a redox-active mediator molecule. Here, we present a protocol to study mediated EET in Lactiplantibacillus plantarum, a probiotic lactic acid bacterium widely used in the food industry, using a bioelectrochemical system. We detail how to assemble a three-electrode, two-chambered bioelectrochemical system and provide guidance on characterizing EET in the presence of a soluble mediator using chronoamperometry and cyclic voltammetry techniques. We use representative data from 1,4-dihydroxy-2-naphthoic acid (DHNA)-mediated EET experiments with L. plantarum to demonstrate data analysis and interpretation. The techniques described in this protocol can open new opportunities for electro-fermentation and bioelectrocatalysis. Recent applications of this electrochemical technique with L. plantarum demonstrated an acceleration of metabolic flux towards producing fermentation end-products, which are critical flavor components in food fermentation. As such, this system has the potential to be further developed to alter flavors in food production or produce valuable chemicals.

Here we present a protocol for characterizing mediated extracellular electron transfer (EET) in lactic acid bacteria using a three-electrode, two-chamber bioelectrochemical system. We illustrate this method with Lactiplantibacillus plantarum and the redox mediator 1,4-dihydroxy-2-naphthoic acid and provide a thorough description of the electrochemical techniques used to evaluate mediated EET.

Many bacteria perform extracellular electron transfer (EET), whereby electrons are transferred from the cell to an extracellular terminal electron ...

Paraffin Embedding and Thin Sectioning of Microbial Colony Biofilms for Microscopic Analysis

Sectioning via paraffin embedding is a broadly established technique in eukaryotic systems. Here we provide a method for the fixation, embedding, and sectioning of intact microbial colony biofilms using perfused paraffin wax. To adapt this method for use on colony biofilms, we developed techniques for maintaining each sample on its growth substrate and laminating it with an agar overlayer, and added lysine to the fixative solution. These optimizations improve sample retention and preservation of micromorphological features. Samples prepared in this manner are amenable to thin sectioning and imaging by light, fluorescence, and transmission electron microscopy. We have applied this technique to colony biofilms of Pseudomonas aeruginosa, Pseudomonas synxantha, Bacillus subtilis, and Vibrio cholerae. The high level of detail visible in samples generated by this method, combined with reporter strain engineering or the use of specific dyes, can provide exciting insights into the physiology and development of microbial communities.

We describe fixation, paraffin embedding, and thin sectioning techniques for microbial colony biofilms. In prepared samples, biofilm substructure and reporter expression patterns can be visualized by microscopy.

Sectioning via paraffin embedding is a broadly established technique in eukaryotic systems. Here we provide a method for the fixation, embedding, and ...

Measuring the Migration and Biofilm Formation of Various Bacteria

As microbes that thrive in the host body primarily have adaptive abilities that facilitate their survival, methods for classifying and identifying their nature would be beneficial in facilitating their characterization. Currently, most studies focus only on one specific characterization method; however, the isolation and identification of microorganisms from the host is a continuous process and usually requires several combinatorial characterization methods. Herein, we describe methods of identifying the microbial biofilm-forming ability, the microbial respiration state, and their chemotaxis behavior. The methods are used to identify five microbes, three of which were isolated from the bone tissue of Sprague-Dawley (SD) rats (Corynebacterium stationis, Staphylococcus cohnii subsp. urealyticus, and Enterococcus faecalis) and two from the American Type Culture Collection (ATCC)-Staphylococcus aureus ATCC 25923&#160;and Enterococcus faecalis V583. The microbes isolated from the SD rat bone tissue include the gram-positive microbes. These microbes have adapted to thrive under stressful and nutrient-limiting environments within the bone matrix. This article aims to provide the readers with the specific know-how of determining the nature and behavior of the isolated microbes within a laboratory setting.

Here, we present a practical method for the isolation and identification of microorganisms within the host. In this way, the physicochemical properties of microorganisms and possible ways of living in the host are clearly described.

As microbes that thrive in the host body primarily have adaptive abilities that facilitate their survival, methods for classifying and identifying their ...

Characterizing Electron Transport through Living Biofilms

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