F.H. acknowledges support by the BMBF (Research Award &#8220;Next generation biotechnological processes - Biotechnology 2020+&#8221;) and the Helmholtz-Association (Young Investigators Group).

As the selection procedure in the bioelectrochemical fed-batch reactor is very specific due to the choice of substrate and anoxic conditions, working in sterile conditions is not necessary for the demonstrated example.
1. Preparation of Growth Medium
<ol>
	<li>Prepare a suitable amount of the growth medium by using a phosphate buffer according to Kim et al.13&#160;(see Table 1). Here, sodium acetate (10 mM) is serving as substrate. The growth medium used in this article is only one example, and depending on the experimental purpose can vary in its components.</li>
	<li>Add 1 ml&#160;of primary waste water per 20&#160;ml&#160;of growth medium as inoculum. The waste water sample is taken from a municipal waste water treatment plant after the mechanical filtration (see for example Figure 2) and can be stored up to three months until usage in a closed container at 4 &#176;C. 
	(Note: Depending on the objective of the experiment the exact initial cell count can be important and then should be determined. In the demonstrated example, the initial cell count is not important, as the biofilm reaches steady-state conditions.)</li>
	<li>Flush the medium including the inoculum with nitrogen for at least 30 min to deaerate it, by inserting a needle in the medium and turn on the nitrogen stream. &#160;The stream should be strong enough that you can observe a complete mixing of the medium. 
	(Note: Depending on the volume of the medium one should even consider extending the time of deaeration. During deaeration setup the fed-batch reactor (steps 2.1-2.2))</li>
</ol>
2. Setup of Fed-batch Reactor
<ol>
	<li>Take a suitable reactor vessel-&#160;here a modified 4-necked 250 ml round-bottom flask is used. 
	(Note: In general the vessel should have at least four inlets. These are necessary for the three electrodes and for sampling/ media exchange. Produced gas volumes can be neglected when the reactor is opened regularly due to sampling or medium change, alternatively a bubble counter should be installed. In order to avoid unwanted phototropic growth, the reactors should be shielded from direct light exposure.)</li>
	<li>For the three-electrode setup (see Figure 3 and section 3 for details) place the reference electrode (RE) (here Ag/AgCl, sat. KCl, 0.197 V vs. SHE), the working electrode (WE) and the counter electrode (CE) in the flask through individual inlets via silicone plugs &#8211; seal air-tight. 
	(Note: As WE and CE graphite rods are used. The wire of the electrodes is pulled through the silicon plug and fixed externally. The reference electrode is put through the silicon plug via a drilled hole. To avoid short-circuits, make sure that there is no physical contact of the electrodes, even while stirring or media recirculation. Place the electrodes that they are completely covered with medium solution. For sealing, wrap the silicon plug of every inlet with Parafilm.)</li>
	<li>Pour the growth medium including the inoculum (here 250 ml) through the sampling port into the reactor while flushing with nitrogen.</li>
	<li>Close the last inlet with a silicone plug and seal it air-tight. Note: Traces of oxygen will not harm the mixed species biofilms. They are able to withstand and even consume low levels of oxygen which in turn may decrease the CE.</li>
</ol>
3. Chronoamperometric Biofilm Growth
<ol>
	<li>Connect the electrodes to the respective cables of the potentiostat. Place the vessel in a temperature controlled chamber, here 35 &#176;C is used31, in order to ensure constant environmental conditions.</li>
	<li>Open the software to control the potentiostat and choose the technique: Chronoamperometry. Set the parameters for example as follows: A constant potential EWE of 0.2 V (vs. Ag/AgCl) for a duration t of 500 hr (&#8776; 20 days). The current is recorded in time steps dti every minute. 
	Note: The choice of the applied potential is dependent on the studied system/ microorganism. Generally anode processes are studied at positive potential and cathode processes at negative potentials. One can also deduce a suitable potential from the standard potentials of the electron donor/ acceptor and formal potential of the expected microbial electron transfer site, here as a starting point, please refer to recent literature.</li>
	<li>Start the chronoamperometry. A graph of the measured current over time is plotted by the software, the observed maximum current (imax) is an important online parameter, as discussed in sections 4 and 5.&#160;</li>
</ol>
4. Biofilm Maintenance
<ol>
	<li>Depending on the environmental conditions, e.g. the temperature, an oxidative current is starting to flow and reaches a plateau (see Figure 4). After reaching zero current again, pause the chronoamperometric measurement and replenish the medium solution. For a faster biofilm growth, replace the medium right after the plateau of the maximum current is reached. Open the sampling port and carefully, without destroying the thin biofilm, discharge the medium. While refilling the medium flush again with nitrogen. Close the sampling port and seal air-tight.</li>
	<li>Repeat the procedure of step 4.1. until the maximum oxidative current reaches a steady-state (see Figure 4). 
	(Note: A constant imax is believed to indicate a stable steady-state biofilm formation.)</li>
	<li>Take 1 ml&#160;samples of the fresh and the exchanged medium solutions for substrate analysis, e.g. by HPLC. The initial substrate (acetate) concentration (c0) as well as the concentration of the remaining substrate solution (c1) are subsequently needed for efficiency calculations (see section Data analysis). 
	Note: To achieve unaltered results, e.g.&#160;due to further substrate consumption, analyze the substrate samples immediately. If this is not possible, store the samples in a freezer at minimum -18 &#176;C for not longer than a month.)</li>
</ol>
5. Application of Cyclic Voltammetry to Study EET
<ol>
	<li>Perform cyclic voltammetry at the maxima and minima of the current flow as indicated (Figure 4). Depending on the question of interest the CV usually is performed when the system has reached steady-state to get comparable results. (Note: The generation of current is termed turnover conditions and related to the microbial oxidation of the substrate and subsequent electron transfer: microbial bioelectrocatalysis; whereas the nonturnover CVs are performed at (nearly) zero current flow in the CA.)</li>
	<li>Choose in the potentiostat controlling software the technique: Cyclic voltammetry, and set the appropriate parameters, here for instance: An initial potential of the EWE to Ei of -0.5 V vs. RE with a scan rate v=dE/dt of 1 mV/sec&#160;to a vertex potential E1 of 0.3 V vs. RE and back to the starting potential E2 = Ei of -0.5 V. 
	Note: When using cyclic voltammetry the applied voltage at the WE is changed linearly with time by the potentiostat (see Figure 5)- Box 2 provides principle rules for choosing the experiment parameters and troubleshooting).</li>
	<li>Start the cyclic voltammetry and record as many cycles as needed to perform a reproducible CV. Usually, at least 3 cycles are needed.&#160; The resulting plot is called a cyclic voltammogram (see also Figure 7). Note: Depending on the measurement conditions, different information can be gained as demonstrated in the next section.</li>
</ol>
6. Data Plotting
Here only the plotting of the data is discussed. A more detailed treatise follows in the next section. 
<ol>
	<li>Use appropriate data analysis software, here all data is exported as text and imported as ASCII file.</li>
	<li>Calculate and use preferentially current densities (j) and not absolute currents (i). 
	(Note: However, as different protocols for calculating j do exist, make clear which one was used. Here the current density per geometric, i.e. projected, surface area of the electrode is used.)</li>
	<li>Chronoamperometry: Plot the measured current density as function of the time.</li>
	<li>Cyclic voltammetry: Plot the measured current density as function of the potential of the WE Ewe.</li>
</ol>

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The authors declare that they have no competing financial interests.

The presented protocol shows an easy and straightforward way for growing electroactive microbial biofilms from waste water inoculate using a three-electrode setup in a fed-batch reactor. Chronoamperometry enables the growth and selection of electroactive microbial biofilms from diverse inocula as well as the fundamental characterization of the biofilms in term of maximum current density and coulombic efficiency. Cyclic voltammetry, being a noninvasive technique, allows performing fundamental studies on the EET thermodynamics of the microbial biofilm and thus to identify possible and actual EET transfer sites. Together, these parameters allow i) characterizing and comparing different microbial biofilms for identical conditions as well as ii) studying the impact of different operational conditions on identical electroactive microbial biofilms. However, these experiments can only be a starting point. On the one hand the multifaceted field of microbial bioelectrochemistry requires further and in-depth exploitation of electrochemical methods and protocols. This includes, for instance, models for extracting kinetic and mechanistic data as well as further methods like electrochemical impedance spectroscopy34,35. On the other hand electrochemical methods can provide no information on the molecular nature, the microbial composition, the spatial microbial organization, etc. Here further techniques being either noninvasive and coupled to electrochemistry or invasive are needed4.&#160; Box 2 provides a short discussion on the critical steps when performing the experiments as well as on initial troubleshooting.&#160;

From the engineering perspective it has to be taken into consideration that the electroactive biofilm anode is only one component of the current microbial bioelectrochemical systems and a multitude of other factors has to be analyzed for full systems36. Yet, we hope that this video-article will help to understand and tailor the core elements of BES, i.e. the microbial electrodes, in future.

The elucidation of the fundamentals of microbial extracellular electron transfer (EET) and its exploitation in engineered systems is a vital and rapidly increasing field of research and development1-3. The study of these electroactive (or bioelectrocatalytic or electrochemically active) microorganisms, including pure strains as well as defined co-cultures and complex consortia, requires a complex arsenal of techniques, methods and protocols4. These methods derive from diverse scientific disciplines, e.g. electrochemistry, materials science, microbiology, and provide insights on different hierarchical levels, i.e. from the entire microbial biofilm to single molecules. Thereby, electrochemistry and electrochemical methods represent the fundament of all activities.
Traditionally, fuel cell type setups were often used for the growth and maintenance of electroactive microbial cultures in the archetype of these engineered systems: microbial fuel cells (MFCs)5. Unfortunately, these types of MFCs often did not allow monitoring or even controlling the potential of a single electrode and thus, only limited insights in the electrode processes were possible-&#160;only the cell voltage was reported. Now it is more and more acknowledged that the individual monitoring and control of the potentials of single electrodes in MFCs represent a clear advantage, not only for fundamental research but also for engineering. Furthermore, with the diversification of the applications of microbial electrochemical technologies that now include, e.g. remediation, desalination, syntheses, and even biocomputing, in so-called microbial bioelectrochemical systems (BES)2,6,7 an external control of individual electrode potentials is often substantial. This control is usually achieved by using external power sources or potentiostats. Thereby the use of potentiostats allows-&#160;in contrast to other types of setups-&#160;an exact adjustment of the individual electrode potential. This is of high importance as the electrode represents the terminal microbial electron acceptor (for anodes) or electron donor (for cathodes) of the extracellular electron transfer. Figure 1 depicts the current state of knowledge on the microbial EET at anodes-&#160;see for details e.g.2,4,6,8,9&#160;The EET at the cathode is still mainly untapped10.
Thus the control of the electrode potential enables not only the use of a reproducible microbial culturing conditions but also its tailoring in terms of EET thermodynamics11. Within this article it is demonstrated how the characteristic parameters of electroactive microbial biofilms can be extracted from fed-batch reactor experiments using chronoamperometry (CA) and cyclic voltammetry (CV). This includes the performance parameters maximum current density (jmax) and coulombic efficiency (CE) as well as EET characteristics. Here the identification of the formal potentials (Ef) of possible and actual electron transfer sites will be in focus. The extraction of these parameters is shown on the example of waste water derived mixed culture biofilms that are dominated by Geobacter spec.12&#160;These waste water derived biofilms are gained by a simple electrochemical selection procedure&#160;as demonstrated by numerous research groups around the world13-16.
Dynamic electrochemical techniques to study microbial extracellular electron transfer
For a thermodynamic characterization of the EET of electroactive microbial cultures dynamic electrochemical techniques are needed. In general these techniques include potential controlled, i.e. voltammetric, and current controlled, i.e. galvanodynamic, techniques17. Thereby the potential controlled techniques are more prevalent and include for instance, square wave voltammetry and linear sweep voltammetry18. Most popular, however, is cyclic voltammetry (CV)19,20. CV is well-known and widespread in different fields of electrochemistry ranging from battery research via materials science to enzyme bioelectrochemistry21. Its application to living microbial cells can only be dated back more than one decade22 and increases significantly during the last years23,24. Especially the combination of CV and spectroscopic methods has provided unprecedented insights in the EET mechanisms and its molecular nature, e.g.25,26&#160;In parallel the theoretical framework for the extraction of thermodynamic and theoretical characteristics of the microbial EET was broadened and deepened27-30. In contrast, however, experimental mishandling or inadequate data-analysis often hamper the appropriate application of CV in the field of microbial bioelectrochemistry. Therefore, in addition to workshops (e.g. at the EU-ISMET 2012) and tutorials19, this video-article shall provide a basic guide for the conduction of cyclic voltammetric experiments on electroactive bacteria. Thereby it is focusing on the basic experimental steps and the fundamental data analysis and is not intended to provide the latest protocols of kinetic analysis. The most important termini for the following discussion are summarized in Box 1 (adapted from Harnisch and Freguia19 and modified).

<table border="0" cellpadding="0" cellspacing="0" width="740">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="29">
			<td height="29" style="height:29px;width:195px;">Primary waste water</td>
			<td style="width:197px;">Waste water treatment plant</td>
			<td style="width:125px;"></td>
			<td style="width:224px;">Taken after mechanical filtration</td>
		</tr>
		<tr height="44">
			<td height="44" style="height:44px;width:195px;">250 ml Round-bottom flasks with three additional inlets</td>
			<td style="width:197px;">Th.Geyer Berlin GmbH, Germany</td>
			<td style="width:125px;">9.011 845</td>
			<td style="width:224px;">Additional inlets were attached by our in-house glassblower</td>
		</tr>
		<tr height="71">
			<td height="71" style="height:71px;width:195px;">Ag/AgCl reference electrode (sat. KCl, 0.195 V vs. SHE) with ceramic diaphragm</td>
			<td style="width:197px;">Sensortechnik Meinsberg, Germany</td>
			<td style="width:125px;">SE 11</td>
			<td style="width:224px;"></td>
		</tr>
		<tr height="47">
			<td height="47" style="height:47px;width:195px;">Graphite rods (&#216;=1 cm, l=3 and 5 cm)</td>
			<td style="width:197px;">CP-Handels-GmbH, Wachtberg, Germany</td>
			<td style="width:125px;"></td>
			<td style="width:224px;">Used as electrode material</td>
		</tr>
		<tr height="48">
			<td height="48" style="height:48px;width:195px;">Stainless steel, No. 1.4301, Wire Fe/Cr18/Ni10</td>
			<td style="width:197px;">Goodfellow GmbH, Bad Nauheim, Germany</td>
			<td style="width:125px;">155-923-05</td>
			<td style="width:224px;"></td>
		</tr>
		<tr height="38">
			<td height="38" style="height:38px;width:195px;">Silicon plugs</td>
			<td style="width:197px;">Th.Geyer Berlin GmbH, Germany</td>
			<td style="width:125px;">9.230 512 and</td>
			<td style="width:224px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:195px;">Parafilm</td>
			<td style="width:197px;">neoLab, Heidelberg, Germany</td>
			<td style="width:125px;">3-1011</td>
			<td style="width:224px;">For sealing to maintain anaerobic conditions</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:195px;">Potentiostat/ Galvanostat Model MPG2</td>
			<td style="width:197px;">BioLogic Science Instruments, France</td>
			<td style="width:125px;"></td>
			<td style="width:224px;"></td>
		</tr>
		<tr height="51">
			<td height="51" style="height:51px;width:195px;">EC-Lab</td>
			<td style="width:197px;">BioLogic Science Instruments, France</td>
			<td style="width:125px;"></td>
			<td style="width:224px;">software for performing electrochemistry measurements</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:195px;">Origin 8.6</td>
			<td style="width:197px;">OriginLab</td>
			<td style="width:125px;"></td>
			<td style="width:224px;">data analysis software</td>
		</tr>
	</tbody>
</table>

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.

1. Chronoamperometry
From Chronoamperometric measurements two operational characteristics can be calculated: the maximum current density (jmax), i.e. the maximum electron flow per second, and the coulombic efficiency (CE), i.e. the total current flow in relation to the consumed substrate. Figure 4 shows a typical chronoamperometric biofilm growth curve of several growth cycles using a fed-batch reactor. After the initial lag-phase a first current density maximum commences. Then the current decreases again due to substrate depletion. At (nearly) zero current flow the medium is exchanged as indicated. After this substrate replenishment the current density increases again and the following maximum current density is higher than the first one. After several growth cycles jmax is constant &#8211; which indicates a steady-state biofilm formation14. As the steady state jmax value is depended on several parameters, e.g. electrode material, reactor temperature and biofilm composition, it is often considered as a characteristic of a certain electroactive microbial biofilm/ electrode system. (In case of flow-through reactors not jmax, but the constant steady-state current density is used.) The second operational characteristic is the coulombic efficiency (CE) (Equation&#160;1). That is the number of electrons recorded as electric current flow, Qp , per fed-batch cycle related to the number of the theoretical maximum number of electrons Qth, which is calculated from substrate consumption during this cycle.
<img alt="Equation 1" fo:content-width="1in" src="/files/ftp_upload/50800/50800eq1.jpg" width="100" />
(Equation&#160;1)
The total transferred charge can be gained by integration of the CA curve (see also Figure 6):
<img alt="Equation 2" fo:content-width="1in" src="/files/ftp_upload/50800/50800eq2.jpg" width="100" />
(Equation&#160;2)
The theoretically possible electric charge is calculated by Faraday&#8217;s law:
<img alt="Equation 3" fo:content-width="1in" src="/files/ftp_upload/50800/50800eq3.jpg" width="100" />
(Equation&#160;3)
Where V is the volume of the growth medium, F the Faraday constant (F&#160;= 96,485.34 C/mol), z the released number of electrons of the oxidized substrate (in case of acetate 8 electrons are released during oxidation to CO2) and &#916;c = c0-c1, which represents the substrate consumption (is determined by chemical analysis methods, e.g. by HPLC).
2. Cyclic voltammetry:
Cyclic voltammograms are recorded for nonturnover conditions and for turnover conditions (as indicated in Figure 4). In both cases different types of information can be gained. In the following it is discussed how possible and actual EET sites can be identified.
2.1 Nonturnover conditions &#8211; identifying possible EET sites: 
Figure 7 shows typical CVs for nonturnover conditions, i.e. in the absence of substrate. Figure 7A shows the CV of a biofilm for high scan rates (here 50 mV/sec) where only one peak pair and thus one formal potential Ef can be identified. In general the formal potential of a redox couple can be calculated from the peak potential of the oxidation peak EpA and the reduction peak of the respective species EpC by forming the arithmetic mean (see also Figure&#160;7C):
<img alt="Equation 4" fo:content-width="1.5in" src="/files/ftp_upload/50800/50800eq4.jpg" width="120" />
(Equation&#160;4)
When applying a low enough scan rate to the identical biofilm (here 1 mV/sec) up to four redox pairs can be identified (the respective formal potentials are indicated in Figure 7B). The reason is that the capacitive background current is decreased when using lower scan rates, see also Box 1 and e.g.17,21 for details. However, the nonturnover CV only shows all redox-active compounds at the electrode and thus the Ef of the possible EET sites.&#160; Only the analysis of the turnover CV provides the Ef of the actual EET transfer sites (see step 2.2.).
When further analyzing the nonturnover CVs different other characteristic parameters like peak separation &#916;Ep (Equation&#160;5) or maximum peak current ipA and minimum peak current ipC can be analyzed (see Figure 7C).
<img alt="Equation 5" fo:content-width="1.5in" src="/files/ftp_upload/50800/50800eq5.jpg" width="120" />
(Equation&#160;5)
These parameters, especially when recorded for different scan rates, can be used for mechanistic and kinetic analysis of electron transfer processes at electrodes. However, this kinetic analysis that is well established for chemical systems is not straightforward for electroactive microbial biofilms, see e.g.19,30

<ol>
	<li value="2.2">Turnover conditions &#8211; identifying actual EET sites:</li>
</ol>
When performing CV measurement in the presence of substrate a turnover CV is obtained (see Figure 8A). After plotting the data you can see the typical s-shaped, so-called bioelectrocatalytic, curve17-19. When further analyzing the data, the maximum of the first derivative, i.e. the inflection point of the CV-curve, provides the formal potential Ef of the actual EET site(s) (see Figure 8B). In the case of a young, i.e. thin, Geobacteraceae-dominated biofilm, here 168 h after starting CA, two inflection points can be observed. Subsequently the derivative shows two maxima corresponding to the formal potential of the bioelectrocatalytic active EET sites: -0.376 V and -0.295 V vs. Ag/AgCl (sat. KCl, 0.197 V vs. SHE). In Figure 7B these sites are denominated as Ef,2 and Ef,3, respectively. This finding also shows that the redox processes associated with the formal potentials Ef,1 and Ef,4 (in the nonturnover CV) are not related to bioelectrocatalysis. As the biofilm grows and gets thicker the CV shows only one inflection point, i.e. meaning one maximum in the derivative curve. This formal potential Ef, here being is -0.32 V vs. Ag/AgCl (sat KCl), is (roughly) equal to the arithmetic mean of the two formal potentials assigned to EET of the thin biofilm (e.g. here Ef,2 and Ef,3) &#8211; see Figure 9. This shows that this very fundamental CV analysis does provide insights into the EET thermodynamics in a fast and noninvasive way, but noteworthy does not allow drawing further mechanistic or kinetic conclusions.
Notably biofilms gained by the specific conditions in this protocol are highly dominated by Geobacter species (not demonstrated here)12,15. Changing selected parameters e.g. substrate, temperature, etc. will also influence the microbial composition and thus the electrochemical properties.
Table 1.&#160;Growth medium according to Kim et al.32
<table border="1" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Component</td>
			<td>(mg/L)&#160;/ (ml/L)&#160;&#160;&#160;</td>
		</tr>
		<tr>
			<td>NaH2PO4&#8226;H2O</td>
			<td>2,690.00</td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>4,330.00</td>
		</tr>
		<tr>
			<td>NH4Cl</td>
			<td>310.00</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>130.00</td>
		</tr>
		<tr>
			<td>sodium acetate</td>
			<td>820.00</td>
		</tr>
		<tr>
			<td>metal solution</td>
			<td>12.50</td>
		</tr>
		<tr>
			<td>vitamin solution</td>
			<td>12.50</td>
		</tr>
	</tbody>
</table>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Table 2. Components of the metal and vitamin solution33.</td>
			<td></td>
		</tr>
		<tr>
			<td>Trace element solution:</td>
			<td>Vitamin solution:</td>
		</tr>
		<tr>
			<td>Component</td>
			<td>mg/L&#160;&#160;&#160;</td>
			<td>Component</td>
			<td>mg/L&#160;&#160;&#160;</td>
		</tr>
		<tr>
			<td>Nitriloacetic acid</td>
			<td>1,500.0</td>
			<td>Biotin</td>
			<td>2.0</td>
		</tr>
		<tr>
			<td>MgSO4&#8226;7H2O&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;</td>
			<td>3,000.0</td>
			<td>Folic acid</td>
			<td>2.0</td>
		</tr>
		<tr>
			<td>MnSO4&#8226;2H2O&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;</td>
			<td>500.0</td>
			<td>Pyridoxine hydrochloride</td>
			<td>10.0</td>
		</tr>
		<tr>
			<td>NaCl&#160;&#160;</td>
			<td>1,000.0</td>
			<td>Thiamine hydrochloride</td>
			<td>5.0</td>
		</tr>
		<tr>
			<td>FeSO4&#8226;7H2O&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;</td>
			<td>100.0</td>
			<td>Riboflavin</td>
			<td>5.0</td>
		</tr>
		<tr>
			<td>CoSO4 or CoCl2&#160;&#160;&#160;&#160;&#160;</td>
			<td>100.0</td>
			<td>Nicotinic acid</td>
			<td>5.0</td>
		</tr>
		<tr>
			<td>CaCl2&#8226;2H2O&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;</td>
			<td>100.0</td>
			<td>DL-Calcium pantothonate</td>
			<td>5.0</td>
		</tr>
		<tr>
			<td>ZnSO4&#160;&#160;&#160;</td>
			<td>130.0</td>
			<td>Vitamin B12</td>
			<td>0.1</td>
		</tr>
		<tr>
			<td>CuSO4&#8226;H2O</td>
			<td>10.0</td>
			<td>p-Aminobenzoic acid</td>
			<td>5.0</td>
		</tr>
		<tr>
			<td>AlK(SO4)2 &#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;</td>
			<td>10.0</td>
			<td>Lipoic acid</td>
			<td>5.0</td>
		</tr>
		<tr>
			<td>H3BO3&#160;</td>
			<td>10.0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Na2MoO4&#8226;2H2O&#160;&#160;&#160;&#160;&#160;</td>
			<td>10.0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Note: Dissolve nitriloacetic acid with KOH to pH 6.5, and then proceed to add minerals.</td>
		</tr>
	</tbody>
</table>
 
<img alt="Box 1" fo:content-width="5in" src="/files/ftp_upload/50800/Box1.jpg" width="600" /> 
<img alt="Box 2" fo:content-width="5in" src="/files/ftp_upload/50800/Box2.jpg" width="600" />

<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig1highres.jpg" src="/files/ftp_upload/50800/50800fig1.jpg" /> 
Figure 1. Scheme of possible microbial extracellular electron transfer mechanisms. <a href="https://www.jove.com/files/ftp_upload/50800/50800fig1highres.jpg" target="_blank">Click here to view larger image.</a>




<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig2highres.jpg" src="/files/ftp_upload/50800/50800fig2.jpg" /> 
Figure 2. Sampling at the waste water treatment plant after the grid collector (waste water treatment plant Germany).&#160;<a href="https://www.jove.com/files/ftp_upload/50800/50800fig2highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig3highres.jpg" src="/files/ftp_upload/50800/50800fig3.jpg" /> 
Figure 3. Three electrode setup.&#160;(A) Scheme, (B) real reactor; CE = counter electrode, RE = reference electrode, WE = working electrode with electroactive microbial biofilm, SP = sampling port. <a href="https://www.jove.com/files/ftp_upload/50800/50800fig3highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig4highres.jpg" src="/files/ftp_upload/50800/50800fig4.jpg" /> 
Figure 4. Plot of a chronoamperometric fed-batch waste water inoculum based biofilm growth. The oxidative current reaches a steady-state after three cycles of medium change. Indicated are the lag-phase and the points of time of medium change and conduction of turnover cyclic voltammetry (CV) and nonturnover CV for steady-state biofilm conditions (raw data from&#160;Liu14). <a href="https://www.jove.com/files/ftp_upload/50800/50800fig4highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 5" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig5highres.jpg" src="/files/ftp_upload/50800/50800fig5.jpg" /> 
Figure 5. Principle potential-time course during one cycle of cyclic voltammetry. CV is starting at the initial potential Ei. The applied voltage is then changed linearly with time with a scan rate of v. At a chosen vertex potential E1 the voltage returns to an end potential E2, here E2 = Ei. <a href="https://www.jove.com/files/ftp_upload/50800/50800fig5highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 6" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig6highres.jpg" src="/files/ftp_upload/50800/50800fig6.jpg" /> 
Figure 6. Typical chronoamperogram of one growth cycle of a waste water derived biofilm; herein the maximum current density jmax, the flown (practical) electric charge Qp, the sampling of the initial substrate concentration c0 and the substrate concentration after one growth cycle c1 are indicated. <a href="https://www.jove.com/files/ftp_upload/50800/50800fig6highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 7" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig7highres.jpg" src="/files/ftp_upload/50800/50800fig7.jpg" /> 
Figure 7. Nonturnover cyclic voltammograms of a G. sulfurreducens biofilm. Scan rate: (A) 50 mV/sec, (B) 1 mV/sec&#160;23; (C) cutout of B, demonstrating the peak current density (jpA) and the respective peak potential (EpA) of the anode at the maxima. Correspondingly at the minima the peak current density (jpC) and the respective peak potential (EpC) of the cathode are shown. The formal potential Ef and the peak separation &#916;Ep result from Equation&#160;4 and Equation&#160;5, respectively (for details see text below). <a href="https://www.jove.com/files/ftp_upload/50800/50800fig7highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 8" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig8highres.jpg" src="/files/ftp_upload/50800/50800fig8.jpg" /> 
Figure 8. (A) Turnover cyclic voltammogram of a metabolizing G. sulfurreducens biofilm. The scan rate was 5 mV/sec. (B) First derivatives of the voltammetric curve23. Indicated are the formal potentials of the actual EET sites: Ef,2 and Ef,3. <a href="https://www.jove.com/files/ftp_upload/50800/50800fig8highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 9" fo:content-width="5in" fo:src="/files/ftp_upload/50800/50800fig9highres.jpg" src="/files/ftp_upload/50800/50800fig9.jpg" /> 
Figure 9. (A) Turnover cyclic voltammogram and (B) first derivatives of a thicker metabolizing G. sulfurreducens biofilm.&#160;The scan rate was 5&#160;mV/sec&#160;(compare Figure 8). <a href="https://www.jove.com/files/ftp_upload/50800/50800fig9highres.jpg" target="_blank">Click here to view larger image.</a>

Watch this Scientific Journal Video about Waste Water Derived Electroactive Microbial Biofilms: Growth, Maintenance, and Basic Characterization at JoVE.com

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

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 ...

waste-water-derived-electroactive-microbial-biofilms-growth

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JoVE Journal

Environment

13.2K Views.  UFZ - Helmholtz-Centre for Environmental Research. 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.

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

Methods for Facilitating Microbial Growth on Pulp Mill Waste Streams and Characterization of the Biodegradation Potential of Cultured Microbes

The kraft process is applied to wood chips for separation of lignin from the polysaccharides within lignocellulose for pulp that will produce a high quality paper. Black liquor is a pulping waste generated by the kraft process that has potential for downstream bioconversion. However, the recalcitrant nature of the lignocellulose resources, its chemical derivatives that constitute the majority of available organic carbon within black liquor, and its basic pH present challenges to microbial biodegradation of this waste material. Methods for the collection and modification of black liquor for microbial growth are aimed at utilization of this pulp waste to convert the lignin, organic acids, and polysaccharide degradation byproducts into valuable chemicals. The lignocellulose extraction techniques presented provide a reproducible method for preparation of lignocellulose growth substrates for understanding metabolic capacities of cultured microorganisms. Use of gas chromatography-mass spectrometry enables the identification and quantification of the fermentation products resulting from the growth of microorganisms on pulping waste. These methods when used together can facilitate the determination of the metabolic activity of microorganisms with potential to produce fermentation products that would provide greater value to the pulping system and reduce effluent waste, thereby increasing potential paper milling profits and offering additional uses for black liquor.

Industrial wastes can be collected and modified to analyze microbial growth. Lignocellulose extraction techniques provide components to analyze specific biodegradation ability. Gas chromatography-mass spectrometry identifies fermentation products of microorganisms grown on pulping waste. These methods determine the metabolic capacity of microorganisms to degrade pulping waste.

The kraft process is applied to wood chips for separation of lignin from the polysaccharides within lignocellulose for pulp that will produce a high ...

Laboratory-determined Phosphorus Flux from Lake Sediments as a Measure of Internal Phosphorus Loading

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) loading from lake sediments can account for a substantial portion of the total P load in eutrophic, and some mesotrophic, lakes. Laboratory determination of P release rates from sediment cores is one approach for determining the role of internal P loading and guiding management decisions. Two principal alternatives to experimental determination of sediment P release exist for estimating internal load: in situ measurements of changes in hypolimnetic P over time and P mass balance. The experimental approach using laboratory-based sediment incubations to quantify internal P load is a direct method, making it a valuable tool for lake management and restoration.
Laboratory incubations of sediment cores can help determine the relative importance of internal vs. external P loads, as well as be used to answer a variety of lake management and research questions. We illustrate the use of sediment core incubations to assess the effectiveness of an aluminum sulfate (alum) treatment for reducing sediment P release. Other research questions that can be investigated using this approach include the effects of sediment resuspension and bioturbation on P release.
The approach also has limitations. Assumptions must be made with respect to: extrapolating results from sediment cores to the entire lake; deciding over what time periods to measure nutrient release; and addressing possible core tube artifacts. A comprehensive dissolved oxygen monitoring strategy to assess temporal and spatial redox status in the lake provides greater confidence in annual P loads estimated from sediment core incubations.

Lake eutrophication is a water quality issue worldwide, making the need to identify and control nutrient sources critical. Laboratory determination of phosphorus release rates from sediment cores is a valuable approach for determining the role of internal phosphorus loading and guiding management decisions.

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) ...

EPA Method 1615. Measurement of Enterovirus and Norovirus Occurrence in Water by Culture and RT-qPCR. I. Collection of Virus Samples

EPA Method 1615 was developed with a goal of providing a standard method for measuring enteroviruses and noroviruses in environmental and drinking waters. The standardized sampling component of the method concentrates viruses that may be present in water by passage of a minimum specified volume of water through an electropositive cartridge filter. The minimum specified volumes for surface and finished/ground water are 300 L and 1,500 L, respectively. A major method limitation is the tendency for the filters to clog before meeting the sample volume requirement. Studies using two different, but equivalent, cartridge filter options showed that filter clogging was a problem with 10% of the samples with one of the filter types compared to 6% with the other filter type. Clogging tends to increase with turbidity, but cannot be predicted based on turbidity measurements only. From a cost standpoint one of the filter options is preferable over the other, but the water quality and experience with the water system to be sampled should be taken into consideration in making filter selections.

EPA Method 1615 uses an electropositive filter to concentrate enteroviruses and noroviruses in environmental and drinking waters. This manuscript describes the procedure for collecting samples for Method 1615 analyses.

EPA Method 1615 was developed with a goal of providing a standard method for measuring enteroviruses and noroviruses in environmental and drinking waters. ...

EPA Method 1615. Measurement of Enterovirus and Norovirus Occurrence in Water by Culture and RT-qPCR. Part III. Virus Detection by RT-qPCR

EPA Method 1615 measures enteroviruses and noroviruses present in environmental and drinking waters. This method was developed with the goal of having a standardized method for use in multiple analytical laboratories during monitoring period 3 of the Unregulated Contaminant Monitoring Rule. Herein we present the protocol for extraction of viral ribonucleic acid (RNA) from water sample concentrates and for quantitatively measuring enterovirus and norovirus concentrations using reverse transcription-quantitative PCR (RT-qPCR). Virus concentrations for the molecular assay are calculated in terms of genomic copies of viral RNA per liter based upon a standard curve. The method uses a number of quality controls to increase data quality and to reduce interlaboratory and intralaboratory variation. The method has been evaluated by examining virus recovery from ground and reagent grade waters seeded with poliovirus type 3 and murine norovirus as a surrogate for human noroviruses. Mean poliovirus recoveries were 20% in groundwaters and 44% in reagent grade water. Mean murine norovirus recoveries with the RT-qPCR assay were 30% in groundwaters and 4% in reagent grade water.

Here we present a procedure to quantify enterovirus and norovirus in environmental and drinking waters using reverse transcription-quantitative PCR. Mean virus recovery from groundwater with this standardized procedure from EPA Method 1615 was 20% for poliovirus and 30% for murine norovirus.

EPA Method 1615 measures enteroviruses and noroviruses present in environmental and drinking waters. This method was developed with the goal of having a ...

Characterization and Application of Passive Samplers for Monitoring of Pesticides in Water

Five different water passive samplers were calibrated under laboratory conditions for measurement of 124 legacy and current used pesticides. This study provides a protocol for the passive sampler preparation, calibration, extraction method and instrumental analysis. Sampling rates (RS) and passive sampler-water partition coefficients (KPW) were calculated for silicone rubber, polar organic chemical integrative sampler POCIS-A, POCIS-B, SDB-RPS and C18 disk. The uptake of the selected compounds depended on their physicochemical properties, i.e., silicone rubber showed a better uptake for more hydrophobic compounds (log octanol-water partition coefficient (KOW) &#62; 5.3), whereas POCIS-A, POCIS-B and SDB-RPS disk were more suitable for hydrophilic compounds (log KOW &#60; 0.70).

A protocol about the characterization and application of five different passive sampling devices is presented.

Five different water passive samplers were calibrated under laboratory conditions for measurement of 124 legacy and current used pesticides. This study ...

A Simple Planting Technique for Re-establishing Trees Where Frequent Inundation Occurs

Many of the Everglades tree islands have lost elevation over the past century and most of their trees have died such that they are now covered with herbaceous plants. This protocol describes a simple, cost-effective tree planting technique needed for restoring degraded Everglade tree islands. The design is patterned after a natural Everglades process that creates floating peat islands, which allows tree survival and growth in flooded conditions and often leads to the development of tree islands. Commercially available peat bags were used as the medium for the growth and establishment of potted native tree saplings. The pop-up configuration floated initially and provided additional elevation to minimize inundation, with a single native tree species sapling and a single tree fertilizer spike. During a 3 year study involving 105 pop-ups, most plants survived (80%) and many thrived. Determining whether this technique can establish trees on a degraded tree island will require longer studies and extensive field tests.

This protocol describes a simple, cost-effective tree planting technique for restoring degraded Everglades tree islands that experience inundation. The design creates an island (pop-up) which floats initially and adds elevation to promote tree survival and growth under flooded conditions.

Many of the Everglades tree islands have lost elevation over the past century and most of their trees have died such that they are now covered with ...

Improving Infrared Spectroscopy Characterization of Soil Organic Matter with Spectral Subtractions

Soil organic matter (SOM) underlies numerous soil processes and functions. Fourier transform infrared (FTIR) spectroscopy detects infrared-active organic bonds that constitute the organic component of soils. However, the relatively low organic matter content of soils (commonly &#60; 5% by mass) and absorbance overlap of mineral and organic functional groups in the mid-infrared (MIR) region (4,000-400 cm-1) engenders substantial interference by dominant mineral absorbances, challenging or even preventing interpretation of spectra for SOM characterization. Spectral subtractions, a post-hoc mathematical treatment of spectra, can reduce mineral interference and enhance resolution of spectral regions corresponding to organic functional groups by mathematically removing mineral absorbances. This requires a mineral-enriched reference spectrum, which can be empirically obtained for a given soil sample by removing SOM. The mineral-enriched reference spectrum is subtracted from the original (untreated) spectrum of the soil sample to produce a spectrum representing SOM absorbances. Common SOM removal methods include high-temperature combustion (&#39;ashing&#39;) and chemical oxidation. Selection of the SOM removal method carries two considerations: (1) the amount of SOM removed, and (2) absorbance artifacts in the mineral reference spectrum and thus the resulting subtraction spectrum. These potential issues can, and should, be identified and quantified in order to avoid fallacious or biased interpretations of spectra for organic functional group composition of SOM. Following SOM removal, the resulting mineral-enriched sample is used to collect a mineral reference spectrum. Several strategies exist to perform subtractions depending on the experimental goals and sample characteristics, most notably the determination of the subtraction factor. The resulting subtraction spectrum requires careful interpretation based on the aforementioned methodology. For many soil and other environmental samples containing substantial mineral components, subtractions have strong potential to improve FTIR spectroscopic characterization of organic matter composition.

SOM underlies many soil functions and processes, but its characterization by FTIR spectroscopy is often challenged by mineral interferences. The described method can increase the utility of SOM analysis by FTIR spectroscopy by subtracting mineral interferences in soil spectra using empirically obtained mineral reference spectra.

Soil organic matter (SOM) underlies numerous soil processes and functions. Fourier transform infrared (FTIR) spectroscopy detects infrared-active organic ...

Optimized Procedure for Determining the Adsorption of Phosphonates onto Granular Ferric Hydroxide using a Miniaturized Phosphorus Determination Method

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric hydroxide (GFH), with little effort and high reliability. The phosphonate, e.g., nitrilotrimethylphosphonic acid (NTMP), is brought into contact with the GFH in a rotator in a solution buffered by an organic acid (e.g., acetic acid) or Good buffer (e.g., 2-(N-morpholino)ethanesulfonic acid) [MES] and N-cyclohexyl-2-hydroxyl-3-aminopropanesulfonic acid [CAPSO]) in a concentration of 10 mM for a specific time in 50 mL centrifuge tubes. Subsequently, after membrane filtration (0.45 &#181;m pore size), the total phosphorus (total P) concentration is measured using a specifically developed determination method (ISOmini). This method is a modification and simplification of the ISO 6878 method: a 4 mL sample is mixed with H2SO4 and K2S2O8 in a screw cap vial, heated to 148-150 &#176;C for 1 h and then mixed with NaOH, ascorbic acid and acidified molybdate with antimony(III) (final volume of 10 mL) to produce a blue complex. The color intensity, which is linearly proportional to the phosphorus concentration, is measured spectrophotometrically (880 nm). It is demonstrated that the buffer concentration used has no significant effect on the adsorption of phosphonate between pH 4 and 12. The buffers, therefore, do not compete with the phosphonate for adsorption sites. Furthermore, the relatively high concentration of the buffer requires a higher dosage concentration of oxidizing agent (K2S2O8) for digestion than that specified in ISO 6878, which, together with the NaOH dosage, is matched to each buffer. Despite the simplification, the ISOmini method does not lose any of its accuracy compared to the standardized method.

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric hydroxide, with little effort and high reliability. In a buffered solution, the phosphonate is brought into contact with the adsorbent using a rotator and then analyzed via a miniaturized phosphorus determination method.

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric ...

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 ...

Construction of a Low-cost Mobile Incubator for Field and Laboratory Use

Incubators are essential for a range of culture-based microbial methods, such as membrane filtration followed by cultivation for assessing drinking water quality. However, commercially available incubators are often costly, difficult to transport, not flexible in terms of volume, and/or poorly adapted to local field conditions where access to electricity is unreliable. The purpose of this study was to develop an adaptable, low-cost and transportable incubator that can be constructed using readily available components. The electronic core of the incubator was first developed. These components were then tested under a range of ambient temperature conditions (3.5 &#176;C - 39 &#176;C) using three types of incubator shells (polystyrene foam box, hard cooler box, and cardboard box covered with a survival blanket). The electronic core showed comparable performance to a standard laboratory incubator in terms of the time required to reach the set temperature, inner temperature stability and spatial dispersion, power consumption, and microbial growth. The incubator set-ups were also effective at moderate and low ambient temperatures (between 3.5 &#176;C and 27 &#176;C), and at high temperatures (39 &#176;C) when the incubator set temperature was higher. This incubator prototype is low-cost (&lt; 300 USD) and adaptable to a variety of materials and volumes. Its demountable structure makes it easy to transport. It can be used in both established laboratories with grid power or in remote settings powered by solar energy or a car battery. It is particularly useful as an equipment option for field laboratories in areas with limited access to resources for water quality monitoring.

This paper describes a method for building an adaptable, low-cost and transportable incubator for microbial testing of drinking water. Our design is based on widely available materials and can operate under a range of field conditions, while still offering the advantages of higher-end laboratory-based models.

Incubators are essential for a range of culture-based microbial methods, such as membrane filtration followed by cultivation for assessing drinking water ...