R.K. and F.H. are grateful for the EU-commision for financial support within the Marie-Sklodowska ITN Biocascades (Nr. 634200).

Caution: Please consult all relevant material safety data sheets (MSDS) before use. Several of the chemicals used in these syntheses are acutely toxic and carcinogenic. All steps involving harmful organic solvents, particularly the extraction of the reaction products and the derivatization of fatty acids must be carried out under a fume hood using personal protective equipment (safety glasses, gloves, lab coat, full-length pants, closed-toe shoes). All operations involving genetically modified organisms in this work require installations that are approved for handling of genetically modified organisms of the safety level S1.
1. Expression of the Recombinant Decarboxylase OleTJE in E. coli
<ol>
	<li>Preparation of the production strain
	<ol>
		<li>Prepare a synthetic gene of OleTJE from Jeotgalicoccus and cloned into the expression vector pASK-IBA37plus as described.8 
		NOTE: To avoid degradation of fatty acids by enzymes from the bacterial metabolism, E. coli strain JW5020 from the Keio collection with a dysfunctional pathway for fatty acid degradation was used.</li>
	</ol>
	</li>
	<li>Cultivation and induction of enzyme expression
	<ol>
		<li>Prepare a pre-culture by inoculating the E. coli JW5020 OleT mutant from an LB-plate (containing 100 &#181;g ml-1 ampicillin) in two 3 ml LB-medium in test tubes. Pipette ampicillin (100 &#181;g ml-1) from a stock solution into the medium for selection and incubate at 37 &#176;C for 15 hr in a shaking incubator at 180 rpm.</li>
		<li>Add 2 ml of the pre-cultures to two 200 ml LB-medium, containing ampicillin (100 &#181;g ml-1), in 1 L shake flasks. Incubation proceeds at 37 &#176;C and 180 rpm.</li>
		<li>Monitor the change in OD600nm after the inoculation. When the OD600 reaches a value between 0.6 and 0.8 induce the expression by adding tetracycline (0.2 &#181;g ml-1). Incubate the cultures for 15 hr at 20 &#176;C and 180 rpm. 
		NOTE: Since OleTJE contains a heme cofactor, addition of &#948;-Amino levulinic acid (0.5 mM) prior to induction is needed to provide the culture with a precursor for cofactor synthesis.</li>
	</ol>
	</li>
	<li>Cell lysis
	<ol>
		<li>Carry out the following steps on ice. Transfer the cultures by decanting or pipetting into centrifuge tubes and use a scale to ensure equal distribution. Centrifuge the cultures for 20 min at 12,000 x g at 4 &#176;C.</li>
		<li>Carefully discard the supernatant and resuspend the pellets in 50 ml buffer (Tris 50 mM, NaCl 200 mM, pH 7.5) by pipetting and transfer the suspension in a conical centrifuge tube.</li>
		<li>After centrifugation for 15 min at 4,000 x g at 4 &#176;C discard the supernatant, resuspend each pellet in 3 ml buffer by pipetting.</li>
		<li>Perform cell lysis by sonicating the solutions on ice (three cycles x 30 sec) leaving a 1 min pause in between the cycles. Centrifuge the solutions for 20 min at 15,000 x g at 4 &#176;C to remove cell debris and transfer the supernatant into a conical centrifugation tube without disturbing the pellet by pipetting. Discard the pellet. 
		NOTE: One solvent fraction will be used directly for biocatalysis after small molecules are removed. The second fraction will be used for purification of the His-tagged OleTJE.</li>
	</ol>
	</li>
	<li>Removal of small molecules of cell-extracts
	<ol>
		<li>Before using the crude extract in biocatalysis remove small molecules which might interfere with hydrogen peroxide formation or electron transfer by pipetting 3 ml of cell-free extract into a centrifuge filter unit with a 10 kDa membrane and centrifuge at 4,000 x g at 4 &#176;C. Resuspend the remaining protein in 3 ml Tris-HCl-buffer.</li>
	</ol>
	</li>
	<li>Purification of OleT
	<ol>
		<li>Apply 3 ml of the second cell extract to His Pur Ni-NTA spin columns, which have been equilibrated before with equilibration buffer (Tris 20 mM, NaCl 300 mM, imidazole 10 mM).</li>
		<li>Seal the columns with the bottom plugs and upper screw-caps and shake them lightly until the resin is distributed equally with the cell-free extract. Incubate the loaded columns for 30 min in an overhead shaker at 4 &#176;C.</li>
		<li>Wash the columns with 1 ml of washing buffer (Tris 20 mM, NaCl 300 mM, imidazole 20 mM, pH 7.4) by centrifuging at 700 x g for 2 min. Repeat this step twice.</li>
		<li>Place the columns into a fresh conical centrifugation tube and add 1 ml of elution buffer (Tris 20 mM, NaCl 300 mM, imidazole 250 mM, pH 7.4) OleTJE . Centrifuge at 700 x g for 2 min and repeat this step two times. 
		NOTE: Imidazole will bind to nickel and thus remove OleTJE from the column.</li>
		<li>Transfer the elution to a centrifuge filter unit (10 kDa membrane) and centrifuge at 4,000 x g at 4 &#176;C to remove imidazole.</li>
		<li>Check the purity of the enzyme by an SDS-Page (15%).9 Mix 3 &#181;l of the elution with a SDS-buffer (final concentration 1x) and incubate the solution at 95 &#176;C for 5 min for denaturation. Subsequently apply the total amount on a SDS-Page. Run the gel at 35 mA using a commercial protein standard. Detect the protein at 50 kDa.</li>
		<li>To determine the protein concentration use a commercial available BSA-kit. Pipette 50 &#181;l of diluted protein samples (1:100, 1:200, 1:500) and BSA standard (0, 20, 30, 40,50, 60, 80, 100 mg ml-1) in a 96-well plate. Add 200 &#181;l of Bradford reagent, measure the absorbance at 595 nm after 15 min with a fluorimeter and calculate the concentrations using the standard curve.</li>
	</ol>
	</li>
</ol>
2. Light-catalyzed Biotransformation
<ol>
	<li>Biocatalytic reactions without hydrogen peroxide addition
	<ol>
		<li>Prepare a 10 ml stock solution of 10 mM stearic acid (Mr 284.5 g mol-1) by adding 10%(v/v) tergitol and 0.0284 g stearic acid to distilled water. Heat the solution in a heating chamber to 60 &#176;C until the fatty acid is completely dissolved.</li>
		<li>Biocatalysis and sampling
		<ol>
			<li>Prepare two 2 ml reaction mixtures containing 50 mM EDTA, 0.01 mM FMN, 0.5 mM stearic acid in Tris-HCl-buffer. Add a magnetic stirring bar for sufficient oxygen supply and place the glass tubes in a water bath, heated to 25 &#176;C.</li>
			<li>Add 200 &#181;g ml-1 of purified enzyme or 10% (v/v) cell-free extract to each of the reaction mixtures and illuminate them with a clear glass light bulb (120 W) in 15 cm distance of the reaction tubes.</li>
			<li>Take 200 &#181;l samples at specific time points (0, 10, 30, 60 and 120 min and over night) stopping the reaction with 20 &#181;l 37% HCl. Add 5 &#181;l myristic acid (10 mM stock solution) as internal standard.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Analysis of reaction products
	<ol>
		<li>For extraction add 500 &#181;l ethyl acetate to the samples twice, invert the tubes and centrifuge for 1 min at 13,000 x g.</li>
		<li>Take off 400 &#181;l of the supernatant and let the solvent completely evaporate.</li>
		<li>Derivatize the carboxylic acids into trimethylsilylcarboxylic acid by adding 200 &#181;l N-methyl-N-(trimethylsilyl)trifluoro acetamide (MSTFA). Incubate the solution at 60 &#176;C for 30 min in order to convert hydroxyl groups to trimethylsilyl ethers.</li>
		<li>Determination of conversion by GC/FID 
		NOTE: Determine the formation of 1-heptadecene (RT: 8.34 min), &#945;- and &#946;-hydroxy stearic acid (RT: 12.05 and 12.1 min) and decrease of the substrate (RT: 11.15 min) using GC/FID.
		<ol>
			<li>Set the temperature profile described below Set the injection temperature to 340 &#176;C. Hold the initial oven temperature is at 90 &#176;C for 3.5 min and subsequently increase with 45 &#176;C min-1. At 220 &#176;C, hold the temperature for 2 min. After a repeated increase of 45 &#176;C min-1 , hold the temperature at 280 &#176;C for 2 min and then increase with 60 &#176;C min-1. 
			NOTE: Maximum temperature of 330 &#176;C is held for 1.44 min.</li>
			<li>Inject 4 &#181;l of the sample, with a split ratio of 5 to get rapid volatilization and homogenous mixing with the carrier gas helium. 
			NOTE: In dependence on the increasing temperature the substrate and products will enter the gas phase. The substances are detected by the GC/FID detector after passing the column at specific retention times and are displayed as peaks on the screen. Observe the peak formation on the monitor.</li>
			<li>Calculate the concentration of formed product by the following formula: 
			<img alt="Equation 1" src="/files/ftp_upload/53439/53439eq1.jpg" /> 
			NOTE: The calibration factor has been determined specifically for this column for each substrate and product by calculating the standard curve from known amounts of the derivatized substances and their corresponding peak area.</li>
		</ol>
		</li>
		<li>Identify olefin and &#946;-hydroxy acid peaks by GC/MS.
		<ol>
			<li>Identify olefin and &#946;-hydroxy acid peaks by GC/MS. Set the temperature profile. Set the injection temperature to 250 &#176;C. Hold the oven temperature at 100 &#176;C for 5 min and subsequently increase with 20 &#176;C min-1. 
			NOTE: Maximum temperature is held for 7.5 min.</li>
			<li>Set the mass spectrometer detector temperature to 250 &#176;C and scan at 50 to 500 m/z in electron impact mode. 
			NOTE: Electron impact ionization removes a single electron resulting in the formation of a radical cation which will be passed forward for mass analysis. Due to the high intramolar energy covalent bonds within the molecule break creating fragments of lower m/z. These fragments form a fingerprint specific for a substance.</li>
			<li>Inject 1 &#181;l of the sample. Detect 1-Heptadecene after 11.4 min retention time by the corresponding fingerprint (Figure 2).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Kourist, R., Dom&#236;nguez de Mar&#236;a, P., Miyamoto, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biocatalytic+strategies+for+the+asymmetric+synthesis+of+profens+-+recent+trends+and+developments.">Biocatalytic strategies for the asymmetric synthesis of profens - recent trends and developments.</a> Green Chem. , 2607-2618 (2011).</li><li>Holtmann, D., Fraaije, M. W., Arends, I. W., Opperman, D. J., Hollmann, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+taming+of+oxygen:+biocatalytic+oxyfunctionalisations.">The taming of oxygen: biocatalytic oxyfunctionalisations.</a> Chemical Comm. 50, 13180-13200 (2014).</li><li>Churakova, E., 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=Specific+photobiocatalytic+oxyfunctionalization+reactions.">Specific photobiocatalytic oxyfunctionalization reactions.</a> Ang. Chem. In. Ed. 123, 10904-10907 (2011).</li><li>Hollmann, F., Arends, I., Buehler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biocatalytic+Redox+Reactions+for+Organic+Synthesis:+Nonconventional+Regeneration+Methods.">Biocatalytic Redox Reactions for Organic Synthesis: Nonconventional Regeneration Methods.</a> ChemCatChem. 2, 762-782 (2010).</li><li>Girhard, M., Kunigk, E., Tihovsky, S., Shumyantseva, V. V., Urlacher, V. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light-driven+biocatalysis+with+cytochrome+P450+peroxygenases.">Light-driven biocatalysis with cytochrome P450 peroxygenases.</a> Biotechnol. Appl. Biochem. 60, 111-118 (2013).</li><li>Bartsch, M., 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=Photosynthetic+production+of+enantioselective+biocatalysts.">Photosynthetic production of enantioselective biocatalysts.</a> Microb. Cell. Fact. 14, 53 (2015).</li><li>Rude, M. 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=Terminal+olefin+(1-alkene)+biosynthesis+by+a+novel+P450+fatty+acid+decarboxylase+from+Jeotgalicoccus+species.">Terminal olefin (1-alkene) biosynthesis by a novel P450 fatty acid decarboxylase from Jeotgalicoccus species.</a> Appl. Environ. Microbiol. 77, 1718-1727 (2011).</li><li>Zachos, I., 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=Photobiocatalytic+decarboxylation+for+olefin+synthesis.">Photobiocatalytic decarboxylation for olefin synthesis.</a> Chem. Comm. 51, 1918-1921 (2015).</li><li>Liu, Y., 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=Hydrogen+peroxide-independent+production+of+&#945;-alkenes+by+OleTJE+P450+fatty+acid+decarboxylase.">Hydrogen peroxide-independent production of &#945;-alkenes by OleTJE P450 fatty acid decarboxylase.</a> Biotechnol. Biofuels. 7, 28 (2014).</li><li>Perez, D. I., Grau, M. M., Arends, I. W., Hollmann, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visible+light-driven+and+chloroperoxidase-catalyzed+oxygenation+reactions.">Visible light-driven and chloroperoxidase-catalyzed oxygenation reactions.</a> Chem. Comm. 40 (44), 6848-6850 (2009).</li></ol>

The authors declare that they have no competing financial interests.

The light-driven generation of hydrogen peroxide can be applied for a range redox transformations, including peroxygenases3, chloroperoxidases10 and P450 monooxygenases5. It is an easy and practicable approach. In the long term, the use of visible light opens up the perspective to utilize sunlight for chemical transformations, which is a sustainable alternative for energy-rich reactions.
The method is applicable with purified enzyme or with cell-free extract. While the latter requires less cost and work, it should be noted that small molecules in the crude extract may interfere with the light catalyzed conversion. A practicable approach is to remove these small components with a micromembrane (i.e., by centrifugation in a Centrifugal filter unit or by dialysis). The concentration of the light-harvesting molecule FMN determines the concentration of the hydrogen peroxide. Depending on the affinity of the oxidoreductase, this concentration is decisive for the enzymatic activity. Another important factor is the concentration of the sacrificial electron donor EDTA. The most important parameter, however, is the operational stability and activity of the enzyme.
The olefinization of fatty acids is an elegant reaction for the conversion of bio-based fatty acids into olefins that belong to the major commodities for the chemical industry. The light-driven biocatalytic decarboxylation can be carried out at room temperature and at neutral pH, which offers clear advantages in terms of sustainability.
Our results show that in situ generation of hydrogen peroxide is a strategy to supply the cofactor without impairing enzyme stability, leading to a high conversion. Present methods for cofactor regeneration use agricultural products or petrol-based chemicals. Light-driven reactions are emerging as renewable alternative. Future research will be dedicated to methods for the substitution of the sacrificial reagent EDTA by cheaper molecules and to reduce the amount of the light-harvesting molecule FMN.

The climate change and the foreseeable depletion of renewable resources poses a serious threat to our society. In this context, enzyme catalysis represents a still not fully exploited potential for the development of sustainable and &#39;greener&#39; chemistry1. Oxidoreductases have the capacity to catalyze the introduction and modification of functional groups under mild reactions conditions and belong to the most important biocatalysts2. Most redox transformations require the supply of external of cofactors such as NAD(P)H. Methods for cofactor regeneration have been applied in industrial scale. However, they still result in high process costs, which limits their application mostly to high-value products. Interestingly, several peroxidases3,4 and P450 monooxygenases5 accept electrons from hydrogen peroxide via the so-called peroxide shunt. While H2O2 is a cheap co-reagent, it is reportedly harmful for many enzymes. A steady in situ formation of low concentrations of hydrogen peroxide is a viable approach to drive the reaction without impairing the operational stability of enzyme.
<img alt="Figure 1" src="/files/ftp_upload/53439/53439fig1.jpg" /> 
Figure 1. Experimental set-up of the photobiocatalytic decarboxylation of fatty acids by OleTJE. <a href="https://www.jove.com/files/ftp_upload/53439/53439fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The use of light as energy source for chemical and biological processes has been receiving increasing attention in the last years6. Light-driven generation of hydrogen peroxide has emerged as an easy and robust method to supply hydrogen peroxide for redox transformations (Figure 1). A photocatalyst such as flavin adenine mononucleotide (FMN) allows the reduction of molecular oxygen to hydrogen peroxide, which then is used as cofactor for the enzymatic oxyfunctionalization reaction. Possible electron donors are ethylenediaminetetraacetic acid (EDTA), ascorbate or the inexpensive formiate. The method is generally applicable for H2O2-dependent enzymes, including peroxidases3,4 and P450 monooxygenases5.
We have recently investigated the application of a novel bacterial decarboxylase7 for the transformation of natural fats into olefins8. This would be a sustainable route for the synthesis of widely used platform chemicals from a bio-based source. The decarboxylase OleTJE from the gram-positive bacterium Jeotgalicoccus sp. catalyzes the oxidative decarboxylation of fatty acids and forms 1-alkenes as products. OleTJE is closely related to bacterial P450 monooxygenases and needs electrons from hydrogen peroxide for the reaction.
Unfortunately, addition of H2O2 to a solution of substrate and enzyme resulted in low conversions and a poor reproducibility of the results, presumably due to a harmful effect of hydrogen peroxide on the stability of OleTJE. Generation of a fusion protein with the NADPH-reductase RhFred made an NADPH-dependent decarboxylation possible.9 Nevertheless, the high price of NADPH and the current limited possibilities for a cost-efficient regeneration prompted us to investigate cheaper electron donors. Inspired by the similarity of OleTJE with P450 monooxygenases, we used the light-catalyzed generation of H2O2. We were pleased to obtain high conversions (up to &#62;95%) using cell-free extracts or purified enzyme solutions.
With the example of fatty acid decarboxylation, we present a general protocol for light-driven enzymatic redox transformations using FMN as photocatalyst and hydrogen peroxide as cofactor. The presented methods include the production of the enzyme in recombinant cell of E. coli, purification of the enzyme, the application for the synthesis of 1-alkenes and the analysis of the reaction products.

<table><tbody><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Ampicillin</td><td>Sigma Aldrich</td><td>69-52-3</td><td></td></tr><tr><td>Bradford reagent</td><td>Roth</td><td>K015.1</td><td></td></tr><tr><td>BSA</td><td>Sigma Aldrich</td><td>90604-29-8</td><td></td></tr><tr><td>DMSO</td><td>Sigma Aldrich</td><td>67-68-5</td><td></td></tr><tr><td>Ethyl acetate</td><td>Fisher Chemical</td><td>141-78-6</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA)</td><td>Roth</td><td>8043.1</td><td></td></tr><tr><td>Riboflavin 5-monophosphate sodium salt hydrate</td><td>Sigma Aldrich</td><td>130-40-5</td><td></td></tr><tr><td>Hydrochlorid acid 37%</td><td>Sigma Aldrich</td><td>7647-01-0</td><td></td></tr><tr><td>Hydrogen peroxide 30%</td><td>Sigma Aldrich</td><td>7722-84-1</td><td></td></tr><tr><td>&amp;delta;-Amino levulinic acid</td><td>Sigma Aldrich</td><td>5451-09-2</td><td></td></tr><tr><td>&lt;em&gt;N&lt;/em&gt;-Methyl-&lt;em&gt;N&lt;/em&gt;-(Trimethylsilyl)trifluoro acetamide (MSTFA)</td><td>Sigma Aldrich</td><td>24589-78-4</td><td></td></tr><tr><td>Myristic acid &gt;99%</td><td>Sigma Aldrich</td><td>208-875-2&amp;nbsp;</td><td></td></tr><tr><td>Imidazole</td><td>Sigma Aldrich</td><td>288-32-4</td><td></td></tr><tr><td>Sodium chloride</td><td>Fisher Chemical</td><td>7647-14-5</td><td></td></tr><tr><td>Stearic acid &gt;99%</td><td>Sigma Aldrich</td><td>57-11-4</td><td></td></tr><tr><td>Tetracycline</td><td>Sigma Aldrich</td><td>60-54-8&amp;nbsp;</td><td></td></tr><tr><td>Tergitol</td><td>Sigma Aldrich</td><td>MFCD01779855&amp;nbsp;</td><td></td></tr><tr><td>Tris(hydroxymethyl)-aminomethan</td><td>Sigma Aldrich</td><td>77-86-1</td><td></td></tr><tr><td>&lt;strong&gt;Device&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Incubator shaker</td><td>G-25CK</td><td>New Brunswick Scientific</td><td></td></tr><tr><td></td><td>Ecotron</td><td>Infors HT</td><td></td></tr><tr><td>Centrifugation</td><td>Labofuge 400R</td><td>Heraeus</td><td></td></tr><tr><td></td><td>RC 5B Plus</td><td>Sorvall</td><td></td></tr><tr><td></td><td>Fresco 17</td><td>Thermo Scientific</td><td></td></tr><tr><td>Centrifugation rotors</td><td>SS34</td><td>Sorvall</td><td></td></tr><tr><td></td><td>SLA</td><td>Sorvall</td><td></td></tr><tr><td>Clean bench</td><td>Envirco</td><td>Ceag Schirp Reinraum technik</td><td></td></tr><tr><td>Column GC-FID&amp;nbsp;</td><td>CP-Sil 5CB (30 m x 0.25&amp;nbsp;mm x 0.25 &amp;micro;m)&amp;nbsp;</td><td>Agilent Technologies</td><td></td></tr><tr><td>Column GC-MS</td><td>FactorFour Capillary Coloumn (VF-5 ms + 5 m EZ Guard)&amp;nbsp;</td><td>Varian</td><td></td></tr><tr><td>GC-FID</td><td>GC-2010 plus</td><td>Shimadzu</td><td></td></tr><tr><td>GC-MS</td><td>IST-40</td><td>Varian</td><td></td></tr><tr><td>Magnetic stirrer</td><td>RCT classic</td><td>IKA</td><td></td></tr><tr><td>pH meter</td><td>SevenEasy</td><td>Mettler toledo</td><td></td></tr><tr><td>Sonicator</td><td>Branson Sonifier 250</td><td>Branson</td><td></td></tr><tr><td>Spectral photometer</td><td>FLUOstar Omega</td><td>BMG Labtech</td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Affinity chromatography column</td><td>His Pur Ni-NTA spin column</td><td>Thermo Scientific</td><td></td></tr><tr><td>Centricon</td><td>Vivaspin turbo 15</td><td>VWR International</td><td></td></tr><tr><td>Microtiter plates</td><td>96 Well Multiply&amp;reg;PCR Plates</td><td>Sarstedt</td><td></td></tr></tbody></table>

light-driven enzymatic decarboxylation

Oxidoreductases belong to the most-applied industrial enzymes. Nevertheless, they need external electrons whose supply is often costly and challenging. Recycling of the electron donors NADH or NADPH requires the use of additional enzymes and sacrificial substrates. Interestingly, several oxidoreductases accept hydrogen peroxide as electron donor. While being inexpensive, this reagent often reduces the stability of enzymes. A solution to this problem is the in situ generation of the cofactor. The continuous supply of the cofactor at low concentration drives the reaction without impairing enzyme stability. This paper demonstrates a method for the light-catalyzed in situ generation of hydrogen peroxide with the example of the heme-dependent fatty acid decarboxylase OleTJE. The fatty acid decarboxylase OleTJE was discovered due to its unique ability to produce long-chain 1-alkenes from fatty acids, a hitherto unknown enzymatic reaction. 1-alkenes are widely used additives for plasticizers and lubricants. OleTJE has been shown to accept electrons from hydrogen peroxide for the oxidative decarboxylation. While addition of hydrogen peroxide damages the enzyme and results in low yields, in situ generation of the cofactor circumvents this problem. The photobiocatalytic system shows clear advantages regarding enzyme activity and yield, resulting in a simple and efficient system for fatty acid decarboxylation.

While the addition of hydrogen peroxide to the reaction mixture resulted in low to moderate conversions (&#60;10%), in situ generation of hydrogen peroxide increased the conversion up to 80% conversion. Analysis by GC/MS shows the formation of olefins from fatty acids (Figure 2).
<img alt="Figure 2" src="/files/ftp_upload/53439/53439fig2.jpg" /> 
Figure 2. (A) Temperature profile for GC/MS measurements. (B) Fingerprint of the 1-heptadecene eluting after 11.4 min. (C) Characteristic secondary CnH2n-1 fragment ions of terminal olefins exemplary shown for 1-heptadecene. <a href="https://www.jove.com/files/ftp_upload/53439/53439fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The light-catalyzed reaction achieved high conversions with cell-free extracts of E. coli. A purified enzyme solution showed a clear correlation between enzyme concentration and conversion. Increasing the concentration of the light-harvesting molecule FMN lead to higher conversions. However, increasing the concentration above 10 mM did not further accelerate the reactions, indicating that the amount of hydrogen peroxide is sufficiently available and no longer the limiting factor.
In addition to the decarboxylation of fatty acids, OleTJE also catalyzes a hydroxylation in &#946;-position. In the conversion of stearic acid, the decarboxylation is about three times faster than the hydroxylation. In a typical experiment using a solution of 0.5 mM stearic acid and 10 &#181;M FMN, 99% of the substrate were converted to a mixture of 1-heptadecene and 2-hydroxystearic acid with a ratio of 3.3:1 (Figure 3)8. An investigation of the substrate spectrum of the light-driven biocatalysis showed that for fatty acids with longer acyl chains, OleTJE preferentially catalyzes the decarboxylation, while the relative amount of hydroxyl-fatty acids in the product mixture increased with shorter chain length. Fatty acids shorter than myristic acid (C14) did not undergo decarboxylation.
<img alt="Figure 3" src="/files/ftp_upload/53439/53439fig3.jpg" /> 
Figure 3. Gas chromatographic analysis of OleTJE-mediated decarboxylation of stearic acid (11.15 min) into 1-heptadecene (8.4 min), &#945;-hydroxy stearic acid (12.04 min) &#946;-hydroxy stearic acid (12.1 min). The change of the peak areas from samples taken over a time-course of 2 hr is shown. <a href="https://www.jove.com/files/ftp_upload/53439/53439fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Surprisingly, the unsaturated acids oleic acid (C18:1d9 cis) and linoleic acid (C18:2d9d12) were not accepted as substrates, indicating that the twisted configuration of cis double bonds cannot be accommodated in a productive binding mode in the enzyme. Addition of oleic acid (10%) also prevented the conversion of stearic acid. Interestingly, stearic acid (C18:1d9 trans) was converted by OleTJE, yet with slightly lower activity then stearic acid.

Watch this Scientific Journal Video about Light-driven Enzymatic Decarboxylation at JoVE.com

Light-driven Enzymatic Decarboxylation

We describe a protocol for the light-catalyzed generation of hydrogen peroxide &#8212; a cofactor for oxidative transformations.

Oxidoreductases belong to the most-applied industrial enzymes. Nevertheless, they need external electrons whose supply is often costly and challenging. ...

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Carboxylic Acids

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Video: Light-driven Enzymatic Decarboxylation

A Toolkit to Enable Hydrocarbon Conversion in Aqueous Environments

This work puts forward a toolkit that enables the conversion of alkanes by Escherichia coli and presents a proof of principle of its applicability. The toolkit consists of multiple standard interchangeable parts (BioBricks)9 addressing the conversion of alkanes, regulation of gene expression and survival in toxic hydrocarbon-rich environments.
A three-step pathway for alkane degradation was implemented in E. coli to enable the conversion of medium- and long-chain alkanes to their respective alkanols, alkanals and ultimately alkanoic-acids. The latter were metabolized via the native &beta;-oxidation pathway. To facilitate the oxidation of medium-chain alkanes (C5-C13) and cycloalkanes (C5-C8), four genes (alkB2, rubA3, rubA4and rubB) of the alkane hydroxylase system from Gordonia sp. TF68,21 were transformed into E. coli. For the conversion of long-chain alkanes (C15-C36), theladA gene from Geobacillus thermodenitrificans was implemented. For the required further steps of the degradation process, ADH and ALDH (originating from G. thermodenitrificans) were introduced10,11. The activity was measured by resting cell assays. For each oxidative step, enzyme activity was observed.
To optimize the process efficiency, the expression was only induced under low glucose conditions: a substrate-regulated promoter, pCaiF, was used. pCaiF is present in E. coli K12 and regulates the expression of the genes involved in the degradation of non-glucose carbon sources.
The last part of the toolkit - targeting survival - was implemented using solvent tolerance genes, PhPFD&alpha; and &beta;, both from Pyrococcus horikoshii OT3. Organic solvents can induce cell stress and decreased survivability by negatively affecting protein folding. As chaperones, PhPFD&alpha; and &beta; improve the protein folding process e.g. under the presence of alkanes. The expression of these genes led to an improved hydrocarbon tolerance shown by an increased growth rate (up to 50%) in the presences of 10% n-hexane in the culture medium were observed.
Summarizing, the results indicate that the toolkit enables E. coli to convert and tolerate hydrocarbons in aqueous environments. As such, it represents an initial step towards a sustainable solution for oil-remediation using a synthetic biology approach.

A sustainable auto regulating bacterial system for the remediation of oil pollutions was designed using standard interchangeable DNA parts (BioBricks). An engineered E. coli strain was used to degrade alkanes via &beta;-oxidation in toxic aqueous environments. The respective enzymes from different species showed alkane degradation activity. Additionally, an increased tolerance to n-hexane was achieved by introducing genes from alkane-tolerant bacteria.

This work puts forward a toolkit that enables the conversion of alkanes by Escherichia coli and presents a proof of principle of its applicability. The ...

Bioengineering

Mapping Metabolism: Monitoring Lactate Dehydrogenase Activity Directly in Tissue

Mapping enzymatic activity in space and time is critical for understanding the molecular basis of cell behavior in normal tissue and disease. In situ metabolic activity assays can provide information about the spatial distribution of metabolic activity within a tissue. We provide here a detailed protocol for monitoring the activity of the enzyme lactate dehydrogenase directly in tissue samples. Lactate dehydrogenase is an important determinant of whether consumed glucose will be converted to energy via aerobic or anaerobic glycolysis. A solution containing lactate and NAD is provided to a frozen tissue section. Cells with high lactate dehydrogenase activity will convert the provided lactate to pyruvate, while simultaneously converting provided nicotinamide adenine dinucleotide (NAD) to NADH and a proton, which can be detected based on the reduction of nitrotetrazolium blue to formazan, which is visualized as a blue precipitate. We describe a detailed protocol for monitoring lactate dehydrogenase activity in mouse skin. Applying this protocol, we found that lactate dehydrogenase activity is high in the quiescent hair follicle stem cells within the skin. Applying the protocol to cultured mouse embryonic stem cells revealed higher staining in cultured embryonic stem cells than mouse embryonic fibroblasts. Analysis of freshly isolated mouse aorta revealed staining in smooth muscle cells perpendicular to the aorta. The methodology provided can be used to spatially map the activity of enzymes that generate a proton in frozen or fresh tissue.

We describe a protocol for mapping the spatial distribution of enzymatic activity for enzymes that generate nicotinatmide adenine dinucleotide phosphate (NAD(P)H) + H+ directly in tissue samples.

Mapping enzymatic activity in space and time is critical for understanding the molecular basis of cell behavior in normal tissue and disease. In situ ...

Biochemistry

Anaerobic Protein Purification and Kinetic Analysis via Oxygen Electrode for Studying DesB Dioxygenase Activity and Inhibition

Oxygen-sensitive proteins, including those enzymes which utilize oxygen as a substrate, can have reduced stability when purified using traditional aerobic purification methods. This manuscript illustrates the technical details involved in the anaerobic purification process, including the preparation of buffers and reagents, the methods for column chromatography in a glove box, and the desalting of the protein prior to kinetics. Also described are the methods for preparing and using an oxygen electrode to perform kinetic characterization of an oxygen-utilizing enzyme. These methods are illustrated using the dioxygenase enzyme DesB, a gallate dioxygenase from the bacterium Sphingobium sp. strain SYK-6.

Here we present a protocol for anaerobic protein purification, anaerobic protein concentration, and subsequent kinetic characterization using an oxygen electrode system. The method is illustrated using the enzyme DesB, a dioxygenase enzyme which is more stable and active when purified and stored in an anaerobic environment.

Oxygen-sensitive proteins, including those enzymes which utilize oxygen as a substrate, can have reduced stability when purified using traditional aerobic ...

Enzymatic Cascade Reactions for the Synthesis of Chiral Amino Alcohols from L-lysine

Amino alcohols are versatile compounds with a wide range of applications. For instance, they have been used as chiral scaffolds in organic synthesis. Their synthesis by conventional organic chemistry often requires tedious multi-step synthesis processes, with difficult control of the stereochemical outcome. We present a protocol to enzymatically synthetize amino alcohols starting from the readily available L-lysine in 48 h. This protocol combines two chemical reactions that are very difficult to conduct by conventional organic synthesis. In the first step, the regio- and diastereoselective oxidation of an unactivated C-H bond of the lysine side-chain is catalyzed by a dioxygenase; a second regio- and diastereoselective oxidation catalyzed by a regiodivergent dioxygenase can lead to the formation of the 1,2-diols. In the last step, the carboxylic group of the alpha amino acid is cleaved by a pyridoxal-phosphate (PLP) decarboxylase (DC). This decarboxylative step only affects the alpha carbon of the amino acid, retaining the hydroxy-substituted stereogenic center in a beta/gamma position. The resulting amino alcohols are therefore optically enriched. The protocol was successfully applied to the semipreparative-scale synthesis of four amino alcohols. Monitoring of the reactions was conducted by high performance liquid chromatography (HPLC) after derivatization by 1-fluoro-2,4-dinitrobenzene. Straightforward purification by solid-phase extraction (SPE) afforded the amino alcohols with excellent yields (93% to &#62;95%).

Chiral amino alcohols are versatile molecules for use as scaffolds in organic synthesis. Starting from L-lysine, we synthesize amino alcohols by an enzymatic cascade reaction combining diastereoselective C-H oxidation catalyzed by dioxygenase followed by cleavage of the carboxylic acid moiety of the corresponding hydroxyl amino acid by a decarboxylase.

Amino alcohols are versatile compounds with a wide range of applications. For instance, they have been used as chiral scaffolds in organic synthesis. ...

Expression, Purification, Crystallization, and Enzyme Assays of Fumarylacetoacetate Hydrolase Domain-Containing Proteins

Fumarylacetoacetate hydrolase (FAH) domain-containing proteins (FAHD) are identified members of the FAH superfamily in eukaryotes. Enzymes of this superfamily generally display multi-functionality, involving mainly hydrolase and decarboxylase mechanisms. This article presents a series of consecutive methods for the expression and purification of FAHD proteins, mainly FAHD protein 1 (FAHD1) orthologues among species (human, mouse, nematodes, plants, etc.). Covered methods are protein expression in E. coli, affinity chromatography, ion exchange chromatography, preparative and analytical gel filtration, crystallization, X-ray diffraction, and photometric assays. Concentrated protein of high levels of purity (&#62;98%) may be employed for crystallization or antibody production. Proteins of similar or lower quality may be employed in enzyme assays or used as antigens in detection systems (Western-Blot, ELISA). In the discussion of this work, the identified enzymatic mechanisms of FAHD1 are outlined to describe its hydrolase and decarboxylase bi-functionality in more detail.

Expression and purification of fumarylacetoacetate hydrolase domain-containing proteins is described with examples (expression in E. coli, FPLC). Purified proteins are used for crystallization and antibody production and employed for enzyme assays. Selected photometric assays are presented to display the multi-functionality of FAHD1 as oxaloacetate decarboxylase and acylpyruvate hydrolase.

Fumarylacetoacetate hydrolase (FAH) domain-containing proteins (FAHD) are identified members of the FAH superfamily in eukaryotes. Enzymes of this ...

An Assay for Measuring the Activity of Escherichia coli Inducible Lysine Decarboxyase

Escherichia coli is an enteric bacterium that is capable of growing over a wide range of pH values (pH 5 - 9)1 and, incredibly, is able to survive extreme acid stresses including passage through the mammalian stomach where the pH can fall to as low as pH 1 - 22. To enable such a broad range of acidic pH survival, E. coli possesses four different inducible amino acid decarboxylases that decarboxylate their substrate amino acids in a proton-dependent manner thus raising the internal pH. The decarboxylases include the glutamic acid decarboxylases GadA and GadB3, the arginine decarboxylase AdiA4, the lysine decarboxylase LdcI5, 6 and the ornithine decarboxylase SpeF7. All of these enzymes utilize pyridoxal-5'-phospate as a co-factor8 and function together with inner-membrane substrate-product antiporters that remove decarboxylation products to the external medium in exchange for fresh substrate2. In the case of LdcI, the lysine-cadaverine antiporter is called CadB. Recently, we determined the X-ray crystal structure of LdcI to 2.0 &#197;, and we discovered a novel small-molecule bound to LdcI the stringent response regulator guanosine 5'-diphosphate,3'-diphosphate (ppGpp) 14. The stringent response occurs when exponentially growing cells experience nutrient deprivation or one of a number of other stresses9. As a result, cells produce ppGpp which leads to a signaling cascade culminating in the shift from exponential growth to stationary phase growth10. We have demonstrated that ppGpp is a specific inhibitor of LdcI 14. Here we describe the lysine decarboxylase assay, modified from the assay developed by Phan et al.11, that we have used to determine the activity of LdcI and the effect of pppGpp/ppGpp on that activity. The LdcI decarboxylation reaction removes the &alpha;-carboxy group of L-lysine and produces carbon dioxide and the polyamine cadaverine (1,5-diaminopentane)5. L-lysine and cadaverine can be reacted with 2,4,6-trinitrobenzensulfonic acid (TNBS) at high pH to generate N,N'-bistrinitrophenylcadaverine (TNP-cadaverine) and N,N&#8242;-bistrinitrophenyllysine (TNP-lysine), respectively11. The TNP-cadaverine can be separated from the TNP-lysine as the former is soluble in organic solvents such as toluene while the latter is not (See Figure 1). The linear range of the assay was determined empirically using purified cadaverine.

An Assay for Measuring the Activity of Escherichia coli Inducible Lysine Decarboxyase

The activity of the inducible lysine decarboxylase is monitored by reacting the substrate L-lysine and the product cadaverine with 2,4,6-trinitrobenzensulfonic acid to form adducts that have differential solubility in toluene.

Escherichia coli is an enteric bacterium that is capable of growing over a wide range of pH values (pH 5 - 9)1 and, incredibly, is able to survive extreme ...

Biology

Metabolic Mapping: Quantitative Enzyme Cytochemistry and Histochemistry to Determine the Activity of Dehydrogenases in Cells and Tissues

Altered cellular metabolism is a hallmark of many diseases, including cancer, cardiovascular diseases and infection. The metabolic motor units of cells are enzymes and their activity is heavily regulated at many levels, including the transcriptional, mRNA stability, translational, post-translational and functional level. This complex regulation means that conventional quantitative or imaging assays, such as quantitative mRNA experiments, Western Blots and immunohistochemistry, yield incomplete information regarding the ultimate activity of enzymes, their function and/or their subcellular localization. Quantitative enzyme cytochemistry and histochemistry (i.e., metabolic mapping) show in-depth information on in situ enzymatic activity and its kinetics, function and subcellular localization in an almost true-to-nature situation.
We describe a protocol to detect the activity of dehydrogenases, which are enzymes that perform redox reactions to reduce cofactors such as NAD(P)+ and FAD. Cells and tissue sections are incubated in a medium that is specific for the enzymatic activity of one dehydrogenase. Subsequently, the dehydrogenase that is the subject of investigation performs its enzymatic activity in its subcellular site. In a chemical reaction with the reaction medium, this ultimately generates blue-colored formazan at the site of the dehydrogenase&#39;s activity. The formazan&#39;s absorbance is therefore a direct measure of the dehydrogenase&#39;s activity and can be quantified using monochromatic light microscopy and image analysis. The quantitative aspect of this protocol enables researchers to draw statistical conclusions from these assays.
Besides observational studies, this technique can be used for inhibition studies of specific enzymes. In this context, studies benefit from the true-to-nature advantages of metabolic mapping, giving in situ results that may be physiologically more relevant than in vitro enzyme inhibition studies. In all, metabolic mapping is an indispensable technique to study metabolism at the cellular or tissue level. The technique is easy to adopt, provides in-depth, comprehensive and integrated metabolic information and enables rapid quantitative analysis.

Here, we present a protocol that can be used to microscopically visualize and quantify activity of dehydrogenases in cells and tissues and its kinetics, function and subcellular localization.

Altered cellular metabolism is a hallmark of many diseases, including cancer, cardiovascular diseases and infection. The metabolic motor units of cells ...

Immunology and Infection

Direct Detection of the Acetate-forming Activity  of the Enzyme Acetate Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of acetate to acetyl phosphate utilizing ATP as a substrate. Acetate kinases are ubiquitous in the Bacteria, found in one genus of Archaea, and are also present in microbes of the Eukarya6. The most well characterized acetate kinase is that from the methane-producing archaeon Methanosarcina thermophila7-14. An acetate kinase which can only utilize PPi but not ATP in the acetyl phosphate-forming direction has been isolated from Entamoeba histolytica, the causative agent of amoebic dysentery, and has thus far only been found in this genus15,16.
In the direction of acetyl phosphate formation, acetate kinase activity is typically measured using the hydroxamate assay, first described by Lipmann17-20, a coupled assay in which conversion of ATP to ADP is coupled to oxidation of NADH to NAD+ by the enzymes pyruvate kinase and lactate dehydrogenase21,22, or an assay measuring release of inorganic phosphate after reaction of the acetyl phosphate product with hydroxylamine23. Activity in the opposite, acetate-forming direction is measured by coupling ATP formation from ADP to the reduction of NADP+ to NADPH by the enzymes hexokinase and glucose 6-phosphate dehydrogenase24.
Here we describe a method for the detection of acetate kinase activity in the direction of acetate formation that does not require coupling enzymes, but is instead based on direct determination of acetyl phosphate consumption. After the enzymatic reaction, remaining acetyl phosphate is converted to a ferric hydroxamate complex that can be measured spectrophotometrically, as for the hydroxamate assay. Thus, unlike the standard coupled assay for this direction that is dependent on the production of ATP from ADP, this direct assay can be used for acetate kinases that produce ATP or PPi.

A method for the determination of acetate kinase activity is described. This assay utilizes a direct reaction for determining enzyme activity and kinetics of acetate kinase in the acetate-forming direction with different phosphoryl acceptors. Furthermore, this method can be utilized for assaying other acetyl phosphate or acetyl-CoA utilizing enzymes.

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of ...

Light-Controlled Fermentations for Microbial Chemical and Protein Production

Microbial cell factories offer a sustainable alternative for producing chemicals and recombinant proteins from renewable feedstocks. However, overburdening a microorganism with genetic modifications can reduce host fitness and productivity. This problem can be overcome by using dynamic control: inducible expression of enzymes and pathways, typically using chemical- or nutrient-based additives, to balance cellular growth and production. Optogenetics offers a non-invasive, highly tunable, and reversible method of dynamically regulating gene expression. Here, we describe how to set up light-controlled fermentations of engineered Escherichia coli and Saccharomyces cerevisiae for the production of chemicals or recombinant proteins. We discuss how to apply light at selected times and dosages to decouple microbial growth and production for improved fermentation control and productivity, as well as the key optimization considerations for best results. Additionally, we describe how to implement light controls for lab-scale bioreactor experiments. These protocols facilitate the adoption of optogenetic controls in engineered microorganisms for improved fermentation performance.

Optogenetic control of microbial metabolism offers flexible dynamic control over fermentation processes. The protocol here shows how to set up blue light-regulated fermentations for chemical and protein production at different volumetric scales.

Microbial cell factories offer a sustainable alternative for producing chemicals and recombinant proteins from renewable feedstocks. However, ...

Loss of Carboxy Group as CO2: Decarboxylation of &beta;-Ketoacids

Carboxylic acids, upon heating, undergo a decarboxylation reaction by releasing carbon dioxide gas. Monocarboxylic acids do not undergo decarboxylation easily. However, a silver salt of carboxylic acid&#160;reacts with bromine or iodine under high temperature to release carbon dioxide gas and forms halide&#160;with one less carbon. This reaction is called the Hunsdiecker reaction.
<img alt="Figure1" src="/files/ftp_upload/12352/12352_PT_Figure_1.svg" style="text-align: center; height: 55px;" />
Unlike monocarboxylic acids, &#42933;-keto acids bearing a keto group at the &#946; position to the carboxyl group, readily undergo decarboxylation under an acid medium to form a ketone and releases carbon dioxide gas.
<img alt="Figure2" src="/files/ftp_upload/12352/12352_PT_Figure_2.svg" style="text-align: center; height: 60px;" />
Decarboxylation reaction is biologically relevant while oxidizing food materials in the tricarboxylic acid (TCA) cycle. In this cycle, oxalosuccinic acid is formed as an intermediate that bears three carboxyl groups and one carbonyl group. However, as the carbonyl group is at the &#42933;-position to only one carboxyl group, only one of the three carboxyl groups undergoes decarboxylation to produce &#945;-ketoglutaric acid by losing CO2.
<img alt="Figure3" src="/files/ftp_upload/12352/12352_PT_Figure_3.svg" style="text-align: center; height: 128px;" />

Loss of Carboxy Group as CO2: Decarboxylation of &amp;#946;-Ketoacids

Carboxylic acids, upon heating, undergo a decarboxylation reaction by releasing carbon dioxide gas. Monocarboxylic acids do not undergo decarboxylation ...

Light-driven Enzymatic Decarboxylation

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