This work was supported by NSF award # 0920274 and South Carolina Experiment Station Project (SC-1700340) to KSS. This paper is Technical Contribution No. 5929 of the Clemson University Experiment Station.

The overall scheme of this protocol is outlined in Figure 1.
1. Solution Preparation for Standard Curves and Assays
<ol>
	<li>Prepare 100 mL of a 2 mol/L solution of hydroxylamine-HCl. Weigh out 13.9 g of hydroxylamine hydrochloride (MW 69.49 g/mol) and dissolve in approximately 50 mL distilled-deionized water (ddH2O). Adjust the pH to 7.0 using potassium hydroxide pellets or a concentrated solution. Bring the final volume to 100 mL. The solution can be stored at room temperature for up to 30 days or at 4 &deg;C for up to 90 days.</li>
	<li>Prepare 100 mL of a ferric chloride/ hydrochloric acid solution. Weigh out 13.5 g ferric chloride (MW 270.32 g/mol) and dissolve in approximately 50 mL ddH2O. Add 41.3 mL concentrated hydrochloric acid (12.1 mol/L) and bring to a final volume of 100 mL. The final concentration of ferric chloride will be 0.5 mol/L and the final concentration of hydrochloric acid will be 5 mol/L. The solution is stored at room temperature.</li>
	<li>Prepare 100 mL of a 1.84 mol/L solution of trichoroacetic acid. Weigh out 30.0 g of trichloroacetic acid (MW 163.39 g/mol), dissolve in ddH2O and bring the final volume to 100 mL. The solution is stored at room temperature.</li>
	<li>Prepare 5 mL of a 0.1 mol/L acetyl phosphate solution. Weigh out 0.092 g acetyl phosphate (MW 184.06 g/mol), dissolve in ddH2O and bring the final volume to 5 mL. Aliquot the solution to microcentrifuge tubes (0.25 mL per tube) and freeze at -20 &deg;C until use. Thawed solution should be placed on ice and can be used for approximately 1 hour. Acetyl phosphate solution that has been thawed should not be refrozen and re-used.</li>
	<li>Prepare 5 mL of a 0.1 mol/L solution of ADP. Weigh out 0.24 g of ADP (MW 472.5 g/mol), dissolve in ddH2O, and bring the final volume to 5 mL. Aliquot the solution in microcentrifuge tubes (0.5 mL per tube) and freeze at -20&deg; C until use. Keep thawed solution on ice until use.</li>
	<li>Prepare 50 mL of a 1 mol/L solution of Tris. Weigh out 6.06 g Tris (MW 121.14 g/mol), and dissolve in 40 mL ddH2O. Adjust the pH of the solution to 7.0 using hydrochloric acid and bring final volume to 50 mL. Store solution at room temperature.</li>
	<li>Prepare 10 mL of a 1 mol/L magnesium chloride solution. Weigh out 2.03 g magnesium chloride (MW 203.31 g/mol), dissolve in ddH2O, and bring final volume to 10 mL. Store the solution at room temperature.</li>
	<li>Prepare 25 mL development solution by mixing equal volumes of the ferric chloride/hydrochloric acid solution and the trichloroacetic acid solution. Development solution should be prepared fresh each day and stored at room temperature.</li>
</ol>
2. Preparation of an Acetyl Phosphate Standard Curve
<ol>
	<li>An acetyl phosphate standard curve is generated using known amounts of acetyl phosphate. A standard curve of six to eight points is suitable. The standards should range from 0.03 &mu;moles to 0.9 &mu;moles per 300 &mu;L, which correlates to final concentrations of 0.1 to 3 mmol/L. Dilute the acetyl phosphate stock solution to the appropriate concentrations in 100 mM Tris solution in a final volume of 300 &mu;L. A control sample without acetyl phosphate should also be prepared.</li>
	<li>Incubate each standard and the control at 37 &deg;C in a heat block for 1 minute.</li>
	<li>Add 50 &mu;L of the hydroxylamine hydrochloride solution to each standard and the control and mix by shaking three times. Place the standard into a 60 &deg;C heat block and incubate for five minutes.</li>
	<li>Add 100 &mu;L of development solution to the standards and the control, mix by shaking three times, and allow color to develop for at least one minute at room temperature.</li>
	<li>Centrifuge all standards and the control in a microcentrifuge at 18,000 x g for 1 minute.</li>
	<li>Measure the samples spectrophotometrically at 540 nm using 1mL plastic cuvettes, using the control as the blank for spectrophotometry.</li>
	<li>Generate a standard curve using appropriate data analysis and graphing software (e.g. Kaleidagraph, Synergy Software). R2 values of 0.99 or greater are desirable. The data is plotted as absorbance at 540 nm versus &mu;moles of acetyl phosphate present.</li>
</ol>
3. Assay for Acetate Kinase Activity
<ol>
	<li>Using the solutions described, prepare a reaction mix containing a final concentration of 0.1 mol/L Tris buffer, 10 mmol/L magnesium chloride, 5 mmol/L ADP, and 2 mmol/L acetyl phosphate. A 10 mL reaction mixture will require 8.2 mL ddH2O, 1.0 mL Tris solution, 0.1 mL magnesium chloride solution, 0.5 mL ADP solution, and 0.2 mL acetyl phosphate solution. Distribute the reaction mix in 300 &mu;l aliquots to microcentrifuge tubes. For determination of kinetic parameters and optimization of substrate concentrations, one substrate can be varied and the other held at a constant concentration in the reactions.</li>
	<li>Pre-incubate the reactions for 1 minute at 37 &deg;C in a heat block.</li>
	<li>Add a desired amount of enzyme to the reaction and immediately return the tube to the 37 &deg;C heat block. Also include a control reaction to which no enzyme is added. If several reactions are performed, add enzyme to the tubes at specific time intervals. Be sure to keep track of time. (NOTE: The amount of enzyme to be used will need to be optimized to ensure that all of the acetyl phosphate or ADP substrate will not be depleted - see Discussion section)</li>
	<li>After 5 minutes, add 50&mu;l of the hydroxylamine hydrochloride solution to each reaction and the control, mix, and place the tubes in a 60 &deg;C heat block. If several reactions are being performed, this should be done at the same time interval as the enzyme was added.</li>
	<li>Incubate the reactions and the control for 5 minutes at 60 &deg;C.</li>
	<li>Add 100 &mu;L of the development solution to each reaction and the control, mix, and place in a tube rack on the lab bench. Allow color to develop for at least 1 minute.</li>
	<li>Centrifuge all tubes in a microcentrifuge at 18,000 x g for 1 minute.</li>
	<li>Measure samples spectrophotometrically at 540 nm using 1mL plastic cuvettes and determine the amount of acetyl phosphate present by comparison to the acetyl phosphate standard curve.</li>
	<li>The amount of acetyl phosphate substrate depleted can be determined by subtracting the amount of acetyl phosphate remaining after the enzymatic reaction from the amount of acetyl phosphate present in the control lacking enzyme. Perform data analysis using appropriate data and graphing software (e.g. Kaleidagraph, Synergy Software). R2 values of greater than 0.97 are desirable.</li>
</ol>
4. Representative Results:
The purpose of this assay is to measure acetate kinase activity in the direction of acetate formation. This is done by measuring consumption of the acetyl phosphate substrate, as there is no easy, direct measurement for acetate production. Acetyl phosphate remaining after a given time during the enzymatic reaction is converted to acetyl hydroxamate and subsequently to ferric hydroxamate complex, which has a reddish color (Figure 2) that can be measured at 540 nm. The standard curve plotting absorbance versus &mu;moles acetyl phosphate present in the standards is used to determine the amount of acetyl phosphate remaining in each reaction. The standard curve should be linear through the zero point. A representative standard curve shown in Figure 3 has a slope of 1.2332 absorbance units per &mu;mole acetyl phosphate present and an R2 value of 0.99, indicating the data fits the linear equation well. The equation for the linear fit to the standard curve data is used for calculating the quantity of acetyl phosphate remaining in the reaction. The &mu;moles of acetyl phosphate consumed is determined as the difference between the &mu;moles of acetyl phosphate present in the control reaction lacking enzyme and the &mu;moles of acetyl phosphate remaining after the enzymatic reaction.
Purified, recombinant M. thermophila acetate kinase was assayed using the described protocol. For the experiment shown in Figure 4, the concentration of ADP was held constant at 5 mmol/L and the concentration of acetyl phosphate in the reaction was varied to determine the Km for acetyl phosphate. As expected, this experiment demonstrates that the enzyme follows standard Michaelis-Menten-like kinetics for each substrate and produced a hyperbolic saturation curve when plotting activity versus acetyl phosphate concentration. The apparent Km value for acetyl phosphate was 0.27 &plusmn; 0.01 mmol/L, which compares favorably to the published Km value of 0.47 &plusmn; 0.01 mmol/L using the coupled assay13.
The direct assay can also be used to measure activity of acetate kinases that utilize substrates other than ATP, which is not possible using the standard coupled assay. Purified recombinant E. histolytica acetate kinase16, which produces PPi rather than ATP in the acetate-forming direction, was subjected to this assay using sodium phosphate in place of ADP. The concentration of sodium phosphate in the reaction was held constant at 0.2 mol/L and the acetyl phosphate concentration was varied. As for the M. thermophila acetate kinase, the E. histolytica acetate kinase followed Michaelis-Menten-like kinetics for acetyl phosphate and a hyperbolic saturation curve was observed (Figure 5). The apparent Km value for acetyl phosphate was 0.50 &plusmn; 0.006 mmol/L. Reeves and Guthrie15 determined a Km value of 0.06 mmol/L for acetyl phosphate for the native enzyme; however, their enzyme was only partially purified and their assay involved four coupling enzymes which were also only partially purified. Thus, it is difficult to directly compare these values.
Protocol Scheme
<ol>
	<li>Generate an acetyl phosphate standard curve</li>
	<li>Dilute enzyme to desired concentration</li>
	<li>Prepare the reaction mixture and aliquot 300 &mu;l to microcentrifuge tubes</li>
	<li>Preincubate reactions at 37 &deg;C for 1 minute</li>
	<li>Start reactions by adding enzyme at specific time intervals</li>
	<li>Add hydroxylamine at the same intervals to stop the reactions and incubate at 60 &deg;C for 5 minutes</li>
	<li>Add ferric chloride/trichloroacetic acid solution for color development</li>
	<li>Centrifuge tubes at 18,000 x g for 1 minute</li>
	<li>Measure absorbance at 540 nm</li>
	<li>Determine amount of acetyl phosphate consumed by comparison to the standard curve</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/3474/3474fig1.jpg" /> 
Figure 1. Protocol Scheme
<img alt="Figure 2" src="/files/ftp_upload/3474/3474fig2.jpg" /> 
Figure 2. Acetyl phosphate standards. Increasing concentrations of acetyl phosphate were reacted with hydroxylamine and color was developed as described. The control lacking acetyl phosphate is bright yellow, and reactions containing increasing amounts of acetyl phosphate are successively darker in color. This color change is measured at 540 nm.
<img alt="Figure 3" src="/files/ftp_upload/3474/3474fig3.jpg" /> 
Figure 3. Sample acetyl phosphate standard curve. Varying amounts of acetyl phosphate were reacted with hydroxylamine and the color development solution and absorbance was measured at 540 nm. The absorbance at 540 nm versus &mu;moles acetyl phosphate was plotted and a linear fit to the data was applied.
<img alt="Figure 4" src="/files/ftp_upload/3474/3474fig4.jpg" /> 
Figure 4. Utilization of acetyl phosphate by the ATP-producing M. thermophila acetate kinase. Enzymatic activity was determined in the direction of acetate formation in the presence of 5 mmol/L ADP with varying concentrations of acetyl phosphate. The amount of acetyl phosphate consumed was determined as the difference between the starting amount of acetyl phosphate in the reaction and the amount of acetyl phosphate remaining after the enzymatic reaction. Assays were performed in triplicate with error bars shown.
<img alt="Figure 5" src="/files/ftp_upload/3474/3474fig5.jpg" /> 
Figure 5. Utilization of acetyl phosphate by the PPi-producing E. histolytica acetate kinase. Enzymatic activity in the direction of acetate formation was determined in the presence of 0.2 mol/L sodium phosphate with varying concentrations of acetyl phosphate. The amount of acetyl phosphate consumed was determined as the difference between the starting amount of acetyl phosphate in the reaction and the amount of acetyl phosphate remaining after the enzymatic reaction. Assays were performed in triplicate with error bars shown.

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No conflicts of interest declared.

The detection of acetyl phosphate in this assay is dependent upon a sufficient concentration of hydroxylamine hydrochloride and the concentration and acidity of the ferric chloride solution. Alteration of the assay volume will require reconsideration of both of these components. The enzymatic reactions described here were performed at 37&deg;C with the hydroxylamine termination performed at 60 &deg;C for 5 minutes. This higher temperature is critical to allow for rapid conversion of the remaining acetyl phosphate to acet...

<table border="1">
 <tr>
 <td width="160">Name of the reagent</td>
 <td width="108">Company</td>
 <td width="134">Catalogue number</td>
 <td width="103">Comments</td>
 </tr>
 <tr>
 <td>Acetyl phosphate</td>
 <td>Sigma Aldrich</td>
 <td>01409</td>
 <td>Lithium Salt (97% )</td>
 </tr>
 <tr>
 <td>Sodium phosphate monobasic (dehydrate)</td>
 <td>ThermoFisher</td>
 <td>S381</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sodium phosphate dibasic (anhydrous)</td>
 <td>ThermoFisher</td>
 <td>S374</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Magnesium chloride (hexahydrate)</td>
 <td>ThermoFisher</td>
 <td>M33</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tris Base</td>
 <td>ThermoFisher</td>
 <td>B152</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Ferric chloride (hexahydrate)</td>
 <td>ThermoFisher</td>
 <td>I88</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Trichloroacetic acid</td>
 <td>ThermoFisher</td>
 <td>A324</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Hydroxylamine hydrochloride</td>
 <td>ThermoFisher</td>
 <td>H330</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Adenosine 5&rsquo;-diphosphate 
 sodium salt</td>
 <td>Sigma Aldrich</td>
 <td>A2754</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Biomate III Spectrophotometer</td>
 <td>ThermoFisher</td>
 <td>142982082</td>
 <td>Standard UV/Vis spectrophotometer</td>
 </tr>
</table>

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.

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

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 ...

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23.0K Views.  Clemson University. 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.

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

Detection of Protein Ubiquitination

Ubiquitination, the covalent attachment of the polypeptide ubiquitin to target proteins, is a key posttranslational modification carried out by a set of three enzymes. They include ubiquitin-activating enzyme E1, ubiquitin-conjugating enzyme E2, and ubiquitin ligase E3. Unlike to E1 and E2, E3 ubiquitin ligases display substrate specificity. On the other hand, numerous deubiquitylating enzymes have roles in processing polyubiquitinated proteins. Ubiquitination can result in change of protein stability, cellular localization, and biological activity. Mutations of genes involved in the ubiquitination/deubiquitination pathway or altered ubiquitin system function are associated with many different human diseases such as various types of cancer, neurodegeneration, and metabolic disorders. The detection of altered or normal ubiquitination of target proteins may provide a better understanding on the pathogenesis of these diseases. &nbsp;Here, we describe protocols to detect protein ubiquitination in cultured cells in vivo and test tubes in vitro. These protocols are also useful to detect other ubiquitin-like small molecule modification such as sumolyation and neddylation.

Ubiquitination is a key posttranslational modification carried out by a set of three enzymes. Mutations of genes involved in this modification are associated with many different human diseases. Here, we describe protocols to detect protein ubiquitination in cultured cells in vivo and test tubes in vitro.

Ubiquitination, the covalent attachment of the polypeptide ubiquitin to target proteins, is a key posttranslational modification carried out by a set of ...

Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect protease activity uses casein labeled with fluorescein isothiocyanate (FITC) as the substrate.
Protease activity results in the cleavage of the FITC-labeled casein substrate into smaller fragments, which do not precipitate under acidic conditions. After incubation of the protease sample and substrate, the reaction is acidified with the addition of trichloroacetic acid (TCA). The mixture is then centrifuged with the undigested substrate forming a pellet and the smaller, acid soluble fragments remaining in solution. The supernatant is neutralized and the fluorescence of the FITC-labeled fragments is measured.
The described kit procedure detects the trypsin protease control at a concentration of approximately 0.5 &mu;g/ml (5 ng of trypsin added to the assay). This sensitivity can be increased with a longer incubation time, up to 24 hours. The assay is performed in microcentrifuge tubes and procedures are provided for fluorescence detection using either cuvettes or multiwell plates.

The Protease Fluorescent Detection Kit is designed for the measurement of protease activity using fluorometry. It is also suitable for detection of trace amounts of protease contamination. The method is based on the proteolytic hydroysis of a proprietary formulation of a FITC-labeled casein substrate.

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect ...

ELIME (Enzyme Linked Immuno Magnetic Electrochemical) Method for Mycotoxin Detection

Immunoassays are a valid alternative to the more expensive and time consuming quantitative HPLC or GC1, 2 methods for the screening detection of hazardous mycotoxins in food commodities. In this protocol we show how to fabricate and interrogate an electrochemical competitive Enzyme linked immunomagnetic assay based on the use of magnetic beads as solid support for the immunochemical chain3 and screen printed electrodes as sensing platform.
Our method aims to determine the total amount of HT-2 and T-2 toxins, mycotoxins belonging to the trichothecenes family and of great concern for human health4. The use of an antibody clone with a cross reactivity of 100% towards HT-2 and T-2 allows to simultaneously detect both toxins with similar sensitivity5.
The first step of our assay is the coating step where we immobilize HT2-KLH conjugate toxin on the surface of magnetic beads. After a blocking step, necessary to avoid non-specific absorptions, the addition of a monoclonal antibody allows the competition between immobilized HT-2 and free HT-2 or T-2 present in the sample or dissolved in a standard solution.
At the end of the competition step, the amount of monoclonal antibody linked to the immobilized HT-2 will be inversely proportional to the amount of toxin in the sample solution.
A secondary antibody labeled with alkaline phosphatase (AP) is used to reveal the binding between the specific antibody and the immobilized HT-2. The final measurement step is performed by dropping an aliquot of magnetic bead suspension, corresponding to a specific sample/standard solution, on the surface of a screen-printed working electrode; magnetic beads are immobilized and concentrated by means of a magnet placed precisely under the screen-printed electrode. After two minutes of incubation between magnetic beads and a substrate for AP, the enzymatic product is detected by Differential Pulse Voltammetry (DPV) using a portable instrument (PalmSens) also able to initiate automatically eight measurements within an interval of few seconds.

ELIME Enzyme Linked Immuno Magnetic Electrochemical Method for Mycotoxin Detection

A protocol to detect trichothecenes (mycotoxins of concern for human health) using a newly developed screening method based on a competitive immunochemical method and a final electrochemical detection is demonstrated.

Immunoassays are a valid alternative to the more expensive and time consuming quantitative HPLC or GC1, 2 methods for the screening detection of hazardous ...

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later stages of embryonic development, and to define how enhancers and promoters regulate gene expression within the embryo. The protocol demonstrated here can be used to efficiently produce transgenic Xenopus laevis embryos. This transgenesis approach involves three parts: 1. Sperm nuclei are isolated from adult X. laevis testis by treatment with lysolecithin, which permeabilizes the sperm plasma membrane. 2. Egg extract is prepared by low speed centrifugation, addition of calcium to cause the extract to progress to interphase of the cell cycle, and a high-speed centrifugation to isolate interphase cytosol. 3. Nuclear transplantation: the nuclei and extract are combined with the linearized plasmid DNA to be introduced as the transgene and a small amount of restriction enzyme. During a short reaction, egg extract partially decondenses the sperm chromatin and the restriction enzyme generates chromosomal breaks that promote recombination of the transgene into the genome. The treated sperm nuclei are then transplanted into unfertilized eggs. Integration of the transgene usually occurs prior to the first embryonic cleavage such that the resulting embryos are not chimeric. These embryos can be analyzed without any need to breed to the next generation, allowing for efficient and rapid generation of transgenic embryos for analyses of promoter and gene function. Adult X. laevis resulting from this procedure also propagate the transgene through the germline and can be used to generate lines of transgenic animals for multiple purposes.

Production of Transgenic Xenopus laevis by Restriction Enzyme Mediated Integration and Nuclear Transplantation

This video protocol demonstrates a method for generating transgenic Xenopus laevis by introduction of transgenes into sperm nuclei followed by nuclear transplantation into unfertilized eggs.

Stable integration of cloned gene products into the Xenopus genome is necessary to control the time and place of expression, to express genes at later ...

Detection of Nitric Oxide and Superoxide Radical Anion by Electron Paramagnetic Resonance Spectroscopy from Cells using Spin Traps

Reactive nitrogen/oxygen species (ROS/RNS) at low concentrations play an important role in regulating cell function, signaling, and immune response but in unregulated concentrations are detrimental to cell viability1, 2. While living systems have evolved with endogenous and dietary antioxidant defense mechanisms to regulate ROS generation, ROS are produced continuously as natural by-products of normal metabolism of oxygen and can cause oxidative damage to biomolecules resulting in loss of protein function, DNA cleavage, or lipid peroxidation3, and ultimately to oxidative stress leading to cell injury or death4. 
Superoxide radical anion (O2&bull;-) is the major precursor of some of the most highly oxidizing species known to exist in biological systems such as peroxynitrite and hydroxyl radical. The generation of O2&bull;- signals the first sign of oxidative burst, and therefore, its detection and/or sequestration in biological systems is important. In this demonstration, O2&bull;- was generated from polymorphonuclear neutrophils (PMNs). Through chemotactic stimulation with phorbol-12-myristate-13-acetate (PMA), PMN generates O2&bull;- via activation of nicotinamide adenine dinucleotide phosphate (NADPH) oxidase5. 
Nitric oxide (NO) synthase which comes in three isoforms, as inducible-, neuronal- and endothelial-NOS, or iNOS, nNOS or eNOS, respectively, catalyzes the conversion of L- arginine to L-citrulline, using NADPH to produce NO6. Here, we generated NO from endothelial cells. Under oxidative stress conditions, eNOS for example can switch from producing NO to O2&bull;- in a process called uncoupling, which is believed to be caused by oxidation of heme7 or the co-factor, tetrahydrobiopterin (BH4)8.
There are only few reliable methods for the detection of free radicals in biological systems but are limited by specificity and sensitivity. Spin trapping is commonly used for the identification of free radicals and involves the addition reaction of a radical to a spin trap forming a persistent spin adduct which can be detected by electron paramagnetic resonance (EPR) spectroscopy. The various radical adducts exhibit distinctive spectrum which can be used to identify the radicals being generated and can provide a wealth of information about the nature and kinetics of radical production9.
The cyclic nitrones, 5,5-dimethyl-pyrroline-N-oxide, DMPO10, the phosphoryl-substituted DEPMPO11, and the ester-substituted, EMPO12 and BMPO13, have been widely employed as spin traps--the latter spin traps exhibiting longer half-lives for O2&bull;- adduct. Iron (II)-N-methyl-D-glucamine dithiocarbamate, Fe(MGD)2 is commonly used to trap NO due to high rate of adduct formation and the high stability of the spin adduct14.

Electron paramagnetic resonance (EPR) spectroscopy was employed to detect nitric oxide from bovine aortic endothelial cells and superoxide radical anion from human neutrophils using iron (II)-N-methyl-D-glucamine dithiocarbamate, Fe(MGD)2 and 5,5-dimethyl-1-pyroroline-N-oxide, DMPO, respectively.

Reactive nitrogen/oxygen species (ROS/RNS) at low concentrations play an important role in regulating cell function, signaling, and immune response but in ...

Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such probes typically consist of variants of CFP and YFP, intervened by a phosphorylatable sequence and a phospho-binding domain. Upon phosphorylation, the probe changes conformation, which results in a change of the distance or orientation between CFP and YFP, leading to a change in FRET efficiency (Fig 1). Several probes have been published during the last decade, monitoring the activity balance of multiple kinases and phosphatases, including reporters of PKA2, PKB3, PKC4, PKD5, ERK6, JNK7, Cdk18, Aurora B9 and Plk19. Given the modular design, additional probes are likely to emerge in the near future10. 
Progression through the cell cycle is affected by stress signaling pathways 11. Notably, the cell cycle is regulated differently during unperturbed growth compared to when cells are recovering from stress12.Time-lapse imaging of cells through the cell cycle therefore requires particular caution. This becomes a problem particularly when employing ratiometric imaging, since two images with a high signal to noise ratio are required to correctly interpret the results. Ratiometric FRET imaging of cell cycle dependent changes in kinase and phosphatase activities has predominately been restricted to sub-sections of the cell cycle8,9,13,14.
Here, we discuss a method to monitor FRET-based probes using ratiometric imaging throughout the human cell cycle. The method relies on equipment that is available to many researchers in life sciences and does not require expert knowledge of microscopy or image processing.

FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such ...

Assaying the Kinase Activity of LRRK2 in vitro

Leucine Rich Repeat Kinase 2 (LRRK2) is a 2527 amino acid member of the ROCO family of proteins, possessing a complex, multidomain structure including a GTPase domain (termed ROC, for Ras of Complex proteins) and a kinase domain1. The discovery in 2004 of mutations in LRRK2 that cause Parkinson's disease (PD) resulted in LRRK2 being the focus of a huge volume of research into its normal function and how the protein goes awry in the disease state2,3. Initial investigations into the function of LRRK2 focused on its enzymatic activities4-6. Although a clear picture has yet to emerge of a consistent alteration in these due to mutations, data from a number of groups has highlighted the importance of the kinase activity of LRRK2 in cell death linked to mutations7,8. Recent publications have reported inhibitors targeting the kinase activity of LRRK2, providing a key experimental tool9-11. In light of these data, it is likely that the enzymatic properties of LRRK2 afford us an important window into the biology of this protein, although whether they are potential drug targets for Parkinson's is open to debate.
A number of different approaches have been used to assay the kinase activity of LRRK2. Initially, assays were carried out using epitope tagged protein overexpressed in mammalian cell lines and immunoprecipitated, with the assays carried out using this protein immobilised on agarose beads4,5,7. Subsequently, purified recombinant fragments of LRRK2 in solution have also been used, for example a GST tagged fragment purified from insect cells containing residues 970 to 2527 of LRRK212. Recently, Dani&euml;ls et al. reported the isolation of full length LRRK2 in solution from human embryonic kidney cells, however this protein is not widely available13. In contrast, the GST fusion truncated form of LRRK2 is commercially available (from Invitrogen, see table 1 for details), and provides a convenient tool for demonstrating an assay for LRRK2 kinase activity. Several different outputs for LRRK2 kinase activity have been reported. Autophosphorylation of LRRK2 itself, phosphorylation of Myelin Basic Protein (MBP) as a generic kinase substrate and phosphorylation of an artificial substrate - dubbed LRRKtide, based upon phosphorylation of threonine 558 in Moesin - have all been used, as have a series of putative physiological substrates including &alpha;-synuclein, Moesin and 4-EBP14-17. The status of these proteins as substrates for LRRK2 remains unclear, and as such the protocol described below will focus on using MBP as a generic substrate, noting the utility of this system to assay LRRK2 kinase activity directed against a range of potential substrates.

Assaying the Kinase Activity of LRRK2 in vitro

Leucine Rich Repeat Kinase 2 is a large multidomain kinase, mutations in which are the most common genetic cause of Parkinson's disease. Analysis of the kinase activity of this protein has proven to be a crucial tool in understanding the biology and dysfunction of this protein. In this paper, in vitro assaying of the kinase activity of LRRK2 and a selection of its mutants is described, providing an experimental system to examine phosphorylation of putative substrates and potential dysfunction of LRRK2 in disease.

Leucine Rich Repeat Kinase 2 (LRRK2) is a 2527 amino acid member of the ROCO family of proteins, possessing a complex, multidomain structure including a ...

GC-based Detection of Aldononitrile Acetate Derivatized Glucosamine and Muramic Acid for Microbial Residue Determination in Soil

Quantitative approaches to characterizing microorganisms are crucial for a broader understanding of the microbial status and function within ecosystems. Current strategies for microbial analysis include both traditional laboratory culture-dependent techniques and those based on direct extraction and determination of certain biomarkers1, 2. Few among the diversity of microbial species inhabiting soil can be cultured, so culture-dependent methods introduce significant biases, a limitation absent in biomarker analysis.
The glucosamine, mannosamine, galactosamine and muramic acid have been well served as measures of both the living and dead microbial mass, of these the glucosamine (most abundant) and muramic acid (uniquely from bacterial cell) are most important constituents in the soil systems3, 4. However, the lack of knowledge on the analysis restricts the wide popularization among scientific peers. Among all existing analytical methods, derivatization to aldononitrile acetates followed by GC-based analysis has emerged as a good option with respect to optimally balancing precision, sensitivity, simplicity, good chromatographic separation, and stability upon sample storage5.
Here, we present a detailed protocol for a reliable and relatively simple analysis of glucosamine and muramic acid from soil after their conversion to aldononitrile acetates. The protocol mainly comprises four steps: acid digestion, sample purification, derivatization and GC determination. The step-by-step procedure is modified according to former publications6, 7. In addition, we present a strategy to structurally validate the molecular ion of the derivative and its ion fragments formed upon electron ionization. We applied GC-EI-MS-SIM, LC-ESI-TOF-MS and isotopically labeled reagents to determine the molecular weight of aldononitrile acetate derivatized glucosamine and muramic acid; we used the mass shift of isotope-labeled derivatives in the ion spectrum to investigate ion fragments of each derivatives8. In addition to the theoretical elucidation, the validation of molecular ion of the derivative and its ion fragments will be useful to researchers using &delta;13C or ion fragments of these biomarkers in biogeochemical studies9, 10.

We describe a method protocol for the GC-based analysis of the aldonitrile acetate derivatives of glucosamine and muramic acid extracted from soil. For elucidation of the chemical mechanism, we also present a strategy to confirm the structure of the derivative and the ion fragments formed upon electron ionization.

Quantitative approaches to characterizing microorganisms are crucial for a broader understanding of the microbial status and function within ecosystems. ...

Measurement of Larval Activity in the Drosophila Activity Monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...

Studying the Activity of Neuropeptides and Other Regulators of the Excretory System in the Adult Mosquito

Studies of insect physiology, particularly in those species that are vectors of pathogens causing disease in humans and other vertebrates, provide the foundation to develop novel strategies for pest control. Here, a series of methods are described that are routinely utilized to determine the functional roles of neuropeptides and other neuronal factors (i.e., biogenic amines) on the excretory system of the mosquito, Aedes aegypti. The Malpighian tubules (MTs), responsible for primary urine formation, can continue functioning for hours when removed from the mosquito, allowing for fluid secretion measurements following hormone treatments. As such, the Ramsay assay is a useful technique to measure secretion rates from isolated MTs. Ion-selective microelectrodes (ISME) can sequentially be used to measure ion concentrations (i.e., Na+ and K+) in the secreted fluid. This assay allows for the measurement of several MTs at a given time, determining the effects of various hormones and drugs. The Scanning Ion-selective Electrode Technique uses ISME to measure voltage representative of ionic activity in the unstirred layer adjacent to the surface of ion transporting organs to determine transepithelial transport of ions in near real time. This method can be used to understand the role of hormones and other regulators on ion absorption or secretion across epithelia. Hindgut contraction assays are also a useful tool to characterize myoactive neuropeptides, that may enhance or reduce the ability of this organ to remove excess fluid and waste. Collectively, these methods provide insight into how the excretory system is regulated in adult mosquitoes. This is important because functional coordination of the excretory organs is crucial in overcoming challenges such as desiccation stress after eclosion and before finding a suitable vertebrate host to obtain a bloodmeal.

This protocol outlines methodologies behind the Ramsay assay, ion-selective microelectrodes, Scanning Ion-selective Electrode Technique (SIET) and in vitro contraction assays, applied to study the adult mosquito excretory system, comprised of the Malpighian tubules and hindgut, to collectively measure ion and fluid secretion rates, contractile activity, and transepithelial ion transport.

Studies of insect physiology, particularly in those species that are vectors of pathogens causing disease in humans and other vertebrates, provide the ...