The authors acknowledge funding for sponsored research provided by CoA Therapeutics, Inc., a subsidiary of BridgeBio LLC, the National Institutes of Health grant GM34496, and the American Lebanese Syrian Associated Charities.

The animal procedure referred to in this protocol was performed according to protocols 323 and 556 and specifically approved by the St. Jude Children&#39;s Research Hospital Institutional Animal Care and Use Committee.
1. Preparation of solutions
NOTE: Use ultrapure water for all solutions and when stated in procedures.
<ol>
	<li>Prepare 1 mM potassium hydroxide (KOH) in water.</li>
	<li>Prepare 0.25 M KOH in water.</li>
	<li>Prepare 1 M Trizma-HCl in water and adjust to pH 8.0.</li>
	<li>Prepare 100 mM monobromobimane (mBBr) in acetonitrile (Optima Grade). Stock solutions of mBBr are stable at -20 &#176;C for many months provided they are kept in the dark as exposure to light can photolyze the monobromobimane to bimane26.</li>
	<li>Prepare Wash Buffer for solid phase extraction (SPE) column: 50% methanol (Optima Grade) + 2% acetic acid.</li>
	<li>Prepare Elution Buffer for SPE column: 95% ethanol (HPLC Grade) containing 50 mM ammonium formate.</li>
</ol>
2. Preparation of CoA-bimane standard
<ol>
	<li>Purchase a high-quality CoA standard from a reputable source (e.g., Avanti Polar Lipids, Inc.).</li>
	<li>Prepare a high concentration stock solution of CoA in 20 mM Tris, pH 8. The amount of CoA in the high concentration stock is determined or confirmed by spectrophotometric absorbance at &#955;260 nm (&#400; = 16,800 M-1&#8729;cm-1). Add 4-fold molar excess of mBBr; vortex on high for 10 s. For example, for 10 mM CoA in buffer, add 40 mM mBBr. The mBBr is added in excess to ensure that all the CoA is derivatized.</li>
	<li>Incubate at room temperature for 4 h in the dark without shaking to derivatize the CoA with mBBr. mBBr reacts with the thiol group of the CoA, and it is pH and temperature dependent27. Addition of the mBBr converts CoA to a water-soluble fluorescent (or ultraviolet absorbant) CoA-bimane (CoA-bimane; Figure 2).</li>
	<li>Add 100 &#956;L acetic acid and vortex on high for 10 s to stop the reaction.</li>
	<li>Centrifuge at 2,000 x g for 15 minutes to remove any precipitant.</li>
	<li>Clean-up the supernatant using an SPE-column to remove the unreacted mBBr (described below). Filtration and centrifugation of the eluate is not necessary after the SPE column clean-up (Section 5.8).</li>
	<li>Collect the eluate from the SPE column containing the CoA-bimane in a pre-weighed tube, dry under nitrogen gas, weigh and construct a standard calibration curve with known quantities of CoA-bimane.
	<ol>
		<li>Check the CoA-bimane standard for purity by HPLC (see below) with absorbance detection at both &#955;260 (adenine moiety) and &#955;393 (bimane moiety) and coincidence of both peaks in the chromatogram.</li>
		<li>Confirm the quantity of the CoA-bimane standard in the high concentration stock using &#955;260 nm (&#400; = 16,800 M-1&#8729;cm-1).</li>
	</ol>
	</li>
	<li>Register the HPLC retention time for identification of CoA-bimane in experimental samples using the CoA-bimane standard.</li>
	<li>Store aliquots of the high concentration stock of the CoA-bimane standard protected from light and stored at -20 &#176;C for up to 2 years. Avoid repeated thawing and re-freezing.</li>
</ol>
3. Extraction and derivatization of CoA in cultured cells
<ol>
	<li>Harvest adherent cells in culture when approaching late subconfluent densities. For example, human HepG2/C3A cells are grown to a density of 6-8 x 106, or HEK 293T cells are grown to a density of ~1.3 x 107 per 100 mm dish.
	<ol>
		<li>Use duplicate or triplicate cultures routinely for CoA determinations. Incubate separate culture dishes in parallel with the sample cell cultures to determine viable cell number. Calculate the amount of CoA per 106-107 cells after HPLC purification.</li>
	</ol>
	</li>
	<li>Aspirate culture medium off the dish. Quickly wash cells on the dish with ice-cold phosphate-buffered saline (PBS) to remove any residual medium and aspirate the PBS off the dish. Quickly wash cells with ice-cold water to remove residual PBS and aspirate the water off the dish quickly. Avoid disturbing the adherent cells when adding the PBS or water.</li>
	<li>Add 1 mL of ice-cold water to the culture dish. Scrape cells into the cold water in the dish and transfer the cell suspension to a glass test tube containing 400 &#956;L of 0.25 M KOH and 1.5 mL of water.
	<ol>
		<li>Mix the suspension vigorously by vortex on high for 10 s. Then cover tightly with paraffin film and incubate at 55 &#176;C for 1 h without shaking in a water bath. The pH of the sample should be &#8805; 12. The high pH is important to hydrolyze the CoA thioesters and can be adjusted with additional aliquots of KOH if necessary.</li>
	</ol>
	</li>
	<li>Harvest cultured cells growing in suspension by centrifugation at low speed: 136 x g for 6 min at 4 &#176;C. Wash once with ice-cold PBS, then briefly wash again with cold water, and finally resuspend in 2.5 mL of cold water plus 400 &#181;L 0.25 M KOH. Mix vigorously and incubate at 55 &#176;C as described above.</li>
	<li>Add 160 &#956;L of 1 M Trizma-HCl and 10 &#956;L of 100 mM mBBr and mix by vortex on high for 10 s. This brings the pH to approximately 8 to support the mBBr reaction with free CoA. Cover and incubate samples at room temperature for 2 h in the dark for the mBBr to react with the thiol of CoA.</li>
	<li>Add 100 &#956;L of acetic acid and mix by vortex on high for 10 s to stop the reaction</li>
	<li>Centrifuge at 2,000 x g for 15 min to remove precipitated cell debris.</li>
	<li>Remove and save supernatant in a glass test tube for the SPE column clean-up that is described below.</li>
</ol>
4. Extraction and derivatization of CoA in tissues
<ol>
	<li>Dissect animal tissue pieces, about 0.5 cm diameter or smaller, and blot briefly (1-2 s) on absorbent paper and then flash freeze in liquid nitrogen and store at -80 &#176;C until processing. CoA in tissues is hydrolyzed or converted to a thioester if not flash frozen. This step is important to obtain the maximum yield of CoA in tissues.</li>
	<li>Weigh frozen tissue pieces (5 &#8211; 60 mg) quickly prior to starting the analysis, and record the weight for each sample. This wet weight determination is used for calculation of the CoA content following HPLC purification. Avoid complete thaw of the tissue pieces.</li>
	<li>Add 2 mL of 1 mM KOH to a glass test tube (e.g., 13 mm x 100 mm disposable) destined for insertion of a probe and tissue disruption with a rotor-stator homogenizer. Keep the tube on ice until use. This is done to ensure that the samples are homogenized at a cold temperature and CoA is not destroyed due to the heat generated during homogenization.</li>
	<li>Transfer and homogenize tissue (30 &#8211; 40 mg) in the glass test tube (containing cold 1 mM KOH) for 30 s. Avoid complete thawing of the tissue.</li>
	<li>Add 500 &#956;L of 0.25 M KOH; vortex on high for 10 s and keep on ice until all samples are homogenized. This will bring the pH of the sample above 12 to hydrolyze the CoA thioesters to yield total CoA (free CoA + hydrolyzed CoA thioesters).</li>
	<li>Incubate samples at 55 &#176;C for 2 h in a water bath without shaking to support hydrolysis.</li>
	<li>Add 150 &#956;L of 1 M Trizma-HCl and 10 &#956;L of 100 mM mBBr; vortex on high for 10 seconds. This brings the pH to about 8 to support the mBBr reaction with free CoA.</li>
	<li>Incubate samples at room temperature for 2 h in the dark to ensure all the CoA is derivatized with mBBr.</li>
	<li>Add 100 &#956;L of acetic acid and vortex on high for 10 s to stop the reaction.</li>
	<li>Centrifuge at 2,000 x g for 15 min to pellet precipitated cell debris.</li>
	<li>Remove and save supernatant in a glass test tube for the SPE column clean-up (described below).</li>
</ol>
5. Sample clean-up with solid phase extraction (SPE) column
<ol>
	<li>At room temperature, equilibrate each disposable 2-(2-pyridyl)-ethyl silica gel column (1 mL size) with 1 mL of Wash Buffer to ensure that the pyridyl functional group is protonated and will function as an anion-exchanger. 2-(2-pyridyl)-ethyl silica gel is a weak anion exchanger which is ideal for CoA-bimane at a basic pH. 2-(2-pyridyl)-ethyl silica gel has a pKa of 6 and the elution is done pH &#8805; 7.</li>
	<li>Add sample supernatant (step 4.11) to column and collect eluate.</li>
	<li>Wash column twice with 1 mL of Wash Buffer to remove any unretained species.</li>
	<li>Wash column once with 1 mL of water.</li>
	<li>Wash column twice with 1 mL of Elution Buffer into a separate glass test tube (12 mm x 75 mm) to collect the CoA-bimane.</li>
	<li>Dry CoA-bimane sample in tube under nitrogen gas to dryness. The dried sample is stable at room temperature until further use. Seal and completely cover the tube and store.</li>
	<li>When ready for HPLC analysis, resuspend sample in 300 &#956;L of water and mix vigorously by vortex on high for 10 s.</li>
	<li>Transfer resuspended sample to a centrifuge tube filter (0.22 &#181;m cellulose acetate, 2 mL size) and centrifuge at 5,000 x g for 10 min to remove any precipitant.</li>
	<li>Transfer filtered sample to a glass vial suitable for HPLC injection.</li>
</ol>
6. HPLC purification and measurement of CoA-bimane
<ol>
	<li>Prepare buffers. Buffer A: 50 mM KH2PO4, pH 4.6; Buffer B: Acetonitrile (Optima Grade).</li>
	<li>Power up the HPLC system. 
	NOTE: The HPLC system in our laboratory is a Waters e2695 Separations Module equipped with a 2489 Ultraviolet-Visible (UV-Vis) absorbance detector and a 2475 fluorescence detector controlled with Empower 3 software. The system also has an automated sample injector.</li>
	<li>Inject each sample (e.g., 20 &#956;L) for separation on a Gemini C18, 3 &#956;m 100 &#197;, 4.6 mm x 150 mm column using the program in Table 1 with a flow rate of 0.5 mL/min. The column temperature is 25 &#176;C. Measure the UV/Visible detector absorbance at &#955;393 nm, and fluorescent detection at &#955;ex = 393 nm, &#955;em = 470 nm. The retention time is determined by running a CoA-bimane standard curve before each set of samples.</li>
	<li>Record the area under the CoA-bimane peak for each sample and compare with the standard curve to calculate pmol of CoA-bimane injected onto the HPLC column. The CoA-bimane absorbance standard curve is used for tissue samples, and the CoA-bimane fluorescence standard curve is used for tissue culture samples.</li>
	<li>As the CoA-bimane values represent total CoA for each sample, normalize to the number of viable cells for culture samples, or mg wet weight for tissue samples (Figure 6). Alternately, calculate the total CoA values following normalization to DNA or protein content for each sample. DNA or protein content can be determined separately using sample aliquots that are removed during sample preparation prior to KOH addition.</li>
</ol>

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MWF, CS and SJ wrote the paper, MWF and CS provided the data and figures, COR designed the experiments and critically reviewed the manuscript. SJ and COR are inventors on a pending patent application (PCT/US17/39037) &#34;Small molecule modulators of pantothenate kinases&#34;&#160;held by St. Jude Children&#39;s Research Hospital that covers the PZ-2891 compound referenced in this article.

Here we demonstrate a reliable, step-by-step procedure for quantifying total CoA in cells and animal tissues with a wide range of linear detection that is accessible in many laboratories that have an HPLC with either an absorbance or fluorescence output detector. Alternatively, mass spectrometry is a common technique for evaluating CoA and CoA thioesters, but is not widely available due to the cost of the instrumentation and the specialized knowledge required for development of methodology and interpretation of data. Iso...

Coenzyme A (CoA) is an essential cofactor in all living organisms and is synthesized from pantothenic acid, also called pantothenate (the salt of pantothenic acid) or vitamin B5. CoA is the major intracellular carrier of organic acids, including short-chain acids such as acetate and succinate, branch-chain acids such as propionate and methylmalonate, long-chain fatty acids such as palmitate and oleate, very long-chain fatty acids such as polyunsaturated fatty acids, and xenobiotics such as valproic acid. The organic acid forms a thioester linkage enzymatically with CoA to enable its use as a substrate in over 100 reactions in intermediary metabolism1. CoA thioesters are also allosteric regulators and transcriptional activators. It is now appreciated2 that the cellular total CoA supply is regulated3,4; thus, CoA availability can be limiting, and that CoA deficiencies can be catastrophic, as exemplified by inherited genetic disorders that impact CoA biosynthesis5. Pantothenate kinase catalyzes the first step in CoA biosynthesis (Figure 1) and Pantothenate Kinase Associated Neurodegeneration, called PKAN, is caused by mutations in the PANK2 gene6. CoA synthase, encoded by the COASYN gene, catalyzes the last two steps in CoA biosynthesis (Figure 1) and COASY Protein-Associated Neurodegeneration, called CoPAN, is caused by a mutation in the COASYN gene7. Both PKAN and CoPAN are inherited neurodegenerative diseases associated with iron accumulation in the brain and CoA deficiencies underly the disease pathologies.
Cellular levels of total CoA vary among tissues8 and total CoA can increase or decrease under a variety of physiological, pathological and pharmacological states. Liver CoA increases during fuel switching from the fed to the fasted state9, and liver CoA levels are abnormally high in leptin-deficient obese mice10. Liver CoA decreases in response to chronic ethanol ingestion11. Brain CoA levels in the Pank2 knockout mouse model are depressed during the perinatal period, but later in the adult stage brain CoA content is equivalent to wild-type levels, indicating an adaptive CoA response during development12. Manipulation of tissue CoA content by transgenesis or gene delivery methods impacts metabolic and neural functions13,14,15. Preclinical development of potential therapies for PKAN or CoPAN includes cell or tissue CoA measurements as indicators of efficacy16,17,18,19,20. Evaluation of all of these conditions and their metabolic or functional consequences requires a quantitative method for measurement of total CoA.
An accurate, reliable assay for measuring CoA in biological samples is a technical challenge in many labs. Unfortunately, there are no probes available to evaluate or quantify CoA or CoA thioesters in intact cells, although analogs of natural CoA thioesters have been widely used as mechanistic probes in studies of CoA ester utilizing enzymes21. The conversion of CoA, with a free sulfhydryl (-SH) moiety, to a CoA thioester (or vice versa) is rapid in cells or animal tissues during transfer to a different environment and during cell lysis. Numerous acyl-CoA synthetases and acyl-CoA thioesterases in cells mediate the interconversions within the CoA pool, and additional enzymes that utilize CoA thioesters as substrates remain active in biological samples until quenched by chemical or physical means. The off-loading of acyl-groups from CoA to carnitine by acyl-transferases is one example within the network of reactions that can alter the CoA/CoA thioester distribution. Radioactive tracers can be used to measure rates of CoA synthesis in cells. Current methods for measuring CoA and CoA derivatives in biological samples have been reviewed22 and include coupled enzymatic spectrophotometric assays, high-pressure liquid chromatography and mass spectrometry-based procedures. However, these methods are often focused on particular CoA molecular species and are blind to variation of the total CoA pool. The coupled enzymatic assays generally require larger amounts of input material due to low detection sensitivities and have a limited range of linearity.
Our laboratory has developed a reliable procedure for quantification of total CoA in cultured cells and animal tissues. The strategy includes hydrolysis of all CoA thioesters to yield only free CoA during sample preparation, rather than making efforts to maintain and analyze the entire spectrum of CoA species. The procedure is a compilation of individual published methods for sample preparation, CoA derivatization, purification and identification following high-pressure liquid chromatography (HPLC), and quantification of the derivatized CoA by absorbance or fluorescence detection23,24,25. The CoA determinations obtained using this procedure have enabled our understanding of CoA regulation and the development of a therapeutic approach for treatment of CoA deficiencies.

<table>
	<tbody>
		<tr>
			<td>2-(2-pyridyl)-ethyl silica gel SPE column</td>
			<td>Millipore-Sigma</td>
			<td>54127-U</td>
			<td></td>
		</tr>
		<tr>
			<td>coenzyme A</td>
			<td>Avanti Polar Lipids</td>
			<td>870700</td>
			<td></td>
		</tr>
		<tr>
			<td>Gemini C18 3 &#956;m 100 &#197; HPLC column</td>
			<td>Phenomenex</td>
			<td>00F-4439-E0</td>
			<td></td>
		</tr>
		<tr>
			<td>monobromobimane</td>
			<td>ThermoFisher Scientific</td>
			<td>M-1378</td>
			<td></td>
		</tr>
		<tr>
			<td>Omni-Tip probe tissue disrupter</td>
			<td>Omni International</td>
			<td>32750H</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Fisher</td>
			<td>S37440</td>
			<td></td>
		</tr>
		<tr>
			<td>PowerGen 125 motorized rotor stator homogenizer</td>
			<td>ThermoFisher Scientific</td>
			<td>NC0530997</td>
			<td></td>
		</tr>
		<tr>
			<td>Spin-X centrifuge tube filter</td>
			<td>CoStar</td>
			<td>8161</td>
			<td></td>
		</tr>
		<tr>
			<td>Trizma-HCl</td>
			<td>Fisher</td>
			<td>T395-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Waters 2475 fluorescence detector</td>
			<td>Waters</td>
			<td>2475</td>
			<td></td>
		</tr>
		<tr>
			<td>Waters 2489 UV-Vis detector</td>
			<td>Waters</td>
			<td>2489</td>
			<td></td>
		</tr>
		<tr>
			<td>Waters e2695 separations module</td>
			<td>Waters</td>
			<td>e2695</td>
			<td></td>
		</tr>
	</tbody>
</table>

quantification of coenzyme a in cells and tissues

Emerging research has revealed that the cellular coenzyme A (CoA) supply can become limiting with a detrimental impact on growth, metabolism and survival. Measurement of cellular CoA is a challenge due to its relatively low abundance and the dynamic conversion of free CoA to CoA thioesters that, in turn, participate in numerous metabolic reactions. A method is described that navigates through potential pitfalls during sample preparation to yield an assay with a broad linear range of detection that is suitable for use in many biomedical laboratories.

A relatively fast and reliable method for the detection of total CoA in cultured cells and tissues has been developed by derivatizing the thiol of CoA to a fluorescent agent using mBBr, and then purifying the derivatized CoA-bimane using reverse phase HPLC. A standard curve is first generated, where known and increasing amounts of the CoA-bimane standard are injected individually and the areas under the peaks in the CoA-bimane chromatograms are plotted as a function of the input CoA-bimane (Figure 4<...

Watch this Scientific Journal Video about Quantification of Coenzyme A in Cells and Tissues at JoVE.com

Quantification of Coenzyme A in Cells and Tissues

This method describes sample preparation from cultured cells and animal tissues, extraction and derivatization of coenzyme A in the samples, followed by high pressure liquid chromatography for purification and quantification of the derivatized coenzyme A by absorbance or fluorescence detection.

Emerging research has revealed that the cellular coenzyme A (CoA) supply can become limiting with a detrimental impact on growth, metabolism and survival. ...

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Biochemistry

8.2K Views.  St. Jude Children's Research Hospital. This method describes sample preparation from cultured cells and animal tissues, extraction and derivatization of coenzyme A in the samples, followed by high pressure liquid chromatography for purification and quantification of the derivatized coenzyme A by absorbance or fluorescence detection.

Video: Quantification of Coenzyme A in Cells and Tissues

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies have revealed that these reversible RNA modifications affect RNA splicing, translation, degradation, and localization. Multiple physiological processes, like circadian rhythms, stem cell pluripotency, fibrosis, triglyceride metabolism, and obesity are also controlled by m6A modifications. Immunoprecipitation/sequencing, mass spectrometry, and modified northern blotting are some of the methods commonly employed to measure m6A modifications. Herein, we present a northeastern blotting technique for measuring m6A modifications. The current protocol provides good size separation of RNA, better accommodation and standardization for various experimental designs, and clear delineation of m6A modifications in various sources of RNA. While m6A modifications are known to have a crucial impact on human physiology relating to circadian rhythms and obesity, their roles in other (patho)physiological states are unclear. Therefore, investigations on m6A modifications have immense possibility to provide key insights into molecular physiology.

A Method for Measuring RNA N6-methyladenosine Modifications in Cells and Tissues

A modified northern blotting method for measuring N6-methyladenosine (m6A) modifications in RNA is described. The current method can detect modifications in diverse RNAs and controls under various experimental designs.

N6-Methyladenosine (m6A) modifications of RNA are diverse and ubiquitous amongst eukaryotes. They occur in mRNA, rRNA, tRNA, and microRNA. Recent studies ...

Quantification of Bacterial Histidine Kinase Autophosphorylation Using a Nitrocellulose Binding Assay

We demonstrate a useful method for quantifying autophosphorylation of purified bacterial histidine kinases. Histidine kinases are known for their involvement in two-component signal transduction, a ubiquitous system through which bacteria sense and respond to environmental stimuli. Two-component signaling features autophosphorylation of a histidine kinase, followed by phosphotransfer to the receiver domain of a response regulator protein, which ultimately leads to an output response. Autophosphorylation of the histidine kinase is responsive to the presence of a cognate environmental stimulus, thereby giving bacteria a means to detect and respond to changes in the environment. Despite their importance in bacterial biology, histidine kinases remain poorly understood due to the inherent lability of phosphohistidine. Conventional methods for studying these proteins, such as SDS-PAGE autoradiography, have significant shortcomings. We have developed a nitrocellulose binding assay that can be used to characterize histidine kinases. The protocol for this assay is simple and easy to execute. Our method is higher throughput, less time-consuming, and offers a greater dynamic range than SDS-PAGE autoradiography.

We report and demonstrate an optimized nitrocellulose binding assay that can be used to quantify autophosphorylation of purified bacterial histidine kinases. Our method has several advantages over traditional SDS-PAGE based techniques, providing a valuable alternative for characterizing these important proteins.

We demonstrate a useful method for quantifying autophosphorylation of purified bacterial histidine kinases. Histidine kinases are known for their ...

Quantification of the Abundance and Charging Levels of Transfer RNAs in Escherichia coli

Transfer RNA (tRNA) is an essential part of the translational machinery in any organism. tRNAs bind and transfer amino acids to the translating ribosome. The relative levels of different tRNAs, and the ratio of aminoacylated tRNA to total tRNA, known as the charging level, are important factors in determining the accuracy and speed of translation. Therefore, the abundance and charging levels of tRNAs are important variables to measure when studying protein synthesis, for example under various stress conditions. Here, we describe a method for harvesting tRNA and directly measuring both the relative abundance and the absolute charging level of specific tRNA species in Escherichia coli. The tRNA is harvested in such a way that the labile bond between the tRNA and its amino acid is preserved. The RNA is then subjected to gel electrophoresis and Northern blotting, which results in separation of the charged and uncharged tRNAs. The levels of specific tRNAs in different samples can be compared due to the addition of spike-in cells for normalization. Prior to RNA purification, we add 5% of E. coli cells that overproduce the rare tRNAselC to each sample. The amount of the tRNA species of interest in a sample is then normalized to the amount of tRNAselC in the same sample. Addition of spike-in cells prior to RNA purification has the advantage over addition of purified spike-in RNAs that it also accounts for any differences in cell lysis efficiency between samples.

Quantification of the Abundance and Charging Levels of Transfer RNAs in Escherichia coli

Here we present a method for directly measuring transfer RNA charging levels from purified Escherichia coli RNA as well as a way to compare relative levels of transfer RNA, or any other short RNA, across different samples based on the addition of spike-in cells expressing a reference gene.

Transfer RNA (tRNA) is an essential part of the translational machinery in any organism. tRNAs bind and transfer amino acids to the translating ribosome. ...

Quantification of Hypopigmentation Activity In Vitro

This study presents laboratory methods for the quantification of hypopigmentation activity in vitro. Melanin, the major pigment in melanocytes, is synthesized in response to multiple cellular and environmental factors. Melanin protects skin cells from ultraviolet damage, but also has biophysical and biochemical functions. Excessive production or accumulation of melanin in melanocytes can cause dermatological problems, such as freckles, dark spots, melasma, and moles. Therefore, the control of melanogenesis with hypopigmentation agents is important in individuals with clinical or cosmetic needs. Melanin is primarily synthesized in the melanosomes of melanocytes in a complex biochemical process called melanogenesis, which is influenced by extrinsic and intrinsic factors, such as hormones, inflammation, age, and ultraviolet light exposure. We describe three methods to determine the hypopigmentation activity of chemicals or natural substances in melanocytes: measurement of the 1) cellular tyrosinase activity and 2) melanin content, and 3) staining and quantifying cellular melanin with image analysis.
In melanogenesis, tyrosinase catalyzes the rate-limiting step that converts L-tyrosine into 3,4-dihydroxyphenylalanine (L-DOPA) and then into dopaquinone. Therefore, the inhibition of tyrosinase is a primary hypopigmentation mechanism. In cultured melanocytes, tyrosinase activity can be quantified by adding L-DOPA as a substrate and measuring dopaquinone production by spectrophotometry. Melanogenesis can also be measured by quantifying the melanin content. The melanin-containing cellular fraction is extracted with NaOH and melanin is quantified spectrophotometrically. Finally, the melanin content can be quantified by image analysis following Fontana-Masson staining of melanin. Although the results of these in vitro assays may not always be reproduced in human skin, these methods are widely used in melanogenesis research, especially as the initial step to identify potential hypopigmentation activity. These methods can also be used to assess melanocyte activity, growth, and differentiation. Consistent results with the three different methods ensure the validity of the effects.

We describe three experimental methods for evaluating the hypopigmentation activity of chemicals in vitro: quantification of 1) cellular tyrosinase activity and 2) melanin content, and 3) measurement of melanin by cellular melanin staining and image analysis.

This study presents laboratory methods for the quantification of hypopigmentation activity in vitro. Melanin, the major pigment in melanocytes, is ...

Dissipative Microgravimetry to Study the Binding Dynamics of the Phospholipid Binding Protein Annexin A2 to Solid-supported Lipid Bilayers Using a Quartz Resonator

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic properties of the absorbed/immobilized mass on sensor surfaces, allowing the measurements of the interaction of proteins with solid-supported surfaces, such as lipid bilayers, in real-time and with a high sensitivity. Annexins are a highly conserved group of phospholipid-binding proteins that interact reversibly with the negatively charged headgroups via the coordination of calcium ions. Here, we describe a protocol that was employed to quantitatively analyze the binding of annexin A2 (AnxA2) to planar lipid bilayers prepared on the surface of a quartz sensor. This protocol is optimized to obtain robust and reproducible data and includes a detailed step-by-step description. The method can be applied to other membrane-binding proteins and bilayer compositions.

Here, we present an experimental protocol that can be employed to determine the binding affinities and mode of interaction of label-free phospholipid-binding protein annexin A2 with immobilized solid-supported bilayers (SLB) by simultaneously measuring the mass uptake and the viscoelastic properties of the protein annexin A2.

The dissipative quartz crystal microbalance technique is a simple and label-free approach to measure simultaneously the mass uptake and viscoelastic ...

Quantification of Protein Interaction Network Dynamics using Multiplexed Co-Immunoprecipitation

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic interactions among multiple proteins in a protein interaction network is technically difficult. Here, we present a protocol for Quantitative Multiplex Immunoprecipitation (QMI), which allows quantitative assessment of fold changes in protein interactions based on relative fluorescence measurements of Proteins in Shared Complexes detected by Exposed Surface epitopes (PiSCES). In QMI, protein complexes from cell lysates are immunoprecipitated onto microspheres, and then probed with a labeled antibody for a different protein in order to quantify the abundance of PiSCES. Immunoprecipitation antibodies are conjugated to different MagBead spectral regions, which allows a flow cytometer to differentiate multiple parallel immunoprecipitations and simultaneously quantify the amount of probe antibody associated with each. QMI does not require genetic tagging and can be performed using minimal biomaterial compared to other immunoprecipitation methods. QMI can be adapted for any defined group of interacting proteins, and has thus far been used to characterize signaling networks in T cells and neuronal glutamate synapses. Results have led to new hypothesis generation with potential diagnostic and therapeutic applications. This protocol includes instructions to perform QMI, from the initial antibody panel selection through to running assays and analyzing data. The initial assembly of a QMI assay involves screening antibodies to generate a panel, and empirically determining an appropriate lysis buffer. The subsequent reagent preparation includes covalently coupling immunoprecipitation antibodies to MagBeads, and biotinylating probe antibodies so they can be labeled by a streptavidin-conjugated fluorophore. To run the assay, lysate is mixed with MagBeads overnight, and then beads are divided and incubated with different probe antibodies, and then a fluorophore label, and read by flow cytometry. Two statistical tests are performed to identify PiSCES that differ significantly between experimental conditions, and results are visualized using heatmaps or node-edge diagrams.

Quantitative Multiplex Immunoprecipitation (QMI) uses flow cytometry for sensitive detection of differences in the abundance of targeted protein-protein interactions between two samples. QMI can be performed using a small amount of biomaterial, does not require genetically engineered tags, and can be adapted for any previously defined protein interaction network.

Dynamic protein-protein interactions control cellular behavior, from motility to DNA replication to signal transduction. However, monitoring dynamic ...

Extracellular Glucose Depletion as an Indirect Measure of Glucose Uptake in Cells and Tissues Ex Vivo

The ongoing worldwide epidemic of diabetes increases the demand for the identification of environmental, nutritional, endocrine, genetic, and epigenetic factors affecting glucose uptake. The measurement of intracellular fluorescence is a widely used method to test the uptake of fluorescently-labeled glucose (FD-glucose) in cells in vitro, or for imaging glucose-consuming tissues in vivo. This assay assesses glucose uptake at a chosen time point. The intracellular analysis assumes that the metabolism of FD-glucose is slower than that of endogenous glucose, which participates in catabolic and anabolic reactions and signaling. However, dynamic glucose metabolism also alters uptake mechanisms, which would require kinetic measurements of glucose uptake in response to different factors. This article describes a method for measuring extracellular FD-glucose depletion and validates its correlation with intracellular FD-glucose uptake in cells and tissues ex vivo. Extracellular glucose depletion may be potentially applicable for high-throughput kinetic and dose-dependent studies, as well as identifying compounds with glycemic activity and their tissue-specific effects.

Extracellular Glucose Depletion as an Indirect Measure of Glucose Uptake in Cells and Tissues Ex Vivo

Extracellular depletion of fluorescently labeled glucose correlates with glucose uptake and could be used for high-throughput screening of glucose uptake in excised organs and cell cultures.

The ongoing worldwide epidemic of diabetes increases the demand for the identification of environmental, nutritional, endocrine, genetic, and epigenetic ...

High-Throughput Quantification of the Synthesis and Distribution of mtDNA

High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells produce most of their ATP in the mitochondria by oxidative phosphorylation. Mitochondria are unique organelles because they have their own genome that is replicated and passed on to the next generation of cells. In contrast to the nuclear genome, there are multiple copies of the mitochondrial genome in the cell. The detailed study of the mechanisms responsible for the replication, repair, and maintenance of the mitochondrial genome is essential for understanding the proper functioning of mitochondria and whole cells under both normal and disease conditions. Here, a method that allows the high-throughput quantification of the synthesis and distribution of mitochondrial DNA (mtDNA) in human cells cultured in vitro is presented. This approach is based on the immunofluorescence detection of actively synthesized DNA molecules labeled by 5-bromo-2'-deoxyuridine (BrdU) incorporation and the concurrent detection of all the mtDNA molecules with anti-DNA antibodies. Additionally, the mitochondria are visualized with specific dyes or antibodies. The culturing of cells in a multi-well format and the utilization of an automated fluorescence microscope make it easier to study the dynamics of mtDNA and the morphology of mitochondria under a variety of experimental conditions in a relatively short time.

Author Spotlight: High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution  

A procedure for studying the dynamics of mitochondrial DNA (mtDNA) metabolism in cells using a multi-well plate format and automated immunofluorescence imaging to detect and quantify mtDNA synthesis and distribution is described. This can be further used to investigate the effects of various inhibitors, cellular stresses, and gene silencing on mtDNA metabolism.

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells ...

Smartphone as an Analytical Device to Quantify Protein Using Bradford Assay

RGBradford: Protein Quantitation with a Smartphone Camera

Protein quantitation is an essential procedure in life sciences research. Amongst several other methods, the Bradford assay is one of the most used. Because of its widespread, the limitations and advantages of the Bradford assay have been exhaustively reported, including several modifications of the original method to improve its performance. One of the alterations of the original method is the use of a smartphone camera as an analytical instrument. Taking advantage of the three forms of the Coomassie Brilliant Blue dye that exist in the conditions of the Bradford assay, this paper describes how to accurately quantify protein in samples using color data extracted from a single picture of a microplate. After performing the assay in a microplate, a picture is taken using a smartphone camera, and RGB color data is extracted from the picture using a free and open-source image analysis software application. Then, the ratio of blue to green intensity (in the RGB scale) of samples with unknown concentrations of protein is used to calculate the protein content based on a standard curve. No significant difference is observed between values calculated using RGB color data and those calculated using conventional absorbance data.

Author Spotlight: An Alternative Approach to Protein Quantification by Bradford Assay Using a Smartphone 

This paper provides a protocol for protein quantification using the Bradford assay and a smartphone as an analytical device. Protein levels in samples can be quantified using color data extracted from a picture of a microplate taken with a smartphone.

Protein quantitation is an essential procedure in life sciences research. Amongst several other methods, the Bradford assay is one of the most used. ...

Quantitative Microtubule Fractionation Technique to Separate Stable Microtubules, Labile Microtubules, and Free Tubulin in Mouse Tissues

Microtubules, composed of &#945;/&#946;-tubulin dimers, are a crucial component of the cytoskeleton in eukaryotic cells. These tube-like polymers exhibit dynamic instability as tubulin heterodimer subunits undergo repetitive polymerization and depolymerization. Precise control of microtubule stability and dynamics, achieved through tubulin post-translational modifications and microtubule-associated proteins, is essential for various cellular functions. Dysfunctions in microtubules are strongly implicated in pathogenesis, including neurodegenerative disorders. Ongoing research focuses on microtubule-targeting therapeutic agents that modulate stability, offering potential treatment options for these diseases and cancers. Consequently, understanding the dynamic state of microtubules is crucial for assessing disease progression and therapeutic effects.
Traditionally, microtubule dynamics have been assessed in vitro or in cultured cells through rough fractionation or immunoassay, using antibodies targeting post-translational modifications of tubulin. However, accurately analyzing tubulin status in tissues using such procedures poses challenges. In this study, we developed a simple and innovative microtubule fractionation method to separate stable microtubules, labile microtubules, and free tubulin in mouse tissues.
The procedure involved homogenizing dissected mouse tissues in a microtubule-stabilizing buffer at a 19:1 volume ratio. The homogenates were then fractionated through a two-step ultracentrifugation process following initial slow centrifugation (2,400 &#215; g) to remove debris. The first ultracentrifugation step (100,000 &#215; g) precipitated stable microtubules, while the resulting supernatant was subjected to a second ultracentrifugation step (500,000 &#215; g) to fractionate labile microtubules and soluble tubulin dimers. This method determined the proportions of tubulin constituting stable or labile microtubules in the mouse brain. Additionally, distinct tissue variations in microtubule stability were observed that correlated with the proliferative capacity of constituent cells. These findings highlight the significant potential of this novel method for analyzing microtubule stability in physiological and pathological conditions.

Microtubules, which are tubulin polymers, play a crucial role as a cytoskeleton component in eukaryotic cells and are known for their dynamic instability. This study developed a method for fractionating microtubules to separate them into stable microtubules, labile microtubules, and free tubulin to evaluate the stability of microtubules in various mouse tissues.

Microtubules, composed of &#945;/&#946;-tubulin dimers, are a crucial component of the cytoskeleton in eukaryotic cells. These tube-like polymers exhibit ...