We are thankful to Dr Stephen Robinson and Ms Hania Czerwinska (Monash University) for the generous supply of astrocyte cultures.

1. Determine Intracellular Ascorbate in Cultured Cells
<ol>
	<li>Cell culture and harvesting
	<ol>
		<li>Grow suspension (e.g.&#160;human erythroleukemia, K562) or adherent cells (e.g.&#160;primary astrocytes) using standard culture procedures19-21,32,38. Note: to ensure cells contain ascorbate, load cultured cells with ascorbate either as ascorbate or DHA33,39.</li>
	</ol>
	</li>
	<li>Create an ascorbate-containing cellular extract
	<ol>
		<li>Incubate an appropriate number of phosphate-buffered saline (PBS)-washed suspension or adherent cells with 450 &#956;l of ice-cold Cell Permeabilization Buffer [CPB; 0.1% (w/v) saponin in PBS; 4 &#176;C]. Note: Determine the number of cells required to obtain an absorbance value within the linear range of the assay (see below) empirically by the end-user. Note: tissue extracts may also be used (see Discussion for further details).</li>
		<li>Agitate cells on ice for 10 min to ensure thorough cellular lysis.</li>
		<li>Create a clarified ascorbate-containing intracellular extract by removing cellular debris by centrifugation of the crude lysate at 16,000 &#215; g for 5 min in a refrigerated microcentrifuge (4 &#176;C).</li>
		<li>Carefully remove 4 x 100 &#956;l aliquots from each sample and add to a horizontal sequence of wells in a 96-well flat bottom plate that contains either 25 &#956;l/well of PBS (&#34;-AO&#34;) only or 25 &#956;l/well of 45.5 U/ml stock solution of L-ascorbate-oxidase (AO) in PBS (&#34;+AO&#34;). Note: this should be done in parallel with the preparation of the standards (see below).</li>
	</ol>
	</li>
	<li>Ascorbate standard-curve construction (must be constructed anew for each assay)
	<ol>
		<li>Prepare a stock solution of 10 mM ascorbic acid in ice-cold PBS. Note: Verify the concentration of ascorbate spectrophotometrically using a quartz cuvette with a 1 cm path-length at 265 nm (extinction coefficient = 14.5 mM-1 cm-1)40.</li>
		<li>Carefully prepare a series of ascorbate standards between 0 and 20 &#956;M.</li>
		<li>In parallel with step 1.2.4, carefully remove 4 &#215; 100 &#956;l aliquots from each standard and add to a horizontal sequence of wells in a 96-well flat bottom plate that contains 25 &#956;l/well of PBS (&#34;-AO&#34;) or a 45.5 U/ml stock solution of AO in PBS (&#34;+AO&#34;). Note: 100 &#956;l of the above ascorbate standards will contain 0-2 nmole of ascorbate. Please note, the purpose of the AO is to control for the contribution of non-ascorbate reductants to ferricyanide reduction.</li>
	</ol>
	</li>
	<li>Determine intracellular ascorbate - Step 1: selectively remove ascorbate and quantitatively oxidize ascorbate with ferricyanide
	<ol>
		<li>Orbitally mix the 96-well plate at 550 rpm at room temperature for 5 min in the dark by covering the plate in foil. Note: This step should oxidize all ascorbate in the &#34;+ AO&#34; wells to DHA. The &#34;-AO&#34; wells should be unaffected.</li>
		<li>Add 50 &#956;l of 3.5 mM potassium ferricyanide (herein referred to as &#34;ferricyanide&#34;) in PBS to all wells. The final [ferricyanide] = 1 mM. Note: Use a multipipettor.</li>
		<li>Orbitally mix the plate at 550 rpm at room temperature for another 5 min in the dark. Note: This step should result in the reduction of ferricyanide to ferrocyanide by ascorbate in a stoichiometric ratio of 2 molecules of ferricyanide reduced per 1 molecule of ascorbate oxidized.</li>
		<li>Immediately add 25 &#956;l of a freshly constructed solution containing 50% (v/v) acetic acid and 30% (w/v) trichloroacetic acid (TCA). Note: Use a multipipettor.</li>
	</ol>
	</li>
	<li>Determine intracellular ascorbate - Step 2: quantitate the amount of ferrocyanide formed37
	<ol>
		<li>Add 100 &#956;l of ferrocyanide determination solution37 to each well. Note: A working solution should be made immediately before use: 2 ml of 3 M Na-acetate (pH 6.0); 0.5 ml of glacial acetic acid (~17.4 M acetic acid); 2 ml of 0.2 M citric acid; 2 ml of 3.3 mM FeCl3 in 0.1 M acetic acid; 1 ml of 30 mM ferene-S. The final volume of this working solution should be 7.5 ml.</li>
		<li>Orbitally mix the plate in the dark at 550 rpm for 30 min at room temperature.</li>
		<li>Read the absorbance values of the wells at 593 nm (i.e.&#160;the absorbance maximum of the Fe(II)(ferene-S)3 complex).</li>
		<li>Calculate the amount of intracellular ascorbate as nmoles ascorbate per million cells by initially subtracting the A593 nm values for the &#39;+ AO&#39; wells from the corresponding &#39;- AO&#39; wells for each sample and then interpolating from the ascorbate standard curve (see step 1.3). When constructing the standard curve, plot this &#34;difference value&#34; for each standard against the amount of ascorbate per well.</li>
	</ol>
	</li>
</ol>
2. Determination of Ascorbate-efflux from Cultured Cells
<ol>
	<li>Cell culture and harvesting
	<ol>
		<li>Grow suspension or adherent cells as above (see step 1.1). Note: carry out the below assay in a 24-well plat format with suspension and adherent cells for best results.</li>
		<li>Resuspend suspension cells, or overlay adherent cells, with 400 &#956;l of pre-warmed HEPES-buffered saline solution, with or without calcium and magnesium, and containing 5 mM D-glucose (HBS/D; pH 7.3, 37 &#176;C) in wells of a 24-well plate. Add the same volume of HBS/D to specified cell-free wells on each 24-well plate to be examined. Note: the latter will serve as cell-free controls for the baseline reaction (see below).</li>
	</ol>
	</li>
</ol>
3. Determine the Amount of Ascorbate Released
<ol>
	<li>The following stock solutions should be prepared beforehand: 120 U/ml AO in HBS/D (prepare fresh); 2.4 mM Ferene-S in HBS/D; 120 &#956;M FeCl3 and 600 &#956;M Na-citrate in HBS/D (prepare immediately from more concentrated stock solutions).</li>
	<li>To start the ferrireduction reaction (final volume per well should be 600 &#956;l), add the following volumes of reagents to individual wells already containing 400 &#956;l of HBS/D:
	<ol>
		<li>Add 50 &#956;l of AO (120 U/ml) or HBS/D to paired wells in triplicate. The final AO concentration should be 10 U/ml. Note: Label paired wells as &#34;- AO&#34; and &#34;+ AO&#34;.</li>
		<li>Mix the plates with gentle orbital mixing for 5 min at 37 &#176;C.</li>
		<li>Add 50 &#956;l of 2.4 mM ferene-S to all wells. The final ferene-S concentration should be 200 &#956;M. Mix the plates as above for 5 min at 37 &#176;C.</li>
		<li>Add 50 &#956;l of freshly prepared 120 &#956;M ferric citrate. The final iron and citrate concentrations should be 10 &#956;M iron and 50 &#956;M citrate. Mix well.</li>
	</ol>
	</li>
	<li>In three triplicate control wells, aspirate the overlying medium and add 600 &#956;l HBS/D containing 0.1% saponin. Note: These will serve as &#34;100% cell-lysis&#34; controls for a lactate dehydrogenase (LDH) release assay to be conducted in parallel (see below).</li>
	<li>Incubate for 60 min in the dark at 37 &#176;C.</li>
	<li>At the end of the ascorbate-efflux assay, rapidly aspirate 500 &#956;l from each well and add to appropriately labelled wells in a 24-well plate. Note: for suspension cells, initially remove the cells by centrifugation at 4 &#176;C.</li>
	<li>Add 300 &#956;l aliquots of the supernatant to a 96-well plate and then read at 593 nm.</li>
</ol>
4. Determination of Extracellular Ascorbate
<ol>
	<li>Read the absorbance values of the wells at 593 nm.</li>
	<li>Calculate the amount of extracellular ascorbate as nmoles ascorbate per mg protein (or per million cells) as described for the determination of intracellular ascorbate in Step 1.5.4. Note: the end user should optimize conditions so that ascorbate release is not limited by intracellular ascorbate.</li>
</ol>
5. Determination of LDH Release
<ol>
	<li>With the remaining 200 &#956;l of extracellular solution from each sample conduct an LDH release assay32,41 to determine the extent of cellular lysis. Note: calculate as a % of total releasable LDH. Determine releasable LDH from the samples that were treated with 0.1% saponin.</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+electron+spin+resonance+spectra+of+radical+intermediates+in+the+oxidation+of+ascorbic+acid+and+related+substances.">The electron spin resonance spectra of radical intermediates in the oxidation of ascorbic acid and related substances.</a> J. Am. Chem. Soc. 94, 9062-9073 (1972).</li><li>Dringen, R., Kussmaul, L., Hamprecht, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detoxification+of+exogenous+hydrogen+peroxide+and+organic+hydroperoxides+by+cultured+astroglial+cells+assessed+by+microtiter+plate+assay.">Detoxification of exogenous hydrogen peroxide and organic hydroperoxides by cultured astroglial cells assessed by microtiter plate assay.</a> Brain Res. Brain Res. Protoc. 2, 223-228 (1998).</li><li>Lane, D. J. R., Lawen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transplasma+membrane+electron+transport+comes+in+two+flavors.">Transplasma membrane electron transport comes in two flavors.</a> Biofactors. 34, 191-200 (2009).</li><li>Lin, S., Lin, D. C., Flanagan, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Specificity+of+the+effects+of+cytochalasin+B+on+transport+and+motile+processes.">Specificity of the effects of cytochalasin B on transport and motile processes.</a> Proc. Natl. Acad. Sci. USA. 75, 329-333 (1978).</li><li>May, J. M., Qu, Z. C., Juliao, S., Cobb, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ascorbic+acid+decreases+oxidant+stress+in+endothelial+cells+caused+by+the+nitroxide+tempol.">Ascorbic acid decreases oxidant stress in endothelial cells caused by the nitroxide tempol.</a> Free Radic. Res. 39, 195-202 (2005).</li><li>Avron, M., Shavit, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+sensitive+and+simple+method+for+determination+of+ferrocyanide.">A sensitive and simple method for determination of ferrocyanide.</a> Anal. Biochem. 6, 549-554 (1963).</li></ol>

The authors have nothing to disclose.

In this paper we present two rapid, specific and relatively sensitive colorimetric microplate assays for the determination of ascorbate derived from the intra- and extracellular compartments in cultured cells. The assays can be completed with access to standard laboratory equipment and reagents. The only moderately costly reagent required for the assay is AO, which is essential as it imparts a high-degree of analyte specificity toward L-ascorbate. The assays are well suited to either suspen...

The discovery of the chemical nature of ascorbic acid (vitamin C), and its identification as the long-sought &#34;anti-scorbutic factor&#34;, by Albert Szent-Gy&#246;rgyi and others in papers published from 1928 to 19341 were landmark events in the history of biochemistry. Indeed, these discoveries contributed to Szent-Gy&#246;rgyi being awarded the Nobel Prize in Physiology or Medicine in 1937. The ever-expanding suite of roles for ascorbate in animal and plant physiology, as well as human health, continue to be the subjects of active scientific investigation and controversy.
L-Ascorbate is an abundant physiological reductant and enzyme cofactor in mammalian systems, and contributes to numerous well-defined enzymatic reactions involving collagen hydroxylation, carnitine and norepinephrine biosynthesis, tyrosine metabolism and peptide hormone amidation2. Intriguingly, mounting evidence suggests that ascorbate plays a role in stimulating other iron-dependent dioxygenases, such as the prolyl and asparaginyl hydroxylases involved in the hydroxylation and targeting of the hypoxia-inducible factors (HIFs) 1&#945; and 2&#945;3. A recent report suggests that ascorbate plays a role in T-cell maturation through affecting chromatin demethylation via its activity in stimulating the nuclear hydroxylases, Jumonji C (JmjC) domain proteins; the latter of which appear to require ascorbate for full activity4. Indeed, the stimulation of such enzymes by ascorbate appears to occur by a similar mechanism to the stimulation by ascorbate of the HIF and collagen hydroxylases. Among other classical effects, ascorbate contributes significantly to cellular antioxidation as a water-soluble chain-breaking radical scavenger5 and to the recycling of plasma membrane &#945;-tocopherol (vitamin E) via the reduction of the &#945;-tocopheroxyl radical6, which is important in protecting against membrane lipid peroxidation7. Importantly, although most mammals are capable of de novo hepatic synthesis of ascorbate from D-glucose, higher primates, guinea pigs and some bats depend on dietary sources of the vitamin8. This is due to inactivation of the GULO gene, the orthologues of which in unaffected mammals encode the enzyme, &#947;-gulono-lactone oxidase9-13. This enzyme is required for the final reaction in ascorbate biosynthesis from glucose13.
Following transporter-mediated absorption from the intestinal lumen in humans, ascorbate is distributed throughout the body by the circulatory system. The vitamin is typically found in its reduced form at millimolar concentrations intracellularly (with the notable exception of erythrocytes in which concentrations are typically similar to the prevailing plasma concentration), and at micromolar concentrations (e.g.&#160;50-200 &#956;M) in most extracellular fluids14,15 .
Under physiological conditions, ascorbate typically undergoes a reversible one-electron oxidation to the ascorbyl free radical (AFR; also known as monodehydroascorbate or semidehydroascorbate). While the AFR is a relatively stable radical16, in the absence of its rapid one-electron enzymatic reduction back to ascorbate, two AFRs can further dismutate to one ascorbate and one dehydroascorbate (DHA)9,13,17. Within the interior of the cell, the two-electron oxidation product of ascorbate, DHA, can be rapidly reduced back to ascorbate by glutathione- and NAD(P)H-dependent enzymatic and non-enzymatic reactions13.
While it is classically accepted that ascorbate&#8217;s only significant role in iron metabolism is to stimulate dietary absorption of non-heme iron18, we and others have provided evidence strongly suggesting that ascorbate plays a greatly expanded role in the metabolism of this metal. First, ascorbate that is released by ascorbate-replete cells appears to play an important role in modulating the uptake of non-transferrin-bound iron by cells19,20, and very recent evidence indicates that ascorbate also modulates the uptake of transferrin-bound iron by cells21, the latter of which corresponds to a major physiological iron-uptake route22.
Ascorbate is essential for normal central nervous system function in mammals23,24. Together with the adrenal cortex, pituitary gland, thymus, retina and corpus luteum, the brain contains high concentrations of ascorbate relative to other body tissues23,25-27. Additionally, the exposure of both astrocytes28,29 and neuron-like cells30 to glutamate is known to trigger the release of ascorbate into the extracellular space, where the ascorbate is thought to help protect neurons against glutamate-induced neuronal dysfunction31. While the exact mechanism of glutamate-induced ascorbate release from astrocytes is unknown, we have recently provided evidence indicating the involvement of cell swelling caused by glutamate uptake by the astrocyte glutamate and aspartate transporter (GLAST; also known excitatory amino acid transporter isoform 1 [EAAT1] in humans) and consequent activation of volume-sensitive osmolyte and anion channels (VSOACs) that are permeable to small organic anions such as ascorbate32. The molecular identities of the plasma membrane conduits involved in VSOAC formation remain to be identified33,34.
Although many assays have been developed for the determination of ascorbate in biological samples, which include spectrophotometric, fluorometric and chromatographic assays35,36, there is much variability in specificity, sensitivity, interference by chemical contaminants, effective linear range and stability of the endpoint analyte. Additionally, other significant factors that influence the choice of assay are rapidity, ease of use and access to relatively specialized equipment such as a high-performance liquid chromatography (HPLC) apparatus.
Here we present a simple and highly specific colorimetric microplate assay for the determination of intracellular ascorbate in cultured cells, as well as a separate assay for the determination of ascorbate-efflux from cultured cells. The latter assay aims to circumvent the problem of underestimation of ascorbate release from cells due to rapid re-uptake of released ascorbate by sodium-dependent ascorbate transporters (SVCTs). Although both of these methods have appeared in some of our previous publications19,20,32,37,38, this manuscript provides an explicit set of instructions and guidelines for their effective execution.

<table><tbody><tr><td>Nunc 96-well flat-bottom plates</td><td>Thermo</td><td>269620</td><td>Any flat-bottom 96-well plate can be used</td></tr><tr><td>Refrigerated benchtop microcentrifuge</td><td>Eppendorf</td><td>5415D</td><td>A non-refrigerated microcentrifuge that has been equilibrated to temperature in a cold room can also be used</td></tr><tr><td>Refrigerated bench-top centrifuge</td><td>Eppendorf</td><td>5810R</td><td>Swing-bucket</td></tr><tr><td>Bio-Rad Benchmark Plus Microplate Spectrophotometer</td><td>Bio-Rad</td><td></td><td>Any microplate spectrophotometer capable of reading at 593 nm can be used and is recommended. If a filter-based plate reader is used, choose the closest wavelength possible and use the standard-curve method.</td></tr><tr><td>Ependorf MixMate (microplate orbital mixer)</td><td>Eppendorf</td><td></td><td>This is a very versatile and reliable microplate mixer and works very well for these assays</td></tr><tr><td>General-purpose buffers</td><td></td><td></td><td></td></tr><tr><td>Phosphate-buffered saline (PBS), pH 7.4</td><td></td><td></td><td></td></tr><tr><td>MOPS-buffered saline (MBS); 137 mM NaCl, 2.7 mM KCl, 15 mM MOPS-Na&lt;sup&gt;+&lt;/sup&gt;, pH 7.3</td><td></td><td></td><td></td></tr><tr><td>MBS + 5 mM D-glucose (MBS/D)</td><td></td><td></td><td></td></tr><tr><td>HEPES-buffered saline + 5 mM D-glucose (HBS/D); 137 mM NaCl, 5.2 mM KCl, 1.8 mM CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;bull;2 H&lt;sub&gt;2&lt;/sub&gt;O, 0.8 mM MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;bull;7 H&lt;sub&gt;2&lt;/sub&gt;O, 5 mM D-glucose, 20 mM HEPES-Na&lt;sup&gt;+&lt;/sup&gt;, pH 7.3)</td><td></td><td></td><td></td></tr><tr><td>Cell permeabilisation buffer (CPB; 0.1% saponin in PBS)</td><td></td><td></td><td></td></tr><tr><td>General chemicals</td><td></td><td></td><td></td></tr><tr><td>L-ascorbic acid or sodium L-ascorbate</td><td>Sigma-Aldrich</td><td></td><td>Highest purity preparations should be obtained</td></tr><tr><td>Dehydro-L-ascorbic acid (DHA) dimer</td><td>Sigma-Aldrich</td><td>30790</td><td>Aqueous solutions theoretically yield 2 moles of DHA monomer per mole of DHA dimer</td></tr><tr><td>Cytochalasin B</td><td>Sigma-Aldrich</td><td>C6762</td><td>Stock solutions prepared in DMSO or ethanol</td></tr><tr><td>Ascorbate oxidase (AO)</td><td>Sigma-Aldrich</td><td>A0157</td><td>Stock solutions (120 U/ml) can be prepared in PBS or MBS and then frozen in aliquots</td></tr><tr><td>Potassium ferricyanide (FIC)</td><td>Sigma-Aldrich</td><td>455989</td><td>Trihydrate</td></tr><tr><td>Ferene-S (3-(2-Pyridyl)-5,6-di(2-furyl)-1,2,4-triazine-5&amp;prime;,5&amp;prime;&amp;prime;-disulfonic acid disodium salt)</td><td>Sigma-Aldrich</td><td>92940</td><td></td></tr><tr><td>Sodium L-glutamate</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>L-glutamine</td><td>Sigma-Aldrich</td><td></td><td></td></tr><tr><td>Saponin</td><td>Sigma-Aldrich</td><td>47036</td><td>Prepare a 0.1% stock solution</td></tr><tr><td>Stock solutions for intracellular ascorbate determination assay</td><td></td><td></td><td></td></tr><tr><td>3 M sodium acetate (pH 6.0)</td><td></td><td></td><td></td></tr><tr><td>Glacial acetic acid</td><td></td><td></td><td></td></tr><tr><td>0.2 M citric acid</td><td></td><td></td><td></td></tr><tr><td>3.3 mM FeCl&lt;sub&gt;3&lt;/sub&gt; in 0.1 M acetic acid</td><td></td><td></td><td></td></tr><tr><td>30 mM ferene-S</td><td></td><td></td><td></td></tr><tr><td>50% (v/v) acetic acid + 30% (w/v) trichloroacetic acid (TCA)</td><td></td><td></td><td></td></tr><tr><td>Stock solutions for ascorbate-efflux assay</td><td></td><td></td><td></td></tr><tr><td>AO (120 U/ml)</td><td></td><td></td><td></td></tr><tr><td>2.4 mM ferene-S</td><td></td><td></td><td></td></tr><tr><td>0.12 mM FeCl&lt;sub&gt;3&lt;/sub&gt; in 0.6 mM sodium-citrate</td><td></td><td></td><td></td></tr></tbody></table>

a rapid and specific microplate assay for the determination of intra- and extracellular ascorbate in cultured cells

Vitamin C (ascorbate) plays numerous important roles in cellular metabolism, many of which have only come to light in recent years. For instance, within the brain, ascorbate acts in a neuroprotective and neuromodulatory manner that involves ascorbate cycling between neurons and vicinal astrocytes - a relationship that appears to be crucial for brain ascorbate homeostasis. Additionally, emerging evidence strongly suggests that ascorbate has a greatly expanded role in regulating cellular and systemic iron metabolism than is classically recognized. The increasing recognition of the integral role of ascorbate in normal and deregulated cellular and organismal physiology demands a range of medium-throughput and high-sensitivity analytic techniques that can be executed without the need for highly expensive specialist equipment. Here we provide explicit instructions for a medium-throughput, specific and relatively inexpensive microplate assay for the determination of both intra- and extracellular ascorbate in cell culture.

Determination of Intracellular Ascorbate in Cultured Suspension Cells
In the first assay (Figure 1), intracellular ascorbate is determined, following ascorbate-specific (i.e.&#160;AO-sensitive) reduction of ferricyanide to ferrocyanide, using the highly sensitive determination of ferrocyanide by a previously published procedure37. The detection of ascorbate is based on the colorimetric chelation of ferrous iron that is generated by the ascorbate-dependent redu...

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A Rapid and Specific Microplate Assay for the Determination of Intra- and Extracellular Ascorbate in Cultured Cells

Ascorbate plays numerous important roles in cellular metabolism, many of which have only come to light in recent years. Here we describe a medium-throughput, specific and inexpensive microplate assay for the determination of both intra- and extracellular ascorbate in cell culture.

Vitamin C (ascorbate) plays numerous important roles in cellular metabolism, many of which have only come to light in recent years. For instance, within ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

13.4K Views.  University of Sydney. Ascorbate plays numerous important roles in cellular metabolism, many of which have only come to light in recent years. Here we describe a medium-throughput, specific and inexpensive microplate assay for the determination of both intra- and extracellular ascorbate in cell culture. 

Video: A Rapid and Specific Microplate Assay for the Determination of Intra- and Extracellular Ascorbate in Cultured Cells

Measuring Trans-Plasma Membrane Electron Transport by C2C12 Myotubes

Trans-plasma membrane electron transport (tPMET) plays a role in protection of cells from intracellular reductive stress as well as protection from damage by extracellular oxidants. This process of transporting electrons from intracellular reductants to extracellular oxidants is not well defined. Here we present spectrophotometric assays by C2C12 myotubes to monitor tPMET utilizing the extracellular electron acceptors: water-soluble tetrazolium salt-1 (WST-1) and 2,6-dichlorophenolindophenol (DPIP or DCIP). Through reduction of these electron acceptors, we are able to monitor this process in a real-time analysis. With the addition of enzymes such as ascorbate oxidase (AO) and superoxide dismutase (SOD) to the assays, we can determine which portion of tPMET is due to ascorbate export or superoxide production, respectively. While WST-1 was shown to produce stable results with low background, DPIP was able to be re-oxidized after the addition of AO and SOD, which was demonstrated with spectrophotometric analysis. This method demonstrates a real-time, multi-well, quick spectrophotometric assay with advantages over other methods used to monitor tPMET, such as ferricyanide (FeCN) and ferricytochrome c reduction.

The goal of this protocol is to spectrophotometrically monitor trans-plasma membrane electron transport utilizing extracellular electron acceptors and to analyze enzymatic interactions that may occur with these extracellular electron acceptors.

Trans-plasma membrane electron transport (tPMET) plays a role in protection of cells from intracellular reductive stress as well as protection from damage ...

Biochemistry

A Comprehensive High-Efficiency Protocol for Isolation, Culture, Polarization, and Glycolytic Characterization of Bone Marrow-Derived Macrophages

Macrophages are among the most important antigen-presenting cells. Many subsets of macrophages have been identified with unique metabolic signatures. Macrophages are commonly classified as M1-like (inflammatory) and M2-like (anti-inflammatory) subtypes. M1-like macrophages are pro-inflammatory macrophages that get activated by LPS and/or pro-inflammatory cytokines such as INF-&#947;, IL-12 &#38; IL-2. M1-like polarized macrophages are involved in various diseases by mediating the host&#39;s defense to a variety of bacteria and viruses. That is very important to study LPS induced M1-like macrophages and their metabolic states in inflammatory diseases. M2-like macrophages are considered anti-inflammatory macrophages, activated by anti-inflammatory cytokines and stimulators. Under the pro-inflammatory state, macrophages show increased glycolysis in glycolytic function. The glycolytic function has been actively investigated in the context of glycolysis, glycolytic capacity, glycolytic reserve, compensatory glycolysis, or non-glycolytic acidification using extracellular flux (XF) analyzers.
This paper demonstrates how to assess the glycolytic states in real-time with easy-to-follow steps when the bone marrow-derived macrophages (BMDMs) are respiring, consuming, and producing energy. Using specific inhibitors and activators of glycolysis in this protocol, we show how to obtain a systemic and complete view of glycolytic metabolic processes in the cells and provide more accurate and realistic results. To be able to measure multiple glycolytic phenotypes, we provide an easy, sensitive, DNA-based normalization method for polarization assessment of BMDMs. Culturing, activation/polarization and identification of the phenotype and metabolic state of the BMDMs are crucial techniques that can help to investigate many different types of diseases.
In this paper, we polarized the na&#239;ve M0 macrophages to M1-like and M2-like macrophages with LPS and IL4, respectively, and measured a comprehensive set of glycolytic parameters in BMDMs in real-time and longitudinally over time, using extracellular flux analysis and glycolytic activators and inhibitors.

This protocol provides detailed and comprehensive methods for the isolation, culture, polarization, and measurement of the glycolytic metabolic state of live bone marrow-derived macrophages (BMDMs). This paper provides step-by-step instructions with realistic visual illustrations for workflow and glycolytic assessment of BMDMs in real-time.

Macrophages are among the most important antigen-presenting cells. Many subsets of macrophages have been identified with unique metabolic signatures. ...

Immunology and Infection

Viability Assays for Cells in Culture

Manual cell counts on a microscope are a sensitive means of assessing cellular viability but are time-consuming and therefore expensive. Computerized viability assays are expensive in terms of equipment but can be faster and more objective than manual cell counts. The present report describes the use of three such viability assays. Two of these assays are infrared and one is luminescent. Both infrared assays rely on a 16&#160;bit Odyssey Imager. One infrared assay uses the DRAQ5 stain for nuclei combined with the Sapphire stain for cytosol and is visualized in the 700 nm channel. The other infrared assay, an In-Cell Western, uses antibodies against cytoskeletal proteins (&#945;-tubulin or microtubule associated protein 2) and labels them in the 800 nm channel. The third viability assay is a commonly used luminescent assay for ATP, but we use a quarter of the recommended volume to save on cost. These measurements are all linear and correlate with the number of cells plated, but vary in sensitivity. All three assays circumvent time-consuming microscopy and sample the entire well, thereby reducing sampling error. Finally, all of the assays can easily be completed within one day of the end of the experiment, allowing greater numbers of experiments to be performed within short timeframes. However, they all rely on the assumption that cell numbers remain in proportion to signal strength after treatments, an assumption that is sometimes not met, especially for cellular ATP. Furthermore, if cells increase or decrease in size after treatment, this might affect signal strength without affecting cell number. We conclude that all viability assays, including manual counts, suffer from a number of caveats, but that computerized viability assays are well worth the initial investment. Using all three assays together yields a comprehensive view of cellular structure and function.

Therapeutic compounds are often first examined in vitro with viability assays. Blind cell counts by a human observer can be highly sensitive to small changes in cell number but do not assess function. Computerized viability assays, as described here, can assess both structure and function in an objective manner.

Manual cell counts on a microscope are a sensitive means of assessing cellular viability but are time-consuming and therefore expensive. Computerized ...

Detecting, Visualizing and Quantitating the Generation of Reactive Oxygen Species in an Amoeba Model System

Reactive oxygen species (ROS) comprise a range of reactive and short-lived, oxygen-containing molecules, which are dynamically interconverted or eliminated either catalytically or spontaneously. Due to the short life spans of most ROS and the diversity of their sources and subcellular localizations, a complete picture can be obtained only by careful measurements using a combination of protocols. Here, we present a set of three different protocols using OxyBurst Green (OBG)-coated beads, or dihydroethidium (DHE) and Amplex UltraRed (AUR), to monitor qualitatively and quantitatively various ROS in professional phagocytes such as Dictyostelium. We optimised the beads coating procedures and used OBG-coated beads and live microscopy to dynamically visualize intraphagosomal ROS generation at the single cell level. We identified lipopolysaccharide (LPS) from E. coli as a potent stimulator for ROS generation in Dictyostelium. In addition, we developed real time, medium-throughput assays using DHE and AUR to quantitatively measure intracellular superoxide and extracellular H2O2 production, respectively.

We adapted a set of protocols for the measurement of reactive oxygen species (ROS) that can be applied in various amoeba and mammalian cellular models for qualitative and quantitative studies.

Reactive oxygen species (ROS) comprise a range of reactive and short-lived, oxygen-containing molecules, which are dynamically interconverted or ...

Measurement and Analysis of Extracellular Acid Production to Determine Glycolytic Rate

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both mitochondrial (respiration) and non-mitochondrial (other redox) reactions consume oxygen, but these reactions can be easily distinguished by chemical inhibition of mitochondrial respiration. However, while mitochondrial oxygen consumption is an unambiguous and direct measurement of respiration rate2, the same is not true for extracellular acid production and its relationship to glycolytic rate 3-6. Extracellular acid produced by cells is derived from both lactate, produced by anaerobic glycolysis, and CO2, produced in the citric acid cycle during respiration. For glycolysis, the conversion of glucose to lactate- + H+ and the export of products into the assay medium is the source of glycolytic acidification. For respiration, the export of CO2, hydration to H2CO3 and dissociation to HCO3- + H+ is the source of respiratory acidification. The proportions of glycolytic and respiratory acidification depend on the experimental conditions, including cell type and substrate(s) provided, and can range from nearly 100% glycolytic acidification to nearly 100% respiratory acidification 6. Here, we demonstrate the data collection and calculation methods needed to determine respiratory and glycolytic contributions to total extracellular acidification by whole cells in culture using C2C12 myoblast cells as a model.

Glycolysis is a defining metabolic marker in multiple biological systems. Monitoring glycolysis by measuring the extracellular flux of H+ is common, but requires correction to be quantitative and unambiguous. Here, we demonstrate how to gather and correct extracellular flux data to distinguish between respiratory and glycolytic sources of extracellular acidification.

Extracellular measurement of oxygen consumption and acid production is a simple and powerful way to monitor rates of respiration and glycolysis1. Both ...

Using Real-Time Cell Metabolic Flux Analyzer to Monitor Osteoblast Bioenergetics

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, prevent fracture. In the interest to both optimize peak bone mass and combat various musculoskeletal diseases (i.e., post-menopausal osteoporosis, anorexia nervosa, type 1 and 2 diabetes mellitus), incredible efforts have been made in the field of bone biology to fully characterize osteoblasts throughout their differentiation process. Given the primary role of mature osteoblasts to secrete matrix proteins and mineralization vesicles, it has been noted that these processes take an incredible amount of cellular energy, or adenosine triphosphate (ATP). The overall cellular energy status is often referred to as cellular bioenergetics, and it includes a series of metabolic reactions that sense substrate availability to derive ATP to meet cellular needs. Therefore, the current method details the process of isolating primary, murine bone marrow stromal cells (BMSCs) and monitoring their bioenergetic status using the Real-time cell metabolic flux analyzer at various stages in osteoblast differentiation. Importantly, these data have demonstrated that the metabolic profile changes dramatically throughout osteoblast differentiation. Thus, using this physiologically relevant cell type is required to fully appreciate how a cell's bioenergetic status can regulate the overall function.

Real-time cell metabolic flux assay measures the oxygen consumption rate and extracellular acidification rate, which corresponds to mitochondrial and glycolytic adenosine triphosphate production, using pH and oxygen sensors. The manuscript explains a method to understand the energy status of osteoblasts and the characterization and interpretation of the cellular bioenergetic status.

Bone formation by osteoblasts is an essential process for proper bone acquisition and bone turnover to maintain skeletal homeostasis, and ultimately, ...

Real-Time Analysis of Bioenergetics in Primary Human Retinal Pigment Epithelial Cells Using High-Resolution Respirometry

Metabolic dysfunction of retinal pigment epithelial cells (RPE) is a key pathogenic driver of retinal diseases such as age-related macular degeneration (AMD) and proliferative vitreoretinopathy (PVR). Since RPE are highly metabolically-active cells, alterations in their metabolic status reflect changes in their health and function. High-resolution respirometry allows for real-time kinetic analysis of the two major bioenergetic pathways, glycolysis and mitochondrial oxidative phosphorylation (OXPHOS), through quantification of the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), respectively. The following is an optimized protocol for conducting high-resolution respirometry on primary human retinal pigment epithelial cells (H-RPE). This protocol provides a detailed description of the steps involved in producing bioenergetic profiles of RPE to define their basal and maximal OXPHOS and glycolytic capacities. Exposing H-RPE to different drug injections targeting the mitochondrial and glycolytic machinery results in defined bioenergetic profiles, from which key metabolic parameters can be calculated. This protocol highlights the enhanced response of BAM15 as an uncoupling agent compared to carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) to induce the maximal respiration capacity in RPE. This protocol can be utilized to study the bioenergetic status of RPE under different disease conditions and test the efficacy of novel drugs in restoring the basal metabolic status of RPE.

The metabolic status of human retinal pigment epithelial cells (H-RPE) reflects their health and function. Presented here is an optimized protocol for examining the real-time metabolic flux of H-RPE using high-resolution respirometry.

Metabolic dysfunction of retinal pigment epithelial cells (RPE) is a key pathogenic driver of retinal diseases such as age-related macular degeneration ...

Use of Minimally Invasive Methods to Assess Fuel Utilization and Circadian Rhythms in Older Adults

Aging is associated with multiple physiological changes that contribute synergistically and independently to physical disability and the risk of chronic disease. Although the etiology of age-related physical disability is complex and multifactorial, the decline in mitochondrial function appears to coincide with the progression of functional decline in many older adults. The reason why there is a decrease in mitochondrial function with aging remains elusive, but emerging science indicates that both fuel metabolism and circadian rhythms can influence mitochondrial function.
Recent studies have established that circadian rhythms become disturbed with aging, and that disrupted circadian rhythms have pathological consequences that impact mitochondrial function and overlap with many age-associated chronic diseases. Current quantitative methods for direct assessment of mitochondrial function are invasive and typically require a muscle biopsy, which can pose difficulties with participant recruitment and study adherence, given the perceived levels of potential pain and risk. Thus, an innovative and relatively noninvasive protocol to assess changes in mitochondrial function at the cellular level and circadian patterns in older adults was adapted. Specifically, a real-time metabolic flux analyzer is used to assess the mitochondrial bioenergetic function of white blood cells under differential substrate availability.
The expression of circadian clock genes in white blood cells to cross-correlate with the mitochondrial bioenergetics and circadian rhythm outcomes are also analyzed. It is believed that these innovative methodological approaches will aid future clinical trials by providing minimally invasive methods for studying mitochondrial substrate preference and circadian rhythms in older adults.

A novel and minimally invasive protocol is presented to evaluate the synergistic impact of fuel utilization and circadian rhythms on aging individuals, utilizing peripheral blood mononuclear cells.

Aging is associated with multiple physiological changes that contribute synergistically and independently to physical disability and the risk of chronic ...

Extracellular Flux Assay: A Method for Measuring Metabolic Profile of Cells

Source: Hlozkova K. et al. <a href="https://jove.unicartagenaproxy.elogim.com/t/58426">Assessment of the Metabolic Profile of Primary Leukemia Cells.</a> J. Vis. Exp. (2018).
This protocol provides a biochemical approach for measuring extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) of primary leukemic cells with extracellular flux assay. The assessment of the metabolic profile of these cells helps to characterize their demands, leading to personalized therapy.

Test strumienia zewnątrzkomórkowego: metoda pomiaru profilu metabolicznego komórek

Source: Hlozkova K. et al. Assessment of the Metabolic Profile of Primary Leukemia Cells. J. Vis. Exp. (2018).
This protocol provides a biochemical ...

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Cancer Research

Leukemia

In vitro techniques & assays

Caco-2 Cell Bioassay: An In Vitro Method for Measuring Iron Bioavailability in Complex Food Sources

Source: Glahn, R. P. <a href="https://appjove.unicartagenaproxy.elogim.com/v/63859">The Caco-2 Cell Bioassay for Measurement of Food Iron Bioavailability.</a> J. Vis. Exp. (2022)
This video demonstrates an in vitro technique to assess iron bioavailability in complex food sources. The technique combines the simulation of human digestion along with iron uptake by differentiated Caco-2 cells. Intracellular ferritin formation acts as a marker for cellular iron uptake.

Test biologiczny komórek Caco-2: metoda in vitro do pomiaru biodostępności żelaza w złożonych źródłach żywności

Source: Glahn, R. P. The Caco-2 Cell Bioassay for Measurement of Food Iron Bioavailability. J. Vis. Exp. (2022)
This video demonstrates an in vitro ...