The project was supported by Ralph and Marian Falk Medical Research Catalyst Award and Kathleen Kelly Award. Other supports included the National Center for Research Resources UL1RR025755 and NCI P30CA16058 (OSUCCC), the NIH Roadmap for Medical Research. The content is solely the responsibility of the authors and does not represent the official views of the National Center for Research Resources or the NIH.

Animal studies were approved by the Institutional Animal Care and Use Committee of The Ohio State University (OSU, protocol 2007A0262-R4).
NOTE: All procedures must be done in a class II biosafety cabinet with the blower on and the lights off.
1. Preparation of materials
NOTE: All materials are listed in the Table of Materials.
<ol>
	<li>Prepare Medium 1, centrifugation Medium 2, and storage and working solutions of fluorescent 2-deoxy-2-[(7-nitro-2,1,3-benzoxadiazol-4-yl) amino]-D-glucose (2-NBDG or FD-glucose) according to Table 1 in a Class II biosafety cabinet. Protect FD-glucose from light throughout the experiment by turning off all the lights under the hood.</li>
	<li>Use ready-to-use cell culture reagents: Trypsin-EDTA, phosphate-buffered saline (PBS), and glucose-free and phenol red-free Dulbecco&#39;s Modified Eagle Medium (hereafter, glucose-free DMEM).</li>
	<li>Use 20 mL of radioimmunoprecipitation assay (RIPA) lysis buffer. 
	&#8203;NOTE: In the following sections, extracellular depletion and intracellular uptake of different FD-glucose doses are compared in 3T3-L1 fibroblasts with and without insulin stimulation (Figure 2).</li>
</ol>
2. 3T3-L1 cell culture and maintenance
<ol>
	<li>Culture 3T3-L1 mouse fibroblasts by plating 1 x 106 cells diluted in 20 mL of Medium 1 in two 96-well plates. Plate 100 &#956;L of cell suspension per well, ensuring to maintain homogeneity of this suspension by mixing. Grow cells for 48 h in a 37 &#176;C incubator (5% CO2 and 95% humidity) without changing media until about 70%-80% confluency.</li>
	<li>Maintain cells confluent and fasted by culturing them in the same medium for 48 h. Vary the incubation time from 24-72 h depending on the desired fasting catabolic state.</li>
</ol>
3. Starvation of 3T3-L1 cells
<ol>
	<li>Decant media from cells in the 96-well plates. Absorb the remaining fluid using sterile paper towels.</li>
	<li>Rinse the 96-well plate with 100 &#956;L of PBS per well and absorb the remaining fluid with sterile paper towels.</li>
	<li>Add 100 &#956;L of glucose-free DMEM per well and incubate for 40 min. Adjust the time of incubation depending on cell type, stimuli, and desired level of fasting. 
	&#8203;NOTE: The time of incubation may require optimization depending on the season.</li>
</ol>
4. Preparation of FD-glucose solutions with different concentrations
NOTE: The experiment in Figure 2 examines extracellular and intercellular media with and without stimulation with insulin.
<ol>
	<li>Use eight replicates (one row in a 96-well plate) for each stimulation condition. Therefore, prepare 1 mL for each stimulation condition (specified in Table 2 and below).</li>
	<li>Use eight stimulation conditions without insulin: FD-glucose of 2.5 &#956;g/mL, 1 &#956;g/mL, 0.5 &#956;g/mL, 0.2 &#956;g/mL, 0.1 &#956;g/mL, 0.05 &#956;g/mL, and 0 &#956;g /mL (control without FD-glucose) in one 96-well plate.</li>
	<li>Use eight stimulation conditions with insulin (10 &#956;g/mL, final concentration): FD-glucose of 2.5 &#956;g/mL, 1 &#956;g/mL, 0.5 &#956;g/mL, 0.2 &#956;g/mL, 0.1 &#956;g/mL, 0.05 &#956;g/mL, and 0 &#956;g/mL (control without FD-glucose) in another 96-well plate.</li>
	<li>To prepare stimulation conditions, dilute 1 &#956;L of 5 mg/mL FD-glucose working solution (Table 1) in 999 &#956;L of glucose-free DMEM to obtain 1 mL of working stock solution (1:1000 dilution or 5 &#956;g/mL).
	<ol>
		<li>Using this working stock solution, prepare 1:2000, 1:5000, 1:10,000, 1:25,000, 1:50,000, and 1:100,000 dilutions of FD-glucose to obtain 2.5 &#956;g/mL, 1 &#956;g/mL, 0.5 &#956;g/mL, 0.2 &#956;g/mL, 0.1 &#956;g/mL, and 0.05 &#956;g/mL (Table 2) in 2 mL tubes immediately before experiments in a biosafety cabinet without lights.</li>
	</ol>
	</li>
	<li>Repeat step 4.4 and add 1 &#956;L of insulin (10 mg/mL, final concentration) to each of the conditions specified above. 
	&#8203;NOTE: The final concentration of insulin in these samples is 10 &#956;g/mL = 1.7 nmol/mL. To achieve homogenization, add insulin to the tubes before adding other solutions.</li>
</ol>
5. Treating starved 3T3-L1 cells
<ol>
	<li>After 40 min of incubation, decant the glucose-free DMEM from each well in 96-well plates and absorb the remaining fluid with sterile paper towels.</li>
	<li>Add 100 &#956;L (per well) each from the various treatment conditions described above to wells along one column, and 100 &#956;L (per well) from the same treatment condition to the wells along the row (eight replicates) in both 96-well plates. Label the plates that utilized the conditions with and without insulin. Add glucose-free DMEM alone to the control wells (n = 8).</li>
	<li>Incubate the plate for 40 min in a cell culture incubator (37 &#176;C, 5% CO2, and 95% humidity) in the dark.</li>
</ol>
6. Extracellular and intercellular measurements for stimulated 3T3-L1 cells
<ol>
	<li>After the cells have been stimulated for 40 min, transfer the stimulation media from both the plates into new 96-well plates maintaining the same experimental layout.</li>
	<li>Decant any remaining solution from the original stimulated 96-well plates with cells on sterile paper towels.</li>
	<li>Optionally, wash cells with PBS. Remove PBS and decant any remaining solution on sterile paper towels.</li>
	<li>Add 100 &#956;L of the RIPA lysis buffer to each well. Optionally, add protease inhibitor to the RIPA buffer to protect proteins. Place plates containing RIPA in a shaker for 30 min.</li>
	<li>Using a microplate reader (Table of Materials), measure fluorescence at excitation (Ex) and emission (Em) wavelengths of 485 and 535 nm, respectively, first in the medium containing extracellular FD-glucose, and then, the plate with RIPA-lysed cells at the end of 30 min incubation (see step 6.4).</li>
</ol>
7. Protein-based normalization of intracellular glucose uptake
NOTE: Intracellular FD-glucose uptake depends on the cell number. Protein levels in cellular lysates are proportional to the cell numbers that allow normalizing intracellular FD-glucose levels to the number of cells in each well.
<ol>
	<li>Perform BCA protein assay according to manufacturer&#39;s instructions (see Table of Materials). Quantify the protein concentrations in each well containing RIPA-lysed cells.</li>
	<li>Use 10 &#956;L of the RIPA homogenate and measure protein concentrations in triplicate to increase the accuracy of measurements in 96-well plates.</li>
	<li>Quantify protein based on protein standards (0, 10, 20, 30, 40, 50, and 60 &#956;g/mL protein) analyzed together with samples on each 96-well plate.</li>
	<li>Measure the absorbance at 562 nm and quantify protein concentrations in each sample based on the linear regression formula obtained for the protein standard.</li>
	<li>Normalize the levels of intracellular FD-glucose by using fluorescent value/protein concentration in each well. Extracellular FD-glucose depletion does not require normalization.</li>
	<li>Optionally, use the average baseline values in control samples for additional normalization (100%).</li>
</ol>
8. Ex vivo measurement of extracellular FD glucose depletion in organs
<ol>
	<li>Pretreat mice with compounds stimulating glucose metabolism before the organ dissection, as follows.</li>
	<li>Use nine-week-old Lepob male mice (n = 11). Feed all mice with a regular chow diet before the experiment.</li>
	<li>Assign mice randomly into three groups for intraperitoneal (i.p.) injections.
	<ol>
		<li>Inject mice from the control Lepob group with 0.1 mL of sterile PBS (n = 4).</li>
		<li>Inject mice from the insulin Lepob group with 0.1 mL of sterile PBS, containing 12 IU human insulin per gram of body weight (BW) (n= 3).</li>
		<li>Inject mice from the AAC2 Lepob group with 0.1 mL of sterile PBS, containing 0.1 nmol AAC2 per gram of body weight (BW) (n = 4).</li>
	</ol>
	</li>
	<li>After 15 min, subject mice to inhalation of 5% isoflurane and exsanguinate blood by cardiac puncture from the anesthetized animals26.</li>
	<li>Dissect visceral epidydimal white adipose tissue (200 mg), liver (200 mg), and whole-brain from each animal. Use a Class II biosafety hood for tissue handling.</li>
	<li>Prepare FD-glucose (0.29 mM) working solution in glucose-free DMEM in a Class II Biosafety cabinet without light.</li>
	<li>Measure the fluorescence of FD-glucose working solution (0.29 mM) at excitation and emission wavelengths of 485 and 535 nm, respectively, using a microplate reader.</li>
	<li>Incubate the harvested tissues/organs in a 6-well plate containing PBS for 1 min. Handle each tissue or organ in a separate well.</li>
	<li>After 1 min, place each tissue on a sterile paper towel to absorb PBS. Transfer tissues/organs into a separate 6-well plate containing 4,000 &#181;L of glucose-free DMEM and incubate for 2 min.</li>
	<li>After 2 min, remove and transfer the tissues into wells of a 6-well plate containing 0.29 mM FD-glucose working solution (1 mL/well). Incubate the 6-well plates containing tissues in FD-glucose working solution at 37 &#176;C (5% CO2 and 95% humidity).</li>
	<li>Collect 100 &#181;L of FD-glucose working solution from each well after 0, 10, 20, 30, 40, 60, 90, and 120 min of incubation, to analyze the kinetics of extracellular FD glucose depletion. Shake before and after collection.</li>
	<li>Transfer 100 &#181;L of FD-glucose working solution into 96-well plates to measure the fluorescence at excitation and emission wavelengths of 485 and 535 nm, respectively, using a microplate reader.</li>
	<li>Normalize fluorescence to the 0 min value (100%) for each organ in each animal (Figure 3).</li>
</ol>

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B., Ziouzenkova, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Osmotic+adaptation+by+Na(+)-dependent+transporters+and+ACE2:+correlation+with+hemostatic+crisis+in+COVID-19.">Osmotic adaptation by Na(+)-dependent transporters and ACE2: correlation with hemostatic crisis in COVID-19.</a> Biomedicines. 8 (11), 460 (2020).</li><li>Ligasova, A., Koberna, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+dyes-highly+sensitive+reporters+of+cell+quantification:+comparison+with+other+cell+quantification+methods.">DNA dyes-highly sensitive reporters of cell quantification: comparison with other cell quantification methods.</a> Molecules. 26 (18), 5515 (2021).</li><li>DeFronzo, R. A., Tobin, J. D., Andres, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glucose+clamp+technique:+a+method+for+quantifying+insulin+secretion+and+resistance.">Glucose clamp technique: a method for quantifying insulin secretion and resistance.</a> American Journal of Physiology. 237 (3), 214-223 (1979).</li></ol>

Authors have no issues to disclose and have no conflict of interests.

The direct comparison of extracellular FD-glucose depletion with normalized intracellular glucose uptake in cells culture showed a high correlation, suggesting that extracellular glucose depletion could be a surrogate measurement for glucose uptake assessment. The measurement of extracellular FD-glucose can use a broad range of FD glucose concentrations, also 0.5-2.5 &#956;g FD-glucose/mL appear to provide the optimal range. Extracellular FD-glucose does not require normalization to cell number or protein concentrations necessary for intracellular FD-glucose uptake.&#160;The anticipated limitation of extracellular FD-glucose measurements is the investigation of reactive oxygen species-inducing agents and the traces of blood within explants that can quench fluorescence. Other environmental factors affecting fluorescence, such as light, should also be considered. Prolonged handling time required while using multiple tissue explants can compromise tissue viability and glucose metabolism. This could be an additional limitation. The excised tissues need to be in a small size range (50-300 mg) to allow the permeability of glucose inside the tissue. Simultaneous handling of tissues by multiple operators can reduce handling time. Although we did not measure the glucose uptake within the tissue explants, the accumulated observations in cultured tissue explants suggest that they survive in the glucose-containing medium for up to 17 days with the progressive loss of functionality33. In this protocol, the short incubation time for explants allows monitoring glucose metabolism in relatively functional tissues. In the tissues with complex cellular organization, such as the brain, the size presents an unresolvable limitation, because the disintegration of complex tissue and resulting cell death influences the glucose uptake. However, the incubation of the brain for 24 h has been previously used for identification of endogenous regulatory mechanism, supporting relative physiological functionality of excised mouse brain in glucose-containing medium34. Regardless, the simplified procedure and easy accessibility of extracellular FD-glucose can allow using this assay in a high-throughput format. The comparison of basic and insulin-stimulated extracellular FD-glucose depletion suggests that this assay could be applied to identify humoral, nutritional, pathogenic, environmental factors, or pharmaceutical drugs regulating glucose uptake. Although this assay simplifies the discovery, the findings required validation with the quantitative radiolabeling20&#160;analytical methods21,22,23 and/or imaging methods5,24&#160;for rigorous mechanistic elucidation of the pathways.
The critical steps in this protocol include optimization of confluence and fasting time that influence both basal and stimulated FD-glucose uptake. These parameters could also be affected by season based on the authors&#39; observations. Moreover, different cell types use specific mechanisms for glucose uptake including different glucose transporters and regulatory hormone/receptor interactions32,35,36. Therefore, positive controls other than insulin need to be considered for the cell cultures representing tissues other than muscle or adipose tissue regulated by insulin/GLUT4 axes. The troubleshooting also includes adjusting cell confluence and optimizing time for fasting/refeeding with FD-glucose. In addition, the sensitivity of extracellular glucose measurement could be improved by reducing the levels of extracellular FD-glucose, taking into consideration the threshold levels required for glucose uptake. The experiment described in Figure&#160;2 can help researchers to customize this assay for use with other cell cultures or troubleshooting.
The intrinsic variability of glucose metabolism also required a higher sample size (n = 5-8). Although extracellular measurements of FD-glucose loss exhibited the same accuracy as intracellular uptake of FD-glucose normalized by protein, the normalization can reduce assay variability. In addition to the measurement of protein concentrations, the measurement of DNA content in the cell can be another reliable surrogate assessment of cell number37.
Extracellular FD-glucose is accessible for kinetic measurements. Here, a simple procedure of kinetic measurements of extracellular FD-glucose has been developed to identify organ-specific responses to mediators of glucose uptake. These ex vitro studies can directly test organ response to various stimuli&#160;in vivo&#160;that is combined with the exposure of organs to FD-glucose and&#160;ex vivo kinetic measurements, as shown in Figure&#160;3. Although ex vivo studies do not fully resemble in vivo conditions, they have multiple advantages. Organ explants maintain complex multicellular primary cell structure, unlike immortal cell cultures exhibiting altered gene expression. Modern drug development utilizes model organisms, indirect assessment of glucose uptake based on transcriptome or metabolome22, comprehensive insulin clamp technique38, or expensive imaging32. Although measurement of extracellular FD-glucose in explanted organs will not replace these technologies, it will provide a rapid assessment of compound, metabolite, and genes to facilitate the discovery of novel therapeutic targets regulating glucose uptake in a tissue-specific manner. The development of safe therapies addressing glucose metabolism in specific tissues can offer a solution for cancer, dementia, aging, diabetes, autoimmune diseases, obesity, and other related applications.

The demand for measuring glucose uptake rises together with the critical need to address an epidemic increase in a multitude of diseases dependent on glucose metabolism. Underlying mechanisms of degenerative metabolic diseases, neurological and cognitive disorders1, inflammatory2 and infectious diseases3, cancer4,5, as well as aging6, depend on glucose metabolism for energy and its storage, anabolic processes, protein, and gene modification, signaling, regulation of genes, and nucleic acids synthesis and replication7,8,9. Diabetes mellitus (DM) is directly related to malfunction of glucose uptake regulation. DM is a spectrum of chronic diseases such as type-1, -2, and -3 diabetes mellitus, gestational diabetes, maturity-onset diabetes of the young, and other types of this disease induced by environmental and/or genetic factors. In 2016, the first WHO Global report on diabetes demonstrated that the number of adults living with the most widespread DM has almost quadrupled since 1980 to 422 million adults10, and this number of DM patients has been rising exponentially for the last few decades. In 2019 alone, an estimate of 1.5 million deaths was directly caused by DM10. This dramatic upsurge is due to the rise in type-2 DM and the conditions driving it, including overweight and obesity10. The COVID-19 pandemic revealed a two-fold increase in mortality in patients with DM compared to the general population, suggesting the profound yet poorly understood role of glucose metabolism in the immune defense3. Prevention, early diagnosis, and treatment of DM, obesity, and other diseases require optimization of measurements of glucose uptake by different tissues, and identification of environmental11, nutritional12, endocrine13, genetic14, and epigenetic15 factors affecting glucose uptake.
In research, intracellular and/or tissue uptake of glucose is commonly measured by fluorescently-labeled glucose (FD-glucose) in vitro16,17,18 and in vivo19. FD-glucose became a preferred method compared to more precise methods using radioactively-labeled glucose20, analytical mass spectroscopy analysis21, metabolomics22, nuclear magnetic resonance methods23, and positron emission tomography/computed tomography (PET/CT)5,24. Unlike FD-glucose uptake, analytical methods requiring more biological material may involve a multi-step sample preparation, expensive instruments, and complex data analysis. Effective and inexpensive measurements of FD-glucose uptake in cell cultures have been utilized in proof-of-concept experiments and may require validation by other methods.
The basis of FD-glucose application for glucose uptake studies is the reduced metabolism of FD-glucose compared to endogenous glucose25. Nonetheless, both endogenous glucose and FD-glucose are dynamically distributed among all cellular compartments for use in anabolic, catabolic, and signaling processes. The compartmentalization and time-dependent processing25 of FD-glucose interfere with the fluorescence measurements, and represent the major limiting factors for the use of this assay in high-throughput screening experiments, kinetic analysis, 3D cell culture, co-cultures, and tissue explant experiments. Here, we provide data demonstrating a high correlation between the extracellular depletion of FD-glucose and its intracellular uptake, suggesting the extracellular depletion of FD-glucose as a surrogate measurement for intracellular glucose uptake. The measurement of extracellular depletion of glucose was applied to validate tissue-specific differences in glucose uptake in mice treated with insulin and an experimental drug18 to provide a proof-of-principle of this method.
The current protocol describes intracellular and extracellular (Figure 1) measurements of FD-glucose uptake in 3T3-L1 cells. Protocol sections 1-7 explain the culture and growth of cells for 48 h; cell starvation, stimulation, and baseline extracellular measurements; and post-stimulation measurements of extracellular FD-glucose and intracellular measurements of FD-glucose and protein. Protocol section 8 describes the ex vivo measurement of extracellular uptake of FD-glucose in tissues dissected from ob/ob mice in the presence and absence of insulin and&#160;amino acid compound 2 (AAC2) described elsewhere18.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3T3-L1 mouse fibroblasts</td>
			<td>ATCC</td>
			<td>CL-173</td>
			<td>Cell line</td>
		</tr>
		<tr>
			<td>96-well plates</td>
			<td>Falcon</td>
			<td>353227</td>
			<td>Plastic ware</td>
		</tr>
		<tr>
			<td>B6.V-Lepob/J male mice</td>
			<td>Jackson Laboratory</td>
			<td>stock number 000632</td>
			<td>Mice</td>
		</tr>
		<tr>
			<td>BioTek Synergy H1 modular multimode microplate reader (Fisher Scientific, US)</td>
			<td>Fisher Scientific, US</td>
			<td>&#160;B-SHT</td>
			<td>Device</td>
		</tr>
		<tr>
			<td>Bovine serum</td>
			<td>Gibco/ThermoFisher</td>
			<td>161790-060</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Calf serum</td>
			<td>Gibco/ThermoFisher</td>
			<td>26010-066</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Cell incubator</td>
			<td>Forma</td>
			<td>Series II Water Jacket</td>
			<td>Device</td>
		</tr>
		<tr>
			<td>Diet (mouse/rat diet, irradiated)</td>
			<td>Envigo</td>
			<td>Teklad LM-485</td>
			<td>Diet</td>
		</tr>
		<tr>
			<td>Dimethylsulfoxide (DMSO)</td>
			<td>Sigma LifeScience</td>
			<td>D2650-100mL</td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium</td>
			<td>Gibco/ThermoFisher</td>
			<td>&#160;11965-092</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sigma Aldrich</td>
			<td>E7023-500mL</td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>Fluorescent 2-deoxy-2-[(7-nitro-2,1,3-benzoxadiazol-4-yl) amino]-D-glucose)</td>
			<td>Sigma</td>
			<td>72987-1MG</td>
			<td>Assay</td>
		</tr>
		<tr>
			<td>Glucose-free and phenol red-free DMEM</td>
			<td>Gibco/ThermoFisher</td>
			<td>A14430-01</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Human insulin 10 mg/mL</td>
			<td>MilliporeSigma, Cat N 91077C</td>
			<td>Cat N 91077C</td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>Isoflurane, 5%</td>
			<td>Henry Schein</td>
			<td>NDC 11695-6776-2</td>
			<td>Anestaetic</td>
		</tr>
		<tr>
			<td>Penicillin/streptomycin (P/S)</td>
			<td>Gibco/ThermoFisher</td>
			<td>15140-122</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Phosphate buffered solution</td>
			<td>Sigma-Aldrich</td>
			<td>DA537-500 mL</td>
			<td>Cell culture</td>
		</tr>
		<tr>
			<td>Pierce bicinchoninic acid (BCA) protein assay</td>
			<td>ThermoFisher</td>
			<td>Cat N23225</td>
			<td>Assay</td>
		</tr>
		<tr>
			<td>Radioimmunoprecipitation assay lysis buffer</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-24948</td>
			<td>Assay</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.05%)</td>
			<td>Gibco/ThermoFisher</td>
			<td>&#160;25300-054</td>
			<td>Cell culture</td>
		</tr>
	</tbody>
</table>

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.

Intracellular intake and extracellular glucose depletion were measured in 3T3-L1 preadipocytes, in response to different concentrations of FD-glucose (Figure&#160;2) with and without insulin stimulation. Figure&#160;2A demonstrates a dose-dependent increase in the intracellular uptake of FD-glucose, which was significantly increased in the presence of insulin. The concomitant decrease in extracellular FD-glucose in the same cells is shown in Figure&#160;2B, where insulin stimulation led to significantly decreased extracellular FD-glucose levels compared to the samples without insulin stimulation. The intracellular intake of FD-glucose correlated with the extracellular depletion of FD-glucose in a dose-dependent manner in samples with and without insulin (Figure&#160;2C). Thus, extracellular depletion of FD-glucose can measure a change in glucose uptake with comparable accuracy (R2 = 0.99, P &#60; 0.001) as intracellular FD-glucose uptake.
Next, an experimental setting was developed to validate extracellular FD-glucose uptake application in kinetic studies examining different tissues upon stimulation with canonic and experimental inducers of glucose uptake (Figure&#160;3). We selected the Lepob mouse model of insulin resistance in peripheral tissues, including visceral white adipose tissue27,28. The well-established responses to insulin in these mice were compared with effects of novel compound AAC2, acting on both peripheral and nervous tissues via leptin receptor and glucose transporter GLUT1 dependent mechanisms18. Lepob mice were i.p. injected with insulin or AAC2. Organs were dissected 15 min after injection. Then, the kinetics of extracellular FD-glucose depletion was studied in visceral fat, liver, and brain ex vivo (Figure&#160;3). In agreement with the reported insulin resistance in peripheral tissues, extracellular FD-glucose was not depleted in non-stimulated control visceral fat during 120 min of incubation (Figure&#160;3A). In contrast, pretreatment of mice with insulin or AAC2 before the dissection led to significant depletion of extracellular FD-glucose within a time interval of 30 min and 60 min, respectively.
Liver uptake of glucose utilizes glucose transporters, including GLUT2, that is independent of insulin stimulation29,30. Accordingly, extracellular FD-glucose was initially not depleted in insulin-stimulated liver explants compared with non-stimulated liver explants (Figure&#160;3B), although significant depletion of FD-glucose was observed after 120 min of incubation in liver explant stimulated with insulin compared with the initial levels (0 min). On the other hand, extracellular FD glucose was significantly decreased in a time-dependent manner in liver explants pretreated with AAC2.
In the brain, the main transporter is GLUT1 among other transporters31. The FD-glucose depletion kinetics was strikingly different in the brain compared to other tissue (Figure&#160;3C). The significant depletion of glucose was observed in the extracellular medium containing the non-treated brain after 60 min of incubation (from 100% at 0 min to 78%). The medium incubated with brains from insulin-treated Lepob mice have shown moderate but a significant linear decrease in FD-glucose (from 100% at 0 min to 95%). AAC2-stimulated brains lead to a profound rapid decrease of extracellular FD-glucose during the first 20 min (from 100% at 0 min to 67.4%). Together, measurements of extracellular glucose depletion support the previously reported differences in glucose uptake in these tissues in vivo32.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63681/63681fig01.jpg" /> 
Figure 1: Principle of the extracellular FD-glucose depletions method. Cells cultured in 96-well format are suitable for this assay. To survive, cells take up glucose and this influx decreases the levels of extracellular glucose. The replacement of glucose with FD-glucose allows monitoring of these changes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63681/63681fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63681/63681fig02.jpg" /> 
Figure 2: Depletion of extracellular FD-glucose correlated with the intracellular uptake in vitro.&#160;3T3-L1 preadipocytes were incubated in a glucose-free medium for 40 min before incubation with different concentrations of FD-glucose with and without insulin (1.7 mM; 40 min incubation). (A, B) Dose-dependent uptake of FD-glucose (A) and extracellular FD-glucose depletion (B) in control (i.e., without insulin) (hatched bars) and insulin-stimulated cells (black bars). FD-glucose uptake was normalized by protein concentrations. Data are shown as % of the value measured in control incubated without FD-glucose (mean SD, n = 8 per condition). Significance was examined by unpaired t-test. (C) Correlation between intracellular and extracellular FD-glucose (%) measures with (squares) or without (circles) insulin in experiments described in (A) and (B). Pearson correlation, P &#60; 0.001. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63681/63681fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63681/63681fig03.jpg" /> 
Figure 3: Kinetics of extracellular FD-glucose depletion in different organs ex vivo. (A-C) Lepob mice were injected with vehicle (PBS, n = 4), insulin (12 IU/kg BW, n = 3), and AAC2 (1 nmol/g BW, n = 3). After 15 min, tissues were dissected and isolated. Explants of (A) visceral fat, (B) liver, and (C) brain were incubated in FD-glucose (0.29 mM). The kinetics of extracellular glucose depletion was measured in aliquots of the medium at different time points. Data (mean SD) are shown as % of fluorescence in each organ at 0 min of incubation. Significance P &#60; 0.05, unpaired t-test. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63681/63681fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Solution/Medium</td>
			<td>Components</td>
		</tr>
		<tr>
			<td>Ethanol:DMSO (1:1/v/v)</td>
			<td>Ethanol (200 &#181;L) and DMSO (200 &#181;L) in 1-1.5 mL tube; use cell culture grade ethanol and DMSO</td>
		</tr>
		<tr>
			<td>Medium 1</td>
			<td>DMEM (89 mL), Penicillin/streptomycin (1%) (1 mL) and Calf serum (10%) (10 mL) in sterile 50 mL tube</td>
		</tr>
		<tr>
			<td>Centrifugation Medium 2</td>
			<td>DMEM (89 mL), Penicillin/streptomycin (1%) (1 mL) and Bovine serum (10%) (10 mL) in sterile 50 mL tube</td>
		</tr>
		<tr>
			<td>Storage FD-glucose solution 5 mg/mL (14.5 mM)</td>
			<td>FD glucose (1 mg) and Ethanol:DMSO (1:1/v/v) (200 &#181;L) in 0.5 mL tube; store at -80 &#176;C, preferentially under argon or nitrogen atmosphere</td>
		</tr>
		<tr>
			<td>Working FD-glucose solution 5 &#181;g/mL (14.5 mM)</td>
			<td>Storage FD glucose solution 5 mg/mL (1 &#181;L), Glucose free DMEM (999 &#181;L); prepare immediately before experiment.</td>
		</tr>
	</tbody>
</table>
Table 1: Preparation of culture media and FD-glucose solutions.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Dilution</td>
			<td>Control</td>
			<td>1:2x103</td>
			<td>1:5x103</td>
			<td>1:1x104</td>
			<td>1:2.5x104</td>
			<td>1:5x104</td>
			<td>1:1x105</td>
		</tr>
		<tr>
			<td>Concentration (&#181;g/mL)</td>
			<td>0</td>
			<td>2.5</td>
			<td>1</td>
			<td>0.5</td>
			<td>0.2</td>
			<td>0.1</td>
			<td>0.05</td>
		</tr>
		<tr>
			<td>Glucose-free DMEM (&#181;L)</td>
			<td>1000</td>
			<td>500</td>
			<td>800</td>
			<td>900</td>
			<td>960</td>
			<td>980</td>
			<td>990</td>
		</tr>
		<tr>
			<td>FD Glucose 5 &#181;g/mL (&#181;L)</td>
			<td>0</td>
			<td>500</td>
			<td>200</td>
			<td>100</td>
			<td>40</td>
			<td>20</td>
			<td>10</td>
		</tr>
	</tbody>
</table>
Table 2: Preparation of FD-glucose solutions with different concentrations for experiments shown in Figure 2.

Watch this Scientific Journal Video about Extracellular Glucose Depletion as an Indirect Measure of Glucose Uptake in Cells and Tissues Ex Vivo at JoVE.com

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.

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 ...

extracellular-glucose-depletion-as-an-indirect-measure-glucose-uptake

Nanyang Technological University

Research

JoVE Journal

Biochemistry

2.6K Views.  The Ohio State University. 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. 

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

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 ...

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate neurotransmission that occurs within synapses because it is highly regulated by dynamic protein-protein interactions and phosphorylation events. Accordingly, when any method is used to study synaptic transmission, a major goal is to preserve these transient physiological modifications. Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments. In other words, the protocol describes a biochemical methodology to carry out protein enrichment from presynaptic, postsynaptic, and extrasynaptic compartments. First, synaptosomes, or synaptic terminals, are obtained from neurons that contain all synaptic compartments by means of a discontinuous sucrose gradient. Of note, the quality of this initial synaptic membrane preparation is critical. Subsequently, the isolation of the different subsynaptic compartments is achieved with light solubilization using mild detergents at differential pH conditions. This allows for separation by gradient and isopycnic centrifugations. Finally, protein enrichment at the different subsynaptic compartments (i.e., pre-, post- and extrasynaptic membrane fractions) is validated by means of immunoblot analysis using well-characterized synaptic protein markers (i.e., SNAP-25, PSD-95, and synaptophysin, respectively), thus enabling a direct assessment of the synaptic distribution of any particular neuronal protein.

Brain Membrane Fractionation: An Ex Vivo Approach to Assess Subsynaptic Protein Localization

Here, we present a brain membrane fractionation protocol that represents a robust procedure to isolate proteins belonging to different synaptic compartments.

Assessing the synaptic protein composition and function constitutes an important challenge in neuroscience. However, it is not easy to evaluate ...

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 ...

Analysis of Endocytic Uptake and Retrograde Transport to the Trans-Golgi Network Using Functionalized Nanobodies in Cultured Cells

Transport of proteins and membranes from the cell surface to the Golgi and beyond is essential for homeostasis, organelle identity and physiology. To study retrograde protein traffic, we have recently developed a versatile nanobody-based toolkit to analyze transport from the cell surface to the Golgi complex, either by fixed and live cell imaging, by electron microscopy, or biochemically. We engineered functionalized anti-green fluorescent protein (GFP) nanobodies &#8212; small, monomeric, high-affinity protein binders &#8212; that can be applied to cell lines expressing membrane proteins of interest with an extracellular GFP moiety. Derivatized nanobodies bound to the GFP reporters are specifically internalized and transported piggyback along the reporters&#39; sorting routes. Nanobodies were functionalized with fluorophores to follow retrograde transport by fluorescence microscopy and live imaging, with ascorbate peroxidase 2 (APEX2) to investigate the ultrastructural localization of reporter-nanobody complexes by electron microscopy, and with tyrosine sulfation (TS) motifs to assess kinetics of trans-Golgi network (TGN) arrival. In this methodological article, we outline the general procedure to bacterially express and purify functionalized nanobodies. We illustrate the powerful use of our tool using the mCherry- and TS-modified nanobodies to analyze endocytic uptake and TGN arrival of cargo proteins.

Retrograde transport of proteins from the cell surface to the Golgi is essential to maintain membrane homeostasis. Here, we describe a method to biochemically analyze cell surface-to-Golgi transport of recombinant proteins using functionalized nanobodies in HeLa cells.

Transport of proteins and membranes from the cell surface to the Golgi and beyond is essential for homeostasis, organelle identity and physiology. To ...

Atomic Absorbance Spectroscopy to Measure Intracellular Zinc Pools in Mammalian Cells

Transition metals are essential micronutrients for organisms but can be toxic to cells at high concentrations by competing with physiological metals in proteins and generating redox stress. Pathological conditions that lead to metal depletion or accumulation are causal agents of different human diseases. Some examples include anemia, acrodermatitis enteropathica, and Wilson&#8217;s and Menkes&#8217; diseases. It is therefore important to be able to measure the levels and transport of transition metals in biological samples with high sensitivity and accuracy in order to facilitate research exploring how these elements contribute to normal physiological functions and toxicity. Zinc (Zn), for example, is a cofactor in many mammalian proteins, participates in signaling events, and is a secondary messenger in cells. In excess, Zn is toxic and can inhibit absorption of other metals, while in deficit, it can lead to a variety of potentially lethal conditions.

Graphite furnace atomic absorption spectroscopy (GF-AAS) provides a highly sensitive and effective method for determining Zn and other transition metal concentrations in diverse biological samples. Electrothermal atomization via GF-AAS quantifies metals by atomizing small volumes of samples for subsequent selective absorption analysis using wavelength of excitation of the element of interest. Within the limits of linearity of the Beer-Lambert Law, the absorbance of light by the metal is directly proportional to concentration of the analyte. Compared to other methods of determining Zn content, GF-AAS detects both free and complexed Zn in proteins and possibly in small intracellular molecules with high sensitivity in small sample volumes. Moreover, GF-AAS is also more readily accessible than inductively coupled plasma mass spectrometry (ICP-MS) or synchrotron-based X-ray fluorescence. In this method, the systematic sample preparation of different cultured cell lines for analyses in a GF-AAS is described. Variations in this trace element were compared in both whole cell lysates and subcellular fractions of proliferating and differentiated cells as proof of principle.

Cultured primary or established cell lines are commonly used to address fundamental biological and mechanistic questions as an initial approach before using animal models. This protocol describes how to prepare whole cell extracts and subcellular fractions for studies of zinc (Zn) and other trace elements with atomic absorbance spectroscopy.

Transition metals are essential micronutrients for organisms but can be toxic to cells at high concentrations by competing with physiological metals in ...

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.

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. ...

Using an Extracellular Flux Analyzer to Measure Changes in Glycolysis and Oxidative Phosphorylation during Mouse Sperm Capacitation

Mammalian sperm acquire fertilization capacity in the female reproductive tract in a process known as capacitation. Capacitation-associated processes require energy. There remains an ongoing debate about the sources generating the ATP which fuels sperm progressive motility, capacitation, hyperactivation, and acrosome reaction. Here, we describe the application of an extracellular flux analyzer as a tool to analyze changes in energy metabolism during mouse sperm capacitation. Using H+- and O2- sensitive fluorophores, this method allows monitoring glycolysis and oxidative phosphorylation in real-time in non-capacitated versus capacitating sperm. Using this assay in the presence of different energy substrates and/or pharmacological activators and/or inhibitors can provide important insights into the contribution of different metabolic pathways and the intersection between signaling cascades and metabolism during sperm capacitation.

We describe the application of an extracellular flux analyzer to monitor real-time changes in glycolysis and oxidative phosphorylation during mouse sperm capacitation.

Mammalian sperm acquire fertilization capacity in the female reproductive tract in a process known as capacitation. Capacitation-associated processes ...

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming increasingly common. Ex vivo liver perfusions allow for a comprehensive analysis of liver metabolism using nuclear magnetic resonance (NMR), in nutritional conditions that can be rigorously controlled. As in silico simulations remain a primarily theoretical means of assessing hormone actions and the effects of pharmaceutical intervention, the perfused liver remains one of the most valuable test beds for understanding hepatic metabolism. As these studies guide basic insights into hepatic physiology, results must be accurate and reproducible. The greatest factor in the reproducibility of ex vivo hepatic perfusion is the quality of surgery. Therefore, we have introduced an organized and streamlined method to perform ex vivo mouse liver perfusions in the context of in situ NMR experiments. We also describe a unique application and discuss common issues encountered in these studies. The overall purpose is to provide an uncomplicated guide to a technique we have refined over several years that we deem the golden standard for obtaining reproducible results in hepatic resections and perfusions in the context of in situ NMR experiments. The distance to the center of the field for the magnet as well as the inaccessibility of the tissue to intervention during the NMR experiment makes our methods novel.

Ex Vivo Hepatic Perfusion Through the Portal Vein in Mouse

The protocol describes a straightforward method of resectioning an intact mouse liver for metabolic studies through portal vein perfusion.

Metabolic diseases such as diabetes, pre-diabetes, non-alcoholic fatty liver disease (NAFLD), and nonalcoholic steatohepatitis (NASH) are becoming ...

Stable Isotope Tracing & LC-MS Analysis to Measure DHODH and SDH Activities

Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in mammalian cells. As oxygen (O2) is the most ubiquitous terminal electron acceptor for the mammalian ETC, the O2 consumption rate is frequently used as a proxy for mitochondrial function. However, emerging research demonstrates that this parameter is not always indicative of mitochondrial function, as fumarate can be employed as an alternative electron acceptor to sustain mitochondrial functions in hypoxia. This article compiles a series of protocols that allow researchers to measure mitochondrial function independently of the O2 consumption rate. These assays are particularly useful when studying mitochondrial function in hypoxic environments. Specifically, we describe methods to measure mitochondrial ATP production, de novo pyrimidine biosynthesis, NADH oxidation by complex I, and superoxide production. In combination with classical respirometry experiments, these orthogonal and economical assays will provide researchers with a more comprehensive assessment of mitochondrial function in their system of interest.

Author Spotlight: Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals  

Here, we present a compilation of assays to directly measure mitochondrial function in mammalian cells independently of their ability to consume molecular oxygen.

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in ...

Exploring Metabolite Exchange in Brown Adipose Tissue

Arteriovenous Metabolomics to Measure In Vivo Metabolite Exchange in Brown Adipose Tissue

Brown adipose tissue (BAT) plays a crucial role in regulating metabolic homeostasis through a unique energy expenditure process known as non-shivering thermogenesis. To achieve this, BAT utilizes a diverse menu of circulating nutrients to support its high metabolic demand. Additionally, BAT secretes metabolite-derived bioactive factors that can serve as either metabolic fuels or signaling molecules, facilitating BAT-mediated intratissue and/or intertissue communication. This suggests that BAT actively participates in systemic metabolite exchange, an interesting feature that is beginning to be explored. Here, we introduce a protocol for in vivo mouse-level optimized BAT arteriovenous metabolomics. The protocol focuses on relevant methods for thermogenic stimulations and an arteriovenous blood sampling technique using Sulzer's vein, which selectively drains interscapular BAT-derived venous blood and systemic arterial blood. Next, a gas chromatography-based metabolomics protocol using those blood samples is demonstrated. The use of this technique should expand the understanding of BAT-regulated metabolite exchange at the inter-organ level by measuring the net uptake and release of metabolites by BAT.

Author Spotlight: Illuminating New Avenues for Adipose Tissue Metabolism and Disease Prevention 

In this protocol, methods relevant for BAT-optimized arteriovenous metabolomics using GC-MS in a mouse model are outlined. These methods allow for the acquisition of valuable insights into BAT-mediated metabolite exchange at the organismal level.

Brown adipose tissue (BAT) plays a crucial role in regulating metabolic homeostasis through a unique energy expenditure process known as non-shivering ...