We are grateful for the partial donations from the Tecnol&#243;gico Nacional de M&#233;xico in the 2023 and 2024 Call for Proposals for Scientific Research, Technological Development (16432.23-P y 19545.24-P). We would like to thank the National Council for Humanities, Sciences, and Technologies (CONAHCyT) for the scholarship received (No. 832315) during the doctoral studies (IHRH), and the participation and collaboration of Wendolyne Monroy-Mart&#237;nez is gratefully acknowledged. Figure 1 was created with BioRender.com.

1. Solution preparation
<ol>
	<li>Prepare 100 mL of 0.1 M phosphate buffer. Weigh 1.28 g of potassium dihydrogen phosphate (KH2PO4) and 0.1304 g of dipotassium phosphate (K2HPO4) and add them to a glass beaker. Now, add 70 mL of deionized water (H2Odi) and adjust the pH to 5.5 using 1 M hydrochloric acid (HCl) or potassium hydroxide (KOH). Transfer the solution to a volumetric flask and adjust the final volume to 100 mL. Then, transfer the solution to an amber container and store the solution at 4-8 &#176;C for up to 3 months. 
	CAUTION: Wear nitrile gloves throughout the preparation process to prevent skin contact and potential contamination, as o-dianisidine dihydrochloride is potentially carcinogenic.</li>
	<li>Prepare a 1% w/v solution of o-dianisidine dihydrochloride. Weigh 0.01 g of o-dianisidine dihydrochloride and place it in a 2.0 mL centrifuge tube. Using a 1,000 mL micropipette, add 1.0 mL of H2Odi to dissolve the compound, then mix slowly by pipetting up and down. Shield the solution from light by wrapping the container in aluminum foil and store it at -20 &#176;C to maintain stability. 
	NOTE: To facilitate support of the tube during measurement on the analytical balance, a 10 mL beaker can be used as additional support. This helps to stabilize the tube, ensuring accurate measurement by minimizing the risk of displacement or tilting that could alter the results obtained on the balance.</li>
	<li>In a 100 mL volumetric flask, add 10 mL of phosphate buffer solution followed by 1 mL of the 1% v/v o-dianisidine dihydrochloride solution. Then, adjust the final volume to 100 mL with phosphate buffer to obtain a final concentration of 0.01% o-dianisidine-buffer mixture. Transfer the solution to an amber flask and wrap it with aluminum foil to protect it from light. Store the solution at 4-8 &#176;C for up to 3-4 months. 
	NOTE: Refrigeration is essential to prevent decomposition, which could compromise the integrity of experimental measurements. To avoid degradation of the o-dianisidine-buffer mixture, a small portion of 13 mL (sufficient for 12 samples) can be placed in a 14 mL plastic tube on ice until use.</li>
	<li>Prepare 50% H2SO4 solution. Place a 100 mL volumetric flask with 30 mL of H2Odi in an ice bath and allow it to cool for 10 min. Then, slowly pour 51 mL of H2SO4 (98% v/v) down the walls of the flask, shaking carefully to homogenize the solution. Wait for the mixture to cool down, as the reaction is exothermic (generates heat). Adjust the final volume to 100 mL with H2Odi and transfer the solution to a glass amber flask. 
	NOTE: Use a fume hood and follow all safety protocols, be sure to wear appropriate protective equipment. Prepare 50 mL of 40% calcium carbonate solution to neutralize any possible spills that may occur.</li>
	<li>Prepare 50 mM sodium acetate solution by dissolving 0.04 g of sodium acetate in 5 mL of H2Odi in a beaker. Adjust the pH to 5.1 using glacial acetic acid. Finally, adjust the volume to 10 mL with H2Odi.</li>
</ol>
2. Preparation of enzymes
<ol>
	<li>In a 2.0 mL sterile microtube, weigh 0.01 g of glucose oxidase and mix with 1 mL of 50 mM sodium acetate, pH 5.0, to obtain a final concentration of 10 mg/mL. Cover the tube with aluminum foil and store at -20 &#176;C. 
	NOTE: The solution can be stored for up to 2 months at -20 &#176;C.</li>
	<li>Calculate the volume of enzyme solution V2 needed to achieve the desired enzyme activity in each plastic cuvette (with a total volume of 1.5 mL) using Eq (1): V1C1 = V2C2, where V1 is 1,500 &#181;L, C1 is 28.5 U of enzyme activity, and C2 (the concentration of the enzyme solution) is 12,970 U/mL (10 mg of the enzyme [129,000 units/g] in 1 mL of H2Odi). 
	V2 = (1,500 &#181;L &#215; 28.5 U)/12,970 U = 3.29 &#181;L&#160; &#160; (1) 
	NOTE: For more accurate pipetting, the value can be rounded to 3.3 &#181;L.</li>
	<li>In a new 2 mL microtube, weigh 0.0031 g of peroxidase enzyme and add 1 mL of phosphate buffer, pH 5.5, to achieve a final concentration of 3.1 mg/mL. Cover the microtube with aluminum foil and store at -20 &#176;C.</li>
	<li>Calculate the volume of the peroxidase solution required to achieve the desired enzyme activity per cuvette using Eq (1). Start with a total cuvette volume of 1,500 &#181;L and a target enzyme activity of 1.25 U. Then, determine the necessary volume of the enzyme solution, given its concentration of 1,551 U. 
	V2 = (1,500 &#181;L &#215; 1.25 U)/1,551 U = 1.208 &#181;L 
	NOTE: For more accurate pipetting, the value was rounded to 1.20 &#956;L.</li>
</ol>
3. Glucose standard
<ol>
	<li>Prepare a 1 g/L D-glucose standard by adding 0.01 g of D-glucose to 10 mL of H2Odi in a volumetric flask. To dilute the concentration to 0.5 g/L, take 500 &#181;L of the stock solution and mix it with 500 &#181;L of H2Odi to achieve a final volume of 1 mL. 
	NOTE: This dilution process is essential for creating accurate standard solutions.</li>
</ol>
4. Standard curve preparation
<ol>
	<li>Using a new 2 mL microtube, add the buffer and glucose volumes indicated in Table 1.</li>
	<li>Set the dry thermobath to 37 &#176;C. Once the temperature is stable, add 3.3 &#181;L of glucose oxidase to all the tubes and then add 1.2 &#181;L of peroxidase to each tube. Incubate the tubes at 37 &#176;C for 20 min.</li>
	<li>After incubation, immediately add 750 &#181;L of 50% H2SO4 (v/v) to each microtube and let the mixture cool in an ice bath for 2 min. This results in a final volume of 1,500 &#181;L. Finally, transfer the solution to a plastic cuvette and measure the absorbance at 529 nm.</li>
</ol>
Table 1: Glucose calibration curve for the GOX-H2SO4 method. Abbreviations: GOD = glucose oxidase; POD = peroxidase; H2SO4 = sulfuric acid. <a href="https://www.jove.com/files/ftp_upload/67126/67126_Table 1.xlsx" target="_blank">Please click here to download this Table.</a>
5. Testing microbiological samples
<ol>
	<li>Obtain the sample by growing bacteria or yeast in a liquid culture medium under specific conditions, including temperature, agitation speed, and incubation time. Once the culture reaches the desired optical density (OD) or cell concentration, collect the culture for further processing. 
	NOTE: Pseudomonas reptilivora was cultured at a constant agitation speed of 300 rpm and 28 &#176;C, while Debaryomyces hansenii was cultured with an agitation speed of 200 rpm at 30 &#176;C in an orbital shaker. In the specific case of D. hansenii, edible canola oil was also used as an inducer for lipase production.</li>
	<li>Clean the work area with a 70% alcohol solution or a suitable cleaning agent according to biosafety protocols. Light a burner to ensure asepsis.</li>
	<li>Transfer 100 &#181;L of the culture media into a 1.5 mL or 2 mL micro tube using sterile tips.</li>
	<li>Centrifuge the samples at 7,500 &#215; g for 7 min at 4 &#176;C, or at 10,000 &#215; g for 7 min without temperature control. After centrifugation, carefully remove the supernatant, which contains glucose in solution, and discard the pellet.</li>
	<li>Use 0.5 &#181;L of the supernatant for glucose concentrations ranging from 1 to 20 g/L, then mix with 749.5 &#181;L of o-dianisidine-buffer, 3.3 &#181;L of GOD, and then 1.2 &#181;L of POD. Incubate the microtubes for 20 min at 37 &#176;C, as detailed in step 4.2 and immediately add 750 &#181;L of 50% H2SO4 and let it cool in an ice bath for 2 min.
	<ol>
		<li>For concentrations above 20 g/L, perform a 1:1 dilution with sterile H2Odi before proceeding.</li>
		<li>If the supernatant samples were previously frozen at -80 &#176;C or -20 &#176;C, thaw them at room temperature and vortex for 30 s before use. If dilution is necessary, use sterile H2Odi.</li>
		<li>For samples with glucose concentrations above 30 g/L, perform a 1:1 dilution with sterile H2Odi, resulting in a dilution factor of 2.</li>
	</ol>
	</li>
	<li>Calculate the total glucose concentration in each sample using Eq (2) in a spreadsheet program. 
	<img alt="Equation 11" src="/files/ftp_upload/67126/67126eq11.jpg" style="margin: auto;" />&#160; &#160; &#160;&#8203;(2) 
	Where: 
	x = concentration of glucose (g/L)&#160; &#160; &#160;&#948; = Final reaction volume (0.0015 L) 
	y = absorbance&#160; &#160; &#160; &#160;M.S. = amount of sample used* 
	NOTE: The b and m values are from the equation that models the trend line of the calibration curve for the method (y = mx + b). *The M.S. value must be based on the amount of sample used to obtain the ratio at the corresponding dilution (i.e., if you take 0.5 &#181;L of the supernatant, M.S. = 0.0000005 L = 0.5 &#215; 10-6 L); if a dilution factor (DF) is used, multiply the obtained absorbance by the DF. 
	For example, if an absorbance of 0.5 is obtained and a DF of 2 was used, the result would be 1.0. Therefore, y = 1.0, and this value should be entered into Eq (2), as previously described.</li>
</ol>
6. Analytical validation
<ol>
	<li>Calculate the analytical validation parameters of the GOX-H2SO4 method according to what is established by the National Metrology Center and the Mexican Accreditation Entity10 (CENAM-ema).
	<ol>
		<li>Calculate precision as a percentage of the coefficient of variation or %CV = (S/x) &#215; 100, where x is the sample group mean and S is the standard deviation with an acceptance value of &#8804;10%.</li>
		<li>Determine linearity by fitting the standard curve to linear model R2 above 0.995.</li>
		<li>Calculate the limit of quantification (LOQ) using the equation: LOQ = yB + 10sB, where yB represents the analyte concentration that produces a signal equivalent to the target signal, and 10sB denotes 10x the standard deviation of the blank</li>
		<li>Calculate the systematic error using this equation: Slant = x - m, where x = average of the experimental concentration and m is the theoretical concentration.</li>
		<li>Determine accuracy as the degree of agreement between a real value and a theoretical value. 
		Recovery% = (CR/CV) &#215; 100 
		Where CR is the average of the experimental concentration and CV is the coefficient of variation of the theoretical concentration.</li>
	</ol>
	</li>
</ol>
7. Interference by other reducing sugars
<ol>
	<li>Evaluate four reducing sugars separately: glucose, fructose, galactose, and xylose from a 0.5 g/L stock solution. Additionally, use trehalose as a negative control. Follow the protocol for all samples and measure the absorbance at 545 and 529 nm.</li>
</ol>

Enzymatic Bergmeyer Glucose Quantification for Microbial and Yeast Analysis

<ol>
	<li>Huang, W. C., Tang, I. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+and+yeast+cultures:+process+characteristics,+products,+and+applications.+Bioprocessing+for+value-added+products+from+renewable+resources.">Bacterial and yeast cultures: process characteristics, products, and applications. Bioprocessing for value-added products from renewable resources.</a> New Technologies and Applications. , 185-223 (2007).</li><li>Hernandez-Lopez, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+reducing+sugars+based+on+the+qualitative+technique+of+Benedict.">Quantification of reducing sugars based on the qualitative technique of Benedict.</a> ACS Omega. 5 (50), 32403-32410 (2020).</li><li>Miller, G. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+dinitrosalicylic+acid+reagent+for+determination+of+reducing+sugar.">Use of dinitrosalicylic acid reagent for determination of reducing sugar.</a> Anal Chem. 31 (3), 426-428 (1959).</li><li>Deshavath, N. N., Mukherjee, G., Goud, V. V., Veeranki, V. D., Sastri, C. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pitfalls+in+the+3,5-dinitrosalicylic+acid+(DNS)+assay+for+the+reducing+sugars:+interference+of+furfural+and+5-hydroxymethylfurfural.">Pitfalls in the 3,5-dinitrosalicylic acid (DNS) assay for the reducing sugars: interference of furfural and 5-hydroxymethylfurfural.</a> Int J Biol Macromol. 156, 180-185 (2020).</li><li>De Abreu, J. R., Santos, C. D. D., De Abreu, C. M. P., Corr&#234;a, A. D., De Oliveira Lima, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sugar+fractionation+and+pectin+content+during+the+ripening+of+guava+cv.+Pedro+Sato.">Sugar fractionation and pectin content during the ripening of guava cv. Pedro Sato.</a> Food Sci Technol. 32 (1), 156-162 (2020).</li><li>Dubois, M., Gilles, K. -. A., Hamilton, J. K., Roberts, P. A., Smith, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+colorimetric+method+for+the+determination+of+sugars.">A colorimetric method for the determination of sugars.</a> Nature. 168 (4265), 167 (1951).</li><li>Gonzalez, N. M., Fitch, A., Al-Bazi, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+RP-HPLC+method+for+determination+of+glucose+in+Shewanella+oneidensis+cultures+utilizing+1-phenyl-3-methyl-5-pyrazolone+derivatization.">Development of a RP-HPLC method for determination of glucose in Shewanella oneidensis cultures utilizing 1-phenyl-3-methyl-5-pyrazolone derivatization.</a> PLoS ONE. 15 (3), e0229990 (2020).</li><li>Galant, A. L., Kaufman, R. C., Wilson, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glucose:+Detection+and+analysis.">Glucose: Detection and analysis.</a> Food Chem. 188, 149-160 (2015).</li><li>Bergmeyer, H. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Methods of enzymatic analysis. 2, (2012).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Gu&#237;a t&#233;cnica sobre trazabilidad e incertidumbre en las mediciones anal&#237;ticas que emplea la t&#233;cnica de espectrofotometr&#237;a de ultravioleta-visible. , (2008).</li><li>Yuen, V. G., McNeill, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+the+glucose+oxidase+method+for+glucose+determination+by+manual+assay+and+automated+analyzer.">Comparison of the glucose oxidase method for glucose determination by manual assay and automated analyzer.</a> J Pharmacol Toxicol Methods. 44 (3), 543-546 (2000).</li><li>Hansen, O. <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+glucose+oxidase+reaction+and+interference+with+quantitative+glucose+oxidase-peroxidase-O-dianisidine+method.">Specificity of glucose oxidase reaction and interference with quantitative glucose oxidase-peroxidase-O-dianisidine method.</a> Scand J Clin Lab Investig. 14 (6), 651-655 (1962).</li><li>Kumar, V., Gill, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimation+of+blood+glucose+levels+by+glucose+oxidase+method.">Estimation of blood glucose levels by glucose oxidase method.</a> Basic concepts in clinical biochemistry: A practical guide. , 57-60 (2018).</li></ol>

The authors declare no conflicts of interest

Quantifying glucose in culture media is essential for understanding microbial growth and metabolic activity, as glucose serves as a primary energy source for many microorganisms. In this study, we employed the GOX-H2SO4 method to accurately measure glucose levels in culture media, aiming to optimize microbial processes in bacteria or yeast fermentations. Our findings are consistent with those of Yuen and McNeill11; however, unlike them, our findings indicate that the readings...

Glucose serves as the primary energy and carbon source for numerous microorganisms, including bacteria, yeasts, and fungi. These microorganisms take up glucose through transporters located on the extracellular membrane, where it undergoes a series of biochemical reactions within the glycolysis metabolic pathway to be converted into pyruvate1. There are various techniques for the quantification of reducing sugars such as glucose. For example, Benedict&#39;s reagent2, the use of 3,5-dinitrosalicylic acid (DNS)3,4, the anthrona method5, or phenol-sulfuric acid6 methods can be employed. However, these methods detect any reducing sugar present in the sample, such as fructose and maltose, which can contribute to the signal and complicate the accurate quantification of a particular sugar.
In addition, they require strict temperature control and the handling of hazardous substances, which can complicate the process and affect the accuracy. These methods can also suffer interference from other compounds present in the sample, making accurate glucose quantification difficult. Conversely, high-performance liquid chromatography (HPLC) offers a more precise alternative, allowing for the discrimination between glucose and other reducing sugars, albeit with higher operational costs. These contrasting methods highlight the ongoing need for the development of accurate and cost-effective glucose quantification techniques7.
The glucose oxidase-peroxidase-sulfuric acid (GOX-H2SO4) method employs two enzymes, glucose oxidase (GOD) and peroxidase (POD). GOD is an oxidoreductase enzyme that is very selective for the oxidation of &#946;-D-glucose8. This complex acts as a catalyst during the reaction that occurs when glucose is in the presence of oxygen, producing gluconic acid and hydrogen peroxide (H2O2). H2O2 is detected by means of a chromogenic oxygen acceptor, o-dianisidine, which, upon interaction with POD, causes its oxidation and H2O2 release, preventing gluconic acid from being converted back to glucose (see Figure 1). The color obtained is stabilized by the addition of sulfuric acid (H2SO4), which also stops the enzymatic reaction, thus avoiding possible interference that could occur if the reaction continues9.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67126/67126fig01.jpg" /> 
Figure 1: Overview of the glucose quantification method using the glucose oxidase enzyme. which reacts with glucose to produce hydrogen peroxide (H2O2) and gluconic acid. Subsequently, the peroxidase enzyme reacts with H2O2 in the presence of O-dianisidine dihydrochloride. In the final step, sulfuric acid (H2SO4) is added to stop the enzymatic reaction, resulting in a pink color that is measured at 529 nm. Abbreviations: POD = peroxidase; GOD = glucose oxidase. <a href="https://www.jove.com/files/ftp_upload/67126/67126fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The GOX-H2SO4 enzymatic method is a technique widely used to measure the concentration of glucose in biological samples, most of which are of clinical interest. However, few studies have investigated a technique that allows the quantification of the amount of glucose in a growth medium to evaluate its consumption. Therefore, in the present protocol, the methodology to quantify glucose by means of the enzymatic reaction of glucose oxidase, peroxidase, o-dianisidine dihydrochloride, and H2SO4 in microbiological liquid samples is presented, which arises as a modification of the work done by Bergmeyer9, by using 50% (v/v) H2SO4 and lower quantities of reagents.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5 mL microtube</td>
			<td>Eppendorf</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>2.0 mL microtube</td>
			<td>Eppendorf</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>CaCO3</td>
			<td>Meyer</td>
			<td>471-34-1</td>
			<td>Calcium Carbonate</td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>Meyer</td>
			<td>50-99-7</td>
			<td>D-Glucose&#160;</td>
		</tr>
		<tr>
			<td>Drybath</td>
			<td>FisherScientific</td>
			<td>11-718-4</td>
			<td>Serial 911NO251. Block WxDxH mm/in: 124 x 76 x 39 / 4.9 x 3.0 x 1.5</td>
		</tr>
		<tr>
			<td>Glucose oxidase&#160;</td>
			<td>Sigma Aldrich</td>
			<td>G6125-50KU</td>
			<td>Enzyme powder</td>
		</tr>
		<tr>
			<td>H2O</td>
			<td>-</td>
			<td>-</td>
			<td>Deionized water</td>
		</tr>
		<tr>
			<td>H2SO4</td>
			<td>Meyer</td>
			<td>7664-93-9</td>
			<td>Sulfuric acid</td>
		</tr>
		<tr>
			<td>HCl</td>
			<td>Meyer</td>
			<td>7647-01-0</td>
			<td>Hydrochloridric acid&#160;</td>
		</tr>
		<tr>
			<td>K2HPO4</td>
			<td>Meyer</td>
			<td>7758-11-4</td>
			<td>Dipotassium phosphate</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Meyer</td>
			<td>7778-77-0</td>
			<td>Monopotassium phosphate</td>
		</tr>
		<tr>
			<td>KOH</td>
			<td>Meyer</td>
			<td>1310-58-3</td>
			<td>Potassium hydroxide</td>
		</tr>
		<tr>
			<td>Lambda 35</td>
			<td>PerkinElmer</td>
			<td>-</td>
			<td>Spectrophotometer</td>
		</tr>
		<tr>
			<td>Microtube centrifuge</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>o-Dianisidine dihydrochloride</td>
			<td>Sigma Aldrich</td>
			<td>D3252</td>
			<td>Chromogenic</td>
		</tr>
		<tr>
			<td>Peroxidase</td>
			<td>Sigma Aldrich</td>
			<td>P8125-50KU</td>
			<td>Enzyme powder</td>
		</tr>
		<tr>
			<td>pH-meter</td>
			<td>Hanna&#160;</td>
			<td>1131</td>
			<td>Hanna 1170</td>
		</tr>
		<tr>
			<td>Sodium acetate</td>
			<td>Meyer</td>
			<td>127-09-3</td>
			<td>Meyer</td>
		</tr>
		<tr>
			<td>Weighing spatulas&#160;</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
	</tbody>
</table>

bergmeyer glucose quantification for microbiological samples

The glucose concentration is a key indicator of cellular metabolism and could indicate the total metabolism rate, an aberration in glucose metabolism, and, in some cases, how cells couple glucose metabolism to energetic metabolism. In addition, intracellular and extracellular glucose levels are indicative of the cellular metabolic status. Enzymatic techniques, such as Bergmeyer glucose quantification, are more accurate and sensitive than other techniques, such as dinitrosalicylic acid or fluorescence methods, which are usually utilized in microbiology. Although mainly used in the clinical area, Bergmeyer glucose quantification can also be applied to any cell but has not been reported in detail for bacteria, fungi, yeasts, or other microorganisms. 
Herein, we present a methodology to quantify glucose from bacteria and yeast samples using the enzymatic Bergmeyer glucose quantification method. The procedure involved the enzymes glucose oxidase, peroxidase, and o-dianisidine dihydrochloride incubated at 37 &#176;C for 20 min, followed by the addition of sulfuric acid. The absorbance is then measured at 545 nm. It is important to highlight that although this technique presents difficulties in measuring high concentrations of glucose (above 60 g/L), it is possible to measure concentrations below 50 g/L using dilution factors. This enzymatic approach is valuable for research and analysis in microbiology and other scientific areas. The precision and sensitivity of the method make it helpful for detecting even low concentrations of glucose in microbiological samples.

The spectrophotometric analysis of GOX-H2SO4 revealed a single maximum absorbance at 529 nm (&#955;max) (Figure&#160;2A), with additional absorbance values observed at 545 nm. By analyzing the absorbance values at different glucose concentrations, we obtained a linearity (R&#178;) of 0.9977 for &#955;529 nm and R&#178; of 0.9967 for &#955;545 nm (Figure 2B). Linearity, within a given range, refers to the ability to provide results direc...

Bergmeyer Glucose Quantification for Microbiological Samples

Bergmeyer glucose quantification is a spectrophotometric enzymatic technique mainly used for clinical tests that accurately and sensitively measure the amount of glucose. Herein, we present a protocol for using this method for microbiological samples.

The glucose concentration is a key indicator of cellular metabolism and could indicate the total metabolism rate, an aberration in glucose metabolism, ...

bergmeyer-glucose-quantification-for-microbiological-samples

Research

JoVE Journal

Biochemistry

273 Views.  Tecnológico Nacional de México/Instituto Tecnológico de Morelia. Bergmeyer glucose quantification is a spectrophotometric enzymatic technique mainly used for clinical tests that accurately and sensitively measure the amount of glucose. Herein, we present a protocol for using this method for microbiological samples.

Video: Bergmeyer Glucose Quantification for Microbiological Samples

Preparation of Homogeneous MALDI Samples for Quantitative Applications

This protocol demonstrates a simple sample preparation to reduce spatial heterogeneity in ion signals during matrix-assisted laser desorption/ionization (MALDI) mass spectrometry. The heterogeneity of ion signals is a severe problem in MALDI, which results in poor data reproducibility and makes MALDI unsuitable for quantitative analysis. By regulating sample plate temperature during sample preparation, thermal-induced hydrodynamic flows inside droplets of sample solution are able to reduce the heterogeneity problem. A room-temperature sample preparation chamber equipped with a temperature-regulated copper base block that holds MALDI sample plates facilitates precise control of the sample drying condition. After drying of sample droplets, the temperature of sample plates is returned to room temperature and removed from the chamber for subsequent mass spectrometric analysis. The areas of samples are examined with MALDI-imaging mass spectrometry to obtain the spatial distribution of all components in the sample. In comparison with the conventional dried-droplet method that prepares samples under ambient conditions without temperature control, the samples prepared with the method demonstrated herein show significantly better spatial distribution and signal intensity. According to observations using carbohydrate and peptide samples, decreasing substrate temperature while maintaining the surroundings at ambient temperature during the drying process can effectively reduce the heterogeneity of ion signals. This method is generally applicable to various combinations of samples and matrices.

A protocol for reducing spatial heterogeneities of ion signals in MALDI mass spectrometry by regulating substrate temperature during sample drying processes is demonstrated.

This protocol demonstrates a simple sample preparation to reduce spatial heterogeneity in ion signals during matrix-assisted laser desorption/ionization ...

Isolation of High-density Lipoproteins for Non-coding Small RNA Quantification

The diversity of small non-coding RNAs (sRNA) is rapidly expanding and their roles in biological processes, including gene regulation, are emerging. Most interestingly, sRNAs are also found outside of cells and are stably present in all biological fluids. As such, extracellular sRNAs represent a novel class of disease biomarkers and are likely involved in cell signaling and intercellular communication networks. To assess their potential as biomarkers, sRNAs can be quantified in plasma, urine, and other fluids. Nevertheless, to fully understand the impact of extracellular sRNAs as endocrine signals, it is important to determine which carriers are transporting and protecting them in biological fluids (e.g., plasma), which cells and tissues contribute to extracellular sRNA pools, and cells and tissues capable of accepting and utilizing extracellular sRNA. To accomplish these goals, it is critical to isolate highly pure populations of extracellular carriers for sRNA profiling and quantification. We have previously demonstrated that lipoproteins, particularly high-density lipoproteins (HDL), transport functional microRNAs (miRNA) between cells and HDL-miRNAs are significantly altered in disease. Here, we detail a new protocol that utilizes tandem HDL isolation with density-gradient ultracentrifugation (DGUC) and fast-protein-liquid chromatography (FPLC) to obtain highly pure HDL for downstream profiling and quantification of all sRNAs, including miRNAs, using both high-throughput sequencing and real-time PCR approaches. This protocol will be a valuable resource for the investigation of sRNAs on HDL.

This protocol describes the isolation and quantification of high-density lipoprotein small RNAs.

The diversity of small non-coding RNAs (sRNA) is rapidly expanding and their roles in biological processes, including gene regulation, are emerging. Most ...

Sheathless Capillary Electrophoresis&#8211;Mass Spectrometry for Metabolic Profiling of Biological Samples

In metabolomics, a wide range of analytical techniques is used for the global profiling of (endogenous) metabolites in complex samples. In this paper, a protocol is presented for the analysis of anionic and cationic metabolites in biological samples by capillary electrophoresis&#8211;mass spectrometry (CE-MS). CE is well-suited for the analysis of highly polar and charged metabolites as compounds are separated on the basis of their charge-to-size ratio. A recently developed sheathless interfacing design, i.e., a porous tip interface, is used for coupling CE to electrospray ionization (ESI) MS. This interfacing approach allows the effective use of the intrinsically low-flow property of CE in combination with MS, resulting in nanomolar detection limits for a broad range of polar metabolite classes. The protocol presented here is based on employing a bare fused-silica capillary with a porous tip emitter at low-pH separation conditions for the analysis of a broad array of metabolite classes in biological samples. It is demonstrated that the same sheathless CE-MS method can be used for the profiling of cationic metabolites, including amino acids, nucleosides and small peptides, or anionic metabolites, including sugar phosphates, nucleotides and organic acids, by only switching the MS detection and separation voltage polarity. Highly information-rich metabolic profiles in various biological samples, such as urine, cerebrospinal fluid and extracts of the glioblastoma cell line, can be obtained by this protocol in less than 1 hr of CE-MS analysis.

Sheathless Capillary Electrophoresis&amp;#8211;Mass Spectrometry for Metabolic Profiling of Biological Samples

A protocol for metabolic profiling of biological samples by capillary electrophoresis&#8211;mass spectrometry using a sheathless porous tip interface design is presented.

In metabolomics, a wide range of analytical techniques is used for the global profiling of (endogenous) metabolites in complex samples. In this paper, a ...

Multimer-PAGE: A Method for Capturing and Resolving Protein Complexes in Biological Samples

There are many well-developed methods for purifying and studying single proteins and peptides. However, most cellular functions are carried out by networks of interacting protein complexes, which are often difficult to investigate because their binding is non-covalent and easily perturbed by purification techniques. This work describes a method of stabilizing and separating native protein complexes from unmodified tissue using two-dimensional polyacrylamide gel electrophoresis. Tissue lysate is loaded onto a non-denaturing blue-native polyacrylamide gel, then an electric current is applied until the protein migrates a short distance into the gel. The gel strip containing the migrated protein is then excised and incubated with the amine-reactive cross-linking reagent dithiobis(succinimidyl propionate), which covalently stabilizes protein complexes. The gel strip containing cross-linked complexes is then cast into a sodium dodecyl sulfate polyacrylamide gel, and the complexes are separated completely. The method relies on techniques and materials familiar to most molecular biologists, meaning it is inexpensive and easy to learn. While it is limited in its ability to adequately separate extremely large complexes, and has not been universally successful, the method was able to capture a wide variety of well-studied complexes, and is likely applicable to many systems of interest.

A method for stabilizing and separating native protein complexes from unmodified tissue lysate using an amine-reactive protein cross-linker coupled to a novel two-dimensional polyacrylamide gel electrophoresis (PAGE) system is presented.

There are many well-developed methods for purifying and studying single proteins and peptides. However, most cellular functions are carried out by ...

Quantification of Hypopigmentation Activity In Vitro

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

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

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

Proteome-wide Quantification of Labeling Homogeneity at the Single Molecule Level

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a fluorescent dye and are spotted by a photodetector following their separation. Single molecule fluorescence imaging can provide ultrasensitive protein detection with its ability for visualizing individual fluorescent molecules. However, the application of this powerful imaging method to electrophoresis assays is hampered by the lack of ways to characterize the homogeneity of fluorescent labeling of each protein species across the proteome. Here, we developed a method to evaluate the labeling homogeneity across the proteome based on a single molecule fluorescence imaging assay. In our measurement using a HeLa cell sample, the proportion of proteins labeled with at least one dye, which we termed &#8216;labeling occupancy&#8217; (LO), was determined to range from 50% to 90%, supporting the high potential of the application of single molecule imaging to sensitive and precise proteome analysis.

Here, we present a protocol to assess the labeling homogeneity for each protein species in a complex protein sample at the single molecule level.

Cell proteomes are often characterized using electrophoresis assays, where all species of proteins in the cells are non-specifically labeled with a ...

RNA Blot Analysis for the Detection and Quantification of Plant MicroRNAs

MicroRNAs (miRNAs) are a class of endogenously expressed non-coding, ~21 nt small RNAs involved in the regulation of gene expression in both plants and animals. Most miRNAs act as negative switches of gene expression targeting key genes. In plants, primary miRNAs (pri-miRNAs) transcripts are generated by RNA polymerase II, and they form varying lengths of stable stem-loop structures called pre-miRNAs. An endonuclease, Dicer-like1, processes the pre-miRNAs into miRNA-miRNA* duplexes. One of the strands from miRNA-miRNA* duplex is selected and loaded onto Argonaute 1 protein or its homologs to mediate the cleavage of target mRNAs. Although miRNAs are key signaling molecules, their detection is often carried out by less than optimal PCR-based methods instead of a sensitive northern blot analysis. We describe a simple, reliable, and extremely sensitive northern method that is ideal for the quantification of miRNA levels with very high sensitivity, literally from any plant tissue. Additionally, this method can be used to confirm the size, stability and the abundance of miRNAs and their precursors.

This method demonstrates use of the northern hybridization technique to detect miRNAs from total RNA extract.

MicroRNAs (miRNAs) are a class of endogenously expressed non-coding, ~21 nt small RNAs involved in the regulation of gene expression in both plants and ...

Visualization and Quantification of TGF&#946;/BMP/SMAD Signaling under Different Fluid Shear Stress Conditions using Proximity-Ligation-Assay

Transforming Growth Factor &#946; (TGF&#946;)/Bone Morphogenetic Protein (BMP) signaling is tightly regulated and balanced during the development and homeostasis of the vasculature system Therefore, deregulation in this signaling pathway results in severe vascular pathologies, such as pulmonary artery hypertension, hereditary hemorrhagic telangiectasia, and atherosclerosis. Endothelial cells (ECs), as the innermost layer of blood vessels, are constantly exposed to fluid shear stress (SS). Abnormal patterns of fluid SS have been shown to enhance TGF&#946;/BMP signaling, which, together with other stimuli, induce atherogenesis. In relation to this, atheroprone, low laminar SS was found to enhance TGF&#946;/BMP signaling while atheroprotective, high laminar SS, diminishes this signaling. To efficiently analyze the activation of these pathways, we designed a workflow to investigate the formation of transcription factor complexes under low laminar SS and high laminar SS conditions using a commercially available pneumatic pump system and proximity ligation assay (PLA).
Active TGF&#946;/BMP-signaling requires the formation of trimeric SMAD complexes consisting of two regulatory SMADs (R-SMAD); SMAD2/3 and SMAD1/5/8 for TGF&#946; and BMP signaling, respectively) with a common mediator SMAD (co-SMAD; SMAD4). Using PLA targeting different subunits of the trimeric SMAD-complex, i.e., either R-SMAD/co-SMAD or R-SMAD/R-SMAD, the formation of active SMAD transcription factor complexes can be measured quantitatively and spatially using fluorescence microscopy.
The usage of flow slides with 6 small parallel channels, that can be connected in series, allows for the investigation of the transcription factor complex formation and inclusion of necessary controls. 
The workflow explained here can be easily adapted for studies targeting the proximity of SMADs to other transcription factors or to transcription factor complexes other than SMADs, in different fluid SS conditions. The workflow presented here shows a quick and effective way to study the fluid SS induced TGF&#946;/BMP signaling in ECs, both quantitatively and spatially.

Here, we establish a protocol to simultaneously visualize and analyze multiple SMAD complexes using proximity ligation assay (PLA) in endothelial cells exposed to pathological and physiological fluid shear stress conditions.

Transforming Growth Factor &#946; (TGF&#946;)/Bone Morphogenetic Protein (BMP) signaling is tightly regulated and balanced during the development and ...

Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This enables quantifications of ultrastructural aspects such as volume fractions, surface area to volume ratios, morphometry, and physical contact sites of different subcellular structures. In the 1970s, a protocol for enhanced staining of glycogen in cells was developed and paved the way for a string of studies on the subcellular localization of glycogen and glycogen particle size using transmission electron microscopy. While most analyses interpret glycogen as if it is homogeneously distributed within the muscle fibers, providing only a single value (e.g., an average concentration), transmission electron microscopy has revealed that glycogen is stored as discrete glycogen particles located in distinct subcellular compartments. Here, the step-by-step protocol from tissue collection to the quantitative determination of the volume fraction and particle diameter of glycogen in the distinct subcellular compartments of individual skeletal muscle fibers is described. Considerations on how to 1) collect and stain tissue specimens, 2) perform image analyses and data handling, 3) evaluate the precision of estimates, 4) discriminate between muscle fiber types, and 5) methodological pitfalls and limitations are included.

A modified post-fixation procedure increases the contrast of glycogen particles in tissue. This paper provides a step-by-step protocol describing how to handle the tissue, conduct the imaging, and use stereological methods to obtain unbiased and quantitative data on fiber type-specific subcellular glycogen distribution in skeletal muscle.

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This ...

An Optimized Single-Molecule Pull-Down Assay for Quantification of Protein Phosphorylation

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to measure protein phosphorylation cannot preserve the heterogeneity in phosphorylation across individual proteins. The single-molecule pull-down (SiMPull) assay was developed to investigate the composition of macromolecular complexes via immunoprecipitation of proteins on a glass coverslip followed by single-molecule imaging. The current technique is an adaptation of SiMPull that provides robust quantification of the phosphorylation state of full-length membrane receptors at the single-molecule level. Imaging thousands of individual receptors in this way allows for quantifying protein phosphorylation patterns. The present protocol details the optimized SiMPull procedure, from sample preparation to imaging. Optimization of glass preparation and antibody fixation protocols further enhances data quality. The current protocol provides code for the single-molecule data analysis that calculates the fraction of receptors phosphorylated within a sample. While this work focuses on phosphorylation of the epidermal growth factor receptor (EGFR), the protocol can be generalized to other membrane receptors and cytosolic signaling molecules.

The present protocol describes sample preparation and data analysis to quantify protein phosphorylation using an improved single-molecule pull-down (SiMPull) assay.

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to ...