The authors gratefully acknowledge the technical support of Ms. Alison Ser and Ms. Hannah Oduro-Obeng, as well as the video editing support by Ms. Janice Fajardo and Mr. Miguel del Rosario.

1. Type of samples
<ol>
	<li>Use five whole grain samples, comprising two cereals (e.g., durum wheat and yellow corn) and three legumes (e.g., Blackeye cowpea bean, soybean, and red kidney bean) for this study.</li>
	<li>Mill 50 g of each grain in triplicates into flour, using a coffee grinder, and pass them through a 500 &#181;m sieve.</li>
	<li>Store them at -20 &#176;C.</li>
</ol>
2. Sample preparation
<ol>
	<li>Determination of dry matter content and expression of dry weight basis 
	NOTE: Determine the dry matter content of each powdered sample according to the method of AOAC (2000)12.
	<ol>
		<li>Turn on a forced-convection oven and set the temperature to 130 &#176;C.</li>
		<li>Dry desiccant (silica gel) in the oven for 30 min to 1 h and transfer the dried silica gel to a desiccator.</li>
		<li>Accurately weigh 2 g of each sample into a clean, pre-dried, and weighed aluminum can.</li>
		<li>Dry the weighed samples at 130 &#176;C for 1 h in a forced convection oven.</li>
		<li>Transfer the dried sample to the desiccator and allow to cool to ambient temperature.</li>
		<li>Weigh the dried, cooled sample and record its weight.</li>
		<li>Calculate the dry matter content of each sample as follows: 
		<img alt="Equation 1" src="/files/ftp_upload/62467/62467eq01v2.jpg" style="margin: auto;" /></li>
		<li>Express each parameter measured in dry weight basis using the following formula: 
		<img alt="Equation 2" src="/files/ftp_upload/62467/62467eq02v2.jpg" style="margin: auto;" /></li>
	</ol>
	</li>
	<li>Phenolic acid extraction 
	NOTE: Extract the soluble free phenolic compounds in the grain samples using a modification of the method of Y. Qiu et al.5 that enables phenolic extraction from milligram quantities of whole grains.
	<ol>
		<li>Accurately weigh 100 mg of the whole grain flour sample directly into an amber-colored 2 mL capacity microcentrifuge tube. The dark color of the tube helps to prevent exposure of the mixture to light.</li>
		<li>Add 1 mL of 80% aqueous HPLC-grade methanol to each of the tubes containing the samples.</li>
		<li>Vortex briefly to mix the methanol solution and the samples.</li>
		<li>Sonicate the samples for 60 min to extract the free soluble phenolic compounds. Put a cover over the samples for the duration of the sonication for added protection from light.</li>
		<li>After sonication, centrifuge the mixture at 20,000 &#215; g for 5 min to sediment the solid residues leaving the supernatant on top. Free phenolic compounds will be present in the supernatant after centrifugation.</li>
		<li>Transfer the supernatant into a clean microcentrifuge tube. 
		NOTE: The supernatant needs to be filtered prior to injecting into the HPLC instrument. To filter the supernatant, remove the plunger of a 3 mL syringe and attach a syringe filter. The filter should have a pore size of no larger than 0.22 &#181;m.</li>
		<li>Pipette approximately 0.4 mL of the supernatant into the top of the syringe. Re-insert the plunger and push the liquid through the filter into an HPLC vial containing a vial insert.</li>
		<li>Once the instrument has been set up to run the method outlined in the manuscript for HPLC analysis, load the vials into the carousel to correspond with the sample list.</li>
		<li>Obtain HPLC chromatograms at 320 nm and 280 nm showing distinct peaks representing different phenolic compounds.</li>
		<li>Using appropriate standard curves, quantify hydroxycinnamic acids at 320 nm since they have maximum absorbance at this wavelength. By the same principle, quantify hydroxybenzoic acids at 280 nm.</li>
		<li>Store the remaining extracts at -20 &#176;C for other analyses.</li>
	</ol>
	</li>
</ol>
3. Phenolic composition
<ol>
	<li>Use a high-pressure liquid chromatograph (Table of Materials) to identify and quantify the extracted phenolic compounds in the samples, based on the method of J. Xiang, F. B. Apea-Bah, V. U. Ndolo, M. C. Katundu, and T. Beta 4.</li>
	<li>Prepare phenolic acid standards (vanillic acid, caffeic acid, p-coumaric acid, ferulic acid, and sinapic acid) to identify and quantify constituent phenolic compounds in the extracts.</li>
	<li>To do this, weigh 1 mg of each standard and dissolve in 1 mL of 50% aqueous methanol to produce 1,000 &#181;g/mL of each standard.</li>
	<li>Mix equal volumes of all five standards in a 2 mL amber centrifuge tube to produce a cocktail of standards, each having a concentration of 200 &#181;g/mL.</li>
	<li>Prepare serial dilutions of the standard cocktail by taking a volume into a new tube and diluting with equal volume of the aqueous methanol solvent.</li>
	<li>Repeat the serial dilutions down to a concentration of 3.125 &#181;g/mL.</li>
	<li>Also, separately dilute each standard 40 times with the solvent to obtain 25 &#181;g/mL concentration of each standard.</li>
	<li>Set the column temperature to 35 &#176;C and the sample oven temperature to 15 &#176;C.</li>
	<li>To prepare mobile phase A (0.1% aqueous formic acid), transfer 1 mL of formic acid into a 1 L capacity volumetric flask and add HPLC-grade water to the 1 L mark. Shake well to mix.</li>
	<li>To prepare mobile phase B (0.1% formic acid in methanol), transfer 1 mL of formic acid into a 1 L capacity volumetric flask and add HPLC-grade methanol to the 1 L mark. Shake well to mix.</li>
	<li>Adjust the volumes to the 1 L mark if necessary.</li>
	<li>Filter both mobile phases through a 0.45 &#181;m hydrophilic filter paper.</li>
	<li>For the analysis, inject 10 &#181;L of each extract or standard (25 &#181;g/mL standards and the standard cocktails) on to the reverse phase column.</li>
	<li>Elute with the mobile phases according to the linear gradient program for a 25 min run as follows: 0-3.81 min, 9%-14% B; 3.81-4.85 min, 14%-15% B; 4.85-5.89 min, 15%-15% B, 5.89-8.32 min, 15%-17% B; 8.32-9.71 min, 17%-19% B; 9.71-10.40 min, 19%-19% B; 10.40-12.48 min, 19%-26% B; 12.48-13.17 min, 16%-28% B; 13.17-14.21 min, 28%-35% B; 14.21-15.95 min, 35%-40% B; 15.95-16.64 min, 40%-48% B; 16.64-18.37 min, 48%-53% B; 18.37-22.53 min, 53%-70% B; 22.53-22.88 min, 70%-90% B; 22.88-25.00 min, 90% B.</li>
	<li>Compare the retention times of chromatographic peaks obtained for the authentic standards of concentrations 25 &#181;g/mL, at 280 nm and 320 nm, with those of the extracts, in order to identify the constituent phenolic compounds in the samples.</li>
	<li>Plot calibration curves for the phenolic acid standard cocktails, with concentration of the standards on the horizontal axis, and the peak area on the vertical axis</li>
	<li>Use the calibration curves to estimate the concentrations of the identified phenolic compounds, by comparing the peak areas on the plot with that of the identified compounds at 280 nm and 320 nm as mentioned in the earlier section.</li>
</ol>
4. Total phenolic content
NOTE: Determine the total phenolic content of the extracts using the Folin-Ciocalteu method described by F. B. Apea-Bah et al.13.
<ol>
	<li>Prepare ferulic acid and gallic acid standards within the concentration range of 0.025 to 0.150 mg/mL to plot calibration curves in comparison, for the estimation of total phenolic content.</li>
	<li>To do this, accurately weigh 1 mg each of ferulic acid and gallic acid standards into 2 mL capacity centrifuge tubes.</li>
	<li>Add 1 mL of 50% aqueous methanol to each standard and vortex to dissolve, producing 1 mg/mL stock of each standard.</li>
	<li>Prepare a series of dilutions from each stock solution in a total of 500 &#181;L.</li>
	<li>Pipette 18.2 &#181;L of each extract or standard into a separate well on a 96-well microplate.</li>
	<li>Add 36.4 &#181;L of 10% (v/v) aqueous Folin-Ciocalteu reagent to each extract or standard.</li>
	<li>Then, add 145.4 &#181;L of 700 mM sodium carbonate to each reaction mixture.</li>
	<li>Incubate the reaction mixtures in the dark at room temperature (20-25 &#176;C) for 2 h.</li>
	<li>Read the absorbance on a microplate reader at 750 nm.</li>
	<li>Plot calibration curves of change in absorbance against concentration of the phenolic acid standards and use them to estimate the total phenolic contents.</li>
	<li>Express the results as milligram ferulic acid equivalents per gram of milled sample (mg FAE/g) and milligram gallic acid equivalents per gram of milled sample (mg GAE/g) on dry weight basis.</li>
</ol>
5. Antioxidant assays
NOTE: Determine the antioxidant capacity of the grain extracts using the following three assays: 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging capacity; 2,2&#39;-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid (ABTS) radical scavenging capacity, which is also called the Trolox equivalent antioxidant capacity (TEAC); and oxygen radical absorbance capacity (ORAC).
<ol>
	<li>Preparation of Trolox standards for standard curves
	<ol>
		<li>Use Trolox, a water-soluble analogue of vitamin E, as a standard to estimate the in vitro antioxidant capacity of the whole grain extracts.</li>
		<li>Accurately weigh 1 mg of Trolox into a 15 mL tube. Dissolve in 4 mL of 50% aqueous methanol. Vortex to dissolve to prepare a stock solution of 1 mM (1000 &#181;M).</li>
		<li>Prepare six concentrations of Trolox, that is, 50, 100, 200, 400, 600, and 800 &#181;M to plot standard curves for the estimation of DPPH radical scavenging capacity and Trolox equivalent antioxidant capacity (TEAC). Similarly, prepare 6.25, 12.5, 25, and 50 &#181;M concentrations of Trolox for estimating oxygen radical absorbance capacity (ORAC). Make up the total volume of each concentration to 500 &#181;L as shown in Table 1.</li>
	</ol>
	</li>
	<li>Dilution of sample extracts
	<ol>
		<li>Dilute the sample extracts with methanol prior to analysis. Here, yellow corn and cowpea extracts were diluted two times, the wheat and kidney bean were diluted five times while the soybean extract was diluted 10 times with methanol.</li>
	</ol>
	</li>
	<li>Trolox equivalent antioxidant capacity (TEAC) assay
	<ol>
		<li>Measure the Trolox equivalent antioxidant capacity (TEAC) of the samples using the method described previously by F. B. Apea-Bah et al.14.</li>
		<li>Weigh 8.23 mg of ABTS into a clean 2 mL capacity amber centrifuge tube.</li>
		<li>Next, weigh 1.62 mg of potassium persulfate into another clean 2 mL capacity amber centrifuge tube.</li>
		<li>Add 1 mL distilled water to each of them and vortex to dissolve them. 
		NOTE: This results in 16 mM ABTS stock solution with 6 mM aqueous potassium persulfate solution.</li>
		<li>Prepare the ABTS stock solution by mixing the ABTS and potassium persulfate solutions in equal volumes. The solution will immediately change to a dark color.</li>
		<li>Incubate the reagent mixture in the dark for 12-16 h.</li>
		<li>Dilute the ABTS stock solution 30 times with 200 mM phosphate buffered saline (PBS), to form the ABTS working solution. To do this, add 58 mL of 200 mM PBS to 2 mL of the ABTS stock solution. The working solution will contain 0.27 mM ABTS and 0.1 mM potassium persulfate.</li>
		<li>For the analysis, place 10 &#181;L of each diluted extract or Trolox into a 96-well microplate.</li>
		<li>Add 190 &#181;L of ABTS working solution to each well and incubate the reaction mixtures for 60 min.</li>
		<li>Measure the absorbance of the reaction mixtures at 750 nm in a microplate reader.</li>
		<li>Use the Trolox standards in a concentration ranging from 100 to 800 &#181;mol/L to plot a calibration curve.</li>
		<li>Estimate the ABTS radical scavenging capacity from the calibration curve.</li>
		<li>Express the results as micromole Trolox equivalents per gram (&#181;mol TE/g) sample on dry weight basis.</li>
	</ol>
	</li>
	<li>DPPH assay 
	NOTE: Determine the DPPH radical scavenging capacity of the samples using the method described previously by F. B. Apea-Bah et al.13. The DPPH antioxidant assay requires a radical-generating compound, DPPH (2,2-diphenyl-1-picrylhydrazyl).
	<ol>
		<li>Accurately weigh 1.2 mg of DPPH into an empty 50 mL capacity centrifuge tube. Dissolve the DPPH in 30 mL of methanol to prepare a 60 &#181;mol/L methanolic solution. 
		NOTE: The DPPH assay tests the ability of the sample extracts to scavenge free radicals produced by DPPH.</li>
		<li>For the analysis, add 5 &#181;L of the sample extracts or Trolox solution into the microplate wells.</li>
		<li>Next, add 195 &#181;L of the 60 &#181;mol/L DPPH methanolic solution and incubate for 60 min.</li>
		<li>Measure the absorbance of the reaction mixture at 515 nm.</li>
		<li>Use the Trolox standards (50-800 &#181;mol/L) to plot a calibration curve, with the change in absorbance on the vertical axis and the Trolox concentrations on the horizontal axis.</li>
		<li>Estimate the DPPH scavenging capacity from the calibration curve.</li>
		<li>Express the results as micromole Trolox equivalents per gram (&#181;mol TE/g) sample on dry weight basis.</li>
	</ol>
	</li>
	<li>Oxygen radical absorbance capacity
	<ol>
		<li>Determine the oxygen radical absorbance capacity (ORAC) of the samples based on the method of F. B. Apea-Bah et al.13.</li>
		<li>To begin, prepare Trolox standards of concentrations 6.25, 12.5, 25, and 50 &#181;M from a 1,000 &#181;M Trolox standard stock solution in 75 mM potassium phosphate (K2HPO4/KH2PO4) buffer.</li>
		<li>To do this, weigh 1 mg Trolox powder and dissolve in 4 mL of the buffer solution under sonication to prepare a 1,000 &#181;M stock solution.</li>
		<li>Then, take 50 &#181;L of the stock solution into a 2 mL capacity centrifuge tube and dilute with 950 &#181;L of buffer solution to obtain 1,000 &#181;L of 50 &#181;M Trolox solution.</li>
		<li>Pipette 500 &#181;L of the 50 &#181;M solution into a new tube and dilute with equal volume of buffer to obtain a 25 &#181;M Trolox solution.</li>
		<li>Repeat the serial dilution of the 25 &#181;M to obtain 12.5 &#181;M, and then similarly repeat the dilution of 12.5 &#181;M to obtain 6.25 &#181;M.</li>
		<li>Prepare appropriate dilutions of the sample extracts.</li>
		<li>Dilute the yellow corn and cowpea extracts 20 times, the wheat and kidney bean extracts 50 times, and the soybean extract 100 times with the buffer solution.</li>
		<li>Transfer 200 &#181;L of each sample or Trolox standard into the wells of a black 96-well microplate for automatic pipetting.</li>
		<li>Then, fill the three reagent tanks provided with the ORAC equipment, with the following: (1) the buffer solution; (2) 0.816 nM fluorescein in the buffer; and (3) 153 mM of 2,2&#8242;-azobis(2-amidinopropane) dihydrochloride (AAPH) in buffer.</li>
		<li>Place them in their receptacles for automatic pipetting.</li>
		<li>Set up the ORAC machine and automatic pipetting system for analysis, based on the standard operating procedure.</li>
		<li>For the analysis, set up the automated pipetting system to transfer 25 &#181;L of each diluted extract or standard to the wells of a clear bottom black 96-well microplate.</li>
		<li>Then, automatically add 150 &#181;L of 0.816 nM fluorescein in buffer.</li>
		<li>Incubate the reaction mixture at 37 &#176;C for 15 min in the ORAC machine.</li>
		<li>Afterwards, automatically add 25 &#181;L of the AAPH to each reaction mixture.</li>
		<li>Incubate them at 37 &#176;C for 50 min in the ORAC machine.</li>
		<li>Set up the ORAC machine to measure the fluorescence decay, over the incubation period, at excitation and emission wavelengths of 485 nm and 520 nm, respectively.</li>
		<li>After the measurements, plot a Trolox standard curve, with Trolox (6.25-50 &#181;M) on the horizontal axis and fluorescence decay on the vertical axis.</li>
		<li>Estimate the oxygen radical absorbance capacity of the extracts from the Trolox standard curve.</li>
		<li>Express the results as &#181;mol TE/g sample on dry weight basis.</li>
	</ol>
	</li>
	<li>Statistical analysis
	<ol>
		<li>Present all the results as means &#177; standard deviation of at least triplicates.</li>
		<li>Perform analysis of variance (ANOVA) to determine the effect of grain type on the response variables.</li>
		<li>Where significant differences exist at p &#60; 0.05, use least significant difference (LSD) to compare the means.</li>
		<li>Perform Pearson correlation analysis to estimate the relationship between the phenolic content and antioxidant capacities.</li>
	</ol>
	</li>
</ol>

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B., Serem, J. C., Bester, M. J., Duodu, K. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenolic+composition+and+antioxidant+properties+of+Koose,+a+deep-fat+fried+cowpea+cake.">Phenolic composition and antioxidant properties of Koose, a deep-fat fried cowpea cake.</a> Food Chemistry. 237, 247-256 (2017).</li></ol>

The authors declare no conflicts of interest.

The whole grains were selected as representative cereal grains and legumes that find wide food applications worldwide. While variations may exist among cultivars of each grain, the focus of this study was to demonstrate a generalized method for free phenolic acid extraction and analysis for whole grains. The extraction method was modified by substantially reducing the amounts of samples and solvents, in order to reduce the amount of chemicals that would be released into the environment when such experiments are conducted...

Phenolic acids are among the most important phytochemicals studied in plants due to the vital role they play in plant defense against herbivory and fungal infection, as well as maintaining structural support and integrity in plant tissues1,2. They are abundant in the bran of cereals and seed coat of legumes3. Structurally, they are divided into two groups: the hydroxybenzoic acids (Figure 1) and hydroxycinnamic acids (Figure 2). The common hydroxybenzoic acids in cereals and legumes include gallic, p-hydroxybenzoic, 2,4-dihydroxybenzoic, protocatechuic, vanillic, and syringic acids, while the common hydroxycinnamic acids include caffeic, p-coumaric, ferulic, and sinapic acids3. Phenolic acids also possess antioxidant properties since they are able to scavenge free radicals, which cause oxidative rancidity in fats, and initiate and propagate radical-induced oxidative stress in physiological systems4,5. Due to this vital physiological role as antioxidants, they are the subject of recent research. This is because when consumed as components of plant foods, they can exert antioxidant protection.
Cereals and cereal products are major carbohydrate food sources for humans and animals worldwide6. Cereals include wheat, rice, corn (maize), barley, triticale, millets, and sorghum. Among them, corn is the most utilized, with an estimated global utilization of 1,135.7 million tonnes in 2019/2020, followed by wheat with an estimated global utilization of 757.5 million tonnes during the same period7. Cereal foods are great sources of energy to consumers since they are rich sources of carbohydrates. They also provide some protein, fat, fiber, vitamins and minerals6. In addition to their nutritional value, cereals are good sources of phytochemical antioxidants, particularly phenolic acids, which have the potential to protect the physiological system from radical-induced oxidative damage3. Legumes are also good sources of nutrients and are generally higher in protein than cereals. They also contain vitamins and minerals and are used in the preparation of various foods8. Additionally, legumes are good sources of a variety of phytochemical antioxidants, including phenolic acids, flavonoids, anthocyanins, and proanthocyanidins9,10. Different varieties of cereals and legumes may have a different phenolic acid composition. There is therefore the need to study the phenolic acid composition of cereals and legumes and their varieties, in order to know their potential health benefits with respect to phenolic antioxidants.
A number of assays have been reported for measuring the quantity of phenolic acids in cereal and legume grains, and determining their antioxidant activities. The most common methods of analysis for whole grain phenolic acids are spectrophotometry and liquid chromatography11. The aim of this work was to demonstrate a generalized high pressure liquid chromatographic method for determining free soluble phenolic acid composition, and spectrophotometric methods for determining total phenolic content and antioxidant capacity of some whole grain cereals and legumes.

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		<tr>
			<td>15 mL Falcon conical centrifuge tubes</td>
			<td>Fisher Scientific</td>
			<td>05-527-90</td>
			<td></td>
		</tr>
		<tr>
			<td>2 mL Amber glass ID Surestop vial</td>
			<td>Thermo Scientific</td>
			<td>C5000-2W</td>
			<td></td>
		</tr>
		<tr>
			<td>2 mL Amber microcentrifuge tubes</td>
			<td>VWR</td>
			<td>20170-084</td>
			<td></td>
		</tr>
		<tr>
			<td>2,2&#8242;-Azobis(2-amidinopropane) dihydrochloride (AAPH)</td>
			<td>Sigma-Aldrich</td>
			<td>440914-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>2,2&#39;-Azino-bis(3-ethylbenzothiazoline-6-sulfonic acid (ABTS) (C18H18N4O6S4) &#8805;98%,</td>
			<td>Sigma Aldrich</td>
			<td>A1888-2G</td>
			<td></td>
		</tr>
		<tr>
			<td>2,2-Diphenyl-1pikrylhydrazyl (DPPH) (C18H12N5O6)</td>
			<td>Sigma Aldrich</td>
			<td>D913-2</td>
			<td></td>
		</tr>
		<tr>
			<td>6-Hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (Trolox) (C14H18O4), &#8805;98%</td>
			<td>Fluka Chemika</td>
			<td>56510</td>
			<td></td>
		</tr>
		<tr>
			<td>9 mm Autosampler Vial Screw Thread Caps</td>
			<td>Thermo Scientific</td>
			<td>60180-670</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well flat bottom plates</td>
			<td>Fisher Scientific</td>
			<td>12565501</td>
			<td></td>
		</tr>
		<tr>
			<td>Agilent BioTek ELx800 microplate reader</td>
			<td>Fisher Scientific</td>
			<td>BT-ELX800NB</td>
			<td></td>
		</tr>
		<tr>
			<td>Agilent BioTek Precision 2000 96/384 Automated Microplate Pipetting System</td>
			<td>Fisher Scientific</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Agilent BioTek FLx800 Microplate Fluorescence Reader</td>
			<td>Fisher Scientific</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Analytical balance SI-114</td>
			<td>Denver Instrument</td>
			<td>SI-114.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Autosampler, Waters 717 Plus</td>
			<td>Waters</td>
			<td>WAT078900</td>
			<td></td>
		</tr>
		<tr>
			<td>BD 3 mL syringe Luer-Lok Tip</td>
			<td>BD</td>
			<td>309657</td>
			<td></td>
		</tr>
		<tr>
			<td>Bransonic ultrasonic cleaner, Branson 5510</td>
			<td>Millipore Sigma</td>
			<td>Z245143</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning LSE Vortex Mixer</td>
			<td>Corning</td>
			<td>6775</td>
			<td></td>
		</tr>
		<tr>
			<td>Durapore Filter (0.45 &#181;m PVDF Membrane)</td>
			<td>Merck Millipore Ltd</td>
			<td>HVLP04700&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Durapore Membrane Filters (0.45 &#181;m HV)</td>
			<td>Merck Millipore Ltd</td>
			<td>HVHP04700</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research&#160;plus, 0.5-10 &#181;L</td>
			<td>Eppendorf</td>
			<td>3123000020</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research&#160;plus, 0.5-5 mL</td>
			<td>Eppendorf</td>
			<td>3123000071</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research&#160;plus, 100-1000 &#181;L</td>
			<td>Eppendorf</td>
			<td>3123000063</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Research&#160;plus, 10-100 &#181;L</td>
			<td>Eppendorf</td>
			<td>3123000047</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl acetate, HPLC grade</td>
			<td>Fisher Chemical</td>
			<td>E195-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Ferulic acid standard</td>
			<td>Sigma Aldrich</td>
			<td>128708-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescein</td>
			<td>Fisher Scientific</td>
			<td>AC119245000</td>
			<td></td>
		</tr>
		<tr>
			<td>Folin &#38; Ciocalteu phenol reagent</td>
			<td>Sigma Aldrich</td>
			<td>F9252</td>
			<td></td>
		</tr>
		<tr>
			<td>Formic acid, 99%</td>
			<td>Acros Organics, Janssen Pharmaceuticalaan 3a</td>
			<td>27048-0010</td>
			<td></td>
		</tr>
		<tr>
			<td>Gallic acid standard</td>
			<td>Sigma</td>
			<td>G7384</td>
			<td></td>
		</tr>
		<tr>
			<td>High performance liquid chromatograph (HPLC), Waters 2695</td>
			<td>Waters</td>
			<td>960402</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol, HPLC grade</td>
			<td>Fisher Chemical</td>
			<td>A452-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro pipet tips, 0.5-10 &#181;L</td>
			<td>Fisherbrand</td>
			<td>21-197-2F</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge Sorvall Legend Micro 21 centrifuge</td>
			<td>Thermo Scientific</td>
			<td>75002435</td>
			<td></td>
		</tr>
		<tr>
			<td>Multichannel micropipette, Proline Plus, 30-300 &#181;L</td>
			<td>Sartorius</td>
			<td>728240</td>
			<td></td>
		</tr>
		<tr>
			<td>Photodiode array detector, Waters 2996</td>
			<td>Waters</td>
			<td>720000350EN</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipet tips, 1000 &#181;L</td>
			<td>VWR</td>
			<td>83007-382</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipet tips, 1-5 mL</td>
			<td>VWR</td>
			<td>82018-840</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium persulfate (K2S2O8), &#8805;99.0%</td>
			<td>Sigma Aldrich</td>
			<td>216224-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate dibasic anhydrous (K2HPO4)</td>
			<td>Fisher Scientific</td>
			<td>P288-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate monobasic (KH2PO4)</td>
			<td>Fisher Scientific</td>
			<td>P285-500</td>
			<td></td>
		</tr>
		<tr>
			<td>PYREX 250 mL Short Neck Boiling Flask, Round Bottom</td>
			<td>Corning</td>
			<td>4321-250</td>
			<td></td>
		</tr>
		<tr>
			<td>Reversed phase C18 Analytical Column (100 x 3 mm) Accucore aQ</td>
			<td>Thermo Scientific</td>
			<td>17326-103030</td>
			<td></td>
		</tr>
		<tr>
			<td>Roto evaporator, IKA RV 10</td>
			<td>IKA</td>
			<td>&#160;0010005185</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium carbonate (NaCO3) anhydrous</td>
			<td>Fisher Chemical</td>
			<td>S263-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride (NaCl)</td>
			<td>Mallinckrodt AR&#174;</td>
			<td>7581</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium phosphate dibasic anhydrous (Na2HPO4)</td>
			<td>Fisher Scientific</td>
			<td>BP332-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium phosphate monobasic anhydrous (NaH2PO4)</td>
			<td>Fisher bioreagents</td>
			<td>BP329-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Standardization pipet tips 0-200&#181;L</td>
			<td>Fisherbrand</td>
			<td>02-681-134</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe Driven Filter unit (0.22 &#181;m)</td>
			<td>&#160;Millex&#174;-GV</td>
			<td>SLGVR04NL</td>
			<td></td>
		</tr>
		<tr>
			<td>Target micro-serts vial insert (400 &#181;L)</td>
			<td>Thermo Scientific</td>
			<td>C4011-631</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultrapure water (Direct Q-3 UV system with pump)</td>
			<td>Millipore</td>
			<td>ZRQSVP030</td>
			<td></td>
		</tr>
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</table>

a generalized method for determining free soluble phenolic acid composition and antioxidant capacity of cereals and legumes

Phenolic acids are a class of organic compounds that bear both a phenolic group, and a carboxylic group. They are found in grains and concentrate in the bran of cereals or seed coat of legumes. They possess antioxidant properties that have generated much research interest in recent years, about their potential antioxidant protective health functions. This work presents a generalized method for the extraction of free soluble phenolic acids from whole grains and analysis of their antioxidant capacity. Five whole grain samples comprising two cereals (wheat and yellow corn) and three legumes (cowpea bean, kidney bean, and soybean), were used. The grains were milled into flour and their free soluble phenolic acids extracted using aqueous methanol. The compounds were then identified using a high-pressure liquid chromatograph (HPLC). The Folin-Ciocalteu method was used to determine their total phenolic content while their antioxidant capacities were determined using the DPPH radical scavenging capacity, Trolox equivalent antioxidant capacity (TEAC) and oxygen radical absorbance capacity (ORAC) assays. The phenolic acids identified included vanillic, caffeic, p-coumaric and ferulic acids. Vanillic acid was identified only in cowpea while caffeic acid was identified only in kidney bean. p-Coumaric acid was identified in yellow corn, cowpea, and soybean, while ferulic acid was identified in all the samples. Ferulic acid was the predominant phenolic acid identified. The total concentration of phenolic acids in the samples decreased in the following order: soybean &gt; cowpea bean &gt; yellow corn = kidney bean &gt; wheat. The total antioxidant capacity (sum of values of DPPH, TEAC and ORAC assays) decreased as follows: soybean &gt; kidney bean &gt; yellow corn = cowpea bean &gt; wheat. This study concluded that HPLC analysis as well as DPPH, TEAC, and ORAC assays provide useful information about the phenolic acid composition and antioxidant properties of whole grains.

Table 2 shows the phenolic acids that were identified in the cereal and legume grains. Based on available authentic standards, four phenolic acids were identified in the samples and they are: vanillic, caffeic, p-coumaric, and ferulic acids. Vanillic acid is a hydroxybenzoic acid while the other three are hydroxycinnamic acids. Vanillic acid was identified only in Blackeye cowpea bean while caffeic acid was identified only in kidney bean. p-Coumaric acid was identified in yellow corn, c...

Watch this Scientific Journal Video about A Generalized Method for Determining Free Soluble Phenolic Acid Composition and Antioxidant Capacity of Cereals and Legumes at JoVE.com

A Generalized Method for Determining Free Soluble Phenolic Acid Composition and Antioxidant Capacity of Cereals and Legumes

Phenolic acids are important phytochemicals that are present in whole grains. They possess bioactive properties such as antioxidant protective functions. This work aimed at reporting on a generalized method for the HPLC identification, total phenolic content estimation, and determination of the antioxidant capacity of phenolic acids in cereals and legumes.

Phenolic acids are a class of organic compounds that bear both a phenolic group, and a carboxylic group. They are found in grains and concentrate in the ...

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6.6K Views.  University of Manitoba. Phenolic acids are important phytochemicals that are present in whole grains. They possess bioactive properties such as antioxidant protective functions. This work aimed at reporting on a generalized method for the HPLC identification, total phenolic content estimation, and determination of the antioxidant capacity of phenolic acids in cereals and legumes.

Video: A Generalized Method for Determining Free Soluble Phenolic Acid Composition and Antioxidant Capacity of Cereals and Legumes

Free Radicals in Chemical Biology: from Chemical Behavior to Biomarker Development

The involvement of free radicals in life sciences has constantly increased with time and has been connected to several physiological and pathological processes. This subject embraces diverse scientific areas, spanning from physical, biological and bioorganic chemistry to biology and medicine, with applications to the amelioration of quality of life, health and aging. Multidisciplinary skills are required for the full investigation of the many facets of radical processes in the biological environment and chemical knowledge plays a crucial role in unveiling basic processes and mechanisms. We developed a chemical biology approach able to connect free radical chemical reactivity with biological processes, providing information on the mechanistic pathways and products. The core of this approach is the design of biomimetic models to study biomolecule behavior (lipids, nucleic acids and proteins) in aqueous systems, obtaining insights of the reaction pathways as well as building up molecular libraries of the free radical reaction products. This context can be successfully used for biomarker discovery and examples are provided with two classes of compounds: mono-trans isomers of cholesteryl esters, which are synthesized and used as references for detection in human plasma, and purine 5',8-cyclo-2'-deoxyribonucleosides, prepared and used as reference in the protocol for detection of such lesions in DNA samples, after ionizing radiations or obtained from different health conditions.

Radical-based biomimetic chemistry has been applied to building-up libraries necessary for biomarker development.

The involvement of free radicals in life sciences has constantly increased with time and has been connected to several physiological and pathological ...

Preparation of Silica Nanoparticles Through Microwave-assisted Acid-catalysis

Microwave-assisted synthetic techniques were used to quickly and reproducibly produce silica nanoparticle sols using an acid catalyst with nanoparticle diameters ranging from 30-250 nm by varying the reaction conditions. Through the selection of a microwave compatible solvent, silicic acid precursor, catalyst, and microwave irradiation time, these microwave-assisted methods were capable of overcoming the previously reported shortcomings associated with synthesis of silica nanoparticles using microwave reactors. The siloxane precursor was hydrolyzed using the acid catalyst, HCl. Acetone, a low-tan &#948; solvent, mediates the condensation reactions and has minimal interaction with the electromagnetic field. Condensation reactions begin when the silicic acid precursor couples with the microwave radiation, leading to silica nanoparticle sol formation. The silica nanoparticles were characterized by dynamic light scattering data and scanning electron microscopy, which show the materials&#39; morphology and size to be dependent on the reaction conditions. Microwave-assisted reactions produce silica nanoparticles with roughened textured surfaces that are atypical for silica sols produced by St&#246;ber&#39;s methods, which have smooth surfaces.

Silica nanoparticles were prepared using acid-catalysis of a siloxane precursor and microwave-assisted synthetic techniques resulting in the controlled growth of nanomaterials ranging from 30-250 nm in diameter. The growth dynamics can be controlled by varying the initial silicic acid concentration, time of the reaction, and temperature of reaction.

Microwave-assisted synthetic techniques were used to quickly and reproducibly produce silica nanoparticle sols using an acid catalyst with nanoparticle ...

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Woody materials are comprised of plant cell walls that contain a layered secondary cell wall composed of structural polymers of polysaccharides and lignin. Layer-by-layer (LbL) assembly process which relies on the assembly of oppositely charged molecules from aqueous solutions was used to build a freestanding composite film of isolated wood polymers of lignin and oxidized nanofibril cellulose (NFC). To facilitate the assembly of these negatively charged polymers, a positively charged polyelectrolyte, poly(diallyldimethylammomium chloride) (PDDA), was used as a linking layer to create this simplified model cell wall. The layered adsorption process was studied quantitatively using quartz crystal microbalance with dissipation monitoring (QCM-D) and ellipsometry. The results showed that layer mass/thickness per adsorbed layer increased as a function of total number of layers. The surface coverage of the adsorbed layers was studied with atomic force microscopy (AFM). Complete coverage of the surface with lignin in all the deposition cycles was found for the system, however, surface coverage by NFC increased with the number of layers. The adsorption process was carried out for 250 cycles (500 bilayers) on a cellulose acetate (CA) substrate. Transparent free-standing LBL assembled nanocomposite films were obtained when the CA substrate was later dissolved in acetone. Scanning electron microscopy (SEM) of the fractured cross-sections showed a lamellar structure, and the thickness per adsorption cycle (PDDA-Lignin-PDDA-NC) was estimated to be 17 nm for two different lignin types used in the study. The data indicates a film with highly controlled architecture where nanocellulose and lignin are spatially deposited on the nanoscale (a polymer-polymer nanocomposites), similar to what is observed in the native cell wall.

The objective of this research was to form synthetic plant cell wall tissue using layer-by-layer assembly of nanocellulose fibrils and isolated lignin assembled from dilute aqueous suspensions.&#160; Surface measurement techniques of quartz crystal microbalance and atomic force microscopy were used to monitor the formation of the polymer-polymer nanocomposite material.

Woody materials are comprised of plant cell walls that contain a layered secondary cell wall composed of structural polymers of polysaccharides and ...

Synthesis of Cd-free InP/ZnS Quantum Dots Suitable for Biomedical Applications

Fluorescent nanocrystals, specifically quantum dots, have been a useful tool for many biomedical applications. For successful use in biological systems, quantum dots should be highly fluorescent and small/monodisperse in size. While commonly used cadmium-based quantum dots possess these qualities, they are potentially toxic due to the possible release of Cd2+ ions through nanoparticle degradation. Indium-based quantum dots, specifically InP/ZnS, have recently been explored as a viable alternative to cadmium-based quantum dots due to their relatively similar fluorescence characteristics and size. The synthesis presented here uses standard hot-injection techniques for effective nanoparticle growth; however, nanoparticle properties such as size, emission wavelength, and emission intensity can drastically change due to small changes in the reaction conditions. Therefore, reaction conditions such temperature, reaction duration, and precursor concentration should be maintained precisely to yield reproducible products. Because quantum dots are not inherently soluble in aqueous solutions, they must also undergo surface modification to impart solubility in water. In this protocol, an amphiphilic polymer is used to interact with both hydrophobic ligands on the quantum dot surface and bulk solvent water molecules. Here, a detailed protocol is provided for the synthesis of highly fluorescent InP/ZnS quantum dots that are suitable for use in biomedical applications.

In this protocol, the synthesis of Cd-free InP/ZnS quantum dots (QDs) is detailed. InP-based QDs are gaining popularity due to the toxicity of Cd2+ ions that may be released through nanoparticle degradation. After synthesis, QDs are solubilized in water using an amphiphilic polymer for use in biomedical applications.

Fluorescent nanocrystals, specifically quantum dots, have been a useful tool for many biomedical applications. For successful use in biological systems, ...

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We demonstrate a method for the synthesis of a water-soluble multimetallic peptidic array containing a predetermined sequence of metal centers such as Ru(II), Pt(II), and Rh(III). The compound, named as a water-soluble metal-organic complex array (WSMOCA), is obtained through 1) the conventional solution-chemistry-based preparation of the corresponding metal complex monomers having a 9-fluorenylmethyloxycarbonyl (Fmoc)-protected amino acid moiety and 2) their sequential coupling together with other water-soluble organic building units on the surface-functionalized polymeric resin by following the procedures originally developed for the solid-phase synthesis of polypeptides, with proper modifications. Traces of reactions determined by mass spectrometric analysis at the representative coupling steps in stage 2 confirm the selective construction of a predetermined sequence of metal centers along with the peptide backbone. The WSMOCA cleaved from the resin at the end of stage 2 has a certain level of solubility in aqueous media dependent on the pH value and/or salt content, which is useful for the purification of the compound.

A potential general method for the synthesis of water-soluble multimetallic peptidic arrays containing a predetermined sequence of metal centers is presented.

We demonstrate a method for the synthesis of a water-soluble multimetallic peptidic array containing a predetermined sequence of metal centers such as ...

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A protocol to avoid the oxidation of metallic substrates during sample transfer from an inhibited acidic solution to an X-ray photoelectron spectrometer is presented.

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Over the last 50 years, Cell-Free Protein Synthesis (CFPS) has emerged as a powerful technology to harness the transcriptional and translational capacity ...

Synthesis Method for Cellulose Nanofiber Biotemplated Palladium Composite Aerogels

Here, a method to synthesize cellulose nanofiber biotemplated palladium composite aerogels is presented. Noble metal aerogel synthesis methods often result in fragile aerogels with poor shape control. The use of carboxymethylated cellulose nanofibers (CNFs) to form a covalently bonded hydrogel allows for the reduction of metal ions such as palladium on the CNFs with control over both nanostructure and macroscopic aerogel monolith shape after supercritical drying. Crosslinking the carboxymethylated cellulose nanofibers is achieved using 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) in the presence of ethylenediamine. The CNF hydrogels maintain their shape throughout synthesis steps including covalent crosslinking, equilibration with precursor ions, metal reduction with high concentration reducing agent, rinsing in water, ethanol solvent exchange, and CO2 supercritical drying. Varying the precursor palladium ion concentration allows for control over the metal content in the final aerogel composite through a direct ion chemical reduction rather than relying on the relatively slow coalescence of pre-formed nanoparticles used in other sol-gel techniques. With diffusion as the basis to introduce and remove chemical species into and out of the hydrogel, this method is suitable for smaller bulk geometries and thin films. Characterization of the cellulose nanofiber-palladium composite aerogels with scanning electron microscopy, X-ray diffractometry, thermal gravimetric analysis, nitrogen gas adsorption, electrochemical impedance spectroscopy, and cyclic voltammetry indicates a high surface area, metallized palladium porous structure.

A synthesis method for cellulose nanofiber biotemplated palladium composite aerogels is presented.The resulting composite aerogel materials offer potential for catalysis, sensing, and hydrogen gas storage applications.

Here, a method to synthesize cellulose nanofiber biotemplated palladium composite aerogels is presented. Noble metal aerogel synthesis methods often ...

NMR-Based Activity Assays for Determining Compound Inhibition, IC50 Values, Artifactual Activity, and Whole-Cell Activity of Nucleoside Ribohydrolases

NMR spectroscopy is often used for the identification and characterization of enzyme inhibitors in drug discovery, particularly in the context of fragment screening. NMR-based activity assays are ideally suited to work at the higher concentrations of test compounds required to detect these weaker inhibitors. The dynamic range and chemical shift dispersion in an NMR experiment can easily resolve resonances from substrate, product, and test compounds. This contrasts with spectrophotometric assays, in which read-out interference problems often arise from compounds with overlapping UV-vis absorption profiles. In addition, since they lack reporter enzymes, the single-enzyme NMR assays are not prone to coupled-assay false positives. This attribute makes them useful as orthogonal assays, complementing traditional high throughput screening assays and benchtop triage assays. Detailed protocols are provided for initial compound assays at 500 &#956;M and 250 &#956;M, dose-response assays for determining IC50 values, detergent counter screen assays, jump-dilution counter screen assays, and assays in E. coli whole cells. The methods are demonstrated using two nucleoside ribohydrolase enzymes. The use of 1H NMR is shown for the purine-specific enzyme, while 19F NMR is shown for the pyrimidine-specific enzyme. The protocols are generally applicable to any enzyme where substrate and product resonances can be observed and distinguished by NMR spectroscopy. To be the most useful in the context of drug discovery, the final concentration of substrate should be no more than 2-3x its Km value. The choice of NMR experiment depends on the enzyme reaction and substrates available as well as available NMR instrumentation.

NMR-Based Activity Assays for Determining Compound Inhibition, IC50 Values, Artifactual Activity, and Whole-Cell Activity of Nucleoside Ribohydrolases

NMR-based activity assays have been developed to identify and characterize inhibitors of two nucleoside ribohydrolase enzymes. Protocols are provided for initial compound assays at 500 &#956;M and 250 &#956;M, dose-response assays for determining IC50 values, detergent counter screen assays, jump-dilution counter screen assays, and assays in E. coli whole cells.

NMR spectroscopy is often used for the identification and characterization of enzyme inhibitors in drug discovery, particularly in the context of fragment ...

Ion Mobility-Mass Spectrometry Techniques for Determining the Structure and Mechanisms of Metal Ion Recognition and Redox Activity of Metal Binding Oligopeptides

Electrospray ionization (ESI) can transfer an aqueous-phase peptide or peptide complex to the gas-phase while conserving its mass, overall charge, metal-binding interactions, and conformational shape. Coupling ESI with ion mobility-mass spectrometry (IM-MS) provides an instrumental technique that allows for simultaneous measurement of a peptide&#8217;s mass-to-charge (m/z) and collision cross section (CCS) that relate to its stoichiometry, protonation state, and conformational shape. The overall charge of a peptide complex is controlled by the protonation of 1) the peptide&#8217;s acidic and basic sites and 2) the oxidation state of the metal ion(s). Therefore, the overall charge state of a complex is a function of the pH of the solution that affects the peptides metal ion binding affinity. For ESI-IM-MS analyses, peptide and metal ions solutions are prepared from aqueous-only solutions, with the pH adjusted with dilute aqueous acetic acid or ammonium hydroxide. This allows for pH dependence and metal ion selectivity to be determined for a specific peptide. Furthermore, the m/z and CCS of a peptide complex can be used with B3LYP/LanL2DZ molecular modeling to discern binding sites of the metal ion coordination and tertiary structure of the complex. The results show how ESI-IM-MS can characterize the selective chelating performance of a set of alternative methanobactin peptides and compare them to the copper-binding peptide methanobactin.

Ion mobility-mass spectrometry and molecular modeling techniques can characterize the selective metal chelating performance of designed metal-binding peptides and the copper-binding peptide methanobactin. Developing new classes of metal chelating peptides will help lead to therapeutics for diseases associated with metal ion misbalance.