Application and Principle
This procedure is used to determine the a-amylase activity, expressed as bacterial amylase units (BAU), of enzyme preparations derived from Bacillus subtilis var., Bacillus licheniformis var., and Bacillus stearothermophilus. It is not applicable to products that contain P-amylase. The assay is based on the time required to obtain a standard degree of hydrolysis of a starch solution at 30 ± 0.1°. The degree of hydrolysis is determined by comparing the iodine colour of the hydrolysate with that of a standard.
Use the Reference Colour Standard, the Comparator, and the Comparator Tubes as described under a-Amylase Activity, Fungal, but use either daylight or daylight-type fluorescent lamps as the light source for the Comparator. (Incandescent lamps give slightly lower results.)
Reagents and Solutions
pH 6.6 Buffer: Dissolve 9.1 g of potassium dihydrogen phosphate (KH2PO4) in sufficient water to make 1000 ml (Solution A). Dissolve 9.5 g of dibasic sodium phosphate (Na2HPO4) in sufficient water to make 1000 ml (Solution B). Add 400 ml of Solution A to 600 ml of Solution B, mix, and adjust the pH to 6.6, if necessary, by the addition of Solution A or Solution B as required.
Dilute Iodine Solution: Prepare as directed under a-Amylase Activity, Fungal.
Special Starch: Use the material described under a-Amylase Activity, Fungal.
Starch Substrate Solution: Disperse 10.0 g (dry-weight basis) of Special Starch in 100 ml of cold water, and slowly pour the mixture into 300 ml of boiling water. Boil and stir for 1 to 2 min, and then cool while continuously stirring. Quantitatively transfer the mixture into a 500 ml volumetric flask with the aid of water, add 10 ml of pH 6.6 Buffer, dilute to volume with water, and mix.
Sample Preparation: Prepare a solution of the sample so that 10 ml of the final dilution will give an endpoint between 15 and 35 min under the conditions of the assay.
Pipet 5.0 ml of Dilute Iodine Solution into a series of 13 x 100-mm test tubes, and place them in a water bath maintained at 30 ± 0.1°, allowing 20 tubes for each assay. Pipet 20.0 ml of the Starch Substrate Solution into a 50 ml Erlenmeyer flask, stopper, and equilibrate for 20 min in the water bath at 30°. At zero time, rapidly pipet 10.0 ml of the Sample Preparation into the equilibrated mixture, and continue as directed in the Procedure under a-Amylase Activity, Fungal, beginning with ‘‘... mix immediately by swirling, stopper the flask....’’
One bacterial amylase unit (BAU) is defined as that quantity of enzyme that will dextrinize starch at the rate of 1 mg/min under the specified test conditions.
Calculate the a-amylase activity of the sample, expressed as BAU, by the formula
in which 40 is a factor (400/10) derived from the 400 mg of starch (20 ml of a 2% solution) and the 10 ml aliquot of Sample Preparation used; F is the dilution factor (total dilution volume/sample weight, in grams); and T is the dextrinizing time, in min.
Application and Principle
This procedure is used to determine the a-amylase activity of enzyme preparations derived from Aspergillus niger var.; Aspergillus oryzae var.; Rhizopus oryzae var.; (and barley malt). The assay is based on the time required to obtain a standard degree of hydrolysis of a starch solution at 30 ± 0.1°. The degree of hydrolysis is determined by comparing the iodine colour of the hydrolysate with that of a standard.
Reference Colour Standard: Use a special a-Amylase Color Disk (Orbeco Analytical Systems, 185 Marine Street, Farmingdale, NY 11735, Catalog No. 620-S5 or similar). Alternatively, prepare a colour standard by dissolving 25.0 g of cobaltous chloride (COCI26H2O) and 3.84 g of potassium dichromate in 100 ml of 0.01 N hydrochloric acid. This standard is stable indefinitely when stored in a stoppered bottle or comparator tube.
Comparator: Use either the standard Hellige comparator (Orbeco, Catalog No. 607) or the pocket comparator with prism attachment (Orbeco, Catalog No. 605AHT) or similar. The comparator should be illuminated with a 100-W frosted lamp placed 6 in. from the rear opal glass of the comparator and mounted so that direct rays from the lamp do not shine into the operator's eyes.
Comparator Tubes: Use the precision-bored square tubes with a 13-mm viewing depth that are supplied with the Hellige comparator. Suitable tubes are also available from other apparatus suppliers.
Reagents and Solutions
Buffer Solution (pH 4.8): Dissolve 164 g of anhydrous sodium acetate in about 500 ml of water, add 120 ml of glacial acetic acid, and adjust the pH to 4.8 with glacial acetic acid. Dilute to 1000 ml with water, and mix.
Я-Amylase Solution: Dissolve into 5 ml of water a quantity of Я-amylase, free from α-amylase activity (Sigma Chemical Co., Catalog No. A7005 or equivalent), equivalent to 250 mg of Я-amylase with 2000° diastatic power.
Special Starch: Use starch designated as ‘‘Starch (Lintner) Soluble’’ (Baker Analyzed Reagent, Catalog No. 4010 or equivalent). Before using new batches, test them in parallel with previous lots known to be satisfactory. Variations of more than ±3° diastatic power in the averages of a series of parallel tests indicate an unsuitable batch.
Buffered Substrate Solution: Disperse 10.0 g (dry-weight basis) of Special Starch in 100 ml of cold water, and slowly pour the mixture into 300 ml of boiling water. Boil and stir for 1 to 2 min, then cool, and add 25 ml of Buffer Solution, followed by all of the Я-Amylase Solution. Quantitatively transfer the mixture into a 500 ml volumetric flask with the aid of water saturated with toluene, dilute to volume with the same solvent, and mix. Store the solution at 30° ± 2° for not less than 18 h but not more than 72 h before use. (This solution is also known as ‘‘buffered limit dextrin substrate.’’)
Stock Iodine Solution: Dissolve 5.5 g of iodine and 11.0 g of potassium iodide in about 200 ml of water, dilute to 250 ml with water, and mix. Store in a dark bottle, and make a fresh solution every 30 days.
Dilute Iodine Solution: Dissolve 20 g of potassium iodide in 300 ml of water, and add 2.0 ml of Stock Iodine Solution. Quantitatively transfer the mixture into a 500 ml volumetric flask, dilute to volume with water, and mix. Prepare daily.
Prepare a solution of the sample so that 5 ml of the final dilution will give an endpoint between 10 and 30 min under the conditions of the assay.
For barley malt, finely grind 25 g of the sample in a Miag-Seck mill (Buhler-Miag, Inc., P.O. Box 9497, Minneapolis, MN 55440 or similar). Quantitatively transfer the powder into a 1000 ml Erlenmeyer flask, add 500 ml of a 0.5% solution of sodium chloride, and allow the infusion to stand for 2.5 h at 30° ± 0.2°, agitating the contents by gently rotating the flask at 20-min intervals.
Caution: Do not mix the infusion by inverting the flask. The quantity of the grist left adhering to the inner walls of the flask as a result of agitation must be as small as possible.
Filter the infusion through a 32-cm fluted filter of Whatman No. 1, or equivalent, paper on a 20-cm funnel, returning the first 50 ml of filtrate to the filter. Collect the filtrate until 3 h have elapsed from the time the sodium chloride solution and the sample were first mixed. Pipet 20.0 ml of the filtered infusion into a 100 ml volumetric flask, dilute to volume with the 0.5% sodium chloride solution, and mix.
Pipet 5.0 ml of Dilute Iodine Solution into a series of 13 x 100-mm test tubes, and place them in a water bath maintained at 30° ±0.1°, allowing 20 tubes for each assay.
Pipet 20.0ml of the Buffered Substrate Solution, previously heated in the water bath for 20 min, into a 50 ml Erlenmeyer flask, and add 5.0 ml of 0.5% sodium chloride solution, also previously heated in the water bath for 20 min. Place the flask in the water bath.
At zero time, rapidly pipet 5.0 ml of the Sample Preparation into the equilibrated substrate, mix immediately by swirling, stopper the flask, and place it back in the water bath. After 10 min, transfer 1.0 ml of the reaction mixture from the 50 ml flask into one of the test tubes containing the Dilute Iodine Solution, shake the tube, then pour its contents into a Comparator Tube, and immediately compare with the Reference Colour Standard in the Comparator, using a tube of water behind the colour disk.
Note: Be certain that the pipet tip does not touch the iodine solution as carry-back of iodine to the hydrolyzing mixture will interfere with enzyme action and will affect the results of the determination.
In the same manner, repeat the transfer and comparison procedure at accurately timed intervals until the -amylase colour is reached, at which time record the elapsed time. In cases where two comparisons 30 s apart show that one is darker and the other lighter than the Reference Colour Standard, record the endpoint to the nearest quarter min. Shake out the 13-mm Comparator Tube between successive readings. Minimize slight differences in colour discrimination between operators by using a prism attachment and by maintaining a 15- to 25-cm. distance between the Comparator and the operator's eye.
One α-amylase dextrinizing unit (DU) is defined as the quantity of α-amylase that will dextrinize soluble starch in the presence of an excess of Я-amylase at the rate of 1 g/h at 30°.
Calculate the α-amylase dextrinizing units in the sample as follows:
in which W is the weight, in grams, of the enzyme sample added to the incubation mixture in the 5 ml aliquot of the Sample Preparation used; T is the elapsed dextrinizing time, in minutes; 24 is the product of the weight of the starch substrate (0.4 g) and 60 min; and M is the percent moisture in the sample, determined by suitable means.
This procedure is designed for the determination of antibacterial activity in enzyme preparation derived from microbial sources.
The assay is based on the measurement of inhibition of bacterial growth under specific circumstances.
Six organisms are tested: Staphylococcus aureus (ATCC 6538); Escherichia coli (ATCC 11229); Bacillus cereus (ATCC 2); Bacillus circulans (ATCC 4516); Streptococcus pyrogenes (ATCC 12344): and Serratia marcescens (ATCC 14041).
Make a test plate of each organism by preparing a 1:10 dilution of a 24 h Trypticase Soya Broth culture in Trypticase Agar (TSA) (for Streptococcus pyrogenes a 1:20 dilution).
Pour 15 ml of plain TSA into a Petri dish and allow the medium to harden. Overlay with 10 ml of seeded TSA and allow to solidify. Place a paper disk prepared according to Disk Preparation of the tested enzyme on each of the six inoculated plates.
Make a 10% solution of the enzyme by adding 1 g of enzyme to 9 ml of sterile, distilled water.
Mix thoroughly with a Vortex mixer to obtain a homogeneous suspension. Autoclave suitable paper disks (for instance, S & S Analytical Filter Papers No. 740-E, 12.7 mm in diameter), then saturate them with the enzyme by application of 0.1 ml (about 3 drops) of a 10% solution of the enzyme to the disk surface. Prepare six disks (one for each of the six organisms) for each enzyme: place one disk on the surface of the six inoculated agar plates.
Keep the six plates in the refrigerator overnight to obtain proper diffusion. Incubate the plates at 37° for 24 h. Examine the plates for any inhibition zones that may have been caused by the enzyme preparation.
A visually clear zone around a disk (total diameter: 16 mm) indicates the presence of antibacterial components in the enzyme preparation. If an enzyme preparation shows obvious antibacterial activity against three (or more) organisms, it is concluded that antimicrobial agents are present.
This procedure is designed for the determination of catalase activity, expressed as Baker Units.
The assay is an exhaustion method based on the breakdown of hydrogen peroxide by catalase, and the simultaneous breakdown of the catalase by the peroxide, under controlled conditions.
Reagents and Solutions
0.250 N Sodium thiosulfate: Dissolve 62.5 g of sodium thiosulfate, Na2S2O3-5H2O in 750 ml of recently boiled and cooled water, add 3.0 ml of 0.2 N sodium hydroxide as a stabilizer, dilute to 1,000 ml with water, and mix. Standardize as directed for 0.1 N Sodium thiosulfate (Volumetric Solutions), and adjust to exactly 0.250 N if necessary.
Peroxide substrate solution: Dissolve 25.0 g of anhydrous dibasic sodium phosphate (Na2HPO4), or 70.8 g of Na2HPO4-12H2O, in about 1,500 ml of water, and adjust to pH 7.0 ± 0.1 with 85% phosphoric acid. Cautiously add 100 ml of 30% hydrogen peroxide, dilute to 2,000 ml, in a graduate, and mix. Store in a clean amber bottle, loosely stoppered. The solution is stable for more than one week if kept at 5° in a full container.
Note: With freshly prepared substrate, the blank will require about 16 ml of 0.250 N sodium thiosulfate. If the blank requires less than 14 ml, the substrate solution is unsuitable and should be prepared fresh again. It is essential that the sample titration is between 50% and 80% of that required for the blank.
Pipet an aliquot of not more than 1.0 ml of the sample, previously diluted to contain approximately 3.5 Baker Units of catalase, into a 200-ml beaker. Rapidly add 100 ml of Peroxide Substrate Solution, previously adjusted to 25°, and stir immediately for 5 to 10 sec. Cover the beaker, and incubate at 25 ± 1 ° until the reaction is completed. Stir vigorously for 5 sec and then pipet 4.0 ml from the beaker into a 50-ml Erlenmeyer flask. Add 5 ml of 2 N sulfuric acid to the flask, mix, then add 5.0 ml of 40% potassium iodide, freshly prepared, and 1 drop of 1 % ammonium molybdate and mix. While continuing to mix, titrate rapidly to a colourless endpoint with 0.250 N Sodium thiosulfate, recording the required volume, in ml, as S. Perform a blank determination with 4.0 ml of Peroxide Substrate Solution, and record the required volume, in ml, as B.
Note: When preparations derived from beef liver are tested, the reaction is complete within 30 min. Preparations derived from Aspergillus and other sources may require up to 1 h. In assaying an enzyme of unknown origin, a titration should be run after 30 min and then at 10 min intervals thereafter. The reaction is complete when two consecutive titrations are the same.
One Baker Unit is that amount of catalase that will decompose 266 mg of hydrogen peroxide under the conditions of the assay. Calculate the activity of the sample by the formula:
in which C is the ml of aliquot of original enzyme preparation added to each 100 ml of Peroxide Substrate Solution, or, when 1 ml of diluted enzyme is used, C is the dilution factor.
Application and Principle
This assay is based on the enzymatic hydrolysis of the interior P-l,4-glucosidic bonds of a defined carboxymethylcellulose substrate at pH 4.5 and at 40°. The corresponding reduction in substrate viscosity is determined with a calibrated viscometer.
Calibrated Viscometer: Use a size 100 Calibrated Cannon-Fenske Type Viscometer, or its equivalent (Scientific Products, Catalog No. P2885-100).
Constant-Temperature Glass Water Bath: (40 ±0.1°) Use a constant-temperature glass water bath, or its equivalent
Stopwatches: Use two stopwatches, Stopwatch No. 1, calibrated In 1/10 min for determining the reaction time (Tr), and Stopwatch No. 2, calibrated in 1/5 s for determining the efflux time (Tt).
Waring Blender: Use a two-speed Waring blender, or its equivalent (Scientific Products, Catalog No. 58350-1).
Reagents and Solutions
Acetic Acid Solution (2 N): While agitating a 1 l beaker containing 800 ml of water, carefully add 116 ml of glacial acetic acid. Cool to room temperature. Quantitatively transfer the solution to a 1 l volumetric flask, and dilute to volume with water.
Sodium Acetate Solution (2 N): Dissolve 272.16 g of sodium acetate trihydrate in approximately 800 ml of water contained in a 1 liter beaker. Quantitatively transfer to a 1 liter volumetric flask, and dilute to volume with water.
Acetic Acid Solution (0.4 N): Transfer 200 ml of Acetic Acid Solution (2 N) into a 1 liter volumetric flask, and dilute to volume with water.
Sodium Acetate Solution (0.4 N): Transfer 200 ml of Sodium Acetate Solution (2 N) into a 1 liter volumetric flask, and dilute to volume with water.
Acetate Buffer (pH 4.5): Using a standardized pH meter, add Sodium Acetate Solution (0.4 N) with continuous agitation to 400 ml of Acetic Acid Solution (0.4 N) in a suitable flask until the pH is 4.5 ±0.05.
Sodium Carboxymethylcellulose: Use sodium carboxymethylcellulose (Hercules, Inc., CMC Type 7HF or equivalent).
Sodium Carboxymethylcellulose Substrate (0.2% w/v): Transfer 200 ml of water into the bowl of the Waring blender. With the blender on low speed, slowly disperse 1.0 g (moisture-free basis) of the Sodium Carboxymethylcellulose into the bowl, being careful not to splash out any of the liquid. Using a rubber policeman to assist, wash down the sides of the glass bowl with water. Place the top on the bowl and blend at high speed for 1 min. Quantitatively transfer to a 500-ml volumetric flask, and dilute to volume with water. Filter the substrate through gauze before use.
Prepare an enzyme solution so that 1 ml of the final dilution will produce a relative fluidity change between 0.18 and 0.22 in 5 min under the conditions of the assay. Weigh the enzyme, and quantitatively transfer it to a glass mortar. Triturate with water and quantitatively transfer the mixture to an appropriate volumetric flask. Dilute to volume with water, and filter the enzyme solution through Whatman No. 1 filter paper before use.
Place the Calibrated Viscometer in the 40 ± 0.1° water bath in an exactly vertical position. Use only a scrupulously clean viscometer. (To clean the viscometer, draw a large volume of detergent solution followed by water through the viscometer by using an aspirator with a rubber tube connected to the narrow arm of the viscometer.) Pipet 20 ml of filtered Sodium Carboxymethylcellulose Substrate and 4 ml of Acetate Buffer into a 50-ml Erlenmeyer flask. Allow at least two flasks for each enzyme sample and one flask for a substrate blank. Stopper the flasks, and equilibrate them in the water bath for 15 min. At zero time, pipet 1 ml of the enzyme solution into the equilibrated substrate. Start stopwatch no. 1, and mix the solution thoroughly. Immediately pipet 10 ml of the reaction mixture into the wide arm of the viscometer. After approximately 2 min, apply suction with a rubber tube connected to the narrow arm of the viscometer, drawing the reaction mixture above the upper mark into the driving fluid head. Measure the efflux time by allowing the reaction mixture to freely flow down past the upper mark. As the meniscus of the reaction mixture falls past the upper mark, start stopwatch no. 2. At the same time, record the reaction time, in minutes, from stopwatch no. 1 (Tr). As the meniscus of the reaction mixture falls past the lower mark, record the time, in seconds, from stopwatch no. 2 (Tt).
Repeat the final step until a total of four determinations are obtained over a reaction time (Tr) of not more than 15 min.
Prepare a substrate blank by pipetting 1 ml of water into 24 ml of buffered substrate. Pipet 10 ml of the reaction mixture into the wide arm of the viscometer. Determine the time (Ts) in seconds required for the meniscus to fall between the two marks. Use an average of five determinations for (Ts).
Prepare a water blank by pipetting 10 ml of equilibrated water into the wide arm of the viscometer. Determine the time (Tw) in seconds required for the meniscus to fall between the two marks. Use an average of five determinations for (Tw).
One Cellulase Unit (CU) is defined as the amount of activity that will produce a relative fluidity change of 1 in 5 min in a defined carboxymethylcellulose substrate under the conditions of the assay.
Calculate the relative fluidities (Fr) and the (Tn) values for each of the four efflux times (Tt) and reaction times (Tr) as follows:
Fr is the relative fluidity for each reaction time;
Ts is the average efflux time, in seconds, for the substrate blank;
Tw is the average efflux time, in seconds, for the water blank;
Tt is the efflux time, in seconds, of reaction mixture;
Tr is the elapsed time, in minutes, from zero time, that is, the time from addition of the enzyme solution to the buffered substrate until the beginning of the measurement of efflux time (Tt);
Tn is the reaction time, in minutes (Tr), plus one-half of the efflux time (Tt), converted to minutes.
Plot the four relative fluidities (Fr) as the ordinate against the four reaction times (Tn) as the abscissa. A straight line should be obtained. The slope of this line corresponds to the relative fluidity change per minute and is proportional to the enzyme concentration. The slope of the best line through a series of experimental points is a better criterion of enzyme activity than is a single relative fluidity value. From the graph, determine the Fr values at 10 and 5 min. They should have a difference in fluidity of not more than 0.22 or less than 0.18. Calculate the activity of the enzyme unknown as follows:
Fr5 is the relative fluidity at 5 min of reaction time;
Fr10 is the relative fluidity at 10 min of reaction time;
1000 is the milligrams per gram;
W is the weight, in mg of enzyme added to the reaction mixture in a 1 -ml aliquot of enzyme solution.
This procedure is designed to detect the presence of ethylenimine in immobilized enzyme preparations containing poly(ethylenimine).
The principle of the method is to react any free ethylenimine which may be present in a sample of immobilized enzyme preparation with an aqueous solution of 1,2-naphthoquinone-4-sulfonate (Folin's reagent) to produce 4-(1-aziridinyl)-1,2-naphthoquinone. This reaction product is extracted into chloroform and the extract analyzed by high performance liquid chromatography (HPLC).
- High performance liquid chromatograph equipped with an ultraviolet detector (254 nm), injection valve and Lichrosorb DIOL column, 5 nm, 4.6-mm i.d. x 25-cm (or equivalent)
- Glass syringe 10 μl
- Separatory funnel, 100 ml
- Pipettes of convenient volumes for the preparation of standard solutions.
Reagents and Solutions
Chloroform with 1 % ethanol as a stabilizer, UV grade, distilled in glass
Hexane, UV grade, distilled in glass
2-propanol, UV grade, distilled in glass
Methyl alcohol, UV grade, distilled in glass
Acetone, UV grade, distilled in glass
1,2-naphthoquinone-4-sulfonic acid, sodium salt
0.1 N sodium hydroxide (NaOH)
0.1 MPotassium dihydrogen phosphate (KH2PO4)
Buffer Solution: pH 7.7; mix 200 ml of 0.1 M KH2PO4 with 93.4 ml of 0.1 NNaOH.
Folin's Reagent: Dissolve 0.40 g of 1,2-naphtoquinone-4-sulfonic acid sodium salt in 100 ml of buffer solution. Dilute to 500 ml with distilled water in a volumetric flask. Wrap the flask in aluminium foil and store in the refrigerator. Discard the reagent after five days.
4-(1 -Aziridinyl)-1 ,2-naphthoquinone
A standard sample of known purity is required. If a commercial source for this standard is not readily available, the substance may by synthesized by the following procedure:
Wrap a separatory funnel with aluminium foil and add 2 g of the sodium salt of 1,2-naphthoquinone-4-sulfonic acid dissolved in 250 ml of distilled water.
Add 25 ml of 0.5 M trisodium phosphate, shake and check that the pH is between 10.5 and 11.5. Add 0.3 ml ethylenimine and shake intermittently for 10 min.
Caution: Ethylenimine has been identified as a carcinogen. Appropriate precautions must be taken in handling the compound to avoid personnel exposure and area contamination.
Extract the 4-(aziridinyl)-1,2-naphthoquinone formed with six 200-ml portions of chloroform.
Place the combined extracts in a 2-liter beaker wrapped in aluminium foil in which three holes have been made.
Evaporate the chloroform at room temperature with a nitrogen purge. Transfer the dry residue to a 50-ml beaker wrapped in aluminium foil.
Add 35 ml of methyl alcohol and 1 ml of chloroform to the residue and stir briefly. Not all of the residue will dissolve.
Place the beaker in an ice-water bath for 10 min and then filter the precipitate through Whatman 42 filter paper.
Rinse the precipitate in the filter with 4 ml of chilled methyl alcohol and discard the filtrates.
Dry the precipitate with a nitrogen purge, transfer it to a brown glass bottle and purge again. Dry the compound overnight in a desiccator containing Drierite. The melting point of the compound is 173-175°. The compound is to be used for making standard solutions for calibration purposes. The compound should be stored in a freezer until standard solutions are to be prepared.
0.5 g/l Standard Solution: Accurately weigh about 125 mg of 4-(1-aziridinyl)-1,2-naphthoquinone into a 250 ml volumetric flask [low actinic glass] and add chloroform to the mark.
0.1 mg/l Standard Solution: By appropriate dilution(s) of the 0.5 g/l Standard Solution, prepare a standard solution which contains 0.1 mg/L (0.1 ng/μl).
Accurately weigh a sample of immobilized enzyme preparation containing about 10 g of dry matter into an aluminium foil-covered beaker. Add 50 ml of Folin's Reagent and agitate the mixture for several minutes. Decant the Folin's Reagent into a separatory funnel and extract with 2 ml of chloroform. Analyze a 20 μl portion of the chloroform extract by the following chromatographic conditions:
Column: Lichrosorb DIOL 5 nm (or equivalent)
Mobile phase: hexane:chloroform (with 1% ethanol) : isopropanol = 59.5 : 40.0 : 0.5 (v/v)
Flow rate: 2 ml/min.
Inject a 20 μl portion of the 0.1 mg/L Standard Solution. The sample response is not greater than that of the 0.1 mg/L Standard Solution. (Another sample containing a standard addition of 4-(1-aziridinyl)-1,2-naphthoquinone to immobilized enzyme preparation should be analyzed to verify that the chromatographic response does not contain interfering substances.)
Application and Principle
This procedure is used to determine Я-Galactosidase activity of enzyme preparations derived from Aspergillus oryzae var. The assay is based on a 15-min hydrolysis of an o-nitrophenyl-b-D-galactopyranoside substrate at 37° and pH 4.5.
Reagents and Solutions
2.0 N Acetic Acid: Dilute 57.5 ml of glacial acetic acid to 500 ml with water. Mix well, and store in a refrigerator.
4.0 N Sodium Hydroxide: Dissolve 40.0 g of sodium hydroxide in sufficient water to make 250 ml.
Acetate Buffer: Combine 50 ml of 2.0 N Acetic Acid and 11.3 ml of 4.0 N Sodium Hydroxide in a 1000-ml volumetric flask, and dilute to volume with water. Verify that the pH is 4.50 6 0.05, using a pH meter, and adjust, if necessary, with 2.0 N Acetic Acid or 4.0 N Sodium Hydroxide.
2.0 mM o-Nitrophenol Stock: Transfer 139.0 mg of o-nitrophenol a 500-ml volumetric flask, dissolve in 10 ml of USP alcohol (95% ethanol) by swirling, and dilute to volume with 1% sodium carbonate.
0.10 mM Standard Solution: Pipet 5.0 ml of the 2.0 mM o-Nitrophenol Stock solution into a 100-ml volumetric flask, and dilute to volume with 1% sodium carbonate solution.
0.14 mM Standard Solution: Pipet 7.0 ml of the 2.0 mM o-Nitrophenol Stock solution into a 100-ml volumetric flask, and dilute to volume with 1% sodium carbonate solution.
0.18 mM Standard Solution: Pipet 9.0 ml of the 2.0 mM o-Nitrophenol Stock solution into a 100-ml volumetric flask, and dilute to volume with 1% sodium carbonate solution.
Substrate: Transfer 370.0 mg of o-nitrophenyl-Я-D-galactopyranoside to a 100-ml volumetric flask, and add 50 ml of Acetate Buffer. Swirl to dissolve, and dilute to volume with Acetate Buffer.
Note: Perform the assay procedure within 2 h of Substrate preparation.
Prepare a solution from the test sample preparation such that 1 ml of the final dilution will contain between 0.15 and 0.65 lactase unit. Weigh, and quantitatively transfer the enzyme to a volumetric flask of appropriate size. Dissolve the enzyme in water, swirling gently, and dilute with water if necessary.
Note: Perform the assay procedure within 2 h of dissolution of the Test Preparation.
Determine the absorbance of the three o-Nitrophenol Standards at 420 nm in a 1-cm cell, using a suitable spectrophotometer. Use water to zero the instrument. Calculate the millimolar extinction, M, for each of the o-Nitrophenol Standards (0.10, 0.14, and 0.18 mM) by the equation
in which An is the absorbance of each o-Nitrophenol Standard at 420 nm and C is the corresponding concentration of onitrophenol in the standard. M for each standard should be approximately 4.60/mM. Perform a linear regression analysis of the absorbance readings of the three o-Nitrophenol Standards versus the o-nitrophenol concentration in each (0.10, 0.14, and 0.18 mM). The r2 should not be less than 0.99. Determine the mean M of the three o-Nitrophenol Standards for use in the calculations below.
For each sample or blank, pipet 2.0 ml of the Substrate solution into a 25 x 150-mm test tube, and equilibrate in a water bath maintained at 37.0 ±0.1° for approximately 10 min. At zero time, rapidly pipet 0.5 ml of the Test Preparation (or 0.5 ml of water as a blank) into the equilibrated substrate, mix by brief (1 s) vortex, and immediately return the tubes to the water bath. After exactly 15 min of incubation, rapidly add 2.5 ml of 10% sodium carbonate solution, and vortex the tube to stop the enzyme reaction. Dilute the samples and blanks to 25.0 ml by adding 20.0 ml of water, and thoroughly mix. Determine the absorbance of the diluted samples and blanks at 420 nm in a 1 -cm cell, using a suitable spectrophotometer. Use water to zero the instrument.
One lactase unit (ALU) is defined as that quantity of enzyme that will liberate o-nitrophenol at a rate of 1 mmol/min under the conditions of the assay. Calculate the activity (lactase activity per gram) of the enzyme preparation taken for analysis as follows:
AS is the average of absorbance readings for the Test Preparation;
B is the average of absorbance readings for the blank;
25 is the final volume, in ml, of the diluted incubation mixture;
ε is the mean absorptivity of the o-Nitrophenol Standards per micromole;
15 is the incubation time, in minutes; and
W is the weight, in grams, of original enzyme preparation contained in the 0.5-ml aliquot of Test Preparation used in the incubation.
Application and Principle
This procedure is used to determine the glucoamylase activity of preparations derived from Aspergillus niger var., but it may be modified to determine preparations derived from Aspergillus oryzae var. and Rhizopus oryzae var. (as indicated by the variations in the text below). The sample hydrolyzes p-nitrophenyl-α-D-glucopyranoside (PNPG) to p-nitrophenol (PNP) and glucose at pH 4.3 and 50°.
Use the quantity of PNP liberated per unit of time to calculate the enzyme activity. Measure the PNP liberated against a quantity of a standard preparation of PNP by measuring the absorbance of the solutions at 400 nm after adjusting the pH of the reaction mixture to pH 8.0.
Note: Use a pH of 5.0 when testing preparations derived from Aspergillus oryzae var. or Rhizopus oryzae var.
Water Bath: Use an open, circulating water bath with control accuracy of at least ±0.1°.
Spectrophotometer: Use a spectrophotometer suitable for measuring absorbances at 400 nm.
Cuvettes: Use 10-mm light-path fused quartz.
Thermometer: Use a partial immersion thermometer with a suitable range, graduated in 1/10°.
Timer: Use a solid-state timer, model 69240 (GCS Corporation, Precision Scientific Group), or equivalent, accurate to ± 0.01 min in 240 min.
Vortex Mixer: Use a standard variable-speed mixer.
Reagents and Solutions
p-Nitrophenol Stock Solution (PNP) (0.001 M): Dissolve 139.11 mg of p-nitrophenol previously dried (60°, maximum 4 h) into water, and dilute to 1000 ml.
Caution: Avoid contact with skin. If contact occurs, wash the affected area with water. Work in a well-ventilated area.
Acetate Buffer Solution: (0.1 M) Dissolve 4.4 g of sodium acetate trihydrate (NaC2H3O2.3H2O) in approximately 800 ml of water, add 4.5 ml of acetic acid (C2H4O2). Adjust to pH 4.5 ± 0.05 by adding either sodium acetate or glacial acetic acid as required. Dilute to 1 l.
Note: Use a pH of 5.0 when testing preparations derived from Aspergillus oryzae var. or Rhizopus oryzae var.
Sodium Carbonate Solution (0.3 M): Dissolve 15.9 g of sodium carbonate (Na2CO3) in water, and dilute to 500 ml.
p-Nitrophenyl-a-D-glucopyranoside Solution (PNPG): Dissolve 100.0 mg of PNPG (Sigma Chemical Co., Catalog No. N1377 or equivalent) in acetate buffer, and dilute to 100 ml.
Dilute three portions of PNP Stock Solution to produce standards for the standard curve. Add 3 ml of the PNP Stock Solution to 125 ml of Sodium Carbonate Solution, and dilute to 500 ml with water to produce the first standard, containing 0.006 mmol/ml. Add 2 ml of PNP Stock Solution to 25 ml of Sodium Carbonate Solution, and dilute to 100 ml with water to produce the second standard, containing 0.02 mmol/ml. Add 5 ml of PNP Stock Solutions to 25 ml of Sodium Carbonate Solution, and dilute to 100 ml with water to produce the third standard, containing 0.05 mmol/ml.
Dilute 1.00 ± 0.01 g of sample in sufficient Acetate Buffer Solution to produce a solution that contains between 0.1 and 0.3 glucoamylase units of activity per ml.
Measure absorbances of each of the three PNP Standard Solutions to calculate the molar extinction coefficient. Equilibrate the PNPG Solution in a 50° water bath for at least 15 min. For active samples, transfer 2.0 ml of the Sample Solution to a test tube. Loosely stopper, and place the tube in the water bath to equilibrate for 5 min. At zero time, add 2.0 ml of PNPG Solution, and mix at moderate speed on a vortex mixer. Return the mixture to the water bath. Exactly 10.0 min later, add 3.0 ml of the Sodium Carbonate Solution, mix on the vortex, and remove from the water bath.
For sample blanks, transfer 2.0 ml of the Sample Solution and 3.0 ml of the Sodium Carbonate Solution into a test tube, and mix. Add 2.0 ml of PNPG Solution, and mix. Measure the absorbance of each sample and the blank versus water in a 10-mm cell.
Note: Determine the absorbance of the sample and blank solutions not more than 20 min after adding Sodium Carbonate Solution.
One unit of glucoamylase activity is defined as the amount of glucoamylase that will liberate 0.1 mmol/min of p-nitrophenol from the PNPG Solution under the conditions of the assay.
Calculate the millimolar extinction of the PNP standards using the following equation:
An is the absorbance of the p-nitrophenol standard, at 400 nm, and
C is concentration, in mmol/ml, of p-nitrophenol.
The averaged millimolar extinction coefficient, M, should be approximately 18.2.
AS is the sample absorbance;
AB is the blank absorbance;
F is the appropriate dilution factor;
W is the weight of sample, in grams; 7 is the final volume of the test solutions;
10 is the reaction time, in minutes; 0.10 is the amount of PNP liberated, in mmol/min/unit of enzyme;
2 is the sample aliquot, in millilitres; and
M is the millimolar extinction coefficient.
Application and Principle
This procedure is used to determine Я-glucanase activity of enzyme preparations derived from Aspergillus niger var. and Bacillus subtilis var. The assay is based on a 15-min hydrolysis of lichenin substrate at 40° and at pH 6.5. The increase in reducing power due to liberated reducing groups is measured by the neocuproine method.
Reagents and Solutions
Phosphate Buffer: Dissolve 13.6 g of monobasic potassium phosphate in about 1900 ml of water, add 70% sodium hydroxide solution until the pH is 6.5 ± 0.05, then transfer the solution into a 2000-ml volumetric flask, dilute to volume with water, and mix.
Neocuproine Solution A: Dissolve 40.0 g of anhydrous sodium carbonate, 16.0 g of glycine, and 450 mg of cupric sulfate pentahydrate in about 600 ml of water. Transfer the solution into a 1000-ml volumetric flask, dilute to volume with water, and mix.
Neocuproine Solution B: Dissolve 600 mg of neocuproine hydrochloride in about 400 ml of water, transfer the solution into a 500-ml volumetric flask, dilute to volume with water, and mix. Discard when a yellow colour develops.
Lichenin Substrate: Grind 150 mg of lichenin (Sigma Chemical Co., Catalog No. L-6133, or equivalent) to a fine powder in a mortar, and dissolve it in about 50 ml of water at about 85°. After solution is complete (20 to 30 min), add 90 mg of sodium borohydride and continue heating below the boiling point for 1 h. Add 15 g of Amberlite MB-3, or an equivalent ion-exchange resin, and stir continuously for 30 min. Filter with the aid of a vacuum through Whatman No. 1 filter paper, or equivalent, in a Buchner funnel, and wash the paper with about 20 ml of water. Add 680 mg of monobasic potassium phosphate to the filtrate, and re-filter through a 0.22-mm Millipore filter pad, or equivalent. Wash the pad with 10 ml of water, and adjust the pH of the filtrate to 6.5 ± 0.05 with 1 N sodium hydroxide or 1 N hydrochloric acid. Transfer the filtrate into a 100-ml volumetric flask, dilute to volume with water, and mix. Store at 2° to 4° for not more than 3 days.
Glucose Standard Solution: Dissolve 36.0 mg of anhydrous dextrose in Phosphate Buffer in a 1000-ml volumetric flask, dilute to volume with water, and mix.
Prepare a solution from the enzyme preparation sample so that 1 ml of the final dilution will contain between 0.01 and 0.02 Я-glucanase units. Weigh the sample, transfer it into a volumetric flask of appropriate size, dilute to volume with Phosphate Buffer, and mix.
Pipet 2 ml of Lichenin Substrate into each of four separate test tubes graduated at 25 ml, and heat the tubes in a water bath at 40° for 10 to 15 min to equilibrate.
After equilibration, add 1 ml of Phosphate Buffer to tube 1 (substrate blank), 1 ml of Glucose Standard Solution to tube 2 (glucose standard), 4 ml of Neocuproine Solution A and 1 ml of the Test Preparation to tube 3 (enzyme blank), and 1 ml of the Test Preparation to tube 4 (sample). Prepare a fifth tube for the buffer blank, and add 3 ml of Phosphate Buffer.
Incubate the five tubes at 40° for exactly 15 min, and then add 4 ml of Neocuproine Solution A to tubes 1, 2, 4, and 5. Add 4 ml of Neocuproine Solution B to all five tubes, and cap each with a suitably sized glass marble.
Caution: Do not use rubber stoppers.
Heat the tubes in a vigorously boiling water bath for exactly 12 min to develop colour, then cool to room temperature in cold water, and adjust the volume of each to 25 ml with water. Cap the tubes with Parafilm, or other suitable closure, and mix by inverting several times. Determine the absorbance of each solution at 450 nm in 1-cm cells, with a suitable spectrophotometer, against the buffer blank in tube 5.
One Я-glucanase unit (BGU) is defined as that quantity of enzyme that will liberate reducing sugar (as glucose equivalent) at a rate of 1mmol/min under the conditions of the assay.
Calculate the activity of the enzyme preparation taken for analysis as follows:
A4 is the absorbance of the sample (tube 4),
A3 is the absorbance of the enzyme blank (tube 3),
A2 is the absorbance of the glucose standard (tube 2),
A1 is the absorbance of the substrate blank (tube 1), 36 is the micrograms of glucose in the Glucose Standard Solution,
106 is the factor converting micrograms to grams,
180 is the weight of 1 µmol of glucose, and
15 is the reaction time in minutes.
This procedure is designed for the determination of glucose isomerase preparations derived from Actinoplanes missouriensis, Arthrobacter globiformis, Bacillus coagulans, Streptomyces olivaceus, Streptomyces olivochromogenes, and Streptomyces rubiginosus.
The assay is based on measurement of the rate of conversion of glucose to fructose in a packed bed reactor.
The procedure as outlined approximates an initial velocity assay method. Specific conditions are: glucose concentration, 45% w/w; pH (inlet) measured at room temperature in the 7.0 to 8.5 range, as specified; temperature, 60.0°; and magnesium concentration, 4 x 10-3 M. The optimum conditions for enzymes from different microbial sources and methods of preparation may vary; therefore, if different pH conditions, buffering systems, or methods of sample preparation are recommended by the manufacturer, such variations in the instructions given herein should be used.
Reagents and Solutions
Glucose substrate: Dissolve 539 g of anhydrous glucose and 1.0 g of magnesium sulfate, MgSO4.7H2O, in 700 ml of water or the manufacturer's recommended buffer, previously heated to 50° to 60°. Cool the solution to room temperature, and adjust the pH as specified by the enzyme manufacturer. Transfer the solution to a 1,000-ml volumetric flask, dilute to volume with water or the specified buffer, and mix. Transfer to a vacuum flask, and de-aerate for 30 min.
Magnesium sulfate solution: Dissolve 1.0 g of magnesium sulfate, MgSO4.7H2O, in 700 ml of water. Adjust the pH to 7.5 to 8.0 as specified by the manufacturer, using 1 N sodium hydroxide, dilute to 1,000 ml with water and mix.
Note: Glucose isomerase activity of the commercial enzyme is usually determined on the enzyme that has been immobilized by binding with a polymer matrix or other suitable material. This method is designed for use with such preparations.
Column Assembly and Apparatus
The column assembly is shown in below.
Note: Make all connections with inert tubing, glass or plastic as appropriate.
Use a 2.5 x 40-cm glass column provided with a coarse sintered glass bottom and a water jacket connected to a constant-temperature water bath, maintained at 60.0° by means of a circulating pump. Connect the top of the column to a variable-speed peristaltic pump having a maximum flow rate of 800 ml per h. The diameter of the tubing with which the peristaltic pump is fitted should permit variation of the pumping volume from 60 to 150 ml per h. Connect the outlet of the column with a collecting vessel.
. Diagram of a column assembly for assay of Immobilized Glucose Isomerase
Transfer to a 500-ml vacuum flask an amount of the sample, accurately weighed in g or measured in ml, as appropriate, sufficient to obtain 2,000 to 8,000 glucose isomerase units (GIc U). Add 200 ml of Glucose Substrate, stir gently for 15 sec and repeat the stirring every 5 min for 40 min. De-aerate by vacuum for 30 min.
Quantitatively transfer the Sample Preparation to the column with the aid of Magnesium Sulfate Solution as necessary. Allow the enzyme granules to settle, and then place a porous disk so that it is even with, and in contact with, the top of the enzyme bed. All of the air should be displaced from the disk. Place a cotton plug about 1 or 2 cm above the disk. (This plug acts as a filter. It ensures proper heating of the solution and traps dissolved gases that may be present in the Glucose Substrate.) Connect the tubing from the peristaltic pump with the top of the column, and seal the connection by suitable means in order to protect the column contents from the atmosphere. Place the inlet tube of the peristaltic pump into the Glucose Substrate solution, and begin a downward flow of the Glucose Substrate into the column at a rate of at least 80 ml per h. Maintain the flow rate for 1 h at room temperature.
Adjust the flow of the Glucose Substrate to such a rate that a fractional conversion of 0.2 to 0.3 will be produced, based on the estimated activity of the sample. The fractional conversion is calculated from optical rotation values obtained on the starting Glucose Substrate and the sample effluent, as specified in Calculations below. After the correct flow rate has been established, run the column overnight (16 h minimum), then check the pH of the Glucose Substrate, and readjust if necessary to the specified pH. Measure the flow rate, and collect a sample of the column effluent. Cover the effluent sample, allow it to stand for 30 min at room temperature, and then determine the fractional conversion of glucose to fructose (see Calculations below). If the conversion is less than 0.2 or more than 0.3, adjust the flow rate to bring the conversion into this range. If a flow rate adjustment is required, collect an additional effluent sample after allowing the column to re-equilibrate for at least 2 h and then determine the fractional conversion.
Measure the flow rate, and collect an effluent sample. Cover the sample, let it stand at room temperature for 30 min, and determine the fractional conversion.
Measure the optical rotation of the effluent sample and of the starting Glucose Substrate at 25.0°, and calculate their specific rotations by the formula:
a is the corrected observed rotation, in degrees,
1 is the length of the polarimeter tube, in dm,
p is the concentration of the test solution, expressed as g of solute per 100 g of solution, and
d is the specific gravity of the solution at 25°
Calculate the fractional conversion, X, by the formula:
αE is the specific rotation of the column effluent,
αs is the specific rotation of the Glucose Substrate,
and αF is the specific rotation of fructose (which in this case has been calculated to be - 94.54).
The enzyme activity is expressed in glucose isomerase units (CIcU, the subscript c signifying column process). One GIcU is defined as the amount of enzyme that converts glucose to fructose at an initial rate of 1 μmol per min, under the conditions specified.
Calculate the glucose isomerase activity by the formula:
F is the flow rate, in ml per min,
S is the concentration of the Glucose Substrate, in μml per ml,
Xe is the fractional conversion at equilibrium, or 0.51, and
W is the weight or volume of the sample taken, in g or ml, respectively.
Application and Principle
This procedure is used to determine glucose oxidase activity in preparations derived from Aspergillus niger var. The assay is based on the titrimetric measurement of gluconic acid produced in the presence of excess substrate and excess air.
Reagents and Solutions
Chloride-Acetate Buffer Solution: Dissolve 2.92 g of sodium chloride and 4.10 g of sodium acetate in about 900 ml of water. Adjust the pH to 5.1 with either dilute acetic acid or dilute sodium hydroxide solution and dilute to 1000.0 ml.
Sodium Hydroxide Solution (0.1 N)
Hydrochloric Acid Solution (0.05 N) Standardized.
Phenolphthalein Solution (2% w/v): Solution in methanol.
Octadecanol Solution: Saturated solution in methanol.
Substrate Solution: Dissolve 30.00 g of anhydrous glucose in 1000 ml of the Chloride-Acetate Buffer Solution.
Dissolve an accurately weighed amount of enzyme preparation in the Chloride-Acetate Buffer Solution, and dilute in the buffer solution to obtain an enzyme activity of 5 to 7 activity units per milliliter.
Transfer 25.0 ml of the Substrate Solution to a 32 x 200-mm test tube. To a second 32 x 200 mm test tube transfer 25.0 ml of the Chloride-Acetate Buffer Solution (blank). Equilibrate both tubes in a 35 ± 0.1° water bath for 20 min. Add 3.0 ml of the Sample Preparation to each test tube, mix, and insert a glass sparger into each tube with a pre-adjusted air flow of 700 to 750 ml/min. If excessive foaming occurs, add 3 drops of the Octadecanol Solution to each tube. After exactly 15 min, remove the sparge and rinse any adhering reaction mixture back into the tube with water. Immediately add 10 ml of the Sodium Hydroxide Solution and 3 drops of the Phenolphthalein Solution to each tube. Insert a small magnetic stirrer bar, stir, and titrate to the phenolphthalein endpoint with the standardized 0.05 N hydrochloric Acid Solution.
One Glucose Oxidase Titrimetric unit of activity (GOTu) is the quantity of enzyme that will oxidize 3 mg of glucose to gluconic acid under the conditions of the assay. Determine the enzyme activity using the following equation:
B is the titration volume, in milliliters, of the blank;
T is the titration volume, in milliliters, of the sample;
N is the normality of the titrant; 180 is the molecular weight of glucose;
F is the sample dilution factor;
3 is from the unit definition; and
W is the weight, in grams, of the enzyme preparation contained in each milliliter of the sample solution.
This procedure is designed to determine the glutaraldehyde carried over into isomerized syrup during isomerization of glucose syrup by the use of immobilized glucose isomerases crosslinked with glutaraldehyde.
The procedure involves sampling the syrup produced during different stages of the enzyme assay "Glucose isomerase activity". Analysis of the sample syrup according to the procedure on page 169 gives the number of mg of glutaraldehyde per kg of syrup. A subsequent calculation gives the amount of glutaraldehyde present per unit of glucose isomerase activity. The enzyme preparation passes the test if the average result is not greater than 0.025.
Samples of syrup during the assay for "Glucose isomerase activity" are taken at steps as prescribed in the following:
Sample 1: 25 ml of syrup is taken out at the step called "Sample preparation" (i.e. syrup decanted off, just after the prescribed 40 min soaking period).
Sample 2: 25 ml of syrup is taken out at the step called "Procedure" (i.e. isomerized syrup from the column outlet just after the flow rate has been adjusted to the correct level).
Sample 3: 25 ml of syrup is taken out at the point of time when samples are taken for determination of the fractional conversion of the glucose to fructose.
As prescribed, this time is at least 16 hours after start-up. In actual practice the time for taking this effluent sample will be in the interval 42-48 hours after start-up.
All three samples (Samples 1, 2, and 3) are subjected to determination for glutaraldehyde as described in "Determination of glutaraldehyde in High Fructose Corn Syrup". As indicated in the text of the assay, it has been determined that the lower detection limit for glutaraldehyde in HFCS (High Fructose Corn Syrup) is 5 mg/kg by this assay.
The relationship between the determination of glutaraldehyde and the determination of activity of the prepared immobilized enzyme can be expressed in the following way:
GA is Glutaraldehyde
GIcU is the activity unit for glucose isomerase in the column process
Interpretation of test results
The enzyme passes test if the average "a" from the three samples tested is not greater than 0.025. (For GA concentrations below the detection limit of 5 mg/kg, the value 5 mg/kg is taken.)
- a = 0.025 is equal to an average GA concentration of 5 mg/kg from 200 GIcU/g enzyme.
- a = 0.025 is equal to an average GA concentration of 7.5 mg/kg from 300 GIcU/g enzyme.
This procedure is designed for the determination of Glutaraldehyde in High Fructose Corn Syrup (HFCS).
The assay is based on a measurement using thin layer chromatography.
TLC plates: Pre-coated TLC plates SIL G-25, available from Macherey-Nagel, Catalog No. 809 013, or equivalent. Activate before use by heating to 100° for at least one h. Use gloves when handling.
Solvent system: Transfer 5.0 ml absolute ethanol to a 100-ml volumetric flask and fill up to the mark with chloroform. Transfer to a 250-ml flask and shake very thoroughly before pouring the mixture into the developing chamber.
Spray reagents: (Sufficient for one TLC plate)
- 1% MBTH: Dissolve 250 mg MBTH (N-methyl-benzothiazolonhydrazon-HCl) in 25 ml water.
- 2% Ferric chloride: Dissolve 0.5 g ferric chloride (FeCl3-6H2O) in 25 ml water.
Glutaraldehyde stock solution (1 mg/ml): Transfer 0.4 ml of 25% glutardialdehyde solution (Merck No. 12179) to a 100-ml volumetric flask. Make up to the mark with water.
Glutaraldehyde solution (25 μg/ml): Dilute 250 μl of glutaraldehyde stock solution to 10.0 ml with water. Dilution to be made freshly before use.
Glutaraldehyde solution (3.75 μg/ml): Dilute 1.50 ml of G - 25 μg/ml to 10.0 ml with water. Dilution to be made freshly before use.
Transfer to 10-ml volumetric flasks:
Assay solution (a): 7.50 g of HFCS sample;
Assay solution (b): 7.50 g of HFCS sample and 1.50 ml of glutaraldehyde solution (25 μg/ml) corresponding to 37.5 μg of glutaraldehyde.
Make both solutions up to volume with water.
Treat the standard and assay solutions for 30 min in an ultra-sonic bath immediately before use.
Spot the TLC plate as follows:
- Spot 1: 150 μl of glutaraldehyde solution (3.75 μg/ml) equivalent to 0.5625 μg glutaraldehyde.
- Spot 2: 150 μl of assay solution (b) equivalent to 0.5625 μg glutaraldehyde plus 0.1125 g HFCS sample.
- Spot 3: 150 μl of assay solution (a) equivalent to 0.1125 g HFCS sample.
The spots should be placed at least 3 cm from the edges of the plate and 5 cm apart. Allow the spots to dry at room temperature. Run the chromatogram until the solvent front has migrated 15 cm (30-40 min). Allow the plate to dry for at least 30 min at room temperature.
Spray with reagent I using a fine nozzle. Approximately 20 ml are needed.
Wait for 10 min and then spray with reagent II until the spots can be seen. Approximately 25 ml are needed.
Estimate the glutaraldehyde content of the assay sample (spot 3) by comparison with the standard (spot 1).
If the intensity of assay sample spot 3 is less than the intensity of standard spot 1, then the HFCS sample contains < 5 mg/kg of glutaraldehyde.
Spot 2 is included as proof that the method can detect 5 mg/kg of glutaraldehyde in HFCS.
This procedure is for the determination of hemicellulase activity of preparations derived from Aspergillus niger, var.
The test is based on the enzymatic hydrolysis of the interior glucosidic bonds of a defined carob (locust) bean gum substrate at pH 4.5 and 40°. The corresponding reduction in substrate viscosity is determined with a calibrated viscometer.
Viscometer: Use a size 100 calibrated Cannon-Fenske Type Viscometer, or its equivalent. A suitable viscometer is supplied as Catalog No. 2885-100 by Scientific Products, 1210 Waukegan Road, McGraw Park, Ill. 60085.
Glass Water Bath: Use a constant-temperature glass water bath maintained at 40 ± 0.1°. A suitable bath is supplied as Catalog No. W3520 10 by Scientific Products.
Reagents and Solutions
Acetate Buffer (pH 4.5): Add 0.2 N sodium acetate, with continuous agitation, to 400 ml of 0.2 N acetic acid until the pH is 4.5 ± 0.05, as determined by a pH meter.
Locust Bean Gum: Use Powdered Type D-200 locust bean gum, or its equivalent, supplied by Meer Corp., 9500 Railroad Avenue, North Bergen, N.J. 07047. Since the substrate may vary from lot to lot, each lot should be tested in parallel with a previous lot known to be satisfactory. Variations of more than ± 5% viscosity in the average of a series of parallel tests indicate an unsuitable lot.
Substrate Solution: Place 12.5 ml of 0.2 N hydrochloric acid and 250 ml of warm water (72° to 75°) in the bowl of a power blender (Waring two-speed, or its equivalent, supplied as Catalog No. 58350-1 by Scientific Products), and set the blender on low speed. Slowly disperse 2.0 g of Locust Bean Gum, on a moisture-free basis, into the bowl, taking care not to splash out any of the liquid in the bowl. Wash down the sides of the bowl with warm water, using a rubber policeman, cover the bowl, and blend at high speed for 5 min. Quantitatively transfer the mixture to a 1,000-ml beaker, and cool to room temperature. Using a pH meter, adjust the mixture to pH 6.0 with 0.2 N sodium hydroxide. Quantitatively transfer to a 1,000-ml volumetric flask, dilute to volume with water, and mix. Filter the substrate through gauze before use.
Prepare a solution of the sample in water so that 1 ml of the final dilution will produce a change in relative fluidity between 0.18 and 0.22 in 5 min under the conditions specified in the Procedure below. Weigh the enzyme preparation, quantitatively transfer it to a glass mortar, and triturate with water. Quantitatively transfer the mixture to an appropriately sized volumetric flask, dilute to volume with water, and mix. Filter through Whatman No. 1 filter paper, or equivalent, before use.
Scrupulously clean the Viscometer by drawing a large volume of detergent solution, followed by water, through the instrument, and place the viscometer, previously calibrated, in the Glass Water Bath in an exactly vertical position. Pipet 20.0 ml of Substrate Solution and 4.0 ml of Acetate Buffer into a 50-ml Erlenmeyer flask, allowing at least two flasks for each enzyme sample and one flask for a substrate blank. Stopper the flasks, and equilibrate them in the water bath for 15 min. At zero time, pipet 1.0 ml of the Sample Preparation into the equilibrated substrate, start timing with a stopwatch (No. 1), and mix thoroughly. Immediately pipet 10.0 ml of this mixture into the wide arm of the Viscometer. After about 2 min, draw the reaction mixture above the upper mark into the driving fluid head by applying suction with a rubber tube connected to the narrow arm of the instrument. Measure the efflux time by allowing the reaction mixture to flow freely down past the upper mark. As the meniscus falls past the upper mark, start the second stopwatch (No. 2), and at the same time record the reaction time (Tr), in min, from stopwatch No. 1. As the meniscus of the reaction mixture falls past the lower mark, record the time (TT), in sec, from stopwatch No. 2. Immediately re-draw the reaction mixture above the upper mark and into the driving fluid head. As the meniscus falls freely past the upper mark, restart stopwatch No. 2, and at the same time record the reaction time (Tr), in min, from stopwatch No. 1. As the meniscus falls past the lower mark, record the time (TT), in sec, from stopwatch No. 2. Repeat the latter operation, beginning with "Immediately re-draw the reaction mixture ..." until a total of four determinations are obtained over a reaction time (TR) of not more than 15 min.
Prepare a substrate blank by pipetting 1.0 ml of water into a mixture of 20.0 ml of Substrate Solution and 4.0 ml of Acetate Buffer, and then immediately pipet 10.0 ml of this mixture into the wide arm of the Viscometer. Determine the time (TS), in sec, required for the meniscus to fall between the two marks. Use an average of five determinations as TS.
Prepare a water blank by pipetting 10.0 ml of water, previously equilibrated to 40 ±0.1°, into the wide arm of the Viscometer. Determine the time (Tw), in sec, required for the meniscus to fall between the two marks. Use an average of five determinations as Tw.
One hemicellulase unit (HCU) is that activity that will produce a relative fluidity change of 1 over a period of 5 min in a locust bean gum substrate under the conditions specified. Calculate the relative fluidities (FR) and T values (see definition below) for each of the four efflux times (Tt) and reaction times (Tr) as follows:
Fr is the relative fluidity for each reaction time;
TS is the average efflux time for the substrate blank, in sec;
Tw is the average efflux time for the water blank, in sec;
TT is the efflux time of the sample reaction mixture, in sec;
TR is the elapsed time from zero time, in min, i.e., the time from addition of the enzyme solution to the buffered substrate, until the beginning of the measurement of the efflux time
TN is the reaction time (TR), in min, plus one half of the efflux time (TT) converted to min.
Plot the four relative fluidities (Fr) as the ordinate against the four reaction times (Tn) as the abscissa. A straight line should be obtained. The slope of the line corresponds to the relative fluidity change per min and is proportional to the enzyme concentration. The slope of the best line through a series of experimental points is a better criterion of enzyme activity than is a single relative fluidity value. From the curve determine the FR values at 10 and 5 min. They should have a difference in fluidity of not more than 0.22 and not less than 0.18. Calculate the activity of the enzyme sample as follows:
FR10 is the relative fluidity at 10 min reaction time;
FR5 is the relative fluidity at 5 min reaction time;
1,000 is mg per g; and
W is the weight, in mg, of the enzyme sample contained in the 1.0-ml aliquot of Sample Preparation added to the equilibrated substrate in the Procedure.
Invertase hydrolyses the non-reducing P-d-fructofuranoside residues of sucrose to yield invert sugar. The invert sugar released is then reacted with 3.5 dinitrosalicylic acid (DNS). The colour change produced is proportional to the amount of invert sugar released, which in turn is proportional to the invertase activity present in the sample. The absorbance is measured at 540 nm and converted into micromoles of reducing sugar produced using a standard curve. One invertase unit is the amount of enzyme which will produce 1 micromole of reducing sugar (expressed as invert sugar) per minute under the conditions specified in this procedure.
Spectrophotometer set at 540 nm
Water bath set at 30±1.0°
Boiling water bath
Ice water bath
Reagents and solutions
0.05 M Sodium acetate buffer, pH 4.7: Adjust the pH of 200 ml of 0.05 M sodium acetate (4.1 g of sodium acetate anhydrous in 1000 ml of water) to pH 4.7 ± 0.05 with 0.05M acetic acid (2.85 ml of glacial acid in 1000 ml of water).
0.3 M Sucrose: 5.13 g sucrose in 50.0 ml of water
20 mM Tris HCl buffer, pH 7.0: Dissolve 2.42 g of tris (hydroxymethyl) aminomethane in about 800 ml of water. Adjust pH to 7.0 using 5% hydrochloric acid (5 ml of conc. hydrochloric acid in 100.0 ml of water).
DNS solution: Weigh 300 g of potassium sodium tartrate tetrahydrate into a one litre conical flask. Add 16 g of sodium hydroxide and 500 ml of water and dissolve by heating gently. When the solution is clear, add slowly 10 g of 3,5-dinitrosalicylic acid (DNS). Keep covered to protect from light until the DNS is totally dissolved. Cool to room temperature and make up to 1 litre with water. Store in a tightly stoppered dark container. Protect from light and carbon dioxide.
Invert sugar standard (0.01M): Dry glucose to constant weight at 105° and dry fructose to constant weight at 70° under vacuum. Dissolve 0.9 g of glucose and 0.9 g of fructose in 1000 ml of 0.1% benzoic acid (1 g of benzoic acid in 1000 ml of water).
Prepare a series of test tubes, in duplicate, according to the table below. The standard curve must include at least four suitable standardsTube No. 1 2 3 4 5 6 Blank Invert sugar standard (ml) 0.1 0.3 0.5 0.8 1.0 1.2 0.0 Water (ml) 2.4 2.2 2.0 1.7 1.5 1.3 2.5 Acetate buffer (ml) 0.5 0.5 0.5 0.5 0.5 0.5 0.5 Content of invert sugar 1.0 3.0 5.0 8.0 10 12 0.0
Reaction and measurement
Mix and incubate for exactly 10 min at 30 ± 0.1°. Add 2.0 ml of DNS solution to each tube, cover tubes and place all tubes in a boiling water bath for exactly 10 min. Cool rapidly in an ice water bath and add 15 ml of water to each tube. Mix thoroughly. Measure the absorbance at 540 nm of each sample using the blank to zero the spectrophotometer. Plot the absorbance against content of invert sugar.
Accurately weigh about 1 g of the sample and dissolve in 10 ml of 20 mM Tris HCl buffer. For powder samples it may be necessary to use a magnetic stirrer for up to 10 min. Dilute the sample with 20 mM Tris HCl buffer to obtain a solution for which the measured absorbance will fall within the linear range of 0.14 and 0.30.
Into each of a series of 30 ml test tubes, pipette, in quadruplicate, 1.4 ml of water, 0.5 ml of acetate buffer and 0.1 ml of diluted enzyme. Equilibrate the tubes in a 30° water bath. Add 1 ml of 0.3 M sucrose solution to 3 of the 4 tubes. Use the fourth tube as an enzyme blank, adding 2 ml of DNS solution before adding 1.0 ml of 0.3 M sucrose solution. Prepare a reagent blank using 0.1ml of water in place of diluted enzyme. Continue as described under 'Reaction and measurement'. Read the respective contents of invert sugar from the standard curve.
CS is Content of invert sugar in sample solution (micromoles)
Cb is Content of invert sugar in enzyme blank solution (micromoles)
W is Weight of sample (g)
CS is Content of invert sugar in sample solution (micromoles)
Cb is Content of invert sugar in enzyme blank solution (micromoles)
W is Weight of sample (g)
S.G. is Specific gravity of sample (g/ml)
This procedure is designed to be applied to enzyme preparations derived from either animal or microbial sources.
The method is based on a visual flocculation endpoint.
Bottle-rotating apparatus: Use a suitable assembly, designed to rotate at a rate of 16 to 18 rpm, such as the Dries-Jacques Associates type model (Available from Dries-Jacques Associates, 1801 East North Avenue, Milwaukee, Wisconsin 53202, USA.) or equivalent
Sample bottles: Use 125-ml squat, round, wide-mouth bottles such as those available as Catalog No. 2-903 from Fisher Scientific Co. (Available from Fischer Scientific, 711 Forbes Av., Pittsburgh, PA 15219, USA.), or equivalent.
Substrate Solution: Dissolve 60 g of low-heat, non-fat dry milk (such as Peake Grade A (Available from Galloway West, Fond du Lac, Wisc. 54935, USA.)), or equivalent in 500 ml of a solution, adjusted to pH 6.3 if necessary, containing in each ml 2.05 mg of sodium acetate (NaC2H3O2) and 1.11 mg of calcium chloride (CaCl2).
Standard Preparation: Use a standard-strength rennet; bovine rennet; milk-clotting enzyme, microbial (E. parasitica); or milk-clotting enzyme, microbial (Mucor species) as appropriate for the preparation to be assayed. Such standards, which are available from commercial coagulant manufacturers, should be of known activity. Dilute the standard-strength material 1 to 200 with water, and mix. Equilibrate to 30° before use, and prepare no more than 2 h prior to use.
Prepare aqueous solutions or dilutions of the sample to produce a final concentration such that the clotting time, as determined in the Procedure below, will be within 1 min of that of the Standard Preparation. Prepare no more than 1 h prior to use.
Transfer 50.0 ml of the Substrate Solution into each of four 125-ml Sample Bottles. Place the bottles on the Bottle-rotating Apparatus, and suspend the apparatus in a water bath, maintained at 30° ± 0.5, so that the bottles are at an angle or approximately 20° to 30° to the horizontal. Immerse the bottles so that the water level in the bath is about equal to the substrate level in the bottles. Begin rotating the apparatus at 16-18 rpm, then add 1.0 ml of the Sample Preparation to each of the two bottles, and record the exact time of addition. Add 1.0 ml of the Standard Preparation to each of the other two bottles, recording the exact time. Observe the rotating bottles, and record the exact time of the first evidence of clotting (i.e. when fine granules or flecks adhere to the sides of the bottle). Variations in the response of different lots of the substrate may cause variations in clotting time; therefore, the test samples and standards should be measured simultaneously on the same substrate. Average the clotting time, in sec, of the duplicate samples, recording the time for the Standard Preparation as Ts and that for the Sample Preparation as Tv.
Calculate the activity of the enzyme preparation by the formula:
in which 100 is the activity assigned to the Standard Preparation, Ds is the dilution factor for the Standard Preparation, and Dv is the dilution factor for the Sample Preparation.
Note: The dilution factors should be expressed as fractions; e.g., a dilution of 1 to 200 should be expressed as 1/200.
This procedure is designed for the determination of protease activity at pH 7.
This assay is based on the enzymatic hydrolysis of the peptide bonds of a defined gelatin substrate at pH 7.0 and 40°. The corresponding reduction in substrate viscosity is determined with a calibrated viscometer. One Viscometric Protease Unit is defined as that activity which will produce a relative fluidity change of 0.01 per sec in a defined gelatine substrate under the conditions of the assay.
Calibrated viscometer: Size 100 Calibrated Cannon-Fenske Type Viscometer, or its equivalent, supplied as Catalog No. P2885-100.
Constant temperature glass water bath (40 ±0.1°): Constant temperature glass water bath, or its equivalent, supplied as Catalog No. W3520-10 (Available from Scientific Products, 1210 Waukegan Rd., McGaw Park, Ill., 60085, USA.).
Stopwatches: Stopwatch calibrated in 1/10 min for determining the reaction time (Tr) and stopwatch calibrated in 1/5 sec for determining the efflux time (Tt).
Reagents and Solutions
Disodium monohydrogen phosphate solution (1 N): Dissolve 47.32 g of anhydrous disodium phosphate in approximately 800 ml of distilled water in a beaker. Quantitatively transfer to a 1,000-ml volumetric flask and dilute to volume with distilled water.
Monosodium dihydrogen phosphate solution (1 N): Dissolve 40.00 g of anhydrous monosodium phosphate in approximately 800 ml of distilled water in a beaker. Quantitatively transfer to a 1,000-ml volumetric flask and dilute to volume with distilled water.
Phosphate buffer (pH 7.0): Using a standardized pH-meter, add disodium monohydrogen phosphate solution (1 N) with continuous agitation to 800 ml of monosodium dihydrogen phosphate solution (1 N) until the buffer is pH 7.0 ± 0.05.
Gelatine substrate (4.0% w/v): With continuous agitation, disperse 20.00 g (moisture-free basis) of gelatin in approximately 400 ml of distilled water in a 1,000-ml Erlenmeyer flask. The dispersion must be free of lumps. Swell the gelatin for 30 min at room temperature with occasional swirling. Place the gelatin solution on a 40 ± 0.1° waterbath. Swirl occasionally until the gelatin is completely solubilized with no particles appearing in solution. Cool to room temperature and quantitatively transfer to a 500-ml volumetric flask and dilute to volume with distilled water.
Enzyme Preparation: Prepare an enzyme solution so that 1 ml of the final dilution will produce a relative fluidity change between 0.18 and 0.22 in 5 min under the conditions of the assay. Weigh the enzyme and quantitatively transfer to a glass mortar. Triturate the enzyme with distilled water and quantitatively transfer to an appropriate volumetric flask. Dilute the volume with distilled water and filter the enzyme solution through Whatman No. 1 filter paper, or equivalent, prior to use.
Place the calibrated viscometer in the 40 ± 0.1 ° water bath in an exactly vertical position. Use only a clean viscometer. Cleaning is readily accomplished by drawing a large volume of detergent solution followed by distilled water through the viscometer. This can be accomplished by using an aspirator with a rubber tube connected to the narrow arm of the viscometer.
Pipet 20 ml of gelatin substrate and 3 ml of phosphate buffer into a 50-ml Erlenmeyer flask. Allow at least two flasks for each enzyme sample and one flask for a substrate blank. Stopper the flasks and equilibrate them in the water bath for 15 min. At zero time pipet 1 ml of the enzyme solution into the equilibrated substrate. Start the stopwatch calibrated in 0.1 min and mix solution thoroughly. Immediately pipet 10 ml of the reaction mixture into the wide arm of the viscometer.
After approximately 2 min apply suction with a rubber tube connected to the narrow arm of the viscometer drawing the reaction mixture above the upper mark into the driving fluid head. Measure the efflux time by allowing the reaction mixture to freely flow down past the upper mark. As the meniscus of the reaction mixture falls past the upper mark, start the other stopwatch. At the same time record the reaction time in min from the first stopwatch (Tr). As the meniscus of the reaction mixture falls past the lower mark, record the time in sec from the second stopwatch (Tt). Immediately redraw the reaction mixture above the upper mark and into the fluid driving head. As the meniscus of the reaction mixture falls freely past the upper mark, restart the second stopwatch. At the same time, record the reaction time in min from the first stopwatch (Tr). As the meniscus of the reaction mixture falls past the lower mark, record the time in sec, from the second stopwatch (Tt).
Repeat from redrawing the reaction mixture above the upper mark, until a total of 4 determinations is obtained over a reaction time (Tr) of not more than 15 min.
Prepare a substrate blank by pipetting 1 ml of distilled water into 24 ml of buffered substrate. Pipet 10 ml of the reaction mixture into the wide arm of the viscometer. Determine the time (Ts) in sec required for the meniscus to fall between the two marks. Use an average of 5 determinations for Ts.
Prepare a water blank by pipetting 10 ml of equilibrated distilled water into the wide arm of the viscometer. Determine the time (Tw) in sec required for the meniscus to fall between the two marks. Use an average of 5 determinations for Tw.
One Viscometric Protease Unit (VPU) is that activity which will produce a relative fluidity change of 0.01 per sec in a defined gelatin substrate under the conditions of the assay.
Calculate the relative fluidities (Fr) and the times (Tn) for each of the four (4) efflux times (Tt) and reaction times (Tr) as follows:
Fr is relative fluidity for each reaction time,
Ts is average efflux time for the substrate blank in sec,
Tw is average efflux time for the water blank in sec,
Tt is efflux time of the reaction mixture in sec,
Tr is elapsed time in min from zero time, i.e. the time from addition of the enzyme solution to the buffered substrate, until the beginning of the measurement of efflux time (Tt),
Tn is reaction time in min (Tr), plus one-half of the efflux time (Tt) converted to min.
Plot the four relative fluidities (Fr) as the ordinate against the four reaction times (Tr) as the abscissa. A straight line should be obtained. The slope of this line corresponds to the relative fluidity change per min and is proportional to the enzyme concentration. The slope of the best line through a series of experimental points is a better criterion of enzyme activity than is a single relative fluidity value. From the graph determine the Fr values at 10 and 5 min. They should have a difference in fluidity of not more than 0.22 nor less than 0.18. Calculate the activity of the enzyme unknown as follows:
Fr5 is relative fluidity at five (5) min of reaction time
Fr10 is relative fluidity at ten (10) min of reaction time
300 is time of relative fluidity change in sec from Fr10 to Fr5
1,000 is milligrams per g
W is weight in milligrams of enzyme added to the reaction mixture in a one (1) ml aliquot of enzyme solution
0.01 is change in relative fluidity per sec per VPU.
This procedure is designed for the determination of protease activity, expressed as PC units.
The assay is based on a 30-min proteolytic hydrolysis of casein at 37° and pH 7.0. Unhydrolyzed casein is removed by filtration, and the solubilized casein is determined spectrophotometrically.
Reagents and Solutions
Casein: Use Hammarsten-grade casein (Available from Nutritional Biochemical Corp., 21010 Miles Ave., Cleveland, Ohio 44128, USA.) or equivalent.
Tris buffer (pH 7.0): Dissolve 12.1 g of enzyme-grade (or equivalent) tris(hydroxymethyl)aminomethane in 800 ml of water, and titrate with 1 N hydrochloric acid to pH 7.0. Transfer into a 1,000-ml volumetric flask, dilute to volume with water, and mix.
TCA solution: Dissolve 18 g of trichloroacetic acid and 19 g of sodium acetate trihydrate in 800 ml of water in a 1,000-ml volumetric flask, add 20 ml of glacial acetic acid, dilute to volume with water, and mix.
Substrate solution: Dissolve 6.05 g of tris(hydroxymethyl)aminomethane (enzyme grade) in 500 ml of water, add 8 ml of 1 N hydrochloric acid, and mix. Dissolve 7 g of Casein in this solution, and heat for 30 min in a boiling water bath, stirring occasionally. Cool to room temperature, and adjust to pH 7.0 with 0.2 N hydrochloric acid, adding the acid slowly, with vigorous stirring, to prevent precipitation of the casein. Transfer the mixture into a 1,000-ml volumetric flask, dilute to volume with water, and mix.
Using Tris Buffer, prepare a solution of the sample enzyme preparation so that 2 ml of the final dilution will contain between 10 and 44 PC units.
Pipet 10.0 ml of the Substrate Solution into each of a series of 25 x 150-mm test tubes, allowing one tube for each enzyme test, one tube for each enzyme blank, and one tube for a substrate blank. Equilibrate the tubes for 15 min in a water bath maintained at 37 ± 0.1°. At zero time, rapidly pipet 2.0 ml of the Sample Preparation into the equilibrated substrate, starting the stopwatch at zero time. Mix, and replace the tubes in the water bath. Add 2 ml of Tris Buffer (instead of the Sample Preparation) to the substrate blank.
After exactly 30 min, add 10 ml of TCA Solution to each enzyme incubation and to the substrate blank to stop the reaction. Caution: Do not use mouth suction for the TCA Solution. Heat the tubes in the water bath for an additional 30 min to allow the protein to coagulate completely.
At the end of the second heating period, shake each tube vigorously, and filter through 11-cm Whatman No. 42, or equivalent, filter paper, discarding the first 3 ml of filtrate.
Note: The filtrate must be perfectly clear.
Determine the absorbance of each sample filtrate in a 1 -cm cell, at 275 nm, with a suitable spectrophotometer, using the filtrate from the substrate blank to set the instrument at zero. Correct each reading by subtracting the appropriate enzyme blank reading, and record the value so obtained in Au.
Transfer 100 mg of L-tyrosine, chromatographic-grade (Available from Calbiochem, La Jolla, Calif. 92037, USA.) or equivalent, previously dried to constant weight, to a 1,000-ml volumetric flask. Dissolve in 60 ml of 0.1 N hydrochloric acid.
When completely dissolved, dilute the solution to volume with water, and mix thoroughly. This solution contains 100 μg of tyrosine in 1.0 ml. Prepare three more dilutions from this stock solution to contain 75.0, 50.0 and 25.0 μg of tyrosine per ml. Determine the absorbance of the four solutions at 275 nm in a 1-cm cell with a suitable spectrophotometer versus 0.006 N hydrochloric acid. Prepare a plot of absorbance versus tyrosine concentration.
One bacterial protease unit (PC) is defined as that quantity of enzyme that produces the equivalent of 1.5 μg per ml of L-tyrosine per min under the conditions of the assay.
From the Standard Curve, and by interpolation, determine the absorbance of a solution having a tyrosine concentration of 60 μg per ml. A figure close to 0.0115 should be obtained. Divide the interpolated value by 40 to obtain the absorbance equivalent to that of a solution having a tyrosine concentration of 1.5 μg per ml and record the value thus derived as As.
Calculate the activity of the sample enzyme preparation by the formula:
22 is the final volume, in ml of the reaction mixture,
30 is the time of the reaction, in min, and
W is the weight of the original sample taken, in g.
This procedure is for the determination of the proteolytic activity, expressed as haemoglobin units on the tyrosine basis (HUT), of preparations derived from Aspergillus oryzae, var., and Aspergillus niger, var., and it may be used to determine the activity of other proteases at pH. 4.7
The test is based on the 30-min enzymatic hydrolysis of a haemoglobin substrate at pH 4.7 and 40°. Unhydrolyzed substrate is precipitated with trichloroacetic acid and removed by filtration. The quantity of solubilized haemoglobin in the filtrate is determined spectrophotometrically.
Reagents and Solutions
Haemoglobin: Use Haemoglobin Substrate Powder (Sigma Chemicals Co., Catalog No. H 262) or a similar high-grade material that is completely soluble in water.
Acetate Buffer Solution: Dissolve 136 g of sodium acetate (NaC2H3O2.3H2O) in sufficient water to make 500 ml. Mix 25.0 ml of this solution with 50.0 ml of 1 M acetic acid, dilute to 1,000 ml with water, and mix. The pH of this solution should be 4.7 ± 0.02.
Substrate Solution: Transfer 4.0 g of the Haemoglobin into a 250-ml beaker, add 100 ml of water, and stir for 10 min to dissolve. Immerse the electrodes of a pH meter in the solution, and adjust the pH to 1.7, stirring continuously, by the addition of 0.3 N hydrochloric acid. After 10 min, adjust the pH to 4.7 by the addition of 0.5 M sodium acetate. Transfer the solution into a 200-ml volumetric flask, dilute to volume with water, and mix. This solution is stable for about 5 days when refrigerated.
Trichloroacetic Acid Solution: Dissolve 14.0 g of trichloroacetic acid in about 75 ml of water. Transfer the solution to a 100-ml volumetric flask, dilute to volume with water, and mix thoroughly.
Dissolve an amount of the sample in the Acetate Buffer Solution to produce a solution containing, in each ml, between 9 and 22 HUT. (Such a concentration will produce an absorbance reading, in the procedure below, within the preferred range of 0.2 to 0.5.)
Pipet 10.0 ml of the Substrate Solution into each of a series of 25 x 150-mm test tubes: one for each enzyme test and one for the substrate blank. Heat the tubes in a water bath at 40° for about 5 min. To each tube except the substrate blank add 2.0 ml of the Sample Preparation, and begin timing the reaction at the moment the solution is added; add 2.0 ml of the Acetate Buffer Solution to the substrate blank tube. Close the tubes with No. 4 rubber stoppers, and tap each tube gently for 30 sec against the palm of the hand to mix. Heat each tube in a water bath at 40° for exactly 30 min, and then pipet rapidly 10.0 ml of the Trichloroacetic Acid Solution into each tube. (Caution: Do not use mouth suction on the pipet.) Shake each tube vigorously against the stopper for about 40 sec, and then allow to cool to room temperature for 1 h, shaking each tube against the stopper at 10 to 12 min intervals during this period. Prepare enzyme blanks as follows: heat, in separate tubes, 10.0 ml of the Trichloroacetic Acid Solution in 10.0 ml of the Substrate Solution, shake well for 40 sec, and to this mixture add 2.0 ml of the preheated Sample Preparation. Shake again, and cool at room temperature for 1 h, shaking at 10 to 12 min intervals.
At the end of 1 h, shake each tube vigorously, and filter through 11-cm Whatman No. 42, or equivalent, filter paper, re-filtering the first half of the filtrate through the same paper. Determine the absorbance of each filtrate in a 1-cm cell, at 275 nm, with a suitable spectrophotometer, using the filtrate from the substrate blank to set the instrument to zero. Correct each reading by subtracting the appropriate enzyme blank reading, and record the value so obtained as AU.
Note: If a corrected absorbance reading between 0.2 and 0.5 is not obtained, repeat the test using more or less of the enzyme preparation as necessary.
Transfer 100.0 mg of L-tyrosine, chromatographic-grade or equivalent (Aldrich Chemical Co.), previously dried to constant weight, to a 1,000-ml volumetric flask. Dissolve in 60 ml of 0.1 N hydrochloric acid. When completely dissolved, dilute the solution to volume with water, and mix thoroughly. This solution contains 100 μg of tyrosine in 1.0 ml. Prepare three more dilutions from this stock solution to contain 75.0, 50.0, and 25.0 μg of tyrosine per ml. Determine the absorbance of the four solutions at 275 nm in a 1-cm cell on a suitable spectrophotometer versus 0.006 N hydrochloric acid. Prepare a plot of absorbance versus tyrosine concentration. Determine the slope of the curve in terms of absorbance per μg of tyrosine. Multiply this value by 1.10, and record it as As. A value of approximately 0.0084 should be obtained.
One HUT unit of proteolytic (protease) activity is defined as that amount of enzyme that produces, in 1 min under the specified conditions, a hydrolysate whose absorbance at 275 nm is the same as that of a solution containing 1.10 μg per ml of tyrosine in 0.006 N hydrochloric acid.
Calculate the HUT per g of the original enzyme preparation by the formula,
22 is the final volume of the test solution,
30 is the reaction time in min, and
W is the weight of the original sample taken, in g.
Note: The value for As, under carefully controlled and standardized conditions, is 0.0084. This value may be used for routine work in lieu of the value obtained from the standard curve, but the exact value calculated from the standard curve should be used for more accurate results and in cases of doubt.
This procedure is for the determination of proteolytic activity, expressed in spectrophotometric acid protease units (SAPU), of preparations derived from Aspergillus niger, var., and Aspergillus oryzae, var.
The test is based on a 30-min enzymatic hydrolysis of a Hammarsten Casein Substrate at pH 3.0 and 37°. Unhydrolyzed substrate is precipitated with trichloroacetic acid and removed by filtration. The quantity of solubilized casein in the filtrate is determined spectrophoto-metrically.
Reagents and Solutions
Casein: Use Hammarsten-grade casein, available from Nutritional Biochemical Corp., 21010 Miles Avenue, Cleveland, Ohio 44128.
Glycine-Hydrochloric Acid Buffer (0.05 M): Dissolve 3.75 g of glycine in about 800 ml of water. Add 1 N hydrochloric acid until the solution is pH 3.0, determined with a pH meter. Quantitatively transfer the solution to a 1000-ml volumetric flask, dilute to volume with water, and mix.
TCA Solution: Dissolve 18.0 g of trichloroacetic acid and 11.45 g of anhydrous sodium acetate in about 800 ml of water, and add 21.0 ml of glacial acetic acid. Quantitatively transfer the solution to a 1000-ml volumetric flask, dilute to volume with water, and mix.
Substrate Solution: Pipet 8 ml of 1 N hydrochloric acid into about 500 ml of water, and disperse 7.0 g (moisture-free basis) of Casein into this solution, using continuous agitation. Heat for 30 min in a boiling water bath, stirring occasionally, and cool to room temperature. Dissolve 3.75 g of glycine in the solution, and adjust to pH 3.0 with 0.1 N hydrochloric acid, using a pH meter. Quantitatively transfer the solution to a 1000-ml volumetric flask, dilute to volume with water, and mix.
Using Glycine-Hydrochloric Acid Buffer: Prepare a solution of the sample enzyme preparation so that 2 ml of the final dilution will give a corrected absorbance of enzyme incubation filtrate at 275 nm ( A, as defined in the Procedure) between 0.200 and 0.500. Weigh the enzyme preparation, quantitatively transfer it to a glass mortar, and triturate with Glycine-Hydrochloric Acid Buffer. Quantitatively transfer the mixture to an appropriately sized volumetric flask, dilute to volume with Glycine-Hydrochloric Acid Buffer, and mix.
Pipet 10.0 ml of Substrate Solution into each of a series of 25 x 150 mm test tubes, allowing at least two tubes for each sample, one for each enzyme blank, and one for a substrate blank. Stopper the tubes, and equilibrate them for 15 min in a water bath maintained at 37° ± 0.1°.
At zero time, start the stopwatch, and rapidly pipet 2.0 ml of the Sample Preparation into the equilibrated substrate. Mix by swirling, and replace the tubes in the water bath. (Note: The tubes must be stoppered during incubation). Add 2 ml of Glycine-Hydrochloric Acid Buffer (instead of the Sample Preparation) to the substrate blank. After exactly 30 min, add 10 ml of TCA Solution to each enzyme incubation and to the substrate blank to stop the reaction. (Caution: Do not use mouth suction for the TCA Solution). In the following order, prepare an enzyme blank containing 10 ml of Substrate Solution, 10 ml of TCA Solution, and 2 ml of the Sample Preparation. Heat all tubes in the water bath for 30 min, allowing the precipitated protein to coagulate completely.
At the end of the second heating period, cool the tubes in an ice bath for 5 min, and filter through Whatman No. 42 filter paper, or equivalent. The filtrates must be perfectly clear. Determine the absorbance of each filtrate in a 1-cm cell at 275 nm with a suitable spectrophotometer, against the substrate blank. Correct each absorbance by subtracting the absorbance of the respective enzyme blank.
Transfer 181.2 mg of L-tyrosine, chromatographic-grade or equivalent (Calbiochem, La Jolla, Calif. 92037), previously dried to constant weight, to a 1,000-ml volumetric flask. Dissolve in 60 ml of 0.1 N hydrochloric acid. When completely dissolved, dilute the solution to volume with water, and mix thoroughly. This solution contains 1.00 μmol of tyrosine in 1.0 ml. Prepare dilutions from this stock solution to contain 0.10, 0.20, 0.30, 0.40, and 0.50 μmol per ml. Determine the absorbance of each dilution in 1-cm cell at 275 nm, against a water blank. Prepare a plot of absorbance versus μmol of tyrosine per ml. A straight line must be obtained. Determine the slope and intercept for use in the Calculation below. A value close to 1.38 should be obtained. The slope and intercept may be calculated by the least squares method as follows:
in which n is the number of points on the standard curve, M is the μmol of tyrosine per ml for each point on the standard curve, and A is the absorbance of the sample.
One spectrophotometric acid protease unit is that activity that will liberate 1 μmol of tyrosine per min under the conditions specified. The activity is expressed as follows:
A is the corrected absorbance of the enzyme incubation filtrate;
I is the intercept of the Standard Curve;
22 is the final volume of the incubation mixture, in ml;
S is the slope of Standard Curve;
30 is the incubation time, in min; and
W is the weight, in g, of the enzyme sample contained in the 2.0-ml aliquot of Sample Preparation added to the incubation mixture in the Procedure.
This procedure is designed for the determination of the proteolytic activity of papain, ficin and bromelain.
The assay is based on a 60 min proteolytic hydrolysis of a casein substrate at pH 6.0 and 40°. Unhydrolyzed substrate is precipitated with trichloroacetic acid and removed by filtration; solubilized casein is then measured spectrophotometrically.
Reagents and Solution
Sodium phosphate solution (0.05 M): Transfer 7.1 g of anhydrous dibasic sodium phosphate into a 1000-ml volumetric flask, dissolve in about 500 ml of water, dilute to volume with water, and mix. Add 1 drop of toluene as preservative.
Citric acid solution (0.05 M): Transfer 10.5 g of citric acid monohydrate into a 1,000-ml volumetric flask, dissolve in about 500 ml of water, dilute to volume with water, and mix. Add 1 drop of toluene as preservative.
Phosphate-cysteine-EDTA buffer solution: Dissolve 7.1 g of anhydrous dibasic sodium phosphate in about 800 ml of water, and then dissolve in this solution 14.0 g of disodium EDTA dihydrate and 6.1 g of cysteine hydrochloride monohydrate. Adjust to pH 6.0 ±0.1 with 1 N hydrochloric acid or 1 N sodium hydroxide, then transfer into a 1,000-ml volumetric flask, dilute to volume with water, and mix.
Trichloroacetic acid solution: Dissolve 30 g of trichloroacetic acid in 100 ml of water.
Casein substrate solution: Disperse 1 g (moisture-free basis) of Hammarsten casein or equivalent in 50 ml of Sodium Phosphate Solution, and heat for 30 min in a boiling water bath, with occasional shaking. Cool to room temperature, and with rapid and continuous shaking, adjust to pH 6.0 ± 0.1 by the addition of citric acid solution.
Note: Rapid and continuous agitation during the addition prevents casein precipitation.
Quantitatively transfer the mixture into a 100-ml volumetric flask, dilute to volume with water, and mix.
Stock standard solution: Transfer 100.0 mg of USP Papain Reference Standard into a 100-ml volumetric flask, dissolve and dilute to volume with Phosphate-Cysteine-EDTA Buffer Solution, and mix.
Diluted standard solutions: Pipet 2, 3, 4, 5, 6 and 7 ml of Stock Standard Solution into a series of 100-ml volumetric flasks, dilute each to volume with Phosphate-Cysteine-EDTA Buffer Solution, and mix by inversion.
Test solution: Prepare a solution from the enzyme preparation so that 2 ml of the final dilution will give an absorbance in the Procedure between 0.2 and 0.5. Weigh the sample accurately, transfer it quantitatively to a glass mortar, and triturate with Phosphate-Cysteine-EDTA Buffer Solution. Transfer the mixture quantitatively into a volumetric flask of appropriate size, dilute to volume with Phosphate-Cysteine-EDTA Buffer Solution, and mix.
Pipet 5 ml of Casein Substrate Solution into each of a series of 25 x 150 mm test tubes, allowing three tubes for the enzyme unknown, six for a papain standard curve, and nine for enzyme blanks. Equilibrate the tubes for 15 min in a water bath maintained at 40 ± 0.1°. At zero time, rapidly pipet 2 ml of each of the Diluted Standard Solutions, and 2-ml portions of the Test Solution, into the equilibrated substrate, starting the stopwatch at zero time. Mix each by swirling, stopper and place the tubes back in the water bath. After 60.0 min. add 3 ml of Trichloroacetic Acid Solution to each tube. (Caution: Do not use mouth suction). Mix each tube immediately by swirling.
Prepare enzyme blanks containing 5.0 ml of Casein Substrate Solution, 3.0 ml of Trichloroacetic Acid Solution, and 2.0 ml of one of the appropriate Diluted Standard Solutions or the Test Solution.
Return all tubes to the water bath, and heat for 30.0 min allowing the precipitated protein to coagulate completely. Filter each mixture through Whatman No. 42, or equivalent, filter paper, discarding the first 3 ml of filtrate. The subsequent filtrate must be perfectly clear. Determine the absorbance of each filtrate in a 1-cm cell at 280 nm with a suitable spectrophotometer, against its respective blank.
One papain unit (PU) is defined in this assay as that quantity of enzyme that liberates the equivalent of 1 μg of tyrosine per h, under the conditions of the assay. Prepare a standard curve by plotting the absorbances of filtrates from the Diluted Standard Solutions against the corresponding enzyme concentrations, in mg/ml. By interpolation from the standard curve, obtain the equivalent concentration of the filtrate from the Test Solution. Calculate the activity of the enzyme preparation taken for analysis as follows:
A is the activity of USP Papain Reference Standard, in PU per mg,
C is the concentration, in mg per ml, of Reference Standard from the standard curve, equivalent to the enzyme unknown,
10 is the total volume, in ml, of the final incubation mixture, and
W is the weight, in mg, of original enzyme preparation in the 2-ml aliquot of Test Solution added to the incubation mixture.
This procedure is designed for the determination of the pullulanase activity. (Pullulan is produced by deep fermentation of food grade hydrolysed starch by Aureobasidium pullulans.)
Pullulanase hydrolyses a 1 -6 glycosidic links in branched poly-saccharides and breaks down pullulan to yield maltotriose only. After the reaction is complete, the reducing sugars formed are estimated by the reaction with dinitrosalicylic acid. Thus one unit of Pullulanase is the activity which will produce reducing sugars equivalent to 1 mg of anhydrous maltose after one min, under the conditions of the assay. (Maltose is used as the standard of comparison, because maltotriose is expensive and not of the highest purity. The method measures the reducing end groups of maltotriose and higher sugars using maltose as a reference.)
Pullulan solution: Add 1 g of standard pullulan to 70 ml of distilled water. Boil for 5 min, cool and add 10 ml of molar acetate buffer pH 5,0 then dilute to 100 ml. Filter if necessary. This solution can be stored up to two weeks in a refrigerator.
3,5-Dinitrosalicylic acid reagent (DNS): Add 1 g of DNS to 16 ml of 10% w/v sodium hydroxide solution. Add 30 g of Rochelle salt (potassium sodium tartrate tetrahydrate) and 50 ml of distilled water and then warm until dissolved. Dilute this solution to 100 ml. It may be kept for 5 days at 5°.
Pipet 1 ml of substrate pullulan solution into a 17 x 1.5 cm test tube and place in a water bath at 50° for 5 min. Add 1 ml of enzyme solution and allow reaction to proceed for exactly 10 min. Stop reaction by adding 2 ml of DNS reagent.
Prepare a blank by adding 2 ml of DNS reagent to substrate before the enzyme is added.
Place the two tubes in a boiling water bath for exactly 5 min and then cool rapidly and add 10 ml of distilled water. Mix solutions well by shaking.
Measure the absorbance of the test solution against the blank using 2-cm glass cells at a wavelength of 540 nm.
The reducing value measured is compared with that of a standard maltose solution. A standard maltose graph is not necessary as, for accurate results, the absorbance produced in the test should be between 0.2 - 0.5. As 1 mg of maltose will give an absorbance of 0.82, for the purpose of the calculation the definition is adjusted to read "0.4 units of activity will produce 0.4 mg of anhydrous maltose equivalent...". Therefore a standard maltose solution is made so that 1 ml contains 0.4 mg of anhydrous maltose and this solution is used for the test in place of the 1 ml of enzyme solution. The absorbance is read as before and should be 0.325. This reading is so constant that, if any difference is found, the wavelength calibration on the spectrophotometer should be checked. This is critical since very small errors in the wavelength can have large effects on the absorbance.
For an unknown sample several dilutions are made up and tested. A graph of absorbance against enzyme concentration is plotted (see ) and the concentration of enzyme which will give an absorbance of 0.325 is found. Then, by definition this concentration of enzyme contains 0.4 Pullulanase units. Thus the activity of Pullulanase preparation is found by:
Pullulanase activity/mg = 1,000 / mg of enzyme in test x (0.4 / 10)Enzyme concentration mg in test Absorbance 0.002% 0.02 0.170 0.003% 0.03 0.245 0.004% 0.04 0.325 0.005% 0.05 0.390 0.006% 0.06 0.465 0.008% 0.08 0.595 0.010% 0.10 0.720
From the graph, an absorbance of 0.325 is given by 0.004% w/v enzyme solution. Therefore the activity equals
(1,000 / 0.04) x (0.4 /10) = 1,000 units per g
. Pullulanase Assay for 50 mg/kg solution
This can now be used to construct a standard graph of absorbance against Pullulanase units for a fixed enzyme concentration. This graph can be used for all further samples. If the 0.005% solution is taken as standard, then its absorbance of 0.39 must give 1,000 units/g (as above). From this, a graph can be constructed for any sample at a concentration of 0.005%Enzyme concentration Absorbance Units/g 0.002% 0.170 400 0.003% 0.245 600 0.004% 0.325 800 0.005% 0.390 1,000 0.006% 0.465 1,200 0.008% 0.595 1,600 0.010% 0.720 2,000
A graph is drawn on absorbance against units/g for a 0.005% enzyme solution.
For an enzyme made up to concentration of 0.0025%, 0.005% and 0.0075%, the absorbances would be:Concentration Absorbance Units g 0.0025% 0,200 480 0.005% 0.375 950 0.0075% 0.553 1,470
Thus the activity is found as follows:
Average = 953 units/g
Xylanase samples are incubated with a remazol-stained wheat arabinoxylan substrate. Unconverted substrate is precipitated with ethanol. The intensity of blue colouring of the supernatant due to unprecipitated remazol-stained substrate degradation products is proportional to the endoxylanase activity. Xylanase activity is measured relative to an enzyme standard and calculated in Farvet Xylanase Units (FXU). The colour profile may vary from enzyme to enzyme.
Thermostatic water bath
10-mlplastic test tubes
Reagents and substrates
Phosphate buffer stock solution, 1.0 M: Dissolve 1210 g sodium dihydrogen phosphate monohydrate and 218.9 g disodium hydrogen phosphate dihydrate in demineralised water. Add 40 ml 4 N NaOH and make up to 10 l with water.
Phosphate buffer, 0.1 M, pH 6.00 ± 0.05: Take 1000 ml phosphate buffer stock solution and adjust the pH to 6.0 ± 0.05 using either 4 N NaOH or 2 N HCl. Make up to 10 L with demineralised water.
Azo-wheat arabinoxylan substrate (Megazyme Ltd., Bray, Ireland) 0.5% w/v pH 6.00 ± 0.05: Weigh 0.500 g Azo-wheat arabinoxylan into a 150-ml beaker. Add about 90 ml of 0.1 M phosphate buffer, and heat to approximately 50°, while stirring. Continue stirring at 50° for a further 20 min. Cool the substrate solution and adjust to pH 6.00 ± 0.05 before transferring to a 100-ml graduated flask. Fill to the mark with phosphate buffer.
Stop reagent: Pipette 6.65 ml 2 N HCl into a 100 ml graduated flask. Fill up to the mark with 99.9% ethanol.
Standard solutions: Reference enzyme stock and working solutions: Accurately weigh approximately 1g FXU standard into a suitable graduated flask, add 0.1 M phosphate buffer to volume and dissolve the standard by stirring for approximately 15 min. Use this stock solution to prepare at least 6 FXU standard working solutions to give a range of activities between 0.2 and 1.4 FXU/ml for the construction of the standard curve. Prepare additional samples of known activity for inclusion at the beginning and the end of each analysis series or at least every 20 samples.
Samples: Samples are diluted on the basis of their anticipated activity so that the activity of the final dilution is between. 0.4-1.4 FXU/ml. Results outside the working range may be used to assess the activity of the sample for the next run. Weigh dry or liquid samples directly into the flask. Granulated products may take a considerable time to dissolve.
Pipette 0.100 ml working standard or sample solution into 10-ml test tubes, add 0.900 ml of the substrate and mix. Incubate the tubes in a 50° water bath for 30 min. Add 5 ml stop reagent and mix for 10-20 sec.
Leave the tubes to stand at room temperature, for 15-60 min and centrifuge at 4000 rpm for 15 min. Measure the absorbance of the supernatant at 585 nm within 20 min.
Use the measurements for the enzyme standards to plot a standard curve. The data may be fitted to a third order polynomial. Determine the corresponding enzyme activity values from the standard curve for the samples. The activity of each sample is calculated as follows:
C is enzyme activity read from the standard curve (FXU/ml)
F is volume of sample (ml)
D is further dilution of sample (e.g. second or third dilution)
W is weight of sample (g)
Xylanase samples are incubated with azurine-crosslinked wheat arabinoxylan substrate. Xylanase hydrolyses the substrate to water-soluble fragments with the concomitant change in colour. The reaction is terminated after a designated time and the optical density (OD) of the reaction mixture is measured at 590 nm (OD590). Xylanase activity is calculated based on the rate of release of the azurine dye. One xylanase unit (XU) is defined as the amount of enzyme that increases the OD590 at a rate of one OD per 10 minutes under standard conditions (pH 5.00; 40°).
Thermostatic water bath
Whatman No. 1 filter paper
Test tubes (15 ml)
Citric acid monohydrate
Disodium hydrogen phosphate dihydrate
TRIS (tris (hydroxyl methyl) amino methane)
Substrate (azurine-crosslinked wheat arabinoxylan: Xylazyme tablets from Megazyme, Ireland)
Note: a new batch of the substrate should be compared with a previous batch by analyzing the same enzyme preparation using both substrates. If a difference in enzymatic activity is noted, an appropriate correction factor should be calculated and applied to the results obtained with the new batch of the substrate.
Reaction buffer (McIlvaine buffer, pH 5.00): Dissolve 10.19 g of citric acid monohydrate and 18.33 g disodium hydrogen phosphate dihydrate in 850 ml distilled water in a 1000-ml volumetric flask. Adjust the pH to 5.00 using either 0.1 M citric acid monohydrate or 0.2 M disodium hydrogen phosphate dihydrate. Add water to 1000 ml. The buffer can be stored for up to 6 months at 2-5°.
Stop solution (2% w/v TRIS, pH 12.0): Dissolve 20 g of TRIS in 850 ml distilled water in a 1000-ml volumetric flask. Adjust the pH to 12.0 with 5 M NaOH. The solution can be stored for up to six months at 2-5°.
Test sample solutions: Accurately weigh a quantity of the enzyme preparation that would give an OD increase within the range of 0.3 - 1.2 in a 100 ml volumetric flask. Add 60 ml of the reaction buffer. Stir the solution using a magnetic stirrer for 10 minutes. Remove the magnet and add the reaction buffer to volume. Transfer the enzyme solution to a glass beaker and let it stand for 5 minutes or until the precipitate settles. Use clear solution for analysis.
Blank: Pre-heat 1.0 ml reaction buffer at 40.0° for 5 min. Add one Xylazyme tablet. After exactly 10 min at 40.0°, add 10.0 ml stop solution and filter the sample through Whatman No.1 filter.
Prepare 3 test tubes for each test sample. Pipette 1 ml of the reaction buffer to each tube and add 50, 75, and 100 microliters of the test sample solution.
- Pre-heat all test sample solutions at 40.0° for 5 min.
- Add one Xylazyme tablet to each tube. Do not stir.
- After 10 minutes (±1 sec), terminate the reaction by adding 10 ml stop solution.
- Filter all solutions through Whatman No. 1 filter paper.
- Measure OD of each test sample solution against the blank at 590 nm.
Perform linear regression on OD590 as a function of test sample volumes (in ml) used in the analysis. Calculate the activity of the enzyme preparation in xylanase units (XU) per gram (g) using the following equation:
S is the slope obtained from linear regression of the OD590 as a function of sample volume in ml
V is the volume of the volumetric flask used to prepare the test sample solution in ml (multiplied by further dilutions, if applicable)
W is the weight of the enzyme preparation in g
Acid value is defined as the number of mg of potassium hydroxide required to neutralize the acids in 1 g of fatty material.
Unless otherwise directed, weigh accurately about 5 g of sample into a 500-ml Erlenmeyer flask, and add 75-100 ml of hot neutral ethanol. Agitation and further heating may be necessary to effect complete solution of the sample. For some samples, it may be necessary to use as the solvent a 1:1 mixture of neutralized diethyl ether/ethanol or petroleum spirit/ethanol. Add 0.5 ml of phenolphthalein TS and titrate immediately, while shaking, with 0.5 N KOH until the pink colour persists for at least 30 sec. (For acidity less than 2% by weight, 0.1 N KOH should be used for the titration; for acidity less than 0.2% by weight, it is necessary, in addition, to first neutralize the carbon dioxide in the reaction vessel.)
T is the titre (ml);
N is the normality of potassium hydroxide solution; and
W is the weight of sample (g).
Determine by Gas Chromatography using the following conditions or equivalent that will elute n-decane before benzene:
ApparatusLiquid phase: Tetracyanoethylated Pentaerythritrol (TCEPE) Length: 30 m i.d.: 0.25 mm Temperatures: Inlet: 275° Detector: 250° Column: 95° Carrier gas: N2 Flow rate: 3 cm3/min Detector Flame ionization Split 100-1
Isooctane: 99 mole percent minimum containing less than 0.05 mole percent aromatic material.
Benzene: 99.5 mole percent minimum.
Internal Standard: n-Decane and either n-undecane or n-dodecane according to the requirement of the System Suitability Test.
Reference Solution A: Prepare a standard solution containing 0.5% by weight each of the Internal Standard and of benzene in isooctane.
Reference Solution B: Prepare a standard solution containing about 0.5% by weight each of n-decane, of Internal Standard, and of benzene in isooctane.
Select the instrument conditions necessary to give the desired sensitivity. Inject a known volume of Reference Solution A, and change the attenuation, if necessary, so that the benzene peak is measured with a chart deflection of not less than 25% or more than 95% of full scale. When choosing the attenuation, consider all unresolved peaks to represent a single compound. There may be tailing of the non-aromatic peak, but do not use any conditions that lead to a depth of the valley ahead of the benzene peak (A) less than 50% of the weight of the benzene peak (B) as depicted in
If there is tailing of the non-aromatic material, construct a baseline by drawing a line from the bottom of the valley ahead of the benzene peak to the point of tangency after the peak (see ). Determine the areas of the benzene peak and the internal standard peak by use of an electronic integrator. Do not use integrators on peaks without a constant baseline, unless the integrator has provision for making baseline corrections with accuracy at least as good as that of manual methods.
Calculate a response factor for benzene (Rb) relative to the Internal Standard by the formula
Ai is the area of the Internal Standard peak in arbitrary units corrected for attenuation;
Wi is the weight percent of Internal Standard in Reference Solution A;
Ab is the area of the benzene peak in arbitrary units corrected for attenuation; and
Wi is the weight percent of benzene in Reference Solution A.
System Suitability Test
Inject the same volume of Reference Solution B as in the Calibration and record the chromatogram. n-Decane must be eluted before benzene, and the ratio of A to B () must be at least 0.5 where A is equal to the depth of the valley between the n-decane and benzene peaks and B is equal to the height of the benzene peak.
. Illustration of A/B Ratio.
. Illustration of A/B Ratio for a Small Component Peak on the Tail of a Large Peak.
Place approximately 0.1 ml of Internal Standard into a tared 25-ml volumetric flask, weigh on an analytical balance, dissolve in and dilute to volume with the sample to be analyzed.
Using the exact instrumental conditions that were used in the calibration, inject the same volume of sample containing the Internal Standard. Before measuring the area of the Internal Standard and benzene peaks, change the attenuation to ensure at least 25% chart deflection.
Measure the area of the Internal Standard and benzene peaks in the same manner as was used for the calibration. Calculate the weight percent of benzene in the sample (WB) by the formula
Ab is the area of the benzene peak corrected for attenuation;
Rb is the relative response factor for benzene;
Wi is the weight, in grams, of Internal Standard added;
Ai is the area of the Internal Standard peak corrected for attenuation; and
S is the weight, in grams of the sample taken.
Retention Times in Minutes for Selected Hydrocarbons Under the given Conditions are:
- Benzene 6.1
- Toluene 7.0
p- and m-Xylenes 8.5
- o-Xylene 10.0
(ASTM D 2502 See Test for Viscosity for Copyright permission)
Determine the kinematic viscosity of the sample at 37.8 and 98.9° as described in the method for Viscosity, 100°. Read the value of H that corresponds to the measured viscosity at 37.8° by the use of Table 1; linear interpolation between adjacent columns may be required. Read a viscosity -molecular weight chart for H and 98.9° viscosity (the chart is available from the American Society for Testing and Materials (ASTM)). A simplified version is shown in for illustration purposes only. Interpolate where necessary between adjacent lines of 98.89° viscosity. After locating the point corresponding to the value of H (ordinate) and the 98.89° viscosity (superimposed lines), read the molecular weight along the abscissa.
Kinematic viscosity, mm2/ at 37.8°
Table1 - Tabulation of H Function0 0.2 0.4 0.6 0.8
-134 -1 9
365 44 52
736 79 85 90 96
1017 106 111 116 120
1248 128 132 136 140
1449 147 151 154 157
16010 163 166
17511 178 180
18812 190 192 195 197 199 13 201 203 206 208 210 14 211 213
21915 212 222
224226 227 16 229 231 231 234
23517 237 238 240 241 243 18 244 245
24919 251 252 253 255 256 20 257 258 259 261
26221 263 264
26722 269 270 271 272 273 23 274 275 276 277
27824 279 280
28225 283 284
28726 288 289
29127 292 293
29528 295 297
29929 300 301 301 302 303 30 304 304
30631 307 308
31032 310 311
31333 314 314
31634 317 317 318 319
31935 320 320 321 322
32236 323 323
32537 325 326
327327 328 38 328 329
33139 331 332
333H 0 1 2 3 4 5 6 7 8 9 40 334 336 339 341 343 345 347 349 352 354 50 355 357 359 361 363 364 366 368 369 371 60 371 374 375 377 378 380 381 382 384 385 70 386 387 388 390 391 392 393 394 395 397 80 398 399 400 401 402 403 404 405 406 407 90 409 409 410 410 411
412413 414 415 416 100 416 417 418 419 420 420 421 422 423 423 110 424 425 423 426 427 428 428 429 430 430 120 431 432 432 433 433 434 435 435 436 437 130 437 438 438 439 439 440 441 441 442 442 140 443 443 444 444 445 446 446 447 447 448 150 448 449 449 450 450 450 451 451 452 452 160 453 453 454 454 455 455 456 456 456 457 170 457 458 458 459 459 460 460 460 461 461 180 461 462 462 463 463 463 464 464 465 465 190 465 466 466 466 467 467 468 468 468 469
H0 10 20 30 40 50 60 70 80 90 200 469 473 476 479 482 485 487 490 492 493 300 497 499 501 503 505 507 509 511 512 514 400 515 517 518 520 521 523 524 525 527 528 500 529 530 531 533 534 535 536 537 538 539 600 540 541 542 543 544 545 546 547 547 548 700 549 550 551 551 552 553 554 555 554 556
800557 557 558 559 559 560 561 562 562 563 900 563 564 565 565 566 566 567 567 568 569 1000 569 574 578 583 587 591 594 597 600 603 2000 605 608 610 614 616 618 620 621 623 625 3000 625 626 628 629 631 632 633 634 636 637 4000 638 639 640 641 642 643 644 645 646 647 5000 648 649 650 651 652 652 653 654
6566000 656 657 658 658 659 660 660 661 662 662 7000 663 664 664 665 665 666 666 667 667 668 8000 668 669 670 670 671 671 671 672 672 673 9000 673 674 674 675 675 676 676 677 677 677 10000 673 681 684 688 691 694 696 699 701 703 20000 705 707 709 711 712 715 715 717 718 719 30000 720 722 723 724 725 726 727 728 729 730 40000 731 732 733 734 735 736 737 738 739 740 50000 739 739 740 741 741 742 743 743 744 744 60000 745 746 746 747 747 748 748 749 749 750 70000 750 751 751 752 752 753 753 753 754 754 80000 755 755 756 756 756 757 758 758 758 758 90000 759 759 759 760 760 760 761 761 761 762 100000 762 762 763 763 763 764 764 764 764 765
. Lines of Constant Viscosity (mm /g) at 98.89°
ASTM D 6352 – 04
Copies of the complete ASTM standard may be purchased directly from ASTM, phone: +1 610-832-9585, fax: +1 610-832-9555, e-mail: ,
"Carbon number" is number of carbon atoms in a molecule. Determine the boiling point distribution of the sample by gas chromatography. Individual hydrocarbons are separated on a non-polar open tubular capillary column using a linear temperature program in the order of their increasing boiling points. Detector response for each paraffin shall be close to unity.
Column: Non-polar wall-coated open tubular column (5 m x 0.53 - 0.75 mm, i.d.) stationary phase, 100% dimethylpolysiloxane, 0.1 µm, or equivalent
Carrier gas: Helium, at a flow rate of 18 ml/min
Detector: FID; temperature 450˚
Oven program: 50˚ - 10˚/min - 400˚
Injector: On-column or temperature programmable vaporizing injector.
Injection volume: 0.5 µl
Calibration mixture: Prepare a mixture of hydrocarbons with known boiling points covering the range of the sample (e.g. from C10 to C90). Each component should be present at approximately 0.5 - 2.0%, dissolved in a suitable viscosity-reducing solvent such as carbon disulfide or cyclohexane.
Column resolution: Resolution is determined using C50 and C52 paraffins from a calibration mixture. Resolution shall be between 2 and 4 for satisfactory column performance.
Sample analysis: Using the schedule and temperature program used for the calibration mixture, cool the column and injector to the initial starting temperature. Inject the sample, diluted to approximately 1% in Carbon disulfide or hexane, and record the chromatogram. Inject a baseline blank, standard mixtures and samples in a predetermined order. Use the baseline blank to determine baseline drift and perform baseline subtraction from runs of samples and standards.
Calculation: Collect data, calculate the sample total area, normalise to area percent after background subtraction. Determine the initial and final boiling points by calculating 0.5% and 99.5% of the area counts, respectively. Use linear interpolation to determine the retention time associated with 5% and read the corresponding boiling temperature from the calibration curve.
Melt in a glass tube (25 mm in diameter and 100 mm in length, the glass being 1 mm in thickness) about 5 g of the sample by heating gently to 15-20° above the expected congealing range. By means of a perforated stopper, fasten the tube in a wide-mouthed bottle of clear glass, approximately 70 mm in diameter and 150 mm in height. Suspend a standard thermometer in the melted sample so that it will serve as a stirrer, cool if necessary, and stir the mass slowly until the mercury remains stationary for 30 sec. Discontinue stirring and allow the thermometer to hang, with the bulb in the centre of the sample, and observe the rise of the mercury column. The highest point to which it rises is the congealing temperature.
(Based on AOCS Method Ca 5a-40)
Unless otherwise directed in the specification monograph, weigh accurately the appropriate amount of the sample, indicated in the table below, into a 250-ml Erlenmeyer flask or other suitable container. Add 2 ml of phenolphthalein TS to the specified amount of hot alcohol, neutralize with alkali to the first faint but permanent pink colour, and then add the hot neutralized alcohol to the sample container. Titrate with the appropriate normality of sodium hydroxide, shaking vigorously, to the first permanent pink colour of the same intensity as that of the neutralized alcohol. The colour must persist for at least 30 sec. Calculate the percentage of free fatty acids (FFA) in the sample by the formula VN/W, in which V is the volume and N is the normality, respectively, of the sodium hydroxide used, W is the weight of the sample, in g, and e is the equivalence factor given in the monograph.FFA Range (%) g of sample ml of alcohol Strength of NaOH 0.00-0.2 56.4 ± 0.2 50 0.1 0.2-1.0
28.2 ±0.250 0.1 1.0-30.0 7.05 ± 0.05 75 0.25 30.0-50.0 7.05 ± 0.05 100 0.25-1.0 50.0-100 3.525 ±0.001 100 1.0
(Based on AOCS Method Cd 13-60)
Note: This method involves use ofpyridine which should be used with appropriate caution.
Hydroxyl value is defined as the number of mg of potassium hydroxide required to neutralize the amount of acetic acid capable of combining by acetylation with 1 g of sample.
Weigh accurately the appropriate amount of sample according to the expected hydroxyl value and transfer it into a 250-ml glass-stoppered Erlenmeyer flask.Hydroxyl value Sample weight (g) 0 to 20 10 20 to 50 5 50 to 100 3 100 to 200 2
Pipet 5.0 ml of pyridine/acetic anhydride TS into the flask. (For samples having a 0-20 hydroxyl value, add an additional 5 ml of pyridine/acetic anhydride TS to the flask.) Thoroughly mix the contents by gently swirling. Pipet 5.0 ml of pyridine/acetic anhydride TS into an empty flask for the reagent blank. (If 10.0 ml of the reagent were used for the acetylation, use a 10.0-ml blank.) Place the flasks on a steam bath, under reflux condensers, and heat for 1 h. To hydrolyze excess acetic anhydride, add sufficient water (not exceeding about 10 ml) through the condensers to the flasks. If the solution separates into two layers, add sufficient pyridine to obtain a homogeneous solution. Heat on a steam bath for 10 min with reflux condensers attached. Add 25 ml of neutralized n-butanol, about half of it through the condensers and the remainder to wash down the sides of the flasks after removal of the condensers. Add 1 ml of phenolphthalein TS and titrate to a faint pink endpoint with 0.5 N ethanolic KOH solution. To correct for free acid, mix about 10 g of the sample, accurately weighed, with 10 ml of pyridine (neutralized to phenolphthalein), add 1 ml of phenolphthalein TS and titrate to a faint pink endpoint with 0.5 N ethanolic potassium hydroxide.
Calculate the hydroxyl value by the formula:
A is ml of KOH solution required for the free acid determination;
B is ml of KOH solution required for the reagent blank;
C is the weight of sample used for the free acid determination;
S is ml of KOH solution required for titration of the acetylated sample;
W is weight of sample used for acetylation; and
N is normality of the ethanolic KOH solution.
Reflux 1 g of sample with 15 ml of 0.5 N ethanolic potassium hydroxide for 1 h. Add 15 ml of water, acidify with dilute hydrochloric acid TS (about 6 ml). Oily drops or a white to yellowish-white solid is produced which is soluble in 5 ml of hexane.
Remove the hexane layer, extract again with 5 ml of hexane and again remove the hexane layer. Collect all the hexane extracts together. The fatty acids thus extracted may be identified by gas-liquid chromatography (see Test A). Carry out the whole of the procedure in a fume cupboard. The aqueous layer is used for Tests B through H.
(Based on AOCS Methods Ce 1-62, Ce 1f-96, Ce 1h-05)
Use a suitable gas chromatograph equipped with a flame ionization detector (FID) and containing a 50-m x 0.25-mm id capillary fused silica column, or equivalent, containing a suitable highly polar stationary phase (0.20 m)film, such as CP™-Sil 88, SP-2650, SP-2340, BPX-70, or SP2560.
Note: For accurate determination of all fatty acids present in non-ruminant animal and vegetable oils and fats, a 100m SP2560 or CP-Sil 88 column is recommended.
The operating conditions may vary with the instrument used, but a suitable chromatogram may be obtained isothermally at temperatures between 170° and 198°, depending on the column stationary phase. Inlet temperature (injector), 250°; detector, 250°; and a suitable hydrogen or helium carrier gas flow.
Column performance is checked using a suitable mixture of fatty acid methyl esters covering the range of fatty acids under investigation Fatty acid methyl esters with a wide range of carbon numbers and double-bond configurations can be purchased. A mixture containing C12:0; 9c-18:1; 11c-18:1, 9c,12c,15c-18:3; 11c-20:1; and an Internal Standard (C21:0) using each carrier gas and column combination. Since commercial GC designs are different, to achieve optimal separation small changes in the sample size, sample concentration or oven temperature may be required. If so, adjust the sample size, sample concentration or oven temperature until the best separation results are obtained. Baseline separation of the various components in both the standard and the sample preparations is desirable.
Sample Preparation (for fats and oils) (Based on AOCS Method Ce 2-66)
Introduce 100 to 1000 mg of the fat into a 50- or 125-mL reaction flask. Add 4 to 10 ml of 0.5 N methanolic sodium hydroxide, and add a boiling chip. Attach a condenser, and heat the mixture on a steam bath until the fat globules go into solution. This step should take 5 to 10 min. Add 5 to 12 ml of 12.5% boron fluoride-methanol reagent (this reagent contains 125 g/l of boron fluoride in methanol and is available commercially) through the condenser, and boil for 2 min. Add 2 to 5 ml of heptane through the condenser, and boil for 1 min longer. Remove from heat, remove condenser, and add about 15 ml of saturated sodium chloride solution. Stopper the flask, and shake vigorously for 15 sec. Transfer about 1 ml of the heptane solution into a test tube and add a small amount of anhydrous sodium sulfate. The dry heptane solution may then be injected directly into a gas chromatograph.
The methyl esters should be analyzed as soon as possible. They may be kept in an atmosphere of nitrogen in a screwcap vial at 2° for 24 h. For longer storage, they should be sealed in a glass ampoule, subjected first to a vacuum and then backfilled with nitrogen and stored at -20° (freezer).
Inject an appropriate volume (1 μl) of sample into the chromatograph. If an automated system is used, follow the manufacturer's instructions; if calculations are to be done manually, proceed as follows:
Calculate the area percent of each component (CN) by the equation
in which AN is the area of the peak corresponding to component CN and TS is the total area for all detected components [TS = ΣAN].
Calculated Iodine Value (Based on AOCS Method Cd 1c-85)
Using the fatty acid composition determined above, calculate the Iodine value (IV) as follows:
- Triglycerides, iodine value = (% hexadecenoic acid x 0.950) + (% octadecenoic acid x 0.860) + (% octadecadienoic acid x 1.732) + (% octadecatrienoic acid x 2.616) + (% eicosenoic acid x 0.785) + (% docosenoic acid x 0.723)
- Free fatty acids, iodine value = (% hexadecenoic acid x 0.9976) + (% octadecenoic acid x 0.8986) + (% octadecadienoic acid x 1.810) + (% octadecatrienoic acid x 2.735) + (% eicosenoic acid x 0.8175) + (% docosenoic acid x 0.7497).
Note: This procedure is not intended to be a rapid method, but rather gives two results from one analysis. For oils with an unsaponifiable content greater than 0.5% (e.g., fish oils), and for materials with low iodine value the calculation tends to be low. Calculated IV based on GC fatty acid composition of non-triglyceride lipid materials such as partial esters of glycerol, partial esters of sorbitol/sorbitan/isosorbide esters, partial esters of polyoxyethylene sorbitol/sorbitan/isosorbide or glycerol, etc. will provide the calculated IV of only the fatty acids used to prepare the partial esters. To obtain the actual IV of partial esters with nonfatty acid poly ol diluents, the chlorinated Wijs Reagent IV method should be used. IV values of partial esters via the Wijs method are lower than those obtained by GC because of the dilution effect of the polyol material.
Transfer about 5 ml of the aqueous layer resulting from the hydrolysis into a dish, add excess calcium carbonate and evaporate until dry. Transfer the major part of the residue into a glass tube. Place a filter paper, moistened with Reagent for acetone (a saturated solution of o-nitrobenzaldehyde in sodium hydroxide TS, freshly prepared) on top of the tube. Heat over a micro flame. The yellow colour of the paper changes into greenish blue by reaction of the Reagent for acetone, with the calcium acetate formed.
Transfer one drop of the aqueous layer resulting from the hydrolysis and a drop of a 0.5% solution of ammonium chloride and several mg of zinc powder into a micro test tube.
The mouth of the tube is covered with a disk of filter paper moistened with a solution in benzene of 5% p-dimethylamino-benzaldehyde and 20% trichloroacetic acid. The bottom of the test tube is heated vigorously with a micro flame for about 1 min. Depending on the amount of succinic acid or succinimide, a red-violet or pink stain appears on the paper.
Transfer 1 ml of the aqueous layer resulting from the hydrolysis with 1 ml of 2 N sodium carbonate into a test tube. Add 2 or 3 drops of 0.1 N potassium permanganate. The solution is promptly discoloured.
Evaporate about 5 ml of the aqueous layer resulting from the hydrolysis in a porcelain dish until dry. Add 2 ml of concentrated sulfuric acid containing 0.5% of pyrogallol and heat on a steam bath. An intense violet colour is produced.
To 3 ml of the aqueous layer resulting from the hydrolysis add a few drops of 1 % potassium permanganate and warm until the colour has disappeared. Then add an excess of bromine TS. A white precipitate (pentabromoacetone) is formed immediately or on cooling.
Evaporate 1 ml of the aqueous layer resulting from the hydrolysis in a porcelain dish, add 1 ml of a mixture of 1 vol acetic anhydride and 5 vol of pyridine into the warm dish. A violet colour is produced. (Tartaric acid produces a green colour.)
Transfer 0.2 ml of the aqueous layer resulting from the hydrolysis and 2 ml of concentrated sulfuric acid into a test tube and place for 2 min in boiling water. Cool and add 1 or 2 drops of a 5% guaiacol solution in ethanol. A red colour is immediately produced.
If tartaric acid is present according to Test E, it must be removed as follows: transfer 3 ml of the aqueous layer resulting from the Hydrolysis and an excess of calcium hydroxide as a powder into a test tube, place in boiling water for 5 min, shaking several times, cool and filter.
Transfer 5 ml of the aqueous layer resulting from the hydrolysis into a test tube. Add excess calcium hydroxide as a powder, place in boiling water for 5 min, shaking several times, cool and filter.
Transfer one drop of the filtrate into a tube and add about 50 mg of potassium hydrogen sulfate. Place a filter paper, moistened with Reagent for acrolein (a 5% solution of disodium pentacyanonitrosylferrate in water and a 20% piperidine solution in water; mix the solutions 1:1 immediately before use) on the top of the tube. Heat over a micro flame. A blue coloured filter paper indicates the presence of glycerol. The colour changes to light red after addition of sodium hydroxide TS.
The test cannot be employed in the presence of ethylene glycol or lactic acid, since they decompose under the prescribed conditions yielding acetaldehyde which reacts with the reagents in the same manner as acrolein.
(Based on AOCS Method Cd 1d-92)
The iodine value (IV) is a measure of unsaturation and is expressed as the number of g of iodine absorbed, under the prescribed conditions, by 100 g of the test substance. For fats and oils, the Iodine Value may be calculated from the results of gas chromatographic quantification of methyl ester (see Methyl Esters of Fatty Acids, above)
Glacial acetic acid
Wijs Solution: this reagent should be purchased commercially.
Potassium iodide TS
N sodium thiosulfate
The appropriate weight of the sample, in g, is calculated by dividing the number 25 by the expected iodine value. Melt the sample, if necessary, and filter it through a dry filter paper. Transfer the accurately weighed quantity of sample into a clean, dry, 500-ml glass-stoppered bottle or flask containing 20 ml of glacial acetic acid/cyclohexane, 1:1, v/v, and pipet 25.0 ml of Wijs Solution into the flask. The excess of iodine should be between 50% and 60% of the quantity added, that is, between 100% and 150% of the quantity absorbed. Swirl, and let stand in the dark for 1.0 h where the iodine value is <150 and for 2.0 h where the iodine value is >150. Add 20 ml of potassium iodide TS and 100 ml of recently boiled and cooled water, and titrate the excess iodine with 0.1 N sodium thiosulfate, adding the titrant gradually and shaking constantly until the yellow colour of the solution almost disappears. Add starch TS, and continue the titration until the blue colour disappears entirely. Toward the end of the titration, stopper the container and shake it violently so that any iodine remaining in solution in the glacial acetic acid/cyclohexane layer may be taken up by the potassium iodide solution. Concomitantly, conduct two determinations on blanks in the same manner and at the same temperature.
Calculate the iodine value by the formula
B - S is the difference between the volumes of sodium thiosulfate required for the blank and for the sample, respectively;
N is the normality of the sodium thiosulfate;
W is the weight, in g, of the sample.
Preparation of Samples
Solid Samples in Flake Form: Mix without melting and take a portion for analysis.
Solid Samples not in Flake Form: Melt at not more than 10° above melting point, mix thoroughly and take a portion for analysis. Do not attempt to test samples which contain so much free glycerol that it separates when the sample solidifies.
Semi-solid and Liquid Samples: Liquefy by heating at not more than 10° above melting point, mix thoroughly, and take a portion for analysis. Do not attempt to test samples which contain so much free glycerol that it separates from the sample when cooled to room temperature.
Caution: The sample must not be subjected to a temperature in excess of that required to melt it, as this may reduce the monoglyceride content if any soap is present.
Procedure for 1-Monoglyceride
Weigh to the nearest mg duplicate samples of 1 g into a 100-ml glass-stoppered volumetric flask. Dissolve in 50 ml of chloroform. Add 25 ml of water and shake vigorously for 30-60 sec. Transfer the aqueous layer to a glass-stoppered 100-ml volumetric flask, using a glass siphon. If an emulsion forms due to the presence of soap in the sample, add 3 or 4 ml of glacial acetic acid to break the emulsion. Extract 3 more times using 25, 25 and 20 ml of distilled water. Add chloroform to the flask until the level of the chloroform coincides with the 100-ml mark. Using the glass siphon, transfer as much as possible of the aqueous layer above the chloroform layer to the flask containing the aqueous extracts. The aqueous extracts in the volumetric flask are saved for the determination of free glycerol.
Pipet 50 ml of acetic periodic acid TS into each of a series of 500-ml glass-stoppered Erlenmeyer flasks. Prepare 3 for blanks, adding 50 ml of chloroform to two and 50 ml of water to the third. The titrations of the water and chloroform blanks are used as a check (within 0.5 ml) on the chloroform. Pipet 50 ml of chloroform sample solution into one the flasks containing 50 ml of acetic periodic acid TS and shake gently to effect thorough mixing. Allow to stand for at least 30 min but not longer than 1.5 h. To each flask add 20 ml of potassium iodide TS. Mix by gentle shaking, allow to stand at least 1 min but not more than 5 min before titrating. Do not allow to stand in strong sunlight. Add 100 ml of distilled water and titrate with 0.1 N sodium thiosulfate. Use a variable speed magnetic stirrer to keep the solution thoroughly mixed. Continue the titration to the disappearance of the brown iodine colour from the aqueous layer. Add 2 ml of starch TS and continue the titration to the disappearance of iodine from the chloroform layer and the disappearance of the blue iodo-starch colour from the aqueous layer.
Calculation of 1-monoglyceride content as pure monostearate:
B is the sodium thiosulfate consumed in the titration of blank containing 50 ml of chloroform;
S is the sodium thiosulfate consumed in the titration of sample;
N is the exact normality of 0.1 N sodium thiosulfate; and
W is the weight of sample, represented by aliquot pipetted for test.
The 1-monoglyceride content may be calculated in terms of a monoester other than the monostearate by dividing the molecular weight of the monoglyceride by 20 and substituting this value for 17.927 in the formula above.
Procedure for free Glycerol
Add distilled water to the combined aqueous extracts from the monoglyceride determination until the volume is 100 ml and mix thoroughly. Pipet 50 ml of acetic periodic acid TS into each of a series of 500-ml glass-stoppered Erlenmeyer flasks. Pipet 50 ml of aqueous sample solution into one of the flasks containing 50 ml of acetic periodic acid TS and shake gently to effect thorough mixing. Continue as described under the procedure for monoglyceride, second paragraph commencing "Allow to stand for at least 30 min...".
Calculation of glycerol content:
B is the sodium thiosulfate consumed in the titration of blank containing 50 ml of water;
S is the sodium thiosulfate consumed in the titration of sample;
N is the exact normality of 0.1 N thiosulfate; and
W is the weight of sample represented by aliquot pipetted for test.
Because of the sensitivity of the test, the possibility of errors arising from contamination is great. It is of the greatest importance, therefore, that all glassware be scrupulously cleaned to remove all organic matter such as oil, grease, detergent residues, etc. Examine all glassware, including stoppers and stopcocks, under ultraviolet light to detect any residual fluorescent contamination. As a precautionary measure it is a recommended practice to rinse all glassware with purified isooctane immediately before use. No grease is to be used on stopcocks or joints. Great care to avoid contamination of wax samples in handling and to assure absence of any extraneous material arising from inadequate packaging is essential. Because some of the polynuclear hydrocarbons sought in this test are very susceptible to photo-oxidation, the entire procedure is to be carried out under subdued light.
Separatory funnels: 250-ml, 500-ml, 1,000-ml, and preferably 2,000-ml capacity, equipped with tetrafluoroethylene polymer stopcocks.
Reservoir: 500 ml capacity, equipped with a 24/40 standard taper male fitting at the bottom and a suitable balljoint at the top for connecting to the nitrogen supply. The male fitting should be equipped with glass hooks.
Chromatographic tube: 180 mm in length, inside diameter to be 15.7 ±0.1 mm, equipped with a coarse, fritted-glass disc, a tetrafluoroethylene polymer stopcock, and a female 24/40 standard tapered fitting at the opposite end. (Overall length of the column with the female joint is 235 mm).
Disc: Tetrafluoroethylene polymer 2-inch diameter disc approximately 3/16-inch thick with a hole bored in the center to closely fit the stem of the chromatographic tube.
Heating jacket: Conical, for 500-ml separatory funnel. (Used with variable transformer heat control).
Suction flask: 250-ml or 500-ml filter flask.
Condenser: 24/40 joints, fitted with a drying tube, length optional.
Evaporation flask (optional): 250-ml or 500-ml capacity all-glass flask equipped with standard taper stopper having inlet and outlet tubes permit passage of nitrogen across the surface of the liquid to be evaporated.
Vacuum distillation assembly: All glass (for purification of dimethyl sulfoxide); 2-l distillation flask with heating mantle; Vigreaux vacuum-jacketed condenser (or equivalent) about 45 cm in length and distilling head with separable cold finger condenser. Use of tetrafluoroethylene polymer sleeves on the glass joints will prevent freezing. Do not use grease on stopcocks or joints.
Spectrophotometric cells: Fused quartz cells, optical path length in the range of 5.000 ± 0.005 cm; also for checking spectrophotometer performance only, optical path length in the range 1.000 ± 0.005 cm. With distilled water in the cells, determine any absorbance differences.
Spectrophotometer: Spectral range 250 - 400 nm with spectral slit width of 2 nm or less, under instrument operating conditions for these absorbance measurements, the spectrophotometer shall, also meet the following performance requirements:
- Absorbance repeatability, ±0.01 at 0.4 absorbance.
- Absorbance accuracy, ±0.05 at 0.4 absorbance.
- Wavelength repeatability, ±0.2 nm.
- Wavelength accuracy, ±1.0 nm.
Nitrogen cylinder: Water-pumped or equivalent purity nitrogen in cylinder equipped with regulator and valve to control flow at 5 p.s.i.g.
Reagents and materials
All solvents used throughout the procedure shall meet the specifications and tests described below. The isooctane, benzene, acetone, and methyl alcohol designated in the list following this paragraph shall pass the following test:
To the specified quantity of solvent in a 250-ml Erlenmeyer flask, add 1 ml of purified n-hexadecane and evaporate on the steam bath under a stream of nitrogen (a loose aluminium foil jacket around the flask will speed evaporation). Discontinue evaporation when not over 1 ml of residue remains. (to the residue from benzene add a 10-ml portion of purified isooctane, re-evaporate, and repeat once to insure complete removal of benzene).
Alternatively, the evaporation time can be reduced by using the optional evaporation flask. In this case the solvent and n-hexadecane are placed in the flask on the steam bath, the tube assembly is inserted, and a stream of nitrogen is fed through the inlet tube while the outlet tube is connected to a solvent trap and vacuum line in such a way as to prevent any flow-back of condensate into the flask.
Dissolve the 1 ml of hexadecane residue in isooctane and make to 25 ml volume. Determine the absorbance in the 5 cm path length cells compared to isoooctane as reference. The absorbance of the solution of the solvent residue (except for methyl alcohol) shall not exceed 0.01 per cm path length between 280 and 400 nm. For methyl alcohol this absorbance value shall be 0.00.
Isooctane (2,2,4-trimethylpentane): Use 180 ml for the test described in the preceding paragraph. Purify, if necessary, by passage through a column of activated silica gel (Grade 12 or equivalent) about 90 cm in length and 5 cm to 8 cm in diameter.
Benzene, reagent grade: Use 150 ml for the test. Purify, if necessary, by distillation or otherwise.
Acetone, reagent grade: Use 200 ml for the test. Purify, if necessary, by distillation.
- 10% benzene in isooctane: Pipet 50 ml of benzene into a 500-ml glass-stoppered volumetric flask and adjust to volume with isooctane, with mixing.
- 20% benzene in isooctane: Pipet 50 ml of benzene into a 250-ml glass-stoppered volumetric flask, and adjust to volume with isooctane, with mixing.
- Acetone-benzene-water mixture: Add 20 ml of water to 380 ml of acetone and 200 ml of benzene, and mix.
n-Hexadecane, 99% olefin-free: Dilute 1.0 ml of n-hexadecane to 25 ml with isooctane and determine the absorbance in a 5-cm cell compared to isooctane as reference point between 280-400 nm. The absorbance per centimeter path length shall not exceed 0.00 in this range. Purify, if necessary, by percolation through activated silica gel or by distillation.
Methyl alcohol, reagent grade: Use 10.0 ml of methyl alcohol. Purify, if necessary, by distillation.
Dimethyl sulfoxide: Pure grade, clear, water-white, m.p. 18° minimum. Dilute 120 ml of dimethyl sulfoxide with 240 ml of distilled water in a 500-ml separatory funnel, mix and allow to cool for 5-10 min. Add 40 ml of isooctane to the solution and extract by shaking the funnel vigorously for 2 min. Draw off the lower aqueous layer into a second 500 ml separatory funnel and repeat the extraction with 40 ml of isooctane. Draw off and discard the aqueous layer. Wash each of the 40 ml isooctane portions three times with 50 ml portions of distilled water. Shaking time for each wash is 1 min. Discard the aqueous layers. Filter the first isooctane portion through anhydrous sodium sulfate prewashed with isooctane (see sodium sulfate below for preparation of filter), into a 250-ml Erlenmeyer flask, or optionally into the evaporating flask. Wash the first separatory funnel with the second 40 ml isooctane portion, and pass through the sodium sulfate into the flask. Then wash the second and first separatory funnels successively with a 10 ml portion of isooctane, and pass the solvent through the sodium sulfate into the flask. Add 1 ml of n-hexadecane and evaporate the isooctane on the steam bath under nitrogen. Discontinue evaporation when not over 1 ml of residue remains. To the residue, add a 10 ml portion of isooctane and re-evaporate to 1 ml of hexadecane. Again, add 10 ml of isooctane to the residue and evaporate to 1 ml of hexadecane to insure complete removal of all volatile materials. Dissolve the 1 ml of hexadecane in isooctane and make to 25 ml volume. Determine the absorbance in 5 cm path length cells compared to isooctane as reference. The absorbance of the solution should not exceed 0.02 per cm path length in the 280-400 nm range.
Note: Difficulty in meeting this absorbance specification may be due to organic impurities in the distilled water. Repetition of the test omitting the dimethyl sulfoxide will disclose their presence. If necessary to meet the specification, purify the water by re-distillation, passage through an ion-exchange resin, or otherwise.
Purify, if necessary, by the following procedure: To 1,500 ml of dimethyl sulfoxide in a 2 l glass-stoppered flask, add 6.0 ml of phosphoric acid and 50 g of Norit A (decolorizing carbon, alkaline) or equivalent. Stopper the flask, and with the use of a magnetic stirrer (tetrafluoro-ethylene polymer coated bar) stir the solvent for 15 min. Filter the dimethyl sulfoxide through four thicknesses of fluted paper (18.5 cm, Schleicher & Schuell, No. 597, or equivalent). If the initial filtrate contains carbon fines, refilter through the same filter until a clear filtrate is obtained. Protect the sulfoxide from air and moisture during this operation by covering the solvent in the funnel and collection flask with a layer of isooctane. Transfer the filtrate to a 2-l separatory funnel and draw off the dimethyl sulfoxide into the 2-l distillation flask of the vacuum distillation assembly and distil at approximately 3 mm Hg pressure or less. Discard the first 200 ml fraction of the distillate and replace the distillate collection flask with a clean one. Continue the distillation until approximately 1 l of the sulfoxide has been collected.
At completion of the distillation, the reagent should be stored in glass-stoppered bottles since it is very hygroscopic and will react with some metal containers in the presence of air.
Phosphoric acid, 85% reagent grade
Sodium borohydride, 98%
Magnesium oxide (Sea Sorb 43, Food Machinery Company, Westvaco Division, distributed by chemical supply firms, or equivalent): Place 100 g of the magnesium oxide in a large beaker, add 700 ml of distilled water to make a thin slurry, and heat on a steam bath for 30 min with intermittent stirring. Stir well initially to insure that all the absorbent is completely wetted. Using a Buchner funnel and a filter paper (Schleicher & Schuell No. 597, or equivalent) of suitable diameter, filter with suction. Continue suction until water no longer drips from the funnel. Transfer the absorbent to a glass trough lined with aluminium foil (free from rolling oil). Break up the magnesia with a clean spatula and spread out the absorbent on the aluminium foil in a layer about 1-2 cm thick. Dry for 24 h at 160±l°. Pulverize the magnesia with mortar and pestle. Sieve the pulverized absorbent between 60-180 mesh. Use the magnesia retained on the 180-mesh sieve.
Celite 545: Johns-Manville Company, diatomaceous earth, or equivalent.
Magnesium oxide-Celite 545 mixture: Place the magnesium oxide (60-180 mesh) and the Celite 545 in 2 to 1 proportions, respectively, by weight in a glass-stoppered flask large enough for adequate mixing. Shake vigorously for 10 min. Transfer the mixture to a glass trough lined with aluminium foil (free from rolling oil) and spread it out on a layer about 1 to 2 cm thick. Reheat the mixture at 160±1° for 2 h, and store in a tightly closed flask.
Sodium sulfate, anhydrous, reagent grade, preferably in granular form: For each bottle of sodium sulfate reagent used, establish as follows the necessary sodium sulfate prewash to provide such filters required in the method: Place approximately 35 g of anhydrous sodium sulfate in a 30 ml coarse, fritted-glass funnel or in a 65 ml filter funnel with glass wool plug; wash with successive 15 ml portions of the indicated solvent until a 15 ml portion of the wash shows 0.00 absorbance per cm path length between 280 nm and 400 nm when tested as prescribed under Organic solvents above. Usually three portions of wash solvent are sufficient.
Before proceeding with the analysis of a sample, determine the absorbance in a 5 cm path cell between 250 nm and 400 nm for the reagent blank by carrying out the procedure, without a wax sample, at room temperature, recording the spectra after the extraction stage and after the complete procedure as prescribed. The absorbance per cm path length following the extraction stage should not exceed 0.040 in the wavelength range from 250 to 400 nm; the absorbance per cm path length following the complete procedure should not exceed 0.070 in the wavelength range from 250 to 299 nm, inclusive, or 0.045 in the wavelength range from 300 nm to 400 nm. If in either spectrum the characteristic benzene peaks in the 250-260 nm region are present, remove the benzene by the procedure under Organic solvents, above, and record absorbance again.
Place 300 ml of dimethyl sulfoxide in a 1-l separatory funnel and add 75 ml of phosphoric acid. Mix the contents of the funnel and allow to stand for 10 min. (The reaction between the sulfoxide and the acid is exothermic. Release pressure after mixing, then keep funnel stoppered). Add 150 ml of isooctane and shake to pre-equilibrate the solvents. Draw off the individual layers and store in glass-stoppered flasks.
Place a representative 1 kg sample of wax, or if this amount is not available, the entire sample, in a beaker of a capacity about three times the volume of the sample and heat with occasional stirring on a steam bath until the wax is completely melted and homogenous. Weigh four 25 ± 0.2 g portions of the melted wax in separate 100 ml beakers. Reserve three of the portions for later replicate analyses as necessary. Pour one weighed portion immediately after re-melting (on the steam bath) into a 500 ml separatory funnel containing 100 ml of the pre-equilibrated sulfoxide-phosphoric acid mixture that has been heated in the heating jacket at a temperature just high enough to keep the wax melted. (Note: In preheating the sulfoxide-acid mixture, remove the stopper of the separatory funnel at intervals to release the pressure).
Promptly complete the transfer of the sample to the funnel in the jacket with portions of the pre-equilibrated isooctane, warming the beaker, if necessary, and using a total volume of just 50 ml of the solvent. If the wax comes out of solution during these operations, let the stoppered funnel remain in the jacket until the wax re-dissolves. (Remove stopper from the funnel at intervals to release pressure).
When the sample is in solution, remove the funnel from the jacket and shake it vigorously for 2 min. Set up three 250 ml separatory funnels with each containing 30 ml of pre-equilibrated isooctane. After separation of the liquid phases, allow to cool until the main portion of the sample-isooctane solution begins to show a precipitate. Gently swirl the funnel when precipitation first occurs on the inside surface of the funnel to accelerate this process. Carefully draw off the lower layer, filter it slowly through a thin layer of glass wool fitted loosely in a filter funnel into the first 250 ml separatory funnel, and wash in tandem with the 30 ml portions of isooctane contained in the 250 ml separatory funnels. Shaking time for each wash is 1 min. Repeat the extraction operation with two additional portions of the sulfoxide-acid mixture, replacing the funnel in the jacket after each extraction to keep the sample in solution and washing each extractive in tandem through the same three portions of isooctane.
Collect the successive extractives (300 ml total) in a separatory funnel (preferably 2-liter), containing 480 ml of distilled water, mix, and allow to cool for a few min after the last extractive has been added. Add 80 ml of isooctane to the solution and extract by shaking the funnel vigorously for 2 min. Draw off the lower aqueous layer into a second separatory funnel (preferably 2-l) and repeat the extraction with 80 ml of isooctane. Draw off and discard the aqueous layer. Wash each of the 80 ml extractives three times with 100 ml portions of distilled water. Shaking time for each wash is 1 min. Discard the aqueous layers. Filter the first extractive through anhydrous sodium sulfate prewashed with isooctane (see Sodium Sulfate above for preparation of filter) into a 250-ml Erlenmeyer flask (or optionally into the evaporation flask). Wash the first separatory funnel with the second 80 ml isooctane extractive and pass through the sodium sulfate. Then wash the second and first separatory funnels successively with a 20 ml portion of isooctane and pass the solvent through the sodium sulfate into the flask. Add 1 ml of n-hexadecane and evaporate the isooctane on the steam bath under nitrogen. Discontinue evaporation when not over 1 ml of residue remains. To the residue, add a 10 ml portion of isooctane, re-evaporate to 1 ml of hexadecane, and repeat this operation once more.
Quantitatively transfer the residue with isooctane to a 25 ml volumetric flask, make to volume, and mix. Determine the absorbance of the solution in the 5 cm path length cells compared to isooctane as reference between 280 nm and 400 nm (take care to lose none of the solution in filling the sample cell). Correct the absorbance values for any absorbance derived from reagents as determined by carrying out the procedure without the sample. If the corrected absorbance does not exceed the limits prescribed in the Characteristics, the sample meets the ultraviolet absorbance specifications. If the corrected absorbance per cm path length exceeds the limits prescribed in the Characteristics, proceed as follows:
Quantitatively transfer the isooctane solution to a 125 ml flask equipped with 24/40 joint and evaporate the isooctane on the steam bath under a stream of nitrogen to a volume of 1 ml of hexadecane. Add 10 ml of methyl alcohol and approximately 0.3 g of sodium borohydride (Minimize exposure of the borohydride to the atmosphere. A measuring dipper may be used). Immediately fit a water-cooled condenser equipped with a 24/40 joint and with a drying tube into the flask, mix until the borohydride is dissolved, and allow to stand for 30 min at room temperature, with intermittent swirling. At the end of this period, disconnect the flask and evaporate the methyl alcohol on the steam bath under nitrogen until the sodium borohydride begins to come out of the solution. Then add 10 ml of isooctane and evaporate to a volume of about 2-3 ml. Again, add 10 ml of isooctane and concentrate to a volume of approximately 5 ml. Swirl the flask repeatedly to assure adequate washing of the sodium borohydride residues.
Fit the tetrafluoroethylene polymer disc on the upper part of the stem of the chromatographic tube, then place the tube with the disc on the suction flask and apply the vacuum (approximately 135 mm Hg). Weigh out 14 g of the 2:1 magnesium oxide-Celite 545 mixture and pour the adsorbent mixture into the chromatographic tube in approximately 3 cm layers. After the addition of each layer, level off the top of the adsorbent with a flat glass rod or metal plunger by pressing down firmly until the adsorbent is well packed. Loosen the topmost few mm of each adsorbent layer with the end of a metal rod before the addition of the next layer. Continue packing in this manner until all the 14 g of the adsorbent is added to the tube. Level off the top of the adsorbent by pressing down firmly with a flat glass rod or metal plunger to make the depth of the adsorbent bed approximately 12.5 cm in depth. Turn off the vacuum and remove the suction flask. Fit the 500 ml reservoir onto the top of the chromatographic column and pre-wet the column by passing 100 ml of isooctane through the column. Adjust the nitrogen pressure so that the rate of descent of the isooctane coming off of the column is between 2-3 ml per min. Discontinue pressure just before the last of the isooctane reaches the level of the adsorbent. (Caution: Do not allow the liquid level to recede below the adsorbent level at any time). Remove the reservoir and decant the 5 ml isooctane concentrate solution onto the column and with slight pressure again allow the liquid level to recede to barely above the adsorbent level. Rapidly complete the transfer similarly with two 5 ml portions of isooctane, swirling the flask repeatedly each time to assure adequate washing of the residue. Just before the final 5 ml wash reaches the top of the adsorbent, add 100 ml of isooctane to the reservoir and continue the percolation at the 2-3 ml per min rate. Just before the last of the isooctane reaches the adsorbent level, add 100 ml of 10% benzene in isooctane to the reservoir and continue the percolation at the aforementioned rate. Just before the solvent mixture reaches adsorbent level, add 25 ml of 20% benzene in isooctane to the reservoir and continue the percolation at 2-3 ml per min until all this solvent mixture has been removed from the column. Discard all the elution solvents collected up to this point.
Add 300 ml of the acetone-benzene-water mixture to the reservoir and percolate through the column to elute the polynuclear compounds. Collect the eluate in a clean 1-l separatory funnel. Allow the column to drain until most of the solvent mixture is removed. Wash the eluate three times with 300 ml portions of distilled water, shaking well for each wash. (The addition of small amounts of sodium chloride facilitates separation). Discard the aqueous layer after each wash. After the final separation, filter the residual benzene through anhydrous sodium sulfate prewashed with benzene (see Sodium sulfate under "Reagents and Materials" for preparation of filter) into a 250-ml Erlenmeyer flask (or optionally into the evaporation flask). Wash the separatory funnel with two additional 20 ml portions of benzene which are also filtered through the sodium sulfate. Add 1 ml of n-hexadecane and completely remove the benzene by evaporation under nitrogen, using the special procedure to eliminate benzene as previously described under Organic Solvents. Quantitatively transfer the residue with isooctane to a 25 ml volumetric flask and adjust the volume. Determine the absorbance of the solution in the 5 cm path length cells compared to isooctane as reference between 250 and 400 nm. Correct for any absorbance derived from the reagents as determined by carrying out the procedure without a wash sample. If either spectrum shows the characteristic benzene peaks in the 250 - 260 nm region, evaporate the solution to remove benzene by the procedure under Organic Solvents. Dissolve the residue, transfer quantitatively, and adjust to volume in isooctane in a 25 ml volumetric flask. Record the absorbance again. If the corrected absorbance does not exceed the limits prescribed in the Characteristics the sample meets the ultraviolet absorbance specifications.
Polyglycerol esters are saponified with alcoholic potassium hydroxide solution and the fatty acids removed by extraction. The polyols are converted to trimethylsilyl (TMS) derivatives and analyzed by gas liquid chromatography.
Preparation of the polyol sample
Weigh about 0.5 g of sample and reflux with 20 ml of ethanolic potassium hydroxide solution (1 N) for 2 h. Reduce the volume of ethanol by evaporation at 45-50° in a stream of nitrogen. Add 10 ml of water and convert the soaps to free fatty acids by acidifying with concentrated hydrochloric acid. Extract the fatty acids from the aqueous phase with successive 20 ml portions of light petroleum (boiling range 40-60°). Wash the combined petroleum extracts with water (20 ml) and combine the wash with the aqueous phase.
Adjust the aqueous polyol solution to pH 7.0 with aqueous potassium hydroxide solution with the aid of a pH-meter. Evaporate to a small volume (2-3 ml) under reduced pressure and extract three times with 30 ml of boiling ethanol. Filter off any residue and evaporate the ethanol under reduced pressure to yield a viscous liquid sample of polyols.
Dissolve a 0.1 g sample of polyol in 0.5 ml of warm pyridine (previously dried over potassium hydroxide) in a 10-ml capped vial. Add 0.2 ml hexamethyl disilazane, shake, add 0.2 ml trimethylchlorosilane and shake again. Place on a warm plate (about 80°) for 3-5 min. Check that white fumes are present indicating an excess of reagent.
Any suitable gas chromatograph equipped as follows:
Stationary phase: 3% OV-1
Carrier gas: Nitrogen
Temperature of injection port: 275°
Column temperature: 90° to 330° at 4-6°/min
Detector type: FID, temperature: 350°
Inject a 2.0 μl sample of TMS derivatives of polyols. The following sequence of peaks are recorded on the resultant chromatogram :Elution sequence of peaks Identity Description 1 Solvent Out of scale 2 Glycerol Single peak 3 Cyclic diglycerols Single peak 4 Diglycerols Single peak 5 Cyclic triglycerols Single peak 6 Triglycerols Single peak 7 Cyclic tetraglycerols Single peak 8 Tetraglycerols Multiple peaks 9 Pentaglycerols Single peak 10 Hexaglycerols Single peak 11 Heptaglycerols Single peak 12 Octaglycerols Single peak 13 Nonaglycerols
Barely discernible in the tail of peak 12
Measure each peak area by a suitable method.
% di-, tri- and tetraglycerols =
(Sum of corrected areas of peaks 3 to 8 x 100) / (Sum of corrected areas of peaks 3 to 13)
% polyglycerols equal to or greater than heptaglycerol =
(Sum of corrected areas of peaks 11 to 13 x 100) / (Sum of corrected areas of peaks 3 to 13)
The sample is hydrolysed. Fatty acids are removed by ion exchange in combination with hexane extraction. The components of the filtrate are separated by thin layer chromatography.
Reflux 1 g of sample with 15 ml of 0.5 N ethanolic potassium hydroxide for 1 h. Add 25 g of strong cation ion exchange resin (such as Amberlite I R 120, H-form), 50 ml of hexane and 25 ml of water. Stir the mixture for about 1 h. Filter off the resin and, after allowing the layers of the filtrate to separate, take the aqueous layer for TLC.
Spot 2 to 5 μl portions of the aqueous layer onto a silica gel G plate and also 2 μl of 5% solutions of glycerol, ethylene glycol and 1,2-propylene glycol.
Develop the chromatogram using chloroform:acetone:5 N ammonia (10:80:10) as the solvent system. After development, dry the plate in a stream of air until the water and ammonia have been removed.
Spray the plate with a solution of lead acetate (1% w/v in toluene) and heat the plate for 5 min at 110°. 1,2-Diols are revealed as white spots on a brown background.
The following are examples of Rf values that may be obtained:
PolyolRf Glycerol 0.35 Ethylene glycol 0.70 1,2-Propylene glycol 0.85
Propylene glycol esters of fatty acids are saponified with alcoholic potassium hydroxide and the fatty acids are removed by extraction. The aqueous polyol fraction is analyzed by gas-liquid chromatography for di- and tripropylene glycol.
Potassium hydroxide, ethanolic solution (56.1 g/l ethanol)
Sodium hydroxide solution (50% w/v in water)
Hydrochloric acid (1 + 1 by volume)
Standards: propylene glycol (1,2-propanediol) dipropylene glycol (1,1-oxydi-2-propanol) tripropylene glycol triethylene glycol
Preparation of polyols
Weigh about 50 g of sample, to the nearest 0.01 g, together with about 2 g to nearest 0.001 g, of triethylene glycol (as internal standard) into a 1-l saponification flask. Add 350 ml of ethanolic potassium hydroxide solution and reflux under an air condenser for 2 h with stirring. Transfer the contents of the flask quantitatively to an 800-ml beaker. Wash the flask and condenser with 200 ml of hot distilled water and evaporate the combined sample solution and washings to about 200 ml on a steam bath. Acidify the hot residue to pH 2 by the dropwise addition, with agitation, of hydrochloric acid (1 + 1). Transfer the hot mixture quantitatively to a 2-l separatory funnel with 200 ml of hexane and shake. Allow the layers to separate. Transfer the lower aqueous layer to a 500-ml separatory funnel and add 200 ml of fresh hexane. Shake, then allow the layers to separate. Draw off the lower aqueous layer into a 600-ml beaker and add the hexane phase to the original 2-l separatory funnel. Wash the 500-ml separator with two further 200-ml portions of hexane and add these to the 2-l separator. Wash the original 800-ml beaker with 100 ml of water and add to the hexane solution. Mix thoroughly and allow the layers to separate. Draw off the aqueous layer into the 600-ml beaker containing the aqueous fractions. Wash the 800-ml beaker once more with 100 ml of water and add the drained aqueous layer to the combined aqueous fractions. Adjust the pH of the combined aqueous solution to pH 7.0-7.05 (using pH-meter) with sodium hydroxide solution and evaporate to about 150 ml on a steam bath. Transfer quantitatively to a 250 ml round bottom flask and concentrate further to about 50 ml by distilling through a vertical Vigreux column to prevent the loss of low boiling glycols. Decant the concentrated polyols from the precipitated salts, through a filter funnel containing Whatman No. 1 paper, into a 100-ml volumetric flask. Wash the salts and flask twice with 20 ml of water and add to the volumetric flask via the filter funnel. Dilute to the mark with water, mix well and use for the GLC polyol analysis. If salts reprecipitate in the volumetric flask refilter before sampling.
Any suitable gas chromatograph equipped with:
Stationary phase: 15% Carbowax 20 M
Carrier gas: Helium
Temperature of injection port: 290°
Column temperature: 150° to 230° at 2°/min
Detector type: FID; temperature: 250°.
Prepare a reference solution of glycols in water by weighing the glycol standards, to the nearest 0.1 mg, into a 100 ml volumetric flask as follows:
- Propylene glycol: 1 g
- Dipropylene glycol: 0.01 g
- Tripropylene glycol: 0.005 g
- Triethylene glycol: 0.01 g
Make up to the mark with water and mix.
Inject aliquots of this reference standard solution and establish the sensitivity setting to yield measurable peaks. Similarly inject the prepared sample solution.
Measure each peak area by a suitable method, such as multiplying the peak height by the peak width at half the peak height, and calculate the % dimer and trimer in the sample as follows:
ADS is the peak area of dipropylene glycol (sample solution);
ADR is the peak area of dipropylene glycol (reference solution);
Ats is the peak area of tripropylene glycol (sample solution);
ATR is the peak area of tripropylene glycol (reference solution);
AIS is the peak area of triethylene glycol (sample solution);
Air is the peak area of triethylene glycol (reference solution);
W is the weight (g) of sample of propylene glycol esters of fatty acids;
Wdr is the weight (g) of dipropylene glycol in the reference solution;
Wtr is the weight (g) of tripropylene glycol in the reference solution;
WIS is the weight (g) of triethylene glycol added to the sample solution; and
Wir is the weight (g) of triethylene glycol in the reference solution.
Corrected for the actual content of polyol (e.g. Wdr is the weight of dipropylene glycol taken x % of assay).
(AOCS Methods Tl 1a-64 and Cd 3-25)
Weigh accurately about 20 g of the sample and subject to alkaline hydrolysis by re fluxing for 2 h with ethanolic potassium hydroxide TS containing a quantity of potassium hydroxide 100% in excess of the calculated amount required to saponify the sample completely. After hydrolysis, convert the ethanolic soap solution to an aqueous solution by the addition of water and evaporation of the alcohol on a steam bath. Acidify the hot aqueous soap solution with sulfuric acid to liberate the fatty acid. Extract the acid solution with 3 portions of petroleum ether to remove the fatty acid. Evaporate the petroleum ether extracts on a steam bath and dry the residue to constant weight under vacuum at 75° to recover the fatty acid. Multiply the weight of recovered fatty acid by 100/W to obtain the yield of fatty acid from a 100-g sample (where W is the exact weight of sample taken). The fatty acid can be identified by determination of the physical and chemical constants, e.g. the solidification temperature, or by gas-liquid chromatography.
Neutralize the aqueous polyol solution to pH 7 with potassium hydroxide. Evaporate the polyol solution to a moist residue on a steam bath and extract the polyol from the salts with 3 portions of hot absolute ethanol. Evaporate off the alcohol on a steam bath and dry the residue to constant weight under vacuum at 75° to yield the polyol moiety of the sample. Multiply the weight of recovered polyol by 100/W to obtain the yield of polyols from a 100-g sample (where W is the exact weight of sample taken).
Saponification value is defined as the number of mg of potassium hydroxide required to neutralize the free acids and saponify the esters in 1 g of test substance.
Melt the sample, if necessary, and filter it through a dry filter paper to remove any traces of moisture. Unless otherwise directed, weigh accurately into a 250-ml flask a sample of such size (usually about 4-5 g) that the titration of the sample solution after saponification will require between 45 and 55% of the volume of 0.5 N hydrochloric acid required for the blank. Add 50.0 ml of ethanolic potassium hydroxide TS from a pipet and allow the pipet to drain for a definite period of time. Prepare and conduct blank determinations simultaneously with the sample and similar in all respects. Connect an air condenser to each flask and boil gently but steadily, with occasional mixing, until the sample is completely saponified. (This usually requires about 1 h for normal samples). After the flasks and condensers have cooled somewhat but not sufficiently for the contents to gel, wash down the inside of the condensers with a few ml of distilled water. Disconnect the condensers, add about 1 ml of phenolphthalein TS to reach flask, and titrate with 0.5 N hydrochloric acid until the pink colour has just disappeared.
A is ml of HCl required for the titration of the blank;
B is ml of HCl required for the titration of the sample;
W is the weight of sample in g; and
N is normality of the HCl.
Sorbitan esters may be assayed by alkaline saponification followed by recovery of the polyol and determination of the isosorbide content by gas-liquid chromatography.
Saponification and recovery of the polyol
Weigh accurately about 25 g of the sample into a 500-ml round-bottomed boiling flask. Add 250 ml of ethanol and a quantity of potassium hydroxide 100% in excess of the calculated amount required for saponification (approximately 7.5 g). Boil the mixture for 2 h under reflux. Transfer the saponification mixture to an 800-ml beaker. Rinse the flask with about 100 ml of water and add to the mixture. Place the beaker on a steam bath to evaporate the alcohol. Add water occasionally to replace the ethanol. When the odour of ethanol can no longer be detected, adjust the volume of the soap solution to approximately 250 ml with hot water.
Acidify the hot soap solution with stirring using sufficient 1:1 sulfuric acid to provide a 10% excess. Heat and stir the mixture until the fatty acid layer separates. Transfer the hot mixture to a 500-ml separating funnel using hot water to rinse the beaker. Cool the contents of the funnel and extract three times with 100-ml portions of petroleum ether. Combine the petroleum ether extracts in a second funnel and wash once with 100 ml of water. Combine the water wash with the aqueous phase in an 800-ml beaker.
Neutralize the polyol solution with 10% aqueous potassium hydroxide solution to pH 7 using a pH meter. Place the beaker in a steam bath and evaporate the solvent to incipient dryness. Extract the residue four times with 150-ml portions of boiling absolute ethanol. Filter the combined extracts into a 1 -l suction flask through a 10-cm Buchner funnel containing a 1 -3 cm bed of silicagel. Wash the funnel with absolute ethanol. Transfer the filtrate and washings to a 1,000-ml volumetric flask. Cool to room temperature and dilute to volume with ethanol. Use this as the sample solution for gas-liquid chromatography.
The experimental operating conditions for the isosorbide analyses are not critical, suitable conditions are listed below . Minor fluctuations in temperature and gas flow rate do not affect resolution or analytical results.
Stationary phase: 15% Carbowax 20 M
Carrier gas: Argon
Temperature of injection port: 295°
Column temperature: 195°
Detector type: FID; 250°
The isosorbide content of an aliquot of the recovered polyol solution is estimated directly from a calibration curve prepared from a standard sorbitan ester or by multiplying the observed peak area by the slope of the curve (μg of isosorbide per unit area).
I is μg of isosorbide found in the aliquot of recovered polyol solution by gas chromatography;
W is g of sorbitan ester taken for analysis; and
f is fractional isosorbide yield from standard sorbitan esters (see Note below).
Note: A known sample of sorbitan ester is treated as described under Saponification and recovery of the polyol above. Suitable aliquots of the solution are subjected to the gas chromato graphic procedure. The fractional yield of isosorbide is calculated from the weight of sample corrected to a dry, fatty-acid-free basis. The procedure is estimated to have an accuracy of 5%.
Note: All reagents used in this test should be reagent grade: water should be of high purity, and gases must be high-purity grade.
The Dohrmann Microcoulometric Titrating System (MCTS-30), or its equivalent as shown in the figure below should be used. It consists of a constant rate injector (A), a pyrolysis furnace (B), a quartz pyrolysis tube (C), a granular tin scrubber (D), a titration cell (E), and a microcoulometer with a digital readout (F).
Granular-Tin Scrubber: Place 5 g of 20/30 mesh granular reagent grade tin between quartz-wool plugs in an elongated 18/8-12/5 standard-taper adaptor which connects the pyrolysis tube and the titration cell.
Microcoulometer: Must have variable attenuation, gain control, and be capable of measuring the potential of the sensing-reference electrode pair, and comparing this potential with a bias potential, amplifying the potential difference, and applying the amplified difference to the working-auxiliary electrode pair so as to generate a titrant. Also the microcoulometer output voltage signal must be proportional to the generating current.
Pyrolysis Furnace: The sample should be pyrolyzed in an electric furnace having at least two separate and independently controlled temperature zones, the first being an inlet section that can maintain a temperature sufficient to volatilize the entire organic sample. The second zone shall be a pyrolysis section that can maintain a temperature sufficient to pyrolyze the organic matrix and oxidize all the organically bound sulfur. A third outlet temperature zone is optional.
Pyrolysis Tube: Must be fabricated from quartz and constructed in such a way that a sample, which is vaporized completely in the inlet section, is swept into the pyrolysis zone by an inert gas where it mixes with oxygen and is burned. The inlet end of the tube shall hold a septum for syringe entry of the sample and side arms for the introduction of oxygen and inert gases. The center, or pyrolysis section, should be of sufficient volume to ensure complete pyrolysis of the sample.
Sampling Syringe: A microliter syringe of 10-μl capacity capable of accurately delivering 1 to 10 μl of sample into the pyrolysis tube. Three-inch x 24-gauge needles are recommended to reach the inlet zone of the pyrolysis furnace.
Titration Cell: Must contain a sensor-reference pair of electrodes to detect changes in triiodide ion concentration, a generator anode-cathode pair of electrodes to maintain constant triiodide ion concentration, and an inlet for a gaseous sample from the pyrolysis tube. The sensor electrode shall be platinum foil and the reference electrode platinum wire in saturated triiodide half cell. The generator anode and cathode half-cell shall also be placed on a magnetic stirrer.
Preparation of Apparatus
Carefully insert the quartz pyrolysis tube into the furnace, attach the tin scrubber, and connect the reactant and carrier-gas lines. Add the Cell Electrolyte Solution (see below) to the titration cell and flush the cell several times. Maintain an electrolyte level of 3.8 cm (1.5 in.) above the platinum electrodes. Place the titration cell on a magnetic stirrer and connect the cell inlet to the tin scrubber outlet. Position the platinum foil electrodes (mounted on the movable cell head) so that the gas-inlet flow is parallel to the electrodes with the generator anode adjacent to the generator cathode. Assemble and connect the coulometer in accordance with the manufacturer's instructions. Double-wrap the adaptor containing the tin scrubber with heating tape and turn the heating tape on. Adjust the flow of the gases, the pyrolysis furnace temperature, the titration cell, and the coulometer to the desired operating conditions. Typical operating conditions are as follows:
Reagent gas flow (oxygen): 200 cm3/min
Carrier-gas flow (Ar, He): 400 cm3/min
Inlet zone: 700° (maximum)
Pyrolysis zone: 800 - 1000°
Outlet zone: 800° (maximum)
Tin-Scrubber flow rate: 200 cm3/min
Titration cell: Stirrer speed set to produce slight vortex
Bias voltage: 160 mV
Constant Rate Injector: 0.25 μl/sec
The tin scrubber must be conditioned to sulfur, nitrogen, and chlorine before quantitative analysis can be achieved. A solution containing 10 mg/kg butyl sulfide, 100 mg/kg pyridine, and 200 mg/kg chlorobenzene in isoctane has proven an effective conditioning agent. With a fresh scrubber installed and heated, two 30-μl samples of this conditioning agent injected at a flow rate of 0.5 μl/sec produces a steady increasing response, with final conditioning indicated by a constant reading from the offset during the second injection.
- Argon or Helium, (Argon preferred) High-purity grade: two-stage regulators must be used.
- Cell Electrolyte Solution: dissolve 0.5 g of potassium iodide and 0.6 g of sodium azide in 500 ml of high-purity water, add 5 ml of glacial acetic acid and dilute to 1 L. Store in a dark bottle or in a dark place and prepare fresh at least every 3 months.
- Oxygen: high-purity grade.
- Iodine: resublimed, 20 mesh or less.
- Sulfur Standard (approximately 100 mg/kg): weigh accurately 0.1569 g of n-butyl sulfide, into a tared 500-ml volumetric flask. Dilute to the mark with isooctane and reweigh. Calculate the sulfur concentration (S), in percent, by the formula:
Ws = weight of n-butyl sulfide, and
Wc = weight of the solution.
Prepare a calibration standard (approximately 5 mg/kg) by pipetting 5 ml of Sulfur Standard into a 10-ml volumetric flask and diluting to volume with isooctane. Fill and clamp the syringe onto the constant rate injector, push the sliding carriage forward to penetrate the septum with the needle, and zero the meter in case of long-term drift in the automatic baseline zero circuitry. Switch S1 automatically starts the stepper-motor syringe drive and initiates the analysis cycle. At 2.5 min (after setting switch S1) set the digital meter with the scan potentiometer to correspond to the sulfur content of the known standard to the nearest 0.01 mg/kg. At the 3-min point, the number displayed on the meter stops, the plunger drive block is retracted to its original position, as preset by switch S2 and a baseline re-equilibration period equal to the injection period must be allowed before a new sample may be injected. Repeat the Calibration step a total of at least four times.
Rinse the syringe several times with sample: then fill it, clamp it onto the constant-rate injector push the sliding carriage forward to penetrate the septum with the needle, and zero the meter. Turn on switch S1 to start the stepper-motor syringe drive automatically and initiate the analysis cycle. After the 3-min hold point, the number displayed on the meter corresponds to the sulfur content of the injected sample.
(ASTM D 445 Adapted, with permission, from the Annual Book of ASTM Standards, copyright American Society for Testing and Materials, 100 Harbor Drive, West Conshohocken, PA 19428. Copies of the complete ASTM standard may be purchased direct from ASTM)
Use a viscometer of the glass capillary type, calibrated and capable of measuring kinematic viscosity with a repeatability exceeding 0.35 % only in one case in twenty. Immerse the viscometer in a liquid bath at the temperature required for the test ±0.1° ensuring that at no time of the measurement will any portion of the sample in the viscometer be less than 20 mm below the surface of the bath liquid or less than 20 mm above the bottom of the bath. Charge the viscometer with sample in the manner dictated by the design of the instrument. Allow the sample to remain in the bath for about 30 min. Where the design of the viscometer requires it, adjust the volume of sample to the mark. Use pressure to adjust the head level of the sample to a position in the capillary arm of the instrument about 5 mm ahead of the first mark. With the sample flowing freely, measure, in seconds (±0.2 s), the time required for the meniscus to pass from the first to the second timing mark. If the time is less than 200 s, select a viscometer with a capillary of smaller diameter and repeat the operation. Make a second measurement of the flow time. If two measurements agree within 0.2 %, use the average for calculating the kinematic viscosity. If the measurements do not agree, repeat the determination after thorough cleaning and drying the viscometer.
C is the calibration constant of the viscometer (mm2/s2), and
t is the flow time (s)
Dissolve about 10 g of sample, accurately weighed, in 50 ml of ethanol, previously neutralized to phenolphthalein TS with 0.1 N sodium hydroxide. Add 1 ml of phenolphthalein TS and titrate with 0.1 N sodium hydroxide until the solution remains faintly pink after shaking for 10 sec, unless otherwise directed. Calculate the Acid Value (AV) by the formula:
S is the number of ml of 0.1 N sodium hydroxide consumed in the titration of the sample, and W is the weight of the sample in g.
For Gas Chromatographic analysis procedures see Gas Chromatography in the Section on Analytical Techniques.
See the Section on Appearance and Physical Properties, under General Methods.
Transfer an accurately weighed quantity of the sample specified in the monograph into a 125-ml Erlenmeyer flask containing a few boiling stones. Add to this flask, and, simultaneously, to a similar flask for a blank test, 25.0 ml of 0.5 N ethanolic potassium hydroxide. Connect each flask to a reflux condenser, and heat the mixtures on a steam bath for exactly 1 h, unless otherwise directed in the monograph. Allow the mixtures to cool, add 10 drops of phenolphthalein TS, to each flask, and titrate the excess alkali in each flask with 0.5 N hydrochloric acid. Calculate the percentage of Ester (E) in the sample by the formula:
b is the number of ml of 0.5 N hydrochloric acid consumed in the titration of the blank;
S is the number of ml of 0.5 N hydrochloric acid consumed in the titration of the sample; and
e is the equivalence factor given in the monograph, and W = the weight of the sample in mg.
See the Section on Appearance and Physical Properties, under General Methods.
See the Section on Appearance and Physical Properties, under General Methods.
Unless otherwise stated in the specification, transfer a 1 ml sample into a calibrated 10-ml glass-stoppered cylinder graduated in 0.1-ml subdivisions, and add slowly, in small portions, ethanol, the concentration and quantity of which are specified in the monograph. Maintain the temperature at 20°. A clear solution free from foreign matter should be obtained.
See the Section on Appearance and Physical Properties, under General Methods.
See the Section on Appearance and Physical Properties, under General Methods.
Note: This determination is done in connection with Water Content (Loss on Drying) for food colours and the result is included in that calculation.
- Potentiometric titration apparatus
- Silver indicator electrode
- Glass body calomel reference electrode or calomel reference electrode with potassium sulfate bridge
- Nitric acid, 1.5 N, reagent grade
- Silver nitrate, 0.1 N, standard solution
Accurately weigh 0.5 - 1.0 g of the colour sample (WS), dissolve in 100 ml of water, and acidify with 5 ml of 1.5 N nitric acid. Place the silver and glass body calomel electrodes in the colour solution. If only a standard calomel reference electrode is available, connect it to the solution by means of the saturated potassium sulfate bridge. (Use of a glass body electrode as the reference electrode eliminates the need for the potassium sulfate bridge; this simplifies the apparatus considerably, and the glass body electrode is sufficiently constant to be used as a reference for this type of titration.)
Determine the chloride content of the solution by titration with the 0.1 N silver nitrate. (Each ml of 0.1 N silver nitrate is equivalent to 0.00585 g of sodium chloride.)
Calculate the chloride content of the sample as percent sodium chloride using the following equation:
- Oven, 0 - 200° range
- Hot plate
- Crucible, fitted with glass fiber disk
- Vacuum flask
- Source of vacuum
- Chloroform, reagent grade
Accurately weigh the quantity of sample indicated in each specification monograph (W1) into a 250-ml beaker. Mix with 100 ml of chloroform (b.p. 61.1°). Stir and heat to boiling on the hot plate in a fume hood. Filter the hot solution through a weighed crucible (W2). Transfer the residue in the beaker to the crucible with chloroform. Wash the residue in the crucible with 10-ml portions of chloroform until the washings are colourless. Place the crucible in the oven at 100 - 150° for 3 h; cool the crucible in the desiccator. Weigh the cooled crucible (W3).
The percent chloroform-insoluble matter in the sample is 100 x (W3 - W2) / W1.
Many of the colours used by food manufacturers are mixtures of colouring matters of the type described in the specification monographs, and some of the mixtures contain added diluents. A simple test to establish whether a powdered sample of colouring matter is a single colouring matter or a mixture of colouring matters is to sprinkle a very small quantity of the powder into each of two beakers, one containing water and the other containing concentrated sulfuric acid. Under these conditions, the specks of individual colouring matters can easily be seen as they dissolve; the test is surprisingly sensitive.
The positive identification of individual food colours is often quite difficult. A large number are the sodium salts of sulfonic acids, which have no precise melting point. In addition, synthesized colours usually contain subsidiary colouring matters, while colouring matters extracted from natural sources generally are mixtures of colours themselves. Identification, therefore, is best achieved by comparison of the observed properties with the properties of authentic commercial samples. The principle techniques in use are chromatography and spectrophotometry. Frequently, both are required, because the presence of subsidiary colouring matters might affect the observed spectra so that positive identification of the principal colour component cannot be made. For this reason, separation of the individual colouring matters by column, paper, or thin layer chromatography is advisable before attempting additional identification by spectrophotometry.
Subsidiary colouring matters are defined as those colouring matters that are produced during the manufacturing process in addition to the principal named colouring matter(s). Paper chromatography has been used for many years for identifying subsidiary colouring matters in water-soluble food colours. The assumption is generally made that spectrophotometric absorbances of subsidiary colouring matters are similar to that of the main colouring matter. Accordingly, standards of individual subsidiary colour matters are not required. The presence of colouring matters other than the principal and subsidiary colouring matters is usually detected on the chromatograms used to determine subsidiary colouring matters. Interpretation of the chromatograms for these colour impurities usually requires additional information.
High-performance liquid chromatography (HPLC) has been used successfully to separate, identify, and quantitate the subsidiary colouring matter contents of various food colours. Standards for individual subsidiary colouring matters are needed for this method. However, the specification limits in the monographs are, unless otherwise stated, linked to the paper chromatographic method and the conditions are provided under "Tests" in the specification.
Paper and thin layer chromatography are often useful in identification of colouring matters and do not require expensive equipment. But it must be kept in mind that the Rf-value of a substance is generally an unreliable quantity because many factors, most of which are beyond the analyst's control, can have a major influence on the Rf-values. These factors include: composition and age of the solvent mixture, concentration of solvent vapour in the atmosphere, quality of the chromatography paper, machine direction of commercially made paper, kind and quality of subsidiary colouring matters, concentration, pH-value of the solution, and temperature. For this reason, comparative chromatography using reference colours should always be used. By simultaneously running several colouring matters of similar concentration a number of these factors are eliminated.
Coincidence of migration distances with a single solvent system should be looked upon only as one criterion of identity and further tests should be made to confirm the finding.
The following table contains examples of the Rf-values that may be expected when 1% aqueous solutions of various colouring matters are subjected to thin layer chromatography on Silica Gel G in the ten solvent systems listed below. The compositions of the solvent systems, all of which must be freshly prepared, are:
Solvent System Number
- iso-Propanol:ammonia (sp.gr. 0.880):water (7:2:1)
- iso-Butanol:ethanol:water:ammonia (sp.gr. 0.880) (10:20:10:1)
- Saturated aqueous potassium nitrate solution
- Phenol:water (4:1, w/v)
- Hydrochloric acid (sp.gr. 1.18):water (23:77)
- Trisodium citrate:ammonia (sp.gr. 0.880):water (2 g:15 ml:85 ml)
- Acetone:2-butanone :ammonia (sp.gr. 0.880):water (60:140:1:60)
- n-Butanol:ethanol:pyridine:water (2:1:1:2)
- iso-Propanol:ammonia (sp.gr. 0.880) (4:1)
- n-Butanol:acetic acid (glacial):water (10:5:6)
Rf Values of Some Water-Soluble Colours (This Table does not indicate the acceptability of the listed colours for food use.) Note: Numbers 1 to 10 refer to solvent systems (see above).REDS C.I. No. INS No. 1 2 3 4 5 6 7 8 9 10 Ponceau 4R or Cochineal Red A 16255 124 0.66 (0.85) 0.75 0.88 0.03 0.95 1.00 0.60 0.90 0.11 0.52 0.00-0.57 Carmosine or Azorubine 14720 122 0.65 (0.77) 0.81 0.00-0.42 0.16 0.00 (0.00-0.32) 1.00 0.65 0.88 0.34 (0.46) 0.63 (0.11-0.70) Amaranth 16185 123 0.62 (0.48, 0.76) 0.75 (0.83) 1.00 (0.00-1.00) 0.04 (0.16) 1.00 1.00 0.40 (0.64, 0.66) 0.90 0.10 (0.41) 0.39, 0.67 Erythrosine RS 45430 127 0.85 (0.68, 0.79) 0.91 (0.86, 0.74, 0.81) 0.00,0.10 0.00-0.90 (0.41) 0.00 0.00-0.95 0.64, 0.66 (0.58) 0.89 0.66 (0.57, 0.43) 1.00 Red2G 18050 — 0.68 0.680 0.37 0.12 0.00-0.71 0.90 0.64 0.90 0.36 0.68 ORANGES Orange G 16230 — 0.71 (0.67, 0.88) 0.80 (0.75) 0.64 (1.00, 0.35) 0.23, 0.15,0.04 0.73 1.00 0.64 (0.62, 0.50, 0.67) 0.91 0.36 (0.32, 0.17) 0.69 (0.46, 0.82) Orange RN 15970 — 0.83 (0.62) 0.88 (0.78) 0.00 (0.00-0.42) 0.42 (0.13) 0.13 (0.38) 0.76 (1.00) 0.68 (0.65) 0.92 0.64 (0.29) 0.82,0.71 Sunset Yellow FCF or Orange Yellow S 15985 110 0.75 (0.68) 0.82 (0.74) 1.00 (0.00-1.00) 0.17, 0.03 1.00 1.00 0.65 (0.48) 0.90 0.34 (0.10, 0.22) 0.67 (0.46) YELLOWS Tartrazine 19140 102 0.66 0.77 0.46-1.00 0.08 0.930 0.930 0.52 0.93 0.14 0.50 Yellow 2G 18965 — 0.63 0.80 0.77 0.21 0.74 0.620 0.62 0.92 0.21 0.75 Quinoline Yellow 47005 104 0.83, 0.88 0.88 (0.82) 0.00-1.00 0.65 (0.21) 0.26-1.00, 0.00-0.38 0.95 (0.35) 0.54 (0.68) 0.88 0.00- 0.31,0.64 0.11-0.75 (0.83) Fast Yellow AB 13015 — 0.77 0.81 0.560 0.14 0.97 0.560 0.56 0.93 0.36 0.66 GREENS, BLUES, AND VIOLETS Green S or Acid Brilliant- Green BS or Lissamine Green 44090 142 0.44 (0.52, 0.68, 0.74) 0.61 (0.67, 0.75, 0.81,0.84) 0.49 (0.24) 0.53 (0.05, 0.36, 1.00) 0.29 (0.43) 1.00 0.46 (0.56, 0.71) 0.75 (0.89, 0.92) 0.07 0.55 Indigo Carmine or Indigotin 73015 132 0.56 (0.70) 0.50-0.76 (0.78) 0.00 (0.05, 0.90, 1.00) 0.09,0.18 (0.52) 0.92 0.94 0.66 (0.71, 0.73) 0.89, 0.84 0.37 (0.00- 0.34) 0.00-0.63 Indanthrene Blue or Solanthrene Blue RS or Anthragen Blue 69800 — 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Brilliant Blue FCF 42090 — 0.64 (0.73) 0.78 0.05 0.45 (0.68) 0.10 0.00-1.00 0.61 (0.68) 0.88 0.30 (0.49, 0.00-0.23)
0.53 (0.64)Patent Blue V 42051 131 0.34- 0.60 0.68 0.05 0.55 0.15 0.95 0.69 (0.72) 0.84 (0.92) 0.00-0.10 0.59 Violet 6B 42640 — 0.73 (0.67, 0.91) 0.80 (0.72) 0.00 (0.00-0.48) 0.62 (0.51-1.00) 0.00-0.37 0.00-1.00 0.67 0.89 (0.62) 0.37, 0.45, 0.70, 0.76 0.64 (0.71) Methyl Violet 42535 — 0.91 (0.80) 0.56, 0.81, 0.90 0.00 (0.00-0.31) 0.79-1.00 0.00-0.80 0.00 (0.00-0.53) 0.00-0.68 (0.11,0.28, 0.53, 1.00) 0.11 (0.90) 0.94, 0.87 (0.00-0.83) 0.75, 0.79 (0.00-0.70) BROWNS AND BLACKS Brown FK — — 0.78, 0.71, 0.66 0.79, 0.86 1.00 (0.00-1.00) 0.69, 0.27, 0.15,0.00 0.00-0.77 1.00 0.59, 0.64, 0.53 (0.37) 0.93 0.34 (0.26, 0.53) 0.00-0.73 Chocolate Brown FB — — 0.00- 0.69 0.00-0.75 0.00-0.82 0.00 (0.00-0.23) 0.00-1.00 0.00-1.00 0.36, 0.51, 0.62 0.87 0.00-0.38 0.00-0.75 Chocolate Brown HT 20285 — 0.00- 0.63 0.74 0.00-1.00 0.00 (0.00-0.16) 0.00-1.00 0.00-1.00 0.34, 0.43, 0.62 0.88 0.00-0.32 0.00-0.73 Black PN or Brilliant-Black BN 28440 151 0.66 (0.47) 0.75 0.00 (0.00-1.00) 0.00 1.00 1.00 0.38 (0.61) 0.85 0.05 0.00-0.43 Black 7984 27755 152 0.62 0.75 1.00 01.00 1.00 1.00 0.38 (0.61) 0.85 0.09 0.00-0.45
Key: C.I. No.: Colour Index Number; Figures in parentheses, ( ), indicate subsidiary spots of lower intensity; "0.xx-0.yy": Streak between spots. Table from Pearson, D. (1973) J. Assoc. Public Anal., 11, 137-138. Reprinted from Environmental Carcinogens - Selected Methods of Analysis, Vol. 4 - Some Aromatic Amines and Azo Dyes in the General and Industrial Environment (IARC Publications No. 40), International Agency for Research on Cancer, Lyon, 1981. Note: E numbers in the original table have been replaced by the appropriate International Numbering System (INS) numbers of the Codex Alimentarius Commission.
Assessment of the colour shade should be made while the chromatograms are still moist with solvent and then again after drying. The shade should be assessed in both incident and transmitted daylight as well as under ultraviolet (UV) light, in which many colours show characteristic colour changes. Furthermore, UV light can often be used to identify the presence of colourless fluorescent impurities. If possible, use two UV emitters which yield different wave lengths; one lamp should emit around 250 nm.
Tests with acids, alkalis and other suitable reagents, in order to confirm the results, should be made. All tests may be carried out with fine capillary pipettes on each colour spot.
The following requirements should be met when identifying the colours in colouring matters by comparing to reference colours:
- equal migration distances in several solvent systems;
- equal shade in daylight and ultraviolet light; and
- equal colour changes with reagents.
Spectrophotometric methods of examination are among the most useful means of identification of colours. The UV, visible, and infrared regions of the electromagnetic spectrum are all employed.
The visible region of the spectrum is ordinarily examined as the first step in attempting to identify an unknown colour. Many colours show characteristic absorption bands in the visible region. Spectra in the UV region may also be of use and should be obtained together with the visible spectrum, if possible.
In the application of UV-visible spectrophotometry, spectra should always be obtained in more than one solvent, or if in a single solvent, under various conditions. Spectra of aqueous solutions should be obtained under neutral (buffered with ammonium acetate), acid (0.1 N hydrochloric acid), and alkaline (0.1 N sodium hydroxide) conditions.
A UV-visible absorption spectrum is ordinarily displayed as a plot of absorbance vs. wavelength. In addition to the wavelength maximum, the most characteristic and useful features of the absorption spectrum can be the "shoulders" or inflection points on the spectral curve. These features often make it possible to distinguish between two or more colouring matters that have absorption maxima at the same wavelength. Many colours can be definitively characterized by observing the extent to which the absorption maxima and other features of the absorption curve are changed by variation in pH or by other changes in the solvent.
Infrared absorption spectra offer another useful means of identification of compounds. An example of their use is in distinguishing Sunset Yellow and Orange GGN. Whereas the UV-visible absorption spectra of these colours are nearly identical, their infrared spectra are quite different in the region of the spectrum in which the sulfonic acid groups absorb strongly.
Infrared spectra of substances can be obtained using various sample preparations; the more commonly used are:
- solutions of the material in suitable solvents;
- suspensions of the material in a suitable liquid;
- potassium bromide pellets (in this technique, a small amount of the colouring matter, usually from 1 to 3 mg, is thoroughly mixed with pure, dry potassium bromide and the mixture is transferred to a suitable die and pressed into a thin pellet by exerting a pressure of 700 to 1,400 kg/cm2).
Spectra are ordinarily displayed as % transmittance vs. wavenumber (cm-1). The salient features of the spectra are the intensities of the absorption peaks, and their shapes.
Detailed discussion of the infrared absorption technique and interpretation of infrared spectra are beyond the scope of this volume. It must be pointed out, however, that some difficulties exist in the practical use of this technique for identifying colours.
The crystal structure or other physical state of the sample may affect the spectra obtained from suspensions or potassium bromide pellets. It is necessary to make certain that the unknown material has been treated in exactly the same manner as was the standard or known sample.
Care must be taken to ensure that absorption bands due to contaminants are identified. All materials to be tested should be free from water or other solvent before an infrared spectrum is obtained because water and all organic solvents absorb infrared radiation. Water-soluble colouring matters can often be prepared for analysis by dissolving samples in water, adding a little acetic acid, evaporating to apparent dryness, and then drying at about 100° to remove the residual water. Infrared spectra should be obtained of the dried solids, as well as blanks.
Other identification techniques
Sometimes chromatographic and spectrophotometric techniques will fail to provide positive identification of colouring matters. In such cases, the problem can often be solved by reducing the colouring matter or otherwise degrading it and identifying the resulting products. This technique is particularly applicable to identifying azo colours. The amine compounds resulting from the reduction can frequently be readily identified by chromatographic and spectrophotometric techniques.
Many other techniques have been applied to the identification of colouring matters. For example, many pigments have a well defined crystalline structure and can be identified by their X-ray diffraction patterns or by X-ray crystallography. Some colouring matters can be converted to crystalline derivatives and similarly identified. The descriptions of these and other available techniques are beyond the scope of this volume.
Total colouring matters content
Two general methods are used for determination of total colouring matters: 'Colouring Matters Content by Spectrophotometry' and 'Colouring Matters Content by Titration with Titanous Chloride.'
When using the spectrophotometric method, the analyst should take into account the accuracy and precision of the spectrophotometer used for the analysis. All colours present in the sample that absorb in the same region as that of the main colour will contribute to the absorbance figure used to calculate the results; subsidiary colouring matters of markedly different hue will not be accounted for by this method. This method uses accepted absorptivity figures obtained from purified standard colours for calculating the total colouring matters content.
The titanous chloride reduction method assumes that isomers and subsidiary colouring matters have the same titanous chloride equivalent as the main colouring matter.
Three experimental procedures are described. Procedure 1 is used for water-soluble colouring matters. Procedure 2 is used for organic solvent-soluble colouring matters, especially the synthetic carotenoids. (The solutions prepared in Procedure 2 are used in the identification tests for the carotenoids.) Procedure 3 is used for lakes.
The absorbance of a solution of the colouring matter is determined at its wavelength of maximum absorption and the total colouring matters content is calculated using a standard absorptivity value quoted in the specification monograph.
- UV-visible range spectrophotometer capable of accurate (± 1% or better) measurement of absorbance in the region of 350 - 700 nm with an effective slit width of 10 nm or less
- Spectrophotometer cells, 1 cm path length
Accurately weigh 0.25 g (± 0.02 g) of the sample (W). Transfer to a 1-liter volumetric flask. Add freshly distilled water or the solvent prescribed in the specification monograph and swirl to dissolve. Make up to volume and mix. Dilute to a solution of suitable strength according to the details given in the specification monograph. Measure the absorbance (A) at the wavelength of maximum absorption in a 1 cm cell, using water or the prescribed solvent as the blank.
Calculate the total colouring matters content of the sample using either of the following equations:
A is the absorbance of the sample solution at the wavelength of maximum absorption;
A1 %1 cm is the specific absorbance of the standard indicated in the specification monograph;
a is the absorptivity of the standard in liter/(g-cm); and
F is the dilution factor (Volume diluted / Volume measured).
- Chloroform, reagent grade, acid free
- Cyclohexane, reagent grade
Accurately weigh 0.08 g (± 0.01 g) of the sample (W) into a 100-ml volumetric flask (V1). Add 20 ml of chloroform and dissolve by swirling briefly. Make sure that the solution is clear. Make up to volume with cyclohexane and mix. Pipet 5.0 ml of the solution (v1) into a second 100-ml volumetric flask (V2) and make up to volume with cyclohexane. Pipet 5.0 ml of this diluted solution (v2) into the final 100-ml volumetric flask (V3) and make up to volume with cyclohexane. Measure the absorbance (A) of the twice-diluted solution at the wavelength of maximum absorption in a 1 cm cell, using cyclohexane as the blank.
Perform this procedure promptly, avoiding exposure to air insofar as possible and undertaking all operations in the absence of direct sunlight.
Calculate the total colouring matters content of the sample using either of the following equations:
A is absorbance of the sample solution at the wavelength of maximum absorption;
A1%1 cm is the specific absorbance of the standard indicated in the specification monograph;
a is the absorptivity of the standard in liter/(g.cm)
V1, V2, and V3 are the volumes of the three volumetric flasks (each 100 ml);
v1 and v2 are the volumes of the two pipets (each 5 ml);
a is absorptivity of the standard in liter/(g-cm); and
10-3 is the correction factor for a in ml/liter.
- Potassium dihydrogen phosphate, reagent grade
- Sodium hydroxide, reagent grade
- Phosphoric acid, reagent grade
- Hydrochloric acid, reagent grade
Prepare pH 7 phosphate buffer as follows: Weigh 13.61 g of potassium dihydrogen phosphate into a 2000-ml beaker, dissolve in 200 ml of water, and dilute to 1,000 ml. Add about 90 ml of 1 N sodium hydroxide. Determine the pH using a pH-meter and adjust the pH to 7.0 using 0.1 N sodium hydroxide or diluted phosphoric acid.
Accurately weigh a quantity of lake which will give an absorbance approximately equal to that of the parent colour when the latter is tested according to Procedure 1, above. Transfer to a 250- ml beaker containing 10 ml hydrochloric acid previously diluted with water to approximately 50 ml. Heat with stirring to dissolve the lake, then cool to ambient temperature. Transfer to a 1-liter volumetric flask, make up to volume with pH 7 phosphate buffer, and mix. Proceed as detailed in Procedure 1, above, and in the specification monograph, using pH 7 phosphate buffer as the spectrophotometric blank.
Titanous chloride (titanium trichloride, TiCl3) reduces the colouring matter to yield titratable reduction products. The method assumes that isomers and subsidiary colouring matters have the same titanium trichloride equivalent as the main colouring matter.
- Titration apparatus (See and below):
- Bottle (borosilicate glass) for titrant (may be up to 5 liter volume, as needed), with 29/42 ground glass center neck (for burette), side arm for inlet gas sparge, side arm stopcock (for gas outlet), and side arm with glass stopper for refilling bottle (Note: bottle may need to be custom-made)
- Digital burette, 25 ml - Brinkmann Digital Burette II ™ or equivalent
- 500-ml conical flasks, sealable with No. 10 rubber stoppers
- Overhead stirrer
- Stopper assembly - No. 10 rubber stopper with five holes for accommodating
- the stirrer rod, burette delivery tip, argon source, gas outlet tubing, and 10 ml
- Glass rod stopper for the pipet inlet in the stopper assembly o Tubing, glass and flexible plastic, for connections
- Hot plate
- Titanium trichloride (20% in HCl), reagent grade
- Hydrochloric acid, reagent grade
- Ferrous ammonium sulfate [Fe(NH4)2(SO4)2.6H2O], reagent grade
- Sulfuric acid, reagent grade
- Potassium dichromate, 0.100 N standard solution
- Ammonium thiocyanate, reagent grade
- Sodium citrate, reagent grade
- Sodium hydrogen tartrate, reagent grade
- Boiling chips
- Argon, UHP compressed gas (Carbon dioxide from a Kipp apparatus may be also be used, but is much less convenient; compressed nitrogen gas may be used provided residual oxygen is removed.)
Procedure for Colouring Matters
Note: A water bubbler should be placed in line between the argon source and the titration apparatus.
Preparation of 0.1 N titanium trichloride
Measure 800 ml of water for each liter of solution required into a beaker of appropriate size. On a hot plate, boil the water vigorously for 1 min, cover with a watch glass, and allow to cool to room temperature. In a fume cupboard, using graduated cylinders, add 90 ml of hydrochloric acid, stir, and add 100 ml of 20% titanium trichloride solution, for each liter of solution required. (Avoid transferring any white precipitate from the titanium trichloride reagent bottle.) Mix the solution and transfer to the titrant bottle. Attach the burette and connect the argon. Pass argon through the solution for 1 -2 h with the sidearm stopcock on the bottle open to maintain ambient pressure. While maintaining a slow flow of gas, draw up titrant into the burette. Drain the burette, discard the titrant, and refill. Drain and refill the burette two more times. Stop the gas flow, close the sidearm stopcock, and store the solution for at least 72 h before use.
Apparatus for titanium chloride titrations
Standardization of 0.1 N titanium trichloride
Drain and refill the burette with 0.1 N titanium trichloride. Use within 1 h. Weigh 3.0 g (± 0.2 g) of ferrous ammonium sulfate into a 500-ml conical flask. Add 200 ml of water. Using a graduated cylinder, add 25 ml of 10 N sulfuric acid. Pipet 20 ml of standard 0.100 N potassium dichromate into the flask. Swirl to mix. Connect the flask securely to the stopper assembly (rubber stopper fitted with stirrer, gas inlet and outlet tubes, burette tip, and glass rod stopper). Gently bubble argon into the flask. Turn on the stirrer and slowly increase the speed until the solution is stirring vigorously without splashing. Wait 1 min before beginning the titration, and continue stirring throughout the procedure.
After adding 15-17 ml of 0.1 N titanium trichloride drop-wise within about 2 min, stop the flow of titrant and reduce the argon flow. Remove the solid glass rod from the stopper assembly, and pipet 10 ml of 50% ammonium thiocyanate (indicator solution) into the flask. The colour of the solution will become brownish-red. Remove the pipet, re-insert the glass rod, and restore the argon flow. Add 0.1 N titanium trichloride dropwise, with 2-3 sec pauses between drops, until a sharp colour change from brownish-red to light green is observed. The endpoint (20-21 ml) is reached when the solution returns to the original light green colour and remains that colour for 20 sec. Stop the argon flow and gradually turn off the stirrer. Record the volume (V) of 0.1 N titanium trichloride used to the nearest 0.05 ml. Perform the titration procedure in triplicate.
Determine the indicator blank by repeating the above procedure without the 0.100 N potassium dichromate. The blank determination should require less than 0.5 ml of 0.1 N titanium trichloride. Record the volume used to the nearest 0.05 ml.
For each titration, the concentration of the titanium trichloride solution is (N x 20)/(V - Vb),
N is the concentration of the standard potassium dichromate solution;
20 is the aliquot (ml) of the potassium dichromate solution;
V is the volume (ml) of titanium trichloride solution required to titrate the aliquot of standard potassium dichromate solution; and
Vb is the volume (ml) of titanium trichloride solution used to determine the blank.
Calculate the concentration of the standard titanium trichloride solution by averaging the three titration results. (Restandardize the solution weekly by performing one titration of 0.100 N potassium dichromate and determining one indicator blank.)
Determination of total colouring matters content of sample
Accurately weigh the quantity of sample indicated in each specification monograph (WS, in mg) into a 500-ml conical flask. Add 10 g of sodium citrate or 15 g of sodium hydrogen tartrate, as specified in each monograph, a few boiling chips, and 150 ml of water. Wash down the walls of the flask with water, cover with a watch glass, and gently swirl to dissolve. In a fume hood, heat the solution to boiling on a hot plate. Boil vigorously for at least 10 sec to remove dissolved oxygen. (Avoid sample decomposition by boiling the solution for no more than 2 min.) Using gloves, remove the flask from the hot plate. Within 2-4 min of removing the flask from the hot plate, remove the watch glass, and connect the flask securely to the stopper assembly (flask might still be hot). Gently bubble argon into the flask. Turn on the stirrer and slowly increase the speed until the solution is stirring vigorously without splashing. Wait 1 min before beginning the titration, and continue stirring throughout the procedure. The colour will act as its own indicator unless otherwise stated in the appropriate monograph.
Rapidly add standardized 0.1 N titanium trichloride dropwise until the colour of the solution begins to change, then stop for 15-20 sec. Continue adding the titrant dropwise, with 1-2 sec pauses between drops. When the solution is close to the final colour, stop again for 20 sec. Continue adding 0.1 N titrant dropwise, with 5-10 sec pauses between drops, until the final colour is observed. The endpoint is reached when the final colour is stable for 20 sec. Stop the argon flow and gradually turn off the stirrer. Record the volume of titrant used to the nearest 0.05 ml
The percent total colouring matters content of the sample is 100 x (V x F x N) / (WS),
V is the ml of standardized titanium trichloride solution required;
F is D/(1.00 ml x 0.1 meq/ml), where D is the weight (mg) of colouring matters equivalent to 1.00 ml of 0.1 N titanium trichloride, quoted in the specification monograph); and
N is the concentration of standardized titanium trichloride solution (in meq/ml).
Procedure for Lakes
Add 150 ml of water to a 500-ml conical flask and dissolve in it the buffer compound specified in the monograph for the parent colour. Accurately weigh a quantity of lake equivalent to 35-40 ml of 0.1 N titanium trichloride and transfer it to the flask. Add a few boiling chips, wash down the walls of the flask with water, and cover with a watch glass. In a fume hood, heat the mixture to boiling or until the lake has completely dissolved. Using gloves, remove the flask from the hot plate. Titrate with standardized 0.1 N titanium trichloride in the manner described under Determination of total colouring matters content of sample, above.
In this method, the subsidiary colouring matters are separated from the main colouring matter by ascending paper chromatography and are extracted separately from the paper. The absorbance of each extract is measured at its wavelength of maximum absorbance by visible spectrophotometry.
Because it is impractical to identify each subsidiary colouring matter and because the subsidiary colouring matters are usually minor components of food colours, the method assumes that the specific absorbance of each subsidiary colouring matter is the same as that of the total colouring matters. The subsidiary colouring matters content is calculated by adding together the absorbances of the extracts in conjunction with the total colouring matters content of the sample.
Chromatography tank and ancillary equipment ( and or equivalent) comprising:
- Glass tank (A) and cover (B)
- Supporting frame (C) for the chromatography paper
- Solvent tray (D)
- Secondary frame (E) for supporting "drapes" of the filter paper
- Whatman No. 1 chromatography grade paper or equivalent, 20 cm x 20 cm sheets
- Microsyringe, capable of delivering 0.1 ml with a tolerance of ± 0.002 ml
- Visible range spectrophotometer
- Spectrophotometer cells, closed, 40 mm path length
- Test tubes
- Filter paper, 9 cm, coarse porosity
Chromatography solvents (all reagent grade)
- Water:ammonia (sp.gr. 0.880):trisodium citrate (95 ml:5 ml:2 g)
- n-Butanol:water:ethanol:ammonia (sp.gr. 0.880) (600:264:135:6)
- 2-Butanone:acetone:water (7:3:3)
- 2-Butanone:acetone:water:ammonia (sp.gr. 0.880) (700:300:300:2)
- 2-Butanone:acetone:water:ammonia (sp.gr. 0.880) (700:160:300:2)
- n-Butanol:glacial acetic acid:water (4:1:5)
Shake for 2 min, allow layers to separate. Use the upper layer as the chromatography solvent.
. Assembly of Chromatography Apparatus
. Components of Chromatography Apparatus
- Acetone, reagent grade
- Sodium hydrogen carbonate, reagent grade
Not less than 2 h before carrying out the determination, arrange the filter-paper drapes in the glass tank and pour over the drapes and into the bottom of the tank sufficient chromatography solvent to cover the bottom of the tank to a depth of approximately 1 cm. Place the solvent tray in position and fit the cover to the tank.
Prepare a 1.0% aqueous solution of the sample. Mark out a sheet of chromatography paper as shown in . Apply 0.10 ml of the sample solution as uniformly as possible within the confines of the 18 cm x 7 mm rectangle, holding the nozzle of the microsyringe steadily in contact with the paper. Allow the paper to dry at room temperature for 1 - 2 h or at 50° in a drying cabinet for 5 min, followed by 15 min at room temperature. Mount the dried sheet, together with a plain sheet to act as a blank on the supporting frame. (If required, several dried sheets may be developed simultaneously.)
. Method of Marking Chromatography Paper
Pour sufficient chromatography solvent into the solvent tray to bring the surface of the solvent about 1 cm below the base line of the chromatography sheets. The volume necessary will depend on the dimensions of the apparatus and should be predetermined. Put the supporting frame into position and replace the cover. Allow the solvent front to ascend the distance above the base line noted in the specification monograph, then remove the supporting frame and transfer it to a drying cabinet at 50-60° for 10-15 min. Remove the sheets from the frame.
Cut each subsidiary band from each chromatogram sheet as a strip, and cut an equivalent strip from the corresponding position of the plain sheet. Place each strip, subdivided into a suitable number of approximately equal portions, in a separate test tube. Add 5.0 ml of water:acetone (1:1 by vol) to each test tube, swirl for 2 - 3 min, add 15.0 ml of 0.05 N sodium hydrogen carbonate solution, and shake the tube to ensure mixing. Filter the coloured extracts and blanks through 9-cm coarse porosity filter papers into clean test tubes and determine the absorbances of the coloured extracts at their wavelengths of maximum absorbance, using 40-mm closed cells, against a filtered mixture of 5.0 ml of water :acetone (1:1 by vol) and 15.0 ml of the 0.05 N sodium hydrogen carbonate solution. Measure the absorbances of the extracts of the blank strips at the wavelengths at which those of the corresponding coloured extracts were measured and correct the absorbances of the coloured extracts with the blank values.
Prepare a standard solution from the 1.0% sample solution, corresponding to L/100% where L is the subsidiary colouring matters limit given in the specification monograph. Apply 0.10 ml of this solution to a sheet of chromatography paper by the technique outlined above, run a chromatogram and a blank, and dry at 50-60° for 10-15 min. Cut the band from the sheet as a strip and cut an equivalent strip from the blank sheet. Proceed as detailed previously and determine the total absorbance (As) of the standard corrected for the blank.
Calculate the percent subsidiary colouring matters in the sample using the following equation:
L is the limit for subsidiary colouring matters given in the specification monograph;
D is the total colouring matters content of the sample;
Aa + Ab + Ac ...An is the sum of the absorbances of the subsidiary colouring matters corrected for the blank values; and
As is the absorbance of the standard solution;
- Upward displacement type liquid/liquid extractor with sintered glass distributor, 500 ml working capacity with a piece of bright copper wire suspended through the condenser
- Distillation flasks: 250 and 500 ml
- Small coils of copper wire (0.5 g) for placing in distillation flasks
- Oven, 0 to 200° range
- Aluminium oxide, powdered, chromatography grade
- Ferrous sulfate, reagent grade
- Ammonium thiocyanate, reagent grade
- Titanium trichloride, 0.1 N, standard solution
- Sodium hydroxide, 2 N and 0.1 N, reagent grade
- Hydrochloric acid, 3 N and 0.1 N, reagent grade
- Ethyl ether or isopropyl ether, freshly distilled or stabilized
Immediately before use, freshly distilled ether should be passed through a 30 cm column of aluminium oxide in order to remove peroxides and inhibitors. Test to ensure the absence of peroxides, as follows:
Prepare a colourless solution of ferrous thiocyanate by mixing equal volumes of 0.1 N solutions of ferrous sulfate and ammonium thiocyanate. Carefully discharge any red colouration, due to ferric ions, with titanium trichloride. To 50 ml of this solution add 10 ml of ether and shake the mixture vigorously for 2-3 min. No red colour should develop.
Alkaline ether extract. Weigh accurately about 5.0 g of the colouring matter sample (WS). (For colouring matters with solubilities of less than 5 g/150 ml, use the lower weights prescribed in the specification monograph under TESTS). Dissolve the sample in 150 ml of water, add 2.5 ml of 2 N sodium hydroxide and transfer the solution to a 500-ml distillation flask; dilute with water to approximately 200 ml. Add 200 ml of ether to the distillation flask and extract for 2 h with a reflux rate of about 15 ml/min. Reserve the colour solution. Transfer the ether extract to a separatory funnel and wash the ether extract with two 25-ml portions of 0.1 N sodium hydroxide and then with water. Transfer to a tared 150-ml distillation flask (W1) containing a clean copper coil and distil off the ether in portions, reducing the volume to about 5 ml.
Acid ether extract. To the colour solution reserved above, add 5 ml of 3 N hydrochloric acid, mix and extract with a further quantity of the ether as above. Wash the ether extract with two 25-ml portions of 0.1 N hydrochloric acid and then with water. Transfer in portions to the flask containing the evaporated alkaline extract and carefully evaporate all the ether. Complete the drying in an oven at 85° for 20 min, then allow the flask to cool in a desiccator for 30 min and weigh. Repeat the drying and cooling until constant weight is obtained (W2).
The percent ether-extractable matter is 100 x (W2-W1)/WS.
- Soxhlet extractor - Suspend a piece of bright copper wire through the condenser. Place a small coil of copper wire (0.5 g) in the distillation flask.
- Ethyl ether or isopropyl ether, freshly distilled or stabilized
Purify the ether and test to ensure the absence of peroxides as directed in Method I.
Weigh accurately about 2 g of the colouring matter sample (WS). Transfer to the Soxhlet thimble and extract with 150 ml ether for 5 h. Concentrate the ether extract on a steam bath to about 5 ml. Dry the residue in a tared evaporating dish (W1) on a water bath and then dry at 105° until a constant weight is obtained (W2).
The percent ether-extractable matter is 100 x (W2-W1)/WS.
- Oven, 0 - 200° range
- Sintered glass crucible, No. 4
- Hydrochloric acid, concentrated
- Hydrochloric acid, 0.5% v/v
Accurately weigh approximately 5 g of the lake (WS) into a 500 ml beaker. Add 250 ml water and 60 ml concentrated hydrochloric acid. Boil until all the colour and alumina have dissolved. Filter through a tared sintered glass crucible (W1). Wash the crucible with hot 0.5% hydrochloric acid until the washings are colourless. Dry the crucible at 135°to constant weight (W2). Cool in a desiccator before weighing.
The percent hydrochloric acid-insoluble matter is 100 x (W2-W1)/WS.
Air is blown through an aqueous solution containing the colouring matter, copper(II) chloride, and dimethylformamide and the solution is analyzed spectrophotometrically. Under these conditions the leuco base is oxidized to the corresponding colouring matters and the increase in absorbance is equivalent to the amount of leuco base originally present.
- Visible range spectrophotometer
- Spectrophotometer cells, 1 cm path lengths (flow-through cells optional)
- Copper (II) chloride [CUCI2.2H2O], reagent grade
- Dimethylformamide (DMF), reagent grade
Note: The entire procedure should be completed as quickly as possible.
Solution A: Weigh 10.0 g of copper (II) chloride and dissolve in 200 ml of DMF. Transfer to a 1-liter volumetric flask and make up to the mark with DMF.
Solution B: Accurately weigh the quantity of sample indicated in the specification monograph (W, in mg). Dissolve in approximately 100 ml water, transfer quantitatively to a 1-liter volumetric flask and make up to the mark with water.
Solution a: Pipet 50 ml DMF into a 250-ml volumetric flask. Cover with Parafilm™ (or equivalent covering) and place in the dark.
Solution b: Accurately pipet 10 ml of Solution B into a 250-ml volumetric flask. Add 50 ml DMF. Cover with Parafilm™ (or equivalent covering) and place in the dark.
Solution c: Pipet 50 ml of Solution A into a 250-ml volumetric flask. Bubble air through the solution for 30 min in the following manner: Insert a 5-ml pipette into flexible tubing attached to a bench air flow source. Slowly, turn on the air, insert the pipette into the solution in the flask and adjust the air flow to a rapid, but controlled, rate. After 30 min remove the pipette from the solution and rinse the sides of the pipette into the flask with water from a wash bottle. Then, turn off the air flow.
Solutions d1 and d2 (duplicates): Accurately pipet 10 ml of Solution B into each of two 250-ml volumetric flasks. Add 50 ml of Solution A to each flask. Bubble air through the solutions for 30 min, using the method given for preparation of Solution c. After stopping the air flow, dilute solutions a- d2 in the five flasks nearly to volume with water and place the flasks in a water bath until they have cooled to room temperature, as heat is evolved when DMF and water are mixed. Do not leave them for longer than necessary; 5-10 min is normally enough. Dilute to volume with water. Immediately measure the absorbances of the solutions by spectrophotometry.
According to the table below, generate the absorbance curves for solutions a, b, c, d1, and d2 between 700 and 500 nm, using solutions a and c as blanks. Rinse cells thoroughly with each sample solution between measurements. When using the flow-through cells, use 3 separate rinses of at least 40 ml of each sample solution to be measured.Curve Blank Solution Comments 1 a a Set zero at 700 nm, run curve; record absorbance at wavelength of maximum absorption for colouring matter standard 2 a b Run curve without readjusting zero setting; record absorbance at wavelength of maximum absorption 3 c c Set zero at 700 nm; record absorbance at wavelength of maximum absorption for colouring matter standard 4a c d1 Run curve without readjusting zero setting; record absorbance at wavelength of maximum absorption 4b c d2 Run curve without readjusting zero setting; record absorbance at wavelength of maximum absorption
Calculate the percent leuco base in the sample using the following equation:
% leuco base = 100 Ч [(A4 - A3) - (A2 - A1)] Ч 1 liter Ч F/(a Ч b Ч W Ч R)
A1.. .A4 are the recorded absorbances of curves 1, 2, 3, and 4, respectively;
F is 250 ml/10 ml (dilution factor);
a is the absorptivity of 100% colouring matters in liter/ (mg-cm);
b is the cell path length (1 cm); and
R is the formula weight of colouring matter/formula weight of leuco base (given in the specification monograph).
For the separation and determination of organic compounds other than colouring matters (i.e., uncoloured impurities, such as uncombined intermediate starting materials), high-performance liquid chromatography (HPLC) has several advantages over other chromatographic techniques, viz. improved separations, speed (it can be automated) and accuracy. When determining named organic compounds, standards of each compound likely to be encountered are needed before any particular colour can be analyzed.
HPLC methods are generally outlined rather than described in detail. Column packing materials, capillary columns, and type and sensitivity of detectors should be chosen for optimum separation and quantitation of impurities currently listed in food colour specifications, as well as other impurities.
The alternative (traditional) method to HPLC is column chromatography (described further below), which involves collecting the eluate in fractions, using ultraviolet spectrophotometry to identify the compounds in each fraction, and calculating their concentrations.
Determination by High Performance Liquid Chromatography
The organic compounds other than colouring matters are separated by HPLC using gradient elution and are quantitatively determined by comparison of their peak areas against those obtained from standards. The conditions prescribed must be treated as guidelines and minor modifications might be needed to achieve the separations. Deviations from the prescribed conditions, such as a different column length, other types of column packing and solvent system, and the use of paired ion procedures, can result in elution characteristics different from those for the conditions given here, such as order of elution and resolution.
- High-performance liquid chromatograph capable of gradient elution with
- pump(s), flow rate 1 ml/min
- auto-sampler with a 20 μl injector
- detector, UV-visible absorption
- Chromatography column, C-18 on silica gel, 5 μm particle size, 250 x 4.6 mm.
- Guard column, C-18 on silica gel, 5 μm particle size, 15 x 4.6 mm
- Methanol, HPLC grade
- Ammonium acetate, HPLC grade
- Reference standards as required
- Injection volume: 20 μl.
- A: 0.2 N ammonium acetate;
- B: methanol
- 0.0 (sample injection)
- 0 to 35 min - 0 to 40% B (analysis)
- 3 5 to 41 min - 100% B (wash)
- 41.1 to 55 min - 100 to 0% B (return to initial gradient composition and equilibrate column)
- Flow rate: 1.0 ml per min
- Temperature: Ambient
- Pump pressure: minimum 300 psi, maximum 4000 psi
- Detector wavelengths: as required
- Integration: peak area
Prepare 0.5% (w/w) colouring matter sample solutions in 0.02 M ammonium acetate. Prepare calibration solutions from standards of impurities named in the specification monograph.
Analyze, following the instructions given for the HPLC chromatograph and detector.
Determination by Column Chromatography
- Chromatography column (see )
- UV range spectrophotometer
- Spectrophotometer cells, 1 cm path length
- Reference standards, as required
. Chromatography Column (dimensions in cm)
- Powdered cellulose, Whatman, or equivalent low iron cellulose
- Ammonium sulfate, reagent grade, very low in iron
Prepare a 25% ammonium sulfate solution for use as the eluent. Prepare a slurry of powdered cellulose in the 25% ammonium sulfate solution, using about 75 g of cellulose to 500 ml of liquid. Place a small disk of stainless steel gauze in the constriction above the tip of the chromatography column. Pour a sufficient volume of the slurry into the column so that the height of the packing is about 5 cm from the top of the column. Tap the column occasionally to ensure efficient packing. Wash the column with 200 ml of the eluent.
Test the column by passing 200 ml of 25% ammonium sulfate solution through it and measuring the UV absorption of the solution by spectrophotometry. The absorption must be sufficiently low to avoid interference with the intended analysis.
Weigh 0.200 g of the colouring matter sample (W) into a 150-ml beaker and dissolve in 20 ml of water. Add approximately 5 g of powdered cellulose. Add 50 g of ammonium sulfate to salt out the colour. Transfer the mixture to the chromatography column, rinse the beaker with the 25% ammonium sulfate solution, and add the washings to the column. Allow the column to drain until flow ceases, or nearly so.
Add 25% ammonium sulfate solution to the column at a rate equal to the flow rate through the column. Collect the effluent in 100-ml fractions. Continue until twelve fractions have been collected. Reserve the column and contents until the last fraction has been examined by spectrophotometry.
Mix each fraction well, and obtain the UV absorption spectrum of each solution from 220 to 400 nm, using the eluent as the blank. If the UV spectrum of the twelfth fraction shows the presence of any compound, continue collecting fractions until the compound is eluted.
Absorptivities of the organic compounds, such as intermediate starting materials, collected in the separate fractions and expected to be present in the colouring matters are used to calculate the percent organic compounds other than colouring matters in the sample and can be found in the specification monograph of the food colour.
Calculate the percent organic compounds other than colouring matters in the sample using the following equation:
A is the total absorbance of eluted fractions corrected for absorbance of blank;
0.100 liter is the volume of one fraction; and
a is the absorptivity in liter/(g-cm).
Note: Usually only one compound is encountered in each eluted fraction. When more than one compound is present in significant quantities in any fraction, the spectrophotometric data will so indicate. In such cases, the amounts of the various compounds must be determined by the procedure customarily used for the spectrophotometric analysis of mixtures of absorbing materials.
Some samples contain small amounts of various materials, particularly inorganic salts, which contribute "background absorption". Correction for this is made as follows: Determine the amount of background absorption of the fraction collected from the column immediately before and of the fraction immediately following those fractions in which the organic compounds are encountered. Subtract one-half of the sum of these two absorbances from the observed absorbance of the fractions containing the organic compounds. The remainder is taken as the absorbance due to inorganic salts.
Note: This determination is done in connection with Water Content (Loss on Drying) for food colours and the result is included in that calculation.
- Sodium chloride, reagent grade, sulfate-free
- Hydrochloric acid, reagent grade
- Barium chloride, reagent grade
Weigh 5.0 g of the colouring matter sample, transfer it to a 250-ml conical flask and dissolve in about 100 ml of water by heating on a water bath. Add 35 g of sulfate-free sodium chloride, stopper the flask, and swirl at frequent intervals for 1 h. Cool the flask, transfer the contents with saturated sodium chloride solution to a 250-ml volumetric flask, allow the solution to cool further to 20°, and dilute to volume. Shake the flask, and filter the solution through a dry filter paper. Pipet 100 ml of the filtrate into a 500-ml beaker, dilute to 300 ml with water and acidify with hydrochloric acid, adding 1 ml in excess. Heat the solution to boiling, and add an excess of 0.25 N barium chloride solution, drop by drop, with stirring. Allow the mixture to stand on a hot plate for 4 h, or leave it overnight at room temperature. Heat the mixture to about 80° and allow the precipitate to settle. Filter off the precipitated barium sulfate, wash with hot water, and ignite at a dull red heat in a tared crucible until a constant weight is obtained. Carry out a blank determination using the above procedure and correct the weight of barium sulfate found.
Calculate the sulfate content of the sample as percent sodium sulfate:
% sodium sulfate =
100 x (2.5 x corrected weight of barium sulfate found x 0.6086)/Weight of sample.
Unsulfonated primary aromatic amines are extracted into toluene from an alkaline solution of the sample, re-extracted into acid, and then determined spectrophotometrically after diazotization and coupling. They are expressed as aniline unless they are known to be some other amine.
Note: This method is not sufficiently sensitive for determining aniline at low mg/kg levels or below.
- Visible range spectrophotometer
- Spectrophotometer cells, 40 mm path length
- Toluene, reagent grade
- Hydrochloric acid, 1 N, reagent grade
- Hydrochloric acid, 3 N, reagent grade
- Potassium bromide, 50% solution, reagent grade
- Sodium carbonate solution, 2 N, reagent grade
- Sodium hydroxide, 1 N, reagent grade
- Sodium hydroxide, 0.1 N, reagent grade
- R salt (2-naphthol-3,6-disulfonic acid, disodium salt) solution, 0.05 N, reagent grade
- Sodium nitrite solution, 0.5 N, reagent grade
- Aniline, reagent grade
Preparation of a Standard Aniline Solution
Weigh 0.100 g of redistilled aniline into a small beaker and transfer to a 100-ml volumetric flask, rinsing the beaker several times with water. Add 30 ml of 3 N hydrochloric acid and dilute to the mark with water at room temperature. Dilute 10.0 ml of this solution to 100 ml with water and mix well; 1 ml of this solution is equivalent to 0.0001 g of aniline. Prepare the standard aniline solution freshly when required.
Preparation of a Calibration Graph
Measure the following volumes of the standard aniline solution into a series of 100-ml volumetric flasks: 5 ml, 10 ml, 15 ml, 20 ml, and 25 ml.
Dilute to 100 ml with 1 N hydrochloric acid and mix well. Pipet 10 ml of each solution into clean, dry test tubes; cool them for 10 min by immersion in a beaker of ice water. To each tube add 1 ml of the potassium bromide solution and 0.05 ml of the sodium nitrite solution. Mix and allow the tubes to stand for 10 min in the ice water bath while the aniline is diazotized. Into each of five 25-ml volumetric flasks, measure 1 ml of the R salt solution and 10 ml of the sodium carbonate solution. Pour each diazotized aniline solution into a separate flask containing R salt solution and sodium carbonate solution; rinse each test tube with a few drops of water. Dilute to the mark with water, stopper the flasks, mix the contents well and allow them to stand for 15 min in the dark.
Measure the absorbance of each coupled solution at 510 nm using 40 mm cells. As a reference solution, use a mixture of 10.0 ml of N hydrochloric acid, 10.0 ml of the sodium carbonate solution, and 2.0 ml of the R salt solution, diluted to 25.0 ml with water. Plot a graph relating absorbance to weight of aniline in each 100 ml of aniline solution.
Preparation and Evaluation of a Test Solution
Weigh, to the nearest 0.01 g, about 2.0 g of the colouring matter sample (W) into a separatory funnel containing 100 ml of water, rinse down the sides of the funnel with a further 50 ml of water, swirling to dissolve the sample, and add 5 ml of 1 N sodium hydroxide. Extract with two 50-ml portions of toluene and wash the combined toluene extracts with 10-ml portions of 0.1 N sodium hydroxide to remove traces of colour. Extract the washed toluene with three 10-ml portions of 3 N hydrochloric acid and dilute the combined extract to 100 ml with water. Mix well. Call this Solution T.
Pipet 10.0 ml of Solution T into a clean, dry test tube, cool for 10 min by immersion in a beaker of ice/water, add 1 ml of the potassium bromide solution and proceed as described above for the preparation of the calibration graph, starting with the addition of 0.05 ml of the sodium nitrite solution.
Measure the absorbance of the coupled test solution at 510 nm using a 40 mm cell. Use a reference solution prepared from 10.0 ml of Solution T, 10 ml of the sodium carbonate solution, and 2.0 ml of the R salt solution diluted to 25.0 ml with water.
From the calibration graph, read the weight of aniline (Wa) corresponding to the observed absorbance of the test solution.
Note: See the methods to determine Chloride as Sodium Chloride and Sulfate as Sodium Sulfate. Specifications for food colours include the results of those tests as part of the calculation of Loss on Drying.
Colouring materials containing -SO3Na or -COONa groups are usually hygroscopic and any water retained from their manufacture (or subsequently absorbed from the atmosphere) is generally present in the form of a hydrate. When such colouring matters are dried at 135° the loss in weight can generally be equated to the total water content, but this is not always the case. For example, Erythrosine and Ponceau 4R each retain one molecule of water of crystallization at 135° and it is normal practice to take this into account when totalling the amounts of main components present in a sample.
- Oven, 0 - 200° range
- Weighing bottle, 50 mm in diameter and 30 mm high, with ground glass stopper
Weigh 2.0 - 3.0 g of the sample (W1) in a tared weighing bottle plus stopper. Heat the unstoppered bottle in the oven at the temperature prescribed in the specification monograph (±5°), until a constant weight is obtained. Cool the crucible and residue in a desiccator before each weighing.
where W2 is the weight of the dried sample. (See Note above)
- Oven, 0 - 200° range
- Porcelain filtering crucible
- Glass micro fiber filter disc, Whatman type GF/C, compliant with BS 1752
Weigh 4.5 - 5.5 g of the sample (WS) into a 250 ml beaker. Add about 200 ml of hot water (80-90°), stir to dissolve, and allow the solution to cool to room temperature. Filter the solution through a tared porcelain crucible and filter disc and wash with cold water until the washings are colourless. Dry the crucible and residue at 135° until a constant weight is obtained. Cool the crucible and residue in a desiccator before weighing.
Wr is the weight of the residue.
- Nitric acid, 1.5 N, reagent grade
- Hydrochloric acid, reagent grade
Accurately weigh 10 g of the sample into a 400 ml beaker. Add 250 ml of water. Stir to wet the sample and then stir occasionally during a period of 30 min. Filter.
Measure 50 ml of the filtrate, add 50 ml water and acidify with 5 ml of 1.5 N nitric acid. Determine the chloride content by the potentiometric method used for soluble colours (see Chloride as Sodium Chloride determination).
Measure 50 ml of the filtrate, dilute to 300 ml with water and acidify with hydrochloric acid, adding 1 ml in excess. Heat the solution to boiling and add an excess of 0.25 N barium chloride, drop by drop, with stirring. Complete the analysis by digesting, filtering, and igniting the precipitate as described in the method used for the determination of sulfate in soluble colours (see Sulfate as Sodium Sulfate determination).
Ring and ball softening point method
The ring-and-ball softening point is defined as the temperature at which a disk of the sample held within a horizontal ring is forced downward a distance of 1 in. (25.4 mm) under the weight of a steel ball as the sample is heated at a prescribed rate in a water or glycerol bath.
The apparatus illustrated in Figures1 and 2 consists of the components described in the following paragraphs.
A brass-shouldered ring conforming to the dimensions shown in Figure 1a should be used. If desired, the ring may be attached by brazing or other convenient manner to a brass wire of about 13 B & S gauge (0.06 to 0.08 in., or 1.52 to 2.03 mm, in diameter) as shown in Figure 2a.
A steel ball, 3/8 in. (9.53 mm) in diameter, weighing between 3.45 and 3.55 g, should be used.
A guide for centering the ball, constructed of brass and having the general shape and dimensions, as illustrated in Figure 1c may be used if desired.
Use a heat-resistant glass vessel, such as an 800-ml low-form Griffin beaker, not less than 3.34 in. (8.5 cm) in diameter and not less than 5 in. (12.7 cm) in depth from the bottom of the flare.
Support for Ring and thermometer
Any convenient device for supporting the ring and thermometer may be used, provided that it meets the following requirements: (1) the ring shall be supported in a substantially horizontal position; (2) when using the apparatus shown in Figure 1d, the bottom of the ring shall be 1.0 in. (25.4 mm) above the horizontal plate below it, the bottom surface of the horizontal plate shall be at least 0.5 in. (13 mm) and not more than 0.75 in. (18 mm) above the bottom of the container, and the depth of the liquid in the container shall be not less than 4.0 in. (10.2 cm); (3) when using the apparatus shown in Figure 1e, the bottom of the ring shall be 1.0 in. (25.4 mm) above the bottom of the container, with the bottom end of the rod resting on the bottom of the container, and the depth of the liquid in the container shall be not less than 4.0 in. (10.2 cm), as shown in Figure 1 a, b and c; and (4) in both assemblies, the thermometer shall be suspended so that the bottom of the bulb is level with the bottom of the ring and within 0.5 in. (13 mm) but not touching the ring.
Depending upon the expected softening point of the sample, use either an ASTM 15C low-softening-point thermometer (-2° to 80°) or an ASTM 16C high-softening-point thermometer (30° to 200°).
Use a suitable mechanical stirrer rotating between 500 and 700 rpm. To ensure uniform heat distribution in the heating medium, the direction of the shaft rotation should move the liquid upward. (See Figure 2d for recommended dimensions.)
Select a representative sample of the material under test consisting of freshly broken lumps free of oxidized surfaces. Scrape off the surface layer of samples received as lumps immediately before use, avoiding inclusion of finely divided material or dust. The amount of sample taken should be at least twice that necessary to fill the desired number of rings, but in no case less than 40 g. Immediately melt the sample in a clean container, using an oven, hot plate, or sand or oil bath to prevent local overheating. Avoid incorporating air bubbles in the melting sample, which must not be heated above the temperature necessary to pour the material readily without inclusion of air bubbles. The time from the beginning of heating to the pouring of the sample shall not exceed 15 min. Immediately before filling the rings; preheat them to approximately the same temperature at which the sample is to be poured. While being filled, the rings should rest on an amalgamated brass plate. Pour the sample into the rings so as to leave an excess on cooling. Cool for at least 30 min, and then cut off the excess material cleanly with a slightly heated knife or spatula. Use a clean container and a fresh sample if the test is repeated.
Materials having softening points above 80°: Fill the glass vessel with glycerol to a depth of not less than 4.0 in. (10.2 cm) and not more than 4.25 in. (10.8 cm). The starting temperature of the bath shall be 32º. For resins (including rosin), the glycerol should be cooled to not less than 45º below the anticipated softening point, but in no case lower than 35º. Position the axis of the stirrer shaft near the back wall of the container, with the blades clearing the wall and with the bottom of the blades 0.75 in. (18 mm) above the top of the ring. Unless the ball-centering guide is used, make a slight indentation in the center of the sample by pressing the ball or a rounded rod, slightly heated for hard materials, into the sample at this point. Suspend the ring containing the sample in the glycerol bath so that the lower surface of the filled ring is 1.0 in. (25.4 mm) above the surface of the lower horizontal plate (see Figure 1d), which is at least 0.5 in. (13 mm) and not more than 0.75 in. (18 mm) above the bottom of the glass vessel, or 1.0 in. (25.4 mm) above the bottom of the container (see Figure 2e). Place the ball in the glycerol but not on the test specimen. Suspend an ASTM high-softening-point thermometer (16C) in the glycerol so that the bottom of its bulb is level with the bottom of the ring and within 0.5 in. (13 mm) but not touching the ring. Maintain the initial temperature of the glycerol for 15 min, and then, using suitable forceps, place the ball in the center of the upper surface of the sample in the ring. Begin stirring, and continue stirring at 500 to 700 rpm until completion of the determination. Apply heat at such a rate that the temperature of the glycerol is raised 5º per min, avoiding the effects of drafts by using shields if necessary.
[Note: The rate of rise of the temperature shall be uniform and shall not be averaged over the test period. Reject all tests in which the rate of rise exceeds ±0.5° for any minute period after the first three.]
Record as the softening point the temperature of the thermometer at the instant the sample touches the lower horizontal plate (see Figure 1d) or the bottom of the container (see Figure 2e). Make no correction for the emergent stem of the thermometer.
Materials having softening points of 80° or below: Follow the above procedure, except use an ASTM low-softening-point thermometer (15C) and use freshly boiled water cooled to 5° as the heating medium. For resins (including rosins), use water cooled to not less than 45° below the anticipated softening point, but in no case lower than 5°.
Apparatus - Ring and Ball Softening Point
(a) Shouldered Ring
(b) Ring Holder
(c) Bell Centering Guide
(d) Assembly Apparatus with Two Rings
Figure 1. Shouldered Ring, Ring Holder Ball-Centering Guide, and Assembly of Apparatus Showing Two Rings
Figure 2. Assembly of Apparatus Showing Stirrer and Single Shouldered Ring.
6th Edition, 2008, p. 1161 (figures 40 and 41).
Reprinted with permission from the US Pharmacopeia, 12601 Twinbrook Parkway, Rockville, MD USA 20852.
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