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'., “$2.3... n. ”a... m ”5.14;. w.- b‘ , .... ."“‘ ~‘u T . k I_ :‘TI‘JJ ,q " . .a. I" n: 'ti £5». 9'. 3‘33 ~ aw. .. “ XI and} :14 t .s. l'r'll m; .3, . r;. ‘ L i‘"' .I. .u - .7 I x . , .. . ‘ . ,_ I . ,1...‘.K".“, . .. . . ‘ . . ”L - 4. 1:313 I t .a ...’I , _ u _ . - _ .- ... I :I n l l w J' I - . g ._ _ V. I ‘ " " I -'-‘ . - '5‘ .nl rhr m 1.- - K- -r .1.- xvi 0012!. . 1;" "n1: .',Il..'.. ;.»._ p... "f- -'-: $7,” :.l..,:j‘:. _ '—-u¢'~‘ ._,..‘ . 1- RSITY LIBRARIES llllllllllllllllllllllllllllllll' “l 3129 This is to certify that the thesis entitled THE STABILITY OF A UV ABSORBER INCORPORATED INTO TRANSPARENT PACKAGES AND ITS EFFECT ON THE PHOTOOXIDATION OF SOYBEAN OIL presented by MELVIN ARTHUR PASCALL has been accepted towards fulfillment of the requirements for MASTER degree in PACKAQING BRUCE R. HARTE, PH.D. Major professor Date NOVEMBER 15 L 1991 0-7639 MS U i: an Affirmative Action/Equal Opportunity Institution ’ l LIBRARY Michigan State University l‘ .J PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on orbefore due due. DATE DUE DATE DUE DATE DUE ;l—_|| l MSU I: An Affirmative Action/Equal Opportunity Institution cMma-M ——_— w __ THE STABILITY OF A UV ABSORBER INCORPORATED INTO TRANSPARENT PACKAGES AND ITS EFFECT ON THE PHOTOOXIDATION OF SOYBEAN OIL. by Melvin Arthur Pascall A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE School of Packaging 1991 ABSTRACT THE STABILITY OF A UV ABSORBER INCORPORATED INTO TRANSPARENT PACKAGES AND ITS EFFECT ON THE PHOTOOXIDATION OF SOYBEAN OIL. by Melvin Arthur Pascall The stability and effectiveness of Tinuvin 326, a UV absorber, dispersed within the regrind layer of coextruded multilayered polypropylene based containers, was studied by investigating the ability of Tinuvin 326 to protect packaged, bleached soybean oil from photooxidation. The level of Tinuvin 326 in the containers was determined using high pressure liquid chromatography and UV spectrophotometric methods respectively. No loss of Tinuvin 326 from the containers to the atmosphere was observed over a 42 day storage period at 21 and 35°C. The migration of Tinuvin 326 from the containers to the oil stored at 35°C for 42 days, was greater than at 21°C for 42 days. The migration level was too low to noticeably reduce Tinuvin 326 levels in the containers. The peroxide value method used to measure lipid oxidation in the bleached oil, determined the effectiveness of Tinuvin 326. Containers with 0.3% Tinuvin 326 were found to have significantly reduced photooxidation of bleached oil exposed to fluorescent light during 35 storage days at 21 and 35°C, as compared to containers with no Tinuvin 326. To my mother Priscilla Pascall "ACKNOWLEDGMENTS" My sincere gratitude is extended to Dr. Bruce R. Harte for his guidance and valuable contribution as major professor and research committee chairman. Sincere appreciation is also extended to Drs. Jack R. Giacin (School of Packaging) and J. Ian Gray, Department of Food Science and Human Nutrition, for their assistance and for serving on my research committee. Special appreciation is extended to Dr. Parvin Hoojjat for her patience and technical assistance during the laboratory experimentations. Thanks to fellow graduate students A. Yvonne Blake-Smith, Virginia Vega-Vargas and Frank Monahan for their support and assistance which helped to make this study successful. Special thanks to The Center for Food and Pharmaceutical Packaging for the financial support they provided. I am grateful to Stephen Webster (Carib Glassworks Limited), for his advice which initiated my interest in studying here at Michigan State University. I would especially like to thank my brothers Ellis and Lionel, and sisters Eunice and Genita for the motivation and encouragement they gave me throughout this course of study. iv TABLE OF CONTENTQ LIST OF TABLES O O O O O O O O O O O O O O O O O O O O Vii LI ST OF FIGURES O O O O O O O O O O O O O O O O O O O O O Xi INTRODUCT ION O O O O O O O O O O O O O O O O O O O O O 1 LITERATURE REVIEW . . . . . . . . . . . . . . . . . . . . .4 Mechanism of lipid oxidation . . . . . . .4 Measurement of lipid oxidation . . . . Types and mechanism of antioxidants . . 11 Effect of light on oxidation . . . . . 15 Effect of packaging on light sensitive foods. . . . 19 UV light absorbers . . . . . . . . . . . . . . . . . 20 Tinuvin 326 . . . . . . . . . . . . . . . . . . . . 29 MATERIALS AND METHODS . . . . . . . . . . . . . . . . . 32 Test packages: . . . . . . . . . . . . . . . . . . . 32 Soybean oil: . . . . . . . . . . . . . . . . . . . . 32 Light source: . . . . . . . . . . . . . . . . . . . 33 EXPERIMENTAL PROCEDURE . . . . . . . . . . . . . . . . 34 Extraction of Tinuvin 326 from sample containers: . 34 HPLC procedure to determine Tinuvin 326 in extract: . . . . . . . . . . . . . . . . . . . 35 UV spectrophotometric procedure to determine Tinuvin 326 in extract: . . . . . . . . . . . . 36 Loss of Tinuvin from the containers: . . . . . . . . 37 Migration of Tinuvin from containers into the oil: . . . . . . . . . . . . . . . . . . . . . 37 Bleaching the soybean oil: . . . . . . . . . 40 Irradiation System to expose the bleached oil to fluorescent light: . . . . . . . . . . . . . . 41 Lipid oxidation determination by peroxide value method: . . . . . . . . . . . . . . . . . . . . 43 RESULTS AND DISCUSSION . . . . . . . . . . . . . . . . 44 Standard curves and absorbance spectra profiles: . . 44 Loss of Tinuvin 326 from the container material: . 52 Migration of Tinuvin from containers into the oil: . . . . . . . . . . . . . . . . . . . . . 63 Lipid Oxidation Determination by Peroxide Value Method: . . . . . . . . . . . . . . . . . . . . 76 Conclusions . . . . . . . . . . . . . . . . . . . . . . . 83 Appendix 1. . . . . . . . . . . . . . . . . . . . . . . 85 Appendix 2. . . . . . . . . . . . . . . . . . . . . . . 86 Appendix 3. . . . . . . . . . . . . . . . . . . . . . . 87 Appendix 4. . . . . . . . . . . . . . . . . . . . . . . 88 Appendix 5. . . . . . . . . . . . . . . . . . . . . . . 94 Appendix 6. . . . . . . . . . . . . . . . . . . . . . . 95 Appendix 7. . . . . . . . . . . . . . . . . . . . . . . 99 Appendix 8. . . . . . . . . . . . . . . . . . . . . . 100 References 0 O O O O O O O O O O O O O O O O O O O O O 102 vi Table 1. Table 2. Table 3. Table 4. Table 5. Table 6. Table 7. Table 8. Table 9. Table 10. Table 11. I T 0 LBS Relative percent of Tinuvin 326 in the multilayer containers stored at 21°C. (HPLC methOd) O O O O O O O O O O O I O O O O O O 0 Relative percent of Tinuvin 326 in the multilayer containers stored at 21°C. (UV spectrophotometric method). . . . . . . . . . Percent Tinuvin 326 in the multilayer containers stored at 35°C. (HPLC method). . Percent Tinuvin 32 6 in the multilayer containers stored at 35°C. (UV spectrophotometric method). . . . . . . . . . HPLC analysis of containers with Tinuvin 326 stored at 21 degrees C. . . . . . . . . . . . HPLC analysis of containers with Tinuvin 326 stored at 35 degrees C. . . . . . . . . . . . Stability studies on containers with Tinuvin 326 stored at 21 degrees C. using UV spectrophotometric method. . . . . . . . . . Stability studies on containers with Tinuvin 326 stored at 35 degrees C. using UV spectrophotometric method. . . . . . . . . . Amount of Tinuvin 326 which migrated from the container wall to the oil at 21°C, (HPLC methOd) O O O O O O O O O I O O O O O O O O 0 Amount of Tinuvin 326 which migrated from the container wall to the oil at 35°C, (HPLC methOd) O O O O O O I O O O O O O O O O Level of Tinuvin 326 in the wall of the containers in which oil was stored at 21°C (HPLC methOd) O O O O C O O O O O O O O O O 0 vii 55 56 57 58 88 91 95 97 66 67 68 Table Table Table Table Table Table Table Table 12. 13. 14. 15. 16. 17. 18. 19. Level of Tinuvin 326 in the wall of the containers in which oil was stored at 35°C (HPLC method). . . . . . . . . . . . . . . . Percent Tinuvin 326 which migrated from the containers wall to the oil at 21°C, (HPLC methOd) O O O O O O O O O O O O O O O O O O 0 Percent Tinuvin 326 which migrated from the containers wall to the oil at 35°C, (HPLC methOd) O O O O O O O C O O O O O O O O O O 0 Percent efficiency of the extraction process and the rota—evaporation concentration procedure O O O O O O O O O O O O O O O O O O Peroxide value (milliequivalents/kg oil) of the soybean oil in the multilayered container with 0% and 0.3% Tinuvin 326, stored at 21°C . . . . . . . . . . . . . . . Peroxide value (milliequivalents/kg oil) of the soybean oil in the multilayered container with 0% and 0.3% Tinuvin 326, stored at 35°C Data used for the standard curve for Tinuvin 3 2 6 at several concentration levels (using HPLC) O O O O O O O O O O O O O O O O O O O 0 Data used for the standard curve for Tinuvin 326 at several concentration levels (using UV spectrophotometry). . . . . . . . . . . . . . viii 69 7O 71 75 78 81 85 86 Fig. Fig. Fig. Fig. Fig. Fig. Fig. Fig. Fig. Fig. Fig. Fig. Fig. 10. 11. 12. 13. LIST OF EIGUREB The mechanism of light energy absorption by a benzotriazole. . . . . . . . . . . . . . . . The structure of Tinuvin 326 . . . . . . . . . Standard curve of Tinuvin 326 concentration vs area response units (HPLC method). . . . . . . Standard curve of Tinuvin 326 concentration vs absorbance (optical density units) (UV spectrophotometric method). . . . . . . . . . Absorbance spectrum of container wall before extraction to remove Tinuvin 326. . . . . . . Absorbance spectrum of container wall after extraction to remove Tinuvin 326. . . . . . . Absorbance spectrum of container wall with 0% Tianin 326 O O O O I O O O O O O O I O O O O 0 Percent Tinuvin 326 in multilayer container wall stored at 21°C. (HPLC method). . . . . . Percent Tinuvin 326 in multilayer container wall stored at 35°C. (HPLC method). . . . . . Percent Tinuvin 326 in multilayer container wall stored at 21°C. (UV spectro. method). . . Percent Tinuvin 326 in multilayer container wall stored at 35°C. (UV spectro. method). . . Percent Tinuvin 326 which migrated from the containers to soybean oil at 21 and 35°C (HPLC method). . . . . . . . . . . . . . . . . Percent Tinuvin 326 in the wall of the containers following oil storage at 21°C (HPLC methOd) O O O O O O O O O O O O O O O C O I 0 ix 29 47 48 49 50 51 59 60 61 62 72 73 Fig. 14. Percent Tinuvin 326 in the wall of the containers following oil storage at 35°C (HPLC methOd) I O O O O O O O O O O O O O O O O O O 0 Fig. 15. Peroxide value of soybean oil in the containers 0% and 0.3% Tinuvin 326 stored at 21°C. . . . Fig. 16. Peroxide value of soybean oil in the containers with 0% and 0.3% Tinuvin 326 stored at 35°C. . 74 79 82 OD 0 An ideal package should provide adequate protection without unfavorable interaction with the product. The package should be attractive, relatively low cost and convenient to use. Various light sensitive products cannot be packaged in transparent containers because of possible deteriorative reactions which are catalyzed by incident light. To protect such products may result in having to conceal them from purchasing consumer. If transparent packages can be made to protect light sensitive products from incident light, and still present the product attractively, then a marketing advantage would be gained. In general, deterioration of fats, oils and foods containing lipids, can occur due to a variety of biological, chemical, and physical factors. Parker et a1. (1952) reported that the chemical changes responsible for spoilage of fats and oils are basically oxidative and hydrolytic processes. Oxidation involves the addition of oxygen to unsaturated fatty acids to form hydroperoxides. These peroxides decompose and. react with water to form aldehydes, ketones, and acids, some of which are responsible for disagreeable odors, flavors, color changes, and nutritive 1 2 losses. Certain metals exert profound effect on the catalytic rate of oxidation of fats and oils. Physical factors associated with fat spoilage are heat, ionizing radiation and light of certain wavelengths. The total exclusion of light from packaged oils would be an obvious solution to photooxidation. This can be accomplished by using metal and other types of packages that are impervious to light. However, in practice this is not employed because of marketing and economic reasons. In the past, amber bottles were used for a significant portion.of the vegetable oil market, but now clear glass and plastic bottles are used by most edible oil processors. Increased store lighting and transparent packaging, increase the likelihood that oils and other fatty food products will develop off flavors while on the grocery shelf. Since transparent plastic packaging is widely used for such foods, modification to improve the stability of oils, oil containing products and light sensitive foods can be beneficial to food processors and consumers . Barriers to light such as metal, paperboard, colored glass and pigmented plastics have been shown by a number of researchers to provide protection to light sensitive foods (Hoskin, 1988; deMan, 1978; Sattar et al., 1976; Nkpa et al., 1990). In this study, however, the effectiveness of a UV absorber, Tinuvin 326, was investigated by incorporating it 3 into coextruded, multilayered polypropylene containers. Theoretically, this would result in a transparent package with the ability to provide protection to light sensitive foods. The stability and duration of activity, as well as the potential migration of Tinuvin 326 into soybean oil was also investigated. Soybean oil is a light sensitive product and degradation can occur due to photooxidation, leading to rancidity and the loss of fat-soluble vitamins such as vitamin E and vitamin A. The specific objectives of this study were: (1) to develop methodology for the quantitation of Tinuvin 326 in the test containers; (2) to evaluate the stability of Tinuvin 326 in the material at various storage temperatures; (3) to investigate the migration of Tinuvin 326 from the package material to the product (soybean oil); and (4) to determine the change in product quality as a function of Tinuvin 326 (in the container wall), light intensity, exposure time, and storage temperature. LIIEBAIEB§_B§!1§! WW Lipid oxidation is a primary cause of food spoilage. In edible oils, and food products containing lipids, this can lead to the development of off flavors, off-odors, decreased nutritive quality and the production of certain potentially toxic, oxidative products. It is generally accepted that autoxidation is the main reaction involved in oxidative deterioration of lipids.. Although photochemical reactions have been known for a long time, only recently has the role of photosensitized oxidation and its interaction with autoxidation begun to be realized (Nawar, 1985). At conditions that normally exist during oxidation of food fats, the primary products of autoxidation are hydroperoxides (Sattar et al., 1976). The reaction is often referred to as autoxidation due to apparent self-catalytic properties. The generalized.mechanism reviewed by Gray 1978, is deplicted as: 1. Initiation RH + 02 » R. + .on 2. Propagation R. + 02 -+ R00. RH + ROO. * ROOH + R. ROOH * R0. + .OH 3. Termination R. + R. * RR R. + ROO. * ROOR ROO. + R00. » ROOR + 02 RH is any unsaturated fatty acid in which the H is labile due to its position on a_carbon atom adjacent to a double bond. R. is a free radical formed by removal of a labile hydrogen (Gray, 1978; Luby, 1982). Frankel (1984) reported that decomposition of lipid hydroperoxides is a very complicated process. Decomposition proceeds by homolytic cleavage of RO-OH to form alkoxy radicals (RO.) . Lipid hydroperoxides can react with oxygen to _ form secondary products as epoxyhydroperoxides, ketohydroperoxides, dihydroperoxides, cyclic peroxides and bicyclic endoperoxides. These secondary products decompose like monohydroperoxides to form volatile breakdown products. Lipid hydroperoxides can condense into dimers and polymers that can also breakdown and produce volatile materials (Hildebrand et al., 1984). The initiation step may occur due to hydroperoxide decomposition, metal catalysis, or by’ exposure to light (Nawar, 1985) . The source of hydroperoxide can be from another system undergoing oxidation. More recently, it has been postulated that singlet oxygen can be the active species involved, with plant and tissue pigments such as chlorophyll 6 or myoglobin, acting as sensitizers (Nawar, 1985). In this process, oxygen becomes activated to the singlet state by transfer of energy from the photosensitizer. The resulting singlet oxygen is extremely reactive (Frankel, 1984). Linoleate is reported to react at least 1500 times faster with singlet oxygen (V5) than with normal oxygen in the triplet ground state (302) Two pathways have been proposed for photosensitized oxidation (Nawar, 1985). In pathway 1, the sensitizer presumably reacts, after light absorption, with substrate (A) to form intermediates, which then react with ground state (triplet) oxygen to yield the oxidation products as shown: Sensitizer + A + hv 4 intermediates - I Intermediates - I + 02-» products + sensitizer In pathway 2, molecular oxygen rather than the substrate is the species that reacts with the sensitizer due to light absorption. Sensitizer + 02 + hv -v intermediates - II Intermediates - II + A » products + sensitizer (Nawar, 1985). Braun and Oliveros (1990) reported that besides sensitization, certain chemical reactions, electric‘discharge and photolytic decomposition of ozone may produce singlet oxygen. They summarized the sensitization mechanism as follows: hv Sensitizer ~ 18ensitizer* isc 1Sensitizer”: -v 3Sensitizer”: at 3Sensitizer* + 02 -v Sensitizer + 102 When the sensitizer is excited by absorption of light (hv), the singlet state of the sensitizer results (ISensitizerir). This singlet state can be converted to triplet state by intersystem crossing (isc) . Depending on the energy (at) of the triplet state, both singlet states of oxygen (12’ and 1A) can be generated, and the sensitizer is deactivated without chemical alteration. However, in practice, the 12’ state is rapidly deactivated, and only the ‘A state acts as the intermediate referred to as singlet oxygen (102) (Braun et al., 1990). There is also increasing evidence that singlet oxygen can react directly with the double bonds of unsaturated fatty acids in fats and oils to produce peroxides. This suggests that photosensitized oxidation may be important in initiating or propagating normal free radical autoxidation of unsaturated fats and oils (Vianni, 1980). Sherwin (1976) reported that the greater the unsaturation of a vegetable oil, the more susceptible it is to oxidative deterioration. Oils containing substantial amounts of linolenic acid, or certain other fatty aCids with more than two double bonds in their molecular structures, may undergo oxidative degradation known as "flavor 8 reversion." In commercial edible oils, the reversion problem is most likely encountered with soybean and rapeseed oils, which have high levels of linolenic acid (Sherwin, 1976). 3 EN P I Various methods have been developed to measure lipid oxidation. These tests may utilize chemical, physical or organoleptic evaluation. Gray (1978) reported that the method chosen depends on the type of information required, the speed of the method, the nature and quantity of the sample, and the test conditions. In sensory evaluation, the method can be as simple as individual tasting or smelling of the product. To minimize bias and human error, a multimembered panel is usually. employed. A statistically designed sampling experiment is then conducted. The disadvantages of this type of analysis are the length of time required and poor reproducibility (Gray, 1978). Various chemical methods are used to measure lipid oxidation. Among these methods the thiobarbituric acid (TBA) test is one of the most popular (Sattar et al., 1976; Gray, 1978; Hoojjat, 1988). This method is based on the assumption that a red color develops because of complexes formed between oxidation products of unsaturated fatty acids and TBA. Sinhuber et al. (1958) reported that the reactive compound in 9 the colorimetric reaction is malonaldehyde. The peroxide value (PV) method has been widely used to measure lipid oxidation in edible oils (Sattar and deMan, 1976; Sattar et al., 1976; Luby et al., 1986; Kiritsakis et al., 1984; Moser et al., 1965; Boki et al., 1989; Warner et al., 1989). In this method the ability of peroxides to liberate iodine from potassium iodine or to oxidize the ferrous ion to the ferric ion is measured. Gray (1978) reported that the PV method has limitations because peroxides undergo transition to other secondary compounds. This method has been revised and updated since its initial development (Gray, 1978). The Kreis test was one of the first tests used commercially to evaluate oxidation of fats (Sattar et al., 1976; Gray, 1978). In this test, development of a red color caused by reaction of phloroglucinol and oxidized products is measured. Mehlenbacher (1962) reported that erroneous results can be obtained as a similar color can develop from reactions not related to rancidity. The oxirane test is another method also used to measure oxidation. Nawar (1976) reported that it is based on the addition of hydrogen halides to the oxirane group. The epoxide content is determined by titrating the sample with hydrogen bromide in acetic acid, in the presence of crystal 10 violet, to a bluish green end point (Sattar et al., 1976). The measurement of carbonyl compounds is another chemical method used to evaluate lipid oxidation. The development of hydrazones, produced by the reaction of 2,4- dinitrophenylhydrazine with the oxidation products of aldehydes and ketones are measured (Henick et al., 1964) . Several methods have been used, but Gray (1978) reported that in the one most commonly employed, the formation of carbonyl compounds occurs in the presence of a trichloroacetic acid catalyst. Gray (1978) reported Lea (1962) criticisms which stated that hydroperoxides can decompose under conditions of this experiment. Physical methods are also used to determine lipid oxidation, these include uv spectrophotometry, fluorescence, infrared spectroscopy, polarography, chromatographic methods and refractometry (Gray 1978) . The uv spectrophotometric method is also referred to as the conjugated diene method. This procedure is based on the principle that oxidation of polyunsaturated fatty acids is accompanied by an increase in their uv absorption. Absorbancy in the region 230 - 375 nm measures conjugated unsaturated fatty acids, while absorbancy at 234 nm and 268 nm is usually measured to monitor oxidation in. diene 'unsaturated. and ‘triene 'unsaturated fatty acids respectively. 11 Gas chromatography has been the most widely used chromatographic system to measure lipid oxidation (Gray, 1978). This method of analysis is based on the measurement of specific compounds which are known to be typically formed during autoxidation. This method has been used by Jarvi et a1. (1971) to measure rancidity in soybean oil. Samples of soybean, sunflower and low erucic acid rapeseed oils were evaluated for flavor and oxidative stability by Warner et al. 1989, using the gas chromatography method. Infrared spectroscopy is used to follow change in functional groups as lipid oxidizes. Profiling the infrared spectra at varying times during oxidation can show a reduction in the number of bands and the consequential development of others. The refractive index method assumes that the. refractive index of reactants increases with increased peroxide formation. '_8 1-! _ 11._ y 0 ‘ 0 ."1’ 1 .V The reaction of oxygen with unsaturated lipids and polymers to cause oxidation, is basically the same. The mechanism is initiated by the removal of hydrogen from a hydrocarbon molecule, which can occur in the presence of metals, light, heat, or hydroperoxides. The resulting free radicals (R.) react with oxygen to form peroxy radicals (ROO.). In the propagation process, ROO. reacts with more RH groups to form hydroperoxides (ROOH), which are the primary 12 products of autoxidation. Antioxidants (AH) can break this chain reaction by reacting with R00. to form stable radicals which are either unreactive or form nonradical products (Frankel, 1984; and Hawkins, 1985). R00. + AH vi ROOH + A. A. + ROO. ------ > } Non radical products A. + A. ------ > Soybean oil is the major edible oil produced in the world (Buck, 1981). An essential characteristic of this oil is its relatively high percentage of unsaturated fatty acids. Oleic, linoleic, and linolenic‘acid, account for approximately 85% of its fatty acid content (Warner et al., 1984; Sherwin, ' 1976,; Erickson, 1983). Its iodine value (IV), (which indicates its ‘total. chemical ‘unsaturation), is. generally between 125-135 IV. Soybean oil also contains the highest level of linolenic acid esters found in any of the major edible vegetable oils (Buck, 1981). Chemically reactive sites are found in the double bonds of oleic, linoleic and linolenic acids present in the oil. These properties make soybean oil very susceptible to oxidative rancidity (Buck, 1981). Susceptibility to oxidation is greatly reduced by the presence of various antioxidants, synergists and quenchers. 13 The antioxidants are those naturally found and others artificially added. Some of the natural antioxidants include tocopherols, lecithin, gum guaiac, nordihydroguaiaretic acid (NDGA) , citric acid, tannins, ascorbic acid and catechol. Widely used commercial antioxidants are butylated hydroxytoluene (BHT) , butylated hydroxyanisole (BHA) , tertiary butyl hydroquinone (TBHQ) , trihydroxybutyrophenone (THBP) , and propyl gallate (PG). Sherwin (1976) reported that certain organic acids react synergistically with primary antioxidants in vegetable oils. Citric, phosphoric, thiodipropionic, ascorbic, and tartaric acids are used in many countries for this purpose. Lecithin has been cleared for use as an antioxidant or a synergist in vegetable oils, in various countries (Sherwin, 1976). Warner et al. (1987) reported that B-carotene acts as a natural quencher by inhibiting singlet oxidation of unsaturated fatty acids during exposure to light in the presence of sensitizers such as chlorophyll. B-carotene occurs naturally in soybean oil where it inhibits singlet oxygen photooxidation, just as tocopherols are naturally occurring inhibitors of radical chain autoxidation (Vianni, 1980) . Singlet oxygen quenchers may function by reacting chemically with singlet oxygen or by acting as screening agents to reduce the ability of incident radiation to transfer 14 energy necessary to form singlet oxygen. In the absence of quenchers, peroxides formed during oxidation.undergo scission to radical species that.accelerate.the rate and.ultimate level of peroxide formation (Kiritsakis et al., 1985). fl-carotene can be destroyed by exposure to incident light. The loss of B-carotene can leave an oil susceptible to photooxidation if exposed to such light. Sattar et al. (1976) studied the effect of fluorescent light on butter, butter fat, corn, rapeseed and soybean oils. The rate of oxidation increased.considerable with corresponding decrease in vitamin A and B-carotene. This was probably due to the photobleaching of the carotene, which was probably a strong inhibitor of lipid oxidation in the early stages of the process (Sattar et al., 1976). Warner et al. (1987) showed that B-carotene in the presence of 6-tocopherol, behaved synergistically to prevent oxidation in olive oil. This lead them to suggest that tocopherols protect fl-carotene from free radical autoxidation. Lehmann and Slover (1976) studied the relative autoxidative and photolytic stabilities of tocopherols, and observed that a-tocopherol was highly effective in protecting y and 6-tocopherols in methyl myristate and methyl linoleate during photolysis studies with UV light at 254 nm. However, there was only slight protection after 24 hours of exposure. 15 Terao et a1. (1980) observed that during irradiation, B-carotene was rapidly lost from soybean oil. They also discovered that 6-tocopherols prevented the loss of B-carotene under these conditions. However, in the presence of chlorophyll, a-tocopherol and the other tocopherols were rapidly destroyed during photooxidation. This helped to speed the loss of B-carotene and thus hasten the oxidation of the oil. This deduction was also made by Carlsson et al. (1976) when they suggested that tocopherols and other phenolic compounds may act as antioxidants under suitable circumstances, and that tocopherols may also quench singlet oxygen. They reported that photooxidation (via singlet oxygen) is not directly controlled by the antioxidants commonly used to inhibit autoxidation. Warner and Frankel (1987) discovered that tocopherols protect B-carotene from free radical autoxidation. Hildebrand et al. (1984) showed that tocopherols acted synergistically with phospholipids to increase the stability of soy oil. Koskas et al. (1984) showed that a-tocopherol can act as a prooxidant at specific concentration levels. At a concentration of 3.8%, a- tocopherol showed prooxidant characteristics, while at 0.38%, it had antioxidant properties. FF C G T N O D ION Clements et al. (1973) studied the photooxidation of refined soybean oil. They investigated whether singlet oxygen-induced oxidation could be observed in refined soybean 16 oil. Oxidation was followed by measuring the conjugated diene absorption at 234nm. The extent of oxidation was found to be linearly dependent on the time of exposure to the light. Formation of conjugated dienes occurs in oxidized soy oil because of the presence of fatty acids with more than one double bond. In linolenic acid, the methylene group at position 11 is activated by the two adjacent double bonds. The loss of hydrogen from this position produces a pentadienyl radical intermediate that can react with molecular oxygen to produce equal mixtures of conjugated 9- and 13-diene hydroperoxides, such as: 13 12 11 10 9 -c = c - c - c = c- ---—> -c c - c. - c II 0 l (1) -c = c - c. - c = c- ----> -c = c - c= c - c. + -c. - c = c - c = c- (2) -c = c - c= c - c. 02 9 + -c. - c = c - c = c- ----- > -c = c - c = c - c- (3) o o H 13 + -c - c = c - c = c- o o H (Nawar, 1985). Clements et al. (1973) tried to determine the major sensitizers in soybean oil by initiating photooxidation, while protecting the oil with a yellow filter. Light with wavelengths less than 500nm was cut off by using this filter. 17 They also found.that.photooxidation induced by the addition of chlorophyll was reduced, but not eliminated by the filter, which indicated that absorbancy of chlorophyll both below and above 500 nm lead to photosensitization (Clements et al., 1973). Du and Armstrong (1970) from the work of several other authors, stated that the mechanism of light-induced oxidation in fatty materials was not clear, but that. oxidation could.be attributed to ultraviolet light, while the effect of visible light was considered of little or no consequence. They observed that the spectral energy output of the commonly used fluorescent light tubes was primarily in the visible region (350-700 nm), and little ultraviolet light was emitted from these lamps. 'They also found that the unsaturated fatty acids in milk exhibit light absorption only in the far ultraviolet range (about 180 rmn. Even after conjugation (during the first stages of autoxidation) and the resulting shifts in locations of the double bonds during resonance, the absorption region still was about 320 nm. Sattar et al. (1976) reported that Chahine and deMan (1971) found that the most rapid oxidation of corn oil occurred during exposure to fluorescent light. Sattar et al. (1976) also reported that Morgan (1935) found blue light and uv light accelerated the development of rancidity, and that Coe and LeClerc (1935) observed that exposure of butter in the 18 wavelength range of 302 to 546.1 nm resulted in rancidity in all cases, except with green light at 546.1 nm. Sattar et al. (1977) studied the effect of light-induced decomposition of vitamin A in chloroform , B-carotene in hexane and both products in milk fat. The loss of B-carotene and vitamin A was reduced markedly in light-exposed foods by blocking wavelengths below 465 nm. In alcohol, B-carotene has two maxima (452 and 481 nm) in the visible-absorption spectra. In petroleum ether, it has strong absorbance at 273, 453, and 481 nm (Isler et al., 1984). Thus B-carotene receives little protection by blocking UV radiation since it absorbs energy mainly in the visible region of the spectrum. Tocopherols on the other hand, should.be protected by blocking UV radiation, since the UV absorption spectrum for a-tocopherol in ethanol shows maxima at 292 and 256nm (Isler and Kienzie, 1984). Therefore, loss of a-tocopherol (and subsequently the other tocopherols) can be a precursor to the loss of B-carotene, which in turn can result in lipid oxidation in oils and other food products exposed to UV radiation. Emmons et al. (1986) reported that wavelengths of 220 to 550 nm from warm-White fluorescent light bulbs, caused a strong oxidative flavor in butter, whereas, wavelengths above 550 nm did not. Since light has such a profound impact on the keeping quality of edible oils and other lipid containing foods, packaging can play an important roll in its shelf-life. 19 Sattar et al. (1976) reported that relatively simple and inexpensive changes in packaging materials result in the retention of flavor quality and light-sensitive vitamins. F C OF G N O L T V . Leo (1983) stated that the container can influence the absorption of light, oxygen, heat and moisture by the product. The light source can be either artificial, incandescent or fluorescent, or natural sunlight. Of special interest was light in the ultra-violet region, with wavelengths up to 390 nm and the visible region of violet and blue having wavelengths of 390-490 nm. Oxygen was the most critical factor affecting oxidative quality, because of its role in the formation of hydroperoxides, the components normally associated with rancid oil. Oxygen can be in the headspace of a container, permeate its walls or be entrapped in the oil, or other lipid containing foods. Nkpa et al. (1990) studied the effect of various packaging materials on the storage stability of crude palm oil. They showed that lacquered metal cans offered the greatest protection against the deleterious effects of sunlight. Colored bottles gave fairly good protection, while transparent plastic and clear glass bottles resulted in the poorest protection of the oils during storage in direct sunlight. Similar results were obtained by Warner and Mounts (1984) after evaluation of the flavor and oxidative stability 20 of soybean oils in amber and clear glass bottles, and in plastic bottles exposed to light. Luby et al. (1986) determined the effect of packaging on the oxidative stability of cholesterol in.butter exposed to fluorescent and daylight, and found that aluminum foil offered better protection than margarine wrap, opaque parchment, wet strength dry wax paper and polyethylene film. Similar results ‘were obtained by Emmons et al. (1986) who determined the light transmission characteristics of wrapping materials in relationship to oxidation of butter by fluorescent light. Bradley (1980) found that milk in glass, polycarbonate, high density polyethylene, blow-molded polyethylene, plastic bags and paperboard containers, developed a characteristic off-flavor during exposure to fluorescent light or sunlight. Kiritsakis et al. (1984) showed that olive oil packaged in transparent glass and polyethylene plastic bottles had little protection from light. Glass provided more protection than the plastic. 'They suggested that the higher oxidation.rate in the plastic might have been due to the intrusion of oxygen as a consequence of the permeability of the plastic. IGHT BSO R In 1985, Fanelli et a1. investigated the effectiveness of visible and UV light screens, compounded in polyethylene, to protect vitamins in milk from photodegradation by 21 fluorescent light. Six light screens (3 pigments and 3 UV absorbers), were selected for the study, all of which had FDA approval for contact with food. The three pigments created red, yellow and blue colored.resin.and.‘were Quinacridone red, FD&C yellow #5 and Ultramarine blue, respectively. The three UV absorbers were Cyasorb 531'", Tinuvin 326T" and Tinuvin 662"K .At the conclusion of the study, they reported that incorporation of Tinuvin 326 and Cyasorb 531 into the resin significantly reduced vitamin A loss. Tinuvin 622 did not have the same protective value, while the FD&C yellow #5, at a concentration of 0.3%, protected vitamin A for at least 24 hours. None of the UV absorbers protected riboflavin and ascorbic acid. After investigating the absorption spectra of the species involved, they showed that vitamin A had a single strong absorption band with a maximum at 325 nm. Tinuvin 326, Cyasorb 531 and FD&C yellow #5 protected vitamin A because these chemicals absorbed energy in the UV region of the spectrum. Consequently, riboflavin with its strong absorption band at approximately 450 nm was not similarly protected, even though it has several bands in the UV and visible range. Ascorbic acid absorbs in the UV range at wavelengths below 300 nm, but was not protected by the UV absorbers. They suggested that the decomposition of ascorbic acid is dependent on the presence of riboflavin, and concluded that riboflavin acts as a photosensitizer with respect to the decomposition of ascorbic acid. Shipe et al. (1983) obtained similar results when samples of pasteurized homogenized low-fat (1%) milk, 22 fortified with vitamin A, was exposed to light in half gallon polypropylene containers impregnated with either Tinuvin 326, Cromophtal yellow 3G or yellow 2 RLTS. Tinuvin 326 was able to reduce vitamin A loss by about 80%. Crompton (1971) reported that Ultra violet absorbers are often used in food packaging materials to protect the plastic material as well as the foodstuff packaged, from the actinic action of ultraviolet radiation. .Actinic effects may cause discoloration of both the plastic material and the foodstuff, and may also occasion changes in taste and loss of vitamins in the food. Several other UV absorbers including Tinuvin 326, have been used in a number of applications where UV screening was necessary to extend the life of certain non-food materials. Dunn et al. (1966) showed that Tinuvin P and an antioxidant referred to as "antioxidant #2246", improved the stability (against photochemical aging) of natural rubber which had been radiation-crosslinked. Colfico (1966) reported that when Tinuvin 326 was incorporated into a coating applied to photographic! reproductions on paper or film, increased protection from the sun and UV radiation resulted. Heller et al. (1962) demonstrated that incorporation of benzotriazoles into cellulose acetate, nylons, polyesters, polymethyl methacrylate, polyvinyl chloride, polyethylene and non-actinic compounds (for human skin), can protect against the damaging 23 effects of ultraviolet radiation, IMaria et al. (1976) utilized Tinuvin to improve the light stability of polyethylene. Evans et al. (1986) showed that dyed wool can be protected from phototendering and dye fading by the 2-( 2 '- hydroxyaryl)-2H-benzotriazoleultravioletabsorber,sulfonated Tinuvin P"K, Kolawole et al. (1982) proved.that.a synergistic effect was obtained when a combination of Tinuvin P and 3,5- ditert.butyl-4-hydroxybenzyl thioglycollate was used to protect acrylonitrile-butadiene-styrene (ABS) from exposure to weathering. Pyong-Nae (1989) reported that ultraviolet light stabilization of polypropylene and other polymers can be achieved by hindered amine light stabilizers (HAL), phenolic antioxidants and phosphites. Also, a synergistic effect is achieved between HALS and UV absorbers in clear coatings, metallic finishes and sparingly pigmented systems. In general, stabilization of polymers to weathering involves retardation or elimination of primary photochemical processes similar to those involved in lipid photooxidation. This is done by preventing the UV radiation in light from reaching the jpolymer by using a coating or a UV screen, by UV absorbers which preferentially absorb and dissipate the UV energy harmlessly, and by addition of a compound which can remove the excited state energy from the polymer before harmful reaction can occur. This last process in known as quenching. 24 Ultraviolet screens function by rendering the polymer opaque to both visible and UV light (Guillet, 1972). This prevents the penetration of UV radiation into the polymer, and thus reduces degradation. The most important of these is carbon black which is one of the most effective polymer stabilizers (Guillet, 1972). Dispersed throughout a polymer (in addition to acting as a UV screen), carbon black can trap radicals produced during photooxidative processes. Heskins and Guillet (1972) have suggested that carbon black may also stabilize polymers by quenching the excited molecules in the polymer through absorption of UV radiation. The terms UV absorbers and UV stabilizers are often used interchangeably. In the Encyclopedia of Chemical Technology (1984), it states that, "ultraviolet stabilizers are colorless or nearly colorless organic substances which protect polymeric and other light-sensitive materials from degradation by sunlight and artificial sources of UV radiation." Several different classes of compounds are used commercially to retard light-induced polymer degradation, including UV absorbers, hindered amines, nickel chelates, hindered phenols and aryl esters (Dexter, 1984). The most widely used stabilizers consist of derivatives of salicylic esters, benzotriazoles andmorthohyroxybenzophenones (Guillet, 1972). Of the known UV-absorbers, the o-hydroxybenzophenones and the o-hydroxyphenylbenzotriazoles are most frequently used in industrial applications. This is 25 due mainly to their extreme light stability in polymeric substrates (Heller et al., 1972). Gugumus (1984) reported that concentrations of about 0.2 - 0.3% benzophenone and benzotriazole types of UV absorbers incorporated into thick sections of HDPE, led to a considerable improvement in UV stability and extended lifetime of the material. However, with the subsequent development and use of Ni-quenchers the stabilization of thin sections of HDPE was similarly improved. A further step forward in the UV stabilization of HDPE was achieved with the hindered amine light stabilizers (HALS) (Gugumus, 1984). An ideal UV absorber should absorb radiation between 290 and 400 run while transmitting all visible light, as absorption directly above 400 nm causes the absorber to impart a yellowing to the substrate (Dexter, 1984). They should also be light stable and not disintegrate during long-term exposure. Dexter (1984), reported that substituted 2-(2- hyroxyphenyl)benzotriazoles approach this ideal most closely since they absorb very strongly throughout most of the UV region and show a rapid decrease in absorbancy approaching 400 nm. The UV stabilizers (preferably derivatives of o -hydroxy-benzophenone or of 2-(2'-hydroxy-5'- methylphenyl)benzotriazole (Tinuvin) transform the absorbed 26 light energy into thermal energy thus preventing all sorts of photochemically initiated reactions (Werner et al., 1981). The photooxidation of polymeric materials can be represented by a simplified reaction sequence as shown by Dexter, (1984). Initiation: ROOH O H RCR O H RCR O H RC. Propagation: R. + 02 R00. +RH Chain branching: ROOH 2 ROOH A or hv 4 R0. + .OH (1) 0* hv 4 RCR (2) photoexcited chromophore O H 4 RC. + .R (3) 4 R. + CO (4) 4 R00. (5) 4 ROOH (6) 4 R0. + .on (7) hv 4 R0. + Roo. + H20 (8) 27 In the presence of excess oxygen, reactions 5 and 6 can be repeated hundreds of times, which increases the concentration of hydroperoxides. These can cleave homolytically by absorption of thermal or actinic energy to yield additional radicals (equation 7) (Dexter, 1984) . The ROOH in reaction 1, can come from another system undergoing oxidation. UV stabilizers can function to retard the initiation of the photooxidation process by absorption of UV energy and by quenching of photoexcited chromophores. Antioxidants can reduce oxidation by scavenging free radicals or destroying hydroperoxides to yield nonradical species. Dexter (1984) reported that many commercially available stabilizers function by more than one mechanism. Thus, 2-hydroxybenzophenones and 2-(2'-hydroxyphenyl)benzotriazoles contribute to polymer stability by absorbing UV radiation, trapping free radicals, and quenching photoexcited chromophores. Their ability to trap free radicals also gives them antioxidant properties. The reaction of oxygen with unsaturated lipids and polymers to cause oxidation, is basically the same. The mechanism is initiated by the removal of hydrogen from a hydrocarbon molecule, which can occur in the presence of metals, light, heat, or hydroperoxides. The mechanism by which 2-(2'-hydroxyphenyl)benzotriazoles dissipate absorbed radiant energy involves tautomeric structures ( 1) and (2) (Werner et al., 1981). 28 CH3 CH3 ‘\\ /N : “x /N -A N(+ ) A \ “—0 H ooooo o(_) (1) (2) Fig 1. The mechanism of light energy absorption by a benzotriazole. In the ground state, the phenolic structure (1) is preferred since the electron density on the oxygen atom is much greater than that on the triazole nitrogen. If UV energy is absorbed, the electron density shifts from the oxygen atom toward the triazole ring. This results in an increase in basicity of the nitrogen atom and causes the proton to jump from oxygen to the nitrogen. Tautomer ( 2) is unstable and rapidly loses its energy as heat, and reverts back to the ground-state structure (1) . This process is efficient and accounts for the excellent light stability of 2-(2 '-hydroxyphenyl) -benzotriazoles as well 29 as their inactivity as photosensitizers (Dexter, 1984). W Tinuvin” 326 is a UV absorber produced by the Ciba Geigy Co. and is chemically known as 2-(3'-tert-butyl-2'- hydroxy-5'-methylphenyl)-5-chlorobenzotriazole. It is a derivative from a benzotriazole. Tinuvin 326, a benzotriazole derivative, has a chloride molecule on the 5th position of the basal benzene ring and a tertiary butyl functional group at the #3 position of the functional benzene ring. Hence the appendices 3'-t-butyl and 5-chloro- are incorporated into the chemical name of the basic 2-(2 '-hydroxyphenyl) -benzotriazole, to give (Tinuvin 326) 2-(3'-t-butyl-2'-hydroxy-5'- methylphenyl)-5-chlorobenzotriazole (Schwemmer, 1990). Capocci 1990 reported. that. Tinuvin 326 shows a :maximum absorbancy at approximately 345 nm. A similar maximum absorbancy was found by Werner et al. 1981 for Tinuvin in a 20:20:1 ethanol/ether/pyridine solution. CH3 CI N ’1’ \ N ‘\~ / N‘ .H—O C(cn-Ia)3 Tinuvin 326 Fig 2. The structure of Tinuvin 326 30 The additional side chains give it various advantageous properties. Heller et al. (1962) suggested that these groups may interfere with its steric hindrance and thus rotation of the molecule in space. Dexter (1984) mentioned that such properties can increase the solubility of the UV absorber in the molten polymer, which would reduce exudation of the additive to the polymer's surface after manufacture of the material. In addition to minimizing volatility, these functional groups help to reduce reaction of the UV absorber with metal ions such as Co“, Zn”, and Cd“. Such reactions would otherwise lead to chelate formation and result in the production of colored complexes. ZPure Tinuvin 326 is a yellow solid, with melting point 137-141°C and a decomposition temperature >220°C (Schwemmer, 1990). Tinuvin 326 was approved by the FDA as a food packaging additives in April 1981 (Title 21 of the Code of Federal Regulations). The incorporation of Tinuvin 326 in containers to package oils and other food products can increase the stability of both the container and the product. This occurs due to its ability to absorb UV radiation and its antioxidant property. By reducing exposure of the oil to UV radiation, lipid oxidation can be reduced, lMigration of Tinuvin 326 from the container to the oil could serve to further inhibit lipid oxidation. This technique has been successfully used in the pasta ‘Hoojjat et al. (1988) showed that the migration of BHT, an antioxidant, from high density polyethylene film into a 31 food product, improved the oxidative stability of the packaged oatmeal cereal. HAIEBIAL§_A!2_!§I§QQE Test packages: The test packages used in this study were 127 ml (4.5 oz), 0.381 mm (15 mil) thick, transparent, coextruded multilayered polypropylene cups which were provided by The Ball Plastics Division, Plastics Packaging Development Center, Evansville, IN. The material profile of the containers from outside to inside was: polypropylene 95.25 pm (3.75 mil) / regrind 80.01 pm (3.15 mil) / adhesive 9.53 pm (0.375 mil) / 44 mol. % ethylene vinyl alcohol copolymer 15.24 pm (0.6 mil) / adhesive 9.53 pm (0.375 mil) / regrind 80.01 um (3.15 mil) / polypropylene 91.44pm (3.60 mil). One set of containers had zero percent, while the other had 0.3% Tinuvin 326 incorporated into the regrind layers, respectively. The Alusuisse‘" "Easy Peel" lid stock was also provided by The Ball Plastics Division. It had a peel force of 1.36 to 1.81 kg. (3.0 to 4.0 pounds) and a break force of 2.04 to 2.9 kg. (4.5 to 6.0 pounds). Soybean oil: Five gallons of refined Soybean salad oil (Mikadom) were obtained from the General Food Stores Michigan State University. The label indicated that the oil contained no added antioxidants. The oil was stored at room temperature 32 33 (21°C) prior to use. Light source: The light source used was a Phillips, cool white, 120 volt fluorescent lamp. It had a power rating of 34 watts and a length of 120 cm. It was positioned in a light box and was used to illuminate the bleached oil samples in the containers with and without Tinuvin 326, prior to determination of the peroxide value of the bleached oil samples. R to Extraction of Tinuvin 326 from sample containers: Analysis for Tinuvin 326 in the containers was carried out using' HPLC and 'UV spectrophotometric ‘methods. The procedure involved cutting the walls of a sample container into approximately 1 cm square pieces. An average sample size of 3 g was used for each analysis. Extraction was carried.out with 110 ml HPLC-grade acetonitrile (Fisher Scientific Inc.), which was mixed with the sample and the sample extracted in a Soxhlet extraction apparatus for 18 hours. On completion, the acetonitrile containing the extracted Tinuvin 326, was made up to 200 ml in a volumetric flask with HPLC grade acetonitrile. The initial sample was extracted two times more, and on each occasion 110 ml of pure HPLC grade acetonitrile was added. For each of these, the extract were made up to 100 ml in a volumetric flasku These three extracts were individually analyzed for Tinuvin 326 using UV spectrophotometric and HPLC procedures, at a wavelength of 347.3 nm. Two replicates were taken for each analytical period, and the analysis duplicated for each replicate when using the HPLC method. The result of the quantification of Tinuvin 326 in each extract was added and the total represented the level of Tinuvin 326 in the sample material. 35 HPLC procedure to determine Tinuvin 326 in extract: An HPLC technique was used to measure the Tinuvin 326 in the containers. A reverse phase system was used with a Perkin-Elmer stainless steel column, 0.24 cm internal diameter and 25 cm in length. The packing material of the column was comprised of ODS-HC sil-x-l. The mobile phase was comprised of 85% acetonitrile/15% distilled water (v/v) at a flow rate of 1 ml/minute. The volume injected was 10 ul using a Hamilton microliter #701-N Syringe. Peak areas and retention times were determined using a computing integrator. The elution 'time of 'Tinuvin 326 was approximately 4.85 minutes. The high pressure liquid chromatograph (HPLC) system consisted of a Perkin-Elmer Series 38 Solvent Delivery System and a LC-100 column oven, with a Perkin-Elmer LC-75 spectrophotometric detector set at 347.3 nm. The detector was interfaced to a Spectra Physics SP4200 Computing Integrator for quantitation. The concentration of Tinuvin 326 in the containers was determined from standard curves constructed by analyzing Tinuvin 326 solutions of known concentrations in acetonitrile. The standards were prepared by dissolving 0.01 g Tinuvin 326 powder in 25 ml acetonitrile and making up to 100 ml in a volumetric flask, to give a concentration of 100 ppm (w/v) Tinuvin 326. The Tinuvin 326 powder was supplied by Ciba- Geigy Corporation, Hawthorne, New York. Serial dilutions from this stock solution were made to give solutions of 1, 10, 60, 36 80 and 100 ppm (w/v), respectively. Peak areas and retention times at these concentrations were used to construct the standard curve. UV spectrophotometric procedure to determine Tinuvin 326 in extract: A Perkin-Elmer Lambda 43 Double Beam UV-visible spectrophotometer was also used to measure the Tinuvin 326 content of the acetonitrile extracts. The optical density at 347.3 nm, and l-cm path length, of each extract, was obtained after pouring approximately 3 ml into a quartz cuvette, then placing it in the cuvette holder of the spectrophotometer. The concentration of Tinuvin 326 in the containers was determined from standard curves constructed by analyzing Tinuvin 326 solutions of known concentrations in acetonitrile. The standards were prepared by dissolving 0.01 g Tinuvin 326 powder in 25 ml acetonitrile and making up to 100 ml in a volumetric flask, to give a concentration of 100 ppm (w/v) Tinuvin 326. Serial dilutions from this stock solution were made to give solutions of 1, 5, 10, and 15 ppm (w/v), respectively. Absorbance in optical density units of these concentrations were used to construct the standard curve. The efficiency of the extraction method was investigated by obtaining an absorbance spectrum, between 37 200 - 400 nm, of the container wall before the Soxhlet extraction. Another spectrum was obtain for the same container sample after the Soxhlet extraction procedure. This was done using a Perkin-Elmer (Oak Brook, Illinois) Lambda 38 UV-visible spectrophotometer which is equipped with an integrating sphere. Three samples from the 0% and three from the 0.3% Tinuvin 326 containers were cut approximately 3.0 cm x 1.5 cm and mounted directly in the sample holder of the Integrating Sphere. This instrument was used to monitor the relative change in concentration of the Tinuvin 326 in the container wall following repetitive extractions. Loss of Tinuvin from the containers: The rate loss of Tinuvin. 326 from the test containers was determined using both HPLC, and UV spectrophotometric methods, by monitoring the change in Tinuvin content in the containers over a period of 42 days, at 21 :1 and 35 11°C. respectively. The containers at 35°C were stored in a Precision Scientific, model 324 oven. The other set of test containers was stored in the laboratory, which had an average ambient temperature of 21°C :2. The containers at both temperatures were stored in the absence of light. Migration of Tinuvin from the containers into the oil: The migration of Tinuvin 326 from the containers was studied over a 42 day period at 21 :1 and 37 11°C. 38 respectively. The samples were prepared by filling 120 ml of refined soybean oil into each of thirty 0.3% Tinuvin 326 containers. Each container was covered and heat sealed with the Alusuisse'" "Easy Peel" aluminum foil. Heat sealing was done at 182.2°C (360°F) and 275.8 kPa (40 psi) for 10 seconds using an Alloyd (Dekalb, Illinois) Blister-pack sealer. Two of the oil filled containers were removed weekly from storage, and the concentration of Tinuvin 326 in the oil determined over a 42 day period. The container material was also analyzed for'rinuvin 326 using both UV spectrophotometric and HPLC methods over the same 42 days period. Analysis for Tinuvin 326 in the oil was carried out using HPLC. The procedure involved removal of the container lids and stirring the oil to ensure a homogeneous mixture. From each sample, 3 ml of oil was removed with a disposable pipette and measured in a 10 ml volumetric cylinder. This was extracted with 15 ml of acetonitrile in a 50 ml separatory funnel. The mixture was shaken vigorously by hand for 20 - 30 seconds. The acetonitrile layer rose to the top after standing for several minutes. This layer was drawn off using a long- nose disposable pipette and transferred to a 100 ml test tube with a rubber cap. This extraction procedure was repeated 4 more times on the same oil sample, using 15 ml of acetonitrile 39 on each occasion. The extractants were combined and then centrifuged. (International. centrifuge, model CM, size 1, centrifuge) at 2000 rpm for 15 minutes to effect further separation of any residual oil. The acetronitrile solution above was drawn off using a long-nose disposal pipette, taking care to leave the small quantity of oil at the bottom of the test tube. The resulting solution (approximately 75 ml) was then concentrated to 5 ml using a rota-evaporator (Buchi Rota vapor, model No. 110813, rota-evaporator). The 5 ml concentrate was then analyzed for Tinuvin 326 using the HPLC procedure at a wavelength of 347.3 nm. The concentration of Tinuvin 326 in the oil was calculated from the standard (Tinuvin 326) curve. The efficiency of the extraction method was investigated by analyzing a 40 ppm solution of Tinuvin 326 in soybean oil. Tinuvin 326 was extracted from the oil by the method described above, and the amount of Tinuvin 326 which was extracted, determined. using the HPLC method. Percent recovery was obtained by comparing the amount remaining in the oil to the initial 40 ppm. The efficiency of the concentration step was investigated by concentrating a 75 ml solution of 10 ppm Tinuvin 326 to 5 ml using the Buchi rota-evaporator. The amount of Tinuvin 326 in the concentrate was determined using the HPLC method. Comparing the calculated concentration to the original 10 ppm, provided the percent recovery. 40 Bleaching the soybean oil: The soybean oil was filtered through an ion exchange column of internal diameter 4.5 cm and a length of 50 cm. Made of glass, the column was prepared by first packing a 2.0 cm layer of glass wool at the bottom. On to this was placed a 2.0 cm layer of MN-Kieselgur G-HR (Macheney, Nagel and Company) , then a 100 g mixture containing 50% activated carbon Darco S-51 grade (American Norit.Company), 35% MN-Kieselgel G- HR. and. 15% Florisil adsorbent for chromatography' (Fluka Chemika). Finally, two successive layers of silicic acid and anhydrous sodium sulfate, each 1.0 cm , were added to complete the packing. The column was packed under reduced pressure using a vacuum pump with a negative pressure of 248 kPa (36 psi), and pressed with a glass rod to ensure proper compaction. The respective packing materials were added to the column in small amounts in slurried form by mixing hexane with the individual materials. After passing 100 ml of hexane through the packed column, 100 g of the soybean oil were mixed with 150 ml of hexane, and poured onto the top of the column. After this mixture was sorbed by the column, 150 ml of hexane were used to elute the oil. The eluate, which contained the bleached oil and hexane, was collected in a 500 ml filtering flask,and transferred to a 500 ml round bottom flask. The hexane was evaporated using a rotary evaporator under vacuum at 30°C. The oil obtained was transferred to an amber glass bottle, flushed with nitrogen and stored at 0°C. 41 The column was regenerated for multiple use by eluting 150 ml each of anhydrous ethyl ether (Em Science), acetonitrile (EM Science), methanol (EM Science), acetonitrile, anhydrous ethyl ether and finally hexane (EM Science) in successive order. This was done after each batch of oil was bleached. Bleaching of the soybean oil was necessary to remove all its natural antioxidants. This was done to insure relatively rapid oxidation of the oil during exposure to fluorescent light. The thoroughness of the bleaching procedure was determined by Kalsec Inc. Kalamazoo, Michigan. Lipid oxidation was determined on three samples of unbleached soybean oil, using' a .Metrohm, Model 679 Rancimat at a temperature of 110°C. For unbleached oil, an induction time of 6 hrs was found. Using the same test for three samples of bleached soybean oil, an induction time of 0.95 hr was found. A shortening' of the induction time indicates a greater susceptibility to oxidation. Irradiation System to expose the bleached oil to fluorescent light: A total of ninety containers with 0% Tinuvin and ninety containers ‘with 0.3% Tinuvin. were used for exposure 'to fluorescent light. Into each of these containers, 15 ml of the bleached soybean oil were added under atmosphere pressure. These were covered and heat sealed with the Alusuisse ”Easy 42 Peel" aluminum foil sheets, using the Alloyd Blister sealer. Heat sealing was done at 182.2°C (360°F) and 275.8 KPa (40 psi), for 10 seconds. During illumination, samples were divided into two equal groups (90 samples each). Ninety samples, composed of forty five containers with 0% Tinuvin 326 and forty five containers with 0.3% Tinuvin 326 were placed into each of two light boxes 91 cm x 61 cm x 53 cm (36" x 24" x 21"). One box was stored at 21 and the other at 35°C for a total of 35 days. The containers were suspended near the inner roof of each box. This was accomplished by hanging them from their heat sealing lip, between rows of strings stretched between the side walls (near the roof) of each box. The fluorescent tubes were placed under the boxes a distance of approximately 64 cm (25") below the containers. The light entered a box through an adjustable slit at the bottom which controlled its intensity. Thus, the oil in the bottom of the containers was exposed to the fluorescent light. A light intensity of 538 Lux (50 foot candles) was selected following a survey of grocery aisles in several retail food stores in the East.Lansing, Michigan area. A General Electric, model J-55, light meter was used to measure the light intensity. The boxes were kept in dark rooms to eliminate light from other sources“ (Over the 35 day period (days 1,2 4,7,14,21,25,28,32 and 35), two samples were removed of each container type, at each temperature, and the extent of lipid oxidation in the oil determined by the 43 peroxide value method (AOAC, 1984). Lipid oxidation determination by peroxide value method: Lipid oxidation of the soybean oil, as a function of storage time and temperature and level of Tinuvin 326 in the containers, was determined using the official AOAC method (1984). Peroxide value is reported as milliequivalents peroxide/kg soybean oil. Approximately 5.0 g of oil accurately weighed, were taken from the sample containers following light exposure, for each analysis . The oil was transferred to a 250 ml Erlenmeyer flask and dissolved in a 30 ml acetic acid - chloroform (3:2) mixture. To this solution was added 0.5 ml of saturated potassium iodide solution, followed by mild agitation for 1 minute. Thirty ml of distilled water were then added and the solution titrated with 0.01N sodium thiosulfate solution. One and a half ml of a 1.0% starch solution were used as the indicator, and the titration carried out until the blue color had just disappeared. The peroxide value (milliequiv. peroxide/kg sample) was obtained by the following formula: = Vol. (m1) sodium thiosulfate titrated x normality x 1000 PV weight of oil (9) BEE!LI§.A§D.DL§Q!§§IQ§ Standard curves and absorbance spectra profiles: Area response units vs concentration of the standard solutions (HPLC method) are summarized in Appendix 1. The standard curve is shown in Figure 3. The absorbance of the standard solutions vs concentration (Perkin-Elmer Lambda 48 Double Beam UV-visible spectrophotometric method) is presented in Appendix 2, while the standard curve is shown in Figure 4. The slope of the regression line for the HPLC standard curve was calculated to be 3.998 x 103. This was used as a conversion factor in calculating the concentration of Tinuvin 326 in the container walls and in the packaged oil, from the area response units obtained using the HPLC method. The slope of the regression line for the UV spectrophotometric standard curve was 5.191 x 10'2. The regression line is described by the following equation: y=S . 191x10" ( .) +2 . 347 x10" * = concentration of Tinuvin 326 in ppm (w/v) This was used to calculate the concentration of Tinuvin 326 in the container walls by converting optical density units from the UV spectrophotometric method into ppm Tinuvin 326 (w/v). 45 The absorbance spectra of the wall of the containers were obtained using a Perkin-Elmer Lambda 3B UV-visible spectrophotometer equipped with an integrating sphere. The absorbance spectra of the wall of the 0.3% Tinuvin 326 containers before and after extraction are shown in Figures 5 and 6, respectivelyu 'The absorbance spectrum of the blank (0% Tinuvin containers) is shown in Figure 7. JFigure:6 and Figure 7 show almost identical absorbance spectrum. Since Figure 6 presents a spectrum of the containers with 0% Tinuvin 326, it was concluded, based on comparison with Figure 5, that the dispersed Tinuvin 326 in the container wall was removed during the Soxhlet extraction process. The wavelength at which maximum absorbance occurred, was determined from the profile illustrated in Figure 5. From this figure, the maximum absorbance of Tinuvin 326 was determined to be at 347.3 nm. At this wavelength, the profile of the containers with 0% Tinuvin 326 shows no absorbance. The profile of the container wall (0.3% Tinuvin) after extraction also shows no absorbance at 347.3 nm. Thus, it was concluded that 347.3 nm ‘was the wavelength of maximum absorbance for Tinuvin 326. Similar spectra were obtained for solutions of acetonitrile containing 0.0% Tinuvin 326 and 10 ppm (w/v) Tinuvin 326. These were obtained using a Perkin-Elmer Lambda 48 Double Beam UV-visible spectrophotometer. Dexter, (1984) 46 reported a similar absorption spectrum for 1.0 mg/100 ml 2- (2'-hydroxy-5-methylphenyl)benzotriazole 1J1 chloroform. Yushkevichyute and Shlyapnikov, (1967) also determined the concentration of Tinuvin 326 in polyethylene using a spectrophotometric method. They found maximum absorbance at wavelengths of 276 and 296 my in a heptane solution. Area response units 47 584-05 49-05- ".3 I 1'0 2'0 3'0 4330 0'0 70 0'0 9'0100 Concentration Tinuvin 326 (ppm) (VI/V) Fig. 3 Standard curve of Tinuvin 326 concentration vs area response units (HPLC method). Absorbance (optical density units) 1.0 48 0.9- 0.84 0.74 0.8- 0.5- 0.4-1 0.3- 002"" 0.1- 0.0 o 5 13 1s r Concentration Tinuvin 326 (ppm) (w/v) Fig. 4 Standard curve of Tinuvin 326 concentration ~ vs absorbance (optical density units) (UV spectrOphotome’tric method). 49 M 1 d l Absorbance (optical density units). 200 360 400 Wavelength (nm). Fig. 5 Absorbance spectrum of container wall before extraction to remove Tinuvin 326. 50 N 1 d I Absorbance (optical density units). 200 360 400 Wavelength (nm). Fig. 6 Absorbance spectrum of container wall after extraction to remove Tinuvin 326. 51 I») 1 up 1 Absorbance (optical density units). 0 r Wavelength (nm). Fig. 7 Absorbance spectrum of container wall with 0.0% Tinuvin 326. 52 Loss of Tinuvin 326 from the container material: In Tables 1 and 2 are summarized the percent Tinuvin 326 remaining in the container material during storage at 21°C., from zero to 42 days using the HPLC and UV spectrophotometric methods, respectively. The percent of Tinuvin 326 remaining in the containers during storage at 35°C., is shown in Tables 3 and 4. Graphs comparing results from the different storage temperatures are shown in Figures 8 and 9 (HPLC) and Figures 10 and 11 (UV spectrophotometric) respectively. The concentration of Tinuvin 326 in the container wall (HPLC method), was calculated from the area response (au) units and the slope of the standard curve. An example of the calculation is presented in Appendix 3. Tables 5 and 6 show the data used to calculate the percentages of Tinuvin 326 at storage temperatures 21 and 35°C respectively. These are summarized in Appendix 4. The concentration of Tinuvin 326 in the container wall was also calculated from the optical density units and the line equation of the standard curve (UV spectrophotometric method). An example of the calculation is presented in Appendix 5. The data used to calculate the percentages of Tinuvin 326, using the UV spectrophotometric method, is shown in Tables 7 and 8 for the containers at storage temperatures 21 and 35°C respectively. These Tables are included in 53 Appendix 6. The results obtained from the HPLC and UV spectrophotometric methods are essentially identical. This indicates that either method could be used to quantify Tinuvin 326 in plastics such as used in this study. The results show little or no loss of Tinuvin 326 from the containers at either temperature during storage for 42 days. Thus, there was a high level of stability of the absorber in the 0.381 mm thick polypropylene containers. UV spectrophotometric analysis was used by Yushkevichyute and Shlyapnikov ( 1967) to quantify levels of Tinuvin 326 (in heptane solutions) in 0.16 mm thick polyethylene. They found that an increase blooming of Tinuvin 326 occurred at the surface of the film at higher temperatures. After approximately 400 hours, the level of Tinuvin 326 stabilized at temperatures of 20, 30, and 40°C. The differences in these results and those obtained in this study may be attributed to the difference in the thickness of the plastic, the material type and the manufacturing process. In this study the Tinuvin 326 was dispersed into the regrind layers of the laminate and not exposed to the surfaces of the material. In the Yushkevichyute and Shlyapnikov (1967 ) study, Tinuvin 326 was incorporated into the entire mass of the plastic. The results from the analysis for Tinuvin 326 in the containers stored at 21 and 35°C, using the HPLC and the UV 54 spectrophotometric methods, show averages of 68.53% recovery for the first Soxhlet extraction, 21.76% for the second and 9.72% recovery for the third Soxhlet extraction. Initially an average concentration of 0.00515% Tinuvin 326 was found in a fourth Soxhlet extraction. Since this represents approximately 1.5% of Tinuvin 326 in the containers, the results from three extractions were used to quantify the levels of Tinuvin 326 in the material. 55 Table 1. Relative percent of Tinuvin 326 in the multilayer containers stored at 21°C. (HPLC method). Storage time : Average % Tinuvin (days) : (absolute) """"m""3"'""""""""STSB """"""""""" 3 0.31 7 0.30 14 0.32 21 0.29 28 0.31 42 0.30 56 Table 2. Relative percent of Tinuvin 326 in the multilayer containers stored at 21°C. (UV spectrophotometric method). """"""""" s ”£55.;Iii;"T"'L222}:E'E'EJEJ'm'"" (days) : (absolute) ------------ — —____0 ————_—- —=— 0.30 __- — 3 0.29 7 0.31 14 0.32 21 0.30 28 0.31 42 0.30 57 Table 3. Percent Tinuvin 326 in the multilayer containers stored at 35°C. (HPLC method). Storage time : Average % Tinuvin (days) l (absolute) --------------------0---------= —_ 0:30 —_— — —_—— 3 0.29 7 0.30 14 0.33 21 0.31 28 0.31 42 0.31 58 Table 4. Percent Tinuvin 326 in the multilayer containers stored at 35°C. (UV spectrophotometric method). Storage time : Average % Tinuvin (days) : (absolute) """"""""""" o” .__-_----..--._-------.o...3.1.---_---___-....--- 3 0.30 7 0.29 14 0.31 21 0.31 28 0.32 42 0.32 Percent Tinuvin in containers 59 0.4 OJW 4 0.2- 0.1d 0.0 T 0 i ' 14 2'1 2'8 3'5 42 Storage time (days) Fig. 8 Percent Tmuvin in containers Stored at 21 degrees C. by HPLC method. Percent Tinuvin 326 in containers. 60 0.4 0.3% i I” 0.2- 0.1 - 0.0 1 I I u 1 0 7 14 21 28 35 42 Storage time (days). Fig 9. Percent Tinuvin 326 in multilayer container wall stored at 35 degrees C. by HPLC method. Percent Tinuvin in containers 61 0.1 - 0.0 ~11 14 2'1 2'8 Storage time (days) 3'5 42 Fig. 10 Percent ‘linuvin in containers stored at 21 degrees C. (UV spectrophotometric method). 62 0.1~ 000 I I r I I 0 7 1 4 21 28 35 42 Storage time (days). Percent 'linuvin 326 in containers. 0 '9 Fig 11. Percent Tinuvin 326 in containers stored at 35 degrees C. by UV spectr0photometric method. 63 Migration of Tinuvin 326 from the container wall into the oil: The amount of Tinuvin 326 which migrated from the ,container wall into the oil at 21 and 35°C respectively, at successive storage times (HPLC method) is illustrated in Tables 9 and 10. In Figure 12, the effect of temperature and storage time on the amount of Tinuvin 326 migrating from the container wall to the oil is shown. Tables 11 and 12 display the level of Tinuvin 326 in the wall of the containers at the beginning of the study, and the level of Tinuvin 326 following storage at 21.and 35°C, respectively (HPLC method). Pictorial representation of Tables 11 and 12 are seen in Figures 13 and 14, respectively. The percent Tinuvin 326 that migrated from the containers to the oil at the two storage temperatures, is shown in Tables 13 and 14. Table 15 shows the percent recovery for extraction of Tinuvin 326 from the oil after the various storage periods. It also shows the efficiency level of the rota-evaporation concentration procedure. The rate of migration of Tinuvin 326 from the container wall to the oil was higher at 35°C than at 21°C. This indicates that at elevated temperatures, Tinuvin 326 is more likely to migrate than at lower temperatures. However, the level of Tinuvin 326 in the container wall remained relatively constant even at the higher temperature. These two sets of results seem to be in conflict at first glance. However, the absolute quantities of Tinuvin 326 which migrated from the (containers to the oil were very low, especially at.21°C. The 64 maximum level of Tinuvin 326 transferred to the oil was 4.77% of the initial level of Tinuvin in the containers. This is calculated as: (9.009x10"g)x4.77= 4.2973x1o“ 100 g Wt. Tinuvin”,5 transferred = 9.009 x10’3 = initial level of Tinuvin326 in the container wall. This occurred after 42 days and at a 35°C storage temperature. At 21°C, no Tinuvin 326 was detected in the oil before seven days of storage. Thus at room temperature, the migration of Tinuvin 326 from the polypropylene container to soybean oil could not be detected by the methods used during the first week of storage. Thus Tinuvin 326 exhibits stability in the plastic containers in the presence of soybean oil. The efficiency of the extraction process to remove Tinuvin 326 from the oil was determined to be 81%. This was determined from the level of Tinuvin 326 recovered from a spiked sample of the oil (40 ppm Tinuvin 326), using the extraction procedure employed to remove migrated Tinuvin 326 from the oil. From this, a conversion factor was used to convert the experimental value to a theoretical value which represents 100% extraction. The efficiency determined for the rota-evaporator (97.78%), was assumed to be equivalent to 65 100%, so no factor was employed in calculation of the amount migrating. This efficiency was determined from the level of 40 ppm Tinuvin 326 recovered from a 75 ml solution which had been concentrated to 5 ml, using the same rota-evaporator. An example of the calculations used to determine the amount of Tinuvin 326 which migrated to the oil after 14 days at 21°C, is shown in Appendix 7. 66 Table 9. Amount of Tinuvin 326 which migrated from the container wall to the oil at 21°C, (HPLC method). storage time Ave. conc. Wt. Tinuvin I I (days) E Tinuvin in oil E 120 ml oil 1 (Vt/V01) l (9) '" 7 ‘ """""""" 5T3 """"""""""" 5 T5" """ 14 1.300 x10'5% 1.550 xlo'sg 21 2.400 x10'5% 2.80 x10'59. 28 7.200 x10'5% 9.300 xlO’Sg 42 5.300 x10'5% 6.400 x10'5g 67 Table 10. Amount of Tinuvin 326 which migrated from the container wall to the oil at 35°C, (HPLC method). storage time Ave. conc. Wt. Tinuvin i l (days) i Tinuvin in oil E 120 ml oil = ‘ WWI) J _‘_9_’__ ............l --------------- 3:00 x10'5%—— 3.750 x10‘59 14 7.600 x10'5% 9.050 x10'5g 21 9.800 x10'5% 11.750 xlo‘sg 28 16.500 x10'5% 19.850 xlO'Sg 42 43.000 x10'5% 35.800 x10'5g 68 Table 11. Level of Tinuvin 326 in the wall of the containers in which oil was stored at 21°C (HPLC method). Storage time i % Tinuvin 326 / 100g l Ave. Wt. Tinuvin 326 (days) H container wall : in the walls of the containers 0 0.30% 9.009 x10"3 7 0.34% 10.271 x10'3 14 0.32% 9.791 x10"3 28 0.31% 9.365 x10'3 9 9 9 21 0.31% 9.567 x10"3 9 9 42 0.33% 9.968 x10‘3 9 69 Table 12. Level of Tinuvin 326 in the wall of the containers in which oil was stored at 35°C (HPLC method). Storage time i % Tinuvin 326 / 100g :* Ave. Wt. Tinuvin 326 (days) : container wall : in the wall of the containers """3 """"""""""""" 5 367mm: _ “9.009 x10:3-_g_- 7 0.31% 9.520 2110’3 g 14 0.32% 9.911 x10‘3 9 21 0.32% 9.600 x10"3 9 28 0.32% 9.790 1110'3 g 42 0.30% 9.113 x10'3 9 * Average of two replicates (two injections per replicate). 70 Table 13. Percent Tinuvin 326 which migrated from the container wall to the oil at 21°C, (HPLC method). storage times : (days) 1 """""""""""" 7 --.._-___-_-----------_-_--_.o...6.;__--------.. 14 0.172% 21 0.311% 28 0.955% 42 0.710% Weight of Tinuvin 326 in the wall of the container storing the oil at zero time = 9.009 x10'3g / container. * Relative % Tinuvin transferred from the initial weight of Tinuvin in the container wall. 71 Table 14. Percent Tinuvin 326 which migrated from the container wall to the oil at 35°C, (HPLC method). storage times : Relative % Tinuvin* transferred (days) 1 """"""""""" 7 ---..-..--------..-----....-..---.o...4.1.;%.--..-----_ 14 1.005% 21 1.304% 28 2.203% 42 4.773% Weight of Tinuvin 326 in the wall of the container storing the oil at zero time = 9.009 xlo’ig / container. * Relative % Tinuvin transferred from the initial weight of Tinuvin in the container wall. Percent linuvin 326 in oil (w/v) 72 Fig. 12 Percent Tinuvin 326 which migrated from the containers to soybean oil at 21 and 35 degrees 0 (HPLC method). Percent 'iinuvin 326 in container walls 73 0.4 0.2- 0.1- 0.0 . r r r . 0 7 14 21 28 3s 42 Storage time (days). Fig. 13 Percent iinuvin 326 in wall of containers after oil storage at 21 degrees C (HPLC method). Percent ‘linuvin 326 in container walls 74 0.4 0.39”""'—'-’—L : i 0.2- 0.1- 0'00 7 1'4 21 2'8 3'5 42 Storage time (days). Fig. 14 Percent Tinuvin 326 in wall of containers after all storage at 35 degrees 0 (HPLC method). 75 Table 15. Percent efficiency of the extraction process and the rota-evaporation concentration procedure. Std. extraction % efficiency % efficiency I I l I trials I (extraction process) I (rota-evaporation) 1 79.66% 97.98% 2 81.12% 97.57% 3 82.03% 97.80% Average 81.00% 97.78% 76 Lipid Oxidation Determination by Peroxide Value Method: The peroxide value method to determine the extent of lipid oxidation in the oil samples, was chosen because of its short analytical time, simplicity and ease of reproducibility. This method of analysis has been used successfully by many researchers to determine the oxidation levels of oil samples (Vianni, 1980; Luby, 1982; Boki, 1989). The peroxide values of the oil packaged in containers with and without Tinuvin 326, and stored at 21°C for a duration of 35 days, are shown in Table 16. Two replicates were used for each. data point. .All observations were duplicated and the values reported are averages. The peroxide values for the oil in containers with and without Tinuvin 326 and stored at 21°C, increased similarly though relatively slowly during the first fourteen days of storage (Fig. 15). After fourteen days, however, the peroxide value for the oil in the containers without Tinuvin 326 increased at a faster rate than for the oil in the containers with Tinuvin 326. Analysis of variance, using a complete randomized design (Appendix 8), shows that there is a significant difference between the peroxide values of the oil stored in the containers with Tinuvin 326, and those stored in containers without Tinuvin 326. This shows that Tinuvin 326 protected the packaged oil from the effects of the fluorescent light at 21°C. During the 0 - 14 day period, the initiation step of the lipid oxidation process probably occurred. During this 77 period, lipid oxidation is relatively slow, but increases dramatically during the next phase (propagation stage). During the propagation stage, it is easier to observe (Fig 15) the effect of the UV absorber as it protects the oil from the influence of the fluorescent light. 78 Table 16. Peroxide value (milliequivalents/kg oil) of the soybean oil in the multilayered container with 0% and 0.3% Tinuvin 326, stored at 21°C Storage time :Peroxide value of oil :Peroxide value of oil (days) (in 0% Tin. containers :in 0.3% Tin. containers 0 0.7985 0.7985 1 1.9462 1.7972 2 2.7208 2.4966 4 4.2665 3.8647 7 5.5849 4.5615 14 8.9848 8.3997 21 16.3081 10.3914 25 18.3367 12.8114 28 20.2853 13.0719 32 22.9514 16.4586 35 25.9298 17.1901 79 70.00 ‘ s—o peroxide value of oil in 0.0:: Tinuvin containers 60 00 H peroxide value of oil in 0.32 iinuvin containers 50.00] Peroxide value (milliequiv./kg oil) T 2'1 28 35 Storage time (days) 0 ; 1'4 Fig. 15 Peroxide value of soybean oil in the containers 0% and 0.3% Tinuvin 326 stored at 21°C. 80 The peroxide values of the oil packaged in containers with and without Tinuvin 326, and stored at 35°C for a duration of 35 days, are shown in Table 17 and presented graphically in Figure 16. The results are similar to the values obtained at 21°C, except, at 35°C, the initiation step seems to be between 0 - 7 days. Tinuvin 326 protected the oil from the effect of the fluorescent light during storage at 35°C. Similar results were obtained by Shipe et al. (1983) and Fanelli et al. (1985) who showed that when.Tinuvin 326 was impregnated into plastic containers, vitamin A in milk was protected. from photooxidation. This protection results because of the UV absorbent characteristic of Tinuvin 326. Guillet (1972) stated that UV absorbers can preferentially absorb light and harmlessly dissipate its energyu This energy lies within the ultra violet region of the electro-magnetic spectrum. Since Tinuvin 326 absorbs light energy between 190 - 390 nm (the ultra violet range of the spectrum), the removal of this energy results in the reduction of photooxidation in the oil exposed to the fluorescent light. 81 Table 17. Peroxide value (milliequivalents/kg oil) of the soybean oil in the multilayered container with 0% and 0.3% Tinuvin 326, stored at 35°C Storage time IPeroxide value of oil I Peroxide value of oil 35 64.6554 (days) Iin 0% Tin. containers I in 0.3% Tin. containers 0 0.7985 0.7985 1 2.7437 2.4940 2 3.9432 3.9652 4 6.9170 5.9054 7 9.0223 8.4222 14 16.7877 13.8682 21 29.7837 21.9952 25 38.7162 25.7162 28 40.1460 30.4720 32 61.4133 39.2034 44.1507 82 70.00 '1 s—e peroxide value of oil in 0.0% Tinuvin containers - 60 00- H peroxide value of oil in 0.32 linuvin containers 1 50.00- Il - 40.00-3 J A 30.00d ' ' .l 20.00- 1 10.00- J —-/ Peroxide value (milliequiv./kg oil) 0.00 e 0 151 21 28 35 Storage time (days) \1-1 Fig. 16 Peroxide value of soybean oil in the containers with 0% and 0.3% Tinuvin 326 stored at 35°C. ON 8 8 The loss of Tinuvin 326 from the container wall of the multilayered polypropylene based cups was measured as a function of time (42 days) and temperature (21 and 35°C). Loss of Tinuvin 326 from the container wall was negligible. The results also showed that both the HPLC and the UV spectrophotometric methods could be used to determine the concentration of Tinuvin 326 in the plastic container system. There was close agreement between the results from these two methods. There was also an acceptable level of precision between the results from each method. Migration of Tinuvin 326 from the containers to the packaged oil was greater at the higher temperature (35°C). The levels migrating to the oil were too low to noticeably affect the stability characteristics.of the UV absorber in the plastic containers. The highest level of migration occurred after 42 days and at 35°C. This was determined to be 4.773% of 9.009 x10'3g Tinuvin 326, and is equivalent to 4.30 x10"g Tinuvin 326. Tinuvin 326 was found to have the ability to reduce the rate of lipid oxidation in soybean oil. Since it absorbs UV radiation and does not itself disintegrate between 190 - 390 83 84 nm, Tinuvin 326 functions as a UV absorber. This study also showed that cool white fluorescent light has energy levels which can contribute to lipid oxidation in oil. The oil in the cups which contained dispersed Tinuvin 326 suffered less lipid oxidation than those in cups containing no Tinuvin 326. This difference was significant at both storage temperatures. "APPENDICES" 85 8229.09.15.1- Table 18. Data used for the standard curve for Tinuvin 326 at several concentration levels (using HPLC). Concentration Tinuvin 326 i Average area ppm (w/v) . Response units 1 4237 10 41844 60 238410 80 319017 100 402316 86 Appendix g. Table 19. Data used for the standard curve for Tinuvin 326 at several concentration levels (using UV spectrophotometry). concentration.of Tinuvin 326 ' Absorbance (optical density in ppm (w/v) units) "" """"" 1 """"""""" ' """""" ' 0'35"; """" " 5 0.263 10 0.516 87 Aoreodiz_11 From the standard curve by HPLC method, slope = 3.998 x 103 au/ppm converting au/ppm to g/au units: 1 ppm 1 x 10’6 g/ml 10 ul injected = 0.01 ml therefore slope (3.998 x 103 au/l x 10‘6 g/ml) + (0.01ml) = 3.998 x 1011 au/g = 2.501 x 10"12 g/au area response units (inject 1) = 136765 area response units (inject 2) = 138170 average area response units = 137467.5 sample weight = 3.0394 g 137467.5 au = 3.438 x 10'7 g Tinuvin/1001 in 200 ml solution = 6.876 x 10’3 g Tinuvin since 200 ml solution has 3.0394g plastic, (6.876 x10’3 g) x100 % Tinuvin 326 + 3.03949 0.226% Tinuvin 326 Appendix 4. Table 5. HPLC analysis of containers with Tinuvin 326 stored at 21 degrees C. TimeI ExtractIRep.'Sample I inj.I Area I % Average daysI No. I I Wt. I No. I responseITinuvin I % 0 1 1 3.0394 1 136765 0 l 1 3.0394 2 138170 0.2263 0 2 1 3.0394 1 71470 0 2 1 3.0394 2 69976 0.0582 0 3 1 3.0394 1 23459 0 3 1 3.0394 2 22518 0.0189 0.303 0 1 2 3.0159 1 122959 0 l 2 3.0159 2 122530 0.2036 0 2 2 3.0159 1 75152 0 2 2 3.0159 2 77449 0.0633 0 3 2 3.0159 1 29894 0 3 2 3.0159 2 29945 0.0248 0.292 Overall average % = 0.298 3 l 1 3.0079 1 162763 3 1 ‘ 1 3.0079 2 160375 0.2687 3 2 1 3.0079 1 23117 i 3 2 1 3.0079 2 22038 0.0375 3 3 1 3.0079 1 9126 3 3 1 3.0079 2 7561 0.0069 0.313 3 1 2 3.0392 1 125725 3 1 2 3.0392 2 131637 0.2118 3 2 2 3.0392 1 47425 3 2 2 3.0392 2 45560 0.0765 3 3 2 3.0392 1 29672 3 3 2 3.0392 2 29771 0.0245 0.313 Overall average % = 0.313 7 l 1 3.0115 1 173451 7 1 1 3.0115 2 158044 0.2753 7 2 1 3.0115 1 27810 7 2 1 3.0115 2 27825 0.0231 7 3 1 3.0115 1 14456 7 3 1 3.0115 2 12893 0.0114 0.310 7 1 2 3.0684 1 141819 7 l 2 3.0684 2 140642 0.2303 7 2 2 3.0684 1 34878 7 2 2 3.0684 2 35769 0.0288 7 3 2 3.0684 1 35129 7 3 2 3.0684 2 34996 0.0286 0.288 Overall average % = 0.299 89 "Table 5 (cont’d)." l4 1 l 3.07 1 129746 14 l l 3.07 2 128912 14 2 l 3.07 1 97623 14 2 l 3.07 2 98108 14 3 1 3.07 1 33303 14 3 l 3.07 2 33667 14 l 2 3.051 1 150929 14 l 2 3.051 2 149863 14 2 2 3.051 1 76121 14 2 2 3.051 2 78097 14 3 2 3.051 1 25109 14 3 2 3.051 2 25161 Overall average % 21 l 1 3.0225 1 102553 21 l 1 3.0225 2 104772 21 2 1 3.0225 1 101230 21 2 1 3.0225 2 102068 21 3 1 3.0225 1 50882 21 3 1 3.0225 2 50727 21 l 2 3.0667 1 108491 21 l 2 3.0667 2 100599 21 2 2 3.0667 1 99601 21 2 2 3.0667 2 102222 21 3 2 3.0667 1 47055 21 3 2 3.0667 2 45970 Overall average % 28 l 1 3.0342 1 137266 28 l 1 3.0342 2 137973 28 2 1 3.0342 1 74518 28 2 1 3.0342 2 76928 28 3 1 3.0342 1 31882 28 3 1 3.0342 2 29446 28 l 2 3.0124 1 128654 28 l 2 3.0124 2 126095 28 2 2 3.0124 1 77901 28 2 2 3.0124 2 73477 28 3 2 3.0124 1 31791 28 3 2 3.0124 2 31315 Overall average % 42 l 1 3.042 1 114032 42 1 1 3.042 2 110820 42 2 1 3.042 1 93766 42 2 1 3.042 2 95446 42 3 1 3.042 1 40824 42 3 1 3.042 2 40753 0.2107 0.0797 0.0273 0.2466 0.0632 0.0206 0.324 0.1716 0.0841 0.0420 0.1705 0.0823 0.0379 0.294 0.2269 0.0624 0.0253 0.2115 0.0628 0.0262 0.308 0.1849 0.0778 0.0335 0.318 0.330 0.298 0.291 0.315 0.301 0.296 90 "Table 5 (cont’d)." 42 l 2 3.0876 1 119432 42 1 2 3.0876 2 118119 0.1924 42 2 2 3.0876 1 95298 42 2 2 3.0876 2 94971 0.0771 42 3 2 3.0876 1 40074 42 3 2 3.0876 2 39376 0.0322 0.302 Overall average % = 0.299 Extract No. = extraction number Rep. = replicate Inj. No. = injection number Table 6. HPLC analysis of containers with Tinuvin 326 stored at 35 degrees C. ExtractIRep.'Sample No. Wt. 9 1 Area % responseITinuvin Average UUUULAUUUUUUU 000000000000 QQQQQQQQQQQQ wUNNl-‘HUUNNHH UUNNHHUUNNHH UUNNHHUUNNHH NPOAJNtOh3HiJF‘HFJP‘ IOAJNIOBJNFJF‘HIHPJH NMNNNNI—‘I—‘l—‘HI—‘H 3.0394 3.0394 3.0394 3.0394 3.0394 3.0394 3.0159 3.0159 3.0159 3.0159 3.0159 3.0159 ~14nahaml4nah1~14niw 136765 138170 71470 69976 23459 22518 122959 122530 75152 77449 29894 29945 Overall average % 3.0742 3.0742 3.0742 3.0742 3.0742 3.0742 3.0663 3.0663 3.0663 3.0663 3.0663 3.0663 NHNHNHNHNHNH 124106 126353 91190 95076 31928 31391 93694 95895 97125 97964 40431 41955 Overall average % 3.0624 3.0624 3.0624 3.0624 3.0624 3.0624 3.0768 3.0768 3.0768 3.0768 3.0768 3.0768 Nt‘k’Hth‘Nh‘h3HtvhJ 129159 130964 39988 40880 44711 45380 138254 129483 50579 50349 72223 74329 Overall average % 0.2263 0.0582 0.0189 0.2036 0.0633 0.0248 0.298 0.2038 0.0758 0.0258 0.1547 0.0796 0.0336 0.287 0.2125 0.0330 0.0368 0.2177 0.0410 0.0596 0.300 0.303 0.292 0.305 0.268 0.282 0.318 92 "Table 6 (cont’d)." 14 l 1 3.0476 1 143807 14 l 1 3.0476 2 145478 14 2 1 3.0476 1 90797 14 2 1 3.0476 2 91023 14 3 1 3.0476 1 31726 14 3 1 3.0476 2 31593 14 1 2 3.0687 1 125261 14 1 2 3.0687 2 123394 14 2 2 3.0687 1 142057 14 2 2 3.0687 2 102060 14 3 2 3.0687 1 36291 14 3 2 3.0687 2 36329 Overall average % 21 l 1 3.0472 1 126568 21 l 1 3.0472 2 123748 21 2 1 3.0472 1 76907 21 2 1 3.0472 2 77842 21 3 1 3.0472 1 42894 21 3 1 3.0472 2 41924 21 l 2 3.0441 1 127132 21 l 2 3.0441 2 124630 21 2 2 3.0441 1 108529 21 2 2 3.0441 2 93468 21 3 2 3.0441 1 35767 21 3 2 3.0441 2 35782 Overall average % 28 l 1 3.0687 1 119549 28 1 1 3.0687 2 127612 28 2 1 3.0687 1 84950 28 2 1 3.0687 2 82620 28 3 1 3.0687 1 41796 28 3 1 3.0687 2 39763 28 1 2 3.0213 1 148498 28 l 2 3.0213 2 148597 28 2 2 3.0213 1 69535 28 2 2 3.0213 2 71865 28 3 2 3.0213 1 19754 28 3 2 3.0213 2 20480 Overall average % 42 1 1 3.0369 1 122922 42 1 1 3.0369 2 122118 42 2 1 3.0369 1 92083 42 2 1 3.0369 2 93110 42 3 1 3.0369 1 36437 42 3 1 3.0369 2 33819 0.2374 0.0746 0.0260 0.2027 0.0995 0.0296 0.335 0.2055 0.0635 0.0348 0.2069 0.0830 0.0294 0.312 0.2015 0.0683 0.0332 0.2460 0.0585 0.0167 0.312 0.2018 0.0763 0.0289 0.338 0.332 0.304 0.319 0.303 0.321 0.307 93 "Table 6 (cont’d)." 42 l 2 3.017 1 125035 42 l 2 3.017 2 124191 0.2066 42 2 2 3.017 1 90538 42 2 2 3.017 2 90694 0.0751 42 3 2 3.017 1 32170 42 3 2 3.017 2 33600 0.0273 0.309 Overall average % - 0.308 Extract No. = extraction number Rep. 8 replicate Inj. No. 3 injection number 94 Appendix 5. absorbance = 0.942 optical density (OD) units sample weight = 3.0210 g dilution factor = 4 corrected absorbance 3.768 optical density (OD) units substitution in y = m* + c y = 5.191 x102 OD/ppm(*) + 2.347 x 104oo/ppm (4) ppm (3.768 00) + (5.191 x 10'2 OD/ppm) - (2.347 x 104 OD/ppm) 72.587 ppm /100 ml solution 72.587 x 10 j g [ml /100 ml solution 72.587 x 104 g Tinuvin 326 In 100 9 plastic, % Tinuvin 326 (sample wt. = 3.0210 g) (72.587 x 104 g) x 100 + 3.0210 g 0.240% Tinuvin 326 95 Appendix 6. Table 7. Stability studies.onicontainers with.Tinuvin 326 at 21 degrees C. using UV spectrophotometric method. Time ISample IExtractI Rep.'Corrected I Conc. Tin. I AVE. days I wt. I No. I IabsorbanceI in plastic I % 0 3.0210 1 1 3.768 0.2403 0 3.0394 2 1 0.925 0.0586 0 3.0159 3 1 0.313 0.0200 0.319 0 3.0210 1 2 3.288 0.2097 0 3.0394 2 2 1.008 0.0639 0 3.0159 3 2 0.389 0.0248 0.298 Overall average % = 0.309 3 3.0079 1 1 4.376 0.2803 3 3.0079 2 1 0.287 0.0184 3 3.0079 3 l 0.1 0.0064 0.305 3 3.0392 1 2 3.464 0.2196 3 3.0392 2 2 0.606 0.0384 3 3.0392 3 2 0.362 0.0229 0.281 Overall average % = 0.293 7 3.0115 1 1 2.934 0.1877 7 3.0115 2 1 1.108 0.0709 7 3.0115 3 1 0.736 0.0471 0.306 7 3.0638 1 2 3.706 0.2330 7 3.0638 2 2 0.926 0.0582 7 3.0638 3 2 0.442 0.0278 0.319 Overall average % = 0.312 14 3.07 l 1 3.352 0.2103 14 3.07 2 1 1.284 0.0806 14 3.07 3 1 0.436 0.0274 0.318 14 3.051 1 2 3.888 0.2455 14 3.051 2 2 0.998 0.0630 14 3.051 3 2 0.331 0.0209 0.329 Overall average % = 0.324 21 3.0225 1 1 2.754 0.1755 21 3.0225 2 1 1.384 0.0882 21 3.0225 3 l 0.67 0.0427 0.306 21 3.0667 1 2 2.77 0.1740 21 3.0667 2 2 1.288 0.0809 21 3.0667 3 2 0.618 0.0388 0.294 Overall average % = 0.300 "Table 7 (cont'd)." 28 3.0342 1 1 3.628 0.2303 28 3.0342 2 1 0.999 0.0634 28 3.0342 3 1 0.376 0.0239 0.318 28 3.0124 1 2 3.44 0.2200 28 3.0124 2 2 1.016 0.0650 28 3.0124 3 2 0.424 0.0271 0.312 Overall average % = 0.315 42 3.042 1 1 3.032 0.1920 42 3.042 2 1 1.268 0.0803 42 3.042 3 l 0.53 0.0336 0.306 42 3.0876 1 2 3.08 0.1922 42 3.0876 2 2 1.255 0.0783 42 3.0876 3 2 0.53 0.0331 0.304 Overall average % = 0.305 Extract No. extraction number Rep. = replicate Conc. Tin. = concentration Tinuvin 326 Ave. = average 97 Table 8. Stability studies on containers with Tinuvin 326 at 35 degrees C. using UV spectrophotometric method. Time ISample IExtractI Rep.'Corrected I Conc. Tin. I AVE. I I days I wt. I No. I IabsorbanceI in plastic % 0 3.0210 1 1 3.768 0.2403 0 3.0394 2 1 0.925 0.0586 0 3.0159 3 1 0.313 0.0200 0.319 0 3.0210 1 2 3.288 0.2097 0 3.0394 2 2 1.008 0.0639 0 3.0159 3 2 0.389 0.0248 0.298 Overall average % = 0.309 3 3.0117 1 l 2.69 0.1721 3 3.0117 2 1 1.194 0.0764 3 3.0117 3 1 0.585 0.0374 0.286 3 3.0742 1 2 3.416 0.2141 3 3.0742 2 2 1.208 0.0757 3 3.0742 3 2 0.397 0.0249 0.315 Overall average % = 0.300 7 3.0861 1 1 2.206 0.1377 7 3.0861 2 1 0.918 0.0573 7 3.0861 3 1 1.31 0.0818 0.277 7 3.0624 1 2 3.37 0.2120 7 3.0624 2 2 1.034 0.0650 7 3.0624 3 2 0.566 0.0356 0.313 Overall average % = 0.295 14 3.0476 1 1 2.582 0.1632 14 3.0476 2 l 1.51 0.0954 14 3.0476 3 l 0.69 0.0436 0.302 14 3.0533 1 2 3.228 0.2037 14 3.0533 2 2 1.344 0.0848 14 3.0533 3 2 0.462 0.0291 0.318 Overall average % = 0.310 21 3.0472 1 l 3.05 0.1928 21 3.0472 2 1 1.027 0.0649 21 3.0472 3 1 0.563 0.0356 0.293 21 3.0441 1 2 3.398 0.2150 21 3.0441 2 2 1.256 0.0795 21 3.0441 3 2 0.484 0.0306 0.325 Overall average % = 0.309 "Table 8 (cont'd)." 28 3.0687 1 1 3.272 0.2054 28 3.0687 2 1 1.117 0.0701 28 3.0687 3 1 0.531 0.0333 0.309 28 3.0213 1 2 3.978 0.2536 28 3.0213 2 2 0.935 0.0596 28 3.0213 3 2 0.287 0.0183 0.332 Overall average % = 0.320 42 3.0369 1 1 3.234 0.2051 42 3.0369 2 1 1.262 0.0801 42 3.0369 3 1 0.478 0.0303 0.316 42 3.017 1 2 3.356 0.2143 42 3.017 2 2 1.2 0.0766 42 3.017 3 2 0.456 0.0291 0.320 Overall average % = 0.318 Extract No. extraction number Rep. = replicate Conc. Tin. = concentration Tinuvin 326 Ave. = average 99 A and 7. Sample calculation for 7 From the standard slope = 1 ppm = 10 pl injected = 0.01 ml therefore slope = (3.998 days storage at 35°C. curve by HPLC method, 3.998 x103 au/ppm 1 x 10‘ g/ml x 103 au/l x 10*5 g/ml) + 0.0 ml = 3.998 x 10" au/g = 2.501 x 10"2 g/au 5 extractions of 15 ml each = Area response units (2.501 x10“ g/au) x 642au = in 5 ml concentrated soln. since extraction efficiency actual weight in the oil (in 3 ml oil) = in 120 m1 oil or a container= % Tinuvin in oil/container 75 ml solution (cone. to 5 ml) 642 an 1.6056 x 10'9 g/lo pl injected (1.6056 x 10* g) 5ml + 0.01ml 8.028 x 10'7 g Tinuvin 326 81.00% 8.028 x 10'7 g x 100 + 81 9.9111 x 10'7 g/5 ml conc. 9.9111 x 10'1 g x 120ml + 3 ml 3.964 x 10* g Tinuvin 326 3.964 x 10* g x 100 + 120 3.3037 x 104 8 (w/v) % Tinuvin transferred from container wall to the oil 3.964 x 10¢ g x 100 + 9.009 x 10'3 g 0.440% Tinuvin 326 100 Appendgx 8. Title: Peroxide value of oil stored at 21 degrees C. Function: FACTOR Experiment Model Number 1: Two Factor Completely Randomized Design Factorial ANOVA for the factors: Replication (Var 1: Replicate) with values from 1 to 2 Factor A (Var 2: Tinuvin 326) with values from 1 to 2 Factor B (Var 3: Time) with values from 1 to 10 Variable 4: Peroxide value (milliequiv./kg oil) Grand Mean = 10.918, Grand Sum.= 436.717, Total Count= 40 A N A L Y S I S O F V A R I A N C E T A B L E Degrees of Sum of Mean F Source Freedom Squares _ Square Value Prob Factor A 1 131.564 131.564 374.6137 0.0000 Factor B 9 1952.747 216.972 617.8050 0.0000 AB 9 106.172 11.797 33.5905 0.0000 Error 20 7.024 0.351 Total 39 2197.507 Coefficient of Variation: 5.43% From the ANOVA table: 1. the observed F value for Factor A (Tinuvin 326) = 374.61 2. the observed F value for Factor A (Tinuvin 326) = 617.81 For the effect of Tinuvin, the F-distribution tables at d.f 1 (numerator) and 9 (denominator) = 10.6 at 1% level of significance For the effect of Time, the F-distribution tables at d.f 1 (numerator) and 9 (denominator) = 10.6 at 1% level of significance Since the observed F value for the effects of time and Tinuvin 326 exceed even the 1% level of significance, the oil in the containers with 0.3% Tinuvin 326 was better protected from the effects of the fluorescent light than the oil in the containers 0% Tinuvin 326 stored at 21%L 101 Title: Peroxide value of oil stored at 35%: Function: FACTOR Experiment Model Number 1: Two Factor Completely Randomized Design Factorial ANOVA for the factors: Replication (Var 1: replicate) with values from 1 to 2 Factor A (Var 2: Tinuvin 326) with values from 1 to 2 Factor B (Var 3: Time) with values from 1 to 10 Variable 4: Peroxide value (milliequiv./Kg oil) Grand Mean = 23.486, Grand Sum = 939.445, Total Count= 40 A N A L Y S I S O F V A R I A N C E T A B L E Degrees of Sum of Mean F Source Freedom Squares Square Value Prob Factor A 1 598.110 598.110 659.7521 0.0000 Factor B 9 13139.301 1459.922 1610.3851 0.0000 AB 9 633.541 70.393 77.6484 0.0000 Error 20 18.131 0.907 Total 39 14389.084 Coefficient of Variation: 4.05% From the ANOVA table: 1. the observed F value for Factor A (Tinuvin 326) = 659.752 2. the observed F value for Factor A (Tinuvin 326) = 1610.385 For the effect of Tinuvin, the F-distribution tables at d.f 1 (numerator) and 9 (denominator) = 10.6 at 1% level of significance For the effect of Time, the F-distribution tables at d.f 1 (numerator) and 9 (denominator) = 10.6 at 1% level of significance Since the observed F value for both the effects of time and Tinuvin 326 exceed even the 1% level of significance, the oil in the containers with 0.3% Tinuvin 326 was better protected from the effects of the fluorescent light than the oil in the containers 0% Tinuvin 326 at 35%L W REFERENCES Association of Official Analytical Chemists. 1984. 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