Evaluation of Modified Amaranth Starch as Shell Material for Encapsulation of Probiotics Reyna Nallely Falfán Cortés,1 Marcela Gaytán Martínez,1 Iñigo Verdalet Guzmán,2 Silvia Lorena Amaya Llano,3 Carlos Raimundo Ferreira Grosso,4 and Fernando Martínez Bustos1,5 ABSTRACT
Cereal Chem. 91(3):300–308
The objective of this study was to develop with thermoplastic extrusion amaranth starch derivatives and to characterize and evaluate their functionality as encapsulating agents of Bifidobacterium breve ATCC 15700 and Lactobacillus casei ATCC 334 during spray drying. The survival of both probiotics during storage at different water activities and at two storage temperatures, their viability in a food model system, and their tolerance to a simulated gastrointestinal tract were determined. Native amaranth starch was chemically modified to obtain phosphorylated, acetylated, and succinylated starch. Starch derivatives were reduced in viscosity, and the solu-
bility in water was increased. In general, the modified amaranth starches and control corn starch did not provide good protection to both probiotics during storage at 25°C. However, there was excellent viability during storage at 4°C for both probiotics. Microcapsules showed a uniform coverage of the cells. Storage for 35 days at 25°C of blends of oat with succinylated amaranth microcapsules with probiotics had a lower reduction. Also, this succinylated amaranth starch containing probiotics showed a higher resistance to simulated gastrointestinal conditions. The results with food model systems supported the applicability of the modified starches.
Probiotics are live microorganisms that confer a beneficial effect on the host when administered in proper amounts (Brown and Valiere 2004). Probiotics are typically Lactobacillus and Bifidobacterium genus members commonly associated with the human gastrointestinal tract. Lactobacillus and Bifidobacterium species used as a probiotic dietary adjunct can have several health and nutritional benefits (Ishibashi and Shimamura 1993). It is of great importance that, when selecting bacteria for their physiological effects, they should stay viable during the whole shelf life of the food product and that there is no decrease in their resistance to the acidic environment of the stomach and to bile salts in the small intestine (Kailasapathy and Rybka 1997; Saarela et al 2000). For probiotics to be therapeutically effective, it has been suggested that products should contain at least 106 colony-forming units (CFU)/g of bacteria at the end of the expiry period (Talwalkar et al 2004). Studies indicate that probiotic bacteria do not survive in high enough quantities when incorporated into food products, mostly in fermented milk-based products (Shah 2000; LourensHattingh and Viljoen 2001). The loss of viability may occur during storage prior to consumption, during processing from oxygen stress, during freezing or drying, or because of the severe gastrointestinal action (Akhiar 2010). The goal of encapsulation is to create a microenvironment in which the bacteria will survive during processing and storage and be released at appropriate sites in the digestive tract (Weinbreck et al 2010). For Bifidobacterium and Lactobacillus species, studies have evaluated the viability in different food matrices, storage conditions (varying temperature, water activity, and packaging material), wall materials used in the encapsulation process, and methods of encapsulation and their tolerance to simulated gastrointestinal conditions (low pH, enzyme activity, and high concentration of bile salts) (Hsiao et al 2004; Crittenden et al 2006; Anal and Singh 2007; Oliveira et al 2007). The main wall materials studied for the encapsulation of these organisms are systems consisting mainly of carrageenan, alginate, cellulose acetate phthalate, gelatin, and chitosan. In almost all cases, gel entrapment was used
with the materials mentioned. However, although promising on a laboratory scale, the developed technologies for producing gel beads still present serious difficulties for large-scale production of food-grade microencapsulated microorganisms. On the other hand, microencapsulation by spray drying is a well-established process that can produce large amounts of material (Anal and Singh 2007). Starch derivatives (succinylated, acetylated, and phosphorylated) have proved effective encapsulating agents when used in the spray drying of flavors and pigments. These materials have demonstrated a high degree of solubility, limited viscosity in solution, good emulsifying properties, and good drying properties (Cai and Corke 2000; Murúa-Pagola et al 2009). Diversity of technological qualities of cereal starches is as widespread as the variety of species, with special emphasis on maize, rice, wheat, and modifications of these basic species. However, this range is expanding, in particular with newly bred or technologically modified species; thus, even pseudocereals such as species of amaranth have become increasingly interesting (Praznik et al 1999). However, no previous work on amaranth modified starches as carrier agents for spray drying of probiotics has been reported. In the search for unconventional new biomaterials, the aim of this work was to study the microencapsulation of Bifidobacterium breve ATCC 15700 and Lactobacillus casei ATCC 334 by spray drying with modified amaranth starches compared with one commercial modified starch.
1 CINVESTAV
unidad Querétaro, Apdo. Postal 1-798, Querétaro, Qro., México. Ciencias Basicas, UV, Col Ind Animas, Xalapa, Veracruz, México. de Química, PROPAC, Centro Universitario s/n. Apdo. Postal 76010, Querétaro, Qro., México. 4 Faculty of Food Engineering, UNICAMP, Campinas, Sao Paulo, Brazil. 5 Corresponding author. Phone +52 (442) 2119905. Fax: +52 (442) 2119938. E-mail:
[email protected] 2 Inst.
3 Facultad
http://dx.doi.org/10.1094 / CCHEM-06-13-0112-R © 2014 AACC International, Inc.
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MATERIALS AND METHODS Materials. Seeds of Amaranthus cultivar Nutrisol were donated by the National Research Institute for Agriculture, Forestry and Livestock, Mexico. N-LOK starch, a commercial waxy corn starch modified with octenyl succinic anhydride (OSA) and enzymatically hydrolyzed after or before chemical modification, was purchased from National Starch and Chemical Co. (Mexico City, Mexico) and was used as the control. In this study, B. breve ATCC 15700 and L. casei ATCC 334, available from the American Type Culture Collection (ATCC, Manassas, VA, U.S.A.), were used as the test organisms. Extraction of Starch. Extraction of starch was performed according to the alkaline wet-milling method described by Radosavljevic et al (1998) with some modifications. Amaranth seeds (1,000 g) were steeped in 2 L of 0.1N NaOH solution for 24 h. After steeping, the steeping solution was decanted, and the seeds were washed with distilled water. The sample was then milled in a stone mill (Fumasa, Querétaro, Mexico). A cold solution of
0.1N sodium bisulfate (NaHSO3) was added during grinding. The ground slurry was screened through U.S. standard sieves (841, 595, 420, 250, 177, 149, 74, and 62.5 μm) for washing the fiber fraction. The starch was isolated by using a centrifuge at 6,000 × g for 20 min. The supernatant was discarded, and the top yellowish layer of protein was removed with a laboratory spatula. The white starch layer was resuspended in distilled water and centrifuged as described earlier. The starch was dried in a convection oven at 40°C for 48 h. Starch had 3.2% protein and 12.4% amylose contents. Starch Hydrolysis. Hydrolysis of the native amaranth starch was performed according to the methodology of Murúa-Pagola et al (2009). A slurry of 40 g of starch (db)/100 mL of distilled water was prepared, and an aqueous HCl solution at a final concentration of 3.4 g of pure HCl/100 g of starch (db) was added. The mixture was placed in a water bath at 50°C with constant stirring for 6 h. The slurry was neutralized to pH 5.0 with 1N NaOH and washed by centrifugation for 10 min at 2,012 × g in a centrifuge (Z2OO A, Hermle, Wehingen, Germany) and dried in a convection oven at 45°C for 48 h. Phosphorylation of Starch by Extrusion. Starch phosphorylation was performed according to the methodology of MurúaPagola et al (2009). A solution of sodium tripolyphosphate (4 g/100 g of starch) was placed in a glass flask and was sprinkled on the dried powder of hydrolyzed starch (the pH was adjusted to 5.0–5.2); the moisture content of starch before extrusion was 25%. Samples were stored in polyethylene bags at 4°C for subsequent extrusion processing. Succinylation of Starch. The modified starch was prepared following the method proposed by Murúa-Pagola et al (2009); a slurry of hydrolyzed starch (45 g [db]/100 mL) was prepared with vigorous stirring. The pH was maintained between 8.5 and 9.0 with an aqueous solution 1.25N NaOH. About 2.0 mL of n-OSA/100 g of starch (db) was added. Then the pH was brought to 5.0 ± 0.2 with 0.5N HCl, and the slurry was centrifuged for 10 min at 2,012 × g. The solid product was washed with water and dried for 24 h in a convection oven at 45°C. The dried powder was milled and sieved in a mesh of 149 μm opening size. After succinylation, the starches were extruded in a single-screw extruder. Acetylation of Starch. Starch acetylation was performed as described by Phillips et al (1999). A mixture of 100 g of starch (db) in 225 mL of distilled water was prepared and stirred for 60 min at 25°C. Acetic anhydride (2.5 g) was added to the mixture, and the slurry was stirred while maintaining a pH range of 8.0–8.4. The slurry was then adjusted to pH 5.0 with 0.5N HCl and centrifuged for 10 min at 2,012 × g. It was washed with distilled water and dried in a convection oven at 45°C for 24 h. After acetylation, the starches were extruded in a single-screw extruder. Extrusion Process. The extrusion process was conducted under the same conditions for all three modifications with a laboratory single-screw extruder designed and manufactured by CINVESTAVIPN, Mexico, with an internal barrel diameter of 20 mm (length/diameter = 20). Barrel temperatures were 50, 130, and 170°C at the feeding, transition, and high-pressure extrusion zones, respectively. The compression ratio of the screw was 2:1, and a 4.0 mm diameter die nozzle was used. Extruded samples were dried in a convection oven (45°C) for 2 h, milled with a hammer mill (model 200, Pulvex, Mexico City, Mexico), and sieved in a mesh of 149 μm opening size. Degree of Substitution (DS)—Starch Phosphates. The DS in starch phosphates was determined following the method proposed by Smith and Caruso (1964) (equation 1). DS was calculated as follows: DS =
162 × %P 3,100 − (124 × %P )
(1)
where %P is the percentage of phosphorus (db) of the phosphorylated starch.
DS—Starch Octenyl Succinates. Octenyl succinylation level (%Succinyl) and DS (equations 2 and 3, respectively) of modified starch were determined with the titrimetric method (Whistler and Paschall 1967). Briefly, 25 mL of an aqueous 0.5N NaOH solution was added to the suspension of 5 g of starch in 50 mL of distilled water and then shaken for 24 h. The excess of alkali was titrated with 0.5N HCL with a Brand burette (error limit 0.05 mL, Wertheim, Germany) in all determinations. A blank with the original unmodified starch was also used. %Succinyl =
(blank − sample )× 0.1 × N HCl × 100 sample weight
(2)
DS was calculated as follows: DS =
(162 × %Succinyl ) 21,00 0 − (209 × %Succinyl )
(3)
where 162 is the molecular weight of the glucose unit, 21,000 = 100 × the molecular weight of octenyl succinyl group, and 209 is the molecular weight of the octenyl succinyl group. DS—Starch Acetates. The percent of acetylation (%Acetyl) and DS (equations 4 and 5, respectively) were determined titrimetrically (Wurzburg 1978). Acetylated starch (1 g) was placed in a 250 mL flask, and 50 mL of 75% ethanol in distilled water was added. The loosely stoppered flask was agitated, warmed to 50°C for 30 min, and cooled, and then 40 mL of 0.5N KOH was added. The excess of alkali was titrated with 0.5N HCL with a Brand burette (error limit 0.05 mL) in all determinations. A blank with the original unmodified starch was also used. %Acetyl =
(blank − sample )× N HCl × 0.043 × 100 sample weight
(4)
Blank and sample were titration volumes in milliliters, and sample weight was in grams. DS was calculated as follows: DS =
(162 × %Acetyl ) 4,300 − (42 × %Acetyl )
(5)
Pasting Properties. The pasting properties of the native and modified starches were measured with a Rapid Visco Analyzer (RVA, model 3C, Newport Scientific, Warriewood, Australia). The analysis was based on the standard program for RVA analysis in AACC International Approved Method 61-02.01. The heating and cooling cycles were programmed in the following manner: the samples were held at 50°C for 1 min, heated to 92°C at a heating rate of 5.6°C/min, held at 92°C for 5 min, cooled to 50°C at a cooling rate of 5.6°C/min, and held at a final temperature of 50°C for 2 min. Total time of analysis was 23 min. Water Solubility Index (WSI) and Water Absorption Index (WAI). WSI and WAI were determined as described by Anderson et al (1969). Starch (2.5 g) was placed in a container (50 mL), and 30 mL of distilled water was added. The tubes were maintained at 30°C with constant agitation for 30 min. Samples were centrifuged at 2,012 × g for 10 min. The gel obtained at the bottom of the tubes was weighed for WAI (g of gel/g of dry sample), and the dried supernatant weight was expressed as WSI (%) (g of dry solids/g of dry sample), with an analytical balance (error limit, 0.0005 g; Explorer, OHAUS, Parsippany, NJ, U.S.A.) as the measuring instrument in all trials. Preparation of Microencapsulated B. breve ATCC 15700 and L. casei ATCC 334. Cultures of B. breve ATCC 15700 and L. casei ATCC 334 were obtained after activation by two successive transfers in lactobacilli MRS broth under anaerobic conditions in anaerobic jars (GasPak EZ anaerobe container system sachets, Becton, Dickinson and Co. [BD], Franklin Lakes, NJ, U.S.A.) at 37°C for 48 h. Cultures in the late-log phase were harVol. 91, No. 3, 2014
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vested by centrifugation at 6,000 × g for 15 min (centrifuge Z200A, Hermle) and washed twice in a sterile solution with 0.9% peptone (BD Bioxon, Mexico City, Mexico). The pellet was resuspended in 10 mL of a sterile solution of 12% reconstituted skimmed milk. The cell suspension (with ≈109 CFU/g as the final bacterial concentration) was mixed with a carrier solution of modified starch (acetylated, succinylated, or phosphorylated). N-LOK commercial modified starch was used as the control; unmodified starch was not tested because it presents too high a viscosity to be used during spray drying. All the carrier solutions were prepared (20 g [db]/100 mL of sterile distilled water) and homogenized in an Ultra Turrax T-25-SI homogenizer (IKA Works, Wilmington, NC, U.S.A.). Microencapsulation was accomplished by spray drying (SD-Basic spray dryer, LabPlant, Huddersfield, U.K.). Drying conditions were as follows: inlet air temperature, 100– 110°C; outlet air temperature, 70–80°C; nozzle diameter, 0.5 mm; and liquid flow rate, 3 mL/min. Dried powder samples were collected from the base of the cyclone. Bacterial Enumeration. The microcapsule powders containing B. breve ATCC 15700 and the microcapsule powders with L. casei ATCC 334 were dispersed (1 g) in 9 mL of a sterile solution of 2% sodium citrate (Golden Bell Reactivos, Mexico, D.F.) with a homogenizer; the dispersed samples were subsequently subjected to serial dilutions with peptone (0.1%, w/v). Each dilution was placed on MRS agar. The colonies were grown under anaerobic conditions at 37°C for 48 h. Enumeration of bacteria was performed in triplicate with the traditional microbial plating method. Viability of Microencapsulated B. breve ATCC 15700 and L. casei ATCC 334 at 25 and 4°C During Storage. The viability during storage was evaluated according to the method proposed by Rascón et al (2010) with some modifications. The microencapsulated cells were evaluated at three water activities (aw) for each temperature: 0.329 (MgCl2), 0.536 (Mg(NO3)2), and 0.765 (NaCl), equilibrated at 25°C; and 0.355 (MgCl2), 0.587 (Mg(NO3)2), and 0.807 (NaCl), equilibrated at 4°C (Greenspan 1977; Labuza et al 1985). The powder containing microencapsulated cells (1 g dry weight) was placed in aluminum containers and stored in desiccators containing the saturated solutions previously mentioned. Viability of cells was determined at 0, 3, 7, 14, 21, 28, and 35 days of storage at both temperatures. Scanning Electron Microscopy (SEM). The morphologies of spray-dried microcapsules were observed with a SEM (environmental SEM with a gaseous secondary electron detector, accelera-
tion voltage of 20 kV, EDAX, Tilburg, The Netherlands) after spray-drying and during storage period. The encapsulated samples were fixed to stubs with double-faced adhesive metallic tape. Tolerance to Simulated Gastrointestinal Conditions in a Food Model System During Storage. The food model system was prepared with 1 g of capsules (B. breve or L. casei), which were mixed with 9 g of commercial precooked oats (db) and homogenized manually. The survival in vitro was evaluated according to the method proposed by Blazenka et al (2000). The encapsulated cells were inoculated in sequence, first in simulated artificial saliva, which then changed to simulated gastric juice and then to simulated intestinal juice. The mixture of encapsulated probiotics and oats (3 g) was resuspended in 3 mL of simulated artificial saliva, which was prepared by diluting of 6.2 g/L of sodium chloride (Baker 3624-01, Mexico D.F., Mexico), 2.2 g/L of potassium chloride (Baker 3040-01), 0.22 g/L of calcium chloride (Baker 1311-01), and 1.2 g/L of sodium bicarbonate (Karal 5010). It was then added to 24 mL of simulated gastric juice, which was prepared with 3 g/L of pepsin (Baker 2844-01), pH 2 (pH was adjusted with 0.1M HCL). The samples were incubated for 2 h at 37°C with constant agitation. Afterward, samples were exposed to the simulated intestinal juice, which was prepared with 1 g/L of pancreatin (P1750, Sigma, St. Louis, MO, U.S.A.) and 1.5 g/L of bile salts (48305, Fluka, Buchs, Switzerland), pH 8 (adjusted with 0.1M NaOH), and incubated for 4 h at 37°C with constant stirring. Cell numbers were determined by plate count. Viability of Encapsulated Microorganisms in a Food Model System. Two blends were prepared with commercial precooked oatmeal as the matrix system, one with microcapsules containing B. breve and the second with microcapsules containing L. casei. The mixture, prepared as described earlier, was homogenized and placed in sealed polyethylene bags, which were stored at 25°C. Viability of bacteria in each blend was evaluated at 0, 3, 7, 14, 21, 28, and 35 days. Statistical Analysis. All experiments were performed in triplicate. Differences between means of treatments were determined with analysis of variance and Tukey’s paired test (P < 0.05) with JMP version 8 software (SAS Institute, Cary, NC). RESULTS AND DISCUSSION DS—Starch Phosphates. In this analysis, the starch phosphorylation was performed by using the extrusion process at pH 5; the phosphorus content was 0.38 ± 0.05%, and the DS was 0.02 ± 0.00. Similar results were found by O’Brien et al (2009). These researchers reported phosphorus content of 0.27–0.43 at pH 9.0 and 0.27–0.33 at pH 11.0 for different types of extruded starch. Landerito and Wang (2005) produced extruded starch phosphates with a combination of sodium tripolyphosphate and sodium trimetaphosphate with waxy maize starch, which resulted in a phosphorous content of 1.63%. The differences reported probably result from the type of starch and type and concentration of reagent, as well as from effects of pH on DS and methods of analysis or DS calculations.
TABLE I Water Absorption Index (WAI) and Water Solubility Index (WSI) of Modified Amaranth Starches After a Chemical Modification and Extrusion Processz Sample
Fig. 1. Viscosity profiles of amaranth starches. A = native starch; B = phosphorylated starch; C = succinylated starch; D = acetylated starch; E = control starch (N-LOK); and F = temperature ramp (°C). 302
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Acetylated starch Succinylated starch Phosphorylated starch N-LOK starch (control) z
WAI (g of gel/ g of dry sample)
WSI (%)
3.6 ± 0.2a 3.5 ± 0.1a 3.1 ± 0.0b 4.0 ± 0.0c
31.8 ± 0.1b 19.8 ± 0.0a 32.2 ± 0.1c 85.3 ± 0.0d
Means ± standard deviations of three replicates. Means with different letters in the same column are significantly different (P < 0.05).
DS—Starch Octenyl Succinates. Succinylation confers a hydrophobic character to starch and weakens the internal bonding that holds the granules together (Bhosale and Singhal 2006). The OSA substitution and DS of the modified amaranth starch were 2.90 ± 0.02% and 0.02 ± 0.00, respectively. Bhosale and Singhal (2006) reported a linear increase of DS values in OSA-modified amaranth and waxy corn starches resulting from increase in the concentration of OSA; they found DS of 0.02 for amaranth starch and 0.02 for waxy corn starch using a 3% OSA reaction mixture. DS—Starch Acetates. The acetyl percentage and DS of acetylated starch were 2.32 ± 0.00% and 0.06 ± 0.00, respectively. Murúa-Pagola et al (2009) reported similar results with a DS of 0.03 for waxy maize starch treated with acetic anhydride. These authors concluded that the hydrophobic acetyl group introduced to starch acetates improved the emulsifying capabilities of the starch derivatives, making them good alternatives for the encapsulation of flavors. Singh-Sodhi and Singh (2005) reported a similar percentage of acetylation (2.26–3.68%) and DS (0.09–0.14) for different varieties of rice starch. These authors mentioned that the differences in acetyl content reported in various studies may be
because of differences in reaction conditions and sources of starch used. Also, Vasanthan et al (1995) reported acetyl group content between 1.01 and 2.80% when using different starch sources (potato, waxy corn, corn, wheat, field pea, and lentil). Pasting Properties. Starch derivatives did not develop viscosity (Fig. 1) in comparison with native amaranth starch, because before the chemical modification they were hydrolyzed with HCl and also extruded. Chang and Lii (1992) reported a very low viscosity of phosphorylated starch after an extrusion process, which can be attributed to disintegration of starch structure resulting from the high shearing, pressure, and temperature conditions during extrusion. Their observations correspond with the results found in this work. Murúa-Pagola et al (2009) also reported that starch phosphates modified by an extrusion process that were hydrolyzed with acid before extrusion did not develop viscosity, and they added that the viscosity for N-LOK starch was lower than 50 cP, with viscosity similar to the modified amaranth starches obtained in this work. Molecular weight (MW) of wall materials used to produce microparticles by spray drying can be important. Recently, Drusch et al (2007) evaluated the oxidative
Fig. 2. Survival of microencapsulated cells in modified amaranth starches during storage for 35 days at 25°C at different water activity (aw) levels. B. breve ATCC 15700: A, aw = 0.765; B, aw = 0.536; and C, aw = 0.329. L. casei ATCC 334: D, aw = 0.765; E, aw = 0.536; and F, aw = 0.329. Error bars indicate standard deviations. Vol. 91, No. 3, 2014
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stability of microparticles produced by spray drying, comparing two OSA-modified starches with different MW. These authors observed that the OSA-modified starch with low MW allowed a denser packaging of the molecules during the drying process and thus provided better protection to the core material. A similar effect, lowering MW of the modified starches, probably occurs during the extrusion step. WAI and WSI. The severe extrusion conditions caused an extensive dextrinization of the starch, resulting in an increased formation of water-soluble products (Harper 1992). WAI and WSI indicate the change in the structure of the starch granule before and after processing. WAI of native amaranth starch was 2.1 g of gel/g of dry sample, and WSI was 6.0%; however, after chemical and mechanical modification, these values were increased. The severity of the thermal treatment in the extruder will progressively increase WSI values, indicating that more starch polymers have been degraded into smaller molecules. Table I shows the WAI and WSI of modified starches. WAI did not show significant differences between succinylated and acetylated starches, but phosphorylated starch was significantly lower than the other
two modified starches. The introduction of hydrophobic groups in OSA-modified starches at low DS levels (0.01–0.1) confers some hydrophobic properties to starch, decreasing the water dispersibility, which could account for the low WSI with respect to hydrophilic modifications (acetylation and phosphorylation). According to various authors, the ideal carrier used to produce microparticles with a spray dryer should have low viscosity at high solid levels and also present high solubility (Reineccius 1989; Murúa-Pagola et al 2009), as showed by the modified starches used in the present work. Viability of Microencapsulated B. breve ATCC 15700 and L. casei ATCC 334 at 25 and 4°C During Storage. The survival of B. breve ATCC 15700 and L. casei ATCC 334 during storage at 25°C was low for all wall materials evaluated. Figure 2 shows the survival of both encapsulated probiotics for 35 days of storage at water activities of 0.765 (Fig. 2A and D), 0.536 (Fig. 2B and E), and 0.329 (Fig. 2C and F) at 25°C. In general, modified amaranth starches and control corn starch did not offer complete protection to B. breve ATCC 15700 and L. casei ATCC 334 until the end of storage (35 days); the surviving population declined rapidly dur-
Fig. 3. Survival of microencapsulated cells in modified amaranth starches during storage for 35 days at 4°C at different water activity (aw) levels. B. breve ATCC 15700: A, aw = 0.807; B, aw = 0.587; and C, aw = 0.355. L. casei ATCC 334: D, aw = 0.807; E, aw = 0.587; and F, aw = 0.355. Error bars indicate standard deviations. 304
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ing storage. Figure 2A, B, and C shows the survival of B. breve ATCC 15700. The control corn starch presented survival of the cells of 1.1 log CFU/g until 21 days when they were stored at aw of 0.765 (Fig. 2A); however, in modified amaranth starches there was no detected viability. When the encapsulated cells were stored at aw of 0.536, the survival in all evaluated wall materials increased in comparison to the cells stored at aw of 0.765 (Fig. 2B). All the evaluated wall materials maintained the best survival of microencapsulated B. breve ATCC 15700 when they were stored at aw of 0.329 at room temperature for 35 days in comparison with other activities evaluated (Fig. 2C). L. casei ATCC 334 (Fig. 2D, E, and F) showed increased survival in the same storage conditions related to B. breve ATCC 15700 at room temperature; succinylated amaranth starch and control corn starch showed viability greater than 4 log CFU/g until the end of storage for all the evaluated water activities. It was observed in both encapsulated organisms that the increase of water activity decreased survival of microencapsulated cells in modified amaranth starches and control corn starch. This result was similar to those reported by O’Riordan et al (2001). These authors evaluated microencapsulation of Bifidobacterium PL1 by using succinylated waxy maize starch as a coating material and found that those starches did not offer any protection to the Bifidobacterium strain in a dry malted beverage powder over 20 days of storage at room temperature (19–24°C). Ying et al (2010) reported that storage under nonrefrigerated conditions, high relative humidity
(70%), and high oxygen atmosphere content decreased the viability of encapsulated probiotic over the storage time. Figure 3 shows the survival of both encapsulated probiotics at 4°C during 35 days of storage at water activities of 0.807 (Fig. 3A and D), 0.587 (Fig. 3B and E), and 0.355 (Fig. 3C and F). The survival at 4°C was different in comparison to 25°C; the survival at 4°C was maintained for all the studied water activities in both bacteria (B. breve and L. casei) (Figs. 2 and 3). There is a substantial interaction between water activity and temperature with respect to their impact on the survival of quiescent probiotics. As the storage temperature is increased, the detrimental impact of moisture is magnified. Although the precise mechanisms of cell death remain unclear, osmotic stresses appear to play a role. Despite clear evidence that very low water activities improve probiotic survival, there may be technological limitations to reducing water activity to very low levels. Maintaining probiotic viability in moderate water activity foods (0.4–0.7) is a major challenge, and solutions such as microencapsulation or incorporation of probiotics resistant to processing or storage can provide improved survival. The results obtained showed that the survival of B. breve ATCC 15700 cells stored at 4°C and aw of 0.807 ranged from 6.88 ± 0.02 to 8.07 ± 0.06 log CFU/g after 35 days of storage (Fig. 3A). At aw of 0.587, succinylated and acetylated starches showed 7.87 ± 0.03 and 7.73 ± 0.01 log CFU/g, respectively; these values were significantly different (P < 0.05) between them and with the control
Fig. 4. Tolerance to simulated gastrointestinal conditions of a food system based on commercial precooked oatmeal containing microencapsulated B. breve ATCC 15700 (A) and L. casei ATCC 334 (B). Error bars indicate standard deviations.
Fig. 5. Viability of encapsulated microorganisms in a food system based on commercial precooked oatmeal B. breve ATCC 15700 (A) and L. casei ATCC 334 (B) during storage at 25°C. Error bars indicate standard deviations. Vol. 91, No. 3, 2014
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corn starch. Phosphorylated starch showed the lowest survival (7.08 ± 0.07 log CFU/g) and was significantly different (P < 0.05) compared with the control corn starch and the other wall materials (Fig. 3B). At aw of 0.355, control corn starch had 8.33 ± 0.01 log CFU/g and showed significant differences (P < 0.05) relative to modified amaranth starches; however, succinylated and acetylated starches (8.07 ± 0.03 and 8.06 ± 0.02 log CFU/g, respectively) did not show significant differences (at P < 0.05) between them. Phosphorylated starch had the lowest survival value (7.81 ± 0.04 log CFU/g) of microencapsulated B. breve ATCC 15700 (Fig. 3C). The water activity affected the survival of microencapsulated B. breve ATCC 15700 (P < 0.05). The best survival at 4°C and aw of 0.355 was for succinylated and acetylated amaranth starches (Fig. 3C). The wall materials used in this study showed
Fig. 6. Scanning electron microscopy image of native amaranth starch.
higher survival of cells at 4°C during all the evaluated storage time; the survival was maintained above 6.0 log CFU/g. The survival of L. casei ATCC 334 cells stored for 35 days at 4°C (Fig. 3D, E, and F) shows that for all the evaluated aw the survival was higher than 6.41 log CFU/g. The best viability for L. casei ATCC 334 was obtained at 4°C during 35 days storage and at aw of 0.355 with succinylated amaranth and control corn starches (8.62 ± 0.02 and 8.54 ± 0.00 CFU/g, respectively) (Fig. 3F). These results showed that it is possible to encapsulate L. casei with high survival values by using modified amaranth starch and spray drying, contrary to the recommendation of Anal and Singh (2007). These authors reported that spray drying is less recommended because of the high mortality of microorganisms from the simultaneous effects of dehydration and thermal inactivation. In this study, the concentration was kept higher than 106– 107 CFU/g for both probiotics, values recommended and necessary for practical applications. However, the resistance of free strains was not evaluated in the present study, because the main purpose was to compare the functionality of the modified starches with commercial modified starch. Because the best storage stabilities were presented by succinylated amaranth and control corn starch for both encapsulated probiotics, these starches were selected as wall materials to be incorporated into a food model system. A food system based on commercial precooked oatmeal containing the microencapsulated microorganisms was used to evaluate the tolerance to gastrointestinal conditions and to evaluate the survival during storage at 25°C. Tolerance to Simulated Gastrointestinal Conditions in a Food Model System. Figure 4 shows the tolerance to simulated gastrointestinal conditions of a food system based on commercial precooked oatmeal containing microencapsulated B. breve ATCC 15700 (Fig. 4A) and L. casei ATCC 334 (Fig. 4B). B. breve ATCC 15700 presented a reduction in survival (initial concentration – final concentration) of 1.42 ± 0.36 log CFU/g (17.3% of initial count) when the control corn starch was used; however, modified amaranth starch (succinylated), showed a lower reduction in survival of 1.29 ± 0.13 log CFU/g (15.5% of initial count). In the case of L. casei ATCC 334, that encapsulated with the control
Fig. 7. Scanning electron microscopy images of microcapsules obtained after spray drying containing encapsulated B. breve ATCC 15700 cells: A, acetylated amaranth starch; B, phosphorylated amaranth starch; C, succinylated amaranth starch; and D, control starch. 306
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corn starch had a reduction in survival of 1.70 ± 0.01 log CFU/g (20% of initial count), whereas succinylated amaranth starch showed a reduction in survival of only 0.36 ± 0.02 log CFU/g (4.4% of initial count), with greater resistance to the evaluated conditions. These results were favorable, because to exert their beneficial effects in the host, it is generally accepted that probiotic bacteria must be alive in the product at the time of consumption and also be capable of reaching the large intestine in high enough quantities to facilitate colonization and proliferation. Sultana et al (2000) microencapsulated Lactobacillus and Bifidobacterium cells with calcium alginate–starch and reported a decrease in survival of 2 log cycles for Lactobacillus and only a slight decrease for Bifidobacterium compared with the initial cell count when particles were exposed to in vitro conditions with a high amount of bile salts to simulate digestion. Petreska et al (2012) obtained symbiotic microparticles that were stable during exposure to simulated gastric and intestinal juices, allowing the release of viable cells above the therapeutic value in the simulated colonic pH. These authors produced the microencapsulation through spray drying combined with polyelectrolyte complexation of alginate, fructooligosaccharides, and chitosan and crosslinking with calcium chloride, followed by freeze-drying. Viability of Encapsulated Microorganisms in a Food Model System During Storage. Figure 5A shows the initial and end viability of B. breve, in which succinylated amaranth and control corn starch at the end of the storage time presented concentrations of 6.17 ± 0.02 and 4.10 ± 0.03 log CFU/g respectively. For the capsules with L. casei, both starches showed final concentrations over 6.0 ± 0.03 log CFU/g (Fig. 5B) during 35 days of storage at 25°C. During storage of blends of oat containing encapsulated probiotics (B. breve and L. casei), no visual changes were observed in the physical appearance of the oatmeal. The beneficial effect of B. breve and L. casei as a probiotic was added to the beneficial properties of oats. Weinbreck et al (2010) produced Lactobacillus rhamnosus GG microencapsulated with whey protein and palm oil as wall materials, evaluating the microencapsulated cells in infant milk formula during storage at 37°C and different water activities. These authors reported that microencapsulation with these materials offered no protection for probiotic microorganism viability during storage in dry products. Also, O’Riordan et al (2001) reported that microcapsules obtained with modified waxy maize starch with probiotics incorporated in dry foods with low water activity were not successful regarding the survival of the microorganisms. However, only a few works have addressed the effects of encapsulation on probiotic survival in dry (low water activity) food products. Morphology and Structure of Microcapsules. SEM photomicrographs of native amaranth starch (Figure 6) showed that the granules were about 1–3 μm in diameter and were angular and polyhedral in shape. As one example, Figure 7 shows microcapsules with encapsulated B. breve ATCC 15700 cells. The morphology of the microcapsules was similar for all types of wall starches (modified amaranth and control). The shape of the microcapsules was characteristic of microcapsules produced by spray drying, with concavities on the surface as a result of fast water evaporation. Similar results were reported by diverse authors (O’Riordan et al 2001; Crittenden et al 2006; Rodriguez-Huezo et al 2007; Ying et al 2010). Although the microparticles’ average size was not determined, the SEM micrographs allowed the estimation that the size of the microcapsules was lower than ≈20 μm for all the modified and control starches; this characteristic can contribute to reducing the impact on the texture when they are incorporated into a food model system. CONCLUSIONS The extrusion process enhanced the fragmentation of starch, producing encapsulating materials with improved characteristics
of solubility and viscosity for use in spray drying. In general, modified amaranth starches and control corn starch did not offer complete protection to both strains during storage at 35 days. Microencapsulated L. casei ATCC 334 stored at aw of 0.329 at room temperature for 35 days maintained the best survival in all the evaluated wall materials. Succinylated amaranth and control corn starches showed viability greater than 4 log CFU/g until the end of storage for all the evaluated water activities. However, the increase in water activity decreased the survival of microencapsulated cells in modified amaranth and control corn starches. Survival of cells of L. casei ATCC 334 was 8.62 ± 0.02 and 8.54 ± 0.00 CFU/g when encapsulated with succinylated amaranth and control corn starches, respectively, and stored at 4°C at aw of 0.355 during 35 days. Succinylated and acetylated amaranth starches had the best survival at 4°C and aw of 0.355. The survival of cells at 4°C during the storage time was maintained above 6.0 log CFU/g in all the evaluated wall materials. A good applicability of derivative starches containing probiotics in a food system based on commercial precooked oatmeal was obtained. ACKNOWLEDGMENTS We thank CINVESTAV-QRO for providing facilities to accomplish the present work, CONACYT for the Ph.D. degree scholarship provided for the first author, and both institutions for financial support for this project. LITERATURE CITED AACC International. Approved Methods of Analysis, 11th Ed. Method 61-02.01. Determination of the pasting properties of rice with the Rapid Visco Analyser. Approved October 15, 1997; reapproved November 3, 1999. AACC International: St. Paul, MN. http://dx.doi.org/10.1094/ AACCIntMethod-61-02.01 Akhiar, M. 2010. Enhancement of probiotics survival by microencapsulation with alginate and prebiotics. Basic Biotechnol. 6:13-18. Anal, A. K., and Singh. H. 2007. Recent advances in microencapsulation of probiotics for industrial applications and targeted delivery review. Trends Food Sci. Technol. 18:240-251. Anderson, R. A., Conway, H. F., Pfeifer, V. F., and Griffin, E. L., Jr. 1969. Gelatinization of corn grits by roll and extrusion cooking. Cereal Sci. Today. 14:4-12. Bhosale, R., and Singhal, R. 2006. Process optimization for the synthesis of octenyl succinyl derivative of waxy corn and amaranth starches. Carbohydr. Polym. 66:521-527. Blazenka, K., Jagoda, S., Jadranka, G., and Srecko, M. 2000. Viability of Lactobacillus acidophilus M92 in simulated gastrointestinal conditions. Food Technol. Biotechnol. 38:121-127. Brown, A. C., and Valiere, A. 2004. Probiotics and medical nutrition therapy. Nutr. Clin. Care 7:56-68. Cai, Y. Z., and Corke, H. 2000. Production and properties of spray-dried Amaranthus betacyanin pigments. J Food Sci. 65:1248-1252. Chang, Y. H., and Lii, C. Y. 1992. Preparation of starch phosphates by extrusion. J. Food Sci. 57:203-205. Crittenden, R., Weerakkody, R., Sanguansri, L., and Augustin, M. 2006. Symbiotic microcapsules that enhance microbial viability during nonrefrigerated storage and gastrointestinal transit. Appl. Environ. Microbiol. 72:2280-2282. Drusch, S., Serfert, Y., Scampicchio, M., Schmidt-Hansberg, B., and Schwarz, K. 2007. Impact of physicochemical characteristics on the oxidative stability of fish oil microencapsulated by spray drying. J. Agric. Food Chem. 55:11044-11051. Greenspan, L. 1977. Humidity fixed points of binary saturated aqueous solutions. J. Res. NBS A Phys. Chem. 81:89-96. Harper, J. M. 1992. Extrusion processing of starch. Pages 37-64 in: Developments in Carbohydrate Chemistry. R. J. Alexander and H. F. Zobel, eds. American Association of Cereal Chemists: St. Paul, MN. Hsiao, H. C., Lian, W. C., and Chou, C. C. 2004. Effect of packaging conditions and temperature on viability of microencapsulated bifidobacteria during storage. J. Sci. Food Agric. 84:134-139. Ishibashi, N., and Shimamura, S. 1993. Bifidobacteria: Research and development in Japan. Food Technol. 47:126-136. Kailasapathy, K., and Rybka, S. 1997. L. acidophilus and Bifidobacterium spp.—Their therapeutic potential and survival in yoghurt. Aust. J. Vol. 91, No. 3, 2014
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[Received June 5, 2013. Accepted December 30, 2013.]
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