Effect of different calcium salts and methods for triggering gelation on the characteristics of microencapsulated Lactobacillus plantarum LIP-1

Abstract

Probiotic Lactobacillus plantarum isolate LIP-1 was microencapsulated in milk protein matrices by means of rennet-induced gelation combined with an emulsification technique. The effect of different calcium salts (CaCO₃ and CaCl₂) and different methods for triggering gelation (acid-triggered versus temperature-triggered) on the characteristics of microencapsulated Lactobacillus plantarum LIP-1 were investigated. The results indicated that microcapsules prepared by CaCO₃ with acid-triggered gelation showed significantly better qualities than microcapsules prepared by CaCl₂ with temperature-triggered gelation. Specifically: microencapsulation efficiency (ME), particle size and size distribution (as observed by optical microscopy), the surface morphology and the microstructure (as observed by scanning electron microscopy), resistance to passage through simulated gastric fluid (SGF), release characteristic in simulated intestinal fluid (SIF) and viability during 4 weeks storage at different temperatures (4 °C, 20 °C and 37 °C) were all better in the microcapsules prepared by CaCO₃ with acid-triggered gelation. For these reasons we propose that CaCO₃ with acid-triggered gelation might be a better choice for crosslinking casein to form microcapsules.

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Introduction

Probiotics are defined as living microorganisms that, when consumed in adequate quantities, confer a health benefit on the host¹ (Homayouni et al., 2007). Consistent with increasing demand, probiotics have been used widely in the production of functional foods, especially dairy foods such as yoghurt and cheese, which represent about 65% of the global functional food market² (Agrawal, 2005).

It is generally believed that the therapeutic minimum level of probiotics should be at least 10⁷ cfu g⁻¹ or mL of live cells in the product³ (Jankovic et al., 2010), but the actual levels detected in probiotic products are often much lower due to adverse conditions during product storage, transportation, marketing and consumption. Factors that influence the stability of probiotics include food composition (e.g. pH, moisture content, hydrogen peroxide concentration, molecular oxygen availability and additives), strains of probiotic bacteria, microbial interactions, heat treatment during the production process, storage temperature and host's own digestive system (e.g. gastric acid, bile salts, enzymes)^4–6 (Malmo et al., 2013; Akalın et al., 2004; Ross et al., 2005).

Studies have shown that microencapsulation techniques can provide protection for probiotics as they represent a physical barrier against adverse environmental conditions, increasing the viability of probiotics in food matrices, increasing stability during storage and protecting against the host's digestive system.^7,8 (Anal & Singh, 2007; Kailasapathy, 2002). There are various types of coatings and mixtures of effective biopolymers that have been used for microencapsulation of probiotics, such as alginate, gellan-gum, xanthan and κ-carrageenan⁹ (Heidebach et al., 2010). However, they are all of non-dairy origin and, therefore, undesirable or prohibited from use in dairy products in some countries¹⁰ (Picot & Lacroix, 2004).

Milk proteins as matrix materials possess well-known physico-chemical properties that could make them appropriate for use as encapsulating agents in food production. Milk proteins are generally recognized as safe (GRAS) raw materials with high nutritional value and good sensory properties¹¹ (Livney, 2010). They also have the properties required for the delivery of functional ingredients. For example, they can bind small molecules and ions; they have self-assembly properties; superb gelation properties; pH-responsive gel-swelling behaviour; they interact with other polymers to form complexes; are biocompatible and biodegradable; and have an excellent buffering capacity against the harsh acid environment of the stomach^12,13 (El-Salam and El-Shibiny, 2012; Doherty et al., 2012) These attributes mean they are capable of controlling the bioaccessibility of bioactive substances. Furthermore, they have good emulsification properties, are highly soluble and, as they have a bland flavour they do not interfere with the sensory properties of the product^9,14 (Heidebach et al., 2010; Heidebach et al., 2009). Another potential benefit associated with protein encapsulation matrices involves hydrolysis of milk proteins by digestive enzymes potentially generating bioactive peptides that may exert a number of physiological effects in vivo^15,16 (Kilara & Panyam, 2003; Korhonen & Pihlanto, 2003). These properties indicate that dairy proteins are good candidates for encapsulating probiotics, and as such they are attracting more research attention² (Agrawal, 2005).

For example, Heidebach et al. (2009)¹⁴ used rennet-induced gelation of milk proteins in reconstituted skimmed milk powder with emulsification to encapsulate probiotics. In their experiment a soluble calcium salt, CaCl₂, was added to the suspension of cold milk protein and probiotic cells during renneting, and the mixture dispersed in cold oil resulting in a water-in-oil-emulsion (W/O) that was further emulsified until a sufficiently small droplet size was achieved (<5 °C). Subsequently the temperature was raised to 40 °C for 15 min to complete the encapsulation reaction. The method was based on a temperature-triggered instant gelation effect; although the κ-casein was cleaved because the renneting was conducted at low temperatures, the resulting micelles did not coagulate, until the temperature was raised to above 18–20 °C, where a gel was formed instantly¹⁷ (Bansal et al., 2007). However, during the rise in temperature, disruption of the emulsion system equilibrium may sometimes cause significant clumping of the microcapsules before they become properly hardened, leading to reduced encapsulation efficiency and stability. The step involving raising the temperature from 5 °C to 40 °C also requires a lot of energy and can cause microbial contamination during industrial production.

Based on the method of Heidebach et al. (2009),¹⁴ the objective of this study was to develop a modified encapsulation process based on emulsification and subsequent acid-triggered gelation by means of cold renneting of skim-milk concentrates and the insoluble calcium salt, CaCO₃, for the production of dairy-based microcapsules of the probiotic Lactobacillus plantarum LIP-1. Furthermore, the characteristics of microencapsules prepared using two different calcium salts and different methods for triggering gelation were compared. Characteristics included microencapsulation efficiency, particle size and size distribution, surface morphology and microstructure, and their ability to protect probiotic cells during subsequent exposure to different environments.

Experimental

Materials

Lactobacillus plantarum LIP-1 originated from the culture collection of the Key Laboratory of Dairy Biotechnology and Engineering, Ministry of Education PRC, Inner Mongolia Agricultural University. Lactobacillus plantarum LIP-1 was identified as a probiotic isolate during screening of lactic acid bacteria from koumiss samples collected in Inner Mongolia; our research showed that L. plantarum LIP-1 had cholesterol-reducing health benefits¹⁸ (Wang et al., 2012).

Skimmed milk powder was obtained from Fonterra Ltd. (Auckland, New Zealand) and the rennet, with a declared activity of 1070 IMCU mL⁻¹, from Shanghai Aijie Biological Technology Co. Ltd. (Shanghai, China). A fresh stock solution of rennet was prepared daily by diluting 1 g rennet preparation in 10 g double-distilled water. Sunflower oil was purchased from JingLongYu Ltd. (Chengdu, China). CaCl₂, CaCO₃, glacial acetic acid and other chemicals used in this study were all of analytical grade and purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China).

Lactobacillus plantarum LIP-1 culture and production of cell suspension

Lactobacillus plantarum LIP-1 was grown in Man, Rogosa and Sharpe (MRS) broth (Oxoid Ltd., Basingstoke, Hampshire, England) at 37 °C for 18 h in an incubator (SANYO Electric Co., Ltd., Japan). The bacteria were subcultured three times prior to being used for the preparation of cell suspensions. They were then centrifuged using a centrifuge (Anhui USTC Zonkia Scientific Instruments Co., Ltd., China) at 1000 g for 10 min. The supernatant was discarded and the cell precipitate was washed thoroughly with sterilized 0.85% NaCl solution. This washing process was repeated two times and the final cell slurry was made up to 2 mL by adding sterilized 0.85% NaCl solution and mixing thoroughly. For enumeration of bacteria, the number of colony forming units (cfu) was determined on MRS agar using the plate count method at 37 °C for 48 h^19,20 (Grosso et al., 2004; Pedroso et al., 2012).

Preparation of the milk-L. plantarum-cell suspension

All glassware used were sterilized at 121 °C for 30 min. Skimmed milk powder was dispersed in double-distilled water to obtain a 30% (w/w) solution and stirred overnight at 4 °C. 2.0 g of the concentrated LIP-1 cell suspension was mixed with 28.0 g of the skimmed milk to achieve a culture with a concentration of approximately 10⁹ to 10¹⁰ cfu mL⁻¹ of probiotic culture.

Microencapsulation

Two microencapsulation methods were compared: In the first method microcapsules were prepared using CaCl₂ according to previously described methods¹⁴ (Heidebach et al. 2009). Firstly, 30 g of the L. plantarum cell suspension was cooled to 4 °C with 400 μL rennet stock-solution (100 IMCU mL⁻¹), and incubated at 4 °C for 60 min until cleavage of κ-casein had finished (renneting). Then 60 μL 10% (w/v) CaCl₂ solution was added to 10 g of the cold-renneted suspension, and the entire mixture added to 100 g of tempered (4 °C) vegetable oil to initiate the encapsulation process. To achieve emulsification the mixture was stirred for 15 min using a magnetic stirring apparatus. The temperature was maintained at 4 °C throughout the process to prevent premature gelling. Following the emulsification process at 4 °C, the temperature was raised to 40 °C over a period of 15 min during which time it was stirred continuously. When the temperature exceeded 18–20 °C the emulsified droplets turned into gel particles instantly. Subsequently, the gelatinized microcapsules were separated from the oil by gentle centrifugation (500 g, 1 min). The supernatant was removed and the sediment was diluted with approximately twice its volume of double-distilled water, shaken for 1 min and then centrifuged again under the same conditions. The supernatant, consisting of water and residual oil was removed. The virtually oil-free aqueous microcapsule-slurry was stored at 4 °C before experimentation.

The second method was a modified encapsulation process using CaCO₃ as follows: After 60 min incubation (4 °C), 1.5 g CaCO₃ was added to 10 g of the cold-renneted suspension. This was then added to 100 g vegetable oil and stirred at 800 rpm for 15 min to emulsify the mixture into the oil. To this 100 μL of glacial acetic acid was added to achieve dissociation of the Ca²⁺ and for the κ-casein to become cross-linked by Ca²⁺ within the emulsion droplets. After allowing the mixture to stand for 30 min, the microcapsules were generated. All steps were done at room temperature except the cleavage of the κ-casein. Subsequent centrifugation and dilution was the same as in the first method, and the resulting microcapsule-slurry was stored at 4 °C prior to experimentation.

Microencapsulation efficiency

Cell viability within both types of microcapsules was determined by counting the number of colony forming units (cfu g⁻¹) developing on MRS agar using a pour-plate technique described previously. Plates were incubated at 37 °C for 48 h before the counts were taken. Microencapsulation efficiency (ME) was calculated using eqn (1):

ME (%) = N/N₀ × 100% (1)

where N is the number of microencapsulated cells released from the microcapsules (log cfu g⁻¹ microcapsule) and N₀ is the number of free cells added to the L. plantarum LIP-1 suspension during the production of the microcapsules (log cfu g⁻¹).

Effect of freeze-drying

The microcapsule slurries produced in the two different ways were each frozen for 24 h at −80 °C before drying under 90 mTorr vacuum in a freeze dryer (SANYO Electric Co., Ltd., Japan)²¹ (Amine et al., 2014), with the cell-free protection agent (10% skimmed milk and 0.1% L-monosodium glutamate) as the control. The survival of encapsulated LIP-1 after freeze-drying was expressed using eqn (2):

Cell survival (%) = N₂/N₁ × 100% (2)

where N₁ is the initial number of microencapsulated cells before freeze-drying (log cfu g⁻¹ microcapsule); and N₂ represents the number of viable cells after the freeze-drying (log cfu g⁻¹ microcapsule).

Analysis of microcapsule morphology and particle size

The morphology of the microcapsules produced using different calcium salts was observed by scanning electron microscope (SEM) (SEM-SU8010, Hitachi, Tokyo, Japan). The samples were fixed to double sided conductive tape on top of SEM stub and then sputter-treated in a metallizer (Agar Sputter Coater) with gold-palladium to reach a thickness of coating of 100 A and then observed in high vacuum mode.

The average particle size and size distribution of the microcapsules were determined by Leica Confocal Laser Scanning Microscope (TCS SP-2, Leica Inc., Wetzlar, Germany). To calculate their diameter, at least 50 particles from each of the different formulations of microcapsules were measured at a magnification of 20ײ² (Krasaekoopt, Bhandari, & Deeth, 2004).

Release of encapsulated LIP-1 under simulated intestinal conditions

The release of encapsulated LIP-1 under simulated intestinal pH conditions was investigated according to previously described methods^23,24 (Mokarram et al., 2009; Mandal et al., 2006), with some modifications. To prepare simulated intestinal fluid (SIF), the pH of a 0.85% NaCl solution was adjusted to 8.0 using NaOH (1 M), then sterilized by autoclaving at 121 °C for 15 min. Trypsin (1 : 250 μ mg⁻¹, High Purity Grade; Biotopped Co., Ltd., Beijing, China) was then suspended in the solution to achieve a final concentration of 0.1% (w/v) and the solution was sterilized by filtration (0.22 mm). Samples (0.5 g) of the two kinds of freeze-dried microcapsules were each suspended in 4.5 mL of sterile SIF and incubated at 37 °C with constant agitation at 100 rpm. Aliquots of 1.0 mL were taken at 0, 30, 60, 90, 120 and 150 min and the survival of encapsulated LIP-1 released in to the SIF was determined using the pour-plate technique as described previously.

Survival of encapsulated and free cells in simulated gastric fluid

Simulated gastric fluid (SGF) were prepared and survival of encapsulated cells in SGF was determined according to previously described methods^25–27 (Dolly et al., 2011; Rajam et al., 2012; Sohail et al., 2011), with slight modification. To prepare simulated gastric fluid (SGF), the pH of a 0.85% NaCl solution was adjusted to 2.0 using 1 M HCl and sterilized by autoclaving at 121 °C for 15 min. Pepsin (1 : 3000 NFU mg⁻¹, High Purity Grade; Biotopped Co., Ltd., Beijing, China) was then suspended in the solution to achieve a final concentration of 0.3% (w/v). Samples (0.5 g) of the two kinds of freeze-dried microcapsules were each suspended in 4.5 mL of sterile SGF and incubated at 37 °C with constant agitation at 100 rpm. The control was 0.5 g of the L. plantarum suspension (free cells). Aliquots of 0.5 mL were taken at 30, 60, 90, 120 and 150 min. The survival of encapsulated L. plantarum released in to SGF was determined using the pour-plate technique as described previously and compared with the free cells control.

Survival of microencapsulated LIP-1 during storage

Samples of each of the two types of freeze-dried microcapsules were placed in sealed glass vials and stored at 4 °C, 20 °C and 37 °C for 28 days. At weekly intervals, the survival of bacterial cells was determined by enumeration on MRS agar, using the pour-plate technique as described previously.

Statistical analysis

All experiments were carried out in triplicate in a completely randomized design and the data were analyzed using SPSS Statistics software, version 22.0 (IBM SPSS Statistics 22). The results were expressed as means ± S.D. Data comparisons were made using the Student Newman Keuls test to determine whether there were significant differences (p < 0.05) between treatment means.

Results and discussion

Microencapsulation efficiency

Microencapsulation efficiency (ME) is an important index for microcapsule evaluation, because it is crucial to maintain high viability within the microcapsules to achieve the full benefits of LIP-1. The ME for viable cells in microcapsules prepared by CaCO₃ (75.6 ± 5.1%) was significantly greater (p < 0.05) than for viable cells in microcapsules prepared by CaCl₂ (58.4 ± 11.3%) (Table 1). The high ME may be attributed to the fact that the entire process involved no detrimental steps such as heat treatment or high shear forces. The ME appeared to be in agreement with the results of Heidebach et al. (2009),¹⁴ who used a similar process with a sodium caseinate emulsion. Shi et al. (2013)²⁸ encapsulated L. bulgaricus in skimmed milk–alginate and an extrusion method, and the ME was around 60%. In the present study, a slight loss of cells may be because the cells present on the droplet surface were lost into the oil phase or drained with the washing water.

Table 1 The microencapsulation efficiency (ME) and size of microcapsules produced in two different ways^a

The type of microcapsules	Microencapsulation efficiency (%)	Microsphere size (μm; n = 50)
a AB means in the same column followed by a different upper case letter are significantly different (P < 0.05).
CaCl2	58.4 ± 11.3^A	109.05 ± 19.53^A
CaCO3	75.6 ± 5.1^B	143.57 ± 29.18^B

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Although the microcapsules produced by insoluble CaCO₃ and acid-triggered gelation had a higher ME than those produced by CaCl₂ and temperature-triggered gelation, the specific mechanism is not entirely clear. It may be because CaCl₂ microcapsules produced by emulsification are formed instantly when the temperature is raised and Ca²⁺ crosslinks rapidly with casein, potentially resulting in some premature gelling that could limit the number of viable cells embedding in the microcapsules. In contrast CaCO₃ releases Ca²⁺ for crosslinking with casein within W/O gelation when the glacial acetic acid is added; in this way the dense structure of the microcapsule enables the immobilized L. plantarum to remain entrapped, thereby increasing the ME.

Particle size and distribution

Particle size and its distribution are important physical properties that directly affect the use of microcapsules into food formulations. The results showed that the mean diameters of microspheres prepared by CaCO₃ (109.05 ± 19.53 μm) were significantly smaller than by CaCl₂ (143.57 ± 29.18 μm) (p < 0.05) (Table 1). Fig. 2 indicated that the beads produced by the two methods exhibited nearly unimodal and normal particle-size distribution, and the size distribution of two microcapsules was significantly different. Microspheres prepared by CaCl₂ exhibited a wide range of sizes varying from 107.86 μm to 172.75 μm diameter, and 78% of the particles were between 120 and 160 μm. While microcapsules prepared by CaCO₃ showed a narrower size distribution ranging from 89.52 μm to 143.84 μm, and 82% of the microspheres were between 90 and 120 μm (Fig. 1 and 2).

Fig. 1 Optical images of two different microencapsulations: (a) prepared by CaCO₃; (b) prepared by CaCl₂ (bar = 100 μm).

Fig. 2 Particle mean size distributions of two different microencapsulations.

It has been known that the size and size distribution of the microspheres could be influenced by the concentration of skimmed milk, the ratio of water to oil (v/v), the stirring speed, and so on²⁹ (Lin et al., 2007) In this study, the microspheres were prepared to ensure the comparability between both methods at given conditions, that is, the same batch of skimmed milk, stirring speed of 700 rpm and the ratio of water to oil of 1 : 10. Therefore the variations in particle size and distribution between two microspheres might be due to the different Ca²⁺ used for encapsulation. For microspheres producted by CaCl₂, Ca²⁺ was distributed in the emulsion system before temperature raised, and the larger mean size was attributed to the collision of the dispersed Ca²⁺ and skimmed milk droplets. The collision could involve two or more droplets, which cause the aggregation of microcapsules³⁰ (Silva et al., 2006) While CaCO₃ releases Ca²⁺ for crosslinking with casein within W/O gelation when the glacial acetic acid is added, so the size distribution of microspheres prepared by CaCO₃ depends on the size of the emulsion droplets, which is determined by a balance between the dispersive and the surface tension forces. So the latter tends to well-dispersed and the former causes coalescence³¹ (Chan, Lee, & Heng, 2002).

Several researchers reported that the optimum microcapsule size is a compromise between the effectiveness of encapsulation and the sensory properties³² (Arup et al., 2011). A minimum diameter of 100 μm has been suggested to offer better protection in gastric juice, and an optimum range of 100–200 μm has been proposed^32,33 (McMaster et al., 2005; Nag et al., 2011). The mean diameter of our two microcapsules were both within this suggested range.

Effect of freeze-drying

Probiotic cells should be capable of being dried so that they can serve as a functional ingredient in food products but also be easily transported. Otero et al. (2007)³⁴ suggested that freeze-drying might be appropriate for this as it is a well-documented technique used to obtain stable cultures in terms of viability and functional activity. However, either freezing or freeze-drying of LIP-1 could reduce its viability due to deterioration of the physiological state of the cells^35,36 (Chávarri et al., 2010; Crittenden, et al., 2006). De Giulio et al. (2005)³⁷ reported that, during freeze-drying, bacteria were subjected to adverse and potentially lethal conditions, such as water crystallization, protein denaturation and bacterial membrane injury. The bead manufacturing process had the most influence on subsequent viability following freeze-drying, but microencapsulated bacteria were also more stable than free cells.

In our study, however, encapsulated LIP-1 cells prepared by either CaCO₃ or CaCl₂ retained significantly greater (P < 0.05) viability than free cells (Table 2). The survival of encapsulated LIP-1 were above 60% while free cell was about 8% after freeze drying for 24 h. There was no significant difference (P > 0.05) in survival after freeze-drying of encapsulated L. plantarum prepared either using CaCO₃ or CaCl₂.

Table 2 Survival rate during freeze drying (%), of LIP-1 as free cells or encapsulated using either CaCO₃ or CaCl₂^a

	CaCO3	CaCl2	Free cell
a ab means in the same row followed by different lower case letters are significantly different from each other (P < 0.05).
Survival rate (%)	64.733 ± 0.210^a	63.867 ± 0.560^a	7.933 ± 0.470^b

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The beneficial effect of microencapsulation prior to freeze-drying has often been reported^38–40 (Homayouni et al., 2008; Shah and Ravula, 2000; Sheu and Marshall, 1993), and so the data from this study are in line with the existing literature.

Overall, cell mortality during the freeze-drying process is known to be mostly due to injuries to the membranes of cells.^41,42 (Carvalho et al.,2003; Saarela et al., 2005) These injuries include structural changes to sensitive proteins resulting from intracellular ice formation, the physical status of membrane lipids, osmotic shock and recrystallization. Consequently, complete protection of the membrane of cells via coating techniques, as we saw in this study (Table 2), could be very effectual. Shah & Ravula (2000)³⁹ also reported that microencapsulation of probiotic bacteria prior to freeze-drying for incorporation into frozen fermented dairy desserts was very effective for improving cell stability during the freeze-drying process. They noted that encapsulation protected the cells from cold shock induced by the process.

Skimmed milk proteins have been reported widely as very effective cell protectants during freeze-drying, a function that has been attributed to the presence of lactose; lactose has the ability to interact with cell membranes retaining their structural integrity⁴³ (Corcoran et al., 2004). Moreover, as a disaccharide, lactose can depress the gel to liquid crystalline state transition temperature of the lipid bilayer of cell membranes, aiding cells to retain their natural structural conformation⁴⁴ (Fu & Chen, 2011). This mechanism could also be extended to other milk protein formulations with similar compositional elements such as whey protein concentrates milk protein concentrates or caseinates. Another mechanism that could also potentially affect the survival rate of LIP-1 during the freeze-drying process, is the ability of proteins to bind water through hydrogen bonds⁴⁵ (Kinsella and Morr, 1984), which can protect cellular membranes from dehydration. From our data, it seems that a synergism between these mechanisms could explain the protection of LIP-1 by skimmed milk protein microcapsules.

Morphology of microcapsules

The optical and SEM morphology of two different microencapsulations were presented in Fig. 1, 3 and 4. As seen in Fig. 1, microcapsules prepared using CaCO₃ were smaller, more homogeneous and showed a narrower size distribution than microcapsules prepared using CaCl₂. Fig. 3 showed the morphology of two different microencapsulations dyed by methylene blue, in which we can see that many LIP-1 were involved in skimmed milk microcapsules. And Fig. 4 showed the morphology of two different microcapsules by SEM.

Fig. 3 Morphology of two different microencapsulations dyed by methylene blue: (a) prepared by CaCO₃ (bar = 50 μm); (b) prepared by CaCl₂ (bar = 50 μm).

Fig. 4 SEM images of two different microencapsulations: (a) prepared by CaCl₂ (bar = 50 μm); (b) prepared by CaCl₂ (bar = 5 μm); (c) prepared by CaCl₂ (bar = 300 μm); (d) prepared by CaCO₃ (bar = 100 μm); (e) prepared by CaCO₃ (bar = 5 μm); (f) the section of microencapsulation prepared by CaCO₃ (bar = 5 μm).

As seen in Fig. 4, microcapsules prepared using CaCl₂ and temperature-triggered gelation, were non-spherical and had an porous organization (Fig. 4a and b). We believe these characteristics may due to the fast sublimation of frozen water from the skimmed milk matrix resulting in the formation of pores in areas where ice crystals had been present and had not enough time to shrink, as has been observed by others⁴⁶ (Smrdel et al., 2008). While microcapsules prepared by CaCO₃ with acid-triggered gelation were smoother (Fig. 4d). When compared at a higher resolution (8000 × ), there was a more compact substructure in microcapsules produced using CaCO₃ (Fig. 4e) than microcapsules produced by CaCl₂ (Fig. 4b). Moreover, some LIP-1 were discovered in the section of microencapsulations in Fig. 4f, which is a stronger evidence that LIP-1 being encapsulated in the microencapsulations.

In the CaCl₂ microcapsules produced by emulsification the crosslinking was achieved instantly by raising the temperature during stirring, and this may be why the microcapsules are always agglomerated (Fig. 4c). By the time the calcium solution contacts the dispersed casein phase, gelling occurs. Thus, the gelation kinetic is heterogeneous, which has often been shown previously to lead to capsules of irregular shape and agglomerated^47,48 (Muthukumarasamy et al., 2006; Sheu et al., 1993). In contrast to this, glacial acetic acid induced Ca²⁺ release into casein suspensions from CaCO₃ underwent a homogeneous internal gelation process, leading to dispersed and homogeneous small microcapsules, which are preferable because there are fewer negative sensorial impacts on the resulting food. So the use of CaCO₃ always induced a compact microcapsule structure, regular shape and a finer more dispersed substructure.

Release of encapsulated probiotics under simulated intestinal condition

The release of cells from microcapsules in the colon is essential for growth and colonization of probiotics. If this does not happen then the probiotics in the microcapsules will be washed out of the body before they have exerting any beneficial effect⁴⁹ (Sabikhi et al., 2010). In our study the encapsulated L. plantarum prepared using CaCO₃ was completely released within 90 min and, in fact, more than 70% was released within 30 min (Fig. 5). In contrast the encapsulated L. plantarum prepared by CaCl₂ took 120 min to be fully released, and by 30 min less than 40% had been released (Fig. 5). The release profile indicated that CaCO₃ microcapsules had significantly faster release properties in SIF than CaCl₂ microcapsules. Many studies have investigated the release of encapsulated probiotics in SIF. For example Sabikhi et al. (2010)⁴⁹ reported that microencapsulated Lactobacillus acidophilus were released after 2.5 h in SIF (pH 7.4 ± 0.2), which was actually slower than in our study. We speculate that differences in speed of release amongst different types of microcapsules, is due to the difference in size and distribution status of those microcapsules⁵⁰ (Chen and Subirade, 2006). In our study the microcapsules produced by CaCO₃ were dispersed better and the particle size was smaller and more homogeneous compared with microcapsules produced using CaCl₂, which were irregular and agglomerated (Fig. 1, and 4). This resulted in a larger specific surface area for enzyme activity, increasing the speed of disintegration.

Fig. 5 Release characteristics of encapsulated cells in SIF. Bars labelled with a different lower case letter (e.g. a, b or c) means in the same microencapsulation methods with different incubation time are significantly different from each other (P < 0.05). Bars labelled with a different upper case letter (e.g. A, B or C) means at the same incubation time with different microencapsulation methods are significantly different from each other (P < 0.05).

Survival of encapsulated and free cells of LIP-1 in simulated gastric juice

Probiotic bacteria are, in general, sensitive to acidic environments, which presents a challenge for the industrial use of these bacteria as well as for consumer effectiveness. In order to exert positive health effects, probiotics should resist the harsh conditions of the stomach to remain active when they reach the colon. One main purpose of encapsulation is to improve the tolerance of probiotics to low pH in the stomach.

In our study free cells of LIP-1 were sensitive to low pH and they lost 2 log cfu g⁻¹ when exposed to acidic conditions for 120 min (Fig. 6). This is in line with many articles reporting that unprotected cells of probiotics are easily damaged by stomach acid. For example, Heidebach et al. (2009)¹⁴ reported that Lactobacillus paracasei isolate F19 lost viability almost entirely within 60 min at a SGF pH of 2.5. In contrast, encapsulation of LIP-1 in skimmed milk microspheres significantly improved survival in SGF (pH 2.0) compared with free cell (p < 0.05) (Fig. 6). Furthermore, microspheres prepared by CaCO₃ with acid-triggered gelation survived significantly longer than microcapsules produced by CaCl₂ (p < 0.05) (Fig. 6). In SGF, the viable counts of LIP-1 that had been encapsulated by either method were greater than 8 log cfu g⁻¹ after 150 min exposure (Fig. 6). Excellent pH tolerance of encapsulated LIP-1 was probably due to the buffering ability of skimmed milk and low porosity on the surface of the microspheres. Our findings agree with those of other studies. Guérin et al. (2003)⁵¹ reported that the buffering ability of whey protein contributed to high survival rates for Bifidobacterium spp. encapsulated in alginate–pectin–whey protein microspheres when exposed to SGF at pH 2.5. Heidebach et al. (2009)¹⁴ reported that viability of L. paracasei and Bifidobacterium lactis exposed to SGF at pH 2.5 was improved when they were encapsulated in casein microcapsule. Gbassi et al. (2009)⁵² reported that viability of L. plantarum isolates was improved when encapsulated in whey protein microspheres.

Fig. 6 Survival of encapsulated L. plantarum in SGF. Bars labelled with a different lower case letter (e.g. a, b or c) means in the same microencapsulation methods with different incubation time are significantly different to each other (P < 0.05). Bars labelled with a different upper case letter (e.g. A, B or C) means at the same incubation time with different microencapsulation methods are significantly different to each other (P < 0.05).

After 150 min of exposure to SGF the viability of cells encapsulated using CaCO₃ had dropped to 8.591 log cfu g⁻¹ compared with cells encapsulated using CaCl₂ where the viability had dropped to 8.190 log cfu g⁻¹ under the same conditions, demonstrating that the microencapsulation process using CaCO₃ provided much better protection for L. plantarum under acidic conditions. This might be explained by the different surface structure of the CaCO₃ microcapsules compared with the CaCl₂ microcapsules. As the surface of the CaCO₃ microcapsules is dense, it is likely to be more resistant to the penetration of SGF (Fig. 4e). In contrast, the surface of the CaCl₂ microcapsules was porous allowing the SGF to enter easily (Fig. 4b).

Survival of microencapsulated LIP-1 during storage

One of the key properties for any microorganism that is being considered as a probiotic is its capacity to survive storage as a formulated product⁵³ (Sathyabama & Vijayabharathi, 2014). Many studies have reported that encapsulation could improve the viability of probiotics during storage. For example, Su et al. (2011)⁵⁴ reported that alginate–human-like-collagen microspheres improved the stability of Bifidobacterium longum isolate BIOMA 5920 during storage for 3 weeks compared to free cells. Brinques & Ayub (2011)⁵⁵ concluded that an alginate–chitosan system had good potential to improve storage stability of L. plantarum isolate BL011, which lost little viability during the storage period evaluated.

In our study the encapsulating material, temperature, and storage time all significantly affected the viability of LIP-1 (Fig. 7). Viability of LIP-1 encapsulated using CaCl₂ with temperature-triggered gelation was significantly lower than when encapsulated by CaCO₃ with acid-triggered gelation (P < 0.05). For CaCO₃-produced microcapsules the average number of viable cells was 7.724 ± 0.328 log cfu g⁻¹ at 4 °C after 28 days, which was significantly higher (p < 0.05) than that of microcapsules stored at 20 °C (7.176 ± 0.414 log cfu g⁻¹) and 37 °C (6.275 ± 0.496 log cfu g⁻¹). For CaCl₂-produced microcapsules, there was also a smaller decrease in viability of L. plantarum (6.362 ± 0.325 log cfu g⁻¹) at 4 °C compared with storage at 20 °C (5.987 ± 0.286 log cfu g⁻¹) and 37 °C (4.886 ± 0.375 log cfu g⁻¹). Regardless of storage temperature, the pattern of reduction in viability was similar for all treatments and, even after 28 days storage, the viability of L. planttarum encapsulated by CaCO₃ remained higher than 6 log cfu g⁻¹ (Fig. 7), the threshold value recommended for a food to exert its probiotic benefits^56,57 (Hamilton-Miller et al., 1999; Reid et al., 2003).

Fig. 7 Survival of encapsulated LIP-1 during storage at different temperatures. Bars labelled with a different lower case letter (e.g. a, b or c) means in the same microencapsulation methods with different incubation time are significantly different to each other (P < 0.05). Bars labelled with a different upper case letter (e.g. A, B or C) means at the same incubation time with different microencapsulation methods are significantly different to each other (P < 0.05).

The maintenance of high viable cell counts after 28 days of storage, as presented in this study, may be attributed to several factors, such as the intrinsic resistance characteristics of the isolate we used, the encapsulating material, the low moisture content of the particles, the type of packaging and storage temperature^58,59 (Morgan et al., 2006; Abe et al., 2009). The barrier characteristics of the packaging material and the structure of microcapsules may have contributed to the microbiological stability of LIP-1 during storage. Milk protein is a good barrier against water vapor, gas and light, thereby preventing photo-oxidation and minimizing the oxygen levels in the interior of the package⁶⁰ (Alves et al., 2008). Furthermore, the milk protein present in the packing may have contributed to minimizing oxygen diffusion to the cell membrane, favouring survival during storage^61,62 (Teixeira et al., 1996; Chávez and Ledeboer, 2007). Rodrigues et al. (2011)⁶³ embedded Lactobacillus acidophilus isolate Ki, L. paracasei isolate L26 and Bifidobacterium animalis isolate BB-12 in whey protein, and the viability was higher than 6 log cfu g⁻¹ after storage for up to 6 months at 5 °C.

From the morphology visible using SEM we know that the microcapsules prepared using CaCl₂ are porous, while CaCO₃-produced microcapsules are compact (Fig. 4). Thus, microcapsules prepared using CaCO₃ provided better protection of LIP-1 than CaCl₂-produced microcapsules, and a higher viability after storage for 28 days. Shi et al. (2013)²⁸ studied storage stability of L. bulgaricus in alginate–milk microspheres, and found that full viability of encapsulated L. bulgaricus could be preserved during storage for 1 month at 4 °C. This was probably due to the dense alginate–milk membrane formed around the microcapsules, which may have provided effective protection. In our study, it is possible that denser skimmed milk membranes were produced in CaCO₃-produced microcapsules than CaCl₂-produced microcapsules and that this provided better protection for LIP-1 during storage.

Conclusions

This investigation compared the effect of CaCl₂ and temperature-triggered gelation with CaCO₃ and acid-triggered gelation on the biological characteristics and stability of microencapsulated LIP-1. The type of calcium salt and the method for triggering gelation significantly affected the surface and microstructure of the resulting microcapsules. The use of CaCO₃ with acid-triggered gelation provided better protection for LIP-1 against freeze-drying and adverse gastrointestinal conditions than CaCl₂ and temperature-triggered gelation. The CaCO₃ encapsulation processing method is not only easy to scale up, but also provides the encapsulated probiotics with better long-term preservation at different temperatures. These results demonstrate that CaCO₃ with acid-triggered gelation might be a better method for the preparation of microencapsulated LIP-1, than the CaCl₂ and temperature-triggered gelation method in order to increase protection against deleterious environmental factors. This study could also be useful for other probiotics used in the production of functional food or other related products.

Acknowledgements

This study was supported financially by Natural Science Foundation of China (Grant No. 31160315), Natural Science Foundation of Inner Mongolia (Grant No. 2015MS0306), the ‘Western Light’ Talent Cultivation Program of CAS and the Key Laboratory of Dairy Biotechnology and Engineering, Ministry of Education PRC, Inner Mongolia Agricultural University. The authors gratefully acknowledge Dr Wang for scientific advice relating to measurements, microbiological testing and for technical assistance.

References

1. A. Homayouni, M. R. Ehsani, A. Azizi, M. S. Yarmand and S. H. Razavi, Effect of lecithin and calcium chloride solution on the microencapsulation process yield of calcium alginate beads, Iran. Polym. J., 2007, 16(9), 597.
2. R. Agrawal, Probiotics: an emerging food supplement with health benefits, Food Biotechnol., 2005, 19(3), 227–246.
3. I. Jankovic, W. Sybesma, P. Phothirath, E. Ananta and A. Mercenier, Application of probiotics in food products—challenges and new approaches, Curr. Opin. Biotechnol., 2010, 21(2), 175–181.
4. C. Malmo, A. La Storia and G. Mauriello, Microencapsulation of Lactobacillus reuteri DSM 17938 cells coated in alginate beads with chitosan by spray drying to use as a probiotic cell in a chocolate soufflé, Food Bioprocess Technol., 2013, 6(3), 795–805.
5. A. S. Akalın, S. Fenderya and N. Akbulut, Viability and activity of bifidobacteria in yoghurt containing fructooligosaccharide during refrigerated storage, Int. J. Food Sci. Technol., 2004, 39(6), 613–621.
6. R. P. Ross, C. Desmond, G. F. Fitzgerald and C. Stanton, Overcoming the technological hurdles in the development of probiotic foods, J. Appl. Microbiol., 2005, 98(6), 1410–1417.
7. A. K. Anal and H. Singh, Recent advances in microencapsulation of probiotics for industrial applications and targeted delivery, Trends Food Sci. Technol., 2007, 18(5), 240–251.
8. K. Kailasapathy, Microencapsulation of probiotic bacteria: technology and potential applications, Curr. Issues Intest. Microbiol., 2002, 3(2), 39–48.
9. T. Heidebach, P. Först and U. Kulozik, Influence of casein-based microencapsulation on freeze-drying and storage of probiotic cells, J. Food Eng., 2010, 98(3), 309–316.
10. A. Picot and C. Lacroix, Encapsulation of bifidobacteria in whey protein-based microcapsules and survival in simulated gastrointestinal conditions and in yoghurt, Int. Dairy J., 2004, 14(6), 505–515.
11. Y. D. Livney, Milk proteins as vehicles for bioactives, Curr. Opin. Colloid Interface Sci., 2010, 15(1), 73–83.
12. M. H. Abd El-Salam and S. El-Shibiny, Formation and potential uses of milk proteins as nano delivery vehicles for nutraceuticals: a review, Int. J. Dairy Technol., 2012, 65(1), 13–21.
13. S. B. Doherty, M. A. Auty, C. Stanton, R. P. Ross, G. F. Fitzgerald and A. Brodkorb, Survival of entrapped Lactobacillus rhamnosus GG in whey protein micro-beads during simulated ex vivo gastro-intestinal transit, Int. Dairy J., 2012, 22(1), 31–43.
14. T. Heidebach, P. Först and U. Kulozik, Microencapsulation of probiotic cells by means of rennet-gelation of milk proteins, Food Hydrocolloids, 2009, 23(7), 1670–1677.
15. A. Kilara and D. Panyam, Peptides from milk proteins and their properties, Crit. Rev. Food Sci. Nutr., 2003, 43(6), 607–633.
16. H. Korhonen and A. Pihlanto, Food-derived bioactive peptides-opportunities for designing future foods, Curr. Pharm. Des., 2003, 9(16), 1297–1308.
17. N. Bansal, P. F. Fox and P. L. McSweeney, Factors affecting the retention of rennet in cheese curd, J. Agric. Food Chem., 2007, 55(22), 9219–9225.
18. J. Wang, H. Zhang, X. Chen, Y. Chen and Q. Bao, Selection of potential probiotic lactobacilli for cholesterol-lowering properties and their effect on cholesterol metabolism in rats fed a high-lipid diet, J. Dairy Sci., 2012, 95(4), 1645–1654.
19. C. R. F. Grosso and C. S. Fávaro-Trindade, Stability of free and immobilized Lactobacillus acidophilus and Bifidobacterium lactis in acidified milk and of immobilized B. lactis in yoghurt, Braz. J. Microbiol., 2004, 35(1–2), 151–156.
20. D. de Lara Pedroso, M. Thomazini, R. J. B. Heinemann and C. S. Favaro-Trindade, Protection of Bifidobacterium lactis and Lactobacillus acidophilus by microencapsulation using spray-chilling, Int. Dairy J., 2012, 26(2), 127–132.
21. K. M. Amine, C. P. Champagne, S. Salmieri, M. Britten, D. St-Gelais, P. Fustier and M. Lacroix, Effect of palmitoylated alginate microencapsulation on viability of Bifidobacterium longum during freeze-drying, LWT--Food Sci. Technol., 2014, 56(1), 111–117.
22. W. Krasaekoopt, B. Bhandari and H. Deeth, The influence of coating materials on some properties of alginate beads and survivability of microencapsulated probiotic bacteria, Int. Dairy J., 2004, 14(8), 737–743.
23. R. R. Mokarram, S. A. Mortazavi, M. H. Najafi and F. Shahidi, The influence of multi stage alginate coating on survivability of potential probiotic bacteria in simulated gastric and intestinal juice, Food Res. Int., 2009, 42(8), 1040–1045.
24. S. Mandal, A. K. Puniya and K. Singh, Effect of alginate concentrations on survival of microencapsulated Lactobacillus casei NCDC-298, Int. Dairy J., 2006, 16(10), 1190–1195.
25. P. Dolly, A. Anishaparvin, G. S. Joseph and C. Anandharamakrishnan, Microencapsulation of Lactobacillus plantarum (mtcc 5422) by spray-freeze-drying method and evaluation of survival in simulated gastrointestinal conditions, J. Microencapsulation, 2011, 28(6), 568–574.
26. R. Rajam, P. Karthik, S. Parthasarathi, G. S. Joseph and C. Anandharamakrishnan, Effect of whey protein–alginate wall systems on survival of microencapsulated Lactobacillus plantarum in simulated gastrointestinal conditions, J. Funct. Foods, 2012, 4(4), 891–898.
27. A. Sohail, M. S. Turner, A. Coombes, T. Bostrom and B. Bhandari, Survivability of probiotics encapsulated in alginate gel microbeads using a novel impinging aerosols method, Int. J. Food Microbiol., 2011, 145(1), 162–168.
28. L. E. Shi, Z. H. Li, Z. L. Zhang, T. T. Zhang, W. M. Yu, M. L. Zhou and Z. X. Tang, Encapsulation of Lactobacillus bulgaricus in carrageenan-locust bean gum coated milk microspheres with double layer structure, LWT--Food Sci. Technol., 2013, 54(1), 147–151.
29. J. Z. Lin, W. T. Yu, X. X. Xu, X. D. Liu and X. J. Ma, Preparation of calcium alginate gel beads by emulsion-internal gelation technology, J. Funct. Mater., 2007, 39, 1879–1882.
30. C. M. Silva, A. J. Ribeiro, I. V. Figueiredo, A. R. Gonçalves and F. Veiga, Alginate microspheres prepared by internal gelation: Development and effect on insulin stability, Int. J. Pharm., 2006, 311(1), 1–10.
31. L. W. Chan, H. Y. Lee and P. W. Heng, Mechanisms of external and internal gelation and their impact on the functions of alginate as a coat and delivery system, Carbohydr. Polym., 2006, 63(2), 176–187.
32. A. Nag, K. S. Han and H. Singh, Microencapsulation of probiotic bacteria using pH-induced gelation of sodium caseinate and gellan gum, Int. Dairy J., 2011, 21(4), 247–253.
33. L. D. McMaster and S. A. Kokott, Micro-encapsulation of Bifidobacterium lactis for incorporation into soft foods, World J. Microbiol. Biotechnol., 2005, 21(5), 723–728.
34. M. C. Otero, M. C. Espeche and M. E. Nader-Macías, Optimization of the freeze-drying media and survival throughout storage of freeze-dried Lactobacillus gasseri and Lactobacillus delbrueckii subsp. delbrueckii for veterinarian probiotic applications, Process Biochem., 2007, 42(10), 1406–1411.
35. M. Chávarri, I. Marañón, R. Ares, F. C. Ibáñez, F. Marzo and M. del Carmen Villarán, Microencapsulation of a probiotic and prebiotic in alginate-chitosan capsules improves survival in simulated gastro-intestinal conditions, Int. J. Food Microbiol., 2010, 142(1), 185–189.
36. R. Crittenden, R. Weerakkody, L. Sanguansri and M. Augustin, Synbiotic microcapsules that enhance microbial viability during nonrefrigerated storage and gastrointestinal transit, Appl. Environ. Microbiol., 2006, 72(3), 2280–2282.
37. B. De Giulio, P. Orlando, G. Barba, R. Coppola, M. De Rosa, A. Sada and F. Nazzaro, Use of alginate and cryo-protective sugars to improve the viability of lactic acid bacteria after freezing and freeze-drying, World J. Microbiol. Biotechnol., 2005, 21(5), 739–746.
38. A. Homayouni, A. Azizi, M. R. Ehsani, M. S. Yarmand and S. H. Razavi, Effect of microencapsulation and resistant starch on the probiotic survival and sensory properties of synbiotic ice cream, Food Chem., 2008, 111(1), 50–55.
39. N. P. Shah and R. R. Ravula, Microencapsulation of probiotic bacteria and their survival in frozen fermented dairy desserts, Aust. J. Dairy Technol., 2000, 55(3), 139–144.
40. T. Y. Sheu and R. T. Marshall, Microentrapment of lactobacilli in calcium alginate gels, J. Food Sci., 1993, 58(3), 557–561.
41. A. S. Carvalho, J. Silva, P. Ho, P. Teixeira, F. X. Malcata and P. Gibbs, Effect of various growth media upon survival during storage of freeze-dried Enterococcus faecalis and Enterococcus durans, J. Appl. Microbiol., 2003, 94(6), 947–952.
42. M. Saarela, I. Virkajärvi, H. L. Alakomi, T. Mattila-Sandholm, A. Vaari, T. Suomalainen and J. Mättö, Influence of fermentation time, cryoprotectant and neutralization of cell concentrate on freeze-drying survival, storage stability, and acid and bile exposure of Bifidobacterium animalis ssp. lactis cells produced without milk-based ingredients, J. Appl. Microbiol., 2005, 99(6), 1330–1339.
43. B. M. Corcoran, R. P. Ross, G. F. Fitzgerald and C. Stanton, Comparative survival of probiotic lactobacilli spray-dried in the presence of prebiotic substances, J. Appl. Microbiol., 2004, 96(5), 1024–1039.
44. N. Fu and X. D. Chen, Towards a maximal cell survival in convective thermal drying processes, Food Res. Int., 2011, 44(5), 1127–1149.
45. J. E. Kinsella and C. V. Morr, Milk proteins: physicochemical and functional properties, Crit. Rev. Food Sci. Nutr., 1984, 21(3), 197–262.
46. P. Smrdel, M. Bogataj, A. Zega, O. Planinšek and A. Mrhar, Shape optimization and characterization of polysaccharide beads prepared by ionotropic gelation, J. Microencapsulation, 2008, 25(2), 90–105.
47. P. Muthukumarasamy, P. Allan-Wojtas and R. A. Holley, Stability of Lactobacillus reuteri in different types of microcapsules, J. Food Sci., 2006, 71(1), M20–M24.
48. T. Y. Sheu, R. T. Marshall and H. Heymann, Improving survival of culture bacteria in frozen desserts by microentrapment, J. Dairy Sci., 1993, 76(7), 1902–1907.
49. L. Sabikhi, R. Babu, D. K. Thompkinson and S. Kapila, Resistance of microencapsulated Lactobacillus acidophilus LA1 to processing treatments and simulated gut conditions, Food Bioprocess Technol., 2010, 3(4), 586–593.
50. L. Chen and M. Subirade, Alginate–whey protein granular microspheres as oral delivery vehicles for bioactive compounds, Biomaterials, 2006, 27(26), 4646–4654.
51. D. Guérin, J. C. Vuillemard and M. Subirade, Protection of bifidobacteria encapsulated in polysaccharide-protein gel beads against gastric juice and bile, J. Food Prot., 2003, 66(11), 2076–2084.
52. G. K. Gbassi, T. Vandamme, S. Ennahar and E. Marchioni, Microencapsulation of Lactobacillus plantarum spp in an alginate matrix coated with whey proteins, Int. J. Food Microbiol., 2009, 129(1), 103–105.
53. S. Sathyabama and R. Vijayabharathi, Co-encapsulation of probiotics with prebiotics on alginate matrix and its effect on viability in simulated gastric environment, LWT--Food Sci. Technol., 2014, 57(1), 419–425.
54. R. Su, X. L. Zhu, D. D. Fan, Y. Mi, C. Y. Yang and X. Jia, Encapsulation of probiotic Bifidobacterium longum BIOMA 5920 with alginate–human-like collagen and evaluation of survival in simulated gastrointestinal conditions, Int. J. Biol. Macromol., 2011, 49(5), 979–984.
55. G. B. Brinques and M. A. Z. Ayub, Effect of microencapsulation on survival of Lactobacillus plantarum in simulated gastrointestinal conditions, refrigeration, and yogurt, J. Food Eng., 2011, 103(2), 123–128.
56. J. M. T. Hamilton-Miller, S. Shah and J. T. Winkler, Public health issues arising from microbiological and labelling quality of foods and supplements containing probiotic microorganisms, Public Health Nutr., 1999, 2(2), 223–229.
57. G. Reid, M. E. Sanders, H. R. Gaskins, G. R. Gibson, A. Mercenier, R. Rastall and T. R. Klaenhammer, New scientific paradigms for probiotics and prebiotics, J. Clin. Gastroenterol., 2003, 37(2), 105–118.
58. C. A. Morgan, N. Herman, P. A. White and G. Vesey, Preservation of micro-organisms by drying; a review, J. Microbiol. Methods, 2006, 66(2), 183–193.
59. F. Abe, H. Miyauchi, A. Uchijima, T. Yaeshima and K. Iwatsuki, Stability of bifidobacteria in powdered formula, Int. J. Food Sci. Technol., 2009, 44(4), 718–724.
60. G. M. Maciel, K. S. Chaves, C. R. F. Grosso and M. L. Gigante, Microencapsulation of Lactobacillus acidophilus La-5 by spray-drying using sweet whey and skim milk as encapsulating materials, J. Dairy Sci., 2014, 97(4), 1991–1998.
61. P. Teixeira, H. Castro and R. Kirby, Evidence of membrane lipid oxidation of spray-dried Lactobacillus bulgaricus during storage, Lett. Appl. Microbiol., 1996, 22(1), 34–38.
62. B. E. Chávez and A. M. Ledeboer, Drying of probiotics: optimization of formulation and process to enhance storage survival, Drying Technol., 2007, 25(7–8), 1193–1201.
63. D. Rodrigues, S. Sousa, T. Rocha-Santos, J. P. Silva, J. S. Lobo, P. Costa and A. C. Freitas, Influence of L-cysteine, oxygen and relative humidity upon survival throughout storage of probiotic bacteria in whey protein-based microcapsules, Int. Dairy J., 2011, 21(11), 869–876.

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