Efficacy of whey protein gel networks as potential viability-enhancing scaffolds for cell immobilization of Lactobacillus rhamnosus GG

Journal of Microbiological Methods 80 (2010) 231–241

Contents lists available at ScienceDirect

Journal of Microbiological Methods j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / j m i c m e t h

Efficacy of whey protein gel networks as potential viability-enhancing scaffolds for cell immobilization of Lactobacillus rhamnosus GG S.B. Doherty a,b, V.L. Gee a, R.P. Ross a,c, C. Stanton a,c, G.F. Fitzgerald b,c, A. Brodkorb a,⁎ a b c

Teagasc, Moorepark Food Research Centre, Fermoy, Co. Cork, Ireland Department of Microbiology, University College Cork, Cork, Ireland Alimentary Pharmabiotic Centre, Cork , Ireland

a r t i c l e

i n f o

Article history: Received 31 July 2009 Received in revised form 4 December 2009 Accepted 15 December 2009 Available online 4 January 2010 Keywords: Cell immobilization Lactobacillus rhamnosus GG Viability Whey proteins Encapsulation Probiotic bacteria

a b s t r a c t This study investigated cell immobilization of Lactobacillus rhamnosus GG in three separate protein products: native, denatured and hydrolysed whey protein isolate (WPI). Treatments were assessed for their ability to enhance probiotic survival during storage, heat stress and ex vivo gastric incubation. Spatial distribution of probiotic cells within immobilized treatments was evaluated by atomic force and confocal scanning laser microscopy, while cell viability was enumerated by plate count and flow cytometry (FACS). Microscopic analysis of denatured treatments revealed an oasis of immobilized cells, phase-separated from the surrounding protein matrix; an environmental characteristic analogous to hydrolysed networks. Cell immobilization in hydrolysed and denatured WPI enhanced survival by 6.1 ± 0.1 and 5.8 ± 0.1 log10 cycles, respectively, following 14 day storage at 37 °C and both treatments generated thermal protection at 57 °C (7.3 ± 0.1 and 6.5 ± 0.1 log10 cfu/ml). Furthermore, denatured WPI enhanced probiotic protection (8.9 ± 0.2 log10 cfu/ml) following 3 h gastric incubation at 37 °C. In conclusion, hydrolysed or denatured WPI were the most suitable matrices for cell immobilization, while native protein provided the weakest safeguard against thermal and acid stress, thus making it possible to envision whey protein gel networks as protective substrates for cell immobilization applications. © 2009 Elsevier B.V. All rights reserved.

1. Introduction The recent escalation in consumer health consciousness has strengthened the global appetite for preventative rather than curative approaches toward diseases, ultimately generating significant growth of functional foods on a global basis (Siro et al., 2008). As more scientific evidence accrues, the dairy industry in particular has been quick to recognise the market potential resonating from the health benefits associated with probiotic bacteria, defined as ‘live microorganisms, which when administered in adequate amounts confer a health benefit on the host’(FAO/WHO, 2001). These microorganisms enhance the population of beneficial bacteria in the gut, suppress pathogens and build-up resistance against intestinal diseases, in addition to providing well-documented clinical benefits (Borchers et al., 2009, Collado et al., 2009, Vasiljevic and Shah, 2008). Lactobacillus rhamnosus GG, a humanderived strain with commercial significance, has recognised benefits in the treatment of diarrhoea (Isolauri et al., 1991) and atopic eczema (Isolauri et al., 1999). Lactobacillus and Bifidobacterium have been accorded the generally recognised as safe (GRAS) status (Salminen et al., 1998) due to their long history of safe use in food; thus, probiotics

⁎ Corresponding author. Tel.: +353 25 42222; fax: +353 25 42340. E-mail address: [email protected] (A. Brodkorb). 0167-7012/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.mimet.2009.12.009

have been highlighted as suitable candidates for the development of functional foods. From a processing point of view, integration of probiotic bacteria into dairy-based food systems presents a pivotal challenge to food manufacturers (Ross et al., 2005), given that optimal probiotic functionality is only accomplished through delivery of sufficient cell concentrations, in an active form, to the target site in the host. The food industry in general has adopted the recommended minimum level of 106 cfu/ml at the time of consumption in order to achieve bacterial populations that are cost effective and technologically manageable (Boylston et al., 2004, Kailasapathy and Chin, 2000). Thus, the prerequisites required for preserving probiotic bioavailability involve the ability to withstand the fluctuating temperatures of dairy processing (Knorr, 1998, Saxelin et al., 1999) and subsequent storage conditions, in addition to the host's natural defense barriers against ingested bacteria, which is primarily high stomach acidity and bile salts (Gardiner et al., 2000). Hence, great demands are imposed upon probiotic product quality, which ultimately catalyzed strong research activity within the dairy industry, in an attempt to improve technological properties of probiotic bacteria for application into novel and non-traditional production processes. From a product standpoint, appropriate selection of the food delivery matrix is of paramount importance during the development of probiotic functional foods. Studies in vitro and in vivo have shown

232

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

that the nature of the food harboring probiotic bacteria has an effect on subsequent viability following consumption (Gardiner et al., 1999). Among the various milk products, yoghurt is regarded as the quintessential vector for probiotic delivery to the consumer. However, Gardiner et al. (1999) revealed that the enhanced buffering capacity and dense matrix provided by Cheddar cheese were the key factors conducive to elevated probiotic survival in gastric juice compared to yoghurt. In this regard, cell immobilization technology may provide the necessary safeguard for probiotics by generating a matrix environment with the aid of a suitable immobilization agent. Recent advances have broadened the use of cell immobilization in the food and biomedical fields. The entrapment of cells within a foodgrade polymeric matrix is highlighted (Heidebach et al., 2009a,b; Nedovic and Willaert, 2005) as the preferred immobilization technique for food applications with the purpose of providing cell protection from hostile environments. For successful immobilization and cultivation of cells, the polymer material must be conducive to cell viability and function (biocompatible) within specific food systems. Whey proteins have interesting physicochemical properties encompassing the ability to form gels, which can be generated by heating the protein solution with subsequent cooling and acidification (Holt, 2000). The net charge of the whey protein molecule is at a minimum at the isoelectric point (pH 5.2), which induces gelation by reducing the electrostatic repulsion between proteins. This mechanism may provide the driving force for aggregation and simultaneous cell-entrapment within whey protein gel systems. In this regard, WPI may be judiciously exploited to formulate biocompatible pH-sensitive matrices for delivery of probiotic bacteria. Kos et al. (2000) investigated the influence of whey proteins on in vitro survival of L. acidophilus M92 cells in simulated gastrointestinal (GI) conditions and demonstrated the efficacy of whey protein as a protector, in the preparation of L. acidophilus M92 for probiotic use. Additionally, dispersion of bifidobacteria in heattreated WPI followed by spray-drying may qualify as a successful GI probiotic delivery technique (Picot and Lacroix, 2004), while immobilization of probiotic bacteria in large heat-denatured WPI beads provided cell protection against acidic conditions (Ainsley Reid et al., 2005, Guerin et al., 2003). Hence, this research advocates WPI as a possible protective ingredient for probiotics. In addition to acidinduced gelation, limited hydrolysis of whey protein solutions provides an alternative for the development of protein microstructures under mild conditions (Doucet et al., 2001). Although research for this probiotic application is limited, whey proteins have the ability to aggregate under different conditions of pH, ionic strength and temperature to form potential scaffolds for enhanced probiotic stability. Thus, the aim of this study was to investigate the potential protective role of whey proteins as matrices for cell immobilization of L. rhamnosus GG during storage, thermal treatment and exposure to porcine gastric contents, in addition to visualizing the matrix interaction operating within optimum treatments. 2. Materials and methods 2.1. Immobilization materials BiPro, a commercial whey protein isolate (WPI) obtained from Davisco Foods International Inc., (MN, U.S.A.) containing 98% (w/w) protein, was utilized for production of cell immobilization matrices. βlactoglobulin (β-lg) and α-lactalbumin (α-la) content were analyzed by reversed-phase HPLC and estimated at 82% and 16%, respectively. Flavourzyme, a protease–peptidase complex produced by submerged fermentation of a selected strain of Aspergillus oryzae, was purchased from Novo Nordisk A/S, Bagsvaerd, Denmark for enzymatic hydrolysis. Highly purified water (MilliQ, BioSciences, Cork, Ireland) was used in all cases for dispersion of samples, culture media and buffer solutions.

2.2. Bacterial strain and culture conditions The probiotic strain Lactobacillus rhamnosus GG (ATCC 53103, L. rhamnosus GG, Valio Ltd., Helsinki, Finland), was sourced from the Moorepark culture collection. Harvested cells were stored as stock solutions in de Man Rogosa Sharpe (MRS) broth (Oxoid Ltd., Hampshire, U.K.) (de Man et al., 1960) containing 50% (v/v) aqueous glycerol at − 20 °C. All tests were performed using subcultures from the same frozen stock, which was routinely checked for purity. The frozen culture was grown in MRS broth at 37 °C under anaerobic conditions, achieved by activation of Anaerocult gas packs (Merck, Darmstadt, Germany). Bacteria for immobilization experiments were propagated from 1% (v/v) inoculations for 18 h at 37 °C. The stationary phase cells (109 cfu/ml) were harvested by centrifugation at 5200 ×g for 10 min at 4 °C (Sorvall, RC-5C Plus, Sorvall Products, Herts, UK), washed and resuspended in a dilute citrate-phosphate buffer (pH 5.2) (Sigma Chemical CO, St Louis, U.S.A.) to obtain a concentrated cell suspension. Ultimately, the cell slurry was either employed within the immobilization process, or utilized (as a control) in a free-cell condition. 2.3. Heat treatment of protein dispersions A protein solution (5% w/v on protein basis) was prepared by dispersion of WPI in sterile water and stirring for 1 h at room temperature; the solution was then stored overnight at 4 °C, to permit complete protein hydration. The solution was subsequently adjusted to pH 7, using NaOH, and filtered through a 0.45 µm Millex, HVLP filter (Millipore Corp., MA, USA) to remove traces of undissolved material. This untreated WPI solution was referred to as native WPI throughout the manuscript. A portion of this material was heated under agitation (75 rpm), in a water bath at 78 °C for 30 min. The suspension of reactive WPI aggregates was subsequently cooled on ice, stored at 4 °C and utilized within 2 h and referred to as heat-treated/denatured WPI thereafter. 2.4. Whey protein hydrolysate and physicochemical characterization An untreated WPI solution (10%, w/v; pH 7) was the preferred substrate for enzymatic hydrolysis, using the protease–peptidase complex, Flavourzyme, which was standardized in Leucine Amino Peptidase Units (LAPU) by the manufacturer. Hydrolysis was performed batchwise in a sterile 4 l thermostated reaction vessel (B.Braun Biotech., Melsungen, Germany). The protein solution was pre-heated (50 °C) and a measured amount of Flavourzyme (1000 LAPU) was pre-suspended in WPI solution and subsequently adjusted to the hydrolysis conditions (pH 7; 50 °C). Hydrolysis was initiated by addition of the enzyme, in order to achieve an enzyme/substrate ratio (E/S) of 1/100 on the basis of total protein content in WPI. The pH was automatically maintained at 7.0 by the continuous addition of 4 N NaOH (Metrohm Ltd., Herisau, Switzerland). Samples were withdrawn from the batch solution between T0 and T4 h at 30 min intervals and the volume of NaOH required to sustain a constant pH value was recorded, thereby providing an approximate value for degree of hydrolysis (DH) (Adler-Nissen, 1979). The reaction was terminated after 4 h by heat treatment at 80 °C for 20 min, to facilitate permanent inactivation of the enzymatic reaction. This was followed by cooling, freeze-drying and storage of the hydrolysate at 4 °C. Size exclusion chromatography was performed using an automated 2695 Waters™ HPLC system (Millipore, Middlesex, UK) equipped with a TSK G2000 SW column (600 × 7.5 mm; Tosoh Corporation, Tokyo, Japan). Samples were filtered (0.2 µm) and eluted at a flow rate of 1 ml/min using 30% acetonitrile containing 0.1% (v/v) TFA. A molecular weight calibration curve was prepared from the average retention time of standard proteins, peptides and amino acids. Following this, hydrolysate samples were subsequently utilized as a cell immobilization milieu. The degree of hydrolysis was assayed directly by quantification of cleaved peptide bonds as assessed by the ο-phthaldialdehyde (OPA)

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

spectrophotometric assay, which involved using N-acetyl-L-cysteine (NAC) as the thiol reagent (Garcia Alvarez-Coque et al., 1989a,b). The OPA-NAC reagent, was prepared as previously described (Spellman et al., 2003) and the reagent was covered and stirred overnight at room temperature. To assay proteolysis, 100 µl of each hydrolysate sample was added to an equal volume of 24% (w/v) trichloroacetic acid (TCA) and allowed to stand at ambient temperature for 10 min. A small aliquot (20 µl) of the supernatant, obtained after centrifugation (14,000 × g for 3 min), was added directly to 3 ml of OPA reagent. The solution was vortexed, incubated at ambient temperature for 30 min and the reaction product was subsequently detected by absorbance at 340 nm. Reversed-phase HPLC confirmed the protein content of the substrate (82% native β-lg and 16% α-la); thus the average molecular mass of protein in WPI was calculated and inserted into the relevant equation (Spellman, et al., 2003), yielding the adjusted formula: DHð%Þ =

Δ Abs: × 1:906 × D C

where Δ Abs. is the absorbance of the test sample at 340 nm— absorbance of unhydrolysed sample at 340 nm, D the dilution factor and C the protein concentration (g/l). 2.5. Immobilization The washed cell suspension of L. rhamnosus GG was blended aseptically with native, heat-treated or hydrolysed whey protein solutions, at ambient temperature with a final protein concentration of 2% (w/v) and a probiotic population corresponding to the stationary phase viable cells at 109 cfu/ml. For comparative purposes, the remaining cell concentrate was diluted accordingly, to generate the free-cell reference. Cold gelation of heat-denatured proteins was induced by slow acidification (100 mM HCl) to pH 5.2; pH adjustment below 5.5 instantly produced a gel, hence, stirring was required to prevent nonhomogenous protein aggregation. Numbers of viable lactobacilli were determined (day 0) prior to initiation of storage and thermal tolerance tests. Samples were agitated gently (200 rpm) for the initial 24 h storage; this action was subsequently halted thereafter. Fig. 1 illustrates the overall sampling and analysis procedure employed during the study.

233

2.6. Determination of probiotic retention and viability in WPI treatments and yoghurt networks The viability and sensitivity of the immobilized bacteria were evaluated within the various protein environments, by storing all treatments at 4, 25 and 37 °C for 2 weeks. The enumeration of viable cells was conducted on days 0, 1, 4, 8 and 14 of storage. Entrapped bacteria were released from their respective protein network using a homogenization procedure (data not shown) to ensure complete liberation of bacteria from the respective protein systems. Control L. rhamnosus GG cells were treated similarly, to maintain consistent treatment conditions. Homogenates were spread-plated on MRS agar and colonies were subsequently counted after 48 h incubation at 37 °C under anaerobic conditions. Plates containing 30–300 colonies were enumerated and recorded as cfu/ml of protein material. Furthermore, cell viability within immobilized and free-cell treatments was evaluated following storage at 4 and 10 °C for 14 days in commercial natural low-fat yoghurt (Irish yoghurts, Cork, Ireland). Fresh treatments were prepared (Section 2.5) and aseptically blended with aliquots of yoghurt (pH 3.98 ± 0.02) to achieve a final probiotic population of 108 cfu/ml of yoghurt. After 14-day storage, treatments were homogenised and the probiotic population was enumerated on Lactobacillus casei (LC) medium, pH 5.1, supplemented with a membrane-filtered sterile solution of 10% (w/v) D (−) ribose (1% final concentration) and 0.2% (v/v) bromocresol green solution (both Sigma Aldrich, Dorset, UK; 0.04% (v/v) final concentration) under anaerobic conditions at 27 °C for 3–4 days (Van de Casteele et al., 2006). Prior to testing, no bacterial growth was detected from blank yoghurt samples (no probiotic), which demonstrated that this medium was selective for L. rhamnosus GG during this test. Streptococcus thermophilus and L. delbrueckii subsp. bulgaricus were also enumerated as previously described (Van de Casteele, et al., 2006) using M17 medium (Difco Laboratories, MI, USA), supplemented with 1% (w/v) lactose (Oxoid) and MRS medium (pH 5.2), respectively. 2.7. Heat challenge assay The heat tolerance of free L. rhamnosus GG cells was determined using stationary phase cultures, harvested and resuspended as

Fig. 1. Procedure for cell immobilization in whey protein products and subsequent treatment analysis.

234

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

previously described. The thermal protection extended to L. rhamnosus GG through immobilization was ascertained directly after sample preparation. Immobilized treatments and free cells were vortexed and aliquots were heated for 5 min at 52, 55, 56 and 57 °C in a gradient PCR thermal block. All samples were immediately cooled and data relating to the thermal protection of L. rhamnosus GG by immobilization in heattreated, native and hydrolysed WPI are all based on duplicate assays. 2.8. Survival of immobilized cells in porcine gastric contents The two best performing immobilization treatments were selected from the thermal tolerance experiment and fresh samples were subsequently evaluated under acidic conditions. Gastric contents from three porcine stomachs were collected and pooled within 2 h of slaughter. The starved animals (12 h prior slaughter) were not prescribed any medicated feed prior to/at the time of collection, gastric juice was subject to centrifugation and filtration, and the final suspension was checked for sterility on brain heart infusion agar (Oxoid Ltd.). Preliminary tests confirmed the absence of indigenous gut microflora within gastric contents; thus selective media was not required. Samples of immobilized bacteria and free-cell references were transferred to aliquots of pre-warmed gastric juice (1:10 dilution; pH 2.5) and incubated at 37 °C for 3 h, while orbital agitation at 100 rpm in a controlled environment incubator was maintained. At appropriate time intervals, samples were withdrawn and L. rhamnosus GG was enumerated on MRS agar under anaerobic incubation at 37 °C for 48 h. The experiment was performed in duplicate and probiotic survival was expressed as a function of gastric incubation time.

integrating the dye concentrations previously optimized during FACS analysis. Samples were imaged as previously described (Auty et al., 2001), using a ×63 magnification objective with a numerical aperture of 1.4. Confocal illumination was provided by an argon laser (488 nm laser excitation) and red-green-blue images (24 bit), 512 × 512 pixels, were acquired using a zoom factor of 2.0, giving a final pixel resolution of 0.2 µm/pixel. Bright-field light microscopy measurements were also carried out using a BX51 light microscope (Olympus, Essex, UK). Samples were deposited on glass slides and analyzed immediately. 2.11. Statistical analysis All experiments were conducted in triplicate (unless stated otherwise) and were independently repeated in duplicate. The average values and the standard deviation (SD) were calculated and are expressed as the mean log count survival as a function of storage time or test temperature. Student's t-tests were performed on all results using Microsoft Excel, assuming two-tailed distribution and equal variance for all experimental data sets. Treatment means were considered significantly different at p ≤ 0.05 unless stated otherwise. 3. Results 3.1. Characterization of protein-probiotic networks by microscopy AFM analysis clearly illustrated clusters of Lactobacillus chains immobilized in heat-treated WPI matrices as illustrated in Fig. 2A

2.9. Probiotic detection by live/dead discrimination using flow cytometry In addition to plate counts, the viability of probiotic cells was assessed by flow cytometry (FACS), using the BD Cell Viability assay (BD Biosciences, Oxford, U.K.). Sample homogenates were diluted to a predetermined cell density and working solutions of the fluorescent stains, Thiazole Orange (TO) and Propidium Iodide (PI) were prepared on the analysis day (unpublished data). Preliminary trials verified the nonspecific binding of protein particles to fluorescent stains, which ultimately interfered with staining and detection of bacteria (Gunasekera et al., 2000). Thus, enzymatic treatment using Protease K was performed in association with fluorescent staining. Data acquisition was performed on a BD FACS Canto II flow cytometer (BD Biosciences, Oxford, U.K.), equipped with 488 nm laser excitation and BD FACS Diva software using a side scatter (SSC) threshold. A sample spiked with a known concentration of live and killed L. rhamnosus GG was prepared to confirm that stained live, injured and dead bacterial populations were sufficiently resolved in gates A3, A2 and A1, respectively. Populations were also assessed by fluorescence microscopy to confirm that the target organisms were stained. 2.10. Microscopy Atomic Force Microscopy (AFM) images were obtained using Asylum Research MFP-3D-AFM (Asylum Research UK Ltd. Oxford, UK) in AC-mode. Prior to imaging, all samples were diluted (×100) in MilliQ H2O and 10 µl aliquots were deposited onto freshly cleaved mica surfaces and subsequently dried in a desiccator. An aluminium reflex coated cantilever with a tetrahedral tip (AC 240), spring constant of 1.8 N/m (Olympus Optical Co. Ltd, Tokyo Japan), working frequency of 50–90 kHz, and scan rate at 1 Hz was used for air-dried samples. The radius of curvature of the tetrahedral tip was 10 (±3) nm. Additional microscopy work was performed using a Leica TCS SP5 confocal scanning laser microscope (CSLM) (Leica Microsystems, Wetzler, Germany). Immobilized treatments were stained using the BD Cell Viability kit (BD Biosciences, San Jose, California), by

Fig. 2. A and B: AFM images illustrating the surface topography of L. rhamnosus GG embedded within heat-treated whey protein (A; 5 µm range) and the presence of distinct regions of bacteria (three cells labeled 1, 2 and 3) immobilized in the protein matrix (B; 1 µm range).

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

using AC-mode height images. Various structural features were identified (Fig. 2B) along the contour of this polymeric protein matrix involving the i) visualization of individual proteins and ii) existence of a micro-phase separation between probiotic cells (labeled 1, 2 and 3 in Fig. 2B) and the surrounding protein network. Further AFM probing of heat-treated samples illustrated that cell surfaces remained spatially uniform and smooth following thermal treatment at 57 °C for 5 min, a surface topography identical to that obtained with hydrolysed treatments under the same experimental condition (Fig. 3A). Importantly, a host of structural changes occurred in free-cell samples as a result of thermal stress at 57 °C, including loss of membrane integrity (Fig. 3B) and significant cell collapse (Fig. 3D), relative to intact healthy bacteria (Fig. 3A and C). Metabolically active cells express a height characteristic b350 nm (Fig. 3C), while injured cells illustrate a depressed surface topography b150 nm (Fig. 3D), which represents a dramatic change in height as a result of cellular injury. The rapid transformation of the cellular dimensions of non-immobilized L. rhamnosus GG illustrated in Fig. 3 represents the cratered product identified in four respective samples. Light and confocal microscopy illustrated an even distribution of bacteria in the serum phase of native WPI with no evidence of whey protein precipitation and bacteria were also found uniformly distributed within the free-cell sample. In contrast to this, bacteria appeared to be aggregated within irregular-shaped gel particles in heat-treated WPI (approx 95% denatured protein) and light microscope images (Fig. 4A and B) clearly illustrate the microstructure at the periphery and the core of the gelled network, respectively. Fig. 4C shows aggregate formation within the hydrolysed system and revealed probiotic enrichment within the aggregated phase and on the aggregate/serum interface (arrows on Fig. 4C). Furthermore, CSLM analysis coupled with a LIVE/DEAD staining technique highlighted the homogenous distribution of viable probiotic cells (Fig. 4D; green rods) immobilized in dense regions of the heat-treated protein matrix (red background). However, the background fluorescence of Fig. 4D, attributable to the homogenous staining of WPI with TO, was beneficial

235

during the present study since it promoted clear observation of immobilized cells within the WPI network. With this said, cellimmobilized matrices of heat-treated WPI (arrows in Fig. 4E) could be differentiated from the continuous protein network of natural yoghurt (dense red background) following 14-day storage of the probiotic treatments at 10 °C due to the high green fluorescent signal emanating from healthy, metabolizing L. rhamnosus GG. 3.2. Physicochemical characterization of whey protein hydrolysate In the present study, the protein hydrolysate was characterized using the OPA method (Spellman, et al., 2003), which established a degree of hydrolysis (DH) of 6.5% ± 0.4%. Size exclusion chromatography was also performed (Table 1) and a predominant exo-peptidase activity was indicated by the production of a peptide fraction b2 kDa coupled with the enhanced liberation of free amino acids (8.19 µmol/ml) after 4 h hydrolysis (data not shown). Analysis also revealed significant enzymatic digestion of the principle whey protein (β-lg) as a function of hydrolysis time; however, a significant proportion of the intact monomeric form continued to prevail at the end of hydrolysis. 3.3. Influence of immobilization matrix on the survival of L. rhamnosus GG during storage When a cell extraction technique was applied in order to combine the release and enumeration of viable cells from protein matrices, no significant reduction in viable probiotic numbers was recorded (p N 0.05), which has previously been reported for lengthy extraction procedures for immobilized cells (Sun and Griffiths, 2000). Fig. 5A and B illustrate the survival of immobilized L. rhamnosus GG in comparison to the relevant reference (free cells in buffer) at 25 and 37 °C, respectively, during 14-day storage. Cell immobilization in native, heat-treated and hydrolysed WPI influenced probiotic survival, albeit to various degrees, generally improving cell viability. Although free cells were not appreciably affected after 2 weeks storage at 4 °C (0.24 ± 0.03 log10

Fig. 3. A and B illustrate AFM images of intact and compromised cell membranes of L. rhamnosus GG after heat treatment at 57 °C in the presence of hydrolysed WPI and for nonimmobilized (free) cells, respectively. C and D represent the height profiles for metabolically active and injured cells, respectively and the red lines illustrate the calculated height measurements.

236

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

Fig. 4. A and B are light microscope images illustrating cell immobilization of L. rhamnosus GG at the periphery and core of a heat-treated WPI network, respectively, and arrows in C indicate regions of high cell density in the hydrolysed WPI system. Confocal laser scanning microscopy (CSLM) clearly exhibits cell immobilization in a denatured WPI matrix (D). Background red fluorescence represents the protein matrix; live bacteria are green; dead bacteria are red (Bar = 25 µm). Cell immobilization within heat-treated WPI matrices (arrows in E) is clearly identified within a dense continuous gel network of natural yoghurt (Bar = 50 µm). Large green dots represent fat globules in the yoghurt system.

cycle reduction), immobilization significantly (p b 0.05) enhanced cell viability by 0.18 and 0.2210 log cycles for heat-treated and hydrolysed treatments, respectively (data not shown). Cell immobilization in

hydrolysed WPI continued to harbor high levels of viable cells after 14 days at 25 °C, expressing approx. 200,000-fold enhanced probiotic viability; however, the survival of free cells declined significantly

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241 Table 1 Size Exclusion Chromatography data for whey protein hydrolysate. Molecular size (kDa)

Retention time (min)

% Area

N 20 20–10 10–5 5–2 2–1 1–0.5 b 0.5

11.23 11.86 13.31 15.46 16.21 16.75 19.68

45.81 7.35 7.65 8.59 7.98 3.78 18.83

(p b 0.001) after 24 h, resulting in a 5.37 ± 0.12 log10 cycle reduction following 14-day storage. Immobilization in heat-treated and hydrolysed WPI continued to show a consistent enhancement of cell viability during storage at 37 °C; however, these improvements were insufficient to prevent significant reductions in cell populations after 14-day storage. While the highest cell survival (9.06 ± 0.04 log10 cycles) occurred in hydrolysed treatments during refrigerated storage, the most dramatic viability augmentation (6.13 ± 0.01 log10 cycles) was reported at 37 °C for cells immobilized in heat-treated WPI. Once again, the sensitivity of L. rhamnosus GG was revealed when free cells, devoid of an immobilization matrix, expressed significant viability losses of 6.63 ± 0.08 log10 cycles after only 24 h agitated storage at 37 °C. After

237

incubation for 4 days at 37 °C, viability losses expressed by native and heat-treated samples (average cell loss approx. 2.37 log10 cycles) were reminiscent of mortality rates observed at 25 °C; however, hydrolysed treatments reported an accelerated cell loss of 2.35 ± 0.13 log10 cycles. Cell immobilization in heat-treated and hydrolysed WPI resulted in good strain persistence throughout storage at 37 °C, with viable cells maintained at 106 cfu/ml after 8 days; however, their continued existence following 14-day storage was significantly different (0.29 log10 cycles; p b 0.001). Moreover, immobilization in native WPI continued to afford the least protection during storage at 37 °C, yielding a viability reduction of 4.6 ± 0.2 log10 cycles. 3.4. Thermal tolerance and product application of immobilized probiotic lactobacilli The effect of immobilization on the survival of L. rhamnosus GG after i) 5 min exposure to elevated temperatures (52, 55, 56 and 57 °C) and ii) 14-day storage in natural yoghurt at 10 °C was investigated. The data presented in Fig. 6 are based on probiotic survival at each test condition. All treatments heat-stressed at 52 °C expressed high heat tolerance (9.3 ± 0.1 log10 cfu/ml) and as cell viability continued to be challenged at 55 °C, the free-cell reference experienced dramatic cell loss of 3.5 log10 cycles, enhancing mortality rates by approx. 1000-fold compared to cell loss at 52 °C. Cell immobilization in hydrolysed or heat-treated WPI provided the greatest safeguards against heat stress at 56 °C; however, significant cell loss was the consequence of a minor temperature rise to 57 °C (p b 0.001), increasing mortality rates by 1.5 ± 0.1 and 1.9 ± 0.1 log10 cfu/ml, respectively. In spite of this, hydrolysed and denatured treatments continued to generate the highest survival rates at 57 °C; enhancing cell viability by 7.3 ± 0.1 and 6.5 ± 0.1 log10 cfu/ml, respectively since the free cells were not detected at this temperature. Furthermore, native treatments failed to prevent dramatic cell loss at 57 °C resulting in final cell counts of 1.55 × 103 cfu/ml. Cell viability in the free-cell reference was not appreciably affected by storage in natural yoghurt for 14-days at 10 °C (Fig. 6); however, statistical analysis revealed a significant difference (p b 0.01) between WPI treatments and the free-cell reference, which corresponded to flow cytometry data (Fig. 8A and C, respectively). 3.5. Effect of selected immobilization matrices on survival of L. rhamnosus GG in porcine gastric contents

Fig. 5. A and B. Log survival counts of immobilized L. rhamnosus GG in whey protein products at 25 °C (A) and 37 °C (B). Stationary phase cultures immobilized in heattreated (■), hydrolysed (●) and native WPI (▲) were stored for 14 days in addition to the free-cell reference (5) (mean ± SD, n = 3). Significant differences in cfu values between heat-treated and hydrolysed treatments are indicated in B (* p b 0.05; *** p b 0.001).

Having demonstrated during thermal tolerance studies that immobilization in hydrolysed and heat-treated WPI significantly enhanced survival of L. rhamnosus GG, the fate of these selected treatments was monitored ex vivo in porcine (as a surrogate for human) gastric juice. Fig. 7 illustrates probiotic survival of immobilized L. rhamnosus GG populations in porcine gastric juice at 37 °C as a function of incubation time. After 3 h incubation, hydrolysed and heat-treated treatments exhibited 3.2 ± 0.2 and 0.1 ± 0.1 log10 cfu/ml cell loss, respectively; however, free L. rhamnosus GG was undetectable after only 15 min exposure. In addition to the traditional plate count technique, FACS analysis also confirmed the dramatic decline in viable counts (Fig. 8B). Following 30 min gastric exposure, the relative decrease in probiotic viability was analogous for hydrolysed and heat-treated WPI (p N 0.05); however, after 120 min exposure cell viability within the former fell unexpectedly, corresponding to a 2.6 log10 cycle viability loss below that expressed by the latter treatment at the same time point. However, heat-treated WPI continued to bestow a controlled influence on probiotic survival following 3 h gastric incubation, representing 8.9 ± 0.2 log10 cfu/ml survival, while hydrolysed treatments expressed intermediate survival abilities (5.6 ± 0.1 log10 cfu/ml). It is noteworthy that both treatments exhibited similar buffering capacities during the 3 h incubation period.

238

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

Fig. 6. Thermal tolerance and product application of immobilized L. rhamnosus GG in whey protein products. Stationary phase cultures were immobilized and subject to heat shock at 52 °C ( ), 55 °C ( ), 56 °C ( ), 57 °C (■) for 5 min in a gradient PCR block. Fresh treatments were also incorporated into a natural yoghurt product ( ) and L. rhamnosus GG was selectively enumerated following 14-day storage at 10 °C (mean ± SD, n = 3). Statistical analysis of immobilized treatments was performed separately for each test temperature (p b 0.05). Columns with the same letters are not significantly different (p N 0.05).

3.6. An alternative approach for live/dead discrimination of probiotic cells The BD Cell Viability assay was adapted to differentiate live and dead bacteria based on plasma membrane permeability. Propidium Iodide (PI) is generally used as a cell death marker because it is excluded from intact plasma membranes and cell viability is estimated using Thiazole Orange (TO), since retention of the fluorescent probe indicates membrane integrity. The dual parameter dot plots of Fig. 8 illustrate the existence of probiotic subpopulations, which consist of dead cells that stained only with PI (gate A1) and live cells that stained with TO (gate A3); while gate A4 represented debris from sample preparation. In addition to plate enumeration, FACS demonstrated high cell viability in heat-treated and hydrolysed treatments after i) 24 h storage at 37 °C and ii) 14-day storage in yoghurt at 10 °C, since the entire treatment populations were encountered in gate A3 (Fig. 8A ). During yoghurt analysis, the freecell reference established a sub-population of injured cells following 14-day incubation (Fig. 8C; gate A2); a characteristic absent from heat-treated or hydrolysed treatments. Furthermore, complete loss of probiotic viability was validated following 4-day storage of free cells at 37 °C, producing a dot plot identical to that obtained for free

Fig. 7. Survival of free and immobilized L. rhamnosus GG in ex vivo porcine gastric juice (pH 2.5; 37 °C). Survival rates for heat-treated (■) and hydrolysed (●) WPI and free cells (▲) during 3 h incubation period (mean ± SD, n = 3). Significant differences in cfu values between hydrolysed and denatured treatments are denoted (** p b 0.01; *** p b 0.001).

cells after only 15 min exposure to porcine gastric juice (Fig. 8B; 37 °C, pH 2.5). 4. Discussion The aim of the present study was to develop viability-enhancing treatments for probiotic bacteria with a focus on practical immobilization methods for product application. Faecal recovery of L. rhamnosus GG has previously been investigated in humans (Saxelin et al., 1991); however the high dosage required for cell recovery from faeces may be reduced by integrating the concept of cell protection into the administration process. Thus, cell immobilization is a plausible mechanism capable of providing probiotic bacteria with the necessary armament to enhance their stability and viability during storage and gastric transit. Heating whey protein at low ionic strength and at pH values far from the isoelectric point (pH 4.6–5.2) induces the formation of soluble polymers (Ju and Kilara, 1998; Moon and M.E., 1997), which can further self-aggregate and subsequently gel upon controlled acidification (Alting et al., 2000) or increase in ionic strength (Barbut and Foegeding, 1993) thereby constructing a three-dimensional protein network. These properties highlighted whey proteins during the quest for a favourable probiotic immobilization matrix. Preliminary work demonstrated that rapid acidification under agitation generated porous, micron-sized gel particles (10 to 200 μm), capable of hosting bacteria, when incorporated prior to acidification. A prerequisite for the use of stationary phase cells instead of log phase cells was established (Lorca and de Valdez, 1999, van de Guchte et al., 2002) and L. rhamnosus GG cultures (109 cfu/ml) were immobilized in whey protein products derived from native, heat-treated and hydrolysed WPI and the effect of respective treatments on probiotic viability was evaluated under different environmental stresses. AFM analysis focused upon high resolution structural arrangements within the immobilized protein systems. The existence of a micro-phase separation between bacteria and heat-treated/hydrolysed protein (Figs. 2B and 3A) diminished confidence in the concept of complete cell encapsulation by protein networks at pH 5.2. However, AFM probing highlighted the retention of cell surface topography in denatured and hydrolysed WPI following thermal stress at 57 °C, while a catalogue of morphological changes were exhibited by free cells under identical conditions. These findings support the potential probiotic safeguard

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

Fig. 8. FACS bivariate dot plot of L. rhamnosus GG immobilized in heat-treated or hydrolysed WPI (A) after i) 24 h storage at 37 °C and ii) 14-day storage in natural yoghurt at 10 °C. B illustrates the free-cell reference after 4-day storage at 37 °C in addition to 15 min exposure to porcine gastric juice (pH 2.5; 37 °C). C: Cell viability of L. rhamnosus GG (free-cell reference) after 14-day storage at 10 °C in yoghurt.

supplied by denatured and hydrolysed WPI and indicate a plausible mechanism of protection involving cell anchorage within a protein milieu. Light microscopy also proved to be valuable tool for assessment of cell immobilization with heat-treated samples expressing high entrapment efficiency of L. rhamnosus GG within a gelled protein micro-environment. This may be a result of the aggregation pattern induced during acidification of denatured protein, which ultimately withdrew probiotic cells from the serum phase. Hence, this mechanism may provide favourable conditions for increased cell survival at

239

the sol/gel interface. Analysis of the hydrolysed system also revealed partial probiotic enrichment within the aggregated phase and on the aggregate/serum interface (arrows on Fig. 4C); thus, the types of interactions in the hydrolysed system may influence the gel distribution homogeneity in the protein network, which may be conducive to probiotic protection during acid and heat stress. Further practical information relating to cell distribution and viability within immobilized treatments was discovered by analysing LIVE/DEAD stained samples by CSLM and the results obtained coincided with light microscope analysis since probiotics were identified throughout the serum phase of native and hydrolysed treatments, while denatured treatments were characterized by complete probiotic enrichment within the aggregated phase. Furthermore, high cell viability in denatured treatments following 14-day yoghurt incubation at 10 °C provided a visual differentiation factor for the detection of immobilization matrices within the dense protein milieu of yoghurt. Cell viability during storage and thermal treatment illustrated that protein treatment was the major factor governing probiotic survival, with denaturation and hydrolysis providing the dominant armament against temperature stress. Immobilization of stationary phase cells in hydrolysed and denatured WPI significantly (pb 0.001) improved probiotic tolerance to heat stress at 57 °C (7.3±0.1 and 6.5±0.1 log10 cfu/ml, respectively), which represents effective probiotic doses according to the recommended probiotic criterion necessary to achieve therapeutic benefits (Boylston, et al., 2004, Shah and Ravula, 2000). In contrast to this, the sensitivity of L. rhamnosus GG was revealed when free cells, devoid of an immobilization matrix, exhibited dramatic viability losses (6.63 ± 0.08 log10 cycles) following 24 h storage at 37 °C. FACS also expressed an injured free-cell population after 14-day storage in yoghurt, which signals the initiation of significant cell damage upon continued storage. It has been demonstrated that microorganisms that have been injured, but not killed by exposure to stress, often become considerably more sensitive to other types of stress (Ananta et al., 2005, Gardiner, et al., 2000, Mauriello et al., 1999, Teixeira et al., 1997). Thus, the rapid acidification during the free-cell reference preparation may be responsible for the enhanced sensitivity of L. rhamnosus GG during the tested environmental conditions. The aggregation behavior of denatured whey proteins at the isoelectric point may be the reason for the protective probiotic effect offered by denatured WPI by providing a superior driving force for cellentrapment compared to that expressed by native or hydrolysed treatments. Preliminary studies suggested an enhanced sedimentation of WPI gel networks during storage at 37 °C (45.5% ±0.7%) relative to refrigerated conditions (33.0% ±1.4%) following 14-day storage. This assumption concurs with previous findings (Beaulieu et al., 2002), which demonstrated that swelling/contraction of denatured WPI molecules was mainly governed by the net charge of the protein molecules. The protective effect illustrated for free cells following 14-day storage in yoghurt at 10 °C is possibly due to the intrinsic gel characteristics encountered within the yoghurt matrix, which averted a dramatic decline in free-cell survival due to a systematic buffering effect. In light of these results, this immobilization technique may potentially provide a novel safeguard for the incorporation of probiotics into a wide spectrum of challenging, non-gelled food environments, with the aim of extracting the greatest yield from these delivery vehicles. Interestingly, hydrolysed treatments performed well throughout the series of storage temperatures, despite the formation of a weaker protein network compared to that produced by heat-treated WPI (Fig. 4). However, it is noteworthy that the gelling properties of proteins can also be modified via enzymatic hydrolysis (Panyam and Kilara, 1996). In most cases, the propensity of the hydrolysed protein to form a gel is less than that of denatured protein (Kester and Richardson, 1984); nonetheless, some cases of improved enzymeinduced gelation of whey proteins have been reported (Doucet, et al., 2001). The size exclusion data reported in Table 1 were forecast since hydrolysis was performed at the optimum pH conditions (pH 7)

240

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241

required for exo-peptidase activity of Flavourzyme. Peptide aggregation within the hydrolysate milieu is plausible, since hydrophobic interactions among peptides with a molecular mass less than 2 kDa were observed to be involved in the formation of aggregates that associated further to form gel particles (Doucet et al., 2003). It is also noteworthy that the denaturation temperature of β-lg at pH 7 is 70– 73 °C (Hoffmann et al., 1997, Morr and Ha, 1993). Thus, since a large fraction of intact monomeric β-lg continued to prevail at the end of hydrolysis, termination of the enzymatic reaction by heat treatment at 80 °C was pertinent to the unfolding of β-lg with concomitant exposure of extra hydrophobic interaction sites within the aqueous environment. This promoted subsequent aggregation of monomeric βlg. Hence, aggregation behavior within the hydrolysate may be endorsed by i) the gelation of denatured monomers and/or hydrolysed peptides and ii) hydrophobic interactions between these components, which ultimately generated a favourable matrix for probiotic entrapment (Fig. 4C). However, the fact that intact WPI was the weakest substrate for cell immobilization may be related to the absence of gel networks in native protein solutions, which may expose cells to environmental stresses similar to that of free cells and ultimately lead to the significant viability losses shown previously. In addition to thermal tolerance, probiotic microorganisms require the ability to tolerate the acidic conditions encountered during gastric transit (Dunne et al., 2001). Thus, knowledge regarding the capacity of ingested bacteria to resist the primary defense mechanism of the host is fundamental for the selection and successful colonic delivery of probiotic bacteria. Since the pH of porcine gastric juice can vary according to the type of diet and duration after feeding (Jonsson and Conway, 1992, Lawrence, 1970), the pooled gastric contents was standardized (pH 2.5) prior to use. Goldin and Gorbach, (1989) reported that L. rhamnosus GG numbers decreased from 108 to b106 cfu/ml after 30 min at pH 2.5 in human gastric juice, although, the present study combined traditional plate counts and FACS to confirm the dramatic decline in viable counts after 30 min incubation. Direct comparisons between such studies are complicated by variations in test parameters including source and pH of gastric juice, physiological age of the cells, cell re-suspension medium and the initial cell population employed. The enhanced stability of L. rhamnosus GG within heat-treated and hydrolysed systems may possibly reside in the buffering capacity of the surrounding micro-environment since this may be the key factor responsible for substantial cell protection by dairy proteins (Heidebach et al., 2009a,b, Kos et al., 2000). However, buffering capacity fails to explain the significant difference expressed between hydrolysed and heat-treated WPI survival rates; cell immobilization in the former significantly enhanced probiotic viability during one lethal condition (e.g. high heat), but weakened survival during another (e.g. low pH), while immobilization in heat-treated WPI consistently offered protection, regardless of the demanding environmental conditions. Hydrolyzate gastric test results may be supported by the fact that digestion of protein hydrolysates is more rapid than that of an equivalent amount of intact protein (Siemensma et al., 1993) thereby leading to the premature release of probiotics from hydrolysed treatments as a result of biodegradation of the protein system. In contrast to this, denatured treatment results may be correlated with the fact that β-lg is resistant to pepsin, an aspartic protease released in the stomach, in its native state due to the positioning of vulnerable hydrophobic amino acids within its compact globular structure (Guo et al., 1995, Morr and Ha, 1993, Schmidt and Vanmarkwijk, 1993). However, it was assumed that heat denaturation of WPI would significantly enhance the susceptibility of whey protein to proteolytic degradation during gastric transit, since non-native monomeric/oligomeric proteins are known to expose hydrophobic residues upon heat treatment at temperatures N70 °C close to neutral pH (Palazolo et al., 2000). Surprisingly, degradation did not proceed according to expectations; thus one may infer that the hydrophobic patches exposed upon heating are buried in the interior of

the aggregate formed during acidification, potentially averting the release of cells from the protein network (Beaulieu, et al., 2002). Hence, cell immobilization in heat-treated WPI may be an effective mechanism of protection for L. rhamnosus GG in harsh acidic environments such as the stomach, ultimately permitting successful colonic delivery of viable probiotics. 5. Conclusion This study evaluated whey protein products as potential cell immobilization matrices through the exploitation of protein physicochemical properties. The results demonstrate that cell immobilization in denatured or hydrolysed whey protein affords protection to probiotic L. rhamnosus GG during storage at elevated temperatures, heat and acid stress. Interestingly, hydrolysed WPI afforded the greatest protection during high heat conditions, while heat-treated WPI provided a probiotic safeguard during porcine gastric incubation and qualified as the primary cell defense model. These findings advocate whey protein, in its denatured state, as a suitable structural scaffold and successive GI delivery device for probiotic bacteria due to the existence of favourable attributes capable of resisting acid and pepsin degradation in porcine gastric fluids. Furthermore, the characterization of immobilization systems, as probed by AFM, provided a significant understanding of the entrapment ability of respective protein systems. Several outstanding issues remain, however, warranting deeper investigation of the mechanism of protection bestowed upon cells immobilized in hydrolysed or heattreated WPI, in addition to the selection of appropriate product environments for food application. Present findings, inclusive of FACS analysis, are of direct significance to the understanding of acid, thermal tolerance and environmental biocompatibility of probiotic cultures within food systems, which ultimately dictates the success of probiotic delivery.

Acknowledgments The technical assistance of Helen Slattery is gratefully acknowledged in addition to the support provided by the staff of the National Food Imaging Centre. The work was funded by the Irish Dairy Research Trust, the National Development Plan 2007–2013 and Science Foundation Ireland (SFI) funds. S. B. Doherty was funded by the Irish Dairy Research Trust under the Teagasc Walsh Fellowship Scheme.

References Adler-Nissen, J., 1979. Determination of the degree of hydrolysis of food protein hydrolysates by trinitrobenzenesulfonic acid. J. Agric. Food Chem. 27, 1256–1262. Ainsley Reid, A., Vuillemard, J.C., Britten, M., Arcand, Y., Farnworth, E., Champagne, C.P., 2005. Microentrapment of probiotic bacteria in a Ca(2+)-induced whey protein gel and effects on their viability in a dynamic gastro-intestinal model. J. Microencapsul. 22, 603–619. Alting, A.C., Hamer, R.J., de Kruif, G.G., Visschers, R.W., 2000. Formation of disulfide bonds in acid-induced gels of preheated whey protein isolate. J. Agric. Food Chem. 48, 5001–5007. Ananta, E., Volkert, M., Knorr, D., 2005. Cellular injuries and storage stability of spraydried Lactobacillus rhamnosus GG. Int. Dairy J. 15, 399–409. Auty, M.A., Gardiner, G.E., McBrearty, S.J., O'Sullivan, E.O., Mulvihill, D.M., Collins, J.K., Fitzgerald, G.F., Stanton, C., Ross, R.P., 2001. Direct in situ viability assessment of bacteria in probiotic dairy products using viability staining in conjunction with confocal scanning laser microscopy. Appl. Environ. Microbiol. 67, 420–425. Barbut, S., Foegeding, E.A., 1993. Ca2+-induced gelation of pre-heated whey-protein isolate. J. Food Sci. 58, 867–871. Beaulieu, L., Savoie, L., Paquin, P., Subirade, M., 2002. Elaboration and characterization of whey protein beads by an emulsification/cold gelation process: application for the protection of retinol. Biomacromolecules 3, 239–248. Borchers, A.T., Selmi, C., Meyers, F.J., Keen, C.L., Gershwin, M.E., 2009. Probiotics and immunity. J. Gastroenterol. 44, 26–46. Boylston, T.D., Vinderola, C.G., Ghoddusi, H.B., Reinheimer, J.A., 2004. Incorporation of bifidobacteria into cheeses: challenges and rewards. Int. Dairy J. 14, 375–387. Collado, M.C., Isolauri, E., Salminen, S., Sanz, Y., 2009. The impact of probiotic on gut health. Curr. Drug Metab. 10, 68–78.

S.B. Doherty et al. / Journal of Microbiological Methods 80 (2010) 231–241 de Man, J.C., Rogosa, M., Sharpe, M.E., 1960. A medium for cultivation of lactobacilli. J. Appl. Bacteriol. 23, 130–135. Doucet, D., Gauthier, S.F., Foegeding, E.A., 2001. Rheological characterization of a gel formed during extensive enzymatic hydrolysis. J. Food Sci. 66, 711–715. Doucet, D., Otter, D.E., Gauthier, S.F., Foegeding, E.A., 2003. Enzyme-induced gelation of extensively hydrolyzed whey proteins by Alcalase: peptide identification and determination of enzyme specificity. J. Agric. Food Chem. 51, 6300–6308. Dunne, C., O'Mahony, L., Murphy, L., Thornton, G., Morrissey, D., O'Halloran, S., Feeney, M., Flynn, S., Fitzgerald, G., Daly, C., Kiely, B., O'Sullivan, G.C., Shanahan, F., Collins, J.K., 2001. In vitro selection criteria for probiotic bacteria of human origin: correlation with in vivo findings. Am. J. Clin. Nutr. 73, 386S–392S. FAO/WHO, 2001. Health and nutrition properties of probiotics in food including powder milk with live lactic acid bacteria. Report from FAO/WHO Expert Consultation, 1–4 October 2001, Cordoba, Argentina. Garcia Alvarez-Coque, M.C., Medina Hernandez, M.J., Villanueva Camanas, R.M., Mongay Fernandez, C., 1989a. Formation and instability of o-phthalaldehyde derivatives of amino acids. Anal. Biochem. 178, 1–7. Garcia Alvarez-Coque, M.C., Medina Hernandez, M.J., Villanueva Camanas, R.M., Mongay Fernandez, C., 1989b. Studies on the formation and stability of isoindoles derived from amino acids, o-phthalaldehyde and N-acetyl-L-cysteine. Anal. Biochem. 180, 172–176. Gardiner, G., Stanton, C., Lynch, P.B., Collins, J.K., Fitzgerald, G., Ross, R.P., 1999. Evaluation of Cheddar cheese as a food carrier for delivery of a probiotic strain to the gastrointestinal tract. J. Dairy Sci. 82, 1379–1387. Gardiner, G.E., O'Sullivan, E., Kelly, J., Auty, M.A., Fitzgerald, G.F., Collins, J.K., Ross, R.P., Stanton, C., 2000. Comparative survival rates of human-derived probiotic Lactobacillus paracasei and L. salivarius strains during heat treatment and spray drying. Appl. Environ. Microbiol. 66, 2605–2612. Goldin, B.R., Gorbach, S.L., 1989. Lactobacillus strain and methods of selection., Official Gazette of the United States Patent and Trademark Office Patents., Vol. US 4839281, U.S. Guerin, D., Vuillemard, J.C., Subirade, M., 2003. Protection of bifidobacteria encapsulated in polysaccharide-protein gel beads against gastric juice and bile. J. Food Prot. 66, 2076–2084. Gunasekera, T.S., Attfield, P.V., Veal, D.A., 2000. A flow cytometry method for rapid detection and enumeration of total bacteria in milk. Appl. Environ. Microbiol. 66, 1228–1232. Guo, M.R., Fox, P.F., Flynn, A., Kindstedt, P.S., 1995. Susceptibility of beta-lactoglobulin and sodium caseinate to proteolysis by pepsin and trypsin. J. Dairy Sci. 78, 2336–2344. Heidebach, T., Först, P., Kulozik, U., 2009a. Microencapsulation of probiotic cells by means of rennet-gelation of milk proteins. Food Hydrocolloids 23, 1670–1677. Heidebach, T., Först, P., Kulozik, U., 2009b. Transglutaminase-induced caseinate gelation for the microencapsulation of probiotic cells. Int. Dairy J. 19, 77–84. Hoffmann, M.A.M., van Miltenburg, J.C., van Mil, P.J.J.M., 1997. The suitability of scanning calorimetry to investigate slow irreversible protein denaturation. Thermochimica Acta 306, 45–49. Holt, C., 2000. Molecular basis of whey protein food functionalities. Aust. J. Dairy Technol. 55, 53. Isolauri, E., Juntunen, M., Rautanen, T., Sillanaukee, P., Koivula, T., 1991. A human Lactobacillus strain (Lactobacillus casei sp strain GG) promotes recovery from acute diarrhea in children. Pediatrics 88, 90–97. Isolauri, E., Salminen, S., Mattila-Sandholm, T., 1999. New functional foods in the treatment of food allergy. Ann. Med. 31, 299–302. Jonsson, E., Conway, P., 1992. Probiotics for pigs. In: Fuller, R. (Ed.), Probiotics: The Scientific Basis. Chapman & Hall, Boundary Hall, London SE1 8HN, pp. 260–317. Ju, Z.Y., Kilara, A., 1998. Textural properties of cold-set gels induced from heat-denatured whey protein isolates. J. Food Sci. 63, 288–292. Kailasapathy, K., Chin, J., 2000. Survival and therapeutic potential of probiotic organisms with reference to Lactobacillus acidophilus and Bifidobacterium spp.Immunol. Cell Biol. 78, 80–88. Kester, J.J., Richardson, T., 1984. Modification of whey proteins to improve functionality. J. Dairy Sci. 67, 2757–2774.

241

Knorr, D., 1998. Technology aspects related to microorganisms in functional foods. Trends Food Sci. Technol. 9, 295. Kos, B., Suskovic, J., Goreta, J., Matosic, S., 2000. Effect of protectors on the viability of Lactobacillus acidophilus M92 in simulated gastrointestinal conditions. Food Technol. Biotechnol. 38, 121–127. Lawrence, T., 1970. Some effects of including differently processed barley in diet of growing pig. 2. In vivo pH gastric changes. Anim. Prod. 12, 151–163. Lorca, G.L., de Valdez, G.F., 1999. The effect of suboptimal growth temperature and growth phase on resistance of Lactobacillus acidophilus to environmental stress. Cryobiology 39, 144–149. Mauriello, G., Aponte, M., Andolfi, R., Moschetti, G., Villani, F., 1999. Spray-drying of bacteriocin-producing lactic acid bacteria. J. Food Prot. 62, 773–777. Moon, J.C.a.M., M.E., 1997. Effects of protein unfolding and soluble aggregates formation on the gel strength of whey proteins. J. Food Sci. Nutr. 2, 281–284. Morr, C.V., Ha, E.Y., 1993. Whey protein concentrates and isolates: processing and functional properties. Crit. Rev. Food Sci. Nutr. 33, 431–476. Nedovic, V., Willaert, R., 2005. Applications of cell immobilization biotechnology. In: Nedovic, V., Willaert, R. (Eds.), Focus on Biotechnology, vol. 8b. Springer, pp. 295–314. Palazolo, G., Rodriguez, F., Farruggia, B., Pico, G., Delorenzi, N., 2000. Heat treatment of beta-lactoglobulin: structural changes studied by partitioning and fluorescence. J. Agric. Food Chem. 48, 3817–3822. Panyam, D., Kilara, A., 1996. Enhancing the functionality of food proteins by enzymatic modification. Trends Food Sci. Technol. 7, 120–125. Picot, A., Lacroix, C., 2004. Encapsulation of bifidobacteria in whey protein-based microcapsules and survival in simulated gastrointestinal conditions and in yoghurt. Int. Dairy J. 14, 505–515. Ross, R.P., Desmond, C., Fitzgerald, G.F., Stanton, C., 2005. Overcoming the technological hurdles in the development of probiotic foods. J. Appl. Microbiol. 98, 1410–1417. Salminen, S., von Wright, A., Morelli, L., Marteau, P., Brassart, D., de Vos, W.M., Fonden, R., Saxelin, M., Collins, K., Mogensen, G., Birkeland, S.E., Mattila-Sandholm, T., 1998. Demonstration of safety of probiotics—a review. Int. J. Food Microbiol. 44, 93–106. Saxelin, M., Elo, S., Salminen, S., Vapaatalo, H., 1991. Dose response colonization of feces after oral administration of Lactobacillus-casei strain GG. Microb. Ecol. Health Dis. 4, 209–214. Saxelin, M., Grenov, B., Svensson, U., Fonden, R., Reniero, R., Mattila-Sandholm, T., 1999. The technology of probiotics. Trends Food Sci. Technol. 10, 387. Schmidt, D.G., Vanmarkwijk, B.W., 1993. Enzymatic-hydrolysis of whey proteins— influence of heat-treatment of alpha-lactalbumin and beta-lactoglobulin on their proteolysis by pepsin and papain. Net. Milk Dairy J. 47, 15–22. Shah, N.P., Ravula, R.R., 2000. Microencapsulation of probiotic bacteria and their survival in frozen fermented dairy desserts. Aust. J. Dairy Technol. 55, 139–144. Siemensma, A.D., Weijer, W.J., Bak, H.J., 1993. The importance of peptide lengths in hypoallergenic infant formulas. Trends Food Sci. Technol. 4, 16–21. Siro, I., Kapolna, E., Kapolna, B., Lugasi, A., 2008. Functional food. Product development, marketing and consumer acceptance—a review. Appetite 51, 456–467. Spellman, D., McEvoy, E., O'Cuinn, G., FitzGerald, R.J., 2003. Proteinase and exopeptidase hydrolysis of whey protein: comparison of the TNBS, OPA and pH stat methods for quantification of degree of hydrolysis. Int. Dairy J. 13, 447–453. Sun, W., Griffiths, M.W., 2000. Survival of bifidobacteria in yogurt and simulated gastric juice following immobilization in gellan-xanthan beads. Int. J. Food Microbiol. 61, 17–25. Teixeira, P., Castro, H., Mohacsi-Farkas, C., Kirby, R., 1997. Identification of sites of injury in Lactobacillus bulgaricus during heat stress. J Appl Microbiol. 83, 219–226. Van de Casteele, S., Vanheuverzwijn, T., Ruyssen, T., Van Assche, P., Swings, J., Huys, G., 2006. Evaluation of culture media for selective enumeration of probiotic strains of lactobacilli and bifidobacteria in combination with yoghurt or cheese starters. Int. Dairy J. 16, 1470–1476. van de Guchte, M., Serror, P., Chervaux, C., Smokvina, T., Ehrlich, S.D., Maguin, E., 2002. Stress responses in lactic acid bacteria. Antonie Van Leeuwenhoek 82, 187–216. Vasiljevic, T., Shah, N.P., 2008. Probiotics—from Metchnikoff to bioactives. Int. Dairy J. 18, 714–728.