Survival of entrapped Lactobacillus rhamnosus GG in whey protein micro-beads during simulated ex vivo gastro-intestinal transit

International Dairy Journal 22 (2012) 31e43

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Survival of entrapped Lactobacillus rhamnosus GG in whey protein micro-beads during simulated ex vivo gastro-intestinal transit S.B. Dohertya, b, M.A. Autya, C. Stantona, c, R.P. Rossa, c, G.F. Fitzgeraldb, c, A. Brodkorba, * a

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

a r t i c l e i n f o

a b s t r a c t

Article history: Received 15 February 2011 Received in revised form 16 June 2011 Accepted 17 June 2011

Cell survival of Lactobacillus rhamnosus GG entrapped in gelled whey protein isolate (WPI) micro-beads was elucidated relative to cells suspended in native WPI and free-cell controls during ex vivo porcine gastro-intestinal incubation. Probiotic gastric tolerance was investigated as a function of pH (2.0e3.4) and time with subsequent intestinal incubation (pH 7.2). Free cells showed no survival after 30 min ex vivo stomach incubation (
pH 3.4), while native WPI enhanced survival by 5.7  0.1, 5.1  0.2 and 2.2  0.2 log10 cfu mL1 following 180 min incubation at pH 3.4, 2.4 and 2.0, respectively. Protein microbeads augmented ex vivo probiotic acid resistance (8.9  0.1 log10 cfu mL1) and demonstrated significant micro-bead adsorption capacity coupled with micro-bead digestion and controlled release of viable, functional probiotics within 30 min intestinal incubation. This technology potentially envisions whey protein micro-beads as efficacious entrapment matrices and binding vehicles for delivery of bioactive ingredients. Ó 2011 Elsevier Ltd. All rights reserved.

1. Introduction Cell encapsulation, immobilization and entrapment epitomize elementary practices for proficient bioactive delivery and controlled release in pharmaceutical, food and flavour industries (de Vos, Faas, Spasojevic, & Sikkema, 2010). Many in vitro studies illustrate the aptitude of entrapment technologies for amplification of probiotic survival during quasi-stomach conditions (Burgain, Gaiani, Linder, & Scher, 2011). However, the selection of entrapment procedures varies according to the research field since protection of cells and bioactive molecules is governed by different physico-chemical and molecular requirements unique to individual bacterial species and compounds. Alginate e the quintessential encapsulation material e is susceptible to disintegration in the presence of excess monovalent ions and Ca2þ chelating agents (Smidsrod & Skjak-Braek, 1990); hence, stable cell entrapment in alginate remains an difficult challenge. According to common credence, research has failed to fabricate a single matrix with all the essential entrapment characteristics; meanwhile, dairy proteins are attracting industrial and academic curiosity as potential alternatives to coated-alginate and composite * Corresponding author. Tel.: þ353 25 42222; fax: þ353 25 42340. E-mail address: [email protected] (A. Brodkorb). 0958-6946/$ e see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.idairyj.2011.06.009

entrapment matrices. Whey proteins, in particular, have a high biological value (Smithers, 2008) partly due to the high content of branched-chain essential amino acids, which stimulate specific intracellular pathways. From a formulation perspective, whey proteins have been exploited as operational scaffolds for drug and cell entrapment due to their aptitude to form emulsions (Beaulieu, Savoie, Paquin, & Subirade, 2002) and gastro-resistant hydro-gels (Ainsley Reid, Champagne, Gardner, Fustier, & Vuillemard, 2007). b-Lactoglobulin (b-Lg), the most abundant whey protein, is a small globular protein with specific affinity for a variety of hydrophobic and amphipathic compounds, including retinol (Kontopidis, Holt, & Sawyer, 2002), phospholipids (Lefevre & Subirade, 2001) and aromatic compounds (Collini, D’Alfonso, & Baldini, 2000). Hence, whey proteins are emblematic of versatile carriers of hydrophobic molecules and probiotic bacteria in controlled release applications. Probiotic bacteria, defined as ‘live micro-organisms, which when administered in adequate amounts, confer a health benefit on the host’, are considered safe for human consumption and illustrate potential for immunomodulation, treatment and prevention of disorders and diseases (Mattila-Sandholm et al., 2002). However, commercially available probiotic products and supplements often provide inadequate cell populations (<107 viable cells) due to harsh processing conditions encountered during manufacture of the carrier system.

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Probiotic research focus has recently shifted due to the realization that probiotic effects are dependent upon the presence of functional ligands or “effector molecules” in the probiotic cell envelope (van Baarlen et al., 2009). These characteristic bioactive molecules are adversely affected by enzymatic action of pepsin and low pH of the stomach with further antagonism associated with antimicrobial activity of bile salts and protease-rich conditions of the intestine (Konstantinov et al., 2008). Hence, effector molecule functionality requires specific conservation requisites during product manufacture and gastro-intestinal (GI) transit; a probiotic functionality consideration more influential than cell survival concentration. In essence, validation of cell survival via plate enumeration will not necessarily demonstrate the preservation of probiotic functional benefits. This indispensable issue has recently catalyzed industrial consciousness for an auxiliary requirement of probiotic functionality. Probiotic bacteria would therefore profit from entrapment ascendancy to avert cell demise for targeted functional delivery to their absorption site. This research focused on developing a cell-entrapment method (Doherty et al., 2011), which validated in vitro probiotic delivery in whey protein micro-beads. Progressive investigations evaluated whey protein matrices as preservation edifices for intestinal delivery of viable, functional cell populations of Lactobacillus rhamnosus GG during ex vivo gastro-intestinal studies. Analysis of whey protein micro-beads and innate cell-surface features were auxiliary elements of this research endeavour for determination of protein matrix compatibility as cell adhesion and functional delivery devices. 2. Materials and methods 2.1. Biochemicals BiPro, a commercial whey protein isolate (WPI) obtained from Davisco Foods International Inc. (Minnesota, U.S.A.) contained 98% (w/w) protein. Native b-Lg and a-lactalbumin (a-La) content in WPI were analyzed by reverse phase-HPLC and estimated at 82% and 16%, respectively. Pepsin (P-7000), sodium acetate and fast green fluorescent dye (product code F7252) were obtained from Sigma Aldrich (Dublin, Ireland) and TweenÒ 20 and AristarÒ Plus grade acetic acid were obtained from VWR International Ltd., Dublin, Ireland. Chemical products acetonitrile (MeCN) and trifluoroacetic acid (TFA), both HPLC grade, were purchased from Fisher Scientific Ltd. (Dublin, Ireland). Milli-Q water (Millipore, Cork, Ireland) was sterilized and utilized in all cases for dispersion of samples, culture mediums and buffer solutions. 2.2. Bacterial strain and culture conditions The probiotic strain Lb. rhamnosus GG (ATCC 53103, LGG, Valio Ltd., Helsinki, Finland), was sourced from the Moorepark culture collection, under a restricted materials transfer agreement. Cells were harvested and stored at 20  C as stock solutions in de Man Rogosa Sharpe (MRS) broth (Oxoid Ltd., Hampshire, U.K.) containing 50% (v/v) aqueous glycerol (Ref. G5516; Sigma Aldrich). Due to the porcine origin of GI digesta, a spontaneous rifampicin-resistant derivative (LGGRif) was required to facilitate selective enumeration of the administered strain during ex vivo studies. Since many lactobacilli demonstrate resistance to vancomycin (Klein et al., 2000), rifampicin-resistant variants were generated according to the method outlined by Gardiner et al. (1999) whereby single colonies were selected and stocked after anaerobic incubation at 37  C for 48 h. Randomly amplified polymorphic DNA-PCR (RAPD-PCR) (Coakley, Ross, & Donnelly, 1996), growth characteristics, heat and acid tolerance was evaluated for both strains to ensure homology

between parent and variant strains. Subcultures were routinely checked for purity using pulse-field gel electrophoresis (PFGE) (Simpson, Stanton, Fitzgerald, & Ross, 2002) and all cell cultures were propagated from 1% (v/v) inoculations for 19 h at 37  C under anaerobic conditions, achieved using Anaerocult gas packs (Merck KGaA, Darmstadt, Germany). This study utilizes stationary phase colony forming units (cfu) since the stress response is generally more resistant to environmental factors (van de Guchte et al., 2002) and all cell cultures were propagated from 1% (v/v) inoculations for 19 h at 37  C under anaerobic conditions. 2.3. Micro-entrapment procedure WPI micro-beads were prepared according to the method outlined by Doherty et al. (2011) whereby a dispersion of WPI (11%, w/ v) was adjusted to pH 7, heated (78  C, 45 min) and the suspension of reactive WPI aggregates was subsequently cooled and refrigerated overnight. A proteineprobiotic blend containing 9% (w/v) WPI and 109 cfu mL1 Lb. rhamnosus GG was aseptically extruded through a 150 mm nozzle into 250 mL of curing media (0.5 M sodium acetate; 0.04% TweenÒ 20; pH 4.6) tempered to 35  C using an Encapsulator Model IE-50R from EncapBioSystem (Greifensee, Switzerland). Micro-bead batches containing 1.7  1010 cfu were polymerized in curing buffer, recovered and subjected to immediate analysis. 2.4. Collection of specimens All ex vivo studies were performed using extracted gastrointestinal (GI) contents (gastric e lower stomach; intestine e ileum and caecum) collected from six finisher pigs starved for 16 h prior to slaughter. Porcine slaughter was performed in compliance with European Union Council Directive 91/630/EEC (outlines minimum standards for the protection of pigs) and European Union Council Directive 98/58/EC (concerns the protection of animals kept for farming purposes). Upon receipt of GI contents, gastric and intestinal digesta were filtered through glass wool, centrifuged (8600  g, 45 min) and clarified using Whatman paper no. 4 (Whatman International Ltd., Kent, U.K.). All samples were checked for sterility using brain heart infusion agar (Merck KGaA) and incubated at 37  C for 48 h, while total Lactobacillus counts were determined on Lactobacillus-selective agar (LBS; Becton Dickinson, Oxford, U.K.) following anaerobic incubation for 5 d at 37  C. GI digesta were also tested on MRSRif plates to detect the presence/ absence of LGG. RAPD analysis was also performed to further investigate the resident microbiota of respective porcine GI regions. All GI contents were analyzed within 6 h of slaughter for (i) pH determination (Mettler Toledo MP220 pH meter; VWR International Ltd.), (ii) protein content according to the Bradford assay (Bradford, 1976) and (iii) activity of respective GI enzymes (see Section 2.5). Following this, stomach and intestinal digesta were individually pooled and stored at 20  C. 2.5. Enzyme assays 2.5.1. Pepsin activity Enzymatic activity of pepsin (EC 3.4.23.2) in porcine gastric contents was evaluated using denatured haemoglobin as a substrate and purified pepsin as the reference material. The reaction was started by the addition of 5 mL gastric juice to an equal volume of acid-denatured haemoglobin solution (20 mg mL1) prepared in 10 mM HCl and incubated at 37  C. Sample reactions were stopped after 10 min by the addition of 10 mL 5% (w/v) trichloroacetic acid (TCA), followed by centrifugation (5000  g, 5 min, 20  C), filtration (0.45 mm) and absorbance measurement at 280 nm using a Cary

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spectrophotometer (JVA Analytical, Dublin, Ireland). The pepsin concentration in gastric contents was estimated from reference samples of known pepsin concentration, which determined the pepsin activity using the following equation:

½A ðsampleÞ  A280nm ðblankÞ  1000  V y ¼ 280nm T  E  EC

(1)

where y ¼ pepsin units mL1 GI contents; V ¼ filtrate volume (mL); T ¼ time of assay (min); E ¼ enzyme in the reaction mixture (mg); EC ¼ extinction coefficient for tyrosine (280 nm) and A280nm ¼ absorbance at 280 nm. One unit (IU) enzyme activity was defined as the amount of enzyme which liberated 1 mmol of tyrosine (min1 mg protein1). Optimum pepsin activity was achieved at ratios of substrate to porcine gastric juice of 1 mg mL1 and results demonstrated 3.9 mg protein per mL gastric juice with 44.5 IU pepsin activity. 2.5.2. Endopeptidase activity in intestinal juice Casein was chosen as the substrate to evaluate the activity of endo-proteases in porcine intestinal contents. Reference protease assays were initially performed for trypsin (EC 3.4.21.4) and a-chymotrypsin (EC 3.4.21.1) standard solutions using casein (prepared by the Hammarstein method; Dunn, 1949) as the substrate during TCA-precipitation reactions. At 30 s intervals after mixing, terminated reactions were held for 1 h at room temperature and the increase in TCA-soluble products was measured at 280 nm. Absorbance values were plotted as a function of time and the slope was converted into mmol tyrosine equivalents released (mg protein min1)1 for standard trypsin and a-chymotrypsin solutions. Following this, sulphanilamide-azocasein (MegazymeÒ, Wicklow, Ireland) was utilized to determine specific trypsin and a-chymotrypsin activity in intestinal contents. Samples were blended with equal volumes of substrate for 10 min at 40  C. Following this, non-hydrolyzed Azocasein was precipitated by the addition of 5% TCA, mixtures were centrifuged (1000  g, 10 min) and casein hydrolysates were assayed spectrophotometrically at 440 nm against the relevant blanks. Analogous to pepsin activity, one unit of trypsin/a-chymotrypsin activity was defined as the amount of enzyme required to hydrolyze 1 mmol tyrosine equivalents (mg protein min1)1 from soluble casein under standard conditions (pH 7.0; 40  C) using the following equation:

y ¼ milliUnits ðassayÞ1 0:001  df

(2)

1

where y ¼ endopeptidase units mL of intestinal contents; milliUnits (assay)1, which referred to the standard curve for enzyme action on casein and df ¼ dilution factor applied to original intestinal samples. Intestinal assays aforementioned were performed in triplicate on five independent porcine GI samples. Data determined enzyme activities of 21.5 and 318.8 IU for trypsin and chymotrypsin, respectively, in ex vivo intestinal juice. Table 1 illustrates amino acid profiles of stomach and intestinal porcine digesta and reveals copious amounts of essential amino acids and protein precursors with total concentrations of 47.6 and 45.4 mg mL1, respectively. 2.6. Viability of encapsulated Lb. rhamnosus GG in ex vivo porcine gastric contents The protective effect of micro-entrapment on LGGRif upon exposure to gastric juice at various natural pH values was assessed as follows: entrapped, suspended and free LGGRif (approx 109 cfu mL1) were incubated in gastric contents (1:10 dilution) for 180 min at 37  C with orbital agitation (150 rpm) in a temperature-controlled environment incubator. To determine probiotic viability, treatment samples from entrapped, suspended and free LGGRif were recovered

33

Table 1 Analysis of ex vivo stomach and intestinal contents based on protein content (mg mL1), enzyme activity (IU units), total and individual free amino acid concentrations (mg mL1).a Characterization assays

Stomach

Intestine

Protein content (mg mL1) Pepsin activity (IU units) Chymotrypsin (IU units) Trypsin (IU units) Total free amino acids (mg mL1) Asp Thr Ser Glu Gly Ala Cys Val Met Ile Leu Tyr Phe His Lys NH3 Arg

3.9 44.5 ND ND 28,926  23 1705  3 2145  2 1362  4 3213  5 2204  2 1870  1 375  1 1892  1 723  3 1562  2 2743  2 1311  3 1351  2 801  1 2707  2 422  2 2540  1

7.8 ND 319.8 21.5 27,603  34 1728  1 2142  2 1439  1 3193  2 2167  3 1851  2 434  1 1881  2 686  1.5 1570  1 2721  2 1050  2 1371  1 770  2 2519  1 264  2 1817  2

a One enzyme unit (IU) is defined as the amount of enzyme that liberated 1 mmol of tyrosine min1 mg protein1 and all amino acid data represent the mean value of five independent tests conducted in triplicate; ND denotes no enzyme detection.

from gastric digesta at pre-determined time points, dispersed in a selective phosphate buffer (0.5 M; pH 7) and stored on ice for 5 min to terminate any residual enzymatic reactions. Samples were homogenized according to a previously validated procedure (Doherty et al., 2010b) as 10-fold dilutions (w/w) using an Ultra-TurraxÒ T10 (IKAÒ Werke, GmbH & Co. KG, Staufen, Germany) to ensure complete liberation of bacteria from the protein matrix with no adverse effect on cell viability. Planktonic (free) cells were treated similarly to maintain consistent treatment conditions. Homogenates were serially diluted in sterile maximum recovery diluent (MRD, Oxoid, Ltd.,) and appropriate dilutions were spread-plated on two media for enumeration of LGGRif and total Lactobacillus. Cell counts for LGGRif were obtained using MRS agar containing rifampicin (Sigma Aldrich) as a selective agent and 50 U mL1 of nystatin (Sigma Aldrich) to inhibit yeasts and moulds after anaerobic incubation for 2 d at 37  C (MRSRif) and total Lactobacillus counts were detected on Lactobacillus-selective agar following anaerobic incubation for 5 d. RAPD-PCR was performed as described by Coakley et al. (1996), which validated the presence of LGGRif on MRSRif plates and all tests were conducted in triplicate and mean log survivor counts were plotted as a function of incubation time. pH values of recovered GI digesta were recorded as a function of incubation time in order to investigate buffer capacity of the micro-bead treatments. 2.7. Release of encapsulated Lb. rhamnosus GG as a function of porcine gastro-intestinal section Entrapped cells (108 cfu mL1) were anaerobically incubated for maximum of 12 h in porcine digesta from different sections of the GI tract (stomach, ileum and caecum). Aliquots of GI digesta were removed at specific time intervals, resuspended in phosphate buffer (0.5 M; pH 7), stored on ice for 5 min and homogenized for subsequent LGGRif and total lactobacilli enumeration as outlined above. 2.8. Live/dead discrimination by flow cytometry In addition to plate counts, viability of encapsulated and free LGGRif suspensions were assessed by flow cytometry (FACS) as

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outlined previously by Doherty et al. (2010b) using voltages and threshold values calibrated for the enumeration of LGGRif. Briefly, samples from gastric and intestinal incubation studies were homogenized to release cells from respective protein matrices, followed by dilution (107 cfu mL1) with working solutions of Thiazole Orange (TO) and Propidium Iodide (PI) fluorescent stains. 2.9. Physico-chemical characterization of micro-bead degradation 2.9.1. Micro-bead digestion Degree of hydrolysis (DH) was also assayed as described by Doherty et al. (2011) using the o-phthaldialdehyde (OPA) spectrophotometric assay. Proteolysis was analyzed in each GI sample for each time point during five independent studies. 2.9.2. Protein analysis To assess the degree of micro-bead degradation, the amount of free protein after GI incubation was determined using the Bradford assay (Bradford, 1976) and reference samples were also prepared by blending micro-beads with 0.2% NaCl solution at neutral pH, instead of GI digesta solution. At specific time points, samples were centrifuged (1000  g, 5 min), supernatant removed, filtered and analyzed using the Bradford assay. Size exclusion chromatography (SEC) was also performed using an automated 2695 WatersÔ HPLC system (Waters, Dublin, Ireland) equipped with a TSK G2000 SW column (600  7.5 mm; Tosu Hass, Japan) according to the method outlined by Doherty et al. (2010a) using 30% acetonitrile containing 0.1% (v/v) TFA. Samples were analyzed in triplicate for each time point for evaluation of pH and enzymatic synergism during microbead disintegration. Free amino acid analysis was also performed in triplicate on recovered GI media at specified time points as demonstrated previously (Doherty et al., 2011) using a Jeol JLC-500/ V amino acid analyser (Jeol (UK) Ltd., Herts, U.K.).

2.10. Hydrophobicity 2.10.1. Micro-bead surface hydrophobicity Surface hydrophobicity (SH) of whey protein micro-beads were determined using the SDS binding method outlined by Kato, Matsuda, Matsudomi, and Kobayashi (1984) with particular adjustment for whey protein profiles. Protein micro-bead batches were suspended in sodium dihydrogen phosphate dihydrate buffer (0.02 M; pH 6.0), while SDS reagent (40.37 mg L1) and methylene blue (24.0 mg L1) were prepared separately in fresh buffer solutions. Individual micro-bead batches (109 cfu mL1, 90 mg WPI mL1) were mixed with SDS reagent (1:2 ratio), incubated for 30 min at 20  C under slight agitation and subsequently dialyzed against the phosphate buffer (ratio 1:25, v/v) for 24 h at 20  C. Mixtures of 0.5 mL of dialysate, 2.5 mL of methylene blue, and 10 mL of chloroform were centrifuged at 2500  g for 5 min. The extinction co-efficient (3 ) of the chloroform phase was assessed at a wavelength of l ¼ 655 nm according to Hiller and Lorenzen (2008). Measurements were performed in triplicate and SH of fresh micro-beads batches were assessed relative to batches procured as a function of ex vivo gastric incubation time. Native and heat-treated WPI represented positive and negative controls, respectively, and all treatments contained equivalent protein concentrations. 2.10.2. Probiotic strain hydrophobicity The apparent hydrophobicity of Lb. rhamnosus GG and LGGRif cell surfaces were evaluated as a function of microbial adhesion to hydrocarbons (MATH) according to a modified method of Rosenburg (1991). Bacterial cells were harvested by centrifugation and resuspended in potassium phosphate buffer (Doherty et al., 2010b), while MATH experiments were performed by spectrophotometer analysis. Analogous to this, cells liberated from microbeads following 3 h ex vivo intestinal incubation at 37  C were recovered and subject to similar hydrophobicity analysis. 2.11. Statistical analysis

2.9.3. Microscopy The dimensions of the whey protein micro-bead were determined by bright-field light microscopy using a BX51 light microscope (Olympus, Essex, U.K.) and all micro-bead batches were examined following staining with fast green fluorescent dye. Additional microscopy work was performed using a Leica TCS SP5 confocal scanning laser microscope (CSLM) (Leica Micro systems, Wetzlar, Germany) for visualization of micro-bead digestion during gastric and intestinal incubation as a function of time. Probiotic micro-beads were stained with BD Cell Viability kit (BD Biosciences, Oxford, U.K.) and imaged as previously described by Auty et al. (2001) while atomic force microscope (AFM) images were also obtained using MFP-3D-AFM instrumentation (Asylum Research UK Ltd., Oxford, U.K.) according to Doherty et al. (2010a). 2.9.4. Electrophoresis The average molecular weights (AMW) of peptides procured during micro-bead digestion in intestinal media were estimated by SDS-PAGE under reducing conditions according to the method described by Laemmli (1970). Treated samples were loaded onto a stacking gel of 4% (w/v) acrylamide (pH 6.8) and separated on a gel containing 15% (w/v) acrylamide (pH 8.8) whereby each gel contained 0.1% SDS. The electrophoresis was performed at a constant voltage of 180 V in a mini Protean II system (Bio-Rad Alpha Technologies, Dublin, Ireland) and gels were stained in a 0.5% Coomassie brilliant blue R-250, 25% iso-propanol, 10% acetic acid solution. The AMW of the protein bands of electrophoretically separated matrix components were estimated by comparison of their mobility to those of standard proteins (Precision Plus ProteinÔ Standards, Bio-Rad Alpha Technologies).

All experimental measurements were conducted in triplicate during five independent studies (unless stated otherwise). Average values and the standard deviation (SD) were calculated and mean log survival counts were plotted as a function of incubation time. Student t-tests were performed 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 (*p < 0.05; **p < 0.01; ***p < 0.001). 3. Results 3.1. Viability of entrapped Lb. rhamnosus GG in ex vivo porcine gastric contents Fig. 1 illustrates the survival of LGGRif in porcine gastric juice at (A) pH 3.4, (B) pH 2.4 and (C) pH 2.0. LGGRif entrapment in whey protein micro-beads significantly (p < 0.001) increased gastric cell survival relative to free cell and native protein suspensions, both of which demonstrated profound (p
0.01) cell loss after 30 min at 37  C. Complementary FACS data further validated cell viability within micro-bead matrices since LGGRif populations were exclusively located in gate A3 of FACS dot plots, indicating the presence of viable cells with intact, functional cell membranes after 180 min gastric incubation at pH 3.4 (Fig. 1A; top inset). In all cases (pH 3.4e2.0), incubation of free LGGRif for 30 min gave complete cell mortality (8.9  0.1 log10 cfu mL1) compared with maximum cell loss of approximately 2 log10 cycles in native protein suspensions during this time. After 60 min at pH 2.4 and 2.0, however, native protein protective properties weakened significantly (p < 0.001)

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Fig. 1. Survival of LGGRif entrapped in micro-beads (C), suspended in native WPI (:) and free-cell controls (-) in ex vivo porcine gastric contents at pH 3.4 (A), 2.4 (B) and 2.0 (C) at 37  C for 3 h with agitation in an orbital shaker at 150 rpm. The insets represent flow cytometry (FACS) dot plots with a specified gating strategy distinguishing live (A3), injured (A2) and dead (A1) cells, while A4 represents debris from respective samples. Confocal scanning laser microscopy (CSLM) images illustrate green and red cells, representing live and dead probiotic bacteria, respectively. Significant differences (*p < 0.05; **p < 0.01; ***p < 0.001) within each treatment are illustrated as a function of respective incubation time periods, i.e., 0e30, 30e60, 60e120 and 120e180 min. Standard deviation is indicated by the vertical bars. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

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expressing 3.4  0.3 and 5.2  0.6 log10 cycle reductions, respectively. Plate enumeration was supported by FACS analysis that illustrated injured and dead cell populations, and mixtures thereof, in native protein and free-cell treatments, respectively (Fig. 1A; insets). Overall, native WPI provided a moderate stabilizing façade for probiotic bacteria during stomach incubation but significant (p < 0.001) cell loss (3.8  0.4 log10 cfu mL1) and cell injury were illustrated after 120 min incubation at pH 2.4, since FACS data revealed the rapid transition of probiotic populations from functional to compromised cell conditions (Fig. 1B; middle inset). After 180 min at pH 2.0, cell mortality (6.8  0.7 log10 cfu mL1) demonstrated the fragile fascia of native WPI, while FACS revealed the emergence of a comprehensively injured cell population (Fig. 1C; middle inset). Hence, FACS analysis revealed acid and peptic sensitivity of LGGRif; an attribute that cannot be fully compensated by the presence of native WPI. Conversely, entrapped treatments revealed a pronounced protective sheath during initial 30 min incubation, which persisted with no significant difference as a function of pH and incubation time. After 180 min at pH 2.4 and 2.0, plate enumeration and FACS demonstrated a distinct amelioration of cell viability in entrapped WPI lattices relative to free-cell controls (8.7  0.1 and 8.4  0.2 log10 cfu mL1, respectively). Confocal microscopy (CSLM) in Fig. 1B, C revealed transverse sections of micro-beads with an apparent abundance of live LGGRif evenly distributed within denatured WPI lattices following 180 min gastric incubation at pH 2.4 and 2.0, respectively. To maintain treatment homology, however, native protein treatments were also extruded through the entrapment system for an objective comparison with micro-bead treatments; hence, all treatments were subject to identical shear forces experienced during the

extrusion process. It is noteworthy that extrusion through a 150 mm nozzle exhibited no significant (p > 0.01) effect on entrapment efficiency of LGGRif in denatured micro-bead matrices (Fig. 2A; live cell indicated by green rods) and after 180 min gastric incubation (Fig. 2C). Meanwhile, native protein suspensions illustrated an apparent entrapment competence during an identical extrusion process (Fig. 2B); however, gastric exposure generated injured cells (Fig. 2D; dead cells indicated by red rods). Probiotic viability was preserved at the micro-bead periphery after 180 min gastric exposure (Fig. 2C), which suggests the favourable absence of pH gradients during acid conditions. The graph in Fig. 2 illustrates cell viability at each step and endorses the acid-susceptible limitation of native proteins relative to denatured protein micro-beads (Fig. 2C, D). Plate enumeration and CSLM corroborated with FACS analysis, which demonstrated a distinct acid-resistant element of micro-bead functionality. (For interpretation of the references to colour in the text, the reader is referred to the web version of this article.) Gastric contents were characterized (Table 1) prior to treatment analysis. Interestingly, 180 min incubation of probiotic-loaded micro-beads in gastric digesta reduced free amino acid concentrations in gastric contents by 308.9  25.7 mmol mL1 (Fig. 3A). The plethora of amino acid residues in gastric digesta expressed similar magnitudes of reduction after 180 min incubation; an exclusive attribute recognized within micro-bead treatments. Hence, amino acid entrapment at the micro-bead periphery or potential penetration to the core may be possible; a hypothesis attributable to electrostatic and/or hydrophobic interactions. Furthermore, chromatography analysis characterized gastric contents during micro-bead incubation (Fig. 3B) and witnessed the

Fig. 2. Viability of LGGRif within micro-beads ( ) and native protein suspensions ( ) before and after micro-bead extrusion and during subsequent gastric studies. Probiotic viability was investigated after (i) preparation of cell-protein amalgams, (ii) micro-bead extrusion and following (iii) ex vivo porcine gastric incubation for 180 min (pH 2.0, 37  C). Confocal scanning laser microscopy (CSLM) revealed high entrapment efficiency of live LGGRif following micro-bead extrusion (A), with subsequent retention of probiotic viability after 180 min gastric incubation (C). Native protein suspensions illustrated acceptable LGGRif entrapment potential (B) during an identical extrusion procedure; however, cell survival was significantly reduced after 180 min gastric incubation (D). Measurement bars for images A and B represent 50 and 10 mm, respectively, while C and D show 25 mm bars. CSLM analysis incorporated Thiazole Orange (TA) and Propidium Iodide (PI) staining to illustrate the presence of live and dead probiotics. Asterisks (***) indicate a significant difference (p < 0.001) within respective treatments relative to the cell concentration in original mixtures.

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600

60

*

*

500

58

*

400

56 300

*

* 54

200 52

*

100 0

Amino acid concentration (mmol mL-1 of intestinal contents)

Amino acid concentration (µmol mL-1 of gastric contents)

A

37

50 0

1

C

3

6

9

12

Time (min)

B Ch romatographic fr action

50%

40%

*** 30%

***

*** ***

20%

*** ***

10%

0% 0

60 Time (min)

180

Fig. 3. Total amino acid concentration is shown (A) for gastric juice (columns) and intestinal digesta (solid line) as a function of incubation time at 37  C; (B) shows the percent peptide distribution, as measured by size exclusion HPLC, in ex vivo porcine stomach digesta during micro-bead incubation. Peptide fragments were evaluated within the following mass range: >10 kDa ( ), 10e2 kDa ( ) and 2e1 kDa ( ) and <1 kDa ( ). Each peptide mass range is expressed in percent as a function of time with errors bars indicating standard deviation of five independent tests performed in triplicate. Significant differences (*p < 0.05; ***p < 0.001) are illustrated relative to the amino/peptide fraction in the original ex vivo digesta extract (t ¼ 0).

production of a broad size range of peptide fragments; possible remnants of the porcine diet prior to starvation, slaughter and dissection. As gastric incubation proceeded, total peptides > 2 kDa demonstrated a pronounced decline of approx 25% with fragments < 1 kDa expressing 10% average reduction after 180 min. 3.1.1. Atomic force microscopy Atomic force microscopy (AFM) revealed a variety of potential proteineprobiotic cohesive mechanisms in operation within denatured whey protein matrices (Fig. 4). Fabrication of an entrapment network at pH 4.6 with probiotic-protective capacity appeared to be characterized by two distinct features: (i) probiotic camouflage via whey protein layers (Fig. 4A) and (ii) progressive engulfment of Lb. rhamnosus GG by aggregates of globular proteins (Fig. 4B) resulting in comprehensive probiotic entrapment by whey protein particles (Fig. 4C; arrow indicating one probiotic cell). These differing probioticeprotein amalgams were visualized at various locations throughout the protein lattice, which revealed diverse and random protein orientations; an attribute possibly associated with the rapid gelation impetus governing matrix generation (Doherty et al., 2011). These findings correlated with zeta potential analysis, which was performed according to the method outlined by Doherty et al. (2011). In brief, micro-beads were homogenized (Ultra-TurraxÒ T10,

IKAÒ Werke, Germany) for 5 min at ambient temperature and zeta potential was derived from the velocity of the protein suspension under an applied electric field of 150 V. Zeta measurements supported the hypothesis relating to electrostatic interactions within entrapment matrices since the net charge of encapsulation matrices (17.5  0.5 mV) revealed no significant (p < 0.001) change after 28 days storage at room temperature. This finding potentially supports the maintenance of probiotice protein alliance during extended storage. Hence, probiotice protein interactions visualized by AFM appear to be valid components of micro-bead matrices as a function of time and environmental conditions. 3.2. Liberation of entrapped Lb. rhamnosus GG in ex vivo porcine GI contents 3.2.1. Cell enumeration and identification The pH-sensitive release of entrapped LGGRif was evaluated during gastric (Fig. 5A) and intestinal (Fig. 5B) incubation at 37  C for 3 h. No probiotic bacteria were released from micro-beads during gastric incubation at pH 3.4; however, trivial concentrations were liberated after 3 h at pH 2.4 and 2.0 (0.2 and 0.5  0.1 log10 cfu mL1, respectively). Moreover, entrapment preserved cell viability after 180 min gastric incubation at pH 2.4 and 2.0 to achieve maximum

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Fig. 4. Visualization of the mechanism of interaction operating within the micro-bead system during probiotic entrapment using atomic force microscopy (AFM). Proteineprobiotic matrices illustrated a range of interactions using 2 mm magnification range. AFM analysis suggested the envelopment of LGGRif by whey protein strata or layers (A), while partial (B) and comprehensive (C) entrapment by whey protein aggregates appeared to be permanent features of proteineprobiotic systems.

LGGRif liberation of 8.9  0.1 log10 cfu mL1 in intestinal contents. Intestinal incubation liberated 6.2  0.1 log10 cfu mL1 after 5 min, while complete cell liberation was achieved following 30 min incubation (Fig. 5B). Random intestinal isolates from MRSRif plates generated the RAPD-PCR profile (Fig. 5C) confirming the selective enumeration of the administered probiotic strain during ex vivo studies. Random colonies from MRSRif plates unveiled nine macro restriction patterns, representing the predicted brand pattern of Lb. rhamnosus GG. Following intestinal liberation, probiotic cells remained vulnerable to the adverse effects of bile and various indigenous intestinal components. However, FACS analysis validated the retention of intact, functional cell membranes after 180 min (Fig. 5D; gate A3) since this methodology was capable of differentiating Lb. rhamnosus GG and LGGRif from background intestinal microbiota. Interestingly, micro-beads incubated in phosphate buffer saline (PBS) (pH 7) under identical conditions failed to

illustrate any signs of degradation after 180 min incubation. Hence, micro-bead degradation responded synergistically to neutral pH and intestinal enzymatic action, which reflects true in vivo scenarios. 3.2.2. Cell-surface hydrophobicity In the selection of probiotic strains with beneficial health effects, adhesion to intestinal mucus is a fundamental criterion for new probiotic strains (Vinderola, Matar, & Perdigon, 2005). Attachment of cells to the intestinal wall is dependent on various phenotypic cellsurface properties including hydrophobicity, which is commonly assessed by microbial adhesion to hydrocarbons (MATH). Hence, the adhesion capacity of fresh Lb. rhamnosus GG and LGGRif to organic solvents elucidated the maintenance of cell-surface hydrophobicity following (i) the generation of a rifampicin-resistant derivative, LGGRif, (ii) micro-bead extrusion and (iii) ex vivo gastro-intestinal

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Fig. 5. Release of viable LGGRif from micro-beads during (A) ex vivo gastric incubation at pH 3.4 (C), pH 2.4 (,) and pH 2.0 (:) and (B) ex vivo intestinal incubation at pH 6.6 (A) for up to 180 min at 37  C. RAPD-PCR analysis (C) further confirmed the presence of LGGRif in random gastric, intestinal and caecum isolates from MRSRif plates of micro-bead treatments. Lanes 1 and 15 contain a molecular ladder (100e1527 bp); Lane 2 ¼ LGG, Lane 3 ¼ LGGRif, Lanes 4e7 ¼ isolates from gastric incubation (180 min); Lanes 8e11 ¼ isolates from intestinal incubation; Lanes 12e14 ¼ caecum isolates. FACS analysis (D) confirmed the maintenance of Lb. rhamnosus GG viability and functionality (gate A3) following stomach incubation (180 min, pH 2.0, 37  C) and subsequent intestinal digestion (180 min, pH 6.6, 37  C).

delivery. Relative to fresh stationary phase cultures, entrapped Lb. rhamnosus GG and LGGRif retained identical and specific adhesion characteristics following cell release from micro-beads (Fig. 6A). Entrapped probiotic populations expressed no significant difference relative to fresh cells, for any hydrocarbon tested, which demonstrated the persistence of hydrophilic, basic cell-surface attributes of LGG following gastric resistance and intestinal liberation. Average hydrophobicity values for fresh and encapsulated LGGRif demonstrated a robust attraction for chloroform (approximately 22%; Fig. 6A i) and weak interaction with n-hexadecane (approximately 1%; Fig. 6A ii) with standard deviations remaining below 1%. This hydrophilic character maybe affiliated with the presence of exopolysaccharide (EPS) produced by Lb. rhamnosus GG since Landersjo, Yang, Huttunen, and Widmalm (2002) identified an EPS containing galactose, rhamnose and N-acetylglucosamine in a molar ratio of 4:1:1. 3.2.3. Micro-bead surface hydrophobicity The surface hydrophobicity (SH) of whey protein micro-beads was determined according to Kato et al. (1984) with slight modifications applied for micro-bead matrices. Anionic SDS molecules were bound to solvent exposed, hydrophobic amino acid residues and quantified spectrophotometrically. The resulting SH values of whey protein micro-beads, heat-treated and native WPI solutions are illustrated in Fig. 6B. In agreement with previous findings (Hiller & Lorenzen, 2008), low SH values were observed for native WPI solutions; which was subsequently maintained during 3 h ex vivo stomach incubation. On the contrary, heat-treated WPI illustrated high SH values, both in solution and in micro-bead matrices, values which declined as a function of ex vivo gastric incubation time. Relative to heat-treated solutions, micro-bead matrices accelerated the reduction of SH by 7.6 SDS units after 3 h in stomach conditions. These results correlate with Hiller and Lorenzen (2008), who demonstrated that heat-treatment increases SH of whey protein isolate. Moreover, these data endorsed the potential existence of hydrophobic amino acid residues on the micro-bead surface, which accelerated interfacial adsorption behaviour of micro-beads during stomach incubation. 3.2.4. Confocal microscopy Following gastric incubation ranging from pH 3.4 to 2.0, microbeads were subsequently washed and resuspended in intestinal

contents (pH 7.2) at 37  C and CSLM analysis (Fig. 7A) visualized micro-bead integrity and degradation as a function of GI incubation time. Protein matrices remained intact following 180 min gastric incubation (Fig. 7A i), with concomitant maintenance of cell viability and functionality, as determined by plate enumeration and FACS analysis, respectively. Matrix biodegradation was initiated after 5 min intestinal exposure (Fig. 7A ii) with probiotic cells progressively discharged from the protein milieu. Protein fragments and aggregates were the initial products of matrix digestion; however, significant cell concentrations remained lodged within the aggregate core. After 15 min, micro-bead digestion progressed from the micro-bead periphery to the core with concomitant liberation of live LGGRif after 30 min (Fig. 7A iii), demonstrating the transition of micro-beads to malleable protein suspensions. Fig. 7A supports plate enumeration and FACS analysis, which confirmed complete intestinal release of viable functional LGGRif populations after 30 min. 3.2.5. Chromatography SEC during intestinal incubation illustrated the acceleration and timely release of proteins, aggregates and peptides (Fig. 7B) during 180 min intestinal incubation. Micro-bead disintegration via proteolytic digestion lead to the release of protein aggregates within the size range 6e67 kDa during initial 60 min incubation (Fig. 7B; black line). Following this, peptides predominantly < 1 kDa, were generated after 180 min incubation (Fig. 7B; blue line) with concomitant disappearance of b-Lg, the principle whey protein. Furthermore, SEC revealed the indigenous amino acid profile of intestinal digesta (Fig. 7B; red line), corresponding to the baseline level following initiation of the intestinal assay. Interestingly, no peptides were released during micro-bead incubation in PBS. 4. Discussion The maintenance of GI homeostasis is considered critical for prevention and development of immune-mediated and metabolicrelated diseases. The positive influence of probiotics on gut homeostasis is achieved by cell interaction with the host, bacterial antagonism and immunomodulation, which may help the healthy host to maintain a ‘physiological state’ of control over inflammatory,

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Fig. 6. Adhesion (A) of LGGRif to hydrocarbons as fresh stationary phase cultures ( ) relative to cell adhesion following ex vivo intestinal incubation ( ) using chloroform, nhexadecane, hexane and diethyl ether. Cell ‘clearing’ was visualized in sample supernatant as a result of (i) LGGRif adhesion to chloroform compared with (ii) cell retention in supernatant in the presence of n-hexadecane. Surface hydrophobicity (SH) of micro-bead batches (B) ( ), heat-treated whey protein ( ) and native ( ) whey protein solutions as effected by ex vivo gastric incubation (pH 2.0, 3 h, 37  C). Vertical bars represent standard deviations from three independent studies performed in triplicate with SH units defined as mg SDS (500 mg WPI)1.

infectious and immunological reactions (Lebeer, Vanderleyden, & De Keersmaecker, 2010). Next to health-promoting characteristics, functional aspects of probiotic bacteria also involve their capacity to reach the colon in a metabolically active state. However, given the recent concerns regarding probiotic delivery and viability in the intestine, this study elucidates probiotic entrapment in whey protein micro-beads as a delivery mechanism capable of transferring functional cell consignments to their target site. Hence, entrapment stability of rifampicin-resistant derivative of Lb. rhamnosus GG (LGGRif) was assessed using ex vivo GI contents to reflect the range of obstacles encountered during in vivo situations. Design of micro-beads for intestinal cell delivery was achieved using ionotropic gelation, while peptic resistance and intestinal delivery of LGGRif in micro-beads, native protein, and free-cell treatments were screened in GI digesta procured from finisher pigs. This approach differentiated probiotic viability pertaining to the presence of whey proteins alone or micro-bead structural effects. Gastric contents varied from pH 3.4 to 2.0, possibly due to (i) the buffering affect of the animal diet or (ii) the adverse effect of pepsin upon stomach cells at acidic pH. Although gastric lumen

has a pH of approx 2.0, the effect of proton release and neutralization by glycoproteins collectively influence the pH gradient in stomach digesta (Campos & Sancho, 2003). Hence, ex vivo studies provide a suitable environment for evaluation of probiotic encapsulation stability during conditions that resemble the stomach environment. Cell survival was assessed via selective plate enumeration, RAPD-PCR, FACS, CSLM and AFM analysis. As a general tendency, free cells demonstrated high susceptibility to low pH and pepsin activity with no apparent ability to survive ex vivo conditions at/below pH 3.4 in the presence of 44.5 IU units of pepsin activity. In contrast to the immediate mortality-motive of free LGGRif, entrapment demonstrated a pronounced augmentation of probiotic acid resistance and continuously supported the metabolic activity of stationary phase cultures during gastric incubation. Research has demonstrated that cellular or culture components of dead probiotic bacteria potentially mediate beneficial effects on the host, in addition to viable metabolizing cells (Ouwehand & Salminen, 1998). In this regard, a validated FACS methodology (Doherty et al., 2010b) was utilized in collaboration

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Fig. 7. Gastric tolerance and progressive dissolution of micro-beads (A) during ex vivo incubation. Micro-bead integrity was maintained following 180 min stomach incubation at pH 2.0 (i), while micro-bead disintegration was initiated after 5 min intestinal (pH 6.6) incubation (ii) with complete cell liberation visualized after 30 min (iii); bar represents 250 mm. Protein and peptide release from whey protein micro-beads is shown in (B), as measured by size exclusion HPLC, after 60 min (black line) and 180 min (blue line) ex vivo intestinal incubation at 37  C. Trace amounts of peptides were identified in the extracted intestinal digesta prior to micro-bead addition and thus represent the baseline reference (red line). The standard curve of protein/peptide standards (A) indicated the molecular masses of eluting ex vivo samples. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

with plate count techniques to evaluate the retention of cell functionality and viability following ex vivo incubation. Regardless of protein structural diversity between treatment groups, FACS analysis complemented plate enumeration via differentiation of cell populations into live, injured and dead sub-groups, providing a real-time assessment of the progressive cell death, injury and viability in free, native protein and entrapped treatments, respectively. Meanwhile, CSLM provided an additional visual aid, which confirmed the maintenance of micro-bead morphology, in addition to the absence of pH gradients during gastric incubation. Micro-bead acid resistance did not appear to be adversely affected by their cratered, porous surface features. In relation to alginate matrices (Krasaekoopt, Bhandari, & Deeth, 2003), whey protein micro-beads represent a favourable matrix material due to their peptic resistance and rigidity during stomach incubation. Moreover, AFM analysis revealed probiotic cell membranes camouflaged by whey protein strata in addition to partial and/comprehensive LGGRif entrapment by globular

protein aggregates. This cohesive attraction between matrix components (Fig. 4) appeared to be a permanent resident within micro-beads potentially free of pH-gradient formation due to the homogenous retention of cell viability throughout the matrix during gastric incubation (CSLM images in Fig. 1). During this study, CSLM validated cell viability and micro-bead integrity following micro-bead extrusion with subsequent gastric incubation (Fig. 2C). Plate enumeration, CSLM and FACS analysis synergistically demonstrated the maintenance of probiotic viability and functionality following gastric incubation, which amplifies the proficiency of the entrapment procedure during challenging conditions. During micro-bead incubation, the surrounding ex vivo gastric media illustrated a time-dependent reduction in free amino acid concentrations, with all amino acid residues demonstrating similar reductions. Since GI extracts expressed a plethora of amino acids with a broad profile of pI values, electrostatic and hydrophobic interactions are both considered interactive mechanisms

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associated with potential amino acid binding to micro-beads. b-Lg present in micro-beads, is known to be a carrier of hydrophobic molecules, particularly retinol (Fox & McSweeney, 2003) and resveratrol from grapes (Liang, Tajmir-Riahi, & Subirade, 2008). Surface hydrophobicity (SH) illustrated a dynamic approach for the investigation of physico-chemical characteristics of microbeads that are sensitive to polarity and hydrophobic entities in the surrounding environment. Hence, binding of hydrophobic compounds to b-Lg can be illustrated via changes in the surface hydrophobicity of micro-beads. SH is defined based on amino acid composition (Bigelow, 1967); however it disregards secondary, tertiary and quaternary structure. Thus, effective hydrophobicity of exposed amino acids residues on the micro-bead periphery may potentially mediate their interfacial adsorption behaviour, particularly during gastric incubation. High SH of heat-treated whey proteins deciphers a strong micro-bead binding capacity; a characteristic associated with heat denaturation of b-Lg (Shpigelman, Israeli, & Livney, 2009). This finding is in agreement with previous work (Moro, Gatti, & Delorenzi, 2001) that demonstrated high SH values for bovine serum albumin and b-Lg, which corresponds to their biological function of transporting surface-bound hydrophobic molecules. The weak SH of native WPI was forecast since the compact globular structure of native protein is frequently associated with low SH (Wagner & Gueguen, 1999). Heat-treated protein suspensions and micro-bead matrices exhibit pronounced SH differentiation, which may reflect stronger hydrophobic interactions between free amino acid residues e originating from gastric media e and unfolded, open micro-bead matrices. Thus, SH reveals potential evidence to reconcile microbead binding capacity and the camouflage of amino acid residues during gastric incubation. Whey protein micro-beads illustrated a dynamic synergism linking pH and enzymatic action, which created optimum microbead functionality via stomach integrity with concomitant encouragement of intestinal proteolytic action. As previously demonstrated by protein systems (Remondetto, Beyssac, & Subirade, 2004), probiotic release from whey protein micro-beads is biphasic, with an initial relatively fast release rate (5 min) representing the proportion of cells entrapped on the micro-bead periphery, followed by slower sustained liberation (after 30 min) associated with retarded liberation of LGGRif enclosed within the micro-bead core. Moreover, characterization of Lb. rhamnosus GG as a presumptive probiotic is based on its ability to persist in the intestinal lumen and epithelium via retention of cell-adhesive capacity post-gastric incubation. Histology was not within the scope of this study; however, FACS analysis and hydrophobicity characterization e a phenotype related to cell-adhesive capacity e both validated the retention of viable, functional and hydrophilic characteristics typical of stationary phase Lb. rhamnosus GG and LGGRif. Taking into consideration that colonization of the GI tract with lactobacilli interferes with colonization of enteropathogenic micro-organisms, the observed retention of probiotic functionality advocates the maintenance and expression of health benefits through competition with Gram-negative pathogens for adhesion sites (Vinderola et al., 2005). To facilitate comparison of enzymatic reactions during ex vivo studies, it was important to consider enzyme activity in the stomach (pepsin) and intestine (trypsin and chymotrypsin). Following microbead transit from stomach to intestinal media, enzyme penetration of the matrix structure occurred within 30 min with subsequent release of the quasi-totality of encapsulated LGGRif. It is noteworthy that bile salts are toxic for many living cells since they can disorganize the structure of the cell membrane and bile salt tolerance is considered another essential property required for probiotic survival in the intestine (Succi et al., 2005); however, extension of intestinal

incubation from 3 to 12 h validated the retention of LGGRif viability and functionality in the presence of bile, generating a FACS result identical to Fig. 5D. Hence, specific probiotic delivery to the intestine in gelled whey protein micro-beads is significantly more effective than cell suspensions in native WPI. Sequential gastric and intestinal ex vivo incubation demonstrated excellent sentinel properties for protein micro-beads with quasi-survival rates of previous in vitro studies (Doherty et al., 2011). Moreover, the release of essential amino acids from whey proteins micro-beads during intestinal proteolysis may enhance the nutritional quality of micro-beads as cell delivery vehicles. These attributes may be useful for site-specific controlled biomolecule delivery with auxiliary promotion of their intestinal absorption. This strategy is widely used in the pharmaceutical field and could find a broad spectrum of applications in the development of innovative bioactive foods. Cell entrapment in whey protein micro-beads epitomizes an interesting alternative to spray-drying due to the high biological value of whey proteins (Smithers, 2008). In contrast to cold-renneting of skim milkconcentrates (Heidebach, Först, & Kulozik, 2009), acetateinduced cross-links generated protein networks with vulnerable hydrophobic patches buried within the globular whey matrix. Hence, micro-beads were fabricated with high resistance to dissolution in enzyme-active environments. Other studies have demonstrated the protective properties associated with spraydrying (Bielecka & Majkowska, 2000) and encapsulation in milk proteins (Ainsley Reid et al., 2007; Beaulieu et al., 2002); however, the present study illustrates micro-beads with sufficient strength to defy ex vivo challenges with concomitant exploitation of the nutritional and functional properties of whey protein for expansion of the bioactive and cell delivery market. A few studies have been performed on cell growth and survival during entrapment in alginate and more rarely in whey protein and no detailed information was specified relating to proteolytic activity, cell functionality or amino acid release during cell liberation. Hence, this study gained specific insights into the structural arrangement post-extrusion, gastric and intestinal incubation using chromatography and image analysis to complement traditional culture techniques. 5. Conclusion Encapsulation matrices were created from denatured whey proteins using an entrapment process devoid of high temperatures, shear forces and cell loss. This dense whey protein gel lattice was capable of offering a micro-milieu for favourable cell entrapment during gastric incubation, with auxiliary fragmentation within intestinal environments. The unique functional properties of whey proteins alleviate the problems associated with capsule size, which is highly important regarding the sensory impact of matrices on the final food products. Hence, it can be concluded that the use of gelled protein matrices is a promising strategy to render milk proteins as valuable entrapment materials for targeted delivery of bioactive compounds in food, beverage and pharmaceutical applications. Acknowledgements The support provided by the National Food Imaging Centre and the advise of John O’Callaghan is gratefully acknowledged. The work was funded by the Irish Dairy Research Trust, the National Development Plan 2007e2013 and Science Foundation Ireland (SFI). S. B. Doherty was funded by the Irish Dairy Research Trust under the Teagasc Walsh Fellowship Scheme.

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