Microbial cells as colloidal particles: Pickering oil-in-water emulsions stabilized by bacteria and yeast

Food Research International 81 (2016) 66–73

Contents lists available at ScienceDirect

Food Research International journal homepage: www.elsevier.com/locate/foodres

Microbial cells as colloidal particles: Pickering oil-in-water emulsions stabilized by bacteria and yeast Hassan Firoozmand, Dérick Rousseau ⁎ Department of Chemistry and Biology, Ryerson University, 350 Victoria St., Toronto, Ontario M5B 2K3, Canada

a r t i c l e

i n f o

Article history: Received 24 July 2015 Received in revised form 15 October 2015 Accepted 19 October 2015 Available online 19 October 2015 Keywords: Pickering emulsions Particles Interfacial adsorption High internal phase emulsions Lactic acid bacteria Yeast

a b s t r a c t Thermally-inactivated baker's yeast (Saccharomyces cerevisiae) and lactic acid bacteria (Lactobacillus acidophilus and Streptococcus thermophilus) were used to generate and stabilize model oil-in-water (O/W) emulsions containing up to 80 wt.% dispersed oil. With optimized compositions, cell-covered dispersed oil droplets were stable against droplet coalescence and bulk phase separation for over four months. From a textural perspective, these emulsions were self-supporting and exhibited a mayonnaise-like consistency. The microbial cells acted as Pickering-type stabilizers by residing at the oil–water interface. The three-phase contact angle of the yeast at the oil–water interface measured using confocal microscopy was 30 ± 9°, demonstrating its ability to stabilize O/W emulsions. These microbial cells may be used in the design of processed food emulsions with an ‘all-natural’ designation as well as for the replacement of common synthetic surfactants to permit clean label declarations. Published by Elsevier Ltd.

1. Introduction Emulsions used in many applications (e.g., chemicals, food, pharmaceuticals, cosmetics and agrochemicals) consist of two or more mutually-insoluble liquids that invariably strive to lower their free energy via bulk phase separation. Oil-in-water (O/W) emulsions are usually stabilized by surface-active species such as proteins or small-molecule surfactants that lower interfacial tension or thickeners that increase the viscosity of the continuous phase. By contrast, Pickering emulsions are stabilized by insoluble solid particles (Pickering, 1907) that often confer superior stability against coalescence compared to molecular surfaceactive agents (Dickinson, 2009). The stabilization mechanism of such emulsions is based on the formation of a steric barrier by interfaciallylodged particles that have a desorption energy orders of magnitude above kT (Dickinson, 2010). Due to biocompatibility and biodegradability requirements for food applications (Lam, Velikov, & Velev, 2014), only a limited number of food-grade Pickering particles have been explored (de Folter, van Ruijven, & Velikov, 2012; Dickinson, 2010; Liu & Tang, 2013; Yusoff & Murray, 2011), namely those based on fat (Poortinga, 2008), βlactoglobulin (Nguyen, Nicolai, & Benyahia, 2013), zein protein (de Folter et al., 2012), hydrophobic cellulose (Wege, Kim, Paunov, Zhong, & Velev, 2008), soy protein (Liu & Tang, 2013), quinoa starch (Rayner, ⁎ Corresponding author. E-mail address: [email protected] (D. Rousseau).

http://dx.doi.org/10.1016/j.foodres.2015.10.018 0963-9969/Published by Elsevier Ltd.

Timgren, Sjöö, & Dejmek, 2012) and whey protein microgels (Destribats, Rouvet, Gehin-Delval, Schmitt, & Binks, 2014). Most of these suffer from key limitations, e.g., fat and starch-based particles may lose their functionality at typical processing temperatures, proteinbased particles can typically be made on a lab-scale only whereas hydrophobic cellulose particles have a relatively broad particle size distribution. As well, production of colloidal particles may be very energy-intensive and expensive or may be too complex to implement in industry settings (Velikov & Pelan, 2008). Previous efforts have shown that certain single-celled microorganisms can disperse oil in water and stabilize O/W emulsions. Decades ago, Srivasta et al. inadvertently formed yeast-stabilized Pickering emulsions while using a mixture of petroleum hydrocarbons, water and mineral salts as a growth medium for feed production, (Srivasta, Singh, Baruah, Krishna, & Iyengar, 1970). This ability to reside at the petroleum hydrocarbon-water interface and stabilize O/W emulsions was also later reported (Dorobantu, Yeung, Foght, & Gray, 2004; Rosenberg & Rosenberg, 1985). More recently, microbial cells have been shown to stabilize O/W emulsions and produce biosurfactants during microbial production of fuels (Balat & Balat, 2010; Heeres, Picone, van der Wielen, Cunha, & Cuellar, 2014; Lim & Teong, 2010; Lin, Cunshan, Vittayapadung, Xiangqian, & Mingdong, 2011; Yusuf, Kamarudin, & Yaakub, 2011), resulting in emulsions resistant to heat treatments and extensive centrifugation (Furtado, Picone, Cuellar, & Cunha, 2015). A number of single-celled microorganisms with the aid of chitosan (Wongkongkatep et al., 2012) and polymerized oleic acid (Zhu et al.,

H. Firoozmand, D. Rousseau / Food Research International 81 (2016) 66–73

2014) have been shown to act as stabilizers in Pickering oil-in-water emulsions. As robust, readily available food-grade micron-sized particles are either scarce, difficult to produce, restricted by legislative regulations and/ or are limited to small/lab-scale applications, we explored yeast and lactic bacteria as the foundation for the production of food-grade Pickering emulsion particles. Saccharomyces cerevisiae is a round/oval yeast b10 μm in diameter important in the baking, winemaking and brewing industries (Randez-Gil, Cocoles-Saz, & Prieto, 2013). Lactobacillus acidophilus is a rod-shaped bacterium measuring b5 μm in length and Streptococcus thermophilus exists as round-shaped cells b1 μm in diameter. Both bacteria are used as starter cultures in fermented dairy products (Carr, Chill, & Maida, 2002). The purpose of this study was to exploit thermally-inactivated S. cerevisiae, L. acidophilus and S. thermophilus as Pickering stabilizers in food-grade O/W emulsions. 2. Materials and methods 2.1. Materials Baker's yeast (S. cerevisiae) (20 billion CFU/g) was supplied by Lesaffre, (Red Star brand) (Milwaukee, WI, USA). L. acidophilus (200 billion CFU/g) and S. thermophilus (400 billion CFU/g) were purchased from Custom Probiotics (Glendale, CA, USA). The cells were received in dry form and used as-is. Distilled water was used throughout. Extra virgin olive oil was purchased from a local supermarket.

67

glass bottles were then left at room temperature to cool prior to centrifugation. In each case, the supernatant was discarded and the sediment was transferred to a 50 ml centrifuge tube and washed 5–7 times with ~30 ml of distilled water by repeated resuspension/centrifugation cycles (5 min for yeast and 30 min for the bacteria, respectively, at 3500 rpm). After the final round of centrifugation, a clear transparent supernatant (water) and homogeneous, uniform cell-containing sediment were obtained.

2.3. Mixing procedure As emulsion formation resulted in increased viscosity, this limited the use of high-pressure valve homogenization for their preparation. We instead used vortexing or impeller-type mixing. Only emulsions containing 50–50 wt.% oil and aqueous phase (i.e., Fig. 1) were prepared via vortexing. With these, the entire aqueous phase (water + cells, total 5 g) was placed in a vial and the oil phase was added in ~1 g increments followed by 20 s of vortexing at 3000 rpm until all of the oil (5 g) was incorporated (total weight 10 g). All other emulsions (Figs. 2-9) were prepared with magnetic stirring, where the entire aqueous phase (water + cells) was placed in a vial and the oil phase was added drop-wise at a rate of ~1 g per 90–100 s to the aqueous phase (starting water + cell volume: 2–6 g) at ~500 rpm. Mixing was continued until the entire oil phase was incorporated (total weight 10 g). Final emulsion compositions are shown in Table 1. All experiments were performed in at least triplicates.

2.2. Cell preparation

2.4. Viscosity

Cell treatment is optional as it is possible to use live (active) cells for emulsion stabilization. However, to prevent their biological activity, the following inactivation protocol was applied. Cells of S. cerevisiae, L. acidophilus or S. thermophilus were suspended in distilled water at 10 wt.% in sealed screw cap glass bottles, placed in a waterbath at 95 °C for 30 min and vigorously shaken every 5 min for 15–20 s. The

A Paar Physica MCR 301 rheometer (Anton Paar GmbH, Graz, Austria) equipped with a Peltier temperature control unit (P-PTD 200) was used to determine apparent viscosity. All measurements were carried out using a parallel plate geometry (PP 25/TG; diam. 25 mm) and an operating gap of 200 μm. For each measurement, each emulsion sample was first pre-sheared at a constant shear rate of 1 s−1 for 60 s and

Fig. 1. The effect of added microbial cells on bulk phase behaviour of 50 wt.% oil and 41–49 wt.% water: (a) 1–9 wt.% S. thermophilus; (b) 1–8 wt.% L. acidophilus, and (c) 1–9 wt.% S. cerevisiae. Sample ‘0’ in (a & c) consists of 50 wt.% water + 50 wt.% oil with no added cells (control). Emulsions were prepared with a vortex mixer.

68

H. Firoozmand, D. Rousseau / Food Research International 81 (2016) 66–73

Fig. 2. CLSM images of single oil droplets in O/W emulsions prepared with (a) S. thermophilus, (b) L. acidophilus and (c) S. cerevisiae.

then allowed to rest for 2 min on the rheometer stage. Apparent viscosity was measured at 25 °C from 0.01 to 100 s−1. 2.5. Droplet/particle size distribution Laser light scattering experiments were conducted with a Malvern Mastersizer 2000 with a Hydro 2000S wet cell attachment (Malvern Instruments, Worcestershire, UK). Samples were dispersed into the instrument's sample cell until an obscuration level of 13–15% was reached. A refractive index of 1.469 for olive oil was used. Results were analysed with the Malvern Mastersizer 2000 software v.5.54. Volumeweighted droplet sizes [D4,3] and representative droplet size distributions (DSDs) are reported. These same emulsions were subjected to rheological measurements.

set at 505 nm. For the colour images, the emission spectra were collected at 505 nm for the microbial cells and 650 nm for the oil. No labelling was required for the cell-containing samples as autofluorescence of the cells and olive oil was sufficient for CLSM image acquisition. All CLSM images were obtained from original undiluted emulsions. For characterization, 10×, 20×, and 63× objectives were used. Images were recorded at 25 °C at a resolution of 1024 × 1024 pixels. Image optimisation was performed using the LSM 510's built-in image analysis software. ImageJ software was used to determine oil–cell–water contact angles (θ), measure the cross-sectional area of the oil droplets and calculate droplet coverage by the cells. Images shown herein are representative of the emulsion morphology seen for a given composition. 3. Results

2.6. Confocal laser scanning microscopy

3.1. Preliminary assessment of microbial cell emulsification capacity

Confocal laser scanning microscopy (CLSM) was performed using an upright Zeiss LSM 510 (Carl Zeiss, Toronto, ON, Canada). The CLSM was operated in fluorescent mode with an Ar laser source (488 nm). For greyscale images, the emission spectra were collected with 1 channel

Fig. 1 shows the phase behaviour of oil–water mixtures as a function of added microbes (wt.%). All images were taken immediately after emulsification. Sample ‘0’ in Fig. 1a and Fig. 1c represents the 50 wt.% water + 50 wt.% oil control with no added cells. It was only with

Fig. 3. i: Phase behaviour of oil–water mixtures consisting of 2 wt.% S. thermophilus and 40–80 wt.% oil and 18–58 wt.% water: (a) 40 wt.% oil + 58 wt.% water; (b) 50 wt.% oil + 48 wt.% water; (c) 60 wt.% oil + 38 wt.% water; (d) 70 wt.% oil + 28 wt.% water; (e) 80 wt.% oil + 18 wt.% water; ii: physical appearance of the emulsion consisting of 80 wt.% oil + 18 wt.% water + 2 wt.% S. thermophilus scooped from vial e; iii: microstructure of the O/W emulsion consisting of 80 wt.% oil + 18 wt.% water + 2 wt.% S. thermophilus scooped from vial e.

Fig. 4. i: Phase behaviour of oil–water mixtures consisting of 2 wt.% L. acidophilus and 40–75 wt.% oil and 23–58 wt.% water: (a) 40 wt.% oil + 58 wt.% water, (b) 50 wt.% oil + 48 wt.% water, (c) 60 wt.% oil + 38 wt.% water, (d) 70 wt.% oil + 28 wt.% water, and (e) 75 wt.% oil + 23 wt.% water; ii: physical appearance of the emulsion consisting of 75 wt.% oil + 23 wt.% water + 2 wt.% L. acidophilus scooped from vial e; iii: microstructure of the O/W emulsion consisting of 75 wt.% oil + 23 wt.% water + 2 wt.% L. acidophilus scooped from vial e.

H. Firoozmand, D. Rousseau / Food Research International 81 (2016) 66–73

Fig. 5. Visual appearance of yeast-stabilized O/W emulsions with different compositions: (a) 60 wt.% oil + 31 wt.% water + 9 wt.% S. cerevisiae; (b) 65 wt.% oil + 27 wt.% water + 8 wt.% S. cerevisiae and (c) 70 wt.% oil + 23 wt.% water + 7 wt.% S. cerevisiae.

8–9 wt.% added S. thermophilus cells that the aqueous phase volume decreased slightly, suggesting limited emulsification by this bacterium (Fig. 1a). Similar results were observed with ≤ 7 wt.% added L. acidophilus whereas at 8 wt.%, a homogeneous one-phase emulsion resulted (Fig. 1b). With yeast, a gradual increase in cell concentration led to a systematic reduction in the bulk aqueous phase volume such that at 9 wt.%, a homogeneous, though coarse, one-phase emulsion was apparent (Fig. 1c). Under the present experimental conditions, the yeast demonstrated the most favourable emulsification capacity followed by L. acidophilus with S. thermophilus showing little ability to emulsify. 3.2. Droplet coverage at constant cell content Fig. 2 shows the CLSM cross-sections of similarly-sized oil droplets with interfacially-adsorbed microbes. Compositions of 50 wt.% water, 48 wt.% oil and 2 wt.% cells provided a clear indication of the behaviour of each microbe at the interface and resulted in large droplets (N 100 μm in diameter). Other than the obvious size difference between the bacteria and yeast, there also existed a clear distinction in droplet surface coverage. S. thermophilus yielded the least effective interfacial adsorption (Fig. 2a circles) with a third of the droplet exterior consisting of bald patches. By contrast, droplet coverage with L. acidophilus was N90% with little evidence of exposed areas at the oil–water interface (Fig. 2b). S. cerevisiae demonstrated a behaviour intermediate to the two bacteria, with a fifth of the droplet surface lacking yeast (Fig. 2c). Differences in aggregation behaviour were apparent, with S. thermophilus demonstrating

69

Fig. 7. Droplet size distributions of emulsions demonstrating long-term kinetic stability: ○ — 2 wt.% S. thermophilus + 80 wt.% oil + 18 wt.% water; ✰ — 2 wt.% L. acidophilus + 75 wt.% oil + 23 wt.% water, and; Δ — 9 wt.% yeast + 60 wt.% oil + 31 wt.% water.

extensive self-aggregation and limited adsorption (Fig. 2a rectangles) compared to L. acidophilus and S. cerevisiae, which were both organized as well-ordered mono- and multilayers. 3.3. Microbial cell emulsion formation and stabilization Fig. 3i shows the ability of S. thermophilus (2 wt.%) to emulsify various mixtures of oil (40–80 wt.%) and water (18–58 wt.%) prepared via magnetic stirring. Samples with 40–70 wt.% oil (vials a–d) existed as two-phase systems with the aqueous subnatant gradually decreasing in volume fraction with an increase in bacteria concentration. Presence of more oil (i.e., 80 wt.% oil + 18 wt.% water) (vial e) resulted in a scoopable one-phase emulsion that resembled a mayonnaise-like gel that could support its own weight (Fig. 3ii). Microstructurally, a closepacked arrangement of oil droplets in a highly-disordered polyhedral array was present, given the high proportion of oil (Fig. 3iii). This emulsion was shelf-stable and did not demonstrate syneresis upwards of four months. Fig. 4i shows the effect of oil content (40–75 wt.%) on emulsion formation with 2 wt.% L. acidophilus. Two-phase systems existed with 40–70 wt.% oil (vials a–d) whereas presence of 75 wt.% oil (vial e) resulted in a scoopable, gel-like one-phase emulsion that demonstrated limited syneresis when scooped (Fig. 4ii). Given its high proportion, this emulsion also revealed a close-packed arrangement (Fig. 4iii), though not as tightly packed as per S. thermophilus, based on the lack of a polyhedral arrangement. At rest, this emulsion was shelf-stable and did not demonstrate any syneresis upwards of four months. However, sampling from the emulsion did result in localized release of the aqueous phase, likely as a result of gel breakdown.

Fig. 6. CLSM of yeast-stabilized O/W emulsions of different compositions: (a) 60 wt.% oil + 31 wt.% water + 9 wt.% S. cerevisiae; (b) 65 wt.% oil + 27 wt.% water + 8 wt.% S. cerevisiae; (c) 70 wt.% oil + 23 wt.% water + 7 wt.% S. cerevisiae.

70

H. Firoozmand, D. Rousseau / Food Research International 81 (2016) 66–73

proportion of oil could also be stabilized with a high proportion of added cells. The key observations from the observations in Sections 3.1–3.3 were as follows: i) irrespective of composition, these three microbial cells acted as Pickering particles; ii) a higher proportion of cells resulted in emulsions with an enhanced kinetic stability and iii) emulsion formation and stabilization was highly-dependent on processing, e.g., with vortexing S. thermophilus cells were not as effective as with magnetic stirring. 3.4. Droplet size distribution of cell-stabilized emulsions

Fig. 8. Viscosity vs. shear rate profile of emulsions of emulsions demonstrating longterm kinetic stability: ○ — 2 wt.% S. thermophilus + 80 wt.% oil + 18 wt.% water; ✰ — 2 wt.% L. acidophilus + 75 wt.% oil + 23 wt.% water, and; Δ — 9 wt.% yeast + 60 wt.% oil + 31 wt.% water.

Fig. 5 presents three S. cerevisiae-containing O/W emulsions with different compositions. All emulsions were kinetically-stable at rest and did not demonstrate visible phase separation or syneresis when sampled. Somewhat paradoxically, in the emulsion with a lower oil content (i.e., 60 wt.%) (Fig. 5a), a higher yeast concentration (9 wt.%) was required to produce a kinetically-stable homogeneous emulsion. By contrast, with 65 wt.% and 70 wt.% oil, 8 wt.% and 7 wt.% yeast, respectively, were required to produce single-phase emulsions stable against phase separation (Fig. 5b and c). When viewed using CLSM, the size range of the emulsified oil droplets widened with a decrease in cell content and increase in oil content (Fig. 6). With 60 wt.% oil, 31 wt.% water and 9 wt.% S. cerevisiae, the oil droplet diameter range was 10–100 μm (Fig. 6a) whereas with 65–70 wt.% oil and 8–9 wt.% S. cerevisiae, oil droplet diameters ranged from 10 to 250 μm (Fig. 6b and c). Similar observations were recorded with bacteria-stabilized emulsions having comparable compositions, implying that O/W emulsions with a high

Fig. 7 shows the DSDs of three representative emulsions with longterm resistance to phase separation: 2 wt.% S. thermophilus + 80 wt.% oil + 18 wt.% water, 2 wt.% L. acidophilus + 75 wt.% oil + 23 wt.% water and 9 wt.% S. cerevisiae + 60 wt.% oil + 31 wt.% water. These emulsions were prepared with magnetic stirring and showed different droplet size distributions. The emulsion with S. thermophilus had a D[4,3] of 204 ± 3 μm and DSD ranging from 100 to 500 μm whereas with L. acidophilus, the D[4,3] was 115 ± 2.8 μm and its DSD ranged from 30 to 300 μm. Finally, the yeast-stabilized emulsion had the lowest D [4,3] value (93 ± 7 μm). Its DSD ranged from 50 to 350 μm, with an added distribution from ~2 to ~15 μm, which was likely related to presence of unabsorbed yeast cells. In general terms, taking into consideration droplet coverage, emulsification and D[4,3] values, S. thermophilus showed the lowest propensity towards Pickering emulsion formation and stabilization. 3.5. Apparent viscosity of cell-stabilized emulsions The flow curves of the three emulsions explored above exhibited three distinct regions: Region 1, where at the lowest shear rates, the emulsion showed a tendency towards increased viscosity. A small shear rate-dependent plateau region was visible (Region 2) as was a decrease in viscosity at higher shear rates (Region 3). In Region 1, the slight increase in viscosity at low shear rates (b0.03 s−1) likely resulted from weak network formation via droplet/cell flocculation/aggregation and weak associative interactions (Cormack, 1999). In Region 2 (0.03– 0.05 s−1), the emulsions showed distinct plateau viscosity values: L. acidophilus — 130 ± 32 Pa s, S. cerevisiae — 71 ± 10 Pa s and S. thermophilus — 51 ± 4 Pa s. Pseudoplastic, i.e., shear-thinning, behaviour was apparent in Region 3. The critical shear rate for the onset of shear-thinning was generally similar for the emulsions such that N0.2 s−1, all emulsions exhibited similar profiles up to 100 s−1. 3.6. Contact angle measurements Microscopy has previously been used to directly measure θ (Hutchins, 1971). Fig. 9 shows a CLSM image of an oil droplet with yeast adsorbed onto its surface. To ascertain individual oil–water-cell θ values, a circle was drawn to represent the periphery of the droplet and tangents were marked where the S. cerevisiae cells intersected the droplet. The θ values of 18 adsorbed S. cerevisiae cells measured across the aqueous phase ranged from 16 to 44° (average: 30 ± 9°). Similar attempts with S. thermophilus and L. acidophilus were unsuccessful due to the smaller size of the bacteria and instrumental limitations. 4. Discussion

Fig. 9. CLSM image of an oil droplet with adsorbed S. cerevisiae cells at the oil–water interface. Contact angle tangents are shown for a number of adsorbed yeast cells. Contact angles were measured through the aqueous phase.

The capacity of bacteria and yeast to adsorb to the oil–water interface as well as promote emulsion formation and kinetic stabilization is multifaceted and relies on many factors including cell surface composition, concentration, size and aggregation behaviour as well as the oil:water ratio. Together, these factors influenced emulsion appearance, texture and longevity.

H. Firoozmand, D. Rousseau / Food Research International 81 (2016) 66–73

71

4.2. Pickering stabilization by microbes

Table 1 Final emulsion compositions; Figs. 1, 3i, 4i and 5. Fig. 1, vial number

1

2

3

4

5

6

7

8

9

Oil wt.% Water wt.% (a) S. thermophilus wt.% (b) L. acidophilus wt.% (c) S. cerevisiae wt.%

50 49 1 1 1

50 48 2 2 2

50 47 3 3 3

50 46 4 4 4

50 45 5 5 5

50 44 6 6 6

50 43 7 7 7

50 42 8 8 8

50 41 9 9

Fig. 3i, vial number

a

b

c

d

e

Oil wt.% Water wt.% S. thermophilus wt.%

40 58 2

50 48 2

60 38 2

70 28 2

80 18 2

Fig. 4i, vial number

a

b

c

d

e

Oil wt.% Water wt.% L. acidophilus wt.%

40 58 2

50 48 2

60 38 2

70 28 2

75 23 2

Fig. 5, vial number

a

b

c

Oil wt.% Water wt.% S. cerevisiae wt.%

60 31 9

65 27 8

70 23 7

4.1. Origin of microbe emulsification ability The ability of these microbes to adsorb to the oil–water interface was related to their cell surface properties. As lactic acid bacteria and yeast live and grow in aqueous environments, they naturally show predominantly hydrophilic behaviour. This is largely due to the dominance of carbohydrates in yeast cell walls (~ 94%) (Dallies, François, & Paquet, 1998) and peptidoglycans in lactic bacteria cell walls (up to 70 wt.%) (Schleifer & Kandler, 1972). Yet, bacteria and yeast have been shown to attach to the oil–water interface via hydrophobic interactions (Dorobantu et al., 2004; van Loosdrecht, Lyklema, Norde, Schraa, & Zehnder, 1987), with such behaviour broadly determined by the composition and conformation of surface-bound proteins, polypeptides and polysaccharides (Chapot-Chartier & Kulakauskas, 2014; Delcour, Ferain, Deghorain, Palumbo, & Hols, 1999; Schär-Zammaretti & Ubbink, 2003). In particular, the presence of (glyco-)proteinaceous material on bacterial cell surfaces greatly contributes to hydrophobicity and by extension, to their propensity towards adhesion (Cuperus et al., 1993; Giaouris, Chapot-Chartier, & Briandet, 2009; Klemm & Schembri, 2000; Mobili, Gerbino, Tymczyszyn, & Gómez-Zavaglia, 2010). As well, in some lactic bacteria, surface-bound (lipo)teichoic acids impart hydrophobicity to the bacterial surface (Schär-Zammaretti & Ubbink, 2003). Similarly, densely-packed fibrous polysaccharides and mannoproteins on yeast cell walls are thought to initiate adhesion to surfaces (Frevert & Ballou, 1985; Hazen, 1989; Hu et al., 2011). Though not tested, heat inactivation of the cells likely denatured cell wall-bound proteins in both bacteria and yeast, which would suggest greater exposure of hydrophobic groups, as observed with biofilms in silicon tubing that become more adherent (Marion-Ferey et al., 2003). Finally, the enhanced ability of L. acidophilus to reside at the interface vs. S. thermophilus may be in part attributed to the existence of pili on the former. Pili are proteinaceous (polymeric) surface-exposed structures with a narrow diameter (1–10 nm) projecting outward from cells up to 1 μm (Kankainen et al., 2009; Mandlik, Swierczynski, Das, & Ton-That, 2008; Proft & Baker, 2009). These are known as a major contributor to cell surface adhesion and to biofilm formation. Though the recorded θ values for the yeast (and presumably the bacteria) supports the existence of surface hydrophilic and hydrophobic groups, the role and conformation of other groups responsible for surface adhesion (e.g., pili) cannot be discounted and may have further promoted the ability of the microbes to adsorb to the oil–water interface. Such ability is the focus of an on-going study.

Key requirements dictating successful emulsion stabilization by Pickering particles include the spatial arrangement, size, surface activity and extent of droplet coverage of the particles. As discussed, both yeast and bacteria behaved as effective Pickering particles (Dorobantu et al., 2004; Ly et al., 2006). In general terms, Pickering stabilization of oil–water interfaces occurs by at least three modes (Fig. 10) (Dickinson, 2013). With monolayer coverage, a single layer of particles englobes the droplet resulting in a thin mechanical barrier around individual droplets (Fig. 10a). Particles may also perform ‘double-duty’ by embedding themselves within the interface of two or more neighbouring droplets thereby arresting coalescence via either monolayer (Fig. 10b) or multilayer (Fig. 10c) bridging. A fourth (non-Pickering) mechanism occurs when particles with little to no affinity for the oil–water interface aggregate to form a network encasing the dispersed aqueous phase (Fig. 10d). As per Fig. 2, the microbial cells englobed the dispersed droplets with either mono- or possibly multilayers, further confirming their Pickering particle-like ability. To permit effective coverage, the diameter of interfacially-adsorbed particles should be much smaller than the droplet size. As noted earlier, S. cerevisiae is b 10 μm in diameter whereas L. acidophilus is b 5 μm in length and S. thermophilus is b 1 μm in diameter. Simply based on geometric constraints, the bacteria were better able to enrobe the droplets than the yeast (Fig. 2), with L. acidophilus yielding the most effective droplet coverage (N90%) followed by S. cerevisiae (~ 80%) and S. thermophilus (~ 65%). The latter bacterium had a greater propensity for self-aggregation than L. acidophilus, which impacted its ability to adsorb to the oil–water interface. Conceptually, effective Pickering stabilization is expected with complete dispersed droplet coverage, however there are numerous instances to the contrary. Midmore was able to obtain ‘stable’ O/W emulsion using colloidal silica particles and hydroxypropyl cellulose as a co-stabilizer (Midmore, 1998). With only 29% surface coverage, it was proposed that the flocculated particles formed a 2-D gel structure at the oil–water interface preventing droplet coalescence. Vignati et al. produced emulsions using fluorescent silica particles with a surface coverage as low as 5% (Vignati, Piazza, & Lockhart, 2003). They proposed that the particles re-distributed on the interface and concentrated near droplet–droplet contact areas, thus hindering coalescence. Horozov and Binks explained the stability of emulsions with partial surface coverage using the bridging particle monolayer mechanism (Horozov & Binks, 2006) further stating that Coulombic repulsion between the most hydrophobic particles resulted in ordered monolayers. The present results suggest a combination of different stabilization mechanisms, including the presence of mono- and multilayers as well as the possibility of localized inter-particle repulsion given the negative zeta potential values of ca. −15 mV (Firoozmand & Rousseau, 2014). Pickering particles should be locally bi-wettable, with a dominant portion of their surface preferentially wetted by the continuous phase. Under optimal conditions, there is a significant free energy change associated with the removal of particles from the oil–water interface, which results in tenacious interfacial anchoring (Dong & Johnson, 2005): ΔE ¼ πR2 γαβ ð1  cos θÞ2

ð1Þ

where ΔE is the free energy of adsorption, R is the radius of the particle, θ is the three-phase contact angle and γαβ is the oil–water interfacial tension. For a preferentially water-wet particle, θ b 90° (Leal-Calderon & Schmitt, 2008). As per Eq. (1), presuming a particle diameter of 1 μm, θ ~ 30° and an oil–water interfacial tension of 32 mN/m, the calculated displacement energy of ~106 kT suggested the effectively irreversible interfacial attachment of the yeast cells. A priori, they should demonstrate a significantly higher detachment energy than either bacteria, given their larger size. However, the situation is not so clear-cut as other factors likely impacted whether or not the microbial cells would

72

H. Firoozmand, D. Rousseau / Food Research International 81 (2016) 66–73

Fig. 10. Schematic representation of modes of emulsion stabilization by particles. a) Particle monolayer coverage; b) droplet–droplet monolayer bridging; c) droplet–droplet multilayer bridging and; d) network stabilization where the dispersed oil phase is encased.

aggregate, e.g., the presence of pili and other cell surface components on certain microbes. 4.3. Oil:water ratio effects on emulsification efficacy The present results indicated that single-phase emulsions were preferentially formed at high oil:water ratios (minimum N 2:1), for example with S. thermophilus (Fig. 3) and L. acidophilus (Fig. 4). Concomitant with the formation of homogeneous kinetically-stable emulsions was their ability to support their own weight and not collapse once removed from their sample vessels (Figs. 3 and 4). Similar observations were generally applicable to the yeast-stabilized emulsions. Presence of a dispersed phase at a higher volume fraction than the continuous phase results in a high internal phase ratio emulsion (Lissant, 1966; Reynolds, Gilbert, & White, 2000; Williams, 1991) alternatively known as a highly-concentrated (Babak & Stebe, 2002; Princen, Aronson, & Moser, 1980) or gel emulsion (Kunieda, Solans, Shida, & Parra, 1987). Such emulsions have practical applications in foods (e.g., mayonnaise), cosmetics (Jager-Lezer et al., 1998), topical drug delivery systems (Riess & Weers, 1996), the extraction of antibiotics and pollutants (Lye & Stuckey, 1998) as well as several other industrial applications (Babak & Stebe, 2002; Krafft & Riess, 1994). The microstructure of the stable one-phase bacteria-based emulsions was strongly dependent on composition. As they consisted of a high oil:water ratio (e.g., 4:1), the oil droplets were arranged in close-packed configuration (Figs. 3 and 4) usually seen above dispersed phase volume fractions of N 0.74, which is the critical volume fraction of closely-packed monodisperse spheres. Hence, the dispersed oil droplets became distorted from their normal, spherical shape with this close packing allowing them to interact strongly with one another. A different picture emerged with the yeast-stabilized emulsions as stable emulsions as lower oil fractions were obtained and a close-packed arrangement was not visible. Rather, these emulsions displayed a polydispersed arrangement typically seen in O/W emulsions, which suggested that the yeast had a greater ability to emulsify than the bacterial cells. As discussed above, cell surface composition was a possible factor. Furthermore, it is possible that given that yeast cells were larger in size than bacterial cells, they may have had a higher propensity to reside at the oil–water

interface than the bacteria (as per Eq. (1)), hence assisting in their ability to produce emulsions with lower proportions of oil. From a rheological standpoint, above threshold compositions, the resulting emulsions were self-supporting, exhibited a thickness similar to many handcreams when rubbed between one's thumb and forefinger (Lukic, Jaksic, Krstonosic, Cekic, & Savic, 2012), and behaved as pseudoplastic fluids exhibiting the typical shear-thinning behaviour of O/W emulsions (Dickinson, 2012). These results demonstrated that composition could be fine-tuned to produce thicker or thinner emulsions, in particular with yeast, which showed the ability to generate more stable emulsions at both lower and higher dispersed oil volume fractions.

5. Conclusion This study has shown that yeast and lactic acid bacteria may be used as micron-sized Pickering-type colloidal particles to produce and stabilize O/W emulsions at different oil:water ratios. With optimized compositions, the dispersed oil droplets covered by cells showed longterm (more than four months) stability against droplet coalescence and bulk phase separation. Of note, these microbial cells were capable of producing one-phase emulsions at high oil volume fractions, with such emulsions consisting of oil droplets in a polyhedral, close-packed arrangement. In terms of stabilization mechanism, the cells adhered to the oil–water interface and prevented oil droplet coalescence via Pickering stabilization. The combined properties of full droplet coverage and the close-packed oil droplet arrangement imparted a ‘stand-up’ ability to the emulsions. This novel approach to emulsion stabilization has many possible food applications, including the replacement of solid fat or synthetic surfactants as well the development of emulsions with an ‘all-natural’ designation. However, as relatively high proportions of microbial cells were required for emulsion formation and stabilization, this suggests that these cells may be used under circumstances where the high surface activity of small-molecule surfactants or proteins is not critical. Studies are underway to further explore the mechanisms of action, usability and range of possible applications of these microbial cells.

H. Firoozmand, D. Rousseau / Food Research International 81 (2016) 66–73

References Babak, V. G., & Stebe, M. J. (2002). Highly concentrated emulsions. II. Real systems. The effect of film thickness and contact angle on the volume fraction in creamed emulsions. Journal of Dispersion Science and Technology, 23(1–3), 1–22. Balat, M., & Balat, H. (2010). Progress in biodiesel processing. Applied Energy, 87(6), 1815–1835. Carr, F. J., Chill, D., & Maida, N. (2002). The lactic acid bacteria: A literature survey. Critical Reviews in Microbiology, 28(4), 281–370. Chapot-Chartier, M. -P., & Kulakauskas, S. (2014). Cell wall structure and function in lactic acid bacteria. Microbial Cell Factories, 13(Suppl. 1: S9), 23. Cormack, D. (1999). The oil–water system. In D. Cormack (Ed.), Response to marine oil pollution, review and assessment (pp. 22–64). Springer Science & Business Media. Cuperus, P. L., Vandermei, H. C., Reid, G., Bruce, A. W., Khoury, A. H., Rouxhet, P. G., & Busscher, H. J. (1993). Physicochemical surface characteristics of urogenital and poultry lactobacilli. Journal of Colloid and Interface Science, 156(2), 319–324. Dallies, N., François, J., & Paquet, V. (1998). A new method for quantitative determination of polysaccharides in the yeast cell wall. Application to the cell wall defective mutants of Saccharomyces cerevisiae. Yeast, 14(14), 1297–1306. Delcour, J., Ferain, T., Deghorain, M., Palumbo, E., & Hols, P. (1999). The biosynthesis and functionality of the cell-wall of lactic acid bacteria. Proceedings of the Sixth Symposium on Lactic Acid Bacteria: Genetics, Metabolism and Applications, Veldhover, Netherlands, 19–23 September 1999, 76. (pp. 159–184). Destribats, M., Rouvet, M., Gehin-Delval, C., Schmitt, C., & Binks, B. P. (2014). Emulsions stabilised by whey protein microgel particles: towards food-grade Pickering emulsions. Soft Matter, 10(10), 6941–6954. http://dx.doi.org/10.1039/C4SM00179F. Dickinson, E. (2009). Hydrocolloids as emulsifiers and emulsion stabilizers. Food Hydrocolloids, 23(6), 1473–1482. Dickinson, E. (2010). Food emulsions and foams: Stabilization by particles. Current Opinion in Colloid & Interface Science, 15(1–2), 40–49. Dickinson, E. (2012). Use of nanoparticles and microparticles in the formation and stabilization of food emulsions. Trends in Food Science & Technology, 24(1), 4–12. Dickinson, E. (2013). Stabilising emulsion-based colloidal structures with mixed food ingredients. Journal of the Science of Food and Agriculture, 93(4), 710–721. Dong, L., & Johnson, D. T. (2005). Adsorption of acicular particles at liquid–fluid interfaces and the influence of the line tension. Langmuir, 21(9), 3838–3849. Dorobantu, L. S., Yeung, A. K. C., Foght, J. M., & Gray, M. R. (2004). Applied and Environmental Microbiology, 70(10), 6333–6336, Stabilization of oil–water emulsions by hydrophobic bacteria [Article]. Firoozmand, H., & Rousseau, D. (2014). Tailoring the morphology and rheology of phaseseparated biopolymer gels using microbial cells as structure modifiers. Food Hydrocolloids, 42(Part 1), 204–214. de Folter, J. W. J., van Ruijven, M. W. M., & Velikov, K. P. (2012). Oil-in-water Pickering emulsions stabilized by colloidal particles from the water-insoluble protein zein. Soft Matter, 8(25), 6807–6815. Frevert, J., & Ballou, C. E. (1985). Saccharomyces-cerevisiae structural cell-wall mannoprotein. Biochemistry, 24(3), 753–759. Furtado, G. F., Picone, C. S. F., Cuellar, M. C., & Cunha, R. L. (2015). Breaking oil-in-water emulsions stabilized by yeast. Colloids and Surfaces B: Biointerfaces, 128, 568–576. Giaouris, E., Chapot-Chartier, M. P., & Briandet, R. (2009). Surface physicochemical analysis of natural Lactococcus lactis strains reveals the existence of hydrophobic and low charged strains with altered adhesive properties. International Journal of Food Microbiology, 131(1), 2–9. Hazen, K. C. (1989). Participation of yeast-cell surface hydrophobicity in adherence of candida-albicans to human epithelial-cells. Infection and Immunity, 57(7), 1894–1900. Heeres, A. S., Picone, C. S. F., van der Wielen, L. A. M., Cunha, R. L., & Cuellar, M. C. (2014). Microbial advanced biofuels production: overcoming emulsification challenges for large-scale operation. Trends in Biotechnology, 32(4), 221–229. Horozov, T. S., & Binks, B. P. (2006). Particle-stabilized emulsions: A bilayer or a bridging monolayer? Angewandte Chemie International Edition, 45(5), 773–776. Hu, S., Kong, J., Sun, Z., Han, L., Kong, W., & Yang, P. (2011). Heterologous protein display on the cell surface of lactic acid bacteria mediated by the s-layer protein. Microbial Cell Factories, 10(86), 13. Hutchins, T. E. (1971). Direct measurement of microscopic contact angle. Thin Solid Films, 8(4), R25-&. Jager-Lezer, N., Tranchant, J. F., Alard, V., Vu, C., Tchoreloff, P. C., & Grossiord, J. L. (1998). Rheological analysis of highly concentrated w/o emulsions. Rheologica Acta, 37(2), 129–138. Kankainen, M., Paulin, L., Tynkkynen, S., von Ossowski, I., Reunanen, J., Partanen, P., ... de Vos, W. M. (2009). Comparative genomic analysis of Lactobacillus rhamnosus GG reveals pili containing a human-mucus binding protein. Proceedings of the National Academy of Sciences, 106(40), 17193–17198. Klemm, P., & Schembri, M. A. (2000). Bacterial adhesins: Function and structure. International Journal of Medical Microbiology, 290(1), 27–35. Krafft, M. P., & Riess, J. G. (1994). Stable highly concentrated fluorocarbon gels. Angewandte Chemie International Edition in English, 33(10), 1100–1101. Kunieda, H., Solans, C., Shida, N., & Parra, J. L. (1987). The formation of gel emulsions in a water nonionic surfactant oil system. Colloids and Surfaces, 24(2–3), 225–237. Lam, S., Velikov, K. P., & Velev, O. D. (2014). Pickering stabilization of foams and emulsions with particles of biological origin. Current Opinion in Colloid & Interface Science, 19(5), 490–500. Leal-Calderon, F., & Schmitt, V. (2008). Solid-stabilized emulsions. Current Opinion in Colloid & Interface Science, 13(4), 217–227. Lim, S., & Teong, L. K. (2010). Recent trends, opportunities and challenges of biodiesel in Malaysia: An overview. Renewable and Sustainable Energy Reviews, 14(3), 938–954. Lin, L., Cunshan, Z., Vittayapadung, S., Xiangqian, S., & Mingdong, D. (2011). Opportunities and challenges for biodiesel fuel. Applied Energy, 88(4), 1020–1031.

73

Lissant, K. J. (1966). The geometry of high-internal-phase-ratio emulsions. Journal of Colloid and Interface Science, 22(5), 462–468. Liu, F., & Tang, C. -H. (2013). Soy protein nanoparticle aggregates as Pickering stabilizers for oil-in-water emulsions. Journal of Agricultural and Food Chemistry, 61(37), 8888–8898. van Loosdrecht, M. C. M., Lyklema, J., Norde, W., Schraa, G., & Zehnder, A. J. B. (1987). The role of bacterial-cell wall hydrophobicity in adhesion. Applied and Environmental Microbiology, 53(8), 1893–1897. Lukic, M., Jaksic, I., Krstonosic, V., Cekic, N., & Savic, S. (2012). A combined approach in characterization of an effective w/o hand cream: The influence of emollient on textural, sensorial and in vivo skin performance. International Journal of Cosmetic Science, 34(2), 140–149. Ly, M. H., Naitali-Bouchez, M., Meylheuc, T., Bellon-Fontaine, M. N., Le, T. M., Belin, J. M., & Wache, Y. (2006). Importance of bacterial surface properties to control the stability of emulsions. International Journal of Food Microbiology, 112(1), 26–34. Lye, G. J., & Stuckey, D. C. (1998). Structure and stability of colloidal liquid aphrons. Colloids and Surfaces A: Physicochemical and Engineering Aspects, 131(1–3), 119–136. Mandlik, A., Swierczynski, A., Das, A., & Ton-That, H. (2008). Pili in Gram-positive bacteria: Assembly, involvement in colonization and biofilm development. Trends in Microbiology, 16(1), 33–40. Marion-Ferey, K., Pasmore, M., Stoodley, P., Wilson, S., Husson, G. P., & Costerton, J. W. (2003). Biofilm removal from silicone tubing: An assessment of the efficacy of dialysis machine decontamination procedures using an in vitro model. Journal of Hospital Infection, 53(1), 64–71. Midmore, B. R. (1998). Preparation of a novel silica-stabilized oil/water emulsion. Colloids and Surfaces A: Physicochemical and Engineering Aspects, 132(2–3), 257–265. Mobili, P., Gerbino, E., Tymczyszyn, E. E., & Gómez-Zavaglia, A. (2010). S-layers in lactobacilli: Structural characteristics and putative role in surface and probiotic properties of whole bacteria. In A. Méndez-Vilas (Ed.), Current Research, technology and education topics in applied microbiology and microbial biotechnology, Vol. 2. (pp. 1224–1234). Spain: Formatex. Nguyen, B. T., Nicolai, T., & Benyahia, L. (2013). Stabilization of water-in-water emulsions by addition of protein particles. Langmuir, 29(34), 10658–10664. Pickering, S. U. (1907). Emulsions. Journal of the Chemical Society, 91, 2001–2021. Poortinga, A. T. (2008). Microcapsules from self-assembled colloidal particles using aqueous phase-separated polymer solutions. Langmuir, 24(5), 1644–1647. Princen, H. M., Aronson, M. P., & Moser, J. C. (1980). Highly concentrated emulsions. 2. Real systems. The effect of film thickness and contact-angle on the volume fraction in creamed emulsions. Journal of Colloid and Interface Science, 75(1), 246–270. Proft, T., & Baker, E. N. (2009). Pili in Gram-negative and Gram-positive bacteria — Structure, assembly and their role in disease. Cellular and Molecular Life Sciences, 66(4), 613–635. Randez-Gil, F., Cocoles-Saz, I., & Prieto, J. A. (2013). Genetic and phenotypic characteristics of Baker's yeast: Relevance to baking. In M. P. Doyle, & T. R. Klaenhammer (Eds.), Annual Review of Food Science and Technology, Vol 4, Vol. 4. (pp. 191–214). Palo Alto: Annual Reviews. Rayner, M., Timgren, A., Sjöö, M., & Dejmek, P. (2012). Quinoa starch granules: A candidate for stabilising food-grade Pickering emulsions. Journal of the Science of Food and Agriculture, 92(9), 1841–1847. Reynolds, P. A., Gilbert, E. P., & White, J. W. (2000). High internal phase water-in-oil emulsions studied by small-angle neutron scattering. Journal of Physical Chemistry B, 104(30), 7012–7022. Riess, J. G., & Weers, J. G. (1996). Emulsions for biomedical uses. Current Opinion in Colloid & Interface Science, 1(5), 652–659. Rosenberg, M., & Rosenberg, E. (1985). Bacterial adherence at the hydrocarbon-water interface. Oil and Petrochemical Pollution, 2(3), 155–162. Schär-Zammaretti, P., & Ubbink, J. (2003). The cell wall of lactic acid bacteria: Surface constituents and macromolecular conformations. Biophysical Journal, 85(6), 4076–4092. Schleifer, K. H., & Kandler, O. (1972). Peptidoglycan types of bacterial cellwalls and their taxonomic implications. Bacteriological Reviews, 36(4), 407–477. Srivasta, S. P., Singh, H. D., Baruah, J. N., Krishna, P. V., & Iyengar, M. S. (1970). Physicochemical studies on water-yeast cells-gas-oil system. [Article]. Journal of Applied Chemistry, 20(4), 105–108. Velikov, K. P., & Pelan, E. (2008). Colloidal delivery systems for micronutrients and nutraceuticals. [Review]. Soft Matter, 4(10), 1964–1980. Vignati, E., Piazza, R., & Lockhart, T. P. (2003). Pickering emulsions: Interfacial tension, colloidal layer morphology, and trapped-particle motion. Langmuir, 19(17), 6650–6656. Wege, H. A., Kim, S., Paunov, V. N., Zhong, Q. X., & Velev, O. D. (2008). Long-term stabilization of foams and emulsions with in-situ formed microparticles from hydrophobic cellulose. Langmuir, 24(17), 9245–9253. Williams, J. M. (1991). High internal phase water-in-oil emulsions — Influence of surfactants and cosurfactants on emulsion stability and foam quality. Langmuir, 7(7), 1370–1377. Wongkongkatep, P., Manopwisedjaroen, K., Tiposoth, P., Archakunakorn, S., Pongtharangkul, T., Suphantharika, M., ... Wongkongkatep, J. (2012). Bacteria interface pickering emulsions stabilized by self-assembled bacteria-chitosan network. [Article]. Langmuir, 28(13), 5729–5736. Yusoff, A., & Murray, B. S. (2011). Modified starch granules as particle-stabilizers of oil-inwater emulsions. Food Hydrocolloids, 25(1), 42–55. Yusuf, N. N. A. N., Kamarudin, S. K., & Yaakub, Z. (2011). Overview on the current trends in biodiesel production. Energy Conversion and Management, 52(7), 2741–2751. Zhu, W., Ma, W., Li, C., Pan, J., Dai, X., Gan, M., ... Zhang, Y. (2014). Magnetic molecularly imprinted microspheres via yeast stabilized Pickering emulsion polymerization for selective recognition of λ-cyhalothrin. Colloids and Surfaces A: Physicochemical and Engineering Aspects, 453, 27–36.