Pea Protein—ĸ-Carrageenan Nanoparticles for Edible Pickering Emulsions

Abstract

Pickering emulsions (PEs) can be utilized as inks for 3D food printing owing to their extensive stability and appropriate viscoelastic properties. This research explores food-grade PEs stabilized with nanoparticles (NPs) based on modified pea protein (PP) isolate and k-carrageenan (KC). NPs are fabricated from solutions with different concentrations of protein and polysaccharide and characterized in terms of size, zeta potential, and wetting properties. The composition of the emulsion is 60% sunflower oil and 40% aqueous phase. Nine emulsion formulations with varying PP and KC concentrations are investigated. The formation of hollow NPs with a hydrodynamic diameter of 120–250 nm is observed. Microscope imaging shows oil droplets surrounded by a continuous aqueous phase, forming homogenous PEs in all formulations that are stable for over 30 days. Further, the oil droplet size decreases with increasing NP concentration while the viscosity increases. Rheologic experiments portray elastic emulsion gels with thixotropic qualities ascribed to the presence of the polysaccharide. The emulsions are subjected to centrifugation in order to compare the original emulsions to concentrated PEs that possess improved capabilities. These emulsions may serve as sustainable and printable saturated fat alternatives due to their composition, texture, stability, and rheological properties. Lastly, PEs are printed smoothly and precisely while maintaining a self-supported structure.

1. Introduction

3D food printing is an innovative technology that enables the creation of a wide variety of precise and personalized food products. This methodology possesses advantages such as convenience, customized design, flexibility, and sustainability. Emulsions have gained interest as functional printing material due to their rheological characteristics that include shear-thinning behavior, high storage modulus, and the ability to be tuned toward an ideal viscosity. These attributes are favorable for obtaining extrusion fluidity, printability, and stability of 3D-printed objects [1,2,3]. A classic emulsion comprises dispersed droplets of one immiscible liquid in another, such as oil and water, stabilized by emulsifiers. In Pickering emulsions (PEs), solid particles replace surfactants, thereby enabling a highly stable, biocompatible dispersion. In particular, high internal phase Pickering emulsions (HIPPEs), which contain an internal phase ratio of over 74% (φ > 0.74), have gained attention in the food industry as a potential substitute for hydrogenated oils and saturated fat [4]. HIPPEs possess many useful characteristics, such as large surface area and controllable flow, which allow for innovative applications in the field of food technology [1]. A facile method to obtain HIPPEs is by centrifugation and concentrating of low internal phase emulsions (pre-emulsions) [5,6]. The emulsifying capacity of many types of particles is limited to certain oil concentrations, and thus, centrifugation may provide a means to prepare HIPPEs with minimal solid concentration [3,7]. However, centrifugation often destabilizes the emulsion due to the coalescing of the oil droplets, leading to phase separation [8]. Therefore, the high stability of emulsions during centrifugation is essential for the successful creation of concentrated emulsions using this method [3,7]. For these stable emulsions, centrifugation is a valuable tool for the formation of biocompatible and food-grade emulsions because it does not require the addition of harmful surfactants or synthetic additives to form the desired emulsions [1,3]. Forcing the oil droplets closer together by means of centrifugation can produce a solid emulsion with gel-like characteristics suitable for 3D printing applications, such as elasticity and high storage modulus [9]. Utilizing plant-based solid particles to stabilize PEs enables the formation of vegan, economic, biocompatible [10], and biodegradable emulsions [11,12]. Pea protein (PP)—one of the most economically accessible natural plant-derived proteins—emerges as a promising substitute for animal-based foods due to its high nutritional value, low allergenicity, and affordability [13,14]. According to previous research, it has been established that protein-based nanoparticles (NPs) are GRAS and pose no risk of toxicity [10,15]. Among legumes, pea is one of the species with the highest protein content, reaching up to 30% protein, and possesses a full amino-acid profile. PP can be classified into four major groups: albumins, globulins, prolamins, and glutelins; the majority are globulins (65–80%) and albumins (10–20%) [14,16]. Despite its advantages, the use of PP in various fields is limited by its hydrophobicity, low solubility, and tendency to aggregate at pH levels close to its isoelectric point (~4.3) or under high ionic strength conditions. Therefore, it is challenging to obtain suitable emulsion stability at the droplet interface using PP as the sole stabilizer [13,17]. Nevertheless, modification of PP through pH shifting, heat shifting, and ultrasound treatment can enhance its solubility, reduce aggregate size, and improve the emulsifying capacity [16].

The modification of plant-based proteins such as PP opens the door to creating sustainable, multifunctional food ingredients and provides solutions to deal with their limitations by changing their physicochemical properties [18]. For example, when a protein is exposed to a thermal treatment, the interior sulfhydryl groups and the hydrophobic side chains that were previously hidden in the core of the native-state structure become exposed due to the unfolding of polypeptide chains [19]. Another approach is ultrasound treatment, which is an efficient, inexpensive, and environment-friendly modification method that can induce structural changes by disrupting non-covalent bonds, although it is inert towards the covalent bonds. Consequently, ultrasound treatment can reduce the aggregate size, improve solubility, and enhance emulsification capacity by partially denaturing the tertiary and quaternary structures of proteins and destroying their secondary structures without significantly altering their fundamental structure [16]. Finally, proteins can undergo structural and functional changes as a result of alkaline treatment. Proteins undergo denaturation and unfold at highly basic pH values, thereby revealing hydrophobic and sulfhydryl regions in their structure and consequently allowing for new protein interactions. pH shifting has been demonstrated to enhance emulsifying and antioxidant activity as well as improve solubility, rheological properties, and surface hydrophobicity of proteins [14,16,18,20]. Applying these methods in the creation of PP NPs can result in superior stability and printability compared to currently researched PP-based PEs.

Polysaccharides have been shown to effectively aid the creation of stable PEs [2,7,17,21,22,23]. Studies that employ NPs based on plant protein in combination with polysaccharides for 3D printing applications are relatively scarce, but several protein-polysaccharide complexes have been reported to stabilize PEs. These include rice protein-carboxymethyl cellulose [24], pea protein-pectin [25], and zein-gum Arabic [26] complexes. In addition, the fabrication of PEs stabilized with complexes of pea protein and k-carrageenan (KC) has only recently been studied [2,27]. Carrageenans are a family of polysaccharides that are obtained from red seaweed. KC (pKa ~4.6) is a sulfated polysaccharide comprising 3,6-anhydro-d-galactopyranosyl and/or d-galactopyranosyl units with a sulfate half-ester group attached to the monosaccharide [28,29]. It is a common food additive used as a thickener, gelling agent, and stabilizer [12,28]. The anionic, hydrophilic, and viscoelastic traits of this hydrocolloid were shown to aid in reducing the size of PP NPs through electrostatic interactions by increasing the functionality and enhancing the emulsification capability of PP alone [30,31,32]. In previous studies, a very low concentration of PP was used along with KC and grape seed oil to enhance the chemical stability of curcumin [27]. In an additional research study, microparticles of PP and KC with different KC concentrations were fabricated and investigated as 3D printing material [2]. These studies demonstrate the promising potential of PP-KC particles to stabilize PE for 3D food printing; however, expanding the range of particle compositions is necessary to improve the understanding of the relationship between the physiochemical properties of the NPs and the emulsifying capacity. In addition, the fabrication of NPs, as compared to microparticles, may improve the stability and rheological performance of the PEs. 3D bioprinting of HIPPEs stabilized with pea protein-pectin-EGCG was demonstrated by Feng et al. [25]. Liu et al. printed curcumin-enriched Pickering emulsions stabilized by pea protein-carrageenan micro-scale complexes [2], hence demonstrating the potential to serve as cake decorations, oral delivery systems, and fat replacements.

In this study, we created biocompatible and food-grade NPs composed of KC and PP in various concentrations and further employed them to stabilize edible Pickering emulsions. Next, we investigated whether centrifugation can be utilized as a method to fabricate concentrated emulsions with improved printing characteristics. We evaluated the properties of the pre-emulsions and the emulsions after centrifugation, as well as their performance in 3D printing. Comparing emulsion characteristics before and after centrifugation may provide a deeper understanding of the requirements for and possibilities of applying PEs in the world of 3D food printing, creating a sustainable, healthier, food-grade fat alternative.

2. Materials and Methods

2.1. Materials

PP (pea protein isolate, F85F) was supplied by Roquette (Lestrem, France). κ-carrageenan (molecular weight—856 kDa [33]), Nile red, NaOH pearls, and HCl were supplied by Sigma-Aldrich Inc. (Rehovot, Israel). Sunflower oil was purchased at a local supermarket. Double distilled water (DDW) was used to prepare all solutions.

2.2. Preparation of the PP-KC NPs

First, PP was modified to enhance its solubility, using a protocol based on previous studies, with certain adjustments [34,35]. PP was dissolved in DDW at a concentration of 2%, 4%, and 6% (w/v) at an ambient temperature and under continuous stirring for 2 h to facilitate the complete hydration of the powders. The pH of the solutions was raised to 12.00 ± 0.02 with 5M NaOH; thereafter, the solutions were heated to 90 °C for 30 min to denature the protein and break its structure into fibrils. The solutions were cooled to room temperature, sonicated at 100 W for 10 min, and finally, the pH was adjusted to 7.00 ± 0.02 with 5M HCl. Each solution was placed in a centrifuge for 30 min at 5000 rpm to remove any insoluble protein, and the supernatant was collected. Next, the soluble PP solutions were mixed at a volume ratio of 1:1 with 0.0%, 0.5%, or 1.0% (w/v) KC solution to create nine NP formulations (see Table 1 below). The final dispersions were heated to 55 °C for 30 min, followed by heating at 95 °C for an additional 40 min to form the PP-KC NPs. Finally, the samples were stored overnight at 4 °C.

2.3. Characterization of the NPs

2.3.1. Dynamic Light Scattering (DLS)

DLS was applied to measure the hydrodynamic sizes and zeta potential of the dispersions. Measurements were performed in triplicates using a Malvern Zetasizer (Nano ZSP, Malvern Instrument Ltd., Worcestershire, UK). The experiments were conducted at 25 °C, with a dilution factor of 100 by volume and an equilibration time of 30 s.

2.3.2. Cryogenic Transmission Electron Microscope (Cryo-TEM)

Samples were cast on a carbon-coated perforated polymer film supported on a 200-mesh TEM grid. The grid was then plunged quickly into liquid ethane at its freezing point (−183 °C). A TEM (Thermo-Fisher Talos F200C, Thermo Fisher Scientific Inc., Waltham, MA, USA, FEG-equipped high-resolution TEM, operated at 200 kV) was utilized to observe the morphology of the PP-KC NPs.

2.3.3. Wettability Measurements and Surface Tension

The three-phase contact angle among oil, water, and NPs (θ_ow) and the surface tension were measured using an Attension Theta Lite tensiometer (Phoenix, AZ, USA). Freeze-dried NP samples were compressed into thin tablets of a thickness of 1 mm and a diameter of 20 mm. Subsequently, each tablet was submerged in sunflower oil in an optical glass cuvette, followed by depositing 7 μL of DDW on the tablet’s surface. After attaining equilibrium, images of the droplet were captured, and the contact angle θ_ow was calculated based on the Young–Laplace equation [27]. The surface tension was measured using the pendant drop method [36]. Further, 7 μL of the NPs’ suspension was deposited into sunflower oil in an optical glass cuvette, followed by the imaging of the submerged droplet. The surface tension was calculated using the Young–Laplace equation [36,37] to determine the surface tension of the droplet based on its shape.

2.3.4. Fourier Transform Infrared Spectroscopy (FTIR)

The FTIR spectra of freeze-dried PP and KC solutions, as well as dried PP-KC NPs, were acquired using an FTIR spectrometer (Bruker Tensor 27 FT-IR, Madison, WI, USA) equipped with a high-sensitivity LN-cooled MCT detector in the range of 4000–400 cm⁻¹ with 32 scans at a resolution of 4 cm⁻¹.

2.4. Preparation of Emulsions

Sunflower seed oil was mixed with PP solution or PP-KC NP dispersions of various concentrations at an oil-dispersion ratio of 3:2. In addition, various oil-dispersion ratios were examined with NPs at a concentration of P2KC0.25. The mixtures were vortexed for a few seconds and immediately homogenized at 24,000 rpm for 2 min using a Bio-Gen PRO200 homogenizer (Oxford, CT, USA) to prepare PEs. The freshly prepared emulsions were centrifuged at 5000 rpm for 20 min, and the redundant aqueous phase formed at the bottom was removed and discarded. After the aqueous bottom phase was collected, the volume fraction of the dispersed phase of the emulsion was calculated according to the following equation:

(1)$\mathrm{Internal} \mathrm{phase} \mathrm{volume} \mathrm{fraction} [\%] = \left({\displaystyle \frac{{\mathrm{V}}_0}{{\mathrm{V}}_{\mathrm{t}}-{\mathrm{V}}_1}}\right)\mathrm{·100}$

where V₀ is the initial oil volume, V_t is the total volume of the initial emulsion, and V₁ is the volume of the aqueous phase withdrawn after centrifugation. The final internal phase rose with an increase in the released water volume. After calculating the final oil fraction, it was established whether HIPPE volume (φ > 0.74) could be achieved as long as phase separation was not induced. The PEs before and after centrifugation were stored in a refrigerator at 2–8 °C.

2.5. Characterization of Emulsions

2.5.1. Stability, Accelerated Stability, and Concentrated Emulsions

An emulsion is considered stable if it is resistant against coalescence over a practical length of time. The stability of the emulsion was observed over a period of 30 days while it was stored at 4 °C to assess if oil was released from the emulsion, a phenomenon which is indicative of phase separation. In addition, accelerated stability tests were performed by centrifugation, which might have induced creaming or sedimentation, flocculation, and eventually coalescence. The test was performed as described by Song et al. (2023) [3]. The emulsions were subjected to centrifugation, as described in Section 2.4, and then inspected to visualize the phase separation.

2.5.2. Confocal Laser Scanning Microscopy (CSLM)

The interfacial structure of the emulsions as-prepared and after centrifugation was analyzed through CLSM using an LSM 880 laser scanning confocal microscope (Carl Zeiss Ltd., Oberkochen, Germany) with a 10 × dry-immersion objective lens. Nile Red (0.1%, w/v) was mixed with the oil phase during emulsion preparation. The stained emulsions were placed on microscope slides without dilution and observed using solid-state lasers with an excitation line of 514 nm for Nile Red. The images were analyzed using Fiji ImageJ software (version 2.14.0) to calculate the droplet size and size distribution.

2.5.3. Rheological Measurements

Rheological experiments were performed by an MCR 302 rheometer (Anton Paar, Kfar Saba, Israel) with a parallel plate geometry of 40 mm and a gap of 1 mm. A range of tests, including an amplitude sweep, was performed in the range of 0.1–100% at a fixed frequency of 10 Hz to determine the linear viscoelastic range (LVR), and frequency sweeps were performed in the range of 1–100 (rad/s) within which subsequent experiments were performed. Further, flow tests were conducted within a shear rate range of 0.1–100 1/s to obtain the viscosity change curve of PEs. The measured viscosity curves were fitted to a power law model, shown in the following equation:

(2)$\eta =K·{\dot{\gamma }}^{\left(n-1\right)}$

where η is viscosity, K is the consistency index, $\dot{\gamma }$ is the shear rate, and n is the flow index; n < 1 indicates shear thinning behavior [38].

To simulate the shear conditions in extrusion 3D printing, oscillatory thixotropy tests were employed to characterize the recoverability of PEs in terms of storage modulus (G′) using a five-step experiment that involved repeating cycles of low strain (0.1% for 40 s) followed by high strain (300% for 20 s). The recoverability (δ) of the PEs was quantified using two parameters. The first is δ₁, which is defined as the average value of G′ obtained during the third step (under low strain) compared to the average value obtained in the first step. The second is δ₂, which is defined as the average value of G′ obtained during the fifth step (under low strain) compared to the average G′ obtained in the third step.

These parameters are defined in the following equations [2,39]:

(3)${\mathsf{\delta }}_1\left[\mathrm{\%}\right]=100·\left({\displaystyle \frac{{{\mathrm{G}}^{\prime }}_3}{{{\mathrm{G}}^{\prime }}_1}}\right)$

(4)${\mathsf{\delta }}_2[\mathrm{\%}]=100·\left({\displaystyle \frac{{{\mathrm{G}}^{\prime }}_5}{{{\mathrm{G}}^{\prime }}_3}}\right)$

where G′₃ and G′₅ are the average storage modulus at low strain in the third and fifth steps, respectively, and G′₁ is the average storage modulus at low strain in the first step. All tests were conducted at 25 °C.

2.6. 3D Printing

PEs were transferred into a 3 mL cartridge (Nordson EFD, Westlake, OH, USA) and loaded into a six-axis printing tool extrusion bioprinter BioAssemblyBot (Advanced Solutions, Louisville, KY, USA). A hollow cylinder structure with an inner diameter of 10 mm and a height of 3 mm, 6 mm, or 9 mm was printed using a 250-µm tapered tip (Nordson EFD, Westlake, OH, USA). A rectilinear grid structure with dimensions of 10 mm × 10 mm and 20 mm × 20 mm was printed using a 410 µm inner diameter blunt end needle (CML supply). The structures were designed using TSIM software (version 1.1.230, Advanced Solution, Louisville, KY, USA). The printing process was performed at a constant speed (11–12 mm/s) and pressure (7 PSI–10 PSI). The printing quality of the PE samples was quantified by calculating the printability index, P_r, according to the following equation:

(5)${\mathrm{P}}_{\mathrm{r}}={\displaystyle \frac{{\mathrm{L}}^2}{16\mathrm{A}}}$

where L is the perimeter, and A is the area of the square. For each printed emulsion with a rectilinear pattern, the perimeter and the area of the internal square were measured using FIJI ImageJ (version 2.14.0), and the P_r index was calculated (n = 5). High geometric accuracy generates P_r = 1, while P_r < 1 or P_r > 1 corresponds to a more round or irregularly shaped geometry, respectively. In addition, the fidelity was assessed by quantifying the deviation between the dimensions of the construct and those of the computer-aided design (CAD) and expressing the results in terms of normalized area (A_r, Equation (6)) and normalized perimeter (D_r, Equation (7)) [40,41].

(6)${\mathrm{A}}_{\mathrm{r}}={\displaystyle \frac{area of printed shape}{area of designed shape}}$

(7)${\mathrm{D}}_{\mathrm{r}}={\displaystyle \frac{perimeter of printed shape}{perimeter of designed shape}}$

where A_r or D_r values close to 1 indicate relatively higher fidelity; moreover, as they diverge from 1, the printing fidelity deteriorates. The fidelity was assessed from five experiments (n = 5).

2.7. Statistical Analysis

Unless otherwise stated, all measurements were conducted in triplicate and reported as the mean values with standard deviation. Statistical analysis was performed using Prism (GraphPad, version 9.1.0). Further, one-way or two-way analysis of variance (ANOVA) with Tukey’s honest significant difference (HSD) test was utilized to evaluate the significant differences at p < 0.05.

3. Results and Discussion

3.1. Characterization of the NPs

3.1.1. Mean Size and Zeta Potential

Here, we describe the fabrication and characterization of NPs based on PP and KC at varying concentrations of both components. As discussed in the following sections, the NPs are further utilized as a solid barrier between oil and water to form stable, food-grade Pickering emulsions. Figure 1A displays the average size of PP-KC NPs for various formulations. The spontaneous formation of the NPs can be attributed to the degradation of the ternary and quaternary structures of the protein, followed by interaction and aggregation among the denatured protein fibrils, as well as interactions between PP and KC [27]. These interactions include weak interactions, such as hydrogen bonds, and hydrophobic interactions, as well as ionic bonds formed between the anionic sulfate groups on the polysaccharide and positively charged local patches (−NH₃⁺ groups, pKa ≈ 11) revealed within the proteins during their modification [27]. As evident from Figure 1A, the hydrodynamic diameter of the NPs ranged from 120 nm to 230 nm, depending on the components’ concentration. The size distribution of the NPs was wide and typically monodispersed (Figure S1). In certain formulations (P1K0.5, P2K0.5), an additional, much smaller, peak appeared in the range of 10–50 nm; in others, a left long-tailed distribution was observed (formulations P1K0, P1K0.25, and P3K0.25). These results are in line with a previous study that attributed the large size variability to the mechanism of complex coacervate formation driven by electrostatic interactions [2]. The formation of self-assembled aggregates driven by hydrophobic interactions as well as ionic interactions with negatively charged KC are spontaneous processes that are not fully controlled; hence, this increases the size variability.

Despite the wide size distribution, clear trends relating to the components’ concentrations to the NP’s average size could be distinguished. When considering varying concentrations of protein at a constant KC concentration, only a slight change in size is observed. The average diameter ranges from 120–165 nm for 0% (w/v) KC, 137–178 nm for 0.25% (w/v) KC, and 217–231 nm at 0.5% (w/v) KC. This finding indicates that the size of the NPs does not strongly depend on the protein concentration within this range. Similarly, the size of NPs prepared from PP solutions without KC does not differ significantly from the size of NPs fabricated from solutions containing PP and 0.25% (w/v) KC for all PP concentrations. In contrast, when increasing the concentration of KC to 0.5% (w/v), the particle size increases, regardless of protein concentration. This trend is observed in all three protein concentrations but is only significant at P1K0.5. The statistical insignificance at higher concentrations of KC and PP could be due to the larger variation in size leading to higher error in size.

The zeta potential of a dispersion is defined as the potential between the liquid superficial layer surrounding the dispersed particle and the remaining volume of the solution. NPs with a zeta potential between −10 mV and +10 mV are considered to be approximately neutral, while NPs with zeta potentials of greater than +30 mV and less than −30 mV are considered strongly cationic and anionic, respectively [42]. Evaluating the zeta potential values of NPs formed from solutions with different PP and KC concentrations may provide a better understanding of their stability, which arises from the surface charge. Zeta potential measurements demonstrated values within a range of −19 mV to −27 mV (Figure 1B). The NP suspension is adjusted to a neutral pH, which is above the PI of the protein and the polysaccharide; therefore, one can expect negatively charged NPs [2]. Electrostatic repulsion among the emulsion droplets could more effectively inhibit the processes of phase separation of emulsions, such as flocculation [27]. In the selected concentration range, neither the PP concentration nor the KC concentration had a significant impact on the zeta potential of the dispersions, with the exception of P3K0.

3.1.2. Cryogenic Transmission Electron Microscope (Cryo-TEM)

Cryo-TEM images reveal hollow spherical particles with a smooth surface (Figure 2A–C). The size of the particles observed in the images aligns with the results obtained by DLS (Section 3.1.1). The formation of spherical NPs was also reported by Wang et al. (2024) [27], where particles were formed at pH 7 from a very low concentration of PP (0.1% w/v); however, these NPs did not appear hollow. In addition, the previous study claimed that the addition of carrageenan affected the roundness [27], but the images in Figure 2 demonstrate that the presence of carrageenan does not impair the shape of the particles. The formation of hollow particles may indicate self-assembly processes due to electrostatic forces between PP aggregates and KC.

3.1.3. Wetting Properties

The influence of PP and KC concentration on the hydrophilicity of the NPs was examined by measuring the three-phase contact angle (θ_ow) of water, sunflower oil, and NPs with different compositions. The wetting properties provide a tool to evaluate the hydrophilicity of the NPs, which in turn influences the emulsion capacity and stability [3,27]. Figure 3A depicts drops of water deposited on tablets of compressed NPs immersed in sunflower seed oil. For all measured NP formulations, the contact angle is larger than 90°, thereby indicating the hydrophobic properties of the NPs [3]. The measured contact angles are similar among all samples, ranging between 100° and 124°, and do not show a significant dependence on the protein or polysaccharide concentration (Figure S2). Moreover, these results do not follow the expectation that the contact angle of the NPs will decrease as the concentration of the hydrophilic KC increases, as seen in the study conducted by Wang et al., (2024) [27]. This may be due to the KC content being relatively small compared to the proportion of PP in each sample, whereas in the study mentioned above, the KC is in excess and, therefore, has a more dominant effect.

Surface tension is a phenomenon that occurs due to cohesive forces between liquid molecules at a certain interface. The tendency of the molecules to interact pulls the molecules toward the bulk; therefore, the value of the surface tension is correlated with the interactions among the molecules in each phase [37,43]. In Figure 3B, the surface tension is depicted as a function of PP and KC concentrations in the solutions used for NP preparation. The surface tension of pure water dropped into sunflower seed oil is 27.3 [N/m]. The surface tension of NP suspensions prepared from solutions containing 1% and 2% (w/v) PP was 28.9 and 30.7 [N/m], respectively, and was not significantly different from the surface tension of water. However, when the protein concentration was raised to 3% (w/v), the surface tension rose to 33.2–34.1 [N/m]. It is evident that the concentration of KC does not have a dominant effect on the measured surface tension, while the protein concentration significantly increases the surface tension when increased beyond 2% (w/v). When characterizing surfactant-based emulsions, the expectation is that the addition of the surfactant will lower the surface tension through interactions with both the water and oil phases. In contrast, in Pickering emulsions, the stability of the emulsion can be derived from an accumulation of solid particles at the oil–water interface. With rising particle concentrations, the Van der Waals force between the accumulated particles increases, which elevates the free energy on the surface and causes the surface tension to increase [36,37]. Since the KC is present in lower proportions compared to PP, the concentration variation effect is minor; however, when increasing the protein concentration above 2% (w/v), the increase in surface tension is significant (p < 0.05), which is similar to the results obtained in the study by Tanvir and Qiao (2012) [36]. The tension among the particles is beneficial for the stability of the emulsion in terms of preventing the coalescence of oil droplets due to the repulsion of the NPs from one another, thereby strengthening the mechanical barrier surrounding the oil droplets [36,44].

3.1.4. Fourier Transform Infrared Spectroscopy (FTIR)

FTIR was applied to examine the interactions between PP and KC in NPs fabricated from solutions at different concentrations. The infrared spectrum of PP (Figure 4) exhibits the characteristic absorption bands of the protein backbone, which is in agreement with previous research [17]. The addition of KC induces spectral shifting as a result of the interaction of polysaccharides with PP. The characteristic peak of O–H stretching bands shifted from 3275 cm⁻¹ in the PP sample to 3286 cm⁻¹ in the NPs, thereby indicating the formation of hydrogen bonds between the components [17,27]. In addition, a peak attributed to N-H stretching vibrations at 3060 cm⁻¹ is only seen in the protein sample and not in the NPs formed from PP-KC, thereby suggesting the formation of hydrogen bonds between KC hydroxyl groups and PP amide groups [27]. The shifting of amide I from 1635 cm⁻¹ in the PP sample to 1639 cm⁻¹ in the NP samples and amide II from 1537 cm⁻¹ in the PP sample to 1527 cm⁻¹ in the NP samples are indicative of hydrophobic and electrostatic interactions, respectively [2,27]. The peak seen at 1396 cm⁻¹ represents amide III [27] (C-N stretching and N-H bending) and does not shift with the addition of KC. Finally, it is possible to observe peaks at 1270–600 cm⁻¹ in the NP samples, which is the fingerprint region for the identification of specific polysaccharides, as analyzed by Şen and Erboz (2010) [45] and depicted in the spectra of KC in Figure 4. Altering the PP concentration did not lead to additional spectral changes (Figure S3).

3.2. Characterization of Emulsions

3.2.1. Stability and Accelerated Stability

The stability of PEs is largely affected by the oil-phase fraction [2]. To evaluate the influence of this parameter, a series of PEs stabilized with P2K25 NPs and with varying oil contents in the range of 20–70% was prepared, and their stability was assessed over time (Figure 5). When the oil fraction was below 40%, the emulsion phase-separated, and a separate oil layer was observed at the top of the samples after one week; this was in addition to the presence of a creaming phase. When the oil content was 40%, the oil layer appeared after approximately two weeks, and at 50–60% oil, the emulsions remained stable for over one month. This suggested the good emulsifying capability and resistance to phase separation of PEs stabilized with PP-KC NPs during storage [27]. Increasing the oil-phase volume fraction is beneficial for enhancing the stability of PE and preventing creaming [2]. When the oil-phase fraction was increased to 70%, the homogeneous structure was destroyed, and obvious sedimentation was observed (Figure 5, circled). In this case, the NP concentration was not sufficiently high to emulsify the high oil content in the sample. The occurrence of flocculation indicates that the NP concentration was insufficient to cover the droplet interface, thus causing the separation of oil and water phases [2]. In order to maintain stable emulsions that do not present creaming, the selected oil concentration in the following sections was set to 60%.

A homogeneous PE was obtained after the preparation of emulsions with 60% oil, regardless of PP or KC concentrations, in the solutions used for the fabrication of NPs. PEs were stable during storage for over one month (Figure 6). The similarity in the stability of the PEs can be attributed to the comparable zeta potential and wettability properties (Section 3.1.1 and Section 3.1.3, respectively) since these characteristics are considered to be indicative of the expected stability of the system. PEs that did not contain KC formed a cream phase after one week (Figure 6B). In agreement with our results, Wang et al. (2024) [27] found that the addition of KC effectively improved the visual appearance of emulsions, which was ascribed to the polysaccharide’s high viscosity and excellent emulsifying properties.

To assess the accelerated stability of the emulsions, PEs were subjected to 20 min of centrifugal force at 5000 RPM. Emulsions with 60% oil stabilized with all studied NP formulations remained stable during centrifugation, thereby yielding a top cream phase without oil release (Figure S4); therefore, the concentrated cream phase was further investigated. Further, concentrated PEs were obtained by removing the bottom aqueous phase. Table 2 presents the effect of PP and KC concentrations in the solutions used for NP fabrication in the oil phase fraction after centrifugation calculated from Equation (1). According to the ANOVA analysis, KC concentration had a significant effect (p < 0.05) on the volume of water released, while PP concentration did not significantly alter the amount of released water. A larger volume of water was released from PEs stabilized with NPs fabricated from PP solutions or solutions containing PP and a low concentration of KC (0.25% w/v). On the contrary, in PEs stabilized with NPs fabricated from solutions with high concentrations of KC (0.5% w/v), the volume fraction did not significantly change after centrifugation (p > 0.05). HIPPEs (φ > 0.74) were obtained for all samples in which the NPs were fabricated from solutions with 1–2% (w/v) PP, except those stabilized with P2K0.5. Among PEs stabilized with NPs prepared from solutions with 3% (w/v) PP, only P3K0 was able to form a HIPPE. Overall, the increase in KC concentration in the NP formulation provided increased resistance toward the creaming of the emulsions. This finding agrees with previous research that attributed enhanced stability to the larger thickness and improved elasticity of the interface layer around oil droplets [7]. In addition, the gel-like appearance of the PEs may indicate that excess KC is present within the water phase of the emulsion, thus further contributing toward preventing the formation of the cream phase and maintaining the original structure of the emulsion without releasing redundant water. Further analysis was performed with microscope imaging, as described in the next section (Section 3.2.2).

3.2.2. Confocal Laser Scanning Microscopy (CSLM)

CSLM was utilized to analyze the size of the oil droplets and the structure of the PEs fabricated using NPs with varying PP and KC concentrations, both before and after centrifugation. The microscope images demonstrate the formation of dense structures within the PEs, with the NPs enveloping the oil droplets to create O/W emulsions, as evidenced by the red fluorescence-stained oil droplets and the non-colored oil–water boundaries and continuous phase (Figure 7). These images demonstrate the ability of PP-KC to provide stability on the micron scale and are in line with the macroscopic stability studies described above (Section 3.2.1). The NPs formed from PP-KC tightly surround the oil droplets, forming a steric elastic barrier at the oil–water interface and providing an obstacle for the accumulation and flocculation of the oil droplets, as well as preventing coalescence and Ostwald ripening [2,27].

For the as-prepared PEs, the size of the oil droplets (red) stabilized by PP-KC NPs became smaller when the KC concentration for all PP concentrations studied was increased (Figure 7A). The higher content of KC may encourage the formation of a denser structure of closely formed oil droplets due to the high viscosity and emulsifying properties of the polysaccharide [2]. Qualitatively, the size of the droplets reduced as the content of KC increased, thereby supporting the suggestion that a denser network is formed within the aqueous phase. The density of the droplets increased with decreasing droplet size, which works to prevent coagulation and bridge flocculation. Smaller droplet sizes lead to improved rheological properties, such as viscosity and storage modulus, and printability of the emulsion by forming a gel-like network [2], thereby increasing the KC concentration and enhancing the emulsion performance (see Section 3.2.3). Similar results are found in the study by Liu et al. (2024) [2]. Increasing the concentration of PP also provided enhanced resistance toward the flocculation of the droplets when centrifugal force was applied compared to lower concentrations (Figure 7B).

As depicted in Figure 7 and Figure 8, both the PP and KC concentrations used for NP fabrication affect the oil droplet size. Figure 8 portrays the average droplet size before and after centrifugation. The smallest drop size was achieved at the highest NP concentration (P3K0.5), 7.8 μm ± 1.3 μm for the as-prepared PEs and 9.9 μm ± 0.2 μm after applying centrifugal forces. In certain cases, the centrifugation causes an increase in the droplet size, which is most likely due to coalesced oil droplets. This phenomenon is most significant when there is no KC, as evident in Figure 8A. For example, the average droplet size for PEs with NP concentration P2K0 increased from 25.1 μm ± 5.1 μm before centrifugation to 85.8 μm ± 6.3 μm after it. The increased droplet size indicates that NPs fabricated solely with PP could not sufficiently stabilize the emulsion and, therefore, centrifugation led to the creation of rather large droplets compared to the emulsions containing the polysaccharide in the NP formulation. While all emulsions did not phase-separate during the 20-min centrifugation, those containing KC were resistant to coalescence; therefore, the average drop size did not significantly vary after centrifugation (Figure 8B,C), which indicates the excellent stability of the emulsions fabricated with KC-PP NPs.

3.2.3. Rheological Measurements

The rheological properties of the emulsions are indicative of their ability to be applied in the field of extrusion-based printing [3]. While an ideal viscous liquid flows under applied stress and an ideal elastic solid deforms, many substances are viscoelastic, and their rheological behavior reflects features in the range between these two distinct types [38]. Oscillatory tests are a powerful tool for examining the viscoelastic behavior of a wide range of materials. The storage modulus (G′) represents the elastic character of the material and its ability to recover its original shape after an applied force, while the loss modulus (G″) is a measure of the deformation energy dissipated or lost by the material—that is, the liquid-state behavior of the sample [46]. For an emulsion to be classified as “printable”, it should display viscoelastic behavior with appropriate shear-thinning characteristics to enable smooth flow through the printer nozzle under applied shear force yet be able to quickly recover to its designated morphology [3,38,47].

Further, a strain amplitude sweep was performed to assess the behavior of the emulsions in the linear viscoelastic range (LVR), which is characterized by low strain values and, hence, represents the emulsion behavior at rest prior to printing [1,2]. G′ transcends over G″ in all samples in the linear range and up to the intersection point, as shown in Figure 9; this is characteristic behavior of a predominantly elastic, solid, gel-like material [1,2]. After the crossover point, G′ decreases quickly, thus indicating permanent structural deformation [27]. The intersection of the storage modulus G′ and the loss modulus G″ represents the change in the behavior of the material from “solid-like” at G′ > G″ to “liquid-like” at G′ < G″ [40]. The crossover point is observed for all studied emulsions. The PE stabilized with P3K0 (containing 3% (w/v) PP and 0% (w/v) KC, Table 1) had a rather low storage modulus of 0.6 [Pa] compared to PE stabilized with P3K0.25 and P3K0.5, which had a storage modulus of 146.5 [Pa] and 539 [Pa], respectively. The addition of KC to the NP formulation increased the elasticity and expanded the storage modulus in samples stabilized with NPs containing 1% and 2% (w/v) PP as well. In addition, the protein concentration in the solution from which the NPs were formed also had an impact on G′. For example, at a KC concentration of 0.5% (w/v), when raising the PP concentration from 1% (w/v) to 3% (w/v), G′ increased from 85.5 [Pa] to 539.0 [Pa].

The effect of centrifugation was also observed. When using NP formulation P2K0.25, centrifugation raised the storage modulus of the PE from 128.5 [Pa] to 669.6 [Pa]; other PP concentrations show a similar increase. The increased storage modulus induced by centrifugation is a consequence of creating a concentrated emulsion due to the removal of water [1,2]. In contrast, the value of G′ changes only slightly after centrifugation for PEs stabilized with NPs prepared from solutions that contain a high concentration of KC (0.5% (w/v)), regardless of PP concentration. As an example, for PEs stabilized with P2K0.5 NPs, G′ remains almost constant—488.7 [Pa] before centrifugation and 481.1 [Pa] afterward. Thus, it can be understood that NP compositions that lead to increased storage modulus after centrifugation are those that enable the elimination of water from the emulsion [1,2]—that is, containing 0.0–0.25% (w/v) KC (Section 3.2.1). Within the LVR, the loss modulus is much lower than the storage modulus for all studied PEs, which is indicative of the dominantly elastic behavior of the emulsion [3]. The loss modulus remains almost unchanged by centrifugal force. For example, the G″ of PE stabilized with NP formulation P2K0.5 was 66.5 [Pa] before centrifugation and 69.9 [Pa] after. Similar observations can be made for other samples [3]. It is noted that the storage modulus of PEs stabilized with NPs based solely on PP was very low, even after centrifugation. For example, the G′ of the emulsion stabilized by P3K0 NPs was raised by two orders of magnitude after centrifugation, from 0.5 [Pa] to 187.5 [Pa], but it was still significantly lower than the storage modulus of the emulsion stabilized by P3K0.5 and was measured to be 490.2 [Pa] (Figure 9A,C,E). It is highly desirable to have sufficiently high G′ for 3D printing, as this enables the material to support its shape and structure post-printing [1,25].

A frequency sweep at a constant and low strain of 0.5% in the linear viscoelastic region was conducted to evaluate the viscoelasticity of the PEs in the frequency range of 0.1–100 rad/s. As depicted in Figure 10, all the PEs presented a dominant elastic behavior (G′ > G″) across the entire studied frequency range. Moreover, it is possible to observe decreasing values of the moduli with decreasing concentrations of PP and KC in the solutions that form the NPs. All samples, both as-prepared and after applying centrifugation, revealed a fairly consistent value of the storage and loss moduli over a long range of angular frequency values and were minorly affected by the applied deformation, which is consistent with the gel-like behavior of the emulsions [7,27]. The emulsions after applying centrifugal force showed an increase in G′ and G″ compared to the as-prepared emulsions across all NP formulations. Before centrifugation, the storage modulus significantly increased with rising PP concentration, regardless of KC concentration. For example, the average storage modulus of emulsions stabilized with G1K0.5 was 88.5 [Pa], while an average value of 190.7 [Pa] was obtained for emulsions stabilized with G3K0.25. After centrifugation, it is apparent that the storage and loss modulus increase with increased PP concentration when KC concentration is 0.5% (w/v) but decrease with increased PP concentration in the solutions used to fabricate the NPs when KC concentration is 0.25% (w/v) (Figure 10D,E). This effect can be attributed to the higher final internal phase volume fraction of emulsions stabilized with NPs that have a low KC concentration, which is caused by removing a larger fraction of the water phase, as seen in Table 2 in Section 3.2.1.

After the viscoelastic properties of the PEs were studied, the flow properties of the emulsions were investigated; therefore, the viscosity of the samples was measured at increasing shear rates. All PEs display clear shear thinning performance and similar flow profiles (Figure 11). The apparent viscosity decreased by several orders of magnitude as the shear rate increased from 0.01 to 100 s⁻¹, which is likely due to the destruction of the internal structure of the emulsion with increasing shear rates. This indicates that the samples are expected to flow efficiently through the fine deposition nozzle at higher shear rates during extrusion printing [1,25]. Both PP and KC concentrations in the solutions used to fabricate the NPs affected the viscosity of the emulsions. The PEs stabilized by particles composed of PP-KC are more viscous than those stabilized by NPs composed of PP alone. For example, the viscosity at a low shear rate of 0.01 [1/s] is 7.9 [Pa·S] for PEs stabilized with P3K0 NPs, while using an NP formulation with 0.25% (w/v) KC increases the viscosity at the same shear rate to 777.3 [Pa·S]. A further increase in KC concentration to 0.5% (w/v) results in a viscosity of 2670.1 [Pa·S]. This finding is in line with a previous study that related the elevated viscosity to the gelling properties of KC and to interactions between PP and KC, which led to the formation of a tighter network of oil droplets within the emulsions [2]. The positive correlation between the concentration of solids used for NP fabrication and the viscosity of PE is likely due to the inhabitation of the free flow of the emulsion resulting from the denser, more rigid network structure and the larger contact surface area of oil droplets at higher NPs concentrations [7]. The rheological properties of the PEs, including high storage modulus and shear thinning, indicate that they are suitable for extrusion-based 3D printing. Additionally, their gel-like behavior may also provide mechanical strength and enhance the emulsions’ ability to resist shear forces, thereby improving their overall stability [2]. Compared to other emulsions stabilized with PP-KC [27], the viscosities observed in the current study are several orders of magnitude higher. This could be attributed to the higher concentration of solids present in the emulsions in this study (up to 3.5% vs. 0.5% (w/v)). When applying centrifugation, the viscosity most significantly increases when the NPs were prepared from solutions with low concentrations of KC or without KC (0–0.25%, w/v). Moreover, emulsions stabilized with NPs fabricated from solutions containing 0.5% (w/v) KC released the smallest amount of water; therefore, the centrifuge only minorly affected the viscosity values measured. This observation can be attributed to less effective water extraction in the presence of hydrophilic KC.

The value of n was obtained by fitting the power law (Equation (2)) to the experimental viscosity data, which is displayed in Table 3. The values of n < 1 were obtained for all measured PEs, thereby indicating that the emulsions portray shear-thinning. Moreover, the decreasing values of n with rising concentration of KC in the NPs formulation are indicative of a gelling effect within the continuous phase of the emulsion as well as interactions between PP and KC within the NPs, thereby enabling better adsorption of solids in the oil–water interface and improving the emulsifying capabilities [27].

For the as-prepared emulsions, altering the protein concentration used to prepare the NPs did not lead to a significant variation in the value of n, while increasing KC concentration decreased the value of n significantly (Table 3). The correlation between decreasing n and increasing KC concentration is evident across all protein concentrations and is in line with the higher viscosity of the samples when increasing KC concentration, thereby suggesting that the KC creates a more gel-like structure, as such a structure often displays dominant shear-thinning capabilities [48]. Despite this, the same trend is not observed in the study by Liu et al. (2024) [2]. After applying centrifugation, the value of n drastically decreases for almost all emulsions, apart from those stabilized with P3K0.25 and P3K0.5 NPs. Further, the difference in n values between these emulsions was non-significant (p > 0.05). In particular, emulsions that become HIPPEs following centrifugation (as seen in Table 2, Section 3.2.1) displayed significantly lower values of n than the corresponding as-prepared PEs, thereby strengthening the assumption that emulsions after centrifugation have valuable advantages in extrusion printing due to their significant shear-thinning characteristics [3]. Although the effect of centrifugation on emulsions stabilized with P2K0.5, P3K0.25, and P3K0.5 NPs is minimal, they still have promising capabilities to be 3D printed because they already display a low n value for as-prepared emulsions.

The thixotropic behavior of the PEs was explored with a five-cycle strain test. Thixotropic inks are highly desirable for extrusion printing. In these inks, the storage modulus is reduced rapidly, thereby causing the PEs to behave like liquids with the application of large amplitude oscillations that disrupt the structure of the PEs. Subsequently, they recover quickly at low shear, regaining their solid-like characteristics at small amplitude oscillations and returning to the initial emulsion properties [38,40]. A fast and reversible elastic response is crucial for emulsions in order to be suitable 3D printing materials, as they could be easily extruded while rapidly recovering mechanical strength to support the next extruded layer and the complete construct [47]. Figure 12 illustrates the shear recovery behavior of the storage and loss moduli of the emulsions under alterations of low (γ = 0.1%) and high (γ = 300%) shear strains. For all studied emulsions, G′ decreased significantly at high strain, while G″ became the dominant modulus, thereby indicating the liquid state of the emulsion. The PEs stabilized rapidly at a low shear strain, reaching a solid-like state (G′ > G″) within no more than five seconds. Applying centrifugation had a dominant effect on the storage modulus of samples with 0.25% (w/v) KC and a minor effect on samples with 0.5% (w/v) KC. For example, for PEs stabilized with P2K0.25 NPs, the storage modulus in the first cycle increased from 159.3 [Pa] when measured as prepared to 777.7 [Pa] after applying centrifugation, which is similar to other emulsions with 0.25% KC (Figure 12A,C). On the contrary, G′ in the first step measured for PEs with P2K0.5 NPs as prepared was 366.6 [Pa], while after centrifugation, the modulus was 409.0 [Pa]. An additional pronounced effect can be observed in the emulsions stabilized NPs fabricated from solutions with PP alone, without KC (Figure 12E), where the as-prepared emulsion displayed no recoverability and low storage modulus with a small dependence on the shear strain; in contrast, after centrifugation, the emulsion displayed a clear solid-to-liquid transition upon application of high strain, with recovery.

Table 4 summarizes the recovery after the first and second high-strain cycles as calculated from Equations (3) and (4). δ₁ is representative of the ability of the emulsion to regain a solid-like state after applying strain from the original state, while δ₂ models a repeated strain that is applied to an already reorganized state and the material’s ability to self-heal [40]. For the as-prepared emulsion samples, the first cycle of strain (δ₁) resulted in recovery values of 50–60%, while the second cycle (δ₂) resulted in significantly higher recovery values (85–90%). Moreover, δ₁ significantly increased after centrifugation compared to the as-prepared emulsions (p < 0.05) when the concentration of KC in the solution used to fabricate the NPs was 0.0–0.25% (w/v), while δ₂ was comparable for PEs before and after centrifugation. No clear correlations could be defined between the recoverability and concentrations of PP and KC in the solutions used to prepare the NPs before applying centrifugation. After centrifugation, there is a prominent increase in thixotropic recovery in the first cycle (δ₁) for emulsions stabilized by NPs with low KC concentration due to the ability to effectively remove water from the emulsion. Recovery values of over 70% are considered suitable for 3D printing, thus strengthening the advantage of the emulsions undergoing centrifugation to promote their recoverability [3].

3.3. 3D Printing

When selecting samples for 3D printing, it is important to consider thixotropic behavior, viscosity, G′ values, and emulsion stability [49]. For the current study, emulsions stabilized with P3K0.5 NPs (fabricated from solutions containing 3% (w/v) PP and 0.5% (w/v) KC, see Table 1) were selected. These compositions showed promising characteristics, including high recovery values (δ₂ = 91–93%), high viscosity at rest (η > 10³ [Pa·s]), shear-thinning characteristics, and appropriate storage modulus to serve as 3D bioink within the LVR (490–540 [Pa]) [25,47,50]. Preliminary experiments revealed that the printing of structures from PEs NPs with lower KC concentration did not have sufficient mechanical strength to support and maintain the printed design (Figure S5). Increasing the concentration of KC beyond 0.5% may improve the printing characteristics but could impair the flavor and texture of the final product as well [51]. The printing performance of as-prepared emulsions was compared to the concentrated emulsions after applying centrifugal force with an aim to observe the effect of the centrifugal force despite the similar rheological properties of these emulsions.

The samples were extruded continuously and smoothly through the printer nozzle due to the shear-thinning properties of the emulsion, thus creating a filament (Figure 13A). Three hollow cylinders with different heights were printed to visually examine the stability of the construct (Figure S6). Printing cylinders with as-prepared emulsions did not show adequate self-support. On the contrary, when using concentrated emulsions, the printed hollow cylinders were stable without post-printing crosslinking. Cylinders with heights lower than 9 mm were stable for several minutes, while taller cylinders collapsed. The stable hollow shapes provide a visual demonstration of the solid-like properties and stability of the construct [2]. Next, in order to obtain quantitative insight regarding the impact of centrifugation on 3D-printed structures, a rectilinear grid pattern was printed. In order to assess the stability and shape fidelity of the different emulsions (Figure 13C,D), the printing index and fidelity indexes were calculated based on Equations (5)–(7). 20 × 20 mm printed grids showed a close similarity to the geometrical template and did not spread or smear when printed, both for the as-prepared PEs and those after centrifugation. On the other hand, the inner squares within the 10·10 mm structure were not measurable without applying centrifugation, indicating the advantages of centrifuged PEs. Therefore, only the 20·20 mm printed grids were quantitively assessed. Using the printing index and fidelity assessments, a measurement of the reproducibility and consistency of the printed emulsion can be achieved [41]. The calculated P_r index and fidelity values were significantly (p < 0.05) closer to one for the concentrated emulsion compared to the as-prepared emulsion. In addition, the as-prepared emulsions exhibited filament breakage in a few samples, while the PEs after centrifugation produced a continuous and uniform structure. While the rheological properties of the samples showed similar recovery and viscosity profiles before and after undergoing centrifugation, the centrifuge improved the printing fidelity and contributed to the visual appearance and shape-precision of the emulsions after printing, highlighting its advantage in creating sustainable, food-grade PEs.

Based on these results, PEs showed the ability to serve as a bioink in food printing technologies. In addition, this work demonstrates the capability of centrifugation to improve the stability and self-support of the emulsion ink. The ability to print large objects onto a hard surface opens new possibilities for utilizing these PEs in 3D printing applications, such as fat substitutions and personalized food products. For example, Song et al. (2023) [3] loaded astaxanthin into PEs and 3D printed them, while C. Li et al. (2023) [52] utilized a similar emulsion as a pork fat replacement. The implementation of these PEs can provide a robust basis for several applications, such as healthier, tunable, and stable delivery systems and fat substitutions. Exploring the potential of the emulsions as carriers of hydrophobic substances in order to increase their bioavailability or incorporating the PEs in vegan meat products as a fat substitute that is not animal derived are some of the possibilities in the scope of future research.

4. Conclusions

This study presented the fabrication and characterization of NPs from PP and KC and investigated the effect of the protein and polysaccharide concentration on the performance of PEs stabilized with those NPs. The morphology study revealed highly stable oil-in-water emulsions with shear-thinning capabilities, dominant storage modulus, and high recoverability. CSLM, surface tension, and rheological experiments provided insight into the important roles of PP and KC NPs in creating a strong physical barrier at the oil–water interface, which works against phase separation of the emulsions and provides a promising basis for the research and development of several necessary, sustainable, food-grade applications such as an animal-free, non-saturated fat replacement. The study further analyzed the effect of centrifugation on the properties and stability of PEs, which had scarcely been methodically compared earlier, revealing the advantages of creating concentrated emulsions. The 3D-printed PEs showed smooth filament extrusions as well as high stability, self-support, and printing fidelity. The outcomes of this paper can be implemented to establish a promising, low-cost, and food-grade saturated fat alternative that can be 3D printed and applied in novel food solutions as well as utilized in loading and controlled release of hydrophobic substances to increase their bioavailability.

Author Contributions

Conceptualization, G.H., S.L. and H.B.-P.; writing—original draft preparation, G.H.; writing—review and editing, G.H., N.H., S.L. and H.B.-P.; supervision, H.B.-P.; project administration, S.L. and H.B.-P.; funding acquisition, S.L. and H.B.-P. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article/Supplementary Material. Further inquiries can be directed to the corresponding author(s).

Conflicts of Interest

The authors declare no conflict of interest.

Footnotes

Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Figures and Tables

Figure 1. (A) Mean hydrodynamic diameter and (B) zeta potential. Different letters within the same graph indicate significant differences (p [less than] 0.05).

Figure 2. Cryo-TEM images: (A,C) P1K0.25, and (B) P2K0.25. Scale bar 100 [nm].

Figure 3. (A) Three-phase contact angle measurements of NPs with different compositions. (B) Surface tension. Different letters within the same graph indicate a significant difference (p [less than] 0.05) in surface tension.

Figure 4. FTIR spectra of NPs formed from solutions with 3% (w/v) PP with 0.25–0.5% KC (w/v), compared to the spectra of PP and KC.

Figure 5. Oil concentration series: (A) Emulsions after preparation, (B) after one week, (C) after two weeks, and (D) after one month. NPs concentration P2K0.25, oil volume fraction φ = 20–70% (v/v). Stored at 4 °C. Emulsions circled in red exhibit phase separation.

Figure 6. Series of the concentration of solids: (A) Emulsions after preparation, (B) after one week, (C) after two weeks, and (D) after one month. Oil volume fraction φ = 60% (v/v), PP = 1%, 2%, and 3% (w/v), KC = 0.0, 0.25, and 0.5% (w/v). Stored at 4 °C.

Figure 7. CLSM images of emulsions of different concentrations (A) as-prepared and (B) after undergoing centrifugation. The scale bar represents 20 μm.

Figure 8. Size analysis of oil droplets at a KC concentration of (A) 0.0% (w/v), (B) 0.25% (w/v), and (C) 0.5% (w/v), as prepared (light blue) and after centrifugation (dark blue). Different letters within the same graph indicate significant differences (p [less than] 0.05).

Figure 9. Amplitude sweeps of PEs as-prepared and after applying centrifugation: G′ (full symbols) and G″ (open symbols) as a function of strain. (A) PEs stabilized P3K0 NPs: as-prepared (light blue) and after centrifugation (dark blue). (B) As-prepared PEs stabilized with P3K0.25 (blue), P2K0.25 (green), or P1K0.25 (gray) NPs, (C) As-prepared PEs stabilized with P3K0.5 (blue), P2K0.5 (green), or P1K0.5 (gray) NPs (D) PEs stabilized with P3K0.25 (blue), P2K0.25 (green), or P1K0.25 (gray) NPs after centrifugation. (E) PEs stabilized with P3K0.5 (blue), P2K0.5 (green), or P1K0.5 (gray) NPs after centrifugation.

Figure 10. Frequency sweep of PEs as-prepared and after applying centrifugation. G′ (full symbols) and G″ (open symbols) as a function of angular frequency. (A) PEs stabilized by P3K0 NPs, as-prepared (light blue) and after centrifugation (dark blue). (B) As-prepared PEs stabilized by P3K0.25 (blue), P2K0.25 (green), or P1K0.25 (gray) NPs. (C) As-prepared PEs stabilized by P3K0.5 (blue), P2K0.5 (green), or P1K0.5 (gray) NPs. (D) PEs stabilized by P3K0.25 (blue), P2K0.25 (green), or P1K0.25 (gray) NPs after centrifugation. (E) PEs stabilized by P3K0.5 (blue), P2K0.5 (green), or P1K0.5 (gray) NPs after centrifugation.

Figure 11. Viscosity of as-prepared emulsions compared to the same formulations after centrifugation (AC). (A) PEs P3K0 (blue), P2K0 (green), or P1K0 (grey) NPs. (B) PEs stabilized by P3K0.25 (blue), P2K0.25 (green), or P1K0.25 (grey) NPs. (C) PEs stabilized by P3K0.5 (blue), P2K0.5 (green), or P1K0.5 (grey) NPs.

Figure 12. Shear recovery of PEs as-prepared and after applying centrifugation: G′ (full symbols) and G″ (open symbols) as a function of time. (A) PEs stabilized with P3K0 NPs: as-prepared (light blue) and after centrifugation (dark blue). (B) As-prepared PEs stabilized P3K0.25 (blue), P2K0.25 (green), or P1K0.25 (gray) NPs. (C) As-prepared PEs stabilized by P3K0.5 (blue), P2K0.5 (green), or P1K0.5 (gray) NPs. (D) PEs stabilized by P3K0.25 (blue), P2K0.25 (green), or P1K0.25 (gray) NPs after centrifugation. (E) PEs stabilized by P3K0.5 (blue), P2K0.5 (green), or P1K0.5 (gray) NPs after centrifugation.

Figure 13. (A) Representation of continuous filament extrusion using PE bioink. (B) 3D printing of rectilinear grids: 10·10 mm (left) and 20·20 mm (right). (C) computer-aided design (CAD) of the printed structures; scale bar 5 mm. (D) The obtained printed structures of as-prepared PE and (E) 3D printed PE after centrifugation. PEs are stabilized with P3K0.5 NPs, scale bar: 10 mm. (F) Calculation of the Pr index and fidelity parameters of as-prepared PEs compared to emulsions after centrifugation (n = 5), * p [less than] 0.05, ** p [less than] 0.01.

Table 1

Concentration abbreviations.

Abbreviation	PP Conc. (% w/v)	KC Conc. (% w/v)
P1K0	1.00	0.00
P1K0.25		0.25
P1K0.5		0.50
P2K0	2.00	0.00
P2K0.25		0.25
P2K0.5		0.50
P3K0	3.00	0.00
P3K0.25		0.25
P3K0.5		0.50

Table 2

Final internal phase volume fraction V_{final, oil} after centrifugation.

Concentration	Vfinal, oil
P1K0	0.84 ± 0.05 ab
P1K0.25	0.84 ± 0.03 ab
P1K0.5	0.68 ± 0.08 cd
P2K0	0.84 ± 0.02 ab
P2K0.25	0.80 ± 0.08 abc
P2K0.5	0.66 ± 0.08 cd
P3K0	0.87 ± 0.03 a
P3K0.25	0.71 ± 0.05 bd
P3K0.5	0.60 ± 0.00 d

Initial oil concentration—60%; different letters indicate a significant difference in V_final,oil values; p < 0.05.

Table 3

Power law exponent (n) for as-prepared emulsions and emulsions after centrifugation.

NP Formulation	n (As-Prepared)	n (After Centrifugation)	Φ > 0.74 (HIPPE)
P1K0	0.70 ± 0.08 a, A	0.10 ± 0.02 b	Yes
P1K0.25	0.48 ± 0.04 a, BC	0.10 ± 0.05 b	Yes
P1K0.5	0.37 ± 0.04 a, CD	0.24 ± 0.02 b	No
P2K0	0.60 ± 0.08 a, AB	0.14 ± 0.03 b	Yes
P2K0.25	0.34 ± 0.02 a, CDE	0.11 ± 0.09 b	Yes
P2K0.5	0.28 ± 0.01 a, DE	0.20 ± 0.04 b	No
P3K0	0.63 ± 0.11 a, AB	0.11 ± 0.04 b	Yes
P3K0.25	0.29 ± 0.02 a, DE	0.19 ± 0.09 a	No
P3K0.5	0.18 ± 0.02 a, E	0.19 ± 0.03 a	No

R² > 0.97 for fitting to the power law, Equation (2). Different lowercase letters in the same row indicate a significant difference in n. Different uppercase letters in the same column indicate a significant difference in n. p < 0.05.

Table 4

Recovery values for as-prepared emulsions and after centrifugation.

NPs Formulation	As-Prepared		After Centrifugation
	δ1 [%]	δ2 [%]	δ1 [%]	δ2 [%]
P1K0.25	50.1 ± 11.3 b, A	101.9 ± 7.5 a, A	92.1 ± 2.9 a, A	95.3 ± 1.9 a, AB
P1K0.5	63.0 ± 6.7 b, A	91.4 ± 3.1 a, A	65.1 ± 16.9 b, C	94.2 ± 7.9 a, AB
P2K0.25	54.0 ± 12.9 b, A	87.1 ± 7.8 a, A	85.6 ± 5.4 a, AB	98.8 ± 0.3 a, A
P2K0.5	61.1 ± 5.9 b, A	91.6 ± 3.8 a, A	53.8 ± 2.8 b, C	88.5 ± 3.2 a, B
P3K0	1.5 ± 0.7 b, B	86.7 ± 14.7 a, A	90.4 ± 3.2 a, AB	97.2 ± 2.0 a, AB
P3K0.25	56.0 ± 5.6 c, A	85.7 ± 1.3 a, A	70.9 ± 6.7 b, BC	95.5 ± 2.3 a, AB
P3K0.5	55.9 ± 8.4 b, A	93.8 ± 1.0 a, A	61.1 ± 1.3 b, C	91.2 ± 0.9 a, AB

Different lowercase letters in the same row indicate a significant difference in recovery. Different uppercase letters in the same column indicate a significant difference in recovery (p < 0.05).

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/polysaccharides6010014/s1, Figure S1: DLS size distributions for NPs formed from PP-KC solutions; Figure S2: Contact angles of different NP formulations; Figure S3: FTIR spectra of NPs samples; Figure S4: As-prepared PEs, PEs after centrifugation and graphic representation; Figure S5: additional 3D printed structures; Figure S6: additional 3D printed structures and CAD of hollow cylinders.

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