Method of encapsulation by electrohydrodynamics

Abstract

The present disclosure relates to a process for electrostatic spray drying of living microorganisms having a specific surface charge using an electrostatic charge provided by a high voltage source, the electrostatic charge being the same as the surface charge of the microorganisms.

Description

Technical Field

[0001] The present disclosure relates to a method for electrostatic spray drying of microorganisms and to microorganisms incorporated into dried particles.

Background Art

[0002] Spray drying is a technique in which a suspension is atomized (or sprayed) and then rapidly dried by use of a hot gas drying medium. The scalability of the manufacturing process enables the formation of particles intended for use in a wide number of industries including food, polymers, biotechnology, pharmaceuticals, and medicine. The choice of atomizer that breaks down the feedstock (suspension) into droplets depends largely on the type of solution and the desired characteristics of the dried particles. Conventional atomizers include rotary atomizers that rely on the use of centrifugal force for droplet formation, hydraulic nozzle atomizers that rely on pressure, and pneumatic nozzle atomizers that rely on kinetic energy. Spray drying has been applied to a wide number of particle types including bacteria and other microorganisms.

[0003] Electrospray is an electrohydrodynamic method used for the encapsulation of compounds or bacterial cells in a polymer matrix. In recent years, electrostatic droplet formation has been disclosed as an improved method for achieving droplet formation. The method relies on electrostatically charging the suspension prior to droplet formation. Charging of the suspension can result in improved drying characteristics of the formed droplets and can facilitate particle collection and aggregation. Electrostatic spray drying offers a significant number of advantages over conventional spray drying, but its use and application for the drying of microorganisms has not yet been successful. This is reasonable since the viability of microorganisms is affected by electrostatic charge and, indeed, high voltage is a common method for sterilizing liquids. Thus, the use of electrostatic spray drying has been mostly concentrated on the drying of inanimate substances.

[0004] The patent publication of International Publication No. 2021152111 (A1) discloses a method for the electrostatic spray drying of living microorganisms. However, there is still a need to provide an improved method for electrostatic spray drying, particularly a method that results in cost savings and / or improved stability of the final product.

Summary of the Invention

Problems to be Solved by the Invention

[0005] It has been observed that the electric field and charge polarity during spray drying can be optimized to improve the encapsulation / immobilization of bioactive compounds such as microorganisms by affecting the nano-microstructure using electrohydrodynamics.

[0006] The present invention provides an improved method for the electrostatic spray drying of living microorganisms by utilizing a high-voltage source of negative charge to encapsulate microorganisms having a negative surface charge.

Means for Solving the Problems

[0007] Microorganisms can inherently have a negative surface charge or can be treated, for example, to obtain a negative surface charge.

[0008] According to a first aspect of the present invention, a process for the electrostatic spray drying of living microorganisms is provided, the process comprising the following steps: a) providing a suspension comprising microorganisms having a surface charge and a formulation aid; b) applying an electrostatic charge to the suspension; c) forming droplets of the suspension; d) forming dry particles by drying the droplets; and e) collecting the dry particles, wherein the electrostatic charge has the same polarity as the surface charge of the microorganisms.

[0009] In one embodiment, the electrostatic charge is applied to the suspension while simultaneously moving the suspension.

[0010] In one embodiment, the collection of the dry particles is achieved using a collector having a charge opposite to the electrostatic charge and the surface charge of the microorganism.

[0011] In one embodiment, the microorganism having a negative surface charge is selected from the group consisting of Bifidobacterium animalis, Lactobacillus acidophilus, Pseudomonas aeruginosa, and Escherichia coli.

[0012] In one embodiment, the microorganism is Bifidobacterium animalis subsp. lactis (DSM15954).

[0013] In one embodiment, the shape of the dry particles is spherical, fibrous, helical, rod-shaped, or hybrid.

[0014] In one embodiment, the formulation aid is a polysaccharide, protein, lipid, or synthetic polymer, or a mixture thereof.

[0015] In one embodiment, the polysaccharide is either positively charged, negatively charged, or neutral.

[0016] In one embodiment, the polysaccharide is selected from the group consisting of maltodextrin, starch, xanthan, gellan, alginic acid, pectin, glucan, and chitosan.

[0017] In one embodiment, the protein is milk protein, non-milk animal protein, plant protein, algal protein, or fermentation product protein.

[0018] According to a second aspect of the present invention, there is provided the use of a high voltage source of positive or negative charge for encapsulating a living microorganism, the microorganism having the same surface charge as the charge of the high voltage source.

[0019] According to a third aspect of the present invention, there is provided a particle comprising a polymer and at least one cell of a living microorganism, the at least one cell being located in the core of the particle, and the composition of the polymer of the particle being uniform and forming a shell around the core substantially free of cells.

[0020] According to a fourth aspect of the present invention, there is provided a particle comprising a living microorganism, which can be obtained by the process according to the first aspect.

[0021] According to a fifth aspect, there is provided the use of the particles according to the third or fourth aspect for producing a food product, a feed product, a dietary supplement, or a pharmaceutical product.

[0022] According to a sixth aspect, there is provided a food product, a feed product, a dietary supplement, or a pharmaceutical product comprising the particles according to the third and fourth aspects.

Brief Description of the Drawings

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[0041] The inventors have found that by utilizing electrohydrodynamics, encapsulation of microorganisms can be achieved, which results in cost savings and / or improvement of process efficiency and / or improvement of the stability of the final product.

[0042] This is achieved through a process for the electrostatic spray drying of live microorganisms, the process comprising providing a suspension comprising microorganisms having a surface charge and a formulation aid, applying an electrostatic charge to the suspension, forming droplets of the suspension, forming dried particles by drying the droplets, and collecting the dried particles. The electrostatic charge should have the same polarity as the surface charge of the microorganisms. This results in particles comprising a polymer and at least one cell of a live microorganism, wherein the at least one cell is located in the core of the particle and the polymer composition of the particle is uniform and forms a shell around the substantially cell-free core. Such a particle structure protects the microorganism and thereby increases the stability of the product.

[0043] It is also possible to utilize electrohydrodynamics to achieve particles having microorganisms close to the surface rather than at the core of the particles. This is achieved by applying an electrostatic charge that is opposite to the surface charge of the microorganisms.

[0044] Preferably, the suspension comprises one or more formulation aids. At least a portion of the formulation aids can be added to the suspension prior to the application of the electrostatic charge to the suspension. The formulation aids are typically provided as cryoprotectants, which function to stabilize the microorganisms within the suspension, droplets, and / or dried particles. Preferably, the cryoprotectant is selected to function to reduce, e.g., prevent, the killing of the microorganisms not only during the electrostatic spray drying process itself but also during subsequent use of the dried particles and / or microorganisms, including storage, transportation, and / or additional processing.

[0045] The present invention further relates to particles comprising viable microorganisms incorporated into a mass of formulation aids. The particles are preferably densely packed such that the total internal void volume is less than 5% of the total volume of the particles. Further, in a preferred embodiment of the present disclosure, the microorganisms are not present on the surface of the particles. Thus, the microorganisms are preferably incorporated within the formulation aids of the dried particles. In an embodiment of the present disclosure, the dried particles comprise, in addition to the microorganisms, a substantially single phase. Thereby, preferably, the dried particles are not layered dried particles, such as dried uniform particles encapsulated by a protective layer. In an exemplary embodiment of the present disclosure, the dried particles have a substantially continuous radial gradient of the formulation aids. Thereby, the dried particles can have a high concentration of the formulation aids, e.g., 100%, at the surface. The concentration of the formulation aids continuously decreases towards the center of the particles, thereby forming a radial gradient of the formulation aids. Preferably, the center of the particles has the highest concentration of the microorganisms.

[0046] In embodiments of the present disclosure, an electrostatic charge is applied such that the polar component is pushed towards the surface of the droplet while the less polar component of the suspension is pushed towards the center of the droplet. Typically, the polarity of the various components affects the resulting electrostatic charge distribution with respect to the formed droplet. For example, the electrons provided to the suspension are typically associated with the more polar component, usually the solvent, and result in an electrostatic repulsion between the polar component that pushes the polar solvent and the polar formulation aid dissolved in the solvent towards the surface of the droplet. A similar effect can further function to push the microorganisms encapsulated by the formulation aid towards the center of the droplet. Pushing the solvent towards the surface typically results in a faster evaporation rate, while incorporating the microorganisms into the center of the droplet results in an improvement in the encapsulation of the microorganisms.

[0047] In embodiments of the present disclosure, the electrostatic charge is applied to the suspension by an electrode in contact with the suspension, and the electrode has a pulsed potential difference with respect to ground. Preferably, the potential difference, i.e., the voltage, has a constant polarity. Thereby, the electrode can be part of a direct current (DC) circuit, the current flows in only one direction, and the electrode is always maintained negative or positive. Thereby, all of the formed droplets can have an overall negative charge or an overall positive charge. Typically, the pulse of the potential difference results in an improvement in the electrostatic charging of the suspension, and further, it can result in an improvement in the properties of the dried particles.

[0048] In embodiments of the present disclosure, the electrostatic charge is applied to the suspension by contacting the suspension with at least one electrode having a potential difference, i.e., a voltage, with respect to ground. Thus, the electrode can be provided in a configuration to apply the voltage to the suspension. To apply the electrostatic charge to the suspension, additional electrodes, for example, two, three, or four, or even more additional electrodes can be provided. Preferably, the two or more electrodes have the same polarity, such as a positive or negative voltage as provided in a DC circuit. The configuration of the electrodes for applying the electrostatic charge is known to those skilled in the art and can include the use of specific materials, surface areas, and shapes for the one or more electrodes.

[0049] In embodiments of the present disclosure, the electrode has a potential difference with respect to ground of less than about 40 kV, such as less than about 35 kV, such as less than about 30 kV, such as less than about 25 kV, such as less than about 20 kV, such as less than about 15 kV, such as less than about 10 kV. Preferably, the voltage is low enough so as not to cause any damage to the microorganisms. Thereby, the voltage should be low enough so as not to kill the living microorganisms.

[0050] In embodiments of the present disclosure, the electrode has a fixed polarity with respect to ground, such as a fixed negative polarity or a fixed positive polarity. Thus, the electrode or electrodes can be configured to apply a direct current (DC) voltage. Preferably, the electrode is configured for continuous supply or continuous discharge of electrons to / from the suspension. Typically, the positive electrode, i.e., the anode, discharges electrons from the suspension, while the negative electrode, i.e., the cathode, supplies electrons to the suspension. The electrode can temporarily have a ground potential, i.e., 0 V.

[0051] In embodiments of the present disclosure, the potential difference of the electrode with respect to ground is constant. Thereby, the voltage of the electrode can be constant, and thus, an electrostatic charge can be applied to the suspension by using a constant voltage.

[0052] In another embodiment of the present disclosure, the potential difference of the electrode with respect to ground changes over time, such as by periodic variations. The periodic variations can be represented by a wave function such as a sine wave, or by a combination of multiple wave functions combined to perform periodic variations. The periodic variations can be represented by two or more voltage levels at which the voltage of the electrode changes in a periodic variation. One of the voltage levels can be ground.

[0053] In embodiments of the present disclosure, the potential difference of the electrode with respect to the ground changes periodically, for example, in a periodic step function. The voltage of the electrode can change according to any function and can further depend on method parameters, for example, the supply rate of the suspension, the droplet size, the content of the suspension, for example, the type of component and the relative ratio of the component, and further, the desired parameters of the dry particles.

[0054] In embodiments of the present disclosure, the potential of the electrode is applied by pulse width modulation such as a rectangular wave. As a result, the voltage of the electrode can change between two or more set levels that form a rectangular wave, and the time between two pulses can be a set value or can change according to the parameters of the processing method as mentioned elsewhere in this specification. The voltage can be provided as pulses between two or more voltage values, the dwell time at each level can be set individually, and one of the voltage levels can be 0V.

[0055] In embodiments of the present disclosure, the components of the suspension are classified within the formed droplets with respect to their polarity, for example, for the improvement of the evaporation of the solvent and / or the improvement of the encapsulation of microorganisms. The classification of the components of the suspension can result in advantageous properties of the formed droplets, for example, the improvement of evaporation, for example, the reduction of the evaporation time and / or a lower water content in the final dry particles, and / or the improvement of the encapsulation of the dry particles. Therefore, the components of the suspension can be selected based on how the components are classified within the formed droplets, for example, electrostatically charged formed droplets.

[0056] In embodiments of the present disclosure, the components of the suspension with higher polarity are classified on the surface of the droplet, and the components of the suspension with lower polarity are classified at the center of the droplet. In embodiments of the present disclosure, at least two components of the suspension have different dielectric properties. Preferably, there is a relationship between the classification of the components of the suspension within the droplet and the dielectric properties and / or effective dielectric properties of the components of the suspension.

[0057] In embodiments of the present disclosure, the microorganism has an effective dielectric property lower than that of the formulation aid and / or the solvent. In embodiments of the present disclosure, the solvent has a higher dielectric constant than the formulation aid, and the formulation aid has a higher dielectric constant than the microorganism. Among the solvent, the formulation aid, and the microorganism, it is preferable that the solvent has the highest dielectric property while the formulation aid has a higher dielectric property than the microorganism. The dielectric property of the microorganism is measured and / or provided as an effective dielectric property given by the overall dielectric property of the microorganism rather than the dielectric properties of individual components such as specific membrane proteins, as is known to those skilled in the art, for example, in "Determination of the Dielectric Properties of Bacterial Cells Using Dielectrophoresis" by Sanchis et al. in Bioelectromagnetics in 2007.

[0058] In embodiments of the present disclosure, the droplets are formed by atomizing a suspension. Atomization and spraying can be used interchangeably herein to refer to the process of forming a plurality of small droplets of a liquid, such as a suspension, from a larger volume such as a feedstock. Droplet formation, i.e., the decomposition of the liquid volume into smaller droplets, requires energy due to the increase in surface area. Typically, the interfacial energy given at the liquid-air interface on the surface of the droplet requires the addition of energy for formation. As is known to those skilled in the art, the energy can be supplied in a wide range of ways.

[0059] In embodiments of the present disclosure, the formation of droplets is carried out by an atomization device, such as an ultrasonic nozzle, a pressure nozzle, a two-fluid nozzle (e.g., using CO2 or N2 or other gas as the atomizing gas), a vibrating nozzle, a vibration nozzle, an electrostatic nozzle, or a rotary atomization device. Various types of nozzles are known to those skilled in the art, and the nozzles can have their respective advantages. Several parameters, including the flow rate of the atomizing gas, the flow rate of the suspension / feedstock, the use of a surfactant, the configuration of the nozzle, the type of the nozzle, and the forces acting on the suspension (gravity, electric force, centrifugal force, or other forces), can be adjusted based on the desired properties of the droplets and thus the dried particles.

[0060] In embodiments of the present disclosure, the formation of droplets is performed by a two-fluid nozzle. The two-fluid nozzle atomizes a liquid such as a suspension by the interaction between a high-speed gas and a liquid such as a suspension. Typically, a compressed gas is used as the atomizing gas, but other gases such as vapor can be used. The two-fluid nozzle can be of an internal mixing type or an external mixing type depending on the mixing point of the gas and liquid flows with respect to the nozzle surface.

[0061] In embodiments of the present disclosure, the formation of droplets in step c) is performed using an atomizing gas. Preferably, the atomization of the droplets is performed by using a two-fluid nozzle configured for droplet formation by the use of the atomizing gas.

[0062] In embodiments of the present disclosure, the atomizing gas is selected from the group consisting of inert gases (such as nitrogen and carbon dioxide), noble gases (e.g., helium, argon, or neon), and alkane gases (such as methane), or mixtures thereof.

[0063] In embodiments of the present disclosure, the atomizing gas comprises or consists of nitrogen, carbon dioxide, and / or atmospheric gas, or mixtures thereof. The gas can be treated prior to use by any suitable method including filtration, sterilization, and / or dehumidification. However, in embodiments of the present disclosure, the atomizing gas is not dehumidified. The use of non-dehumidified gas can be advantageous in several aspects including providing simpler operation, reducing cost and time, and further resulting in better droplet and / or dry particle characteristics.

[0064] In embodiments of the present disclosure, the atomizing gas has a moisture content of less than about 1000 ppm, such as less than about 500 ppm, such as less than about 100 ppm, such as less than about 50 ppm, such as less than about 10 ppm.

[0065] In embodiments of the present disclosure, the droplet formation step (e.g., the spraying step) is performed using a nebulized gas inlet temperature that is at most about 200°C, for example, in the range from about 20°C to about 200°C, for example, in the range from about 40°C to about 150°C, or for example, in the range from about 40°C to about 120°C, for example, from about 40°C to about 90°C, for example, from about 50°C to about 90°C, for example, from about 60°C to about 85°C, for example, about 80°C. It may be desirable to have a temperature higher than standard room temperature for optimal droplet formation, solvent evaporation, and / or dry particle properties. However, at the same time, a high temperature may be unfavorable to components of the suspension such as microorganisms. Thus, the optimal temperature can be based not only on the simplest mode of performing the process, such as room temperature, but instead on a plurality of factors including the ease of performing the process, rapid evaporation, viability of living microorganisms, specific properties of the dry particles, such as optical encapsulation of microorganisms, a sufficiently small volume of gas-filled voids (compactness of the dry particles) within the dry particles, and other parameters. Thus, the optimal temperature is not trivial and is based on several factors. Preferably, the temperature is above room temperature, for example, for rapid evaporation, but preferably below a temperature unfavorable to microorganisms, taking into account the specific temperature experienced by the microorganisms when they are thermally isolated during drying by being incorporated into the formulation aid and at least temporarily into the solvent. Thus, the temperature can be in the range from about 20°C to about 200°C.

[0066] In embodiments of the present disclosure, the nebulized gas has an inlet pressure in the range from about 1 kPa to about 500 kPa, for example, in the range from about 5 kPa to about 500 kPa, for example, in the range from about 5 kPa to about 300 kPa, for example, in the range from about 5 kPa to about 100 kPa, for example, about 60 kPa, for example, about 70 kPa, or for example, about 80 kPa, or for example, in the range from about 100 kPa to about 400 kPa, for example, about 120 kPa, about 150 kPa, about 200 kPa, about 250 kPa, about 300 kPa, or about 350 kPa.

[0067] In embodiments of the present disclosure, the atomized gas has an inlet pressure in the range from about 50 kPa to about 400 kPa. Typically, the inlet pressure of the atomized gas is defined as the pressure of the atomized gas before being supplied to a nozzle such as a two-fluid nozzle. Typically, the inlet pressure of the atomized gas is the pressure of the atomized gas supplied to a spray drying apparatus such as an electrostatic spray drying apparatus. The gas can be supplied from a gas tank comprising at least one pressure regulator that controls the pressure provided to the spray dryer and / or the nozzle. Preferably, the pressure supplied to the nozzle is substantially the same as the inlet pressure. After droplet formation, the pressure of the atomized gas typically decreases due to the high resistance in the atomizing nozzle combined with the large cross-section of the drying chamber.

[0068] In embodiments of the present disclosure, the dried particles have a size in the range from about 1 micrometer to about 800 micrometers, for example from about 5 micrometers to about 800 micrometers, for example from about 10 micrometers to about 600 micrometers, for example from about 10 micrometers to about 300 micrometers, for example from about 10 micrometers to about 200 micrometers, for example from about 10 micrometers to about 50 micrometers, or for example from about 50 micrometers to about 200 micrometers, for example from about 50 micrometers to about 100 micrometers, for example about 75 micrometers, or for example from about 100 micrometers to about 200 micrometers, for example about 150 micrometers, as measured by the Dv50 value. As is known to those skilled in the art, Dv50 is typically used to indicate the center of the volume distribution.

[0069] In embodiments of the present disclosure, the size distribution of the dried particles is substantially unimodal.

[0070] In embodiments of the present disclosure, the dried particles are substantially dry. A small amount of solvent such as a trace amount of solvent may be present in the dried particles. Thus, in embodiments of the present disclosure, the process includes a drying step of drying the formed droplets.

[0071] In embodiments of the present disclosure, the process includes a drying step in which the drying of the droplets is carried out under reduced pressure. The reduced pressure can be used to improve evaporation, for example, to increase the evaporation rate, and further to control or direct the dried particles into a dedicated collector at the outlet end of the drying chamber. The collector can be in the form of a filter or a sieve. The reduced pressure can have additional advantages such as recirculation of the atomizing gas.

[0072] In embodiments of the present disclosure, the process includes a drying step of the formed droplets (wet particles), and the water activity (a w ) of the dried particles is less than about 1.0, for example, in the range from about 0.01 to about 0.6, for example, from about 0.05 to about 0.5, for example, from about 0.1 to about 0.5, for example, about 0.2, for example, about 0.3, or for example, about 0.4. The water activity (a w ) is the ratio of the vapor pressure of water in the substance to the vapor pressure of water in the standard state. The standard state is defined as the vapor pressure of pure water at the same temperature. In one embodiment of the present disclosure, the water activity is selected based on the type of microorganism. For example, the level of water activity is set to a value at which the microorganism of that type is advantageously dried, for example, the viability is not significantly affected.

[0073] In embodiments of the present disclosure, the process includes a drying step of the formed droplets (wet particles), and the solvent (water) content of the dried particles is less than about 20% by weight, for example, less than about 15% by weight, for example, less than about 10% by weight, for example, less than about 5% by weight, for example, less than about 3% by weight, for example, less than about 1% by weight, for example, less than about 0.1% by weight, based on the total weight of the dried particles.

[0074] In embodiments of the present disclosure, the solvent (e.g., water) content of the dried particles is less than about 10% by weight (preferably less than about 5% by weight or less than about 1% by weight) based on the total weight of the dried particles. Thus, a small amount of water can be present after the drying of the formed droplets, which can even be advantageous in that the water can be beneficial for improving the viability of the microorganisms.

[0075] In embodiments of the present disclosure, the dried particles are collected at the outlet end of the drying chamber. The outlet end may further comprise a collector for collecting the dried particles, such as a filter, a container, and / or a sieve.

[0076] In embodiments of the present disclosure, the dried particles are collected at the outlet end of the drying chamber using a filter (such as an electrostatic filter) or a sieve. The steric interaction preferably hinders the passage of the particles, while a gas such as the atomized gas can pass through the collector.

[0077] In embodiments of the present disclosure, the dried particles are collected at the outlet end of the drying chamber using a sieve having an aperture diameter of less than about 500 micrometers, such as in the range from about 40 micrometers to about 300 micrometers, such as in the range from about 50 micrometers to about 250 micrometers, such as about 50 micrometers, such as about 100 micrometers, such as about 150 micrometers, such as about 200 micrometers, or such as about 250 micrometers. Preferably, the aperture diameter is less than the particle size given by Dv50, more preferably less than Dv30, even more preferably less than Dv10, and still even more preferably less than Dv5, such as less than Dv3, such as less than Dv1. The apertures can be provided in a suitable shape such as round holes and / or slits (grids). Generally, the minimum dimension of the holes is a suitable size for hindering the particles. Thereby, for the slits, the aperture diameter refers to the slit width, and the same reasoning applies to holes of other shapes.

[0078] In embodiments of the present disclosure, the dried particles are collected at the outlet end of the drying chamber using a sieve having an aperture diameter in the range from about 40 micrometers to about 300 micrometers.

[0079] Deposit and specialized solutions The Applicant requests that, until the date of patent grant, samples of the deposited microorganisms described below be made available only to experts in accordance with the available regulations administered by the industrial property offices of the Contracting States to the Budapest Treaty.

[0080] The deposit was made with Leibniz Institute DSMZ - German Collection of Microorganisms and Cell Cultures Inhoffenstr.7B, 38124 Braunschweig, Germany, a depository institution having obtained the status of an international depository authority under the Budapest Treaty on the International Recognition of the Deposit of Microorganisms for the Purposes of Patent Procedure. The accession number and details of the deposit are shown in Table 1.

Table 1

Examples

[0081] Example 1 Materials and Methods Maltodextrin (dextrose equivalent value 12), trade name Glucidex IT12, was obtained from Roquette (Lestrem, France). Sodium chloride (NaCl) (Kenilworth, New Jersey, USA, Merck) and tryptone (Hampshire, UK, Oxoid) isotonic solution was prepared for bacterial dilutions. Measurement of bacterial viability was performed using the double - staining LIVE / DEAD™ BacLight™ Bacterial Viability Kit (Eugene, Oregon, USA, Molecular Probes, L - 7012). Bifidobacterium animalis DSM15954, a probiotic cell at a concentration of 3×10 11 cells / gram (hereinafter referred to as bifido), was provided by Chr. Hansen A / S (Helsingholm, Denmark) and stored at - 80 °C until further use.

[0082] Electrohydrodynamic drying Probiotics (3×10 11 cells / gram) stored at -80 °C were thawed at room temperature and transferred (5 g) to a 55 mm × 80 mm aluminum sample holder. The sample was placed inside a polycarbonate sealed chamber (Figure 1), where the temperature and relative humidity (RH = 20 ± 3%) were controlled by introducing air or nitrogen gas through a diffuser.

[0083] The EHD drying mechanism consists of a unipolar DC high voltage generator (Gamma High Voltage Research, Florida, USA) connected to a computer that executes an algorithm to control the DC current applied to three metal needles (0.1 mm in diameter and 20 mm in length). The three metal needles were spaced 20 mm apart and arranged vertically above the sample holder where the bifid sample was placed.

[0084] The sample was subjected to an external electric field with an intensity ranging from 1.5 kV / mm to 6.6 kV / mm, while the current was automatically adjusted (AA) from the power supply.

[0085] The effects of the applied polarity, voltage, gas, distance between the needle tip and the sample holder, addition of maltodextrin as an excipient, and temperature inside the chamber were investigated in the EHD drying of bifid. The tested conditions are summarized in Table 2.

Table 2

[0086] Mechanism 2 was developed to optimize Mechanism 1. A mobile compartment (M) was used, where a nylon filter membrane (MF) with a pore size of 0.45 μm and a diameter of 90 mm (Sigma-Aldrich, Steinheim, Germany) was placed on a sample holder of a metal mesh. The current intensity was always constant at 200 μ using a computer algorithm, while the voltage was automatically adjusted (AA). The influence of the number of needles and the distance between the needle tip and the sample holder was tested (samples MF2 - MF5 described in Table 2).

[0087] For both Mechanisms 1 and 2, bifid samples were either subjected to an external electric field for 1 hour and 2 hours (EHD drying - S) or not (reference sample - R).

[0088] Lyophilization Probiotics stored at -80 °C (3×10 11 cells / gram) were thawed at room temperature and transferred (5 g) to a sample holder of 55 mm × 80 mm aluminum (same protocol as EHD drying). Bifid samples were frozen at -20 °C for 24 hours and then lyophilized for 2 hours (at 1,030 millibars (about 103 kPa)) using a bench-scale lyophilizer (Chist, Beta LMC1 - 8, Germany), sample FDM (Table 2). The experiment was performed twice.

[0089] Characterization of samples Water loss (%) The initial weight of the sample and the subsequent weight of the sample after 1 hour and 2 hours of EHD drying were measured with a digital scale (Mettler Toledo, Greifensee, Switzerland) with an accuracy of 0.001 g. The % water loss (WL) of the sample drying in the experiment was calculated based on the following formula.

Equation

[0090] Cell viability (flow cytometry) Dry probiotics were stained with the double-staining kit BacLight™ composed of the fluorescent dyes SYTO9 (excitation / emission 485 / 498 nm) and propidium iodide (PI) (excitation / emission: 535 / 617 nm). The BD Accuri™ C6 Plus system (BD Biosciences, Franklin Lakes, NJ, USA) was utilized for flow cytometry analysis regarding the viability of bacteria. All analyses were performed at a low flow rate (14 μL / min), and the maximum volume of the collected sample for fixation was performed at 10 μL. The concentration of bacteria in the sample was adjusted between approximately 1×10 3 cells / mL and approximately 5×10 6 cells / mL, and appropriate gates were defined for SYTO9- and PI-stained bacteria classified by the dot plot of FL1 vs. FL3 (BD-Biosciences, 2012). The experiment was performed three times.

[0091] FTIR Infrared transmittance measurements were performed at room temperature (25 °C) with an FT-IR spectrometer (Thermo Fisher Scientific, Nicolet iS50, Waltham, MA, USA). Spectra were recorded after 32 scans at a resolution of 4 cm -1 from 4,000 cm -1 to 400 cm -1 in the wavenumber range.

[0092] Surface charge of bacterial cells Approximately 10 8Bifidobacterium dried cells at a concentration of cells / mL were suspended in 10 mM KH2PO4 (Steinheim, Germany, Sigma-Aldrich) and transferred to a capillary cell (DTS1070 cell, Malvern Panalytical Ltd, Malvern, UK) with a gold-plated beryllium / copper electrode. The electrophoretic mobility of the sample was measured using a Zeta-sizer (Malvern Panalytical Ltd, Malvern, UK) and converted to zeta potential (mV) using the Helmholtz-Schmoluchowski equation. Each sample was analyzed 5 times, and all measurements were performed at 25 °C.

[0093] Bacterial cell surface hydrophobicity The hydrophobic / hydrophilic properties of the surface of Bifidobacterium cells after EHD drying and freeze-drying were evaluated by analysis of microbial adhesion to hexadecane (MATH) (Deepika et al., 2009). Bifidobacterium probiotic cells were suspended in 10 mM KH2PO4 until an absorbance (OD600) of approximately 0.8. An equal volume of 2 mL of bacterial cell suspension and hexadecane (Thermo Fisher Scientific, Waltham, Massachusetts, USA) were vortexed for 1 minute and left to stand for 20 minutes until complete phase separation. The OD400 of the aqueous phase was carefully removed and measured.

[0094] The percentage of microbial adhesion to hexadecane (or % hydrophobicity) was calculated using the following equation.

Equation

[0095] Statistical analysis Results are presented as mean ± standard deviation (SD). To analyze the differences between them, single factor ANOVA was utilized. Statistically significant different sample results were considered for p-values less than 5% (p < 0.05).

[0096] Results EHD mechanism 1 Influence of polarity and gas type The influence of polarity on EHD drying has been investigated and is shown in Fig. 2. Since an increase in WL water loss (WL) was observed over time for all samples after 2 hours, Fig. 2A shows that WL is an EHD drying parameter that is time-dependent.

[0097] Sample S1 (negative polarity) lost approximately 38.7 ± 3.2% and 62.8 ± 5.3% of water after 1 hour and 2 hours, respectively (statistically significant, P < 0.05), while S2 (positive polarity) lost approximately 9.1 ± 1.3% and 18.2 ± 2.4% of water after 1 hour and 2 hours, respectively (not statistically significant).

[0098] Sample S3 (WL = 38.8 ± 3.2% (1 hour), WL = 64.3 ± 0.3% (2 hours)) was also dried under negative polarity and lost a greater amount of water compared to sample S4 (WL = 7.7 ± 0.4% (1 hour), WL = 17.5 ± 1.6% (2 hours)) dried under positive polarity (P < 0.05). Nevertheless, no significant difference was shown in terms of bacterial viability (V) when compared to samples dried using either negative (samples S1 and S3) or positive (samples S2 and S4) polarity.

[0099] Negative corona results in a higher corona current than positive corona for the same voltage (Lai and Wong, 2003), which has been demonstrated to justify the greater water loss (Figure 2) observed for samples dried with negative polarity. Furthermore, the ionic wind generated by negative corona exhibits a higher velocity than that associated with positive corona at low voltages (less than 10 kV / mm) (S. Chen, Van Den Berg, and Nijdam, 2018), which may also affect the water evaporating from the EHD-dried samples. The lower dielectric breakdown strength of air (1.15 times lower than nitrogen) enabled an increase in voltage up to 10 kV for samples S5 and S6. Sample S5 dried under the influence of negative corona discharge also showed significantly higher water loss (WL = 67 ± 5.4% (1 hour), WL = 81.3 ± 1.6% (2 hours)) and similar probiotic viability (V = 66.3 ± 39.6 (1 hour), V = 25.7 ± 18.3% (2 hours)) compared to sample S6 dried under the influence of positive corona discharge (WL = 9.3 ± 1.9% (1 hour), WL = 18.7 ± 2.2% (2 hours), and V = 83.3 ± 16.6% (1 hour), V = 26.1 ± 16.3 (2 hours)). Here, a higher contribution of negative polarity compared to positive polarity was confirmed for EHD drying when either nitrogen or air was the ambient gas. Samples dried using positive polarity showed similar water loss compared to the reference sample (R0) (almost no drying), highlighting the higher efficiency of EHD drying using negative polarity.

[0100] Influence of Voltage Figure 3A demonstrates that an increase in voltage from 3.7 kV to 4.7 kV increased the water loss in samples S7 and S3 from 11.1 ± 1.6% to 38.9 ± 3.2% and 11.1 ± 1.6% to 38.9 ± 3.2% respectively after 1 hour of drying. After 2 hours of drying, samples S7 and S3 lost approximately 19.5 ± 2.1% and 64.32 ± 0.3% of water respectively. A similar trend was shown for samples S8 (3.7 kV) and S5 (10 kV), where for sample S8, the sample lost approximately 10.7 ± 0.3% (1 hour) and 21.9 ± 0.6% (2 hours), while for sample S5, the water loss was approximately 67 ± 5.4% (1 hour) and 81.3 ± 1.6% (2 hours). The increase in applied voltage was demonstrated to enhance the electric field by increasing the charge density, current, and velocity of the ionic wind, and thus improve the drying rate (Anukiruthika, Moses, and Anandharamakrishnan, 2021; Defraeye and Martynenko, 2019; S. Chen, Nobelen, and Nijdam, 2017).

[0101] When nitrogen was the ambient gas, no significant difference in bacterial viability was shown between samples S7 (3.7 kV), S3 (4.7 kV), and the reference sample R0 (Figure 3B). For sample S7, the probiotic viability decreased from 99.9 ± 4.1% (1 hour) to 95.9 ± 3.2% (2 hours), while in sample S3, the % of viable cells decreased from 84.4 ± 12.1% (1 hour) to 53 ± 30.8% (2 hours). The reference sample showed a viability of approximately 99% for both test times.

[0102] When comparing samples S8 (3.7 kV) and S5 (10 kV) with the reference sample (R0), a significant difference was found in the percentage of viable probiotics. For sample S8, viability decreased from 96.3 ± 3.0% (1 hour) to 86.6 ± 15.3% (2 hours), while for sample S5, viability decreased from 71.5 ± 26.8% (1 hour) to 16.2 ± 13.4% (2 hours). Sample S5 had the highest evaporation rate among the samples tested in Figure 3, contrary to slightly dried samples S7 and S8, and thus had a lower viability.

[0103] Similar to other drying techniques, during EHD drying, when water is removed from the cells and their surrounding environment, bacteria are subjected to dehydration and osmotic stress. Therefore, it can be expected that an increase in voltage will improve the evaporation of water and decrease the viability of bacteria.

[0104] Effect of the distance between the needle and the sample holder The effect of the distance between the high-voltage electrodes for each pair of samples is shown in Figure 4. Sample S7 with a distance of 25 mm did not show a significant difference from the reference R0 in terms of water loss (WL = 11.1 ± 1.6% (1 hour), WL = 19.5 ± 2.1% (2 hours)), but it was found that sample S1 with a distance of 15 mm had more than three times the water loss (WL = 38.7 ± 3.2% (1 hour), WL = 62.8 ± 5.3% (2 hours)) than S7 and R0 after 2 hours of drying. However, the viability of bacteria was not statistically significant for both samples S7 (V = 99.9 ± 4.1% (1 hour) and V = 95.9 ± 3.2% (2 hours)) and S1 (V = 78.3 ± 27.6% (1 hour), V = 55.8 ± 33.6% (2 hours)), and no statistical difference was shown from the reference sample (R0).

[0105] Samples S5 (15 mm) and S9 (25 mm), dried under air conditions, showed a statistically significant difference in their water loss at the first hour (WL = 67.0 ± 5.4% (S5), WL = 57.3 ± 3.2% (S9)), but did not show a statistically significant difference at the second hour of EHD drying (WL = 81.0 ± 1.6% (S5), WL = 79.6 ± 0.9% (S9)). Most of the water evaporation occurred at the first hour, while during the second hour of EHD drying, the dehydration of bacteria in S5 (V = 66.4 ± 39.6% (1 hour), V = 25.7 ± 18.3% (2 hours)) and S9 (V = 66.3 ± 39.5% (1 hour), V = 26.1 ± 18.1% (2 hours)) reached a critical point where the viability decreased significantly compared to the reference sample R0.

[0106] Similar to the increase in voltage, the decrease in electrode distance is another parameter that affects the corona discharge, forming a large ion current and increasing the ion wind speed, thus accelerating the drying rate (Tirtha R. Bajgai et al., 2006). This result showed that, conversely, the decrease in electrode distance affected the water removal from the sample.

[0107] Effect of temperature The effect of nitrogen temperature on the EHD drying of samples S11, S14, and S16 is shown in (Figure 5). The increase in nitrogen temperature from 20 °C to 30 °C and 40 °C increased the water loss of S11 (WL = 35.4 ± 2.7% (1 hour)), S14 (57.2 ± 10.2% (1 hour)), and S16 (76.3 ± 3.6% (1 hour)), respectively, while their corresponding viabilities decreased (V = 85.9 ± 18.5% (1 hour), V = 64.9 ± 28.5% (1 hour), and V = 19.3 ± 19% (1 hour)).

[0108] The combination of EHD drying and heated nitrogen at 30 °C for sample S14 showed no statistically significant difference in probiotic viability compared to S11 dried at ambient temperature (20 °C) at the same electric field strength. However, the water evaporation of S14 reached the same loss as that of S11 after 2 hours (WL = 66.9 ± 4.8% (1 hour)) from the first hour of drying. Therefore, it can be concluded that the combination of EHD drying with a high gas temperature can increase water loss due to convective and heat diffusion effects from the core to the surface of the sample without compromising bacterial viability at least during the first hour of drying. Sample S16 showed an acceleration of water loss from the first hour of EHD drying at 40 °C. However, it can be observed that there was no significant difference between the water loss after 1 hour of drying (WL = 76.3 ± 3.6% (1 hour)) and after 2 hours of drying (WL = 81.4 ± 0.1% (2 hours)), and the drying rate decreased. However, it is clearly evident that such rapid water loss significantly decreased probiotic viability (V = 19.3 ± 19% (1 hour), V = 4.9 ± 3.7% (2 hours)).

[0109] EHD mechanism 2 Effect of the number of needles and the distance between the needle and the sample holder To improve probiotic drying, a moving section was added for the periodic movement of the flat electrode (mechanism 2). Furthermore, a mesh electrode and a nylon filter were utilized (Mujumdar and Xiao, 2019; Iranshahi, Martynenko, and Defraeye, 2020) to facilitate the passage of the air stream (under the probiotics).

[0110] The influence of the number of needles and the distance between the needles and the sample was also investigated, and the results are shown in (Figure 6). Among the water losses of samples MF4 (WL = 37 ± 5.3% (1 hour), WL = 68 ± 2.3% (2 hours)), MF5 (WL = 42.9 ± 5.6% (1 hour), WL = 74 ± 0.1% (2 hours)), MF2 ((WL = 39.3 ± 2.5% (1 hour), WL = 72.5 ± 3.5% (2 hours)), and MF3 (WL = 41.5 ± 3.8% (1 hour), WL = 73.5 ± 3.4 (2 hours)), and also for the viability of the corresponding probiotics (for MF4, V = 89.2 ± 4.8% (1 hour), V = 60.8 ± 20.8% (2 hours), for MF5, V = 84.9 ± 10.8% (1 hour), V = 48.1 ± 28.5% (2 hours), for MF2, V = 88.4 ± 4.9% (1 hour), V = 33.3 ± 23.9% (2 hours), for MF3, V = 90.5 ± 4.9% (1 hour), V = 43.8 ± 23.9% (2 hours)), it was observed that neither the number of needles nor the electrode gap resulted in a significant difference. Nevertheless, a significant difference was shown between all test samples and the reference sample R0. This experimental setup did not show a significant difference by increasing the number of needles and the electrode gap, and thus, sample MF4 was selected for further study.

[0111] Comparison of EHD with freeze dryer Water loss and viability The influence of the adopted 2-hour drying process on water loss and the viability of probiotic bacteria is shown in (Figure 7). Overall, the EHD-dried sample MF4 with the optimized "mechanism 2" showed the highest water loss (WL = 76.4 ± 2.2% (2 hours)) together with the freeze-dead sample FD (WL = 76.8 ± 5.5% (2 hours)), while no significant difference was shown between them in terms of the viability of bacteria (for MF4, V = 60.8 ± 20.8% (2 hours), for FD, V = 45.5 ± 11.3% (2 hours)).

[0112] Both MF4 and FD showed the highest water losses among the dry samples, while subsequently, sample S1 had less water loss of 62.8 ± 5.3% (2 h). The EHD-dried samples S1 and MF4 showed statistically different water loss results, indicating that the filter mesh electrodes did promote the air flow and improve water evaporation (Mujumdar and Xiao, 2019; Iranshahi, Martynenko, and Defraeye, 2020).

[0113] The addition of maltodextrin decreased the water loss of SM1 (WL = 23.7 ± 3.3% (2 h)) and FDM (WL = 48.5 ± 3.5% (2 h)) compared to the corresponding MF4 or FD samples without maltodextrin. Using single-drop convective drying experiments with various sugars, Adhikari (Adhikari et al., 2004) suggested that the addition of maltodextrin decreased its drying rate because water diffusion through maltodextrin molecules was difficult. Similarly, Gianfrancesco et al. also reported a decrease in the water flux towards the maltodextrin amorphous matrix that limited the evaporation rate and water loss during the selected drying time (Gianfrancesco et al., 2012).

[0114] Nevertheless, SM1 (V = 72 ± 10.6% (2 h)) showed an improvement in viability compared to both lyophilized samples. It is known that protective sugars such as maltodextrin decrease the membrane transition temperature (Tm) and maintain the membrane integrity of bacterial cells while being dehydrated (Mille et al., 2004; Strasser et al., 2009). Nevertheless, no statistical difference was shown between SM1 and S1 (V = 55.8 ± 33.6% (2 h)). Similarly, the viability of probiotics for sample FD without maltodextrin (V = 45.5 ± 11.3% (2 h)) and sample FDM with maltodextrin (V = 48.7 ± 10.2% (2 h)) showed no significant difference.

[0115] Fourier transform infrared spectrometer The Fourier transform infrared spectrometer (FTIR) spectrum of bifidum (Figure 8) shows the main characteristic bands of probiotic cells. For non-dried bifidum, the band at about 3280 cm representing the stretching of the hydroxy group bond (-OH) appeared more strongly because its sample was not as dried as the sample treated by electrohydrodynamics or lyophilization. -1 appeared more strongly.

[0116] 3000 cm -1 and 2800 cm -1 The fatty acid region between and was not present in the spectrum of non-dried bacteria, yet it appeared for all dried samples. More specifically, the bands at 2966 cm, 2936 cm, and 2876 cm indicate that the asymmetric CH3 stretching, asymmetric CH2 stretching, and symmetric CH3 stretching of the non-polar part of the phospholipid bilayer appeared after the evaporation of water from the cells (Santos et al., 2015; Shakirova et al., 2010; Dianawati, Mishra, and Shaha, 2012). The aforementioned groups were present in fresh probiotics and thus appeared after the evaporation of the external fluid present in the fatty acid region. -1 2936 cm -1 and 2876 cm -1 respectively, indicating that the asymmetric CH3 stretching, asymmetric CH2 stretching, and symmetric CH3 stretching of the non-polar part of the phospholipid bilayer appeared after the evaporation of water from the cells (Santos et al., 2015; Shakirova et al., 2010; Dianawati, Mishra, and Shaha, 2012). The aforementioned groups were present in fresh probiotics and thus appeared after the evaporation of the external fluid present in the fatty acid region.

[0117] The transmittance in the bacterial protein region occurs at frequencies between 1660 cm and 1500 cm. More precisely, the carbonyl stretching of the secondary amide (amide I) was found at 1639 cm, and the broad peak found at 1545 cm indicated the N-H bond of amide II. Since no shift for amide II was observed, there is no clear evidence that the secondary structure of the protein was affected by the application of the EHD or lyophilization process. However, the peak at 1639 cm was stronger for fresh probiotics, while the peak at 1545 cm -1 ~1500 cm -1 The transmittance in the bacterial protein region occurs at frequencies between 1660 cm and 1500 cm. More precisely, the carbonyl stretching of the secondary amide (amide I) was found at 1639 cm, and the broad peak found at 1545 cm indicated the N-H bond of amide II. Since no shift for amide II was observed, there is no clear evidence that the secondary structure of the protein was affected by the application of the EHD or lyophilization process. However, the peak at 1639 cm was stronger for fresh probiotics, while the peak at 1545 cm -1 was found at 1545 cm -1 indicated the N-H bond of amide II. Since no shift for amide II was observed, there is no clear evidence that the secondary structure of the protein was affected by the application of the EHD or lyophilization process. However, the peak at 1639 cm was stronger for fresh probiotics, while the peak at 1545 cm -1 was stronger for fresh probiotics, while the peak at 1545 cm -1Note that the peaks in were more intense for the freeze-dried sample FD and MF4 of EDH drying. The EDH-dried sample S1 showed similar intensities for both peaks. Based on Bozkurt et al., an increase in the contact of bifid cells with water molecules increases the amide I absorption band, as observed in this result (Bozkurt et al., 2019). Interestingly, the intensity of the amide II absorption also correlated with the water loss of the sample, as it appeared more intensely in the drier FD and MF4, less intensely in S1, and was much reduced in fresh bifid. The P=O vibration of PO2- can be identified at 1250 cm -1 while broad carbohydrate and phosphate bands were observed for all samples at a frequency of 1100 cm -1 ~980 cm -1 The maximum transmittance was shown at 1070 cm -1 and 1040 cm -1 by the valence vibration of the C-O-C group with respect to the polar part of the phospholipid bilayer. The peaks at 1070 cm -1 and 1040 cm -1 decreased for fresh bacteria due to the expansion of contact with water molecules, as shown by Bozkurt et al. (Bozkurt et al., 2019).

[0118] The comparison between EHD and freeze-dried samples with and without maltodextrin is shown in (Figure 9).

[0119] Samples containing maltodextrin did not show a peak at 2966 cm -1 and the intensity of the carbonyl stretching of the secondary amide (amide I) at 1639 cm -1 was more intense than the N-H bending of amide II at 1548 cm -1 . The above may be due to the fact that samples containing maltodextrin are characterized by a higher water content, and an increase in the contact of bifid with water molecules can increase the amide I absorption band, similar to fresh bifid samples. 1394 cm -1 ~1310 cm -1a broader peak, as well as an additional small peak at 1210 cm -1 and the relatively intense peak at 1150 cm -1 are due to the chemical structure of maltodextrin. The latter peak represents C-O-C and C-O bonds dominated by the vibration of the carbohydrate ring. Furthermore, the interaction between PO2 of the bacterial envelope and maltodextrin is shown by a change in frequency from 1040 cm -1 for dry bifidobacteria to 1020 cm -1 for the sample with bifidobacteria and maltodextrin. The latter is confirmed by noting the shift of the maltodextrin band from 995 cm -1 to 1020 cm -1 . Although it is a hydrogen bond, such an interaction between sugar and the phospholipid moiety of cell lipids has been previously suggested and described in the literature (Oldenhof et al., 2005; Leslie et al., 1995; Dzuba, Leonov, and Surovtsev, 2020). FTIR technology has been widely used in investigations regarding the protective effects of sugars, such as maltodextrin. However, it should be mentioned that the protective effects regarding such high molecular weight molecules are indirect by promoting the replacement of water on the cell surface from smaller sugars such as glucose (Santos et al., 2015).

[0120] Zeta potential Bacteria had a negative charge in all samples, as previously suggested for other Bifidobacterium strains (Gomez Zavaglia et al., 2002). Such a negative cell surface charge at neutral pH may indicate the predominance of anionic domains such as strong acids (phosphate-based (lipo)teichoic acids) as well as weak acids (acidic polysaccharides and proteins) (Deepika et al., 2009).

[0121] According to Fig. 10A, no significant difference was shown between the zeta potential values of the non-dried sample and the dried sample. Sample R0, as well as the EHD or freeze-dried samples (S1 and FD) without maltodextrin, showed lower absolute values of zeta potential compared to the corresponding samples (R17, SM1, and FDM) containing maltodextrin. The latter result can confirm the indication from the FTIR spectrum that maltodextrin molecules interacted with the bacterial membrane surface. When the absolute zeta potential value increases in a colloidal system, more stable clusters are formed during electrophoresis. Therefore, it can be assumed that the interaction of sugar with the bacterial surface can stabilize the bacterial surface (Klodzinska et al., 2010).

[0122] Hydrophobicity The adhesion test of microorganisms to hydrocarbons is usually used to evaluate the cell surface hydrophobicity in Bifidobacterium (Pelletier et al., 1997; Gomez Zavaglia et al., 2002). When bacteria are hydrophobic, the bacteria show a high adhesion (%) to hexadecane. As shown in Fig. 10B, all samples showed similar values of hydrophobicity (Fig. 10B). Interestingly, the samples of the EHD-dried samples in Mechanism 1 showed statistically different results with lower hydrophobicity compared to the control of EHD2 and the freeze-dried samples.

[0123] Furthermore, the addition of maltodextrin in the same sample increased the hydrophobicity of bifidobacteria. According to Shakirova et al. (2010), cell surface hydrophobicity was associated with a higher content of bifidobacterial surface proteins (Shakirova et al., 2010). A number of other studies have suggested that the presence of (glycol)protein materials and polysaccharides is responsible for higher hydrophobicity and hydrophilicity, respectively (Collado, Meriluoto, and Salminen, 2008; Boonaert and Rouxhet, 2000; Van Der Mei et al., 2003). Thus, as indicated by both the negative charge nature and high hydrophobicity of bifidobacteria, and as seen in the FTIR spectrum, a protein-rich bacterial surface is suggested.

[0124] Conclusion Electrohydrodynamic drying can be used to dry probiotic cells. The EHD drying process was found to depend on several parameters such as the polarity and voltage of the ionic wind, the type of collector, the ambient temperature and gas, the probiotic concentration, and the use of excipients. The optimized conditions using mechanism 1 enabled reaching 70% drying and a maximum 34% cell viability. Probiotic viability can be increased up to 50% after 2 h by dispersing the probiotics using maltodextrin. However, the addition of maltodextrin decreased the water evaporation rate by 30%. The EHD performance (cell viability and water loss) was improved using a flat electrode of a moving mesh under the probiotics so as to facilitate the air flow of drying, which is called mechanism 2. Using this mechanism 2, water evaporation and probiotic cell viability increased up to 78% and 70%, respectively, compared to mechanism 1. Furthermore, the comparison of EHD drying to the lyophilization process revealed that the probiotic survival and water evaporation rate were similar while the cell surface characteristics remained the same as those of non-dried cells.

[0125] Example 2 Materials and Methods The probiotic cell Bifidobacterium animalis, DSM15954 (hereinafter referred to as Bifid) was provided by Chr-Hansen A / S (Horsholm, Denmark). Maltodextrin with a dextrose equivalent value of 12 (Roquette, Glucidex IT12, Restron, France) was used for the encapsulation of Bifid. Sodium chloride (NaCl) (Merck, Kenilworth, New Jersey, USA) and tryptone (Oxoid, Hampshire, UK) were used for the preparation of the isotonic solution. KH2PO4 (VWR International, Leuven, Belgium) was used as a suspension medium for the evaluation of the bacterial surface properties, and hexadecane (Thermo Fisher Scientific, Waltham, Massachusetts, USA) was used to perform the hydrophobicity measurement. The viability of the bacteria was determined using the double staining LIVE / DEAD™ BacLight™ Bacterial Viability Kit (Molecular Probes, Eugene, Oregon, USA, L-7012). The fluorescent dye thiazole orange (Sigma-Aldrich, USA) with a dye content of approximately 90% was used for the staining of Bifid cells.

[0126] Evaluation of the Physicochemical Properties of the Bacterial Cell Surface Surface Charge of Bacterial Cells Approximately 3*10 8Bifidobacterium probiotic cells at a concentration of cells / mL were suspended in 10 mM KH2PO4 to obtain an optical density (OD600) of approximately 1.0. The pH of the solution was adjusted to 1, 2, 4, 6, and 8 using 1 M HCl or 1 M NaOH. The electrophoretic mobility was determined using a Zeta-sizer (Malvern Panalytical Ltd, Malvern, UK). A 1 mL volume sample was injected into a folded capillary cell (Malvern Panalytical Ltd, Malvern, UK, DTS1070 cell) with gold-plated beryllium / copper electrodes. The measurements were carried out at 25 °C and each sample was analyzed 5 times. The electrophoretic mobility was converted to zeta potential using the Helmholtz-Schmoluchowski equation.

[0127] Hydrophobic / hydrophilic properties of the bacterial cell surface Analysis of microbial adhesion to hexadecane (MATH) was employed to evaluate the hydrophobic properties of the surface of Bifidobacterium cells at different pHs (1, 2, 4, 6, 8). Probiotic cells were suspended in 10 mM KH2PO4 to obtain an optical density (OD400) of approximately 0.8. Two milliliters of the bacterial cell suspension was mixed with an equal volume of hexadecane, vortexed for 1 minute, and left standing for 20 minutes to ensure complete phase separation. The aqueous phase was carefully removed after equilibration and the OD400 was measured.

[0128] The percentage of microbial adhesion to hexadecane was calculated using the following equation.

Equation

[0129] Here, A0 is the initial absorbance regarding the bacterial suspension, and A1 is the absorbance (OD400) after 20 minutes of culture and phase separation.

[0130] Each measurement was performed 3 times.

[0131] Electrospray of Probiotic Solution Maltodextrin (75% w / v) was dispersed in Millipore water and stirred until a homogeneous solution was obtained. Then, Bifidobacterium probiotic cells (10% w / v) were added and stirred until the Bifidobacterium probiotic cells were completely dispersed.

[0132] The electrospray mechanism included a high-voltage generator (Gamma High Voltage Research, Inc., ES50P-10W, United States of America) for providing a voltage of 15 kV to 40 kV and a syringe pump (New Era Pump Systems, Inc., Farmingdale, New York State, United States of America) for supplying the maltodextrin / Bifidobacterium dispersion at a flow rate of 0.03 mL / min. The maltodextrin Bifidobacterium capsules were horizontally collected on a steel plate covered with aluminum foil, which was placed at a distance of 10 cm from the tip of the needle (27 gauge, 12 mm in length, 0.21 mm in inner diameter). The electrospray mechanism was placed inside a chamber with a nitrogen flow, and the temperature and relative humidity were 23 °C and 22 ± 3%, respectively.

[0133] The polarity of the electrode connected to the tip of the syringe needle is positive or negative. Then, for the sake of simplicity, the samples are each said to be electrosprayed under positive or negative polarity. Furthermore, the charge applied to the tip of the needle is first represented, and the charge applied to the collector is represented. For example, when -15 kV is applied to the nozzle tip and +5 kV is applied to the collector, it is then referred to as (-15 kV)(+5 kV), and the absolute value of the potential is shown as |20 kV|.

[0134] Confocal Laser Scanning Microscope An Inverted Zeiss LSM-710 confocal laser scanning microscope (Jena, Germany, Carl Zeiss MicroImaging GmbH) equipped with a diode laser (405 nm), argon lasers (458 nm, 488 nm, and 514 nm), two HeNe lasers (543 nm and 633 nm), and three detectors and one transmission detector was used. Probiotic cells were stained with the fluorescent dye thiazole orange (Sigma-Aldrich, United States of America) with a dye content of approximately 90% and washed with 0.85% w / v NaCl before encapsulation. The dye was in powder form and was therefore dissolved in dimethyl sulfoxide (DMSO) at a concentration of 42 μmol / L. For image acquisition, the samples were scanned by using the Z-series mode, and confocal fluorescence photographs were taken using 20x, 40x, and 60x objective lenses. All images were acquired with an excitation wavelength of 488 nm and an emission bandpass filter of 505 nm - 550 nm. The images were analyzed using ZEN software.

[0135] Fourier transform infrared spectrophotometer The Fourier transform infrared spectrophotometer (FTIR) spectra of the samples were recorded in the range of 400 cm -1 ~4000 cm -1 using a Nicolet iS50 spectrometer (Thermo Scientific, New York, United States of America) in the attenuated total reflection (ATR) mode. All spectra were recorded in transmission mode at room temperature (25 °C) with a scan resolution of 4 cm -1 and 32 scans. Spectral analysis was performed using Omnic software (Thermo).

[0136] (Using no maltodextrin as the encapsulating compound) Bifidobacterium pure cultures were electrosprayed using negative (-15 kV) (+5 kV) polarities and positive (+15 kV) (-5 kV) polarities, and 200 μL of the electrosprayed samples were evaluated by ATR-FTIR. The same volume of Bifidobacterium not treated with an external electric field was also evaluated as a control.

[0137] Differential Scanning Calorimetry Differential Scanning Calorimetry (DSC) experiments were conducted to determine the glass transition temperature (Tg) of the electrosprayed microcapsules. The thermograms of the samples were obtained using a DSC250 (TA Instruments, Newcastle, Delaware, USA) equipped with a Refrigerated Cooling System 90. The instrument was calibrated in terms of heat flow and temperature using distilled water (melting point (m.p.) = 0 °C; ΔHm = 334 J / g) and indium (m.p. = 156.5 °C; ΔHm = 28.5 J / g). Nitrogen was used as the carrier gas at a flow rate of 50 mL / min.

[0138] Approximately 5 mg of the electrosprayed microcapsules (collected immediately after treatment) were placed in pre-weighed standard DSC aluminum pans (TA Instruments, Tzero aluminum Hermetic pans, USA) and hermetically sealed. An empty hermetically sealed aluminum pan was used as a reference. The pans were first equilibrated at 10 °C for 5 minutes and then followed by a heating ramp of 3 °C / min up to 120 °C. The run settings were set at least 30 °C lower and higher than the predicted Tg of the electrosprayed microcapsules based on the Tg of maltodextrin DE12.

[0139] The obtained DSC thermograms were analyzed using Trios software interfaced with the DSC, and the Tg was determined from the starting point, midpoint, and end point of the shift in the curve. All measurements were performed three times, and the Tg values were averaged.

[0140] Stability of Microencapsulated Bifidobacterium during Storage Microcapsules containing bifidobacteria electrosprayed with polarities of negative (-15 kV)(+5 kV) and positive (+15 kV)(-5 kV), and non-encapsulated Bifidobacterium animalis as a reference sample were stored in a culture cabinet (Ninolab A / S, Solrod Strand, Denmark, Model KB8400F) at 25 °C with a relative humidity of 35 ± 2% RH. The viability of encapsulated and non-encapsulated bifidobacteria cells as a function of time (0 days to 14 days) was determined by colony-forming unit (CFU) analysis. Specifically, the electrosprayed microcapsules were dispersed in a 0.85% w / v sodium chloride (NaCl) and 0.1% w / v peptone (tryptone) solution and homogenized by vortexing. Then, 0.1 mL of the appropriate 10-fold dilution was spread on a de Man Rogosa Sharpe (MRS) agar plate supplemented with 0.05 w / V% L-cysteine hydrochloride monohydrate (Merck, Kenilworth, NJ, USA) and plated. The MRS agar plates were cultured at 37 °C for 3 days under anaerobic conditions (Oxoid, AnaeroGen, Hampshire, UK), and cell viability was calculated as the average of 4 plates.

[0141] The encapsulation efficiency (EE) of the microcapsules was determined by dividing the number of viable cells (N) inside the capsules by the number of viable cells (N0) in the initial solution, as expressed using the following formula.

Equation

[0142] Statistical analysis The results are presented as mean ± standard deviation (SD). A one-way ANOVA was used to analyze the differences between the samples compared at each time. Statistically significant differences between the results of the samples were considered for p-values less than 5% (p < 0.05).

[0143] Results and discussion Surface charge of bacterial cells and hydrophobic / hydrophilic properties of the cell surface The zeta potential of bifidobacteria in potassium dihydrogen phosphate buffer as a function of pH is shown in Fig. 11. The cell surface electronegativity showed negative zeta potential values above pH 2 and a slightly positive value at pH 1, and decreased with decreasing pH. A similar zeta potential profile has been previously reported for other bifidobacterium strains (Bifidobacterium bifidum and Bifidobacterium pseudolongum) (Gomez Zavaglia et al., 2002), due to the fact that the cell surface composition is dominated by anionic domains containing strong acids (phosphate-based (lipo)teichoic acids) as well as weak acids (acidic polysaccharides and surface proteins) (Deepika et al., 2009). The progressive protonation of the aforementioned chemical groups can justify the decrease in bacterial electronegativity with decreasing pH (Pelletier et al., 1997).

[0144] The hydrophobicity of the bifid cell surface was evaluated by studying the adhesion of bifidobacteria to hexadecane (Fig. 12). At pH values of 1 - 6, bifidobacteria showed a similar affinity (80 ± 6.9%) for hexadecane, suggesting the nature of a hydrophobic surface. At higher pH values (pH = 8), bifidobacteria showed a lower hydrophobicity property, with a percentage of affinity for non-polar hexadecane of less than 40%. Covalently bound proteins, S-layer proteins, and fatty acids present on the surface of bifid cells (Dianawati and Shah, 2011; Shakirova et al., 2010, 2013) have been suggested to promote adhesion to hydrophobic solvents and contribute to high bacterial hydrophobicity.

[0145] Effect of electric field polarity and voltage on the “self - organization” of cells within microcapsules The effect of the negative and positive polarities of an electric field on the distribution of probiotic cells - pH 6 bifid capsules within electrosprayed maltodextrin was studied by confocal laser scanning microscopy (CLSM) (Fig. 13).

[0146] The negative polarity in the needle mostly brought about the "self-organization" of probiotic cells in the core of the electrosprayed capsules. This was due to the negatively charged bifid cells (at pH 6) being electrostatically repelled by the electric field of the negative charges on the needle wall and the inner surface of the Taylor cone. Similar "self-organization" of bifid cells in the core of the electrosprayed capsules was observed by using different voltages with the negative polarity in the needle (Figure 13). On the other hand, when the positive polarity was applied to the needle, the cells were dispersed towards the outer surface of the electrosprayed capsules (Figure 13).

[0147] Figure 14 shows the CLSM of the electrosprayed maltodextrin / bifid dispersions at different pH values. Similar to the results observed at pH 6, the negatively charged bifid cells at pH 4 - 8 "self-organized" in the core of the capsules when the electric field of the negative polarity was applied by the needle, while the cells at pH 1 (slightly positively charged) were dispersed closer to the outer surface of the capsules. Therefore, the encapsulation of bifid in the core and outer surface of the capsules is due to the movement of charged cells by the Coulomb and electrophoretic forces within the DC electric field. Furthermore, since the probiotics showed similar hydrophobic properties at pH 1 - pH 6 (Figure 12), it should be noted that the "self-organization" of the cells within the electrosprayed capsules was generated by the surface charge of the cells, rather than by the hydrophobic / hydrophilic properties of the cell surface.

[0148] Glass transition temperature of electrosprayed maltodextrin / probiotic capsules The influence of the voltage and polarity of the electric field on the glass transition temperature (Tg) of the electrosprayed capsules of maltodextrin - bifid was also evaluated. Tg is related to the moisture content of the electrosprayed capsules (Tg increases with the decrease in moisture content), as well as the subsequent stability and viability of the encapsulated cells (Drake et al., 2018; Haque and Roos, 2004; Terpou et al., 2019). To achieve stability during the storage of cells in the dry state, sufficient water removal and the formation of an amorphous glassy solid matrix are required (Drake et al., 2018).

[0149] The glass transition temperature (Tg) of electrosprayed maltodextrin capsules without probiotic cells changed slightly with the increase of the electrospray voltage for both positive and negative polarities. Slightly higher Tg values of maltodextrin capsules were observed using a positive-polarity electric field. On the other hand, electrosprayed maltodextrin-bifidobacterium capsules at pH 6 (Figure 15) showed a maximum Tg value at |20 kV| ((-15 kV)(+5 kV)), and then the Tg decreased with the increase of the electrospray voltage. Furthermore, maltodextrin-bifidobacterium capsules treated using a negative polarity were characterized by much higher Tg values than the capsules produced using a positive polarity. In particular, maltodextrin-bifidobacterium capsules produced using a negative polarity of (-15 kV)(+5 kV) were characterized by a Tg of 68.21 ± 1.66 °C, which is a significantly higher value compared to other electrosprayed capsules treated using a negative polarity. When a positive polarity of (+15 kV)(-5 kV) was used, the Tg was approximately 10 °C lower than that of the capsules produced using a negative polarity of the same voltage (-15 kV)(+5 kV). As suggested above, at pH 6, probiotic cells (with negative surface charge) electrosprayed using a negative-polarity electric field repelled and "self-organized" in the core of the capsules, while the solvent accumulated near the droplet surface. Therefore, the improved evaporation of the solvent during electrospray started from the surface of the highly charged droplets when a negative-polarity electric field was applied compared to when a positive one was applied. This improved evaporation resulted in a lower moisture content and subsequently a higher Tg value for electrosprayed maltodextrin-bifidobacterium capsules with a negative polarity compared to the corresponding capsules treated using a positive polarity. Therefore, different electric field voltages and polarities affect the thermal properties and glass transition of electrosprayed capsules due to the presence of surface-charged probiotics.

[0150] Fourier transform infrared spectrophotometer (FT-IR spectra of (non-encapsulated) electrosprayed bifidobacteria under the influence of different electric field polarities and of bifidobacteria not treated with an external electric field are shown in (Figure 16). The band at about 3280 cm -1 represents the stretching of the hydroxy group bond (-OH) and is associated with the water-related bonds of the sample. The band appeared more intense for the non-treated bifidobacteria, while the lowest hydroxy group stretching intensity was shown for the bifidobacteria electrosprayed at a negative polarity (-20 kV) (Figure 17A). The increase in the -OH band intensity after FTIR measurement of the spin-coated bifidobacteria biofilm in a moisture environment has been reported previously in the literature (Bozkurt et al., 2019).

[0151] The difference in the intensity of the bands in the fatty acid region between 3000 cm -1 and 2800 cm -1 was evident between the differently treated samples and was more intense in the spectra of the electrosprayed samples (Figure 16). The bands at 2961 cm -1 , 2930 cm -1 , and 2876 cm -1 show, respectively, the asymmetric CH3 and CH2 stretching and the symmetric CH3 stretching of the apolar parts of the phospholipid bilayer (Dianawati et al., 2012; Santos et al., 2015; Shakirova et al., 2010). Higher transmittance intensities were observed for the bifidobacteria electrosprayed using a negative polarity compared to a positive polarity electric field. The presence of water has been suggested to interfere with the intensity of these groups (Dianawati et al., 2012), showing that less water was present in the bacteria electrosprayed at -20 kV, more water was present at +20 kV, and even more was present in the non-treated bacteria sample, where the interferogram of the non-treated bacteria sample completely lacked these peaks.

[0152] The main protein amide bands for biological samples are typically at about 3200 cm -1 (amide A), 1660 cm -1 ~1500 cm -1(Amide I and Amide II), and those found at frequencies of 1310 cm -1 ~1240 cm -1 (Amide III) (Gautier et al., 2013; Krimm and Bandekart, 1986). For non-electrosprayed bifidobacteria, the carbonyl stretching vibration of the peptide bond from the secondary amide (at about 1640 cm -1 ) was found to be more prominent compared to the peak of Amide II at about 1550 cm -1 . Nevertheless, for electrosprayed bifidobacteria, a less prominent peak for Amide I compared to Amide II was observed, and this peak also appeared with the lowest intensity for bifidobacteria electrosprayed using negative polarity (Figure 16). Bozkurt et al. studied the effect of relative humidity on the surface structure of spin-coated bifidobacteria biofilms and concluded that the increase in the absorption of Amide I indicates an increase in the contact of bifidobacteria with water molecules (Bozkurt et al., 2019). Therefore, the decrease in the peak intensity of Amide I in the sample electrosprayed with negative polarity suggests a drier sample.

[0153] Furthermore, differences in the intensity of the bands at the polar sites of the cell's phospholipid bilayer, represented by the frequencies of 1070 cm -1 and 1040 cm -1 due to the valence vibration of the C-O-C group, were also observed between samples (Figure 6). The decrease in the transmittance intensity for non-electrosprayed bifidobacteria indicates an expansion of the cell's contact with water molecules, while the highest intensity found in the sample electrosprayed with negative polarity suggests that there was the least amount of water in the bifidobacteria electrosprayed using negative polarity compared to the sample electrosprayed using positive polarity (Bozkurt et al., 2019).

[0154] Stability and Cell Viability of Microencapsulated Probiotics during Storage As shown in Fig. 17, probiotics encapsulated using a negative polarity showed significantly higher viability (about 7.3 times higher, with a logarithmic loss of 0.06) compared to probiotics encapsulated using a positive polarity (logarithmic loss of 0.44), even after 3 days of storage. Similar trends in viability (about 2.4 times and 1.5 times higher) were observed even after 7 days and 14 days for the stability study. Non-encapsulated bifido had viability 29.5 times, 9 times, and 5.5 times lower than the cells encapsulated using a negative polarity after 3 days, 7 days, and 14 days of storage, respectively. It should be noted that the encapsulation efficiency of the microcapsules was similar for both polarities at about 92% (92.9% ± 2.1 and 92.4% ± 1.2 for electrosprayed with positive and negative polarities, respectively).

[0155] Therefore, encapsulated maltodextrin-bifido electrosprayed using a negative polarity showed higher viability up to about 14 days of storage compared to cells electrosprayed using a positive polarity. This results from the "self-organization" of the cells in the core of the capsules electrosprayed using a negative polarity and the lower water content (and water activity) of the capsules, thereby ensuring a better stability and viability state for the encapsulated cells. Thus, the stability and viability of encapsulated probiotics can be effectively controlled by utilizing the polarity of the external electric field.

[0156] Conclusion In this study, the encapsulation of Bifidobacterium lactis, DSM15954, a Gram-positive bacterium, in maltodextrin capsules using electrospray needles with both negative and positive charges was investigated. The charge polarity of the needles in the DC electric field directed the surface-charged probiotic cells either near the core or the outer surface of the capsules, as recorded by confocal microscopy images. This is a new "self-assembly" process that results in the formation of different "compositions" regarding the core or the outer surface of the microcapsules during the electrospray process. This can further be utilized to develop electrosprayed capsules with tailored core and outer surface compositions based on the interaction of the applied electric field and the dispersed charged molecules. The charge polarity applied to the electrospray needle controls not only the "self-assembly" of the probiotic cells in the liquid jet but also mass transfer, solvent evaporation, and the physicochemical properties of the capsules (e.g., glass transition).

[0157] The probiotics encapsulated using the negative polarity showed a significantly (by about 10 °C) higher glass transition value than the capsules produced using the positive polarity and exhibited far superior (about 7.3 times higher) viability even after 3 days of storage at 30% RH and 25 °C compared to the probiotics encapsulated using the positive polarity.

[0158] When a low-conductivity or neutral maltodextrin matrix was employed, the charge density at the bacterial surface was sufficient for the "self-organization" of the bacteria. A high-charge density biopolymer (i.e., a polyelectrolyte) that changes the charge density of the solution can further enhance the functionality of cell self-organization and distribution. The polarity of the electric field can also be used to modify the molecular interactions between polyelectrolytes and between polyelectrolytes and charged probiotic cells. Thus, it can be advantageous to use a polyelectrolyte that has the same charge as the cells, a different charge compared to the cells, or a neutral charge. An increase in the conductivity of the solution through the addition of salts that increase the surface charge density can also be used to regulate cell encapsulation. In addition, selecting probiotic cells with a sufficiently strong surface charge (at a pH not close to the isoelectric point of the probiotic cells) further contributes to the selective localization of encapsulated cells.

[0159] Furthermore, this study essentially evaluates a method for improving the protection and viability of live probiotic cells encapsulated by electrospraying. The viability of encapsulated cells was substantially improved when the cells were placed in the core of the capsule compared to when the probiotics were dispersed near the outer surface of the capsule.

Claims

1. A process for electrostatic spray drying of live microorganisms, consisting of the following steps: a) A step of providing a suspension containing a microorganism having a surface charge and a compounding aid, b) A step of applying an electrostatic charge to the suspension, c) A step of forming droplets of the suspension, d) A step of forming dried particles by drying the droplets, and e) A step of collecting the dried particles, A process comprising the same polarity as the surface charge of the microorganism.

2. The process according to claim 1, wherein the electrostatic charge is applied to the suspension while simultaneously moving the suspension.

3. The process according to claim 1 or 2, wherein the collection of the dried particles is performed using a collector having a charge opposite to the electrostatic charge and the surface charge of the microorganism.

4. The process according to claim 1, wherein the microorganism having a surface charge has a negative surface charge and is selected from the group consisting of Bifidobacterium animalis, Lactobacillus acidophilus, Pseudomonas aeruginosa, and Escherichia coli.

5. The process according to claim 4, wherein the microorganism is Bifidobacterium animalis subspecies lactis (DSM15954).

6. The process according to claim 1, wherein the shape of the dried particles is spherical, fibrous, spiral, rod-shaped, or hybrid.

7. The process according to claim 1, wherein the compounding agent is a polysaccharide, protein, lipid, or synthetic polymer, or a mixture thereof.

8. The process according to claim 7, wherein the polysaccharide is in a state of positive, negative, or neutral charge.

9. The process according to claim 8, wherein the polysaccharide is selected from the group consisting of maltodextrin, starch, xanthan gum, guerane, alginic acid, pectin, glucan, and chitosan.

10. The process according to claim 7, wherein the protein is a milk protein, an animal-derived non-milk protein, a plant-derived protein, an algal protein, or a fermentation product protein.

11. Use of a positively or negatively charged high-voltage source for encapsulating a living microorganism having the same surface charge as the charge of the high-voltage source.

12. A particle comprising a polymer and at least one living microbial cell, wherein the at least one living microbial cell is located in the core of the particle, the composition of the polymer in the particle is uniform, and a shell is formed around the core where substantially no cells are present.

13. Particles containing live microorganisms, obtainable by the process described in claim 1.

14. Use of the particles according to claim 12 for manufacturing food products, feed products, nutritional supplements, or pharmaceutical products.

15. Use of the particles according to claim 13 for manufacturing food products, feed products, nutritional supplements, or pharmaceutical products.

16. A food product, a feed product, a nutritional supplement, or a pharmaceutical product comprising the particles described in claim 12.

17. A food product, feed product, nutritional supplement, or pharmaceutical product comprising the particles described in Claim 13.

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