Encapsulation method by electrohydrodynamics

Abstract

The present disclosure relates to a method for electrostatically spray-drying a living microorganism having a specific surface charge using an electrostatic charge provided by a high voltage power source, wherein the electrostatic charge is the same as the surface charge of the microorganism.

Description

Field of the Invention

[0001] The present disclosure relates to methods for electrostatic spray drying of microorganisms and microorganisms embedded in dry particles. Background of the Invention

[0003] Spray drying is a technique in which a suspension is atomized (or sprayed) and then rapidly dried using a gaseous hot drying medium. The scalability of this manufacturing process allows the formation of particles that are intended for use in a wide range of industries, including food, polymers, biotechnology, pharmaceuticals or medical. The choice of an atomizer to break up the feedstock (suspension) into droplets depends largely on the type of solution and the characteristics of the desired dry particles. Traditional atomizers include rotary atomizers that rely on the use of centrifugal force to form droplets, hydraulic nozzle atomizers that rely on pressure, and pneumatic nozzle atomizers that rely on kinetic energy. Spray drying has been applied to a large number of particle types, including bacteria and other microorganisms.

[0004] Electrospray is an electrohydrodynamic method for encapsulating compounds or bacterial cells in a polymer matrix. Electrostatic droplet formation has recently been disclosed as an improved method for achieving droplet formation. This method relies on electrostatically charging the suspension before the droplets form. Charging the suspension can improve the drying characteristics of the formed droplets and can promote the collection and aggregation of particles. Although electrostatic spray drying provides a large number of advantages over traditional spray drying, its use and application in microbial drying has not yet been successful. This is logical, because the vitality of known microorganisms can be affected by electrostatic charge, and in fact, high voltage is a commonly used liquid disinfection method. Therefore, the use of electrostatic spray drying mainly focuses on drying non-living substances.

[0005] Patent publication WO2021152111A1 discloses a method for electrostatic spray drying of live microorganisms. However, there is still a need to provide improved electrostatic spray drying methods, and in particular methods that result in cost savings and / or improved stability of the final product. SUMMARY OF THE INVENTION

[0007] It has been observed that the electric field and charge polarity during spray drying can be optimized by utilizing electrohydrodynamics to affect nanostructures to improve the encapsulation / immobilization of bioactive compounds such as microorganisms.

[0008] The present invention provides an improved method for electrostatic spray drying of live microorganisms by encapsulating microorganisms having negative surface charges using a negatively charged high voltage power source.

[0009] The microorganisms may inherently have a negative surface charge, or the microorganisms may be treated to acquire a negative surface charge.

[0010] According to a first aspect of the present invention, there is provided a method for electrostatically spray drying live microorganisms, the method 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) drying the droplets to form dry particles; and e) collecting the dry particles; wherein the electrostatic charge has the same polarity as the surface charge of the microorganisms.

[0011] In one embodiment, an electrostatic charge is applied to the suspension while the suspension is being moved.

[0012] In one embodiment, collection of dry particles is achieved using a collector having an opposite charge compared to the electrostatic charge and the surface charge of the microorganisms.

[0013] 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.

[0014] In one embodiment, the microorganism is Bifidobacterium animalis subsp. lactis (DSM 15954).

[0015] In one embodiment, the shape of the dry particles is spherical, fibrous, spiral, rod-like, or a mixture of shapes.

[0016] In one embodiment, the formulation aid is a polysaccharide, a protein, a lipid or a synthetic polymer, or a mixture thereof.

[0017] In one embodiment, the polysaccharide is positively charged, negatively charged, or neutral.

[0018] In one embodiment, the polysaccharide is selected from the group consisting of maltodextrin, starch, xanthan gum, gellan gum, alginate, pectin, dextran and chitosan.

[0019] In one embodiment, the protein is a dairy protein, a non-dairy animal protein, a plant protein, an algal protein, or a fermentation product protein.

[0020] According to a second aspect of the present invention, there is provided the use of a positively or negatively charged high voltage power source to encapsulate living microorganisms, said microorganisms having a surface charge identical to the charge of the high voltage power source.

[0021] 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, wherein the at least one cell is located in the core of the particle, and wherein the polymer composition of the particle is uniform and forms a substantially cell-free shell around the core.

[0022] According to a fourth aspect of the present invention, there is provided a particle, the particle comprising living microorganisms, the particle being obtainable by the method according to the first aspect.

[0023] According to a fifth aspect, there is provided use of the particle according to the third aspect or the fourth aspect for manufacturing a food, a feed product, a dietary supplement or a medicine.

[0024] According to a sixth aspect, there is provided a food, 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

[0026] Figure 1 is a schematic diagram of the EHD drying setup.

[0027] Figure 2 are two graphs showing the effect of polarity on water loss (A) and viability (B) of EHD-dried BIFIDO probiotics. Each pair of samples (S1-S2, S3-S4, S5-S6) showing statistically significant differences (p<0.05) between the results and the reference sample (R0) is represented by a different letter.

[0028] Figure 3 are two graphs showing the effect of the distance between the needle and the sample holder on the water loss (A) and viability (B) of EHD-dried BIFIDO probiotics. Each pair of samples (S1-S7, S5-S9) showing a statistically significant difference (p<0.05) between the results and the reference sample (R0) is represented by a different letter.

[0029] Figure 4 are two graphs showing the effect of temperature on water loss (A) and viability (B) of EHD-dried BIFIDO probiotics. Samples (S11, S14, S16) showing statistically significant differences (p<0.05) between the results and their corresponding reference samples (R0, R13, R15) are indicated by different letters.

[0030] Figure 5 Two graphs showing the effect of the number of needles and the distance between the needles and the mobile filter on the water loss (A) and viability (B) of EHD-dried BIFIDO probiotics. Samples (MF4, MF5, MF2, MF3) showing statistically significant differences (p<0.05) between the results and the reference sample (R0) are indicated by different letters.

[0031] Figure 6 Two graphs showing the effect of the number of needles and the distance between the needles and the mobile filter on the water loss (A) and viability (B) of EHD-dried BIFIDO probiotics. Samples (MF4, MF5, MF2, MF3) showing statistically significant differences (p<0.05) between the results and the reference sample (R0) are indicated by different letters.

[0032] Figure 7 Two graphs showing the effect of different drying settings on the water loss (A) and viability (B) of BIFIDO probiotics dried for 2 hours. Samples (S1, SM1, MF4, FD, FDM) showing statistically significant differences (p<0.05) between the results and the reference samples (R0, R17) are indicated by different letters.

[0033] Figure 8 is a graph showing the FTIR spectrum of BIFIDO probiotics dried for 2 hours. Samples S1, MF4 and FD were dried in different settings, and the reference sample R0 was non-dried BIFIDO.

[0034] Fig. 9 is a graph showing the FTIR spectra of BIFIDO probiotics dried for 2 hours with and without the addition of maltodextrin. Samples S1, SM1 and FD, FDM were dried in different settings.

[0035] Fig.10 Two graphs showing the effect of different drying settings on the zeta potential (A) and hydrophobicity (B) of BIFIDO probiotics dried for 2 hours. Samples (S1, SM1, MF4, FD, FDM) showing statistically significant differences (p<0.05) between the results and the reference samples (R0, R17) are indicated by different letters.

[0036] Fig.11 It is a KH display of probiotics at different pH values. 2 PO 4 Diagram of the zeta potential in .

[0037] Fig.12 is a graph showing the cell surface hydrophobicity (expressed as hexadecane adhesion %) of probiotic cells under different pH conditions.

[0038] Fig.13 are confocal laser scanning microscopy micrographs showing the effects of voltage and polarity on the "self-organization" of cells within microcapsules.

[0039] Fig.14 is a confocal laser scanning microscopy micrograph showing the distribution of cells within electrosprayed microcapsules.

[0040] Fig.15 are two graphs showing the effect of electric field polarity and voltage on the glass transition temperature (Tg) of electrospray microcapsules. Fig.15 A shows data for cells with 75% w / v maltodextrin and 10% v / v, and Fig.15 B shows data for cells with 75% w / v maltodextrin.

[0041] Fig.16 are FTIR spectra of probiotics with and without the influence of an external electric field.

[0042] Fig.17 is a graph showing the loss of viability of encapsulated cells and non-encapsulated cells (free cells) using negative and positive polarity electrospray. DETAILED DESCRIPTION OF THE INVENTION

[0044] The present inventors have found that by utilizing electrohydrodynamics, encapsulation of microorganisms can be achieved resulting in cost savings and / or improved process efficiency and / or improved stability of the final product.

[0045] This is achieved by a method for electrostatically spray drying live microorganisms, the method comprising the steps of 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; drying the droplets to form dry particles; and collecting the dry particles. The polarity of the electrostatic charge should be the same as the surface charge of the microorganism. This produces particles comprising a polymer and at least one cell of the live microorganism, wherein the at least one cell is located in the core of the particle, and wherein the polymer composition of the particle is uniform and forms a shell substantially free of cells around the core. Such a particle structure protects the microorganism, thereby improving the stability of the product.

[0046] Electrohydrodynamics can also be used to obtain particles with microorganisms close to the particle surface rather than in the particle core. This is achieved by applying an electrostatic charge that is opposite to the surface charge of the microorganisms.

[0047] Preferably, the suspension comprises one or more formulation aids. At least part of the formulation aids can be added to the suspension before the suspension is applied with an electrostatic charge. Formulation aids are usually provided in the form of drying protective agents, wherein the effect of the drying protective agent is to stabilize the microorganisms in the suspension, droplets and / or dry particles. Preferably, the drying protective agent is selected so that its effect is in the electrostatic spray drying process itself, and also in the process of using the dry particles and / or microorganisms subsequently, including storage, transportation and / or additional processing, reducing, for example, preventing the killing of the microorganisms.

[0048] The present invention also relates to particles comprising live microorganisms embedded in a large amount of formulation aids. The particles are preferably dense so that the particles have an internal total void volume less than 5% of the total volume of the particles. In addition, in a preferred embodiment of the present disclosure, the microorganisms are not present on the surface of the particles. Therefore, the microorganisms are preferably embedded in the formulation aids of the dry particles. In an embodiment of the present disclosure, the dry particles basically contain a single phase except the microorganisms. Preferably, the dry particles are therefore not layered dry particles, such as dry homogeneous particles encapsulated in a protective layer. In a typical embodiment of the present disclosure, the dry particles have a substantially continuous radial gradient of formulation aids. Therefore, the dry particles can have a high concentration of formulation aids, such as 100%, on the surface. The concentration of the formulation aid can be continuously reduced toward the center of the particle, thereby forming a radial gradient of the formulation aid. Preferably, the center of the particle has the highest concentration of microorganisms.

[0049] In the embodiment of the present disclosure, applying electrostatic charge makes polar components be forced towards the droplet surface, and the component with less polarity in the suspension is then forced towards the droplet center.Usually, the polarity of different components has an impact on the resulting electrostatic charge distribution of the formed droplet.For example, the electronics provided to the suspension are usually relevant to the component with stronger polarity (usually solvent), thereby producing electrostatic repulsion between the polar components, forcing the polar solvent and the polar formulation auxiliary agent dissolved in the solvent towards the droplet surface.Similar effects can further force microorganisms towards the droplet center, encapsulated by formulation auxiliary agents.Forcing the solvent to cause faster evaporation rate usually towards the surface, and embedding microorganisms in the droplet center then causes improved microorganism encapsulation.

[0050] In an embodiment of the present disclosure, an electrostatic charge is applied to the suspension by an electrode in contact with the suspension, and wherein the electrode has a pulsed potential difference relative to ground. Preferably, the potential difference, i.e., the voltage, has a constant polarity. Therefore, the electrode can be part of a direct current (DC) circuit, in which current flows in only one direction, and the electrode remains negative or positive at all times. Therefore, the droplets formed can each have an overall negative charge or an overall positive charge. Typically, the pulsation of the potential difference results in an improved charging of the suspension, and further, it can also result in an improvement in the properties of the dry particles.

[0051] In an embodiment of the present disclosure, an electrostatic charge is applied to the suspension by contacting the suspension with at least one electrode having a potential difference, i.e., a voltage, relative to ground. Therefore, electrodes can be provided in a configuration for applying the voltage to the suspension. Additional electrodes, such as two or three or four or even more additional electrodes, can be provided for applying the electrostatic charge to the suspension. Preferably, two or more electrodes have the same polarity, such as, for example, a positive voltage or a negative voltage provided in a DC circuit. Electrode configurations for applying an electrostatic charge are known to those skilled in the art and may include the use of specific materials, surface areas, and shapes of one or more electrodes.

[0052] In an embodiment of the present disclosure, the electrode has a potential difference relative 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 not to cause any damage to the microorganisms. Therefore, the voltage should be low enough not to kill live microorganisms.

[0053] In an embodiment of the present disclosure, the electrode has a fixed polarity relative to grounding, such as a fixed negative polarity or a fixed positive polarity. Therefore, an electrode or more than one electrode can be configured to apply a direct current circuit (DC) voltage. Preferably, the electrode is configured to continuously supply electrons to the suspension or continuously discharge electrons from the suspension. Typically, the positive electrode (anode) discharges electrons from the suspension, and the negative electrode (cathode) supplies electrons to the suspension. The electrode can temporarily have a ground potential, i.e. 0V.

[0054] In an embodiment of the present disclosure, the potential difference of the electrodes relative to ground is constant. Therefore, the voltage of the electrodes can be constant, and therefore, electrostatic charge can be delivered to the suspension by using a constant voltage.

[0055] In another embodiment of the present disclosure, the potential difference of the electrode relative to ground changes over time, such as periodically. The periodic variation can be described by a wave function (e.g., a sine wave) or a combination of more than one wave function, which produces the periodic variation. The periodic variation can be described by two or more voltage levels, and the voltage of the electrode changes in the form of periodic variation. One of the voltage levels can be grounded.

[0056] In an embodiment of the present disclosure, the potential difference of the electrode relative to ground is varied periodically, for example, in a periodic step function. The voltage of the electrode may be varied according to any function and may depend on the parameters of the process, such as the feed rate of the suspension, the droplet size, the contents of the suspension, such as the types of components and their relative proportions, and further, the desired parameters of the dried particles.

[0057] In an embodiment of the present disclosure, the potential of the electrode is applied by pulse width modulation, for example, by a square wave. Therefore, the voltage of the electrode can change between two or more set levels, forming a square wave, wherein the time between two pulses can be a set value, or can vary depending on the parameters of the treatment method mentioned elsewhere herein. The voltage can be provided as a pulse between two or more voltage values, wherein the dwell time at each level can be set individually, and wherein one of the voltage levels can be 0V.

[0058] In embodiments of the present disclosure, components of the suspension are distributed in the formed droplets according to their polarity, e.g., to increase evaporation of the solvent and / or to increase encapsulation of the microorganisms. The distribution of the suspension components can result in the formed droplets having advantageous properties, e.g., increased evaporation, e.g., reduced evaporation time and / or less water content in the final dried particles, and / or improved encapsulation of the dried particles. Thus, the suspension components can be selected based on how they are distributed in the formed droplets (e.g., electrostatically charged formed droplets).

[0059] In an embodiment of the present disclosure, components of the suspension with higher polarity are distributed to the surface of the droplets, while components of the suspension with lower polarity are distributed to the center of the droplets. In an embodiment of the present disclosure, at least two components of the suspension have different dielectric properties. Preferably, there is a correlation between the distribution of the components of the suspension in the droplets and the dielectric properties and / or effective dielectric properties of the components of the suspension.

[0060] In an embodiment of the present disclosure, the microorganism has an effective dielectric property lower than that of a formulation aid and / or a solvent. In an embodiment of the present disclosure, the solvent has a dielectric constant higher than that of the formulation aid, and the formulation aid has a dielectric constant higher than that of the microorganism. Among solvents, formulation aids and microorganisms, preferably the solvent has the highest dielectric property, and the formulation aid has a dielectric property higher than that of the microorganism. The dielectric property of the microorganism can be measured and / or provided as an effective dielectric property, wherein the overall dielectric property of the microorganism is provided, rather than the dielectric property of individual components such as specific membrane proteins, which is known to those skilled in the art, for example, in Sanchis et al., Dielectric characterization of bacterial cells using dielectrophoresis, Bioelectromagnetics, 2007.

[0061] In an embodiment of the present disclosure, droplets are formed by atomizing a suspension. Atomization and spraying are used interchangeably herein and refer to the process of forming a plurality of droplets from a liquid (e.g., a suspension) of a larger volume (e.g., a raw material). Due to the increase in surface area, the formation of droplets (i.e., the liquid volume is decomposed into smaller droplets) requires energy. Interfacial energy (usually given at the liquid-gas interface of the droplet surface) requires additional energy to form. As known to those skilled in the art, energy can be supplied in a variety of ways.

[0062] In embodiments of the present disclosure, droplets are formed by an atomizing device such as an ultrasonic nozzle, a pressure nozzle, a two-fluid nozzle (e.g., using CO 2 or N 2 or other gases as atomizing gas), vibrating nozzles, frequency nozzles, electrostatic nozzles; or rotary atomizing devices. Different types of nozzles are known to those skilled in the art, and they may have their own advantages. Several parameters may be adjusted, including the flow rate of atomizing gas, the flow rate of suspension / raw material, the use of surfactants, the configuration of the nozzle, the type of nozzle, the force acting on the suspension (gravity, electricity, centrifugal force or other force), depending on the desired properties of the droplets and therefore the desired properties of the dry particles.

[0063] In an embodiment of the present disclosure, the formation of droplets is carried out by a two-fluid nozzle. The two-fluid nozzle atomizes a liquid (e.g., a suspension) by the interaction between a high-speed gas and a liquid (e.g., a suspension). Compressed air is usually used as an atomizing gas, but other gases such as steam may also be used. The two-fluid nozzle may be an internal mixing type or an external mixing type, depending on the mixing point of the airflow and liquid flow relative to the nozzle face.

[0064] In an embodiment 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 to form droplets by using the atomizing gas.

[0065] In an embodiment of the present disclosure, the atomizing gas is selected from the group consisting of an inert gas (eg, nitrogen and carbon dioxide), a noble gas (eg, helium, argon, or neon), and an alkane gas (eg, methane), or a mixture thereof.

[0066] In an embodiment of the present disclosure, the atomizing gas comprises or consists of nitrogen, carbon dioxide and / or atmospheric gases or mixtures thereof. The gas may be treated in any suitable manner prior to use, including filtering, sterilizing and / or dehumidifying. However, in an embodiment of the present disclosure, the atomizing gas is not dehumidified. The use of a non-dehumidified gas may be advantageous in a number of ways, including providing simpler operation, reducing cost and time, and may further produce better droplet and / or dry particle properties.

[0067] In embodiments of the present disclosure, the atomizing gas has a water 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.

[0068] In embodiments of the present disclosure, the droplet formation step (e.g., spraying step) is performed at an atomizing gas inlet temperature of up to about 200°C, such as a range of about 20°C to about 200°C, such as a range of about 40°C to about 150°C, or such as a range of about 40°C to about 120°C, such as about 40°C to about 90°C, such as about 50°C to about 90°C, such as about 60°C to about 85°C, such as about 80°C. It may be desirable to have an elevated temperature relative to normal room temperature for optimal droplet formation, solvent evaporation, and / or dry particle properties. However, elevated temperatures may also be detrimental to components of the suspension (e.g., microorganisms). Therefore, the optimal temperature may not only be based on the simplest mode of performing the method (e.g., at room temperature), but may be a contribution of multiple factors, including the ease of performing the method, rapid evaporation, viability of live microorganisms, specific properties of the dry particles (e.g., optical packaging of microorganisms), sufficiently low volume of gas-filled voids within the dry particles (compactness of the dry particles), and other parameters. Therefore, the optimal temperature is important and based on multiple factors. Preferably, the temperature is above room temperature (e.g., to allow for rapid evaporation), but should be below a temperature that is unfavorable to the microorganisms, given the specific temperatures experienced by the microorganisms, since the microorganisms are preferably thermally isolated during the drying process by being embedded in the formulation aids and solvents (at least temporarily). Thus, the temperature may be in the range of about 20°C to about 200°C.

[0069] In an embodiment of the present disclosure, the atomizing gas has an inlet pressure in the range of about 1 kPa to about 500 kPa, for example, in the range of about 5 kPa to about 500 kPa, for example, in the range of about 5 kPa to about 300 kPa, for example, in the range of about 5 kPa to about 100 kPa, for example, about 60 kPa, or for example, about 70 kPa, or for example, about 80 kPa, or for example, in the range of about 100 kPa to about 400 kPa, for example, about 120 kPa, or about 150 kPa, or about 200 kPa, or about 250 kPa, or about 300 kPa, or about 350 kPa.

[0070] In an embodiment of the present disclosure, the atomizing gas has an inlet pressure in the range of about 50kPa to about 400kPa. Typically, the inlet pressure of the atomizing gas is defined as the pressure of the atomizing gas before being supplied to a nozzle (e.g., a two-fluid nozzle). Typically, the inlet pressure of the atomizing gas is the pressure of the atomizing gas supplied to a spray drying device (e.g., an electrostatic spray drying device). Gas can be supplied by a gas tank that includes at least one pressure regulator for controlling 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 atomizing gas can be reduced, which is typically due to the high resistance in the atomizing nozzle combined with the large cross-section of the drying chamber.

[0071] In embodiments of the present disclosure, the dry particles have a size of about 1 micron to about 800 microns, for example, about 5 microns to about 800 microns, for example, about 10 microns to about 600 microns, for example, about 10 microns to about 300 microns, for example, about 10 microns to about 200 microns, for example, about 10 microns to about 50 microns, or for example, about 50 microns to about 200 microns, for example, about 50 microns to about 100 microns, for example, about 75 microns, or for example, about 100 microns to about 200 microns, for example, about 150 microns. As known to those skilled in the art, Dv50 is typically used to represent the median value of volume distribution.

[0072] In embodiments of the present disclosure, the particle size distribution of the dry particles is substantially unimodal.

[0073] In an embodiment 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 an embodiment of the present disclosure, the method comprises a drying step for drying the formed droplets.

[0074] In an embodiment of the present disclosure, the method includes a drying step, and wherein the drying of the droplets is performed under reduced pressure. Reduced pressure may be used to improve evaporation, for example to increase the evaporation rate, and further to control or direct the dry particles into a dedicated collector at the outlet end of the drying chamber, wherein the collector may be in the form of a filter or a sieve. Reduced pressure may have other advantages, such as facilitating the recirculation of the atomizing gas.

[0075] In an embodiment of the present disclosure, the method comprises a drying step of the formed droplets (wet particles), and wherein the water activity (a w ) is less than about 1.0, for example in the range of about 0.01 to about 0.6, for example about 0.05 to about 0.5, for example about 0.1 to about 0.5, for example about 0.2, or for example about 0.3, or for example about 0.4. Water activity (a w ) is the partial vapor pressure of water in a substance divided by the standard state partial vapor pressure of water. Wherein the standard state is defined as the partial 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, wherein the level of water activity is set to a value where the type of microorganism is advantageously dried, for example, where viability is not significantly affected.

[0076] In an embodiment of the present disclosure, the method includes a drying step of the formed droplets (wet particles), and wherein the solvent (e.g., water) content of the dry particles is less than about 20% by weight, such as less than about 15% by weight, such as less than about 10% by weight, such as less than about 5% by weight, such as less than about 3% by weight, such as less than about 1% by weight, such as less than about 0.1% by weight, relative to the total weight of the dry particles.

[0077] In embodiments of the present disclosure, the solvent (e.g., water) content of the dry particles is less than about 10% by weight (preferably less than about 5% by weight, or less than about 1% by weight) relative to the total weight of the dry particles. Therefore, after the formed droplets are dried, a small amount of water may be present, and the small amount of water may even be advantageous because it may be beneficial to increase the viability of the microorganisms.

[0078] In an embodiment of the present disclosure, the dried particles are collected at the outlet end of the drying chamber. The outlet end may also be equipped with a collector for collecting the dried particles, such as a filter, a container and / or a sieve.

[0079] In an embodiment of the present disclosure, a filter (eg, an electrostatic filter) or a screen is used to collect the dried particles at the outlet end of the drying chamber. The steric interaction preferably blocks the passage of particles, while gases (such as atomizing gases) are allowed to pass through the collector.

[0080] In an embodiment of the present disclosure, a sieve is used to collect dry particles at the outlet end of the drying chamber, the sieve having an aperture of less than about 500 microns, such as in the range of about 40 microns to about 300 microns, such as in the range of about 50 microns to about 250 microns, such as about 50 microns, such as about 100 microns, such as about 150 microns, such as about 200 microns or such as about 250 microns. Preferably, the aperture is lower than the particle size provided according to Dv50, more preferably lower than Dv30, even more preferably lower than Dv10, still even more preferably lower than Dv5, such as lower than Dv3, such as lower than Dv1. The hole can be provided in a suitable shape, such as a circular opening and / or a slit (grid). Typically, the minimum size of the opening is the relevant size that hinders the particle. Therefore, for a slit, the aperture refers to the width of the slit, and the same reasoning applies to openings of other shapes.

[0081] In an embodiment of the present disclosure, the dried particles are collected at the outlet end of the drying chamber using a screen having a pore size ranging from about 40 microns to about 300 microns.

[0082] Preservation and Expert Solutions

[0083] At the applicant's request, only samples of the deposited microorganisms described below may be provided to the expert, subject to the applicable regulations administered by the industrial property office of a Contracting State to the Budapest Treaty until the date of grant of the patent.

[0084] The deposit was made in accordance with the Budapest Treaty on the International Recognition of the Deposit of Microorganisms for the Purposes of Patent Procedure with a depository that has obtained the status of an international depository institution: Leibniz Institute DSMZ - German Collection of Microorganisms and Cell Cultures, Inhofenstrasse 7B, 38124 Braunschweig, Germany. The deposit number and deposit details are shown in Table 1.

[0085] Table 1. Deposit information

[0086] Example

[0087] Example 1

[0088] Materials and methods

[0089] Maltodextrin (Dextrose Equivalent 12), commercially available as Glucidex IT12, was obtained from Roquette (Lestrem, France). Sodium chloride (NaCl) (Merck, Kenilworth, NJ, USA) and tryptone (Oxoid, Hampshire, UK) isotonic solutions were prepared for bacterial dilution. Dual staining LIVE / DEAD TM BacLightTM Bacterial viability was measured using a bacterial viability kit (L-7012, Molecular Probes, Eugene, OR, USA). The concentration was 3x10 11 The probiotic cells of Bifidobacterium animalis DSM 15954 (hereinafter referred to as BIFIDO) were obtained from Chr. Hansen A / S ( Denmark) and stored at −80°C until further use.

[0090] Electrohydrodynamic drying

[0091] Probiotics (3x10 11 Cells / g) were thawed and transferred (5 g) to a 55 mm x 80 mm aluminum sample holder. The samples were placed in a closed polycarbonate chamber ( Figure 1 ), wherein the temperature and relative humidity (RH = 20 ± 3%) are controlled by introducing air or nitrogen through a diffuser.

[0092] The EHD drying apparatus consisted of a monopolar DC high voltage generator (Gamma High Voltage Research, Florida, USA) connected to a computer that executed an algorithm to control the DC current applied to three metal needles (0.1 mm in diameter and 20 mm in length) that were 20 mm apart and placed vertically on a sample holder on which the BIFIDO sample was placed.

[0093] The samples were subjected to external electric field strengths ranging from 1.5 kV / mm to 6.6 kV / mm, while the current was automatically regulated by the power supply (AA).

[0094] The effects of applied polarity, voltage, gas, distance between the needle tip and the sample holder, addition of maltodextrin as an excipient, and room temperature were investigated during the EHD drying of BIFIDO. Table 2 summarizes the conditions tested.

[0095] Table 2. Test parameters

[0096]

[0097] "Setup 2" was developed to optimize "Setup 1". A mobile compartment (M) was used, in which a nylon filter membrane (MF) (Sigma-Aldrich, Steinheim, Germany) with a pore size of 0.45 μm and a diameter of 90 mm was placed on a metal mesh sample holder. Using a computer algorithm, the current intensity was always constant at 200 μ, 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).

[0098] For setups 1 and 2, the BIFIDO samples were subjected to an external electric field (EHD dried—S) or not (reference sample—R) for 1 and 2 hours.

[0099] Freeze drying

[0100] Probiotics (3x10 11 Cells / gram) were thawed and transferred (5 g) to a 55 mm x 80 mm aluminum sample holder (same as the EHD drying protocol). The BIFIDO samples were frozen at -20 °C for 24 h and then freeze-dried for 2 h (1,030 mbar) using a benchtop freeze dryer (Beta LMC 1-8, Chist, Germany), sample FDM (Table 2). The experiment was performed in duplicate.

[0101] Sample characterization

[0102] Water loss (%)

[0103] The initial weight of the sample and its subsequent weight after 1 and 2 h of EHD drying were measured using a digital balance with an accuracy of 0.001 g (Mettler Toledo, Greifensee, Switzerland). The water loss (WL) percentage of the sample drying experiment was calculated according to the following equation:

[0104]

[0105] Where Wo and W are the initial weight of the sample and the weight of the sample after EHD drying, respectively.

[0106] Cell viability (flow cytometry)

[0107] Using the double staining kit BacLight TM The kit was used to stain the dried probiotics, which consisted of the fluorophores SYTO9 (excitation / emission: 485 / 498 nm) and propidium iodide (PI) (excitation / emission: 535 / 617 nm). TM Bacterial viability was analyzed by flow cytometry using a C6Plus system (BD Biosciences, Franklin Lakes, NJ, USA). All analyses were performed at a low flow rate (14 μL / min), and the maximum volume of collected samples was fixed at 10 μL. The bacterial concentration of the samples was adjusted to approximately 1 × 10 3 Up to 5x10 6cells / ml and defined appropriate gates for SYTO9 and PI stained bacteria for sorting on FL1 vs FL3 dot plots (BD-Biosciences, 2012). Experiments were performed in triplicate.

[0108] FTIR

[0109] Infrared transmittance measurements were performed at room temperature (25°C) using a FT-IR spectrometer (Nicolet iS50, Thermo Fisher Scientific, Waltham, MA, USA). -1 The wave number range is 4cm -1 The spectra were recorded after 32 scans at a resolution of 1.34 nm.

[0110] Bacterial cell surface charge

[0111] The concentration is about 10 8 10 cells / mL of BIFIDO dried cells suspended in 10 mM KH 2 PO 4 (Sigma-Aldrich, Steinheim, Germany), and then transferred to a capillary cell with gold-plated beryllium / copper electrodes (DTS1070 cell, Malvern Panalytical Ltd, Malvern, UK). The electrophoretic mobility of the samples was measured using a Zeta-sizer (Malvern Panalytical Ltd, Malvern, UK) and converted into zeta potential (mV) using the Helmholtz-Schmoluchowski equation. Each sample was analyzed in quintuplicate, and all measurements were performed at 25°C.

[0112] Bacterial cell surface hydrophobicity

[0113] The hydrophobic / hydrophilic properties of the BIFIDO cell surface after EHD drying and freeze drying were evaluated by the microbial adhesion hexadecane (MATH) assay (Deepika et al. 2009). BIFIDO probiotic cells were suspended in 10 mM KH 2 PO 4 , until the absorbance (OD600) is about 0.8. An equal volume of 2 mL of bacterial cell suspension and hexadecane (Thermo Fisher Scientific, Waltham, MA, USA) was vortexed for 1 min and then allowed to stand for 20 min until the phases were completely separated. The OD400 of the aqueous phase was carefully removed and measured.

[0114] The percentage of microorganisms adhering to hexadecane (or percentage hydrophobicity) was calculated using the following equation:

[0115]

[0116] Wherein, A0 is the initial absorbance of the bacterial suspension, and A1 is the absorbance at 600 nm after phase separation. The samples were prepared in triplicate.

[0117] Statistical analysis

[0118] The results are expressed as mean ± standard deviation (SD). To analyze the differences between samples, one-way ANOVA was used. When the p value was less than 5% (p<0.05), the sample results were considered to be statistically significantly different.

[0119] result

[0120] EHD Setup 1

[0121] Effect of polarity and gas type

[0122] The effect of polarity on EHD drying was studied and Figure 2 Shown in. Figure 2 A shows that water loss is a time-dependent EHD drying parameter, since after 2 h, an increase in WL water loss (WL) was observed for all samples over time.

[0123] Sample S1 (negative polarity) lost approximately 38.7±3.2% and 62.8±5.3% of water after 1 h and 2 h, 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 h and 2 h, respectively (statistically not significant).

[0124] Sample S3 dried under negative polarity (WL = 38.8 ± 3.2% (1 h), WL = 64.3 ± 0.3% (2 h)) lost a higher amount of water (P < 0.05) compared to sample S4 dried under positive polarity (WL = 7.7 ± 0.4% (1 h), WL = 17.5 ± 1.6% (2 h)). However, no significant differences were observed in terms of bacterial viability (V) when comparing samples dried using negative polarity (samples S1 and S3) or positive polarity (samples S2 and S4).

[0125] It has been shown that for the same voltage, negative corona produces higher corona current than positive corona (Lai and Wong 2003), which may explain the greater water loss observed for samples dried at negative polarity ( Figure 2). In addition, at low voltages (less than 10 kV / mm), the ion wind generated by the negative corona exhibits a higher velocity than the positive corona (S. Chen, Van Den Berg, and Nijdam 2018), which can also affect the evaporation of water from the EHD dried samples. The lower dielectric breakdown strength of air (1.15 times lower than nitrogen) allows the voltage of samples S5 and S6 to be increased to 10 kV. Sample S5 dried under the influence of negative corona discharge also showed significantly higher water loss (WL = 67 ± 5.4% (1 h), WL = 81.3 ± 1.6% (2 h)) and similar probiotic viability (V = 66.3 ± 39.6 (1 h), V = 25.7 ± 18.3% (2 h)) compared to sample S6 dried under the influence of positive corona discharge (WL = 9.3 ± 1.9% (1 h), WL = 18.7 ± 2.2% (2 h) and V = 83.3 ± 16.6% (1 h), V = 26.1 ± 16.3 (2 h)). This paper confirms that negative polarity contributes more to EHD drying than positive polarity when nitrogen or air is used as the surrounding gas. Compared with the reference sample (R0) (almost no drying), the samples dried using positive polarity showed similar water loss, highlighting the higher efficiency of EHD drying using negative polarity.

[0126] Effect of voltage

[0127] Figure 3 A shows that the water loss of samples S7 and S3 after drying for 1 hour increased from 11.1±1.6% to 38.9±3.2% as the voltage increased from 3.7 kV to 4.7 kV. After drying for 2 hours, samples S7 and S3 lost about 19.5±2.1% and 64.32±0.3% of water, respectively. Similar trends were observed for samples S8 (3.7 kV) and S5 (10 kV), where the water loss of sample S8 was about 10.7±0.3% (1 hour) and 21.9±0.6% (2 hours), while for sample S5, the water loss was about 67±5.4% (1 hour) and 81.3±1.6% (2 hours). An increase in the applied voltage has been shown to enhance the electric field by increasing the charge density current and the speed of the ion wind, thereby increasing the drying rate (Anukiruthika, Moses, and Anandharamakrishnan 2021; Defraeye and Martynenko 2019; S. Chen, Nobelen, and Nijdam 2017).

[0128] When nitrogen was used as the surrounding gas, no significant difference in bacterial viability was observed between samples S7 (3.7 kV), S3 (4.7 kV) and reference sample R0 ( Figure 3B). For sample S7, the viability of the probiotics decreased from 99.9±4.1% (1 h) to 95.9±3.2% (2 h), while in sample S3 the percentage of viable cells decreased from 84.4±12.1% (1 h) to 53±30.8% (2 h). The viability of the reference sample at both tested hours was about 99%.

[0129] When comparing samples S8 (3.7 kV), S5 (10 kV) with the reference sample (R0), significant differences were found in the percentage of live probiotics. For sample S8, the viability decreased from 96.3 ± 3.0% (1 h) to 86.6 ± 15.3% (2 h), while for sample S5, the viability decreased from 71.5 ± 26.8% (1 h) to 16.2 ± 13.4% (2 h). Figure 3 Of the samples tested, sample S5 had the highest evaporation rate and was therefore less active, in contrast to samples S7 and S8 which showed little drying.

[0130] Similar to other drying techniques, during EHD drying, bacteria experience dehydration and osmotic stress as water is removed from the cells and their surroundings. Therefore, increasing the voltage can be expected to enhance water evaporation and reduce bacterial viability.

[0131] Effect of the distance between the needle and the sample holder

[0132] Figure 4 The effect of the distance between the high voltage electrodes on each pair of samples is shown. Note that the sample S7 with a distance of 25 mm did not show a significant difference in water loss from the reference R0 (WL = 11.1 ± 1.6% (1 h), WL = 19.5 ± 2.1% (2 h)), while the sample S1 with a distance of 15 mm lost three times more water after two hours of drying (WL = 38.7 ± 3.2% (1 h), WL = 62.8 ± 5.3% (2 h)). However, the bacterial viability of samples S7 (V = 99.9 ± 4.1% (1 h) and V = 95.9 ± 3.2% (2 h)) and S1 (V = 78.3 ± 27.6% (1 h), V = 55.8 ± 33.6% (2 h)) was not statistically significant, indicating no statistical difference from the reference sample (R0).

[0133] Samples S5 (15 mm) and S9 (25 mm) (dried under air conditions) showed statistically significant differences in water loss in the first hour of EHD drying (WL = 67.0 ± 5.4% (S5), WL = 57.3 ± 3.2% (S9)), but not in the second hour of EHD drying (WL = 81.0 ± 1.6% (S5), WL = 79.6 ± 0.9% (S9)). Most of the water evaporation occurred in the first hour, while during the second hour of EHD drying, bacterial dehydration of S5 (V = 66.4 ± 39.6% (1 h), V = 25.7 ± 18.3% (2 h)) and S9 (V = 66.3 ± 39.5% (1 h), V = 26.1 ± 18.1% (2 h)) reached a critical point, where viability was significantly reduced compared to reference sample R0.

[0134] Similar to increasing voltage, reduction in electrode distance is also a parameter that affects corona discharge, forming a large ion current and increasing ion wind speed, thereby increasing the drying rate (Tirtha R.Bajgai et al.2006). The present results show that reduction in electrode spacing has an adverse effect on the removal of water from the sample.

[0135] Effect of temperature

[0136] The effect of nitrogen temperature on the EHD drying of samples S11, S14 and S16 is shown in ( Figure 5 ). Increasing the nitrogen temperature from 20°C to 30°C and 40°C increased the water loss of S11 (WL = 35.4 ± 2.7% (1h)), S14 (57.2 ± 10.2% (1h)) and S16 (76.3 ± 3.6% (1h)), respectively, while their corresponding activities decreased (V = 85.9 ± 18.5% (1h), V = 64.9 ± 28.5% (1h) and V = 19.3 ± 19% (1h)).

[0137] The combination of EHD drying and heated nitrogen at 30°C for sample S14 did not show a statistically significant difference in probiotic viability compared to S11 dried under the same electric field strength and ambient temperature (20°C). However, water evaporation from S14 reached the same amount of loss as S11 after two hours in the first hour of drying (WL = 66.9 ± 4.8% (1h)). Therefore, it can be concluded that combining EHD drying with increased gas temperature can increase water loss (due to convection and thermal diffusion effects from the core to the surface of the sample) without compromising bacterial viability (at least in the first hour of drying). Sample S16 showed accelerated water loss from the first hour of EHD drying at 40°C. However, it can be noted that the drying rate decreased, indicating that there was no significant difference in water loss after one hour of drying (WL = 76.3 ± 3.6% (1h)) and two hours of drying (WL = 81.4 ± 0.1% (2h)). However, it can be clearly observed that such rapid water loss significantly reduced the viability of the probiotics (V = 19.3 ± 19% (1 h), V = 4.9 ± 3.7% (2 h)).

[0138] EHD Setup 2

[0139] The influence of the number of needles and the distance between the needle and the sample holder

[0140] To improve the drying of probiotics, a moving compartment was added to periodically move the flat electrode (Setup 2). In addition, a mesh electrode and a nylon filter (located below the probiotics) were used to facilitate airflow (Mujumdar and Xiao 2019; Iranshahi, Martynenko, and Defraeye 2020).

[0141] The effects of the number of needles and the distance between the needles and the sample were also studied, and the results are presented in ( Figure 6) is shown in the figure. It was observed that neither the number of needles nor the electrode spacing resulted in significant differences in the water loss of samples MF4 (WL = 37 ± 5.3% (1 h), WL = 68 ± 2.3% (2 h)), MF5 (WL = 42.9 ± 5.6% (1 h), WL = 74 ± 0.1% (2 h)), MF2 (WL = 39.3 ± 2.5% (1 h), WL = 72.5 ± 3.5% (2 h)) and MF3 (WL = 41.5 ± 3.8% (1 h), WL = 73.5 ± 3.4 (2 h)). Nor did it lead to significant differences in the viability of the corresponding probiotics (for MF4, V = 89.2 ± 4.8% (1h), V = 60.8 ± 20.8% (2h); for MF5, V = 84.9 ± 10.8% (1h), V = 48.1 ± 28.5% (2h); for MF2, V = 88.4 ± 4.9% (1h), V = 33.3 ± 23.9% (2h); for MF3, V = 90.5 ± 4.9% (1h), V = 43.8 ± 23.9% (2h)). Nevertheless, significant differences were observed between all tested samples and the reference sample R0. The current experimental setup did not show significant differences by increasing the number of needles and the electrode spacing, so sample MF4 was selected for further study.

[0142] Comparison of EHD and freeze drying

[0143] Water loss and vitality

[0144] The effect of the two-hour drying process on the water loss and activity of probiotics was ( Figure 7 ) are shown in Figure 2. Overall, the samples MF4 (WL = 76.4 ± 2.2% (2 h)) and freeze-dried samples FD (WL = 76.8 ± 5.5% (2 h)) subjected to EHD drying using the optimized "setting 2" showed the highest water loss, however, no significant differences in their bacterial viability were observed (for MF4, V = 60.8 ± 20.8% (2 h); for FD, V = 45.5 ± 11.3% (2 h)).

[0145] Among the dried samples, the highest water loss was observed for MF4 and FD, followed by sample S1 with a lower water loss of 62.8 ± 5.3% (2 h). EHD dried samples S1 and MF4 showed statistically different water loss results, which means that the filter-mesh electrode indeed facilitated airflow and enhanced water evaporation (Mujumdar and Xiao 2019; Iranshahi, Martynenko, and Defraeye 2020).

[0146] The addition of maltodextrin reduced 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 different sugars, Adhikari (Adhikari et al. 2004) proposed that the addition of maltodextrin reduced the drying rate due to the difficulty of water diffusion through maltodextrin molecules. Similarly, Gianfrancesco et al. also reported that the water flux towards the surface of the amorphous maltodextrin matrix was reduced, thereby limiting the evaporation rate and the amount of water lost within the selected drying time (Gianfrancesco et al. 2012).

[0147] Nevertheless, SM1 (V = 72 ± 10.6% (2h)) showed enhanced viability compared to both freeze-dried samples. Protective sugars such as maltodextrin are known to reduce the membrane phase transition temperature (Tm) and maintain the membrane integrity of bacterial cells while dehydrating (Mille et al. 2004; Strasser et al. 2009). Despite this, no statistical difference was observed between SM1 and S1 (V = 55.8 ± 33.6% (2h)). Similarly, the probiotic viability of sample FD without maltodextrin (V = 45.5 ± 11.3% (2h)) and sample FDM with maltodextrin (V = 48.7 ± 10.2% (2h)) did not show significant differences.

[0148] Fourier transform infrared spectroscopy

[0149] Fourier transform infrared spectroscopy (FTIR) of BIFIDO ( Figure 8 ) shows the main characteristic bands of probiotic cells. For non-dried BIFIDO, at about 3280 cm -1 The band at (representing the stretching of the hydroxyl bonds (-OH)) appears more intense because the sample was not dried like the samples processed by electrohydrodynamics or freeze drying.

[0150] The 3000 cm-1 band does not exist in the spectrum of undried bacteria. -1 and 2800cm -1 , but all dried samples showed this region. More specifically, 2966 cm -1 、2936cm -1 and 2876cm -1The bands at represent the asymmetric CH3 stretch, asymmetric CH2 stretch, and symmetric CH3 stretch at the nonpolar site of the phospholipid bilayer, respectively, which are revealed after evaporation of water from the cells (Santos et al. 2015; Shakirova et al. 2010; Dianawati, Mishra, and Shaha 2012). The above groups are revealed after evaporation of the additional water present in fresh probiotics and are therefore present in the fatty acid region.

[0151] The transmittance of bacterial protein region appears in 1660-1500cm -1 More precisely, at 1639cm -1 The carbonyl stretching of the secondary amide (amide I) was found at 1545 cm -1 The broad peak found at 1639 cm indicates the NH bending of amide II. There is no clear evidence that the secondary structure of the protein was affected by the application of EHD drying or freeze drying process, as no amide II shift was observed. However, it should be noted that the fresh probiotic has a peak at 1639 cm -1 The peak at 1545 cm is stronger for freeze-dried samples FD and EHD-dried MF4. -1 The peak at 1250 cm is stronger. The EHD dried sample S1 shows similar intensities for both peaks. According to Bozkurt et al., increased contact of BIFIDO cells with water molecules would increase the absorption band of amide I, as observed in the current results (Bozkurt et al. 2019). Interestingly, the amide II adsorption intensity may also be related to the amount of water lost from the sample, as the amide II adsorption intensity of FD and MF4, which are shown to be drier, is stronger, that of S1 is not as strong, and that of fresh BIFIDO is greatly reduced. -1 The P=O vibration of PO2- was identified at 1100-980 cm -1 Broad carbohydrate and phosphate bands were observed at 1070 cm -1 and 1040cm -1 The maximum transmittance is observed at 1070 cm-1, which is due to the vibration of the COC group at the polar site of the phospholipid bilayer. -1 and 1040cm -1 The peak at is decreased, which is due to the prolonged contact with water molecules, as shown by Bozkurt et al. (Bozkurt et al. 2019).

[0152] Comparison of EHD dried samples with and without maltodextrin to freeze dried samples ( Fig. 9 ) is displayed.

[0153] The sample containing maltodextrin was at 2966 cm -1 There is no peak at 1639cm -1 The carbonyl stretching strength of the secondary amide (amide I) is 1548 cm - The NH bending of amide II at 1 is stronger. This may be attributed to the fact that the samples containing maltodextrin are characterized by a higher water content and similar to the fresh BIFIDO samples, the increased contact of BIFIDO with water molecules increases the absorption band of amide I. 1394-1310 cm -1 The broad peak at 1210 cm -1 Additional small peaks at 1150cm -1 The relatively strong peak at 1040 cm is attributed to the chemical structure of maltodextrin. The latter peaks represent the COC and CO bands dominated by the vibration of carbohydrate rings. In addition, the interaction between PO2 of bacterial envelope and maltodextrin was confirmed by the 1040 cm peak of dry BIFIDO. -1 to 1020 cm for samples with BIFIDO and maltodextrin -1 The latter can be seen by noting that the maltodextrin band shifts from 995 cm -1 Displacement to 1020cm -1 This interaction between the phospholipid sites of cellular lipids and sugars (although hydrogen bonds) has been proposed before and described in the literature (Oldenhof et al. 2005; Leslie et al. 1995; Dzuba, Leonov, and Surovtsev 2020). FTIR technology has been widely used to study the protective effect of sugars such as maltodextrin. However, it should be noted that the protective effect of such high molecular weight molecules is indirect by promoting the replacement of water on the cell surface with smaller sugars such as glucose (Santos et al. 2015).

[0154] Zeta potential

[0155] Bacteria in all samples were negatively charged, as previously suggested for other Bifidobacterium strains (Gómez Zavaglia et al. 2002). This negative cell surface charge at neutral pH may indicate the dominance of anionic domains such as strong acids (phosphate-based (lipo)teichoic acids) and weak acids (acidic polysaccharides and proteins) (Deepika et al. 2009).

[0156] according to Fig.10A, No significant differences were observed between the zeta potential values ​​of the non-dried and dried samples. Sample R0 as well as the EHD-dried or freeze-dried samples without maltodextrin (S1 and FD) showed lower absolute values ​​of zeta potential compared to the corresponding samples with maltodextrin (R17, SM1 and FDM). The latter results confirm that the FTIR spectra indicate that the maltodextrin molecules have interacted with the bacterial membrane surface. When the absolute zeta potential value of the colloidal system increases, more stable clusters are formed during electrophoresis. It can therefore be assumed that the interaction of sugars with the bacterial surface stabilizes the bacterial surface (Kodzińska et al. 2010).

[0157] Hydrophobicity

[0158] Microbial adhesion hydrocarbon test is often used to evaluate the cell surface hydrophobicity of bifidobacteria (Pelletier et al. 1997; Gómez Zavaglia et al. 2002). When bacteria are hydrophobic, they show high adhesion to hexadecane (%). Fig.10 As shown in B, all samples showed similar hydrophobicity values ​​( Fig.10 B). Interestingly, samples using setting 1, i.e. EHD dried samples, showed statistically significant results, being less hydrophobic compared to EHD 2, control and freeze dried samples.

[0159] Furthermore, the addition of maltodextrin in the same sample increased the hydrophobicity of BIFIDO. According to Shakirova et al. (2010), cell surface hydrophobicity is associated with a higher protein content on the surface of BIFIDO (Shakirova et al. 2010). Many other studies have shown that the presence of (glyco)protein substances and polysaccharides are responsible for the higher hydrophobicity and hydrophilicity, respectively (Collado, Meriluoto, and Salminen 2008; Boonaert and Rouxhet 2000; Van Der Mei et al. 2003). Therefore, as indicated by both the negatively charged characteristics and the high hydrophobicity of BIFIDO, it is suggested that the bacterial surface is rich in proteins, which is also observed in the FTIR spectrum.

[0160] in conclusion

[0161] Electrohydrodynamic drying can be used to dry probiotic cells. It was found that the EHD drying method depends on multiple parameters, such as the polarity and voltage of the ion wind, the type of collector, the ambient temperature and gas, the probiotic concentration, and the use of excipients. The optimized conditions using setting 1 allowed to achieve 70% drying and up to 34% cell viability. By dispersing the probiotics with maltodextrin, the viability of the probiotics could be increased by up to 50% after 2 hours. However, the addition of maltodextrin reduced the water evaporation rate by 30%. The performance of EHD (cell viability and water loss) was improved by promoting the drying airflow with a moving mesh flat-end electrode located below the probiotics (called "setting 2"). Compared with "setting 1", using this "setting 2", the water evaporation and cell viability of the probiotics were increased to up to 78% and 70%, respectively. In addition, a comparison of the EHD drying method with the freeze-drying method revealed that the survival and water evaporation rates of the probiotics were similar, while the cell surface properties were similar to those of non-dried cells.

[0162] Example 2

[0163] Materials and methods

[0164] Probiotic cells Bifidobacterium animalis, DSM 15954 (hereinafter referred to as BIFIDO), were obtained from Chr-Hansen A / S ( Maltodextrin with a dextrose equivalent of 12 (Glucidex IT12, Roquette, Lestrem, France) was used to encapsulate BIFIDO. Sodium chloride (NaCl) (Merck, Kenilworth, NJ, USA) and tryptone (Oxoid, Hampshire, UK) were used to prepare isotonic solutions. KH 2 PO 4 (VWR International, Leuven, Belgium) was used as the suspension medium for the evaluation of bacterial surface properties and hexadecane (Thermo Fisher Scientific, Waltham, MA, USA) was used for hydrophobicity measurements. Bacterial viability was determined using dual staining LIVE / DEAD TM BacLight TM Bacterial Viability Kit (L-7012, Molecular Probes, Eugene, OR, USA) was used for the staining of BIFIDO cells. Fluorescent dye Thiazole Orange (dye content is about 90%) (Sigma-Aldrich, USA) was used for staining of BIFIDO cells.

[0165] Assessing the physicochemical properties of bacterial cell surfaces

[0166] Bacterial cell surface charge

[0167] The concentration is about 3*10 8 BIFIFO probiotic cells were suspended in 10 mM KH 2 PO 4 , to obtain an optical density (OD600) of about 1.0. The pH of the solution was adjusted to 1, 2, 4, 6 and 8 with 1M HCl or 1M NaOH. Electrophoretic mobility was determined using a Zeta-sizer (Malvern Panalytical Ltd, Malvern, UK). The sample volume of 1 mL was injected into a folded capillary cell (DTS1070 sample cell, Malvern Panalytical Ltd, Malvern, UK) with a gold-plated beryllium / copper electrode. The measurement was carried out at 25 ° C, and each sample was analyzed in quintuplicate. Utilizing the Helmholtz-Schmoluchowski equation, electrophoretic mobility was converted into zeta potential.

[0168] Hydrophobic / hydrophilic properties of bacterial cell surfaces

[0169] The microbial adhesion hexadecane (MATH) assay was used to evaluate the hydrophobic properties of the BIFIDO cell surface at different pH values ​​(1, 2, 4, 6, and 8). The probiotic cells were suspended in 10 mM KH 2 PO 4 50 mL of the bacterial cell suspension was mixed with an equal volume of hexadecane, vortexed for 1 min, and allowed to stand for 20 min to ensure complete phase separation. After equilibrium, the aqueous phase was carefully removed and the OD400 was measured.

[0170] The percentage of microorganisms adhering to hexadecane was calculated using the following equation:

[0171]

[0172] Wherein, A0 is the initial absorbance of the bacterial suspension, and A1 is the absorbance after 20 min of incubation and phase separation (OD400).

[0173] Each measurement was performed in triplicate.

[0174] Electrospraying of probiotic solutions

[0175] Maltodextrin (75% w / v) was dispersed in Millipore water and stirred until a homogenous solution was obtained, then BIFIDO probiotic cells (10% v / v) were added and stirred until they were fully dispersed.

[0176] The electrospray device includes a high voltage generator (ES50P-10W, Gamma High Voltage Research, Inc., USA) to provide a voltage of 15-40 kV and a syringe pump (New Era Pump Systems, Inc., Farmingdale, NY, USA) for providing maltodextrin / BIFIDO dispersion at a flow rate of 0.03 mL / min. The maltodextrin BIFIDO capsules are collected horizontally on a steel plate covered with aluminum foil, which is placed at a distance of 10 cm from the end of a needle (27 gauge, 12 mm long, 0.21 mm inner diameter). The electrospray device is placed in a nitrogen-circulated room, and the temperature and relative humidity are 23° C. and 22±3%, respectively.

[0177] The polarity of the electrode connected at the tip of the syringe needle is positive or negative, and for simplicity, the sample is referred to as being electrosprayed under positive or negative polarity conditions, respectively. In addition, the charge applied to the needle tip is shown first, followed by the charge applied to the collector. For example, when -15 kV is applied to the nozzle tip and +5 kV is applied to the collector, it is referred to as (-15 kV)(+5 kV), and the absolute value of the potential is expressed as |20 kV|.

[0178] Confocal laser scanning microscopy

[0179] An inverted Zeiss LSM-710 confocal laser scanning microscope (Carl Zeiss MicroImaging GmbH, Jena, Germany) was used, which was equipped with a diode laser (405 nm), an argon laser (458, 488 and 514 nm), two HeNe lasers (543 and 633 nm) and three detectors and a transmission detector. Before encapsulation, the probiotic cells were stained with the fluorescent dye thiazole orange (dye content was about 90% (Sigma-Aldrich, USA)) and washed with NaCl 0.85% w / v. The dye was in powder form, so it was dissolved in dimethyl sulfoxide (DMSO) at a concentration of 42 μmol / L. To collect images, the samples were scanned using the Z series mode, and confocal fluorescence images were taken using 20x, 40x and 60x objectives. All images were collected at an excitation wavelength of 488 nm and an emission bandpass filter of 505-550 nm. Images were analyzed using ZEN software.

[0180] Fourier transform infrared spectroscopy

[0181] A Nicolet iS50 spectrometer (Thermo Scientific, NY, USA) was used in attenuated total reflectance (ATR) mode at 400–4000 cm -Fourier transform infrared (FTIR) spectra of the samples were recorded in the range of 1. All spectra were recorded in transmission mode at room temperature (25 °C) with a scanning resolution of 4 cm -1 The samples were scanned 32 times. Spectral analysis was performed using Omnic software (Thermo).

[0182] BIFIDO pure culture (without maltodextrin as encapsulation compound) was electrosprayed with negative (-15 kV) (+5 kV) and positive (+15 kV) (-5 kV) polarity and 200 μL of electrosprayed sample was evaluated by ATR-FTIR. The same volume of BIFIDO not treated with external electric field was also evaluated as a control.

[0183] Differential Scanning Calorimetry

[0184] Differential scanning calorimetry (DSC) experiments were performed to determine the glass transition temperature (Tg) of the electrospray microcapsules. Thermograms of the samples were obtained using a DSC 250 (TA Instruments, New Castle, Delaware, USA) equipped with a Refrigerated Cooling System 90. The heat flow and temperature of the instrument were calibrated using distilled water (melting point (mp) = 0°C; DHm = 334 J / g) and indium (mp = 156.5°C; DHm = 28.5 J / g). Nitrogen was used as a carrier gas at a flow rate of 50 mL / min.

[0185] About 5 mg of electrospray microcapsules (collected immediately after treatment) were placed in a pre-weighed standard DSC aluminum pan (Tzero aluminum sealed pan (Tzero aluminum Hermetic pans, TA Instruments, USA) and sealed. An empty sealed aluminum pan was used as a reference. The pan was first equilibrated at 10°C for 5 min and then heated to 120°C with a 3°C / min ramp. According to the Tg of maltodextrin DE12, the run settings were set to be at least 30°C lower and higher than the expected Tg of the electrospray microcapsules.

[0186] The obtained DSC thermograms were analyzed using Trios software interfaced with the DSC and the Tg was determined from the start, midpoint and end point of the curve shift. All measurements were performed in triplicate and the Tg values ​​were averaged.

[0187] Stability of microencapsulated BIFIDO during storage

[0188] Microcapsules containing BIFIDO electrosprayed with negative (-15 kV) (+5 kV) and positive (+15 kV) (-5 kV) polarity and non-encapsulated animal Bifidobacterium as a reference sample were stored at 25°C in an incubator with a relative humidity of 35 ± 2% RH (Model KB 8400F, A / S Ninolab, Strand, Denmark). The viability of encapsulated and non-encapsulated BIFIDO cells was determined over time (0-14 days) by colony forming unit (CFU) analysis. In particular, electrospray microcapsules were dispersed in 0.85% w / v sodium chloride (NaCl) and 0.1% w / v peptone (tryptone) solution, homogenized by vortexing, and then 0.1 mL of appropriate ten-fold serial dilutions were coated on de Man Rogosa Sharpe (MRS) agar plates supplemented with 0.05 w / V% L-cysteine ​​hydrochloride monohydrate (Merck, Kenilworth, NJ, USA). MRS agar plates were incubated at 37 ° C under anaerobic conditions (AnaeroGen, Oxoid, Hampshire, UK) for 3 days, and cell viability was calculated as the average of 4 plates.

[0189] The encapsulation efficiency (EE) of the microcapsules was determined by dividing the number of viable cells (N) within the capsules by the number of viable cells (N0) in the initial solution and was expressed by the following equation:

[0190]

[0191] Statistical analysis

[0192] The results are expressed as mean ± standard deviation (SD). One-way ANOVA was used to analyze the sample differences for each comparison. When the p value was less than 5% (p < 0.05), the sample results were considered to have statistically significant differences.

[0193] Results and discussion

[0194] Bacterial cell surface charge and cell surface hydrophobic / hydrophilic properties

[0195] Fig.11The zeta potential of BIFIDO in potassium dihydrogen phosphate buffer is shown as a function of pH. The electronegativity of the cell surface decreases with decreasing pH, showing negative zeta potential values ​​above pH 2 and slightly positive values ​​at pH 1. Similar zeta potential characteristics have been previously reported for other Bifidobacterium strains (Bifidobacterium bifidum and Bifidobacterium pseudolongum) (Gómez Zavaglia et al., 2002), due to the fact that the surface components of the cells are dominated by anionic domains including strong acids (phosphate-based (lipoteichoic acids)) and weak acids (acidic polysaccharides and surface proteins) (Deepika et al., 2009). The gradual protonation of the above chemical groups can explain the decrease in the electronegativity of bacteria with decreasing pH (Pelletier et al., 1997).

[0196] The hydrophobicity of the BIFIDO cell surface was evaluated by studying the adhesion of BIFIDO to hexadecane ( Fig.12 ). At pH 1-6, BIFIDO showed similar affinity for hexadecane (80 ± 6.9%), indicating the hydrophobic nature of the surface. At higher pH (pH = 8), BIFIDO showed lower hydrophobicity, with the affinity percentage for non-polar hexadecane being less than 40%. The covalently bound proteins, S-layer proteins, and fatty acids present on the surface of BIFIDO cells (Dianawati & Shah, 2011; Shakirova et al., 2010, 2013) have been proposed to promote adhesion to hydrophobic solvents and contribute to the increased hydrophobicity of the bacteria.

[0197] Effects of electric field polarity and voltage on cell self-organization in microcapsules

[0198] The effects of negative and positive electric field polarity on the probiotic cell distribution in electrosprayed maltodextrin-BIFIDO capsules at pH 6 were investigated by confocal laser scanning microscopy (CLSM). Fig.13 ).

[0199] Negative polarity at the needle resulted in probiotic cell "self-organization" primarily at the core of the electrospray capsule. This was due to the electrostatic repulsion of the negatively charged surface BIFIDO cells (pH 6) by the negatively charged electric field of the needle wall and the inner surface of the Taylor cone. Similar BIFIDO cell "self-organization" at the core of the electrospray capsule was observed by using different voltages with negative polarity at the needle ( Fig.13 ). In contrast, when positive polarity is applied at the needle, cells are distributed toward the outer surface of the electrospray capsule ( Fig.13 ).

[0200] Fig.14 CLSM of electrosprayed maltodextrin / BIFIDO dispersions at different pH values ​​are shown. Similar to the results observed at pH 6, when a negative electric field was applied at the needle, the BIFIDO cells with negative surface charges “self-organized” at the capsule core at pH 4-8, while the cells (with a slightly positive surface charge) were distributed closer to the outer surface of the capsule at pH 1. Therefore, the encapsulation of BIFIDO in the core and outer surface of the capsule is due to the movement of the charged cells under the action of Coulombic and electrophoretic forces in the DC electric field. In addition, it is worth noting that since the probiotics exhibit similar hydrophobic properties at pH 1-6 ( Fig.12 ), so the “self-organization” of cells within the electrospray capsule is due to their surface charge rather than the hydrophobic / hydrophilic properties of the cell surface.

[0201] Glass transition temperature of electrosprayed maltodextrin / PROBIO capsules

[0202] The effects of voltage and electric field polarity on the glass transition temperature (Tg) of maltodextrin-BIFIDO electrospray capsules were also evaluated. Tg is related to the water content of the electrospray capsules (Tg increases with decreasing water content) and to the subsequent stability and viability of the encapsulated cells (Drake et al., 2018; Haque & Roos, 2004; Terpou et al., 2019). In order to achieve stability during storage of cells in a dry state, sufficient water needs to be removed and a non-crystalline glassy solid matrix needs to be formed (Drake et al., 2018).

[0203] The glass transition temperature (Tg) of electrosprayed maltodextrin capsules without probiotic cells slightly changed with increasing electrospray voltage for both positive and negative polarity. It was observed that the Tg value of maltodextrin capsules was slightly higher using positive polarity electric field. In contrast, electrosprayed maltodextrin-BIFIDO capsules ( Fig.15) showed a maximum Tg value at |20kV|((-15kV)(+5kV)), and then the Tg value decreased with the increase of the electrospray voltage. In addition, the maltodextrin-BIFIDO capsules treated with negative polarity are characterized in that their Tg values ​​are much higher than those of the capsules produced using positive polarity. In particular, the maltodextrin-BIFIDO capsules produced using the negative polarity of (-15kV)(+5kV) are characterized in that their Tg is 68.21±1.66°C, which is significantly higher than other electrospray capsules treated with negative polarity. When using positive polarity (+15kV)(-5kV), Tg is about 10°C lower than that of the capsules produced using the negative polarity (-15kV)(+5kV) of the same voltage. As indicated above, at pH 6, the probiotic cells (negatively charged on the surface) electrosprayed using a negative polarity electric field are repelled and "self-organized" in the core of the capsule, while the solvent accumulates near the surface of the droplets. Therefore, the evaporation of the solvent from the highly charged droplet surface during electrospraying was enhanced when a negative electric field was applied compared to when a positive electric field was applied. This enhancement of evaporation resulted in lower moisture content and subsequently higher Tg values ​​for negatively electrosprayed maltodextrin-BIFIDO capsules compared to the corresponding capsules treated with positive polarity. Therefore, different electric field voltages and polarities can affect the thermal properties and glass transition of electrosprayed capsules due to the presence of surface-charged probiotics.

[0204] Fourier transform infrared spectroscopy

[0205] FT-IR spectra of (non-encapsulated) electrosprayed BIFIDO and BIFIDO not treated with an external electric field under different electric field polarities ( Fig.16 ) is shown in the figure. At about 3280cm -1 The band at represents the stretching of the hydroxyl bond (-OH) and is critically related to the water phase of the sample. The band of untreated BIFIDO appears to be stronger, while the lowest hydroxyl stretching intensity ( ) is observed for BIFIDO electrosprayed with negative polarity (-20 kV). Fig.17 A). It has been previously reported in the literature (Bozkurt et al., 2019) that the intensity of the -OH band increased after FTIR measurement of spin-coated BIFIDO biofilms in a water-wet environment.

[0206] Samples with different treatments at 3000 cm -1 and 2800cm -1 The difference in the intensity of the fatty acid bands between the two samples is obvious, with the spectrum of the electrospray sample being stronger ( Fig.16 ). 2961, 2930 and 2876cm -1The bands at represent asymmetric CH3 and CH2 stretching and symmetric CH3 stretching at nonpolar sites of the phospholipid bilayer, respectively (Dianawati et al., 2012; Santos et al., 2015; Shakirova et al., 2010). A higher transmission intensity was observed for BIFIDO using negative polarity electrospray compared to positive polarity electric field. The presence of water has been shown to hamper the intensity of these groups (Dianawati et al., 2012), meaning that less water was present in bacteria electrosprayed at -20 kV, more water in bacteria electrosprayed at +20 kV, and even more water in untreated bacterial samples, which were completely absent from the interferogram.

[0207] The main protein amide band of biological samples usually appears at a frequency of about 3200 cm -1 (Amide A), 1660-1500cm -1 (amide I and amide II) and 1310-1240cm -1 (amide III) (Gautier et al., 2013; Krimm & Bandekart, 1986). For non-electrospray BIFIDO, the -1 Compared with the amide II peak at 1640 cm -1 However, for electrosprayed BIFIDO, it was observed that the peak of amide I was less distinct than that of amide II, and the intensity of this peak was also minimal for BIFIDO using negative polarity electrospray ( Fig.16 ). Bozkurt et al. investigated the effect of relative humidity on the surface structure of spin-coated BIFIDO biofilms and concluded that an increase in amide I absorption indicates increased contact between BIFIDO and water molecules (Bozkurt et al., 2019). Therefore, a decrease in the amide I peak intensity of samples electrosprayed at negative polarity indicates a drier sample.

[0208] In addition, differences in the intensity of the band at the polar sites of the cellular lipid bilayer (at 1070 cm due to the vibration of the COC group) were observed between the samples. -1 and 1040cm -1 ( Figure 6 The transmittance intensity of non-electrosprayed BIFIDO decreased, indicating expanded contact between cells and water molecules, while the highest intensity found in samples electrosprayed at negative polarity indicated that the least amount of water was present in BIFIDO using negative polarity electrospray compared to samples electrosprayed using positive polarity (Bozkurt et al., 2019).

[0209] Stability and cell viability of microencapsulated probiotics during storage

[0210] like Fig.17 As shown, compared with the probiotics encapsulated with positive polarity (0.44 logarithmic loss), the probiotics encapsulated with negative polarity showed significantly higher viability (about 7.3 times higher, 0.06 logarithmic loss) even after 3 days of storage. Even after 7 and 14 days of stability study, similar viability trends were monitored (about 2.4 times and 1.5 times higher), respectively. After storage for 3, 7 and 14 days, non-encapsulated BIFIDO had 29.5 times, 9 times and 5.5 times lower viability than cells encapsulated with negative polarity. In addition, it should be noted that the encapsulation efficiency of microcapsules of both polarities is similar, both of which are about 92% (92.9% ± 2.1 and 92.4% ± 1.2 for electrospraying with positive and negative polarity, respectively).

[0211] Thus, encapsulated maltodextrin-BIFIDO using negative polarity electrospray showed higher viability when stored up to about 14 days compared to cells using positive polarity electrospray. This is due to the "self-organization" of cells in the capsule core using negative polarity electrospray and the lower water content (and water activity) of these capsules, which of course provides better stability and survival conditions for encapsulated cells. Therefore, the stability and viability of encapsulated probiotics can be effectively controlled using the polarity of the external electric field.

[0212] in conclusion

[0213] In this study, the encapsulation of the Gram-positive bacteria Bifidobacterium lactis (DSM 15954) in maltodextrin capsules using negatively and positively charged electrospray needles was investigated. The charge polarity of the needle of the DC electric field directs the surface-charged probiotic cells to the vicinity of the core or outer surface of the capsule, as shown in confocal microscopy images. This is a novel "self-organization" process that results in different "compositions" being formed on the core or outer surface of the microcapsules during the electrospray process. This can be further developed to develop electrospray capsules with customized core and outer surface compositions based on the interactions between the applied electric field and the charged molecules in the dispersion. The charge polarity applied to the electrospray needle not only controls the "self-organization" of the probiotic cells in the liquid jet, but also controls the mass transfer, solvent evaporation, and physicochemical properties (e.g., glass transition) of the capsules.

[0214] Probiotics encapsulated using negative polarity showed significantly higher glass transition values ​​(close to 10°C) compared to capsules produced using positive polarity, and showed much higher viability (about 7.3 times higher) compared to probiotics encapsulated using positive polarity even after storage at 30% RH and 25°C for 3 days.

[0215] When a low conductivity or neutral maltodextrin matrix is ​​used, the charge density on the bacterial surface is sufficient for its "self-organization". Biopolymers (i.e., polyelectrolytes) with high charge density will change the charge density of the solution, which can further functionalize the self-organization and distribution of the cells. The polarity of the electric field can also be used to change the molecular interactions between polyelectrolytes and between polyelectrolytes and charged probiotic cells. Therefore, it can be advantageous to use polyelectrolytes with the same charge as the cells, or with a different charge or a neutral charge compared to the cells. Increasing the conductivity of the solution by adding salts that increase the surface charge density can also be used to adjust the encapsulation of the cells. In addition, selecting probiotic cells with a sufficiently strong surface charge (at a pH that is not close to its isoelectric point) will further promote the selective positioning of the encapsulated cells.

[0216] Furthermore, this study essentially evaluated how to improve the protection and viability of live probiotic cells encapsulated via electrospray. The viability of the encapsulated cells was significantly improved when the cells were located in the core of the capsule compared to when the probiotics were distributed near the outer surface of the capsule.

Claims

1. A method for electrostatic spray drying of live microorganisms, the method 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) drying the droplets to form dry particles; as well as e) collecting the dried particles; Wherein, the electrostatic charge has the same polarity as the surface charge of the microorganism.

2. The method of claim 1, wherein the electrostatic charge is applied to the suspension while the suspension is moved.

3. The method according to any one of claims 1 or 2, wherein the collection of the dry particles is achieved using a collector having an opposite charge compared to the electrostatic charge and the surface charge of the microorganisms.

4. The method according to any one of the preceding claims, wherein the microorganism having a negative surface charge is selected from the group consisting of Bifidobacterium animalis, Lactobacillus acidophilus, Pseudomonas aeruginosa and Escherichia coli.

5. The method according to claim 4, wherein the microorganism is Bifidobacterium animalis subsp. lactis (DSM 15954).

6. The method according to any one of the preceding claims, wherein the dry particles are in the shape of spheres, fibers, spirals, rods or mixed shapes.

7. The method according to any one of the preceding claims, wherein the formulation auxiliary is a polysaccharide, a protein, a lipid or a synthetic polymer, or a mixture thereof.

8. The method of claim 7, wherein the polysaccharide is positively charged, negatively charged or neutral.

9. The method of claim 8, wherein the polysaccharide is selected from the group consisting of maltodextrin, starch, xanthan gum, gellan gum, alginate, pectin, dextran and chitosan.

10. The method of any one of claims 7-9, wherein the protein is a dairy protein, a non-dairy animal protein, a plant protein, an algal protein, or a fermentation product protein.

11. Use of a positively or negatively charged high voltage power source to encapsulate living microorganisms having a surface charge which is the same as the charge of the high voltage power source.

12. A particle comprising a polymer and at least one cell of a living microorganism, wherein the at least one cell is located in the core of the particle, and wherein the polymer composition of the particle is uniform and forms a substantially cell-free shell around the core.

13. A particle comprising living microorganisms, the particle being obtainable by the method according to any one of claims 1 to 10.

14. Use of the particles according to any one of claims 12 or 13 for the manufacture of food, feed products, dietary supplements or medicines.

15. A food, feed product, dietary supplement or pharmaceutical product comprising the particles according to any one of claims 12 or 13.

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