Microbe delivery composite, methods and uses thereof

Abstract

The present disclosure provides a composite includes biocompatible particles includes: a biocompatible polymer, one or more microbes and a growth media for the microbes, and a mucoadhesive hydrogel, where the particles are embedded in the hydrogel.

Description

MICROBE DELIVERY COMPOSITE, METHODS AND USES THEREOFCROSS REFERENCE TO RELATED APPLICATIONS

[0001] This application claims the benefit of priority of U.S. Provisional Patent Application No. 63 / 564,245 filed March 12, 2024; and of U.S. Provisional Patent Application No. 63 / 724,544 filed November 25, 2024, which are hereby incorporated by reference.FIELD

[0002] This specification relates to composites for delivery of one or more microbes to a mucosal membrane.BACKGROUND

[0003] The following paragraphs are not an admission that anything discussed in them is prior art or part of the knowledge of persons skilled in the art.

[0004] A. Toiviainen and others disclose lozenges with Lactobacillus rhamnosus GG and Bifidobacterium animalis subsp. lactis BB-12 in Clin Oral Invest (2015) 19:77-83. The 1-g lozenges were compressed from 50% xylitol and 46% sorbitol. Each probiotic lozenge contained LGG 4.4* 108and BB-12 4.8* 108.

[0005] A. Ranjith and others disclosed a mouth rinse in Int J Dent Hygiene (2022) 20:415-421. The mouth rinse was made by dissolving 1 gram of powder of 1.25 billion freeze-dried combination of a mixture of L. acidophilus, L. rhamnosus, B. longum and S. boulardii in 20 mL of water.

[0006] A. Aysha Jebin and others disclosed chewable tablets containing 0.5 billion CFU of Lactobacillus reuteri UBLRu-87 in Contemp Clin Dent (2021) 12: 121-7.

[0007] S. Saha and others disclosed carboxymethyl cellulose (CMC) films that incorporate Lactobacillus fermentum NCIMB 5221 (6.75 x 108cells / film) in Expert Opin. Drug Deliv. (2013) 10(11): 1471-1482. The films were prepared by dissolving CMC in water, incorporating an overnight bacterial culture at 0.1 g of bacterial pellet / mLof CMC solution, homogenizing the mixture, and allowing a film of the mixture to dry for 12 hours at 25°C.

[0008] A. Abruzzo and others disclosed in Pharmaceutics (2020) 12:241 hydroxypropylmethyl-cellulose-based films prepared with a casting method that loaded L. brevis CD2 on one side of the film using a capsule-based dry powder dispenser to dispense a pure lyophilized form of the bacteria.INTRODUCTION

[0009] The following introduction is intended to introduce the reader to this specification but not to define any invention. One or more inventions may reside in a combination or sub-combination of the apparatus elements or method steps described below or in other parts of this document. The inventors do not waive or disclaim their rights to any invention or inventions disclosed in this specification merely by not describing such other invention or inventions in the claims.

[0010] Delivery of microbes to a mucosal membrane may be beneficial in the treatment or prevention of a health disorder, such as a periodontal condition (for example dental carries) or a gynaecological disease (for example bacterial vaginosis).

[0011] In one aspect, the present disclosure provides a composite that includes: (1) biocompatible particles that include: a biocompatible polymer, one or more microbes, and a growth media for the microbes; and (2) a mucoadhesive hydrogel, where the particles are embedded in the hydrogel.

[0012] In another aspect, the present disclosure provides a mixture of biocompatible particles where the mixture includes: a first portion of biocompatible particles, and a second portion of biocompatible particles. The first and second portions of biocompatible particles each include at least one microbe, and a growth media. The growth media in the two portions of particles are different.

[0013] In yet another aspect, the present disclosure provides a biocompatible particle that includes (1) one or more microbes in the form of a biofilm and (2) a growth media for the microbes.

[0014] In a still further aspect, the present disclosure provides a combination that includes: (1) particles that include: one or more microbes and a growth media for the microbes; (2) a polymer solution; and (3) a crosslinker; where the polymer solution and the crosslinker form a mucoadhesive hydrogel when mixed.

[0015] In still another aspect, the present disclosure provides a method of microbial transplantation in a subject. The method includes applying a composite according to the present disclosure to a mucosa of the subject, such as oral, nasal, or vaginal mucosa.

[0016] In still another aspect, the present disclosure provides a method of making a composite according to the present disclosure. The method may include (A) providing biocompatible particles, providing a mucoadhesive hydrogel, and mixing the particles with the hydrogel; or (B) providing biocompatible particles, mixing the particles with a polymer solution to form a particle-polymer mixture, and mixing a crosslinker with the particle-polymer mixture to form a mucoadhesive hydrogel. In either exemplary method, the particles include a biocompatible polymer, one or more microbes, and a growth media for the microbes.BRIEF DESCRIPTION OF THE DRAWINGS

[0017] FIG. 1 illustrates a flow channel design for determining mucoadhesive strength of a hydrogel according to the present disclosure.

[0018] FIG. 2A illustrates a flow chamber design for determining mucoadhesive strength of a hydrogel according to the present disclosure.

[0019] FIG. 2B is a photo showing the constructed flow chamber illustrated in FIG. 2A.

[0020] FIG. 3 illustrates a schematic diagram of an experimental setup for measuring hydrogel attachment time.

[0021] FIG. 4 is a graph of a COMOL simulation of the flow chamber design shown in FIG. 2A with the maximum velocity as a function of the x-axis (length).

[0022] FIG. 5 is a graph of a COMOL simulation of the flow chamber design shown in FIG. 1 with the shear stress as a function of the y-axis (width).

[0023] FIG. 6 illustrates contour surface maps of the maximum velocity at the center plane, and shear stress at the bottom of the channel.

[0024] FIG. 7 illustrates an updated flow chamber design for determining mucoadhesive strength of a hydrogel according to the present disclosure.

[0025] FIG. 8 A is a scanning electron microscopy image (lOOOx magnification) of unseeded agar particles.

[0026] FIG. 8B is a scanning electron microscopy image (lOOOx magnification) of agar particles according to the present disclosure seeded with L. Brevis.

[0027] FIG. 8C is a scanning electron microscopy image (lOOOx magnification) of agar particles according to the present disclosure seeded with S. Salivarius.

[0028] FIG. 9 A is a scanning electron microscopy image (lOOOx magnification) of unseeded agar particles prepared using mashed agar.

[0029] FIG. 9B is a scanning electron microscopy image (lOOOx magnification) of agar particles according to the present disclosure prepared using mashed agar and seeded with L. Brevis.

[0030] FIG. 9C is a scanning electron microscopy image (lOOOx magnification) of agar particles according to the present disclosure prepared using mashed agar and seeded with S. Salivarius.

[0031] FIG. 10A is an image of an agar microsphere according to the present disclosure at time = 0.

[0032] FIG. 10B is an image of the agar microsphere of FIG. 10A at time = 24 hours.

[0033] FIG. 11 is a graph illustrating the polydispersity of agar microspheres according to the present disclosure.

[0034] FIG. 12A is a graph illustrating biofilm cultivation of Sepharose beads according to the present disclosure.

[0035] FIG. 12B is a graph illustrating biofilm cultivation of agar microspheres according to the present disclosure.

[0036] FIG. 13 is a graph illustrating biofilm cultivation of mashed agar and oil- emulsified microspheres according to the present disclosure.

[0037] FIG. 14A is a graph illustrating S. salivarius biofilm attachment to agar microspheres according to the present disclosure.

[0038] FIG. 14B is a graph illustrating L. brevis biofilm attachment to agar microspheres according to the present disclosure.

[0039] FIG. 15 is a graph illustrating results of a time sweep experiment to determine a suitable incubation period for biofilm formation in both S. salivarius and L. brevis.

[0040] FIG. 16 is a graph illustrating the bioadhesion force of alginate, CMC (90 kDa) and CMC (250 kDa) calculated using rheological synergism.

[0041] FIG. 17A is a graph illustrating results of time sweep experiments of 2% alginate with 0% CMC in formulations according to the present disclosure.

[0042] FIG. 17B is a graph illustrating results of time sweep experiments of 2% alginate with 0.5% CMC in formulations according to the present disclosure.

[0043] FIG. 17C is a graph illustrating results of time sweep experiments of 2% alginate with 0.5% CMC in formulations according to the present disclosure.

[0044] FIG. 17D is a graph illustrating results of time sweep experiments of 2% alginate with 1% CMC in formulations according to the present disclosure.

[0045] FIG. 17E is a graph illustrating results of time sweep experiments of 2% alginate with 1% CMC in formulations according to the present disclosure.

[0046] FIG. 18A is a graph illustrating results of strain sweep experiments of 2% alginate with 0% CMC in formulations according to the present disclosure.

[0047] FIG. 18B is a graph illustrating results of strain sweep experiments of 2% alginate with 0.5% CMC in formulations according to the present disclosure.

[0048] FIG. 18C is a graph illustrating results of strain sweep experiments of 2% alginate with 0.5% CMC in formulations according to the present disclosure.

[0049] FIG. 18D is a graph illustrating results of strain sweep experiments of 2% alginate with 1% CMC in formulations according to the present disclosure.

[0050] FIG. 18E is a graph illustrating results of strain sweep experiments of 2% alginate with 1% CMC in formulations according to the present disclosure.

[0051] FIG. 19A is a graph illustrating results of frequency sweep experiments of 2% alginate with 0% CMC in formulations according to the present disclosure.

[0052] FIG. 19B is a graph illustrating results of frequency sweep experiments of 2% alginate with 0.5% CMC in formulations according to the present disclosure.

[0053] FIG. 19C is a graph illustrating results of frequency sweep experiments of 2% alginate with 0.5% CMC in formulations according to the present disclosure.

[0054] FIG. 19D is a graph illustrating results of frequency sweep experiments of 2% alginate with 1% CMC in formulations according to the present disclosure.

[0055] FIG. 19E is a graph illustrating results of frequency sweep experiments of 2% alginate with 1% CMC in formulations according to the present disclosure.

[0056] FIG. 20 is a graph illustrating bioadhesion force of formulations according to the present disclosure.

[0057] FIG. 21 A is a graph showing the Fourier transform infrared spectroscopy (FTIR) spectra of polymer solutions alone and polymer solutions combined with mucins, which illustrate hydrogel interactions.

[0058] FIG. 2 IB is a graph showing FTIR spectra of polymer solutions combined (Alg- CMC) with and without mucins and crosslinked hydrogel with and without mucins.

[0059] FIG. 22A is a graph illustrating the velocity profile across the channel of the chamber depicted in FIG. 2A.

[0060] FIG. 22B is a graph illustrating the viscous force profile over the tissue slot at the bottom of the flow chamber depicted in FIG. 2A.

[0061] FIG. 23 is a graph illustrating surface contour analysis of flow properties through the flow channel depicted in FIG. 1 using COMSOL Multiphysics.

[0062] FIG. 24 is a graph illustrating surface contour analysis of flow properties through the flow channel depicted in FIG.1 using COMSOL Multiphysics.

[0063] FIG. 25 is a graph illustrating attachment times to porcine esophagus mucosa under constant shear in the flow chamber depicted in FIG. 1 for hydrogel formulations according to the present disclosure.

[0064] FIG. 26A is a graph illustrating degradation in phosphate buffered saline (PBS) of 2% alginate with 0% CMC in formulations according to the present disclosure.

[0065] FIG. 26B is a graph illustrating degradation in phosphate buffered saline (PBS) of 2% alginate with 0.5% CMC in formulations according to the present disclosure.

[0066] FIG. 26C is a graph illustrating degradation in phosphate buffered saline (PBS) of 2% alginate with 1% CMC in formulations according to the present disclosure.

[0067] FIG. 27A is a graph illustrating degradation profiles of hydrogels according to the present disclosure in simulated saliva.

[0068] FIG. 27B is a graph illustrating degradation profiles of hydrogels according to the present disclosure in simulated saliva.

[0069] FIG. 27C is a graph illustrating degradation profiles of hydrogels according to the present disclosure in simulated saliva.

[0070] FIG. 27D is a graph illustrating degradation profiles of hydrogels according to the present disclosure in simulated saliva.

[0071] FIG. 27E is a graph illustrating degradation profiles of hydrogels according to the present disclosure in simulated saliva.

[0072] FIG. 28A is a graph illustrating microbial survival of L. brevis seeded on hydrogels according to the present disclosure.

[0073] FIG. 28B is a graph illustrating microbial survival of S. salivarius seeded on hydrogels according to the present disclosure.

[0074] FIG. 28C is a graph illustrating microbial release of L. brevis seeded on hydrogels according to the present disclosure into artificial saliva.

[0075] FIG. 28D is a graph illustrating microbial release of S. salivarius seeded on hydrogels according to the present disclosure into artificial saliva.

[0076] FIG. 29A is a graph illustrating the concentration of viable monocultures of S. salivarius and of L. brevis in a hydrogel according to the present disclosure over 24 hours.

[0077] FIG. 29B is a graph illustrating the concentration of viable co-cultures of S. salivarius and L. brevis in a hydrogel according to the present disclosure over 24 hours.DETAILED DESCRIPTION

[0078] In one aspect, the present disclosure provides a composite that includes biocompatible particles and a mucoadhesive hydrogel. The particles include: a biocompatible polymer, one or more microbes, and a growth media for the microbes. The particles are embedded in the hydrogel.

[0079] The biocompatible polymer or polymers of a particle and the mucoadhesive hydrogel are selected to have little to no adverse effects (such as little to no bactericidal or bacteriostatic effects) towards the one or more microbes present in the particle.

[0080] In some examples, at least some of the particles include a plurality of different microbes. For example, the particles could include Lactobacillus acidophilus, Lactobacillus gensenii, Lactobacillus gasseri, Lactobacillus crispatus, and a growth media such as De Man-Rogosa-Sharpe (MRS) media.

[0081] In some examples, the particles may be a mixture of different particles. For example, the particles may include a first portion of particles with a first growth media, and a second portion of particles with a second growth media that is different from the first growth media.

[0082] The microbes may be: in the form of a biofilm, such as in the form of particles of a biofilm, and / or in a planktonic state. The microbes may be: embedded in the particles, and / or on an outer surface of the particles. In the context of the present disclosure, the term "biofilm" should be understood to refer to an assemblage of microbial cells enclosed in an extracellular polymeric matrix. In the context of the present disclosure, the term "planktonic state" should be understood to refer to microbes that free-living, nonadherent microbes. For example, the particles may include: (1) microbes in the form of abiofilm that are embedded in the particles, (2) microbes in a planktonic state that are embedded in the biofilm, (3) microbes in the form a biofilm that are on an outer surface of the particles, (4) microbes in a planktonic state that are on an outer surface of the particles, (5) any combination thereof.

[0083] The biofilm or planktonic state of a microbe can be determined by differential gene expression profiling. The identification of genes that are differentially expressed in biofilm compared with planktonic cells may be achieved as discussed by M. Shemesh et al in "Differential gene expression profiling of Streptococcus mutans cultured under biofilm and planktonic conditions" Microbiology (2007) 153:5, pp 1307-1317 (https: / / doi.Org / 10.1099 / mic.0.2006 / 002030-0), which is incorporated herein by reference. The identified differential gene expression may then be used to determine whether a microbe that is part of a particle according to the present disclosure is in the form of a biofilm or in a planktonic state.

[0084] In some examples, the particles may include (i) a first portion of particles that include a microbe in the form of a biofilm and a first growth media, (ii) a second portion of particles that include a microbe in a planktonic state and a second growth media that is different from the first growth media. The microbes in the first portion of particles may be the same species as the microbes in the second portion of particles.

[0085] The biocompatible polymer may include a gel-forming polymer, such as agar or gelatin.

[0086] The particles may be smaller than about 1000 pm, for example the particles may have an average diameter less than about 500 pm, such as an average diameter that is from about 100 pm to 200 pm.

[0087] The mucoadhesive hydrogel may include alginate, and optionally carboxymethyl cellulose (CMC). The CMC may have a molecular weight from about 90 kDa to about 700 kDa, such as from about 90 kDa to about 250 kDa, preferably about 250 kDa. The alginate may have a molecular weight from about 10 kDa to about 600 kDa, such as from about 12 kDa to about 40 kDa. The alginate and the CMC may be present in a wt ratio from about 8: 1 to about 1 : 1 (alginate:CMC). The alginate may be present in the hydrogelat a concentration from about 0.5 wt% to about 5 wt%, and / or the CMC may be present in the hydrogel at a concentration from about 0.1 wt% to about 2 wt%. The hydrogel may further include a crosslinker, such as a divalent cationic crosslinker, for example Ca2+, preferably calcium chloride at a concentration of from about 0.5 mg / mL to about 4 mg / mL.

[0088] The hydrogel may have a mucoadhesive strength sufficient to withstand a fluid shear stress of up to 0.01, 0.05, 0.1, 0.2, 0.4, 0.6, 0.8, or 1.0 dyne / cm2. Hydrogels used on oral mucosal surfaces may require a mucoadhesive strength sufficient to withstand a fluid sheer stress up to 0.1 dyne / cm2, while hydrogels used on gut mucosal surfaces may require a mucoadhesive strength sufficient to withstand a fluid shear stress of up to 0.8 dyne / cm2.

[0089] The microbes may be a bacteria, such as Lactobacillus, for example Lactobacillus brevis, Lactobacillus reuteri UBLRu-87 , Lactobacillus acidophilus, Lactobacillus rhamnosus, Lactobacillus gensenii, Lactobacillus gasseri, Lactobacillus crispatus, or Lactobacillus fermentum, Bifidobacterium, for example Bifidobacterium longum o Bifidobacterium animalis subsp. lactis BB-12, Streptococcus, for example Streptococcus Salivarius, or any combination thereof. The microbes may be a yeast, such as Saccharomyces, for example Saccharomyces boulardii. The microbes may be a fungus, such as Candida albicans. The microbes in the composite may be any combination thereof, such as a combination of different species of bacteria, or a combination of bacteria and yeast.

[0090] When a combination of microbes can be grown on the same growth media, the mixture of the different microbes may be provided on the same particles. This may be illustrated as microbes A to D being provided on a particle with growth media #1 (particle "ABCDE1"). This may be alternatively illustrated as microbes A to D being provided on separate particles for each of the different microbes (for example: particles “Al”, “Bl”, “Cl”, “DI” and "El"). When the different microbes require different growth media, the mixture of different microbes may be provided on separate particles. This may be illustrated as particles “Al”, “B2”, “G3”, “H4” and “15”. The expression “differentmicrobes” may include microbes of the same species but different growth states that require different growth media, illustrated as “Bl” and “B2”. It should be understood that other combinations are also contemplated herein, such as a mixture that includes particles "ABC1", “DI” , “B2” and “15”.

[0091] In specific examples, the particles include: a first portion of particles includes Lactobacillus brevis and De Man-Rogosa-Sharpe (MRS) medium, a second portion of particles includes Streptococcus Salivarius and brain heart infusion (BHI) medium, and a third portion of particles includes Saccharomyces cerevisiae and BactotmPeptone and BactotmYeast Extract. In other specific examples, the particles include a first portion of particles with Lactobacillus acidophilus and MRS medium, a second portion of particles with Lactobacillus gensenii and MRS medium, a third portion of particles with Lactobacillus gasseri and MRS medium, and a fourth portion of particles with Lactobacillus crispatus and MRS medium. In yet other specific examples, the particles include a mixture of Lactobacillus acidophilus, Lactobacillus gensenii, Lactobacillus gasseri, and Lactobacillus crispatus, and MRS medium.

[0092] It should be understood that the microbes, particles, polymers, crosslinking solutions, hydrogels, and other features discussed above can be used in any of the aspects discussed below when such subject matter does not conflict with the requirements outlined by that specific aspect. For example, the discussion above with respect to microbes being in a planktonic state would not apply to an aspect discussed below that required the microbe to be in the form of a biofilm.

[0093] In another aspect, the present disclosure provides a mixture of biocompatible particles where the mixture includes: a first portion of biocompatible particles, and a second portion of biocompatible particles. The first and second portions of biocompatible particles each include at least one microbe, and a growth media. The growth media in the two portions of particles are different.

[0094] The first portion of the particles may include a first microbe, and the second portion of the biocompatible particles may include a second microbe that is different from the first microbe.

[0095] As discussed above, the microbes may be in the form of a biofilm, such as in the form of particles of a biofilm, and / or in a planktonic state; and the microbes may be embedded in the particles, and / or the microbes may be on an outer surface of the particles.

[0096] In still another aspect, the present disclosure provides a biocompatible particle that includes one or more microbes in the form of a biofilm, such as in the form of particles of a biofilm, and a growth media for the microbes. The microbes may be embedded in the particles, and / or the microbes may be on an outer surface of the particles.

[0097] In yet another aspect, the present disclosure provides a combination that includes: (1) particles, (2) a polymer solution, and (3) a crosslinker. The particles include one or more microbes and a growth media for the microbes. The polymer solution and the crosslinker form a mucoadhesive hydrogel when mixed. The crosslinker may be a divalent cationic crosslinker, for example Ca2+. In particular examples, such as when the polymer solution includes alginate and optionally CMC, the crosslinker may be calcium chloride at a concentration of from about 0.5 mg / mL to about 4 mg / mL. In the combination, the particles and the polymer solution may be in a mixture. Alternatively, in the combination, the particles, the polymer solution, and the crosslinker may be separate and combining the components results in a composite discussed herein.

[0098] In a still further aspect, the present disclosure provides a method of microbial transplantation in a subject. The method includes applying a composite according to the present disclosure to a mucosa of the subject, such as oral, nasal, or vaginal mucosa.

[0099] In still another aspect, the present disclosure provides a method of making a composite according to the present disclosure. The method may include providing biocompatible particles, providing a mucoadhesive hydrogel, and mixing the particles with the hydrogel. Alternatively, the method may include providing biocompatible particles, mixing the particles with a polymer solution to form a particle-polymer mixture, and mixing a crosslinker with the particle-polymer mixture to form amucoadhesive hydrogel. In either exemplary method, the particles include a biocompatible polymer, one or more microbes, and a growth media for the microbes.

[0100] Providing the particles may include forming the particles from a biocompatible particle-forming material, and inoculating the particles with the microbes. Alternatively, providing the particles may include mixing the microbes with a biocompatible particleforming material, and forming the particles from the mixture of particle-forming material and microbes.

[0101] For a composite that includes more than one portion of particles, the different portions of particles may be made using different methods. For example, one portion of particles may be made in a process that includes forming the particles from a particleforming material and inoculating the particles with the microbes; and another portion of particles may be made in a process that includes mixing the microbes with a biocompatible particle-forming material, and forming the particles from the mixture of particle-forming material and microbes. The two portions of particles may, for example, be mixed with the hydrogel. The two portions of particles may be mixed with the hydrogel sequentially or at the same time.

[0102] The particle-forming material may include a gel-forming polymer that has little to no adverse effects (such as little to no bactericidal or bacteriostatic effects) towards the one or more microbes, such as agar or gelatin. Forming the particles may include drop casting or microfluidic particle generation. Drop casting may include pouring a warm 4% agar solution into cold canola oil using a single needle, or a concentric needle assembly with agar as core flow and oil as sheath flow fluid, to form an emulsion of agar particles, and removing the oil from the particles.

[0103] Surface inoculation of the particles with the microbes may include incubating the particles with a bacterial culture for a period of time, such as for about 12, 24 or 48 hours, optionally while shaking the mixture of particles and bacterial culture. The particles may be seeded in a bacterial culture having an optical density (OD) of about 0.05 OD to about 0.4 OD. For surface seeded biofilms, inoculated particles may beincubated for an additional 12 to 48 hours post seeding in either static or shaking condition.

[0104] Planktonic microbes may also be embedded into the particles by mixing the planktonic microbes with liquid agar, followed by fabrication of the particles as described above. Seeding dosage may be from about 0.05 OD to about 1 OD of microbe suspension before mixing with the agar.

[0105] For embedded biofilm seeding, established biofilm on tissue culture polytene Petri dishes may be harvested by physical scaping and light vortexing to produce a suspension of biofilm fragments. These may be mixed with liquid agar followed by fabrication of the particles as described above. Seeding dosage may be defined by biofilm area per unit volume of agar solution.

[0106] Providing a mucoadhesive hydrogel may include mixing a polymer solution with a crosslinking solution to form the mucoadhesive hydrogel. The crosslinking solution may be a divalent cationic crosslinker, such as calcium chloride. The crosslinking solution may preferably include CaCh at a concentration from about 1 mg / mL to about 8 mg / mL.

[0107] The polymer solution may include alginate, and optionally carboxymethyl cellulose (CMC). The alginate may be present in the polymer solution at a concentration from about 1 wt% to about 10 wt%, and / or the carboxymethyl cellulose may be present in the hydrogel at a concentration from about 0.2 wt% to about 4 wt%.

[0108] Materials and Methods

[0109] Sepharose microsphere preparation. Sepharose 6B beads (Sigma Aldrich 6B100), supplied suspended in ethanol, were prepared by pipetting the slurry into a 15 mL Falcon tube. The tube was centrifuged at 600 x g for 5 minutes to separate the ethanol from the beads. After centrifugation, the ethanol supernatant was carefully removed, and the beads were resuspended in sterile water. The suspension was centrifuged again at 600 x g for 5 minutes, and the process of resuspending the beads in sterile water and centrifugation was repeated a total of five times to ensure the completeremoval of ethanol. After the final rinse, the beads were resuspended in sterile water at half the volume of the beads to create a concentrated slurry.

[0110] Agar microsphere preparation. Agar solutions were prepared by dissolving agar powder (Sigma- Aldrich A 1296) in Brain Heart Infusion (BHI) broth (Millipore 110493) and Difco Lactobacilli deMan Rogosa Sharpe (MRS) broth (Fisher Scientific 288130) at concentrations of 2% (w / v), 4% (w / v), and 6% (w / v). The solutions were then autoclaved to ensure sterilization. To create an emulsion, 150 mL of a cold non-polar solution was placed in a 250 mL beaker and stirred using a magnetic stirrer at maximum speed. The non-polar solutions used included dichloromethane (DCM) with and without 2% (v / v) Tween 80, mineral oil, and canola oil (No Name 100% Pure Canola Oil).[OHl] While continuously stirring, 50 mL of the hot agar solution was slowly added to the non-polar solution to form an emulsion. The emulsion was then transferred into four 50 mL Falcon tubes. The tubes were centrifuged at 894 x g for 5 minutes to separate the agar microspheres from the non-polar solution. After centrifugation, the supernatant was carefully removed. The agar microspheres were rinsed with sterile water and poured over a 500-micron sieve placed above a 600 mL beaker to reduce the polydispersity of the microspheres. The microspheres collected in the beaker were then transferred back into four new 50 mL Falcon tubes, with additional sterile water to rinse. The tubes were centrifuged again at 894 x g for 5 minutes, and the supernatant was discarded. The contents of all four tubes were combined into a single 50 mL Falcon tube, and the centrifugation and rinsing process was repeated three more times to ensure thorough cleaning of the agar microspheres. After the final centrifugation, the supernatant was removed, resulting in approximately 5 mL of agar microspheres. The final volume was adjusted to 7.5 mL with sterile water to create a concentrated microsphere slurry.

[0112] Measuring particle size distribution of agar microspheres. To determine the particle size distribution (PSD) of the agar microspheres, a Malvern Mastersizer 3000 laser diffraction particle size analyzer was utilized following the manufacturer's guidelines. Particles, sieved to less than 500 pm, were suspended in distilled water to achieve an obscuration value between 1-3% for the suspension. Blue (1 = 470 nm) andred (1 = 632.8 nm) lasers were employed to measure the bead suspension. Each sample was measured five times, and the PSD data were reported as the mean diameters Dx90, Dx50, and DxlO.

[0113] Biofilm preparation on microspheres.

[0114] Bacteria strains and culture conditions. Streptococcus salivarius strain Ml 8 (5. salivarius) and Levilactobacillus brevis strain Bbl 4 (Z. brevis), were purchased form commercially (ATCC) and used to assess the viability of the proposed delivery platform. S. salivarius (ATCC 14869) was originally isolated from a human oral cavity. Frozen stocks of this bacteria were stored in BHI broth containing 25% (v / v) glycerol stored at - 80 C. L. brevis (ATCC 14869) frozen stocks were stored in MRS broth containing 25% (v / v) glycerol stored at -80 °C. For use of the S. salivarius in experiments, frozen stocks were streaked on 1.5% (w / v) BHI agar (Sigma- Aldrich) plates and incubated overnight at 37 °C, for 16 to 18 hours. L. brevis frozen stocks were streaked on MRS on 1.5% (w / v) MRS agar plates and incubated at 37 °C for approximately 36 to 48 hours.

[0115] An overnight culture was then established for Z. brevis and S. salivarius in MRS and BHI broth respectively; an isolated colony from each strain was obtained from a freshly streaked agar plate, no more than a week old, using a sterile loop to inoculate 5 mL of the broth of preference of each bacteria. A negative control was kept at all times for each broth; in the same incubation process an extra 5 mL tube with the broth was kept and it was inoculated with a loop without bacteria to guarantee a lack of external bacterial contamination neither in the broth nor in loops. The 5 mL broth tubes were placed in a shaking incubator at 200 rpm and 37 °C for 16 to 18 hours.

[0116] Biofilm cultivation. To cultivate biofilm on the surface of the microspheres the bacterial concentration of the starting stock solutions was first obtained by reading their optical density (ODeoo), which was read at a wavelength of 600 nm. S. salivarius culture was then diluted to a 2:5 ratio in BHI, while Z. brevis was diluted to a 3:5 ratio in MRS. Both strains were left to further incubate for 3 hours in the same conditions to ensure both strains of bacteria were in the log phase of their growth curve which was developed by a previous student in the lab. After this period, S. Salivarius and Z. Brevis werediluted to the desired ODeoo for seeding onto the agar microspheres. These dilutions were calculated using Equation 1 given the initial ODeoo reading from the diluted overnight cultures.

[0117] CiV2= C2V2(1)

[0118] To inoculate the microspheres (Sepharose and agar) 2 mL of the washed microsphere slurries were added to the overnight bacterial cultures diluted to the desired ODeoo of 0.1 (BHI microspheres were added to S. Salivarius and MRS microspheres were added to L. Brevis). The inoculated microspheres were then incubated for 24 hours in an incubator set to 37 °C.

[0119] To assess if the oil from the emulsion was impacting the surface properties of the agar microspheres and disrupting the biofilm attachment, dry agar was mashed up by hand and inoculated using the conditions discussed herein. 4 % (w / v) solutions of agar in distilled water were autoclaved for sterilization and poured into a petri dish. Once the agar solidified it was transferred into a 500-micron sieve and forced through the mesh using a syringe plunger. These agar fragments were then suspended in autoclaved water and pipetted into the diluted overnight cultures and incubated for 24 hours in an incubator set to 37 °C.

[0120] Biofilm quantification. After incubation the inoculated microspheres were centrifuged at 600 x g for 5 minutes and the supernatant was discarded. The microspheres were then resuspended with sterile water to rinse, and then recentrifuged. This rinsing process was repeated five times. 100 pl, 200 pl, or 400 pl of agar microspheres were then resuspended in 400 pl of sterile water to simulate the hydrogel. The samples were then serially diluted with PBS in a 96-well plate. Samples from monoculture conditions were diluted from 10'1to 10'8and 20 pl of each dilution was spot-plated onto BHI agar (for S. Sal) and MRS agar (for L. Brevis). BHI agar plates were incubated for 16 to 18 hours and MRS agar plates were incubated for 36 - 48 hours, both at 37°C and 5% CO2. Colonies were counted for colony-forming unit (CFU) determination. Equation 2 was used to calculate the concentration of viable bacteria within each condition. Where N is theconcentration of viable bacteria in CFU / ml, C is the number of colonies counted, V is the volume of sample plated in ml, and D is the dilution factor.

[0121] N = C / (V x D) (2)

[0122] Scanning electron microscopy. The remaining biofilm-loaded microspheres were fixed in 2.5% glutaraldehyde prepared in 0.1 M sodium cacodylate buffer for a minimum of 2 hours to stabilize their structure. Following fixation, the microspheres were rinsed three times with 0.1 M sodium cacodylate buffer for at least 10 minutes per rinse to remove residual glutaraldehyde. Secondary fixation was performed using 1% osmium tetroxide for 2 hours to enhance contrast. The microspheres were then quickly rinsed with distilled water and agitated to resuspend. Samples were pipetted onto Poly-L- Lysine coated 13 mm diameter glass coverslips and allowed to settle for 2 hours, ensuring the samples did not dry out during this period. Coverslips were gently rinsed with 0.1 M sodium cacodylate buffer and stored at 4°C.

[0123] Dehydration of the samples was carried out using a graded series of ethanol: 50% for 10 minutes, followed by two rinses in 70% ethanol for 10 minutes each, two rinses in 95% ethanol for 10 minutes each, and finally two rinses in 100% ethanol for 10 minutes each, followed by two additional rinses in 100% ethanol dried for 10 minutes each. The dehydrated samples were then dried using a critical pointer dryer (Leica EM 300). Following drying, the coverslips were attached to 13 mm diameter SEM stubs using double-sided carbon adhesive tabs. The stubs were subsequently sputter-coated with gold / palladium to a thickness of 5 nm using the Leica EM Ace 600 vacuum coater. The samples were examined using a Zeiss SIGMA 300 FESEM equipped with a secondary electron detector.

[0124] Biofilm cultivation for agar microspheres. To determine suitable conditions for the protocol described above in “Biofilm cultivation” for the agar microspheres, the cultures were tested under four conditions: low optical density (ODeoo = 0.1) with static conditions, high optical density (ODeoo = 0.5) with static conditions, low optical density (ODeoo = 0.1) with non-static conditions, and high optical density (ODeoo = 0.5) with non-static conditions. For static conditions, the 15 mL Falcon tubes containing theinoculated agar microspheres were placed upright in an incubator set to 37 °C to simulate a static environment. For non-static conditions, the 15 mL Falcon tubes were placed in a tissue culture roller rotator (New Brunswick TC-7) at 28 rpm inside an incubator set to 37 °C. These were incubated for 24 hours and assessed for biofilm formation using methods described in above in “Biofilm Quantification” and “Scanning Electron Microscopy” to determine suitable combinations of optical density and environmental condition. Based on these results, suitable conditions were selected for further evaluation. A time sweep was then conducted using the same methods of biofilm evaluation, with 200 pL samples taken at 6, 12, 24, and 48 hours to identify the suitable incubation time for biofilm formation under the identified conditions.

[0125] Hydrogel preparation. The hydrogel formulations investigated were a combination of sodium alginate (Sigma-Aldrich W201502) and either 90 kDa sodium CMC (Sigma-Aldrich 419273) or 250 kDa sodium CMC (Sigma-Aldrich 419281) crosslinked with CaCh as outlined in Table 1. Stock polymer solutions were prepared at a concentration of 6 % (w / v) sodium alginate with either 0 % (w / v) CMC, 1.5 % (w / v) CMC, or 3 % (w / v) CMC in sterile water. To sterilize the sodium alginate and CMC, the powder was placed into a 50 ml Falcon tube and the tube was placed horizontally into an ultraviolet (UV) light sterilizing chamber for 60 minutes (with the tube being rotated once every 20 minutes). Stock calcium chloride solutions were prepared by solubilizing calcium chloride in sterile water at a concentration of 40 mg / ml. For use within the hydrogels, the higher concentration stock CaCh solution was diluted with sterile water directly before use to concentrations of 3 mg / ml, 4.5 mg / ml, and 6 mg / ml. The calcium chloride solutions were sterilized using filter sterilization with a 0.22 pm filter. The polymer solutions were first diluted at a 1 : 1 ratio using sterile water to simulate the agar microsphere solution and mixed using luer lock-connected syringes. The diluted polymer solutions were then crosslinked with CaCh at a 2: 1 ratio using the same method of mixing as previously described producing the formulations in Table 1, below.

[0126] Viscoelastic characterization of hydrogels. The storage (elastic, G’) and loss (viscous, G”) moduli of the potential mucoadhesive hydrogel formulations weremeasured. As described above in “Hydrogel Preparation” the formulations consisted of 2% (w / v) alginate combined with varying concentrations of CMC (0% (w / v), 0.5% (w / v), 1% (w / v)) and CaCL crosslinker (1 mg / ml, 1.5 mg / ml, 2 mg / ml). A controlled- stress / controlled-rate rheometer (HR10, TA Instruments) with a 40 mm diameter flat plate geometry was used to measure the moduli. To conduct the measurements, the hydrogels were first prepared as outlined above, with 5 ml of each hydrogel being prepared in a syringe. For each measurement, 1.2 ml of the hydrogel was deposited onto the plate directly from the syringe. The temperature was kept constant at 20 °C. To determine the time for the hydrogel to become stable and fully crosslinked, a time sweep was conducted for 200 seconds immediately after crosslinking using 1% strain at an angular frequency of 10 rad / s. Strain sweeps were performed to determine the linear viscoelastic region (LVR) of each hydrogel. Strain sweeps were performed from 0.1% to 100% strain at an angular frequency of 10 rad / s. Frequency sweeps were then performed to determine the frequency-dependent storage modulus and loss modulus of each hydrogel. Frequency sweeps were performed from 0.1 to 100 rad / s at a strain found within the LVR of each hydrogel, which was chosen to be 1%. For both the strain and frequency sweeps, hydrogels were deposited 5 minutes after mixing to allow the hydrogels to fully crosslink. Each hydrogel formulation was tested in triplicate across three batches, resulting in nine runs per formulation.

[0127] Mucoadheisve assessment using rheological synergism. Rheological synergism, indicative of mucoadhesion, was determined by comparing the complex viscosity of the polymer-mucin mixtures with the sum of the viscosities of the individual components. An increase in complex viscosity beyond the additive values of the individual solutions was considered indicative of mucoadhesive interactions as disclosed by E. E. Hassan and J. M. Gallo in “A simple rheological method for the in vitro assessment of mucin-polymer bioadhesive bond strength,” Pharm. Res., vol. 7, no. 5, pp. 491-495, May 1990, doi: 10.1023 / a: 1015812615635. This was determined using Equation 3 where Ar|* is the increase in complex viscosity due to mucoadhesive interactions, r|*mix is the complex viscosity of mucins combined with the polymersolution, is the complex viscosity of the polymer solution alone, and complex viscosity of the mucin solution alone.W = ifmix - p + ’Zm)

[0128] Sodium alginate and CMC solutions were prepared at a concentration of 3% (w / v) by dissolving the respective polymers in distilled water. A porcine gastric mucin (Type II, Sigma-Aldrich M2378) solution was also prepared as a 3% (w / v) solution in distilled water. Initially, the complex viscosities of the 3% (w / v) alginate, 3% (w / v) CMC (90 kDa and 250 kDa), and 3% (w / v) mucin solutions were measured separately.Following this, 6% (w / v) solutions of each were prepared as previously described, and equal volumes of each polymer solution were mixed with the mucin solution using luer lock-connected syringes. The complex viscosity was measured over a frequency range of 0.1 to 100 rad / s to assess the viscoelastic properties of the solutions using a controlled- stress / controlled-rate rheometer (HR10, TA Instruments) with a 40 mm diameter flat plate geometry. The temperature was maintained at 37°C to simulate physiological conditions.

[0129] A similar approach was taken to measure the mucadhesion of the crosslinked hydrogels where the complex viscosity values of the hydrogels alone were taken from the frequency sweep described above in “Viscoelastic characterization of hydrogels” and the complex viscosity of the mucins was taken from the previous study looking at the polymers alone. To combine the mucins with the hydrogels, the hydrogels were prepared as described above in “Hydrogel preparation” but initially diluted with a 9 % (w / v) mucin solution prepared in distilled water. The complex viscosities of the hydrogel-mucin mixtures were then measured as previously described with the polymers.

[0130] Mucoadhesion assessment using Fourier transform infrared spectroscopy.Fourier Transform Infrared Spectroscopy (FTIR) analysis was used to investigate the chemical interactions between mucins and both the individual polymers and the crosslinked hydrogels as disclosed by G. H. Shin and J. T. Kim, in “Comparative Study of Chitosan and Oligochitosan Coatings on Mucoadhesion of CurcuminNanosuspensions,” Pharmaceutics, vol. 13, no. 12, Art. no. 12, Dec. 2021, doi: 10.3390 / pharmaceuticsl3122154. Polymer and mucin solutions were prepared individually as described above in “Mucoadheisve assessment using rheological synergism”. Combined polymer-mucin solutions and combined hydrogel-mucin solutions were also prepared following the protocols in “Mucoadheisve assessment using rheological synergism”. Hydrogel formulations were prepared separately as described in “Hydrogel preparation”. FTIR spectra were obtained using a Bruker Vertex 70 FTIR spectrometer equipped with a Specac Golden Gate Attenuated Total Reflectance (ATR) accessory. The ATR crystal was cleaned with a Kimwipe and ethanol before each measurement to ensure a contaminant-free surface. To account for water peaks present in the samples, a background scan was performed with a drop of distilled water placed on the ATR crystal. The background scan was conducted using the OPUS software (Bruker) with the following settings: a scanner velocity of 40 kHz, a spectral resolution of 8and data acquisition in the mid-infrared range from 4000 cm1to 500 cm After the background scan, the water was removed, and the ATR crystal was cleaned again using a Kimwipe and ethanol. For sample measurement, a drop of the sample was placed directly onto the ATR crystal. Spectra were recorded using the OPUS software under the same settings as the background scan. A total of 50 scans were performed per sample to ensure signal-to-noise ratio optimization. The collected scans were subsequently averaged to produce a final spectrum for each sample.

[0131] Mucoadhesive assessment of hydrogels using fluid shear. To assess the mucoadhesive properties of the hydrogel, a shear-based method was employed using porcine esophagus mucosa as the substrate. This approach simulates the mechanical forces encountered by mucosal tissues in physiological conditions, providing an accurate evaluation of the hydrogel’s adhesive strength as discussed by A. G. Mikos and N. A. Peppas in “Bioadhesive analysis of controlled-release systems, iv. an experimental method for testing the adhesion of microparticles with mucus,” J. Controlled Release, vol. 12, no. 1, pp. 31-37, Mar. 1990, doi: 10.1016 / 0168-3659(90)90180-2. The porcine esophagus was selected due to its structural and compositional similarities to human 1mucosal tissues, making it a suitable model for this study. The hydrogel was applied to the mucosal surface, and shear forces were systematically applied to quantify the adhesive interaction between the hydrogel and the mucosa. This method enables the determination of both the initial adhesion strength and the durability of the adhesive bond under continuous shear stress.

[0132] COMSOL simulation of fluid shear stress in flow chamber. COMSOL Multiphysics (Version 6.0) was used to analyze the flow properties through the flow channel. A 3D space dimension was selected, and the Laminar Flow physics package was applied. The geometry of the flow channel, including the entrance and exit tubing, was designed as shown in FIG. 1, with water chosen as the material for the fluid domain. The inlet boundary condition was set to the experimentally determined flow rate of the fluid, while the outlet boundary condition was set to a pressure of 0 Pa. A coarse mesh size was initially chosen and then progressively refined until no further changes in the simulation results were observed, ensuring mesh independence. A stationary study was conducted to compute the velocity profile along the x-axis. This allowed for the determination of the entry length, identifying the region where the flow became fully developed and where the tissue should be placed within the flow channel. Additionally, the shear force along the bottom wall of the channel was calculated to assess the shear stress exerted on the hydrogel. Both surface plots and ID plots were generated to visualize the velocity distribution and shear stress along the flow channel.

[0133] Flow chamber design. The flow chamber was designed in Onshape according to specific dimensional and functional requirements determined through the previously described COMSOL simulations. The final base design measured 70 mm in length and 25 mm in width. The flow channel was engineered with a rectangular cross-section, featuring a width of 7 mm and a height of 3.175 mm. To prevent fluid stagnation, the channel incorporated gradual expansions and contractions at both the entry and exit points. A 5 mm by 5 mm square slot, with a depth of 3 mm, was positioned 50 mm from the entry point of the flow channel for the placement of the tissue sample. A surrounding rectangular slot was integrated into the design to accommodate a Buna-N gasket,ensuring a leak-proof assembly. Eight screw holes with a 3-48 thread were incorporated around the channel to secure the top of the flow chamber assembly. The final CAD design of the base is shown in FIG. 2A. The base of the flow chamber was 3D printed using an Elegoo Mars Pro 2 resin printer with transparent ABS-like resin. The printed base was then paired with a 1 / 16 inch thick piece of transparent polycarbonate, which was CNC milled (Sain Smart Genmitsu 3018-MX3) to include the eight corresponding screw holes. This polycarbonate piece served as a lid, allowing visualization through a microscope during experiments. The final constructed flow chamber is shown in FIG. 2B.

[0134] Porcine esophagus tissue preparation. Before conducting experiments all animal work was approved by the Dalhousie University committee for laboratory animals (Protocol# 123-42). Fresh porcine esophagi were purchased from D'Aubin's Family Meats (Halifax, NS) and immediately transported on ice to the laboratory at Dalhousie University. Upon arrival, the esophagi were dissected to extract the mucosal lining. The mucosa was carefully spread out on a wax board and rinsed with PBS to remove any residual debris. Using a scalpel, the mucosa was cut into 5 mm by 5 mm squares. These tissue squares were then attached to 5 mm by 5 mm polycarbonate squares, each 1 / 16 inch thick using epoxy resin (Gorilla Glue Epoxy). The prepared mucosal samples were subsequently placed into the tissue slot within the flow chamber for mucoadhesion testing. After each trial, the mucosal samples were removed from the flow chamber and disposed of according to the approved ethics protocol.

[0135] Measuring hydrogel attachment time. Hydrogel formulations were prepared separately as described above in “Hydrogel preparation”. The initial dilution included 100-micron dark blue polystyrene beads suspended in distilled water, which served as an indicator of when the hydrogel fully detached from the mucosa. To assess the hydrogel attachment 20 pL of hydrogel was pipetted directly onto the mucosa surface, and the chamber lid was secured by screwing it in place. Tygon tubing (ADF02002) was attached to both sides of the chamber. One side was connected to a syringe pump (Chemyx Fusion 200) equipped with a 50 mL syringe filled with distilled water, while the other side directed the runoff into a waste beaker. The initial flow rate was set at 35 mL / min, basedon values for similar experiments disclosed by A. G. Mikos and N. A. Peppas in “Bioadhesive analysis of controlled-release systems, iv. an experimental method for testing the adhesion of microparticles with mucus,” J. Controlled Release , vol. 12, no. 1, pp. 31-37, Mar. 1990, doi: 10.1016 / 0168-3659(90)90180-2. The flow rate was incrementally increased, and the design of the flow chamber was modified iteratively to accommodate higher flow rates, as determined by COMSOL simulations. The hydrogel detachment flow rate was identified when the hydrogel was fully removed from the mucosa, which occurred at 50 mL / min. Due to the limited capacity of the syringe pump (50 mL), the experiment could only be run for 1 minute at this flow rate. To allow for longer flow durations, a custom reservoir was created using a 1000 mL sterile water container (Baxter JF7624). A hole was drilled in the bottom of the container to attach the tubing, and the height of the reservoir was calculated to generate a flow rate of 50 mL / min. This flow rate was verified by collecting runoff in a beaker after 3 minutes, and the volume was measured and averaged over 10 trials. The final flow rate was determined to be 53 mL / min. The flow was initiated by opening a tube clamp near the entrance of the chamber. The time from flow initiation to hydrogel detachment was observed using a Trinocular Stereo Zoom Microscope (3.5X - 180X, AmScope). A schematic diagram of the experimental setup up is shown in FIG. 3. Hydrogel formulations were pipetted onto the mucosa surface within the flow chamber, which was connected to a custom reservoir system using a 1000 mL sterile water container via Tygon tubing. Hydrogel detachment was observed using a Trinocular Stereo Zoom Microscope. The experiment was repeated five times for each hydrogel formulation, and the mucosal tissue was replaced for each experiment.

[0136] Hydrogel degradation. Hydrogels were prepared following the protocol described above in “Hydrogel preparation”. A 40-micron cell strainer was initially weighed to obtain its baseline weight. A volume of 400 pL of the prepared hydrogel was then deposited into the cell strainer, which was weighed again to determine the combined weight of the hydrogel and strainer. The mass of the hydrogel was calculated by subtracting the initial weight of the dry empty cell strainer from the weight of the strainerwith the hydrogel. The cell strainer containing the hydrogel was placed into a well of a 6- well plate, and 8 mL of either artificial simulated saliva (BioChemazone BZ109) or PBS was pipetted into each well to ensure complete submersion of the hydrogel within the strainer, with PBS serving as a control condition. The weight of the cell strainer containing the hydrogel was measured at time points of 0, 30 minutes, 1 hour, 2 hours, 4 hours, and 8 hours to monitor any changes in hydrogel mass due to swelling or degradation in the simulated saliva and PBS. After each measurement the simulated saliva and PBS was replaced to avoid polymer saturation. For each hydrogel formulation, three replicates were conducted.

[0137] Assessing hydrogel biocompatibility with bacteria. Bacteria-loaded hydrogels were prepared following the protocol described above in “Hydrogel preparation” in a 3 mL syringe where the first dilution consisted of 1 mL of S. Salivarius coated microspheres, 1 mL of L. Brevis coated microspheres, or 500 pL of each. A volume of 400 pL of the prepared bacteria-loaded hydrogel and was then deposited into a 1.5 mL microcentrifuge tube and left at room temperature for 24 hours. In parallel, identical volumes of microspheres were loaded into a 3 mL syringe filled with PBS and 400 pL was deposited into a 1.5 mL microcentrifuge tube and left at room temperature for 24 hours. At time points of 0 hours and 24 hours, 200 pL of the bacteria loaded hydrogel and PBS was deposited into a 96-well plate and serial diluted as outlined above in the section entitled “Biofilm quantification”. The samples containing both S. Salivarius and L. Brevis were spot-plated onto both BHI and MRS agar plates as it was determined no L. Brevis colonies formed after 24 hours on BHI plates and no colonies of S. Salivarius were seen at countable dilutions (10-100 colonies) after 48 hours on MRS plates. For each hydrogel, three replicates were conducted and the entire experiment was repeated three times using a freshly streaked plate.

[0138] Assessing microbial release in simulated saliva. Bacteria-loaded hydrogels consisting of both S. Salivarius and L. Brevis were prepared as described above in “Assessing hydrogel biocompatibility with bacteria”. In parallel, 500 pL of both S. Salivarius and L. Brevis coated microspheres were diluted with the same volume of PBS(2 mL) to act as a control. The microbial release was assessed using a similar approach to the degradation study described above in “Hydrogel degradation”. A volume of 400 pL of the prepared bacteria-loaded hydrogels and the bacteria loaded PBS was deposited directly into each cell strainer. The cell strainers were then placed into a well of a 6-well plate, and 8 mL of artificial simulated saliva was added to each well to completely submerge the hydrogels. At time points of 0, 30 minutes, 1 hour, 2 hours, 4 hours, and 8 hours, 200 pL samples of the simulated saliva were withdrawn from each well to assess microbial release from the hydrogel using spot plating as described in “Assessing hydrogel biocompatibility with bacteria”. After each time point the cell strainer was disposed of, therefore, a hydrogel sample was created for each time point. For each hydrogel, three replicates were conducted and the entire experiment was repeated three times using a freshly streaked plate.

[0139] Example 1

[0140] Different hydrogels were prepared by combining dried sodium alginate powder with dried sodium CMC powder in a beaker and allowing the mixture to solubilize in water overnight. The polymer solutions were made at double the final concentration and diluted by half on mixing with the crosslinking solution at a 1 : 1 ratio. Mixing the polymer solution and the crosslinking solution was achieved by connecting two syringes together using a luer-lock syringe couplers and pushing the mixture back and forth between the syringes. The preliminary hydrogel formulations that were selected for tested for muco-adhesion are shown Table 1.

[0141] Table 1

[0142] Physical characterization of the hydrogels was conducted using rheology to obtain the viscoelastic properties of each gel. The storage modulus, G’, and loss modulus, G” were determined using a TA HR10 rheometer. Amplitude sweeps were conducted first where the frequency was kept constant, and the strain was increased incrementally until the structure of the sample starts to disintegrate. Amplitude sweeps were used to determine the linear viscoelastic (LVE) region of each hydrogel, which is the range of strains that can be used on the sample before disintegration. With a known LVE region, time sweeps, and frequency sweeps were completed for each hydrogel to determine the storage and loss moduli. Frequency sweeps were measurements taken where the strain amplitude was kept constant, and the frequency is increased incrementally. In a timesweep, both the strain and frequency were kept constant to verify if the hydrogels had any time dependent properties. The constant strain was selected within the LVE region during the amplitude sweeps.

[0143] The results indicated that increasing CMC concentration leads to a decrease in moduli where the higher molecular weight CMC showed a larger decrease. This results in a decrease in complex viscosity and an increase of spreadability.

[0144] Table 2 shows the results of a mucoadhesion study using 3% mucins and the individual polymer components. The viscosity of the mucins (r|m), polymers (r|t) and both combined (qt) were measured and the increase in viscosity was determined to be the viscosity due to mucoadhesion (r|b). Using the following equations, which are equivalent to equation (3) discussed above but use different nomenclature,the mucoadhesive force was determined by multiplying the mucoadhesive viscosity by the frequency. The results suggest 250 kDa CMC has the strongest interactions.

[0145] Table 2

[0146] Mucoadhesive strength was measured in an in-vitro flow system using porcine esophagus tissue. Using COMSOL simulation, the flow system was simulated to determine the entrance length and the shear stress along the bottom surface of the channel based on a flowrate of 25 mL / min. The design of the flow system is illustrated in FIG. 1, where a gasket and polycarbonate sheet will be screwed on top to seal the device. FIG. 2 shows the maximum velocity along the x-axis of the channel, which shows that it a reached steady state maximum velocity of 0.056 m / s at approximately 50 mm. Therefore, the mucosal tissue will be placed 50 mm into the channel. FIG. 3 shows the viscous force (shear stress) profile along the y axis of the channel, resulting in a maxshear stress of 0.065 N / m2. Based on this profile the mucosal tissue will be placed 1 mm away from the walls on either side to reduce the hydrogel closer to the walls from being subjected to significantly lower shear than the center. The matching contour maps of the maximum velocity at the center of the channel and the shear stress at the bottom of the channel are shown in FIG. 4. Based on these results the final design was developed to provide substantially consistent flow over top of the hydrogel attached to the mucosa (FIG. 5).

[0147] Agar particles were prepared by pouring a 4 wt% agar solution into cold canola oil to create an emulsion. The particles were then spun down and washed 5 times to remove the oil. The particles were then poured over a 500-micron sieve and spun down again to remove the supernatant. The particles were then placed in an overnight culture of S. Salivarius with an OD of 0.2 or of L. Brevis with an OD of 0.15 and incubated for 24 hours. Following incubation, the particles were imaged using SEM to determine if the particles did support biofilm growth. The SEM images in FIG. 6A to FIG. 6C show that the agar particles were able to support biofilm growth as the images show bacteria plumes attached to the surface of the particle. FIG. 6A shows the SEM image of unseeded agar particles. FIG. 6B shows the SEM image of agar particles seeded with L. Brevis. FIG. 6C shows the SEM image of agar particles seeded with S. Salivarius. Each was imaged at lOOOx magnification, 5 kV electron height tension, and 11.3 mm working distance. All the samples were examined using a Zeiss SIGMA 300 FESEM equipped with a secondary electron detector.

[0148] Inoculation of microbes onto the surface of the particles was tested using different incubation conditions, including shaking vs. static, optical density, and incubation time. First the optical density was varied from high to low at both shaking and static conditions. The number of colony forming units (CFU) / mL was determined using spot plating and images were taken using SEM. The SEM images proved to be ineffective quantifying bacteria growth but useful just to visualize biofilm attachment, which each scenario demonstrated. The CFU results shown in Table 3 show that shaking vs. static incubation has the largest impact, where shaking significantly improves attachment. Theresults also show that seeding the particles at a lower optical density result in higher biofilm attachment. Based on these results, suitable conditions are shaking with 0.1 OD, and a time sweep was conducted using these conditions and incubating for 6, 12, 24, and 48 hours. Both results are similar to the growth curves of the respective bacteria. The results showed that suitable S. Salivarius biofilm attachment is between 12 and 24 hours, and suitable L. Brevis attachment is passed 24 hours.

[0149] Table 3 - CFU / mL of biofilm on 100, 200, and 400 pL of agar particles after 24 hours with varying growth conditions; CFU / mL of biofilm on 200 pL of agar particles after 6, 12, 24, and 48 hours with 0.1 initial optical density and shaking growth conditions.

[0150] The described protocol for developing particles with canola oil was repeated with mashed agar (no oil) to compare attachment and investigate if the oil inhibits biofilm attachment. FIG. 7 A to FIG. 7C compare the SEM images between the two protocols using the same incubation conditions. FIG. 7A shows the SEM image of unseeded mashed agar particles. FIG. 7B shows the SEM image of agar particles seeded with L. Brevis. FIG. 7C shows the SEM image of agar particles seeded with S. Salivarius. Each was imaged at 250x magnification, 5 kV electron height tension, and 11.3 mmworking distance. All the samples were examined using a Zeiss SIGMA 300 FESEM equipped with a secondary electron detector. The CFU / mL measurements are shown in Table 4. Both the SEM images and CFU show no significant difference in biofilm attachment, suggesting that the oil does not inhibit biofilm growth.

[0151] Table 4 - CFU / mL for 200 pL of agar particles formed using either the oil emulsion method or the mashed agar method, and seeded with either L. Brevis or S. Salivarius.

[0152] Oil emulsion agar particles seeded at OD 0.1 with S. Salivarius and / or L.Brevis were loaded into selected mucoadhesive hydrogel formulations (Gel A, E and I) according to the present disclosure. For monoculture, 200pL of beads with either S.Salivarius or L. Brevis were loaded. For co-culture, 100 pL of S. Salivarius and 100 pL of L. Brevis seeded beads were loaded into the gels. Table 5 shows the CFU / ml immediately after seeding (0 hr) and at 24 hr post-seeding.

[0153] Table 5

[0154] Agar microsphere production. The results for agar microspheres prepared using different agar concentrations in the selected non-polar solvent are discussed in the following three sections. First, the formation and stability of the agar emulsion solution were evaluated for optimal microsphere production. Followed by the percent swelling of the microspheres after 24 hours, and the polydispersity to assess size uniformity.

[0155] Emulsion fluid and agar concentration. The formation of agar microspheres was assessed using agar concentrations of 2%, 4%, and 6% in different non-polar solvents. Specific to the experimental conditions used here, the 2% agar solution failed to form distinct microspheres and instead produced a soft gelatinous mass, indicating that the concentration was too low to maintain structural integrity in the emulsion. In contrast, both 4% and 6% agar successfully formed microspheres; however, the 6% agar solidified too quickly during the emulsification process, making it difficult to work with. Based on these observations, the 4% agar concentration was chosen for further analysis due to its handling properties and successful microsphere formation. Changing the viscosity of the solvent, mixing speed of the emulsion and temperature differential between agar and solvent as well as fabrication technique, such as the use of microfluidic bead generators, can allow additional agar concentrations to form usable beads.

[0156] Agar microsphere percent swelling. The swelling behavior of the agar microspheres was assessed to determine the structural stability of the microspheres over 24 hours. Percent swelling was determined to be minimal across all three trials, with slight increases in diameter after 24 hours of incubation. In the first trial (FIG. 10A (time = 0) and 10B (time = 24 hours)), the microsphere diameter increased from 190.77 pm to 197.65 pm, resulting in a percent swelling of 3.6%. The second trial showed a diameter increase from 313.25 pm to 329.81 pm, corresponding to 5.3% swelling. Finally, the third trial exhibited the largest swelling, with the diameter increasing from 93.66 pm to 99.95 pm, resulting in a 6.5% swelling. The agar microspheres were imaged using the EVOS™ FL Auto 2 Imaging System at lOx magnification. Diameters were measured using Image J software.

[0157] Mineral oil and canola oil were both tested as the non-polar solvent, and both supported the formation of stable microspheres. Canola oil was identified as a suitable non-polar solvent since canola oil is edible and therefore safer for use in oral applications.

[0158] Agar microsphere polydispersity. The polydispersity of the agar microspheres was assessed by measuring the mean diameters to confirm a uniform particle size,consistent with the sieving process of washing the microspheres through a 500-micron sieve to achieve minimal size variation. The microspheres were measured using a Malvern Mastersizer 3000 laser diffraction particle size analyzer. The microspheres, sieved to less than 500 pm, were suspended in distilled water with an obscuration value between 1-3%. Blue (1 = 470 nm) and red (1 = 632.8 nm) lasers were used to measure the suspension. The sample was measured five times and the PSD is given as the mean diameter (size class). A uniform particle size distribution, as shown in FIG. 11, was determined by the measured diameters that ranged from 100 pm to 500 pm. Specifically, the mean diameter for Dx(10) was 101 pm, indicating that 10% of the microspheres were smaller than this size. The mean diameter for Dx(50) was 255 pm, representing the median size, while the mean diameter for Dx(90) was 500 pm, signifying that 90% of the microspheres were smaller than this size.

[0159] Biofilm cultivation. To assess the relationship between microsphere volume and bacterial load, the CFU / ml was evaluated for different volumes of microspheres (Sepharose and agar). The CFU / ml doubled with increasing microsphere volumes, showing a proportional relationship between microsphere volume and bacterial concentration as shown in FIGs. 12 A and 12B. (FIG. 12 A. Concentration of biofilm formed (CFU / mL) of S. salivarius (left axis) and Z. brevis (right axis) on 100, 200, and 400 pL of Sepharose beads. FIG. 12B. Concentration of biofilm formed (CFU / mL) of S. salivarius (left axis) and L. brevis (right axis) on agar microspheres at the same volumes. Black circles represent the value of each biological replicate, and each bar indicates the mean biofilm concentration (n = 3).)

[0160] Resuspending 100 pl, 200 pl, or 400 pl of microspheres resulted in progressively higher CFU / ml values, with the 400 pl condition yielding twice the bacterial load of the 200 pl condition, and the 200 pl condition doubling that of the 100 pl sample.Additionally, biofilm adherence was stronger on agar microspheres (FIG. 12B) compared to Sepharose beads (FIG. 12 A), as indicated by higher CFU / mL for both S. salivarius and L. brevis. Furthermore, given that this platform is designed to degrade on its own and be ingested, all components must be safe for human consumption. To ensurethe safety of the Sepharose 6B beads, Cytiva, the manufacturer, was contacted to inquire about their suitability for ingestion. In their response, Cytiva clarified that Sepharose beads are strictly intended for chromatography use and should not be ingested, as the cross-linking process involves potential carcinogens. Given this, it was decided to move forward with the agar microspheres produced using the oil emulsion method, as these offer a safer alternative for oral delivery applications.

[0161] To assess if the oil in the emulsion was impacting the surface properties of the agar microspheres and disrupting biofilm attachment, the CFU / ml was compared between 200 pL of mashed agar and oil-emulsified agar microspheres. The CFU / ml results shown in FIG. 13 (each bar indicates the mean biofilm concentration of three biological replicates (n = 3)) indicate no significant difference between the two conditions. Both mashed agar and oil-emulsified microspheres yielded similar colony counts for S. salivarius and L. brevis, confirming that the oil in the emulsion did not disrupt biofilm formation or bacterial attachment to the agar surface.

[0162] SEM imaging was performed to visually confirm the biofilm formation of S. salivarius and L. brevis and assess any differences between biofilms formed on oil emulsion microspheres and the mashed agar. Additionally, SEM was used to inspect the amount of planktonic bacteria remaining after the washing steps. The images shown in FIGs 8 and 9 confirm that biofilms of S. salivarius and L. brevis were successfully formed on both types of agar, with no discernible differences in bacterial attachment or biofilm structure. Furthermore, the minimal presence of planktonic bacteria in the background confirmed that the washing procedure was effective in removing unattached cells. Based on these biofilm quantification results and SEM images, the oil emulsion process will be utilized in future experiments.

[0163] Biofilm attachment on agar microspheres. To test biofilm attachment to the agar microspheres, various conditions were tested to identify suitable environments for biofilm formation. The previously observed correlation between increasing microsphere volume and a corresponding increase in CFU / mL was also observed in these experiments as shown in FIGs. 14A and 14B. (FIG. 14 A. Concentration of biofilm formed (CFU / mL)of S. salivarius on 100, 200, and 400 pL of agar microspheres. FIG. 14B. Concentration of biofilm formed (CFU / mL) of L. brevis on 100, 200, and 400 pL of agar microspheres at the same volumes. Black circles represent the value of each technical replicate, and each bar indicates the mean biofilm concentration (n = 1).)

[0164] Similar to the previous experiments, the CFU / mL values consistently doubled as the microsphere volumes increased from 100 pL to 200 pL and then to 400 pL. This trend held across all tested conditions, regardless of the optical density or environmental condition (static vs. dynamic), reaffirming that the bacterial load correlates with microsphere volume. In addition, the species-specific attachment results indicated that L. brevis (Figure 14B) consistently exhibited higher CFU / mL than S. salivarius (Figure 14 A) across all conditions tested.

[0165] As for the incubation conditions, the impact of dynamic conditions on biofilm attachment was more pronounced compared to static conditions. Samples incubated under dynamic conditions showed higher CFU / mL counts than their static counterparts, indicating that agitation and the microspheres being in suspension facilitated biofilm formation. These results are consistent with the formation of a distinct visual layer of biofilm across the top of the settled microspheres in static conditions. Optical density had a smaller but noticeable effect; lower optical density (ODeoo = 0.1) consistently resulted in higher CFU / mL than the high optical density (ODeoo = 0.5), though the effect was less dramatic than the difference caused by changing environmental conditions. Low optical density and dynamic incubation conditions were used for future experiments.

[0166] A time sweep experiment was then conducted to determine a suitable incubation period for biofilm formation in both S. salivarius and L. brevis (FIG. 15). Concentration of viable S. salivarius and L. brevis (CFU / mL) on 200 pL of agar microsphere after 6, 12, 24, and 48 hours using an OD of 0.1 and dynamic incubating conditions. Each bar indicates the mean biofilm concentration between three biological replicates (n = 3).

[0167] For S. salivarius, CFU / mL increased from 6 hours to 12 hours of incubation. However, the CFU / mL remained relatively stable after 24 hours, with no significant increase, and then declined after 48 hours. In contrast, L. brevis showed a steady increasein CFU / mL from 6 hours to 24 hours, after which the CFU / mL remained relatively unchanged through the 48-hour mark. Based on these trends, 24 hours was chosen as the optimal incubation time for future experiments to ensure consistency between both strains, as this time point yielded the highest CFU / mL for both L. brevis and S. salivarius.

[0168] Selection of mucoadhesive polymers based on rheological synergism.Rheological synergism was assessed by measuring the increase in complex viscosity of polymer-mucin mixtures compared to the sum of the viscosities of the individual components. FIG. 16 illustrates the bioadhesion force of each polymer calculated using rheological synergism. The bars span from the minimum to the maximum value with the line within the bars representing the average value from three independent runs (n = 3). P values were obtained using a one-way ANOVA. *p < 0.05, **p < 0.01, ***p < 0.001.

[0169] As shown in FIG. 16, the 250 kDa CMC exhibited the largest increase in complex viscosity when combined with mucins, demonstrating a statistically significant difference between both alginate and 90 kDa CMC. This suggests that 250 kDa CMC is the most mucoadhesive polymer tested. In contrast, there was no statistically significant difference between the mucoadhesion of 90 kDa CMC and alginate; however, the trend indicated that 90 kDa CMC had the smallest increase in complex viscosity, implying it was the least mucoadhesive of the three. Based on these findings, only formulations containing alginate alone (Table 1, A-C) and alginate combined with 250 kDa CMC (Table 1, G-I and M-O) will be pursued for further investigation.

[0170] Selection of mucoadhesive hydrogels based on viscoelastic characterization. The viscoelastic properties of the nine remaining hydrogel formulations, consisting of varying concentrations of CMC (0%, 0.5%, 1%) and CaCL (1 mg / ml, 1.5 mg / ml, 2 mg / ml) combined with 2% alginate, were analyzed using time sweep measurements. These measurements were conducted to assess the stability and crosslinking behavior of each formulation.

[0171] FIGs. 17A to 17E illustrate time sweeps of potential mucoadhesive hydrogels containing: (FIG. 17 A) 2% alginate (formulations A-C), (FIG. 17B) 2% alginate with0.5% CMC (formulations D-F), (FIG. 17C) 2% alginate with 0.5% CMC (formulations G-I), (FIG. 17D) 2% alginate with 1% CMC (formulations J-L), and (FIG. 17E) 2% alginate with 1% CMC (formulations M-O); each assessed for 200 seconds immediately after crosslinking using 1% strain at an angular frequency of 10 rad / s. Values are the averages from three independent hydrogel mixtures (n = 3).

[0172] As shown in FIGs. 17A to 17E, the time sweep results indicated no significant changes in the viscoelastic properties across all hydrogel formulations. Both G’ and G” remained constant throughout the 200-second period, confirming that the crosslinking process occurred instantaneously and the hydrogels stabilized immediately after preparation. This rapid stabilization allows for the hydrogels to be tested right away without the need for additional waiting time.

[0173] An amplitude sweep was conducted to determine the LVR for each of the fifteen hydrogel formulations, where the LVR represents the range of strain within which the hydrogel's G’ remains constant. Beyond the LVR, the hydrogel begins to deform, indicating structural breakdown. On a plot, the LVR appears as a flat, horizontal section where the G’ remains unchanged as strain increases, indicating elastic behavior. Once the strain surpasses the LVR, the G’ begins to deviate, marking the onset of structural deformation. FIGs 18A to 18E illustrate strain sweeps of potential mucoadhesive hydrogels containing: (FIG. 18 A) 2% alginate (formulations A-C), (FIG. 18B) 2% alginate with 0.5% CMC (formulations D-F), (FIG. 18C) 2% alginate with 0.5% CMC (formulations G-I), (FIG. 18D) 2% alginate with 1% CMC (formulations J-L), and (FIG. 18E) 2% alginate with 1% CMC (formulations M-O) subjected to strain sweeps from 0.1% to 100% strain at an angular frequency of 10 rad / s. Values are the averages from three independent hydrogel mixtures (n = 3).

[0174] As shown in FIGs 18A to 18E, the LVR of each hydrogel extended past 1% strain, therefore, a strain of 1% was selected for the frequency sweeps. Furthermore, the results indicated that increasing the CMC concentration resulted in an expanded LVR while decreasing the CaCL concentration also led to an increase in the LVR range.

[0175] A frequency sweep was conducted to evaluate the frequency-dependent behavior of the hydrogel formulations, measuring both G’ and G” across a range of angular frequencies (0.1 to 100 rad / s) at 1% strain.

[0176] FIGs 19A to 19E illustrate frequency sweeps of potential mucoadhesive hydrogels containing: (FIG. 19A) 2% alginate (formulations A-C), (FIG. 19B) 2% alginate with 0.5% CMC (formulations D-F), (FIG. 19C) 2% alginate with 0.5% CMC (formulations G-I), (FIG. 19D) 2% alginate with 1% CMC (formulations J-L) and (FIG. 19E) 2% alginate with 1% CMC (formulations M-O) subjected to frequency sweeps from 0.1 to 100 rad / s at a strain of 1%, chosen within the linear viscoelastic region (LVR) of each hydrogel. Values are the averages from three independent hydrogel mixtures (n = 3).As shown in FIGs 19A to 19E, the addition of CMC resulted in a decrease in both G’ and G” across all formulations. In contrast, increasing the concentration of the CaCh led to an increase in both G’ and G”. Furthermore, in all cases, both G’ and G” increased with increasing angular frequency. The frequency sweep results are summarized in Table 6, where G’ and G” were measured at an angular frequency of 1 rad / s. Using these values, the tan 6 was calculated for each hydrogel formulation. Where Tan 6 is the ratio of the loss modulus to the storage modulus (tan 6 = G” / G’), and it provides insight into the viscoelastic behavior of the hydrogels. A higher tan 6 indicates a more viscous or liquidlike material, while a lower tan 6 signifies a more elastic or solid-like material. This parameter is particularly important when evaluating the spreadability of a hydrogel. Hydrogels with a higher tan 6 have more viscous properties, allowing them to spread more easily, which is crucial for effective mucoadhesion and application in oral delivery systems. The results revealed that gels A and B had particularly low tan 6 values (0.148 and 0.135), which is a characteristic of a predominantly solid-like hydrogel. This was also visually observed, as both gels formed a coagulated, dense structure after preparation, further confirming their lack of spreadability. Because of this, gels A and B are excluded from further consideration, as their inability to spread effectively limits their potential for mucoadhesive applications.Table 6. Viscoelastic properties of potential mucoadhesive hydrogel formulations. Values were measured at an angular frequency of 1 rad / s and a strain of 1%. Values are the average from three independent hydrogel mixtures (n = 3).

[0177] Selection of mucoadhesive hydrogel based on rheological synergism.Rheological synergism was again used to evaluate mucoadhesive strength by comparing the complex viscosities of hydrogel-mucin mixtures to the sum of the viscosities of the individual components. As illustrated in FIG. 20, Hydrogels A, B, H, and O demonstrated positive bioadhesion force values indicated by an increase in complex viscosity after mucin addiction suggesting mucoadhesive capability; while gels C, G, I, M, and N resulted in negative bioadhesion values, meaning these gels experienced a decrease in the complex viscosity upon the addition of mucins to the hydrogels. (The bars span from the minimum to the maximum value with the line within the bars representing the average value from three independent runs (n = 3).) Based on these results there is no noticeable trend correlating the bioadhesion values with specific hydrogel formulations.

[0178] Assessing hydrogel interactions with mucins using Fourier transform infrared spectroscopy. FTIR analysis was used to investigate the chemical interactions between mucins and both the individual polymers and the crosslinked hydrogels. Spectra were obtained using a Bruker Vertex 70 FTIR spectrometer equipped with a Specac Golden Gate Attenuated Total Reflectance (ATR) accessory. Each spectrum represents the average of 50 scans recorded in the mid-infrared range from 4000 cm1to 500 cmfollowing background correction with distilled water.

[0179] The FTIR spectra of the polymer solutions alone and polymer solutions combined with mucins are shown in FIG. 21 A. The absorption bands for the functional groups of both alginate and CMC were identified. Bands at 1583 cm1and 1412 cm1were assigned to the asymmetric and symmetric COO stretching vibrations of the carboxylate groups present in both polymers as discussed by R. Pereira, A. Tojeira, D. Vaz, A. Mendes, and P. B.rtolo in “Preparation and Characterization of Films Based on Alginate and Aloe Vera,” Int. J. Polym. Anal. Charact. - vol. 16, pp. 449-464, Oct. 2011, doi: 10.1080 / 1023666X.2011.599923; and by S. Hidayat, P. Ardiaksa, N. Riveli, and I. Rahayu in “Synthesis and characterization of carboxymethyl cellulose (CMC) from salak-fruit seeds as anode binder for lithium-ion battery,” J. Phys. Conf. Ser., vol. 1080, no. 1, p. 012017, Aug. 2018, doi: 10.1088 / 1742-6596 / 1080 / 1 / 012017. Additionally,strong bands around 3410 cm1correspond to O-H stretching vibrations of the hydroxyl groups, while the band at 1050 cm1can be attributed to C-0 stretching from hydroxyl groups in the polysaccharide structures as discussed in the references identified above. Finally, in the FTIR spectrum of CMC, a distinct absorption band at 1327 cm1was observed, corresponding to the symmetrical deformation of the CFF groups present in the carboxymethyl substituents, which are not present in the alginate structure as discussed by S. Hidayat and others in the reference identified above.

[0180] The FTIR spectra of the polymer solutions combined (Alg-CMC) with and without mucins and the crosslinked hydrogel with and without mucins are shown in FIG. 21B. Overall, there were no change in peaks when adding alginate to CMC aside from the absence or decrease in intensity of the band at 1327 cm'1that was observed in CMC. Similarly, when CaCh was added to the Alg-CMC composite there were no change in peaks aside from the asymmetric and symmetric stretching vibrations of carboxylic groups observed at 1583 and 1412 cm'1were shifted to 1590 and 1415 cm'1, respectively. Adding mucin to either CMC or Alg, two new big absorption bands appear at about 2919 and 2850 cm'1, which were assigned to C-H stretching and aromatic C-H bending of the mucins, respectively, as discussed by D. Petrash, S. Lalonde, M. Gingras, and K. Konhauser in “A surrogate approach to studying the chemical reactivity of burrow mucous linings in marine sediments,” Palaios, vol. 26, pp. 594-600, Sep. 2011, doi:10.2110 / palo.2010. pl0-140r. These peaks are less prominent when mucin is added to the CMC-Alg composite spectra, which could indicate that interactions of the two polymers together decrease their ability to interact with mucin. However, when mucin was added to the crosslinked Alg-CMC, these peaks reappeared. Finally, adding mucin to either CMC or alginate, two new small absorption bands appear at about 1263 and 798 cm'1, which were assigned to C-N stretching of amide groups and aromatic C-H of out plane bending, respectively, as discussed by Z. Hern.ndez-Nolasco, Ma. A. R.os-Corripio, J. V. Hidalgo-Contreras, P. H. Castellano, E. Rubio-Rosas, and A. S. Hern.ndez-C.zares in “Optimization of sodium alginate, taro starch and lactic acid based biodegradable films: Antimicrobial effect on a meat product,” LWT, vol. 192, p. 115718, Jan. 2024, doi:10.1016 / j.lwt.2023.115718; and by P. Darshani et al. in “Chemically synthesized butein and butin: Optical, structure and electrochemical redox functionality at electrode interface,” J. Photochem. Photobiol. B, vol. 182, Apr. 2018, doi: 10.1016 / j.jphotobiol.2018.04.001. However, these peaks do not appear or are less intense when mucin is added to CMC-Alg composite spectra, which again could indicate that interactions of the two polymers together decrease their ability to interact with mucin.

[0181] Selection of mucoadhesive hydrogels based on mucosal attachment time. To further assess the mucoadhesive properties of the hydrogel, a shear-based approach was implemented with porcine esophagus mucosa as the substrate. The flow chamber for this assessment was developed using an iterative design process, with each version refined and validated through simulations in COMSOL. Hydrogel formulations were applied directly to the mucosal tissue in the flow chamber, with a gravity-driven flow set to 53 mL / min to determine the point of detachment. Hydrogel attachment time was measured using a microscope, and mucoadhesive capabilities were compared based on this attachment duration.

[0182] Flow chamber COMSOL simulation. COMSOL simulations were conducted to investigate the flow characteristics in the designed chamber as the flow rate was incrementally increased to achieve hydrogel detachment. The mesh of the simulation was refined iteratively until minimal variation in results was observed, ultimately resulting in the use of a coarse mesh. The entry length required for the flow to become fully developed was then analyzed, as this increases proportional to the flow rate. A velocity plot (with fluid velocity (m / s) along the x-axis (m) of the flow chamber, illustrating the velocity profile across the channel) is shown in FIG. 22A, and was used to determine the optimal placement of the tissue slot within the channel. In the final design, with a flow rate of 53 mL / min, fully developed flow was achieved at approximately 40 mm from the chamber inlet, which guided the placement of the tissue slot at the 50 mm mark to maintain consistent flow conditions during testing. Additionally, the width of the flow channel was examined to minimize the effects of side walls on the shear profile over the tissue surface. FIG. 22B (with viscous force (N / m3) along the y-axis (mm) of the flowchannel over the tissue slot at the bottom of the flow chamber) shows the side wall effects led to variations in shear stress distribution, with higher shear occurring near the center of the channel. To mitigate this, the channel width was determined to be 7 mm, with a 1 mm clearance on either side of the tissue slot, which allowed for a more uniform shear distribution over the hydrogel.

[0183] Surface contour plots were used to evaluate the velocity and shear profiles within the flow chamber. FIGs 23 and 24 illustrate surface contour analysis of flow properties through the flow channel using COMSOL Multiphysics. The black squares represent the tissue slot.

[0184] FIG 23 (2D surface contour plot of fluid velocity in the middle of the flow chamber (z = 1.5875 mm), illustrating the velocity distribution across the channel) illustrates the uniform velocity profile over the tissue slot, demonstrating consistent flow conditions. The maximum velocity over the hydrogel surface was determined to be 0.08 m / s. This velocity profile directly influenced the shear forces experienced by the hydrogel.

[0185] FIG. 24 (2D surface contour plot of shear force at the bottom of the flow chamber (z = 0 mm), showing the shear stress exerted on the hydrogel) presents a surface contour plot of the bottom of the flow channel, showing a maximum shear stress of 0.11 N / m2over the hydrogel. These results indicate that the flow design achieved the intended uniform distribution of shear stress, which is crucial for accurately evaluating the mucoadhesive properties of the hydrogel.

[0186] Hydrogel attachment duration under constant shear. Using the designed flow chamber system, developed with the aid of the COMSOL simulations, the hydrogels were tested for attachment time on porcine esophagus mucosa. FIG. 25 presents the attachment times to porcine esophagus mucosa under constant shear in the flow chamber for all hydrogel formulations.

[0187] FIG 25 shows that gels with a storage modulus (G') between 100 and 200 Pa exhibited the longest attachment times, though this outcome may result from the experimental technique itself. More elastic gels with more structure (G' > 200 Pa) couldexperience greater perpendicular force, causing them to detach more quickly, while gels with lower elasticity (G' < 100 Pa) might simply wash away.

[0188] Selection of mucoadhesive hydrogels based on degradation. All hydrogels were tested for degradation in PBS to investigate any trends related to CMC and CaCh concentration. FIG. 26A illustrates results with hydrogels containing 2% alginate (A-C). FIG. 26B illustrates results with hydrogels containing 2% alginate and 0.5% CMC (G-I). FIG. 26C illustrates results with hydrogels containing 2% alginate and 1% CMC (M-O) were submerged in PBS to assess degradation. The values are the averages from six independent hydrogel mixtures (n = 6).

[0189] FIGs. 26A to 26C presents the degradation times of each hydrogel, highlighting their swelling behavior and degradation profiles. The hydrogels containing alginate alone exhibited the highest degree of swelling, with their mass increasing up to double their original weight. In contrast, as the concentration of CMC increased, the extent of swelling decreased, suggesting an inverse relationship between CMC content and swelling. Additionally, an increase in the concentration of CaCh crosslinker also led to reduced swelling. Specifically, Hydrogel A, which contained no CMC and 1 mg / mL CaCh, showed the most significant swelling, doubling its weight to approximately 0.8 g. On the other hand, Hydrogel O, with 1% CMC and 2 mg / mL CaCL, swelled the least, reaching a final weight of approximately 0.6 g. These results indicate that both CMC concentration and crosslinking density play a role in determining the hydrogels' ability to swell. Overall, most hydrogels fully degraded within the 8-hour testing period, except for Hydrogels B, H, and N. These three hydrogels shared a mid-level crosslinker concentration of 1.5 mg / mL CaCL, which appeared to contribute to their resistance to complete degradation. This finding suggests that an intermediate level of crosslinking may allow for prolonged hydrogel stability in PBS.

[0190] Gels A to O were tested for degradation in simulated saliva. FIGs. 27A to 27E illustrate a comparison of degradation of hydrogels A to O in simulated saliva.

[0191] FIGs 27A to 27E highlight the influence of crosslinking concentration on degradation rate in artificial saliva. Here, a lower concentration of crosslinker generallyresulted in more swelling, which also matched observations in PBS trials. As for degradation rates, lower crosslinking tended to degrade more slowly, with one exception: hydrogels containing 1% 250 kDa CMC, in which the middle CaCh concentration degraded the slowest, which was observed across all formulations in the PBS experiments.

[0192] Bacteria survival and release from microbe-seeded agar microspheres embedded in different hydrogel formulations

[0193] Microbe-seeded agar microspheres (either L. brevis or S. salivarius), prepared as disclosed above, were incorporated into the hydrogel formulations (Hydrogels A to O) following methods described in the paragraph above entitled “Assessing hydrogel biocompatibility with bacteria”, and cultured for 5 hrs. Microbial survival (FIG. 28A - L. brevis and FIG. 28B - S. salivarius), measured using CFU assay at 5 hr post seeding, were normalized against CFU measurement at 0 hr and expressed as fold change. Microbial release from hydrogel into artificial saliva were also measured in the saliva phase (FIG. 28C - L. brevis and FIG. 28D - S. salivarius) and expressed as normalized fold change, against 0 hr measurements.

[0194] Bacteria growth and viability when co-cultured in mucoadhesive hydrogel. Microbe-seeded agar microspheres, prepared as discussed above, were incorporated into the polymer blend of Hydrogel H as either monocultures or co-culture prior to crosslinking. After crosslinking, bacterial viability was assessed over 24 hours. FIGs. 29A and 29B illustrate the concentration of viable S. salivarius and L. brevis in hydrogel H over 24 hours. Each bar represents the average of three biological replicates (n = 3).

[0195] FIG. 29 A (monoculture of S. salivarius or L. brevis in hydrogel H and PBS) presents the CFU / ml for S. salivarius and L. brevis separately loaded as monocultures into Hydrogel H. The results indicate that overall, the CFU / mL was higher in PBS compared to the hydrogel for both bacterial species. Additionally, both S. salivarius and L. brevis showed a decrease in CFU / mL by two orders of magnitude after 24 hours in both the hydrogel and PBS conditions.

[0196] FIG. 29B (co-culture of S. salivarius and L. brevis in hydrogel H and PBS) illustrates the CFU / mL of S. salivarius and L. brevis when loaded together as co-culture into Hydrogel H. Consistent with the individual tests, the results showed that the CFU / mL values were higher in PBS for both bacterial species compared to the hydrogel. For S. salivarius, the CFU / mL in the hydrogel decreased by two orders of magnitude after 24 hours, whereas in PBS, there was only a decrease of one order of magnitude. For L. brevis, the CFU / mL in the hydrogel decreased by only 50%, while in PBS, there was a reduction of one order of magnitude. Overall, these results suggest that the hydrogel is neither completely inhibiting bacterial viability nor promoting bacterial growth when compared with PBS.

Claims

CLAIMSWhat is claimed is:

1. A composite comprising: biocompatible particles comprising: a biocompatible polymer, one or more microbes, and a growth media for the microbes; and a mucoadhesive hydrogel; wherein the particles are embedded in the hydrogel.

2. The composite of claim 1, wherein at least some of the particles comprise a plurality of different microbes; or the particles comprise at least a first portion and a second portion, wherein the first portion of the particles comprise a first growth media, and the second portion of the particles comprise a second growth media that is different from the first growth media.

3. The composite of claim 2, wherein the first portion of the particles comprise a first microbe, and the second portion of the particles comprise a second microbe that is different from the first microbe.

4. The composite of any one of claims 1 to 3, wherein the microbes are: in the form of a biofilm, such as in the form of particles of a biofilm; and / or in a planktonic state.

5. The composite of any one of claims 1 to 4, wherein: the microbes are embedded in the particles, and / or the microbes are on an outer surface of the particles.

6. The composite of any one of claims 1 to 5, wherein the biocompatible polymer comprises a gel-forming polymer, such as agar or gelatin.

7. The composite of any one of claims 1 to 6, wherein the particles are smaller than about 1000 pm, for example wherein the particles have an average diameter less than about 500 pm, such as an average diameter that is from about 100 pm to 200 pm.

8. The composite of any one of claims 1 to 7, wherein the mucoadhesive hydrogel comprises alginate, and optionally carboxymethyl cellulose (CMC).

9. The composite of claim 8, wherein the CMC has a molecular weight from about 90 kDa to about 700 kDa, such as from about 90 kDa to about 250 kDa, preferably about 250 kDa.

10. The composite of claim 8 or 9, wherein the alginate has a molecular weight from about 10 kDa to about 600 kDa, such as from about 12 kDa to about 40 kDa.

11. The composite of any one of claims 8 to 10, wherein the alginate and the CMC are present in a wt ratio from about 8: 1 to about 1 : 1 (alginate: CMC).

12. The composite of any one of claims 8 to 11, wherein the alginate is present in the hydrogel at a concentration from about 0.5 wt% to about 5 wt%, and / or the carboxymethyl cellulose is present in the hydrogel at a concentration from about 0.1 wt% to about 2 wt%.

13. The composite of any one of claims 8 to 12, wherein the hydrogel further comprises a crosslinker, such as a divalent cationic crosslinker, for example Ca2+, preferably calcium chloride at a concentration of from about 0.5 mg / mL to about 4 mg / mL.

14. The composite of any one of claims 1 to 13, wherein the hydrogel has a mucoadhesive strength sufficient to withstand a fluid shear stress of up to 0.01, 0.05, 0.1, 0.2, 0.4, 0.6, or 0.8 dyne / cm2.

15. The composite of any one of claims 1 to 14, wherein the microbes are a bacteria, such as Lactobacillus, for example Lactobacillus brevis, Lactobacillus reuteri UBLRu-87 , Lactobacillus acidophilus, Lactobacillus rhamnosus, Lactobacillusgensenii, Lactobacillus gasseri, Lactobacillus crispatus, or Lactobacillus fermentum; Bifidobacterium, for example Bifidobacterium longum or Bifidobacterium animalis subsp. lactis BB-12; Streptococcus, for example Streptococcus Salivarius,' or any combination thereof; a yeast, such as Saccharomyces, for example Saccharomyces boulardii; a fungus, such as Candida albicans,' or any combination thereof.

16. The composite of claim 3, wherein the particles comprise:1) a first portion of particles comprising Lactobacillus brevis and De Man- Rogosa-Sharpe (MRS) medium, a second portion of particles comprising Streptococcus Salivarius and brain heart infusion (BHI) medium, and a third portion of particles comprising Saccharomyces cerevisiae and BactotmPeptone and BactotmYeast Extract;2) a portion of particles comprising Lactobacillus acidophilus and MRS medium, and a second portion of particles comprising Lactobacillus gensenii and MRS medium, a third portion of particles comprising Lactobacillus gasseri and MRS medium, and a fourth portion of particles comprising Lactobacillus crispatus and MRS medium; or3) Lactobacillus acidophilus, Lactobacillus gensenii, Lactobacillus gasseri, Lactobacillus crispatus, and MRS medium.

17. A mixture of biocompatible particles, the mixture comprising: a first portion of biocompatible particles comprising at least one microbe and a first growth media, and a second portion of biocompatible particles comprising at least one microbe and a second growth media that is different from the first growth media.

18. The mixture of claim 17 wherein the first portion of the biocompatible particles comprise a first microbe, and the second portion of the biocompatible particles comprise a second microbe that is different from the first microbe.

19. The mixture of claim 17 or 18 wherein the microbes are:in the form of a biofilm, such as in the form of particles of a biofilm; and / or in a planktonic state.

20. The mixture of any one of claims 17 to 19 wherein: the microbes are embedded in the particles, and / or the microbes are on an outer surface of the particles.

21. A biocompatible particle comprising one or more microbes in the form of a biofilm, such as in the form of particles of a biofilm, and a growth media for the microbes.

22. The biocompatible particle of claim 21, wherein: the microbes are embedded in the particles, and / or the microbes are on an outer surface of the particles.

23. A combination comprising: particles comprising: one or more microbes and a growth media for the microbes; a polymer solution; and a crosslinker, such as a divalent cationic crosslinker, for example Ca2+, preferably calcium chloride at a concentration of from about 0.5 mg / mL to about 4 mg / mL; wherein the polymer solution and the crosslinker form a mucoadhesive hydrogel when mixed.

24. The combination of claim 23 wherein the particles and the polymer solution are in a mixture.

25. The combination of claim 23 or 24, wherein combining the particles, the polymer solution, and the crosslinker results in a composite according to any one of claims 1 to 16.

26. A method of microbial transplantation in a subject, the method comprising: applying the composite according to any one of claims 1 to 16 to a mucosa of the subject, such as oral, nasal, or vaginal mucosa.

27. A method comprising:A) providing biocompatible particles comprising: a biocompatible polymer, one or more microbes and a growth media for the microbes; providing a mucoadhesive hydrogel; and mixing the particles with the hydrogel; orB) providing biocompatible particles comprising: a biocompatible polymer, one or more microbes, and a growth media for the microbes; mixing the particles with a polymer solution to form a particle-polymer mixture; and mixing a crosslinker with the particle-polymer mixture to form a mucoadhesive hydrogel.

28. Use ofA) biocompatible particles comprising: a biocompatible polymer, one or more microbes and a growth media for the microbes, and a mucoadhesive hydrogel; orB) biocompatible particles comprising: a biocompatible polymer, one or more microbes and a growth media for the microbes, a polymer solution, and a crosslinker, for forming a composite for microbial transplantation to a mucosa of a subject, such as oral, nasal, or vaginal mucosa.

29. The method of claim 27 or the use of claim 28, wherein at least some of the particles comprise a plurality of different microbes; or the particles comprise at least a first portion and a second portion, wherein the first portion of the particles comprise a first growth media, and the second portion of the particles comprise a second growth media that is different from the first growth media.

30. The method of claim 27 or 29, or the use of claim 28 or 29, wherein the microbes are in the form of a biofilm, such as in the form of particles of a biofilm; and / or are in a planktonic state.

31. The method of any one of claims 27, 29 and 30, or the use of any one of claims 28 to 30, wherein the microbes are a bacteria, such as Lactobacillus, for example Lactobacillus brevis, Lactobacillus reuteri UBLRu-87 , Lactobacillus acidophilus, Lactobacillus rhamnosus, Lactobacillus gensenii, Lactobacillus gasseri, Lactobacillus crispatus, or Lactobacillus fermentum; Bifidobacterium, for example Bifidobacterium longum or Bifidobacterium animalis subsp. lactis BB-12; Streptococcus, for example Streptococcus Salivarius,' or any combination thereof; a yeast, such as Saccharomyces, for example Saccharomyces boulardii; a fungus, such as Candida albicans,' or any combination thereof.

32. The method of any one of claims 27 and 29 to 31, wherein providing the particles comprises: forming the particles from a biocompatible particle-forming material, and inoculating the particles with the microbes; or mixing the microbes with a biocompatible particle-forming material, and forming the particles from the mixture of particle-forming material and microbes.

33. The method of claim 32, wherein the particle-forming material comprises a gelforming polymer, such as agar or gelatin.

34. The method of claim 32 or 33, wherein forming the particles comprises drop casting or microfluidic particle generation.

35. The method of claim 34, wherein drop casting comprises pouring a hot 4% agar solution into cold canola oil to form an emulsion of agar particles, removing the oil from the particles.

36. The method of any one of claims 32 to 35, wherein inoculating the particles with the microbes comprises incubating the particles with a bacterial culture for a period of time, such as for about 12, 24 or 48 hours, optionally while shaking the mixture of particles and bacterial culture.

37. The method of claim 36, wherein the particles are seeded in a bacterial culture having an optical density (OD) of about 0.05 OD to about 0.4 OD.

38. The method of any one of claims 27 and 29 to 37, wherein providing a mucoadhesive hydrogel comprises mixing a polymer solution with a crosslinking solution, such as a solution comprising a divalent cationic crosslinker, for example Ca2+, to form the mucoadhesive hydrogel.

39. The method of claim 38, wherein the divalent cationic crosslinker is calcium chloride, preferably wherein the crosslinking solution comprises CaCh at a concentration from about 1 mg / mL to about 8 mg / mL.

40. The method of any one of claims 27 and 29 to 39, or the use of any one of claims 28 to 31, wherein the polymer solution comprises alginate, and optionally carboxymethyl cellulose (CMC).

41. The method or use of claim 40, wherein: the alginate is present in the polymer solution at a concentration from about 1 wt% to about 10 wt%, and / or the carboxymethyl cellulose is present in the hydrogel at a concentration from about 0.2 wt% to about 4 wt%.

42. Use of the composite according to any one of claims 1 to 16 for microbial transplantation to a mucosa of a subject, such as oral, nasal, or vaginal mucosa.

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