Title:  Bioadhesive microspheres and their use as drug delivery and imaging systems

United States Patent:   6,197,346

Assignee:   Brown Universtiy Research Foundation (Providence, RI)

Filed:  April 24, 1992

Bioadhesive polymers in the form of, or as a coating on, microcapsules containing drugs or bioactive substances which may serve for therapeutic, diagnostic, or diagnostic purposes in diseases of the gastrointestinal tract, are described. The polymeric microspheres all have a bioadhesive force of at least 11 mN/cm² (110 N/CM²). Techniques for the fabrication of bioadhesive microspheres, as well as a method for measuring bioadhesive forces between microspheres and selected segments of the gastrointestinal tract in vitro are also described. This quantitative method provides a means to establish a correlation between the chemical nature, the surface morphology and the dimensions of drug-loaded microspheres on one hand and bioadhesive forces on the other, allowing the screening of the most promising materials from a relatively large group of natural and synthetic polymers which, from theoretical consideration, should be used for making bioadhesive microspheres.

DETAILED DESCRIPTION OF THE INVENTION

In general terms, adhesion of polymers to tissues may be achieved by (i) physical or mechanical bonds, (ii) primary or covalent chemical bonds, and/or (iii) secondary chemical bonds (i.e., ionic). Physical or mechanical bonds can result from deposition and inclusion of the adhesive material in the crevices of the mucus or the folds of the mucosa. Secondary chemical bonds, contributing to bioadhesive properties, consist of dispersive interactions (i.e., Van der Waals interactions) and stronger specific interactions, which include hydrogen bonds. The hydrophilic functional groups responsible for forming hydrogen bonds are the hydroxyl (--OH) and the carboxylic groups (--COOH).

Adhesive microspheres have been selected on the basis of the physical and chemical bonds formed as a function of chemical composition and physical characteristics, such as surface area, as described in detail below. These microspheres are characterized by adhesive forces to mucosa of greater than 11 mN/cm².

Classes of Polymers Useful in Forming Bioadhesive Microspheres.

Suitable polymers that can be used to form bioadhesive microspheres include soluble and insoluble, nonbiodegradable and biodegradable polymers. These can be hydrogels or thermoplastics, homopolymers, copolymers or blends, natural or synthetic. A key feature, however, is that the polymer must have a bioadhesive force of between 110 N/m² (11 mN/cm²) and 5000 N/m² to a mucosal membrane of a patient.

Two classes of polymers appear to have potentially useful bioadhesive properties: hydrophilic polymers and hydrogels. In the large class of hydrophilic polymers, those containing carboxylic groups (e.g., poly[acrylic acid]) exhibit the best bioadhesive properties. One could infer that polymers with the highest concentrations of carboxylic groups should be the materials of choice for bioadhesion on soft tissues. Other promising polymers were: sodium alginate, carboxymethylcellulose, hydroxymethylcellulose and methylcellulose. Some of these materials are water-soluble, while others are hydrogels. Hydrogels have often been used for bioadhesive drug delivery; however, one big drawback of using hydrogels is the lack of long-term stability during storage which is a problem for therapeutic applications.

Rapidly bioerodible polymers such as poly[lactide-co-glycolide], polyanhydrides, polyorthoesters--which would expose carboxylic groups on the external surface as their smooth surface erodes--are excellent candidates for bioadhesive drug delivery systems in the gastrointestinal tract. Biodegradable polymers are more stable than hydrogels. In addition, polymers containing labile bonds, such as polyanhydrides and polyesters, are well known for their hydrolytic reactivity. Their hydrolytic degradation rates can generally be altered by simple changes in the polymer backbone.

Representative natural polymers are proteins, such as zein, serum albumin, or collagen, and polysaccharides, such as cellulose, dextrans, and alginic acid. Representative synthetic polymers include polyamides, polycarbonates, polyalkylenes, polyalkylene glycols, polyalkylene oxides, polyalkylene terephthalates, polyvinyl alcohols, polyvinyl ethers, polyvinyl esters, polyvinyl halides, polyvinylpyrrolidone, polyglycolides, polysiloxanes, polyurethanes, alkyl cellulose, hydroxyalkyl celluloses, cellulose ethers, cellulose esters, nitrocelluloses, polymers of acrylic and methacrylic esters, poly[lactide-co-glycolide], polyanhydrides, polyorthoester blends and copolymers thereof. Specific examples of these polymers include cellulose acetate, cellulose propionate, cellulose acetate butyrate, cellulose acetate phthalate, carboxymethyl cellulose, cellulose triacetate, cellulose sulphate, poly(methyl methacrylate), poly(ethyl methacylate), poly(butyl methacrylate), poly(isobutyl methacrylate), poly(hexyl methacrylate), poly(isodecyl methacrylate), poly(lauryl methacrylate), poly(phenyl methacrylate), poly(methyl acrylate), poly(isopropyl acrylate), poly(isobutyl acrylate), poly(octadecyl acrylate), polyethylene, polypropylene, poly(ethylene glycol), poly(ethylene oxide), poly(ethylene terephthalate), poly(vinyl alcohols), poly(vinyl acetate), poly(vinyl chloride), polystyrene and polyvinylpyrrolidone, polyurethane, polylactides, poly(butyric acid), poly(valeric acid), poly[lactide-co-glycolide], polyanhydrides, polyorthoesters, poly(fumaric acid), and poly(maleic acid).

These polymers can be obtained from sources such as Sigma Chemical Co., St. Louis, Mo., Polysciences, Warrenton, Pa., Aldrich, Milwaukee, Wis., Fluka, Ronkonkoma, N.Y., and BioRad, Richmond, Calif.

In the studies detailed below, a variety of polymer microspheres were compared for adhesive force to mucosa. Negatively charged hydrogels, such as alginate and carboxymethylcellulose, that expose carboxylic groups on the surface, were selected, as well as some positively-charged hydrogels, such as chitosan. The rationale behind this choice is the fact that most cell membranes are actually negatively charged and there is still no definite conclusion as to what the most important property is in obtaining good bioadhesion to the wall of the gastrointestinal tract. Thermoplastic polymers: (a) non-erodible, neutral polystyrene, and (b) semicrystalline bioerodible polymers that generate carboxylic groups as they degrade--polylactides and polyanhydrides, were also selected. Polyanhydrides are good candidates for bioadhesive delivery systems since, as hydrolysis proceeds, more and more carboxylic groups are exposed to the external surface. Polylactides erode by bulk erosion; furthermore, the erosion is slower. In designing these systems as bioadhesive polymers, polymers that have high concentrations of carboxylic acid were preferred. This was done by using low molecular weight polymers (Mw 2000), since low molecular weight polymer contain high concentration of carboxylic acids at the end groups.

Measurement of Bioadhesive Studies Using a Tensile Technique

The adhesive forces between polymer microspheres and segments of intestinal rat tissue can be measured using the Cahn DCA-322, as shown in FIG. 1. Although this piece of equipment is designed for measuring contact angles and surface tensions using the Wilhelmy plate technique, it is also an extremely accurate microbalance. The DCA-322 system includes a microbalance stand assembly, a Cahn DACS computer, and an Okidata Microline 320 dot matrix printer. The microbalance unit consists of stationary sample and tare loops and a moving stage powered by a stepper motor. The balance can be operated with samples weighing up to 3.0 g, and has a sensitivity rated at 0.001 dynes. The stage speed can be adjusted from 20 to 264 .mu.m/sec using the factory installed motor, or from 2-24 .mu.m/sec using the optional slow motor. Adhesive forces were measured by attaching a polymer sample to one of the sample loops and placing an adhesive substrate 10, intestinal tissue, below it on the moving stage 20.

For adhesive measurements, 1.5 cm sections are cut from the excised intestine. These were then sliced lengthwise and spread flat, exposing the lumen side. The samples were then placed in a temperature-regulated chamber 30, clamped 32 at their edges, and covered with approximately 0.9 cm high level of phosphate buffer saline, as shown in FIG. 1. Physiologic conditions were maintained in the chamber. The chamber was then placed in the microbalance enclosure and a microsphere, mounted on a wire and hung from the sample loop of the microbalance, was brought in contact with the tissue. The microspheres were left in contact with the tissue for seven minutes with an applied force of approximately 0.25 mN and then pulled vertically away from the tissue sample while recording the required force for detachment. The contact area was estimated to be the surface area of the spherical cap defined by the depth of penetration of the bead below the surface level of the tissue. The force values were normalized by the projected area of this cap (Area=.pi.R² -.pi.(R-a)², where R is the microsphere radius and a is the depth of penetration. For microspheres larger than 800 .mu.m, a=400 m was used, for smaller microspheres a=R was used.

Graphs of force versus distance as well as force versus time were studied. FIG. 2 shows a typical graph of force versus stage position for the P(CPP-SA) 20:80 microspheres. Point A in FIG. 2 indicates the applied force, which can be varied in each experiment, and which indirectly affects the degree of penetration into the tissue. Portion AB indicates the adhesion time, the time the sphere is left to interact with the tissue before movement of the stage is started to separate the surfaces. Segment BC indicates the elevation of the sphere to 0 mg applied force (point C). During the early part of the tensile experiment (CD), the force increases as a function of stage position, while the contact area between the sphere and the mucus is assumed to be constant and equal to the surface of the immersed sphere. The next portion of this curve (DE) indicates a period where partial detachment of the polymeric device from the mucus occurred with some changes in the contact area. The last point (E) is the detachment of the sphere from the mucus. In some cases, a detachment does not occur until the microsphere has been moved to a height of 4 mm above the initial level of contact.

From these graphs it is possible to determine the maximum force applied to the sample, the maximum adhesive force, the distance required for detachment of the samples and the work of adhesion (the surface under the force versus stage position curves CDE). More importantly, it allows quantification of the adhesive forces of a variety of individual microspheres and correlation of these forces with physical and chemical properties of the polymers.

Modification of Bioadhesive Polymers to Increase Bioadhesive Force.

The polymers are selected from commercially available polymers based on their adhesive properties using the method described above to determine those polymers forming microspheres (either as solid polymer or as a polymeric coating on a different material) having an adhesive force greater than 11 nN/mg². The microspheres are then formed having an appropriate surface area to provide the desired adhesive forces. The polymers (or polymeric surface) can also be modified as described below to increase the bioadhesive properties of the polymer.

For example, the polymers can be modified by increasing the number of carboxylic groups accessible during biodegradation, or on the polymer surface. The polymers can also be modified by binding amino groups to the polymer.

The attachment of polyethyleneimine or polylysine-coated acrylamide beads to intestine is probably due to the electrostatic attraction of the cationic groups coating the beads to the net negative charge of the mucus. The mucopolysaccharides and mucoproteins of the mucin layer, especially the sialic acid residues, are responsible for the negative charge coating. Any ligand with a high binding affinity for mucin could also be covalently linked to most microspheres with the appropriate chemistry, such as CDI, and be expected to influence the binding of microspheres to the gut. The ligand affinity need not be based only on electrostatic charge, but other useful physical parameters such as solubility in mucin or else specific affinity to carbohydrate groups.

The covalent attachment of any of the natural components of mucin in either pure or partially purified form to the microspheres would decrease the surface tension of the bead-gut interface and increase the solubility of the bead in the mucin layer. The list of useful ligands would include but not be limited to the following: sialic acid, neuraminic acid, n-acetyl-neuraminic acid, n-glycolylneuraminic acid, 4-acetyl-n-acetylneuraminic acid, diacetyl-n-acetylneuraminic acid, glucuronic acid, iduronic acid, galactose, glucose, mannose, fucose, or else any of the partially purified fractions prepared by chemical treatment of naturally occurring mucin, e.g., mucoproteins, mucopolysaccharides and mucopolysaccharide-protein complexes.

The covalent attachment of lectins to microspheres would also increase the affinity of the spheres to components of the mucin and mucosal cell layer. Useful lectin ligands include lectins isolated from: Abrus precatroius, Agaricus bisporus, Anguilla anguilla, Arachis hypogaea, Pandeiraea simplicifolia, Bauhinia purpurea, Caragan arobrescens, Cicer arietinum, Codium fragile, Datura stramonium, Dolichos biflorus, Erythrina corallodendron, Erythrina cristagalli, Euonymus europaeus, Glycine max, Helix aspersa, Helix pomatia, Lathyrus odoratus, Lens culinaris, Limulus polyphemus, Lysopersicon esculentum, Maclura pomifera, Momordica charantia, Mycoplasma gallisepticum, Naja mocambique, as well as the lectins Concanavalin A and Succinyl-Concanavalin A.

Formation of Microspheres.

As used herein, microspheres includes microparticles and microcapsules (having a core of a different material than the outer wall), having a diameter in the nanometer range up to 1 mm. The microsphere may consist entirely of bioadhesive polymer or have only an outer coating of bioadhesive polymer.

Microspheres have been fabricated from the different polymers. Polylactic blank microspheres were fabricated by using two methods: solvent evaporation, as described by E. Mathiowitz, et al., J. Scanning Microscopy, 4, 329 (1990); L. R. Beck, et al., Fertil. Steril., 31, 545 (1979); and S. Benita, et al., J. Pharm. Sci., 73, 1721 (1984); and hot-melt microencapsulation, as described by E. Mathiowitz, et al., Reactive Polymers, 6, 275 (1987). Polyanhydrides made of bis-carboxyphenoxypropane and sebacic acid with molar ratio of 20:80 (P(CPP-SA) 20:80) (Mw 20,000) were prepared by hot-melt microencapsulation. Poly(fumaric-co-sebacic) (20:80) (Mw 15,000) blank microspheres were prepared by hot-melt microencapsulation. Polystyrene microspheres were prepared by solvent evaporation.

Hydrogel microspheres were prepared by dripping the solution from a reservoir though a 250 microliter pipet tip into a stirred ionic bath. The specific conditions for alginate, chitosan, alginate/polyethylenimide (PEI) and carboxymethyl cellulose (CMC) are listed in Table 1.

a. Solvent Evaporation. In this method the polymer is dissolved in a volatile organic solvent, methylene chloride. The drug (either soluble or dispersed as fine particles) is added to the solution, and the mixture is suspended in an aqueous solution that contains a surface active agent such as poly(vinyl alcohol). The resulting emulsion is stirred until most of the organic solvent evaporates, leaving solid microspheres. Several different polymer concentrations will be used (0.05-0.20 g/ml). The solution will be loaded with a drug and suspended in 200 ml of vigorously stirred distilled water containing 1% (w/v) poly(vinyl alcohol) (Sigma). After 4 hours of stirring, the organic solvent will have evaporated from the polymer, and the resulting microspheres are washed with water and dried overnight in a lyophilizer. Microspheres with different sizes (1-1000 microns) and morphologies can be obtained by this method. This method is useful for relatively stable polymers like polyesters and polystyrene.

However, labile polymers, such as polyanhydrides, may degrade during the fabrication process due to the presence of water. For these polymers, the following two methods, which are performed in completely organic solvents, are more useful.

b. Hot Melt Microencapsulation. In this method, the polymer is first melted and then mixed with the solid particles of the dye or drug that have been sieved to less than 50 microns. The mixture is suspended in a non-miscible solvent (like silicon oil), and, while stirring continuously, heated to 5^oC. above the melting point of the polymer. Once the emulsion is stabilized, it is cooled until the polymer particles solidify. The resulting microspheres are washed by decantation with petroleum ether to give a free-flowing powder. Microspheres with sizes between one to 1000 microns can be obtained with this method. The external surfaces of spheres prepared with this technique are usually smooth and dense. This procedure is used to prepare microspheres made of polyesters and polyanhydrides. However, this method is limited to polymers with molecular weights between 1000-50000.

c. Solvent Removal. This technique was primarily designed for polyanhydrides. In this method, the drug is dispersed or dissolved in a solution of the selected polymer in a volatile organic solvent like methylene chloride. This mixture is suspended by stirring in an organic oil (such as silicon oil) to form an emulsion. Unlike solvent evaporation, this method can be used to make microspheres from polymers with high melting points and different molecular weights. Microspheres that range between 1-300 microns can be obtained by this procedure. The external morphology of spheres produced with this technique is highly dependent on the type of polymer used.

d. Hydrogel Microspheres. Microspheres made of gel-type polymers, such as alginate, are produced by dissolving the polymer in an aqueous solution, suspending the barium sulphate or any other active material in the mixture and extruding through a microdroplet forming device, producing microdroplets which fall into a hardening bath, that is slowly stirred. The advantage of these systems is the ability to further modify the surface of the microspheres by coating them with polycationic polymers, like polylysine after fabrication. Microsphere particles are controlled by using various size extruders. Table 1 summarizes the various hydrogels and the concentrations that were used to manufacture them.

Claim 1 of 11 Claims

We claim:

1. A method for delivering a compound to a patient comprising administering to a mucosal membrane of a patient in need thereof an effective amount of a compound within a microparticle fabricated using a method yielding a morphology and particle diameter with a polymeric surface with an adhesive force of between 110 N/m² and 5000 N/M² as measured on living rat intestine, wherein the polymer is selected from the group consisting of synthetic polymers and hydrophilic proteins

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