Biocompatible member

Abstract

The purpose of the present invention is to provide a biocompatible member which is easily integrated with a biological tissue. This biocompatible member has a surface of which at least a portion is composed of a porous body, wherein the porous body is an aggregate of polymer particles.

Description

Biocompatible member

[0001] The present invention relates to a biocompatible member.

[0002] Conventionally, biocompatible members such as artificial blood vessels and artificial skin have been used. As biocompatible members, in addition to tubular ones such as artificial blood vessels and sheet-like ones such as artificial skin, those used as constituent members of devices equipped with subcutaneous implantable access ports as described in Patent Document 1 below are also known.

[0003] For materials for constructing this type of biocompatible member, polymers with excellent biocompatibility such as polytetrafluoroethylene (PTFE) and silicone-based polymers are adopted. Among them, a material made porous by stretching polytetrafluoroethylene is widely used under the name of ePTFE and the like.

[0004] Japanese Patent No. 7142007

[0005] If the biocompatible member is such that the contact state with the living body is not sufficiently good and gaps where air, body fluids, etc. can stay are formed between the living body tissue, bacteria may multiply in the gaps, so it is desired to easily integrate with the living body tissue. However, such a desire has not been fully satisfied. Therefore, an object of the present invention is to provide a biocompatible member that can easily integrate with living body tissue.

[0006] In order to solve the above problems, the present invention provides a biocompatible member in which at least a part of the surface is composed of a porous body, and the porous body is an aggregate of polymer particles.

[0007] According to the present invention, a biocompatible member that can easily integrate with living body tissue can be provided.

[0008] Figure 1 is a schematic perspective view showing an artificial blood vessel. Figure 2 is a diagram (scanning electron microscope image) showing a cross-section of a porous material constituting a biocompatible component. Figure 3a is a schematic diagram showing a mold used in the manufacture of corrugated tubes that make up an artificial blood vessel. Figure 3b is a schematic diagram showing the manufacturing process of corrugated tubes that make up an artificial blood vessel. Figure 4a is a schematic perspective view showing the state of the segmented molds that can be used in the manufacture of corrugated tubes that make up an artificial blood vessel before assembly. Figure 4b is a schematic perspective view showing the state of the segmented molds that can be used in the manufacture of corrugated tubes that make up an artificial blood vessel after assembly. Figure 4c is a schematic perspective view showing the disassembly of the segmented molds after the manufacture of the corrugated tubes. Figure 5 is a partially cutaway schematic front view showing a vascular access device. Figure 6 is a schematic diagram of the rat implantation evaluation. Figure 7a is an image (SEM image) of a cross-section of a sample made of a sintered body of polyester polymer particles observed with a scanning electron microscope. Figure 7b is a diagram showing the Si element mapping in the observation area in Figure 7a. Figure 8 shows the state of a sample for rat implantation evaluation. Figure 9a shows the overall state of sample 1-1 that was evaluated with implantation. Figure 9b shows the details of sample 1-1 that was evaluated after being planted. Figure 9c shows the details of sample 1-1 that was evaluated after being planted. Figure 9d shows the details of sample 1-1 that was evaluated after being planted. Figure 9e shows the details of sample 1-1 that was evaluated after being planted. Figure 10a shows the overall appearance of sample 1-2 that was evaluated after being planted. Figure 10b shows the details of sample 1-2 that was evaluated after being planted. Figure 10c shows the details of sample 1-2 that was evaluated after being planted. Figure 10d shows the details of sample 1-2 that was evaluated after being planted. Figure 10e shows the details of sample 1-2 that was evaluated after being planted. Figure 11a shows the overall appearance of sample 1-3 that was evaluated after being planted. Figure 11b shows the details of sample 1-3 that was evaluated after being planted. Figure 11c shows the details of sample 1-3 that was evaluated after being planted. Figure 11d shows the details of sample 1-3, which was evaluated after being planted. Figure 11e shows the details of sample 1-3, which was evaluated after being planted. Figure 12a shows the overall appearance of sample 1-4, which was evaluated after being planted.Figure 12b shows the details of sample 1-4 that was evaluated after being planted. Figure 12c shows the details of sample 1-4 that was evaluated after being planted. Figure 12d shows the details of sample 1-4 that was evaluated after being planted. Figure 12e shows the details of sample 1-4 that was evaluated after being planted. Figure 13a shows the overall appearance of sample 2-1 that was evaluated after being planted. Figure 13b shows the details of sample 2-1 that was evaluated after being planted. Figure 13c shows the details of sample 2-1 that was evaluated after being planted. Figure 13d shows the details of sample 2-1 that was evaluated after being planted. Figure 13e shows the details of sample 2-1 that was evaluated after being planted. Figure 14a shows the overall appearance of sample 2-2 that was evaluated after being planted. Figure 14b shows the details of sample 2-2 that was evaluated after being planted. Figure 14c shows the details of sample 2-2 that was evaluated after being planted. Figure 14d shows the details of sample 2-2 that was evaluated after being planted. Figure 14e shows the details of sample 2-2 that was evaluated after being planted. Figure 15a shows the overall appearance of sample 2-3 that was evaluated after being planted. Figure 15b shows the details of sample 2-3 that was evaluated after being planted. Figure 15c shows the details of sample 2-3 that was evaluated after being planted. Figure 15d shows the details of sample 2-3 that was evaluated after being planted. Figure 15e shows the details of sample 2-3 that was evaluated after being planted. Figure 16a shows the overall appearance of sample 2-4 that was evaluated after being planted. Figure 16b shows the details of sample 2-4 that was evaluated after being planted. Figure 16c shows the details of sample 2-4 that was evaluated after being planted. Figure 16d shows the details of sample 2-4 that was evaluated after being planted. Figure 16e shows the details of sample 2-4 that was evaluated after being planted. Figure 17a shows the overall appearance of sample 3-1, which was evaluated after being planted. Figure 17b shows the details of sample 3-1, which was evaluated after being planted. Figure 17c shows the details of sample 3-1, which was evaluated after being planted. Figure 17d shows the details of sample 3-1, which was evaluated after being planted. Figure 17e shows the details of sample 3-1, which was evaluated after being planted. Figure 18a shows the overall appearance of sample 3-2, which was evaluated after being planted. Figure 18b shows the details of sample 3-2, which was evaluated after being planted.Figure 18c shows the details of sample 3-2 that was evaluated after being planted. Figure 18d shows the details of sample 3-2 that was evaluated after being planted. Figure 18e shows the details of sample 3-2 that was evaluated after being planted. Figure 19a shows the overall appearance of sample 3-3 that was evaluated after being planted. Figure 19b shows the details of sample 3-3 that was evaluated after being planted. Figure 19c shows the details of sample 3-3 that was evaluated after being planted. Figure 19d shows the details of sample 3-3 that was evaluated after being planted. Figure 19e shows the details of sample 3-3 that was evaluated after being planted. Figure 20a shows the overall appearance of sample 3-4 that was evaluated after being planted. Figure 20b shows the details of sample 3-4 that was evaluated after being planted. Figure 20c shows the details of sample 3-4 that was evaluated after being planted. Figure 20d shows the details of sample 3-4 that was evaluated after being planted. Figure 20e shows the details of sample 3-4, which was evaluated after being planted. Figure 21a shows the overall appearance of sample 4-1, which was evaluated after being planted. Figure 21b shows the details of sample 4-1, which was evaluated after being planted. Figure 21c shows the details of sample 4-1, which was evaluated after being planted. Figure 21d shows the details of sample 4-1, which was evaluated after being planted. Figure 21e shows the details of sample 4-1, which was evaluated after being planted. Figure 22a shows the overall appearance of sample 4-2, which was evaluated after being planted. Figure 22b shows the details of sample 4-2, which was evaluated after being planted. Figure 22c shows the details of sample 4-2, which was evaluated after being planted. Figure 22d shows the details of sample 4-2, which was evaluated after being planted. Figure 22e shows the details of sample 4-2, which was evaluated after being planted. Figure 23a shows the overall appearance of sample 4-3, which was evaluated after being planted. Figure 23b shows a detailed view of sample 4-3 that was evaluated after being planted. Figure 23c shows a detailed view of sample 4-3 that was evaluated after being planted. Figure 23d shows a detailed view of sample 4-3 that was evaluated after being planted. Figure 23e shows a detailed view of sample 4-3 that was evaluated after being planted. Figure 24a shows an overall view of sample 4-4 that was evaluated after being planted. Figure 24b shows a detailed view of sample 4-4 that was evaluated after being planted. Figure 24c shows a detailed view of sample 4-4 that was evaluated after being planted.Figure 24d shows the details of sample 4-4 that was evaluated after being planted. Figure 24e shows the details of sample 4-4 that was evaluated after being planted.

[0009] Embodiments of the present invention will be described below with reference to the drawings. Two embodiments, the first and second embodiments, will be described below. Specifically, as examples of the biocompatible member of the present invention, examples of use in artificial blood vessels and examples of use in vascular access devices will be given below. However, the specific embodiments of the biocompatible member of the present invention are not limited to these.

[0010] (First Embodiment) First, a first embodiment of the biocompatible member will be described with reference to Figure 1. Figure 1 is an artificial blood vessel 1, which is a biocompatible member of the first embodiment. The artificial blood vessel 1 illustrated in the figure comprises a main vessel 11 and branch vessels 12 branched from the main vessel 11. The artificial blood vessel 1 illustrated in the figure comprises a plurality of branch vessels 12, including a first branch vessel 121, a second branch vessel 122, and a third branch vessel 123.

[0011] The artificial blood vessel 1 is made of a polymer composition with excellent flexibility, and has excellent elasticity and bendability. The main tube 11 and the branch tubes 12 are straight when placed on a horizontal surface without any stress applied (natural state). The three branch tubes 12 are straight in their natural state and are smaller in diameter than the main tube 11. The main tube 11 and the branch tubes 12 are corrugated tubes, and have a bellows-like shape with large diameter sections 11L and 12L that define their outer diameters and small diameter sections 11S and 12S that are smaller in diameter than the large diameter sections 11L and 12L and define the inner diameters of the main tube 11 and the branch tubes 12, respectively, arranged alternately in the longitudinal direction. The corrugated pipe may be a ring-type corrugated pipe in which the larger diameter sections 11L, 12L and the smaller diameter sections 11S, 12S are each independent ring-shaped and arranged alternately in the longitudinal direction, or it may be a spiral-type corrugated pipe in which the larger diameter sections 11L, 12L and the smaller diameter sections 11S, 12S are spiral-shaped and extend radially so that they form a double helix. The main pipe 11 and the branch pipe 12 not only exhibit elasticity and flexibility due to the flexibility of the material itself, but also exhibit excellent elasticity and flexibility from a structural standpoint.

[0012] The three branch tubes 12 of the artificial blood vessel 1 branch off in a direction perpendicular to the longitudinal direction of the main tube 11, roughly in the longitudinal center of the main tube 11, and each branch extends in the same direction, with the branching points arranged in a line from one end 11a to the other end 11b of the main tube 11.

[0013] The artificial blood vessel 1, as illustrated in Figure 1, can be used, for example, to substitute for the aortic arch by curving the main vessel 11 into an arc. More specifically, the artificial blood vessel 1 illustrated in Figure 1 can be used to substitute for the aortic arch by having one end 11a of the main vessel 11 connected to the ascending aorta a and the other end 11b connected to the descending aorta a. In this artificial blood vessel 1, the branching point of the first branch 121 is located closer to the one end 11a of the main vessel 11 than the second branch 122 and the third branch 123, and can be used, for example, to connect to the brachiocephalic artery c. The branching point of the third branch 123 is located closer to the other end 11b of the main vessel 11 than the first branch 121 and the second branch 122, and can be used, for example, to connect to the left subclavian artery d. The second branch canal 122 is positioned such that its branching point is between the first branch canal 121 and the third branch canal 123, and can be used, for example, to connect to the left common carotid artery e.

[0014] In the artificial blood vessel 1 illustrated in Figure 1, the main tube 11 and the branch tubes 12 are made of a common polymer composition. The artificial blood vessel 1 is a porous body 100 made of a polymer composition. The porous body 100 is a aggregate of polymer particles, and more specifically, a sintered body obtained by heating polymer particles to a temperature close to their melting point and sintering them. In the artificial blood vessel 1 illustrated in Figure 1, the voids between the polymer particles constituting the porous body 100 are open to the outer surface 1s1 and the inner surface 1s2, respectively.

[0015] The artificial blood vessel 1 is constructed by individually manufacturing four corrugated tubes (porous materials) that each constitute one main tube 11 and three branch tubes 12, and then connecting them together.

[0016] Figure 2 is an SEM image taken with a scanning electron microscope (SEM) of a cross-section of a porous body 100, which is composed of polyester polymer particles, the same as the one used as an evaluation sample in the later examples. As shown in this figure, the porous body 100 is a solidified body in which polymer particles are solidified into a three-dimensional network. The porous body 100 has a polymer portion P composed of solidified polymer particles and void portions V which are the spaces between the polymer particles. The artificial blood vessel 1 has excellent elasticity and flexibility due to its macroscopic structural characteristics, such as its bellows-like shape, and also exhibits excellent elasticity and flexibility in terms of its microscopic structural characteristics, such as the fact that the polymer portion P is a three-dimensional network structure.

[0017] Each of the multiple polymer particles constituting the porous body 100 is bonded to some of the polymer particles of adjacent polymer particles in a direction parallel to the outer surface 1s1 (planar direction) or in a direction perpendicular to the outer surface 1s1 (depth direction), leaving a wide gap between it and the remaining polymer particles. These gaps between particles are continuous in the planar and depth directions, forming interconnected pores. Moreover, these interconnected pores are formed to have meandering and branching characteristics, and in a complementary relationship with the polymer portion P, which is a three-dimensional network structure, they form a three-dimensional network-like void portion V that fills a single space. That is, the void portion V is formed in a complex, interwoven state and is continuous in the depth direction from the surface of the porous body 100.

[0018] In conventional biocompatible materials, ePTFE has voids formed between fibrils parallel to the stretching direction, which are elongated in the stretching direction. That is, the opening shape of the voids in ePTFE is a slit shape with an extremely short width relative to its length. Also, the voids in ePTFE are relatively monotonous in the depth direction. The porous body 100 constituting the artificial blood vessel 1 of this embodiment is formed by the bonding of polymer particles, and the opening shape is wide, making it easy for cells to penetrate. For example, as shown by the double arrows No. 1 to No. 3 in Figure 2, the width of the voids in the cross-section of the porous body 100 (the distance between the cross-sections of adjacent polymer parts P) is several tens of micrometers to several hundred micrometers. Furthermore, in a polymer particle aggregate, the voids V are interconnected in an intricate manner into the depths, making it difficult for cells that have penetrated to escape from the voids V. For example, if biological tissue in contact with the outer surface 1s1 of the artificial blood vessel 1 is subjected to stress in a direction that moves it away from the outer surface 1s1, if the voids are formed linearly in the depth direction, cells that have already invaded may be pulled back, and in some cases may even fall out completely. However, in the artificial blood vessel 1 of this embodiment, the voids V are interconnected in an intricate manner that extends deep into the vessel, so even in such cases, changes that would reduce the depth of cell invasion are less likely to occur. Therefore, the artificial blood vessel 1 of this embodiment, which is composed of a polymer particle aggregate, can integrate with the biological tissue it comes into contact with more quickly than conventional artificial blood vessels composed of ePTFE.

[0019] The ratio of the polymer portion P to the total volume of the void portion V and the polymer portion P (the apparent volume calculated from the external shape of the porous body 100) is advantageous if it is below a certain level, considering the ease with which cells can penetrate. This ratio is, for example, 90 vol% or less. This ratio may also be 80 vol% or less, 70 vol% or less, or 60 vol% or less. On the other hand, considering the strength, it is considered advantageous if this ratio is above a certain level. This ratio is, for example, 25 vol% or more. This ratio may also be 30 vol% or more, or 35 vol% or more.

[0020] As shown in Figure 2, where the circle is labeled X, if the diameter D of the largest circle that can be drawn in a location within the cross-section of the porous body 100 such that it encloses only the cross-section of the void portion V without enclosing the cross-section of the polymer portion P is taken as the opening diameter of the void portion V at that location, then considering the ease with which cells can penetrate, it may be advantageous for the opening diameter to be above a certain level. On the other hand, in terms of exhibiting excellent sealing properties to suppress leakage and bacterial permeation, it is considered advantageous for the opening diameter (D) to be below a certain level. The opening diameter (D) is, for example, 5 μm or more and 500 μm or less. The opening diameter (D) may be, for example, 10 μm or more, or 15 μm or more. The opening diameter (D) may be, for example, 400 μm or less, or 300 μm or less.

[0021] Preferably, the polymer portion P and the void portion V have a shape that allows multiple (two or more) circles of the above-mentioned size to be drawn within a randomly set square area with sides of 1 mm in the cross-section of the porous body 100 without overlapping. The polymer portion P and the void portion V in the porous body 100 may have a shape that allows three or more circles of the above-mentioned size to be drawn within a 1 mm square area, or it may have a shape that allows four or more circles to be drawn, or it may have a shape that allows five or more circles to be drawn. The number of circles that can be drawn within a square area with sides of 1 mm may be, for example, 100 or less, or 50 or less.

[0022] The polymer particles that make up the porous body 100 may consist of a single polymer species, may contain multiple types of polymers, or may contain additives other than polymers. The porous body 100 may consist of a solidified body in which polymer particles are directly joined, or it may contain a binder that adheres the polymer particles together. The porous body 100 may also be a sintered body in which polymer particles are heat-fused together.

[0023] The type of polymer contained in the polymer particles is not particularly limited as long as it can be used in biocompatible materials, but examples include polyethylene such as ultra-high molecular weight polyethylene; polypropylene such as homopolypropylene and copolymerized polypropylene (e.g., block copolymers or random copolymers of propylene and ethylene); and polyolefin-based thermoplastic elastomers with polyethylene or polypropylene as hard segments and ethylene-propylene rubber as soft segments. Other examples of polymers contained in the polymer particles include polyesters such as polyethylene terephthalate; and polyester-based thermoplastic elastomers with polybutylene terephthalate or polybutylene isophthalate as hard segments and polyethers such as polytetramethylene glycol as soft segments. Polystyrene-based polymers such as polystyrene, poly-α-methylstyrene, acrylonitrile-styrene-butadiene copolymer (ABS), and styrene-based thermoplastic elastomers which are block copolymers of polystyrene blocks and polyolefins are also possible. The polymers contained in the polymer particles may also be silicone-based polymers; polysulfone; polytetrafluoroethylene; polymethyl methacrylate; polyvinyl chloride, etc. The polymer contained in the polymer particles may be polyamide-based polymers such as polyamide; amide-based thermoplastic elastomers with polyamide as the hard segment and polyester, polyether, etc. as the soft segment. Other polymers that can be contained in the polymer particles include, for example, polyurethane; polyurethane-based thermoplastic elastomers with polyurethane containing short-chain diols and polyisocyanates as repeating units as the hard segment and polyurethane containing long-chain diols and polyisocyanates as repeating units as the soft segment.

[0024] The polymer contained in the polymer particles may, for example, exhibit a melt viscosity of 300 Pa·s or more at 200°C. The melt viscosity of the polymer at 200°C may be 400 Pa·s or more, 500 Pa·s or more, or 600 Pa·s or more. The melt viscosity of the polymer contained in the polymer particles at 200°C may be, for example, 2000 Pa·s or less. The melt viscosity of the polymer can be measured, for example, by heating the polymer sufficiently to the measurement temperature (200°C) using a flow tester equipped with a die with a diameter of 1 mmφ and a length of 10 mm, and then applying a load of 10 kgf. The melt viscosity of the polymer can be determined as the arithmetic mean of multiple measurements (for example, 5 times).

[0025] Among the polymers exemplified above, ultra-high molecular weight polyethylene and thermoplastic elastomers (olefin-based thermoplastic elastomers, ester-based thermoplastic elastomers, styrene-based thermoplastic elastomers, amide-based thermoplastic elastomers, urethane-based thermoplastic elastomers, etc.) are suitable because their high melt viscosity makes it difficult for the polymer particles to soften excessively when preparing a solidified body (sintered body) under heating and pressure, thus making it easier to secure the desired void space. The porous body is preferably configured to contain one or more of the aforementioned polymer particles selected from the group consisting of polyethylene particles, polypropylene particles, polyester-based polymer particles, polystyrene-based polymer particles, polyetheretherketone particles, and polyurethane particles.

[0026] Of the polymers mentioned above, for silicone-based polymers, for example, all units except the two ends of the molecule are substantially D units (R ２ SiO ２／２ It may consist only of D units and T units (RSIO ３／２ ) and Q units (SiO ４／２ ) may also contain such as. In other words, the silicone polymer may have a chain-like molecular structure, commonly referred to as silicone oil, or a branched molecular structure, commonly referred to as silicone resin. Regarding what siloxane units are contained in such silicone polymers,２９ This can be determined by Si-NMR. Examples of substituents (R) in the siloxane unit include alkyl groups such as methyl groups and aryl groups such as phenyl groups.

[0027] For example, the silicone oil may be a dimethyl silicone oil with dimethylsiloxane as the main repeating unit; a methylphenyl silicone oil with dimethylsiloxane and diphenylsiloxane as the main repeating units; or a methylhydrogen silicone oil with dimethylsiloxane and monomethylsiloxane as the main repeating units. Furthermore, the silicone oil may be a modified product, for example, a modified product that has been modified to impart reactive functional groups such as amino modification, epoxy modification, carboxy modification, carbinol modification, methacrylic modification, mercapto modification, or phenol modification, or a modified product that has been modified to impart non-reactive functional groups such as polyether modification, methylstyryl modification, alkyl modification, higher fatty acid ester modification, or fluorine modification.

[0028] As a binder used to bind polymer particles together, for example, a polymer solution obtained by dissolving the polymer in an organic solvent or the like, or a precursor of the polymer, can be used. For example, silicone polymers can be obtained by curing a one-component curable silicone or a two-component curable silicone by an addition reaction or a condensation reaction, and such a curable silicone may be used as a binder. The curable silicone may be a low-viscosity liquid fluid that flows by gravity alone without the application of external force at room temperature (25°C), or it may be a high-viscosity paste-like fluid that does not exhibit fluidity by gravity alone.

[0029] By incorporating such a coagulant into the raw materials of the porous body 100 together with polymer particles, it is possible to suppress the scattering of the raw materials of the porous body 100 or their adhesion to unwanted areas due to static electricity, thereby improving the manufacturing efficiency of the porous body 100.

[0030] When a curable silicone is used as a binder, the curable silicone may be an addition-reaction type having a polysiloxane skeleton in its main chain and vinyl groups bonded to the main chain or side chains. When a curable silicone is used as a binder, the curable silicone may be a condensation-reaction type having hydroxyl groups or alkoxy groups instead of vinyl groups in the addition-reaction type curable silicone.

[0031] On the surface of a porous body 100 made using curable silicone and particles of a polymer other than silicone-based polymers, such as polyester particles, both the silicone-based polymer, which is the cured product of the curable silicone, and the polyester particles will be exposed. That is, the surface of the porous body 100 may be formed with a first region composed of silicone-based polymers and a second region formed by the exposure of polyester particles on the surface. While it is usually possible to finely disperse common polymers such as polyester and silicone-based polymers, they do not typically dissolve at the molecular level. As a result, a microphase separation structure consisting of the first region and the second region is observed on the surface of the porous body 100. Depending on the type of polymer other than the silicone-based polymer and the ratio of curable silicone, one of the first and second regions may constitute a continuous phase (matrix), and the other may constitute a dispersed phase (domain).

[0032] Microphase separation on the surface of the porous body 100 as described above can be achieved not only when using curable silicone, but also by forming the porous body 100 with polymer particles containing multiple types of polymers, such as silicone-based polymers like silicone oil and silicone resin, and polymers other than silicone-based polymers. Furthermore, the formation of a first region formed of silicone-based polymer and a second region formed of polymers other than silicone-based polymers on the surface of the porous body 100 can also be achieved, for example, by forming the porous body 100 using multiple types of polymer particles, such as silicone-based polymer particles and polymer particles other than silicone-based polymers.

[0033] The porous body 100 comprises a first polymer, which is a silicone-based polymer, and a second polymer other than a silicone-based polymer. By exposing both the first and second polymers on the surface of the porous body 100, which forms the outer surface 1s1 and inner surface 1s2 of the artificial blood vessel 1, the biocompatibility of each polymer can be expressed, resulting in good biocompatibility. In the porous body 100, it is preferable that the silicone-based polymer (first region) constitutes a dispersed phase, and the second polymer other than a silicone-based polymer (second region) constitutes a continuous phase.

[0034] When using a curable silicone as the raw material for the porous body 100, a condensation-curable silicone that hardens by a condensation reaction is preferable to an addition-curable silicone that hardens by an addition reaction because it is easier to control the reactivity during hardening. Since condensation-curable silicones generate water and alcohol as byproducts in the hardening reaction, it is possible to interpose water and alcohol between the particles during heat molding to suppress excessive fusion between polymer particles, which is advantageous for ensuring the continuity of the voids (forming continuum).

[0035] Even when the first polymer in the raw materials of the porous body 100 is not a cured product of curable silicone but rather a silicone oil, the silicone oil can suppress excessive fusion between polymer particles, which is advantageous in ensuring the continuity of the voids (forming continuum pores).

[0036] The artificial blood vessel 1 of this embodiment can be manufactured by general molding methods such as extrusion molding using an extruder or mold molding using a mold, as will be described later. The artificial blood vessel 1 of this embodiment, which is a porous body 100 with a silicone polymer exposed on its surface, exhibits excellent release properties in extrusion molding and mold molding, and therefore also offers excellent production efficiency.

[0037] The artificial blood vessel 1 of this embodiment can be manufactured by sequentially performing, for example, (S1) a raw material preparation step of preparing raw materials containing polymer particles, (S2) a molding step of producing molded products (corrugated tubes) that will become the main tube and branch tubes of the artificial blood vessel by molding using the raw materials prepared in the raw material preparation step, and (S3) a post-processing step of performing post-processing on the molded products produced in the molding step.

[0038] In the raw material preparation process, for example, raw materials may be prepared using only one type of polymer particle, raw materials containing multiple types of polymer particles, raw materials containing a binder in addition to polymer particles, or raw materials containing silicone oil that functions as a mold release agent in addition to polymer particles.

[0039] Raw materials containing a coagulant can be prepared by mixing polymer particles and the coagulant using a common mixing and stirring device such as a Henschel mixer or ribbon blender. A specific method for this is to place the polymer particles in the mixing and stirring device, stir (fluidize) the polymer particles with a stirring blade or similar device, add a predetermined amount of coagulant, and continue stirring to uniformly disperse the coagulant. The coagulant can be added using a feeder or spray nozzle. It is preferable to spray the coagulant onto the polymer particles, as this makes it easier to disperse the coagulant relatively uniformly.

[0040] Raw materials containing a release agent can also be prepared using general mixing and stirring equipment such as a Henschel mixer or ribbon blender to prepare raw materials containing polymer particles. A specific method in this case is to place the polymer particles in the mixing and stirring equipment, stir (fluidize) the polymer particles with a stirring blade or the like while adding a predetermined amount of release agent, and then continue stirring to uniformly disperse the release agent. The release agent can be added using a feeder or spray nozzle. It is preferable to spray the release agent onto the polymer particles because it is easier to disperse the release agent relatively uniformly.

[0041] The polymer particles used as raw materials can be, for example, those having an average particle diameter of 5 μm or more and 500 μm or less. The average particle diameter of the polymer particles may be, for example, 10 μm or more, or may be 20 μm or more. The average particle diameter of the polymer particles may be, for example, 300 μm or less, or may be 100 μm or less. The average particle diameter of the polymer particles can be determined, for example, as the median diameter (D50) based on volume in the laser diffraction scattering method.

[0042] The shape of the polymer particles used as raw materials may be, for example, spherical, needle-like, plate-like, or irregular. The polymer particles are preferably spherical or irregular in terms of being advantageous for the formation of the void portion V, and more preferably irregular.

[0043] The molding process may be carried out, for example, by dividing it into a primary molding for molding the raw material into a straight tubular body and a secondary molding for molding the tubular body obtained in the primary process into a bellows shape. As such a method, a method using an apparatus similar to the manufacturing apparatus for corrugated tubes used for flexible electric wires or the like can be considered. Specifically, for the corrugated tubes constituting the main pipe and the branch pipes, for example, a primary molding in which the raw material is extruded from an extruder having an annular discharge port so as to be straight tubular, and a secondary molding in which a bellows shape is imparted by a corrugating machine provided immediately after the discharge port of the extruder are carried out, and the obtained long corrugated tube can be cut into a predetermined length for manufacturing.

[0044] The corrugated pipe forming the main pipe and branch pipes can be manufactured by using a molding die instead of the above method. When the corrugated pipe is a spiral corrugated pipe, as shown in FIGS. 3a and 3b, a molding die M1 having a thread groove-shaped molding surface MP corresponding to the shape of the outer surface of the corrugated pipe, and a core pin (inner die M2) which is an inner die slightly smaller in diameter than the cavity MV which is the space defined by the molding surface MP of the molding die M1 and has a thread crest-shaped surface shape facing the molding surface MP can be used for manufacturing. In this method, the core pin (inner die M2) is set in the cavity MV of the molding die M1 so that a molding space corresponding to the corrugated pipe is formed between the molding surface MP and the core pin (inner die M2), the raw material 100' containing polymer particles is accommodated in the molding space, and the corrugated pipe can be manufactured by heating the raw material through the molding die M1 or the core pin (inner die M2). The molding die M1 shown in FIGS. 3a and 3b can be made up of a plurality of split dies with a plane containing the central axis CX of the cavity MV as the mating surface, making it easy to take out the formed corrugated pipe. Also, the core pin (inner die M2) can be axially withdrawn by rotating relative to the manufactured corrugated pipe around the central axis CX.

[0045] When the corrugated pipe forming the main pipe and branch pipes is a ring-shaped corrugated pipe, using the previous molding die M1 as the outer die, an inner die M2 which is a split die as shown in FIGS. 4a, 4b, and 4c and is smaller than the outer die by the thickness of the corrugated pipe is formed, the inner die M2 is set in the cavity MV, and the raw material of the corrugated pipe is filled in the gap between the inner die M2 and the outer die for molding and manufacturing. Incidentally, regarding this inner die M2, it can also be made up of split dies M21, M22, M23, M24, M25 as shown in FIGS. 4a, 4b, and 4c, making it easy to take out the inner die M2 from the formed corrugated pipe.

[0046] The corrugated pipe can also be produced, for example, by powder injection molding.

[0047] In this molding process for producing corrugated tubes from raw materials containing polymer particles, the shape and volume ratio of the voids V can be adjusted by controlling temperature and pressure conditions, thereby preparing the corrugated tubes to be suitable for early integration with biological tissue.

[0048] In the post-processing step after the molding process, the artificial blood vessel 1 is fabricated by drilling holes for branching to the branch pipes 12 in the corrugated pipe for the main pipe 11 manufactured in the molding process, or by connecting the corrugated pipe for the branch pipes 12 to these holes. Various methods can be used to connect the corrugated pipes, such as bonding with adhesive, heat fusion, and ultrasonic welding. Generally, it is not easy to bond silicone-based polymers with high adhesive strength. Therefore, as mentioned above, forming a continuous phase on the surface of the corrugated pipe with a polymer other than a silicone-based polymer makes it easier to connect the main pipe 11 and the branch pipes 12.

[0049] Thus, in this embodiment, an artificial blood vessel 1 with excellent biocompatibility that can integrate with biological tissue at an early stage can be easily fabricated.

[0050] Next, an embodiment different from the artificial blood vessel 1 described in the first embodiment will be described with reference to Figure 5. (Second Embodiment) Figure 5 illustrates biocompatible components used as constituent members of the vascular access device 2. The vascular access device 2 illustrated in Figure 5 is an in-vivo implantable device used by being implanted in the body (biological LO) of the device wearer, and more specifically, it is a subcutaneous implantable device used by being implanted under the epidermis SK of the device wearer.

[0051] The vascular access device 2 comprises an access port 21, which is a container for storing a drug solution to be supplied to the device wearer's blood vessel BV (more specifically, a vein), and a catheter 22, which is a highly flexible tube. The vascular access device 2 is able to supply the drug solution to the device wearer's blood flow by connecting the storage space 21a of the access port 21, which contains the drug solution, with the inside of the blood vessel BV via the catheter 22.

[0052] The access port 21 is used by being embedded subcutaneously, with its position in the depth direction D1 determined so that the entire access port is not exposed from the body surface, when the direction normal to the epidermis SK (body surface) of the device wearer is defined as the depth direction D1, and the direction perpendicular to the depth direction D1 is defined as the planar direction D2. The housing space 21a of the access port 21 is a flattened cylindrical shape that extends a short distance in the depth direction D1, and its dimensions in the depth direction D1 are shorter than its dimensions (diameter) in the planar direction D2.

[0053] The access port 21 has a bottomed cylindrical housing 211 that defines the bottom and side edges of the accommodation space 21a. The housing 211 is positioned so that its upper opening faces the body surface of the device wearer. The access port 21 further comprises a disc-shaped septum 212 that closes the upper opening of the housing 211 and defines the upper edge of the accommodation space 21a. In the vascular access device 2 illustrated in the figure, the septum 212 is fitted inside the opening of the housing 211.

[0054] The access port 21 is embedded subcutaneously with the upper opening of the housing 211 facing the body surface of the device wearer, and has a nozzle portion 21b extending in a planar direction at its bottom, at which the catheter 22 is connected. In other words, the catheter 22 is connected to the access port 21 at one end in the longitudinal direction, and the other end opposite to the one end is inserted into the blood vessel BV, and the catheter 22 connects the blood vessel BV and the containment space 21a, forming a flow path for the drug solution contained in the containment space 21a to the blood vessel BV.

[0055] In this embodiment, the housing 211 comprises an inner housing 211a that defines the containment space 21a and constitutes the part that comes into contact with the liquid medicine, and an outer housing 211b that is arranged to cover the bottom of the inner housing 211a. The opening into which the septum 212 is fitted has a single-layer structure of the inner housing 211a, while the bottom of the nozzle portion 21b has a double-layer structure of the inner housing 211a and the outer housing 211b.

[0056] The upper end of the inner housing 211a, which constitutes the opening of the housing 211, is provided with a circumferential groove for fitting the septum 212. Specifically, the inner housing 211a is provided with a larger diameter portion 2111 located deeper than the opening edge 211e, which is larger in diameter than the opening diameter at the opening edge, and is provided with a circumferential groove into which the septum 212 can be fitted.

[0057] The vascular access device 2 is configured to allow the supply of a drug solution to the access port 21 from outside the body, and the septum 212 is made of a material through which an injection needle can be inserted. The septum 212 constitutes the needle-insertion portion into which an injection needle for supplying a drug solution to the vascular access device 2 is inserted, and is a porous body with excellent elastic deformability composed of a polymer particle aggregate, similar to the artificial blood vessel 1 in the first embodiment.

[0058] The septum 212 will be described in detail below as a biocompatible member in the second embodiment, but the biocompatible member composed of a polymer particle aggregate is not limited to the septum 212, and may also be an inner housing 211a, an outer housing 211b, a catheter 22, etc.

[0059] The septum 212 is positioned such that the direction of insertion of the injection needle is in the thickness direction. The front surface of the septum 212, relative to the direction of insertion of the injection needle, is the biological contact surface that comes into contact with the biological LO, while the opposite end surface, in the direction of insertion of the injection needle, is the drug solution contact surface that comes into contact with the drug solution supplied by the injection needle.

[0060] Furthermore, the septum 212 constituting the needle insertion site of the vascular access device 2 is required to have elastic deformation recovery that can close the needle mark (through hole) after puncture, and to have self-occluding properties after puncture. The self-occluding properties of the septum 212 can be confirmed, for example, by applying water pressure (e.g., 100 mm hydrohead pressure) from the biological contact surface side to the site after withdrawing an injection needle that has been inserted so as to penetrate in the thickness direction, and confirming that water does not pass through the puncture mark to the drug solution contact surface side.

[0061] The self-occluding properties of septum 212 may be exhibited, for example, with a 16G diameter needle or a thinner needle at room temperature (25°C). The self-occluding properties of septum 212 may also be exhibited, for example, with a 23G diameter or larger needle at room temperature (25°C). The self-occluding properties of septum 212 may also be exhibited with a 22G diameter or larger needle, a 21G diameter or larger needle, a 20G diameter or larger needle, a 19G diameter or larger needle, or a 18G diameter or larger needle.

[0062] The porous material constituting the septum 212 preferably exhibits the above-described self-occluding properties at a thickness of 3.0 mm, more preferably at a thickness of 2.5 mm, even more preferably at a thickness of 2.0 mm, and particularly preferably at a thickness of 1.5 mm.

[0063] In this embodiment, the septum 212 can, for example, have a diameter larger than the opening edge 211e of the inner housing 211a when viewed in the depth direction, and less than or equal to the diameter of the enlarged portion 2111 of the inner housing 211a. The septum 212 may also have a diameter larger than the diameter of the enlarged portion 2111 of the inner housing 211a into which it is fitted. In that case, the septum 212 will receive a pressing force from the inner housing 211a toward its radial center, thereby more reliably exhibiting self-closing properties.

[0064] If the circumference of the large diameter portion 2111 of the inner housing 211a is "L1 (mm)", and the circumference of the septum 212 at the portion in contact with the large diameter portion 2111 (circumference in its natural state, not compressed by the inner housing 211a) is "L2 (mm)", and the ratio of the circumference of the septum 212 (L2) to the circumference of the large diameter portion 2111 (L1) (L2 / L1) is defined as the compression ratio, then the septum 212 can have a size that allows it to be fitted into the inner housing 211a with a compression ratio of, for example, 1.01 or higher. The septum 212 may also have a size that allows it to be fitted into the inner housing 211a with a compression ratio of, for example, 1.02 or higher, or 1.05 or higher, or 1.10 or higher. The septum 212 may have a size that allows it to be fitted into the inner housing 211a with a compression ratio of 1.30 or less.

[0065] The septum 212 is the same as the biocompatible member in the first embodiment in that it has voids that open to the biological contact surface and the drug solution contact surface. The preferred ratio of the voids to the polymer portion and the constituent materials, such as polymer particles and coagulants, are also the same as the biocompatible member in the first embodiment, so a detailed explanation will not be repeated here. Furthermore, the fact that the septum 212 has a complex three-dimensional network of voids extending in the thickness direction from the biological contact surface facilitates early integration with biological tissue, which is also the same as described in the first embodiment, so a detailed explanation will not be repeated here. In addition, the septum 212 can be manufactured by methods such as mold molding and powder injection molding, which is also the same as the artificial blood vessel 1, so a detailed explanation will not be repeated here.

[0066] (Other Embodiments) The need for early integration between the contacting biological tissue and the component is not limited to this vascular access device 2 or the aforementioned artificial blood vessel 1. Other applications of biocompatible components composed of porous materials, which are aggregates of polymer particles, include, for example, artificial skin.

[0067] Although the above provides a detailed explanation of biocompatible components with specific examples, the present invention is not limited in any way to the above-mentioned examples. Furthermore, this specification includes the following disclosures.

[0068] (1) A biocompatible member in which at least a portion of the surface is composed of a porous body, wherein the porous body is a aggregate of polymer particles.

[0069] (2) The biocompatible member according to (1), wherein the porous body is a sintered body of polymer particles.

[0070] (3) The biocompatible member according to (1) or (2), wherein the porous body comprises a first polymer which is a silicone polymer and a second polymer which is not a silicone polymer, and both the first polymer and the second polymer are exposed on the surface.

[0071] The porous body comprises one or more polymer particles selected from the group consisting of polyethylene particles, polypropylene particles, polyester polymer particles, polystyrene polymer particles, polyether ether ketone particles, and polyurethane particles, according to any one of (1) to (3).

[0072] The present invention will now be described in more detail with reference to examples, but the present invention is not limited to these examples.

[0073] To evaluate the integration performance of biocompatible materials with biological tissues, implantation tests were conducted in rats. The samples prepared for the implantation tests were as follows:

[0074]

[0075] Each sample was sterilized using EO (electrolysis) and then implanted in a rat. Samples 1-1 and 2-4 were both disc-shaped samples with a diameter of 20 mm and a thickness of 3.5 mm, made using the same ePTFE. However, as shown in Figure 6, the implantation positions in the rats were different. The same applies to samples 3-1 and 4-4. Samples 3-4 and 4-1 were subjected to the same implantation test as samples 1-1 and 2-4, except that the sterilization method was autoclave sterilization. Samples 1-2 and 2-3 were prepared by sewing polyester threads onto the ePTFE disc-shaped samples used in sample 1-1, with different polyester threads being used for samples 1-2 and 2-3. Sample 1-4 was prepared by compressing the ePTFE to reduce the volume ratio of the voids compared to samples 1-1 and others. Sample 3-2 was a porous sintered sheet (porosity approximately 60%, aperture diameter approximately 20 μm) made by molding ultra-high molecular weight PE particles in a mold. A disc-shaped sample with a diameter of 20 mm and a thickness of 3 mm was prepared. Sample 4-2 was tested in the same manner as sample 3-2, except that the sample thickness was changed from 3 mm to 2 mm and the embedding position was changed as shown in Figure 6. Sample 3-3 was a porous sintered sheet (porosity approximately 60%, aperture diameter approximately 20 μm) made by molding polyester polymer particles in a mold. A strip-shaped sample with a length of 20 mm, a width of 5 mm, and a thickness of 3 mm was prepared. This sample was prepared by adjusting the material to contain a small amount of silicone oil. Sample 3-3 was prepared by first creating a thick sample and then cutting the thick sample to the above thickness. At that time, the cut surface was heated to harden it.

[0076] For sample 3-3, the cross-section was observed using a scanning electron microscope (SEM). The obtained SEM image is shown in Figure 7a. From this figure, it was observed that in sample 3-3, which is a sintered body of polymer particles, the particles are joined together to form a three-dimensional network structure, and large voids with a wide aperture area are intricately interwoven in the depth direction. Furthermore, Si element mapping was performed in the same field of view using energy-dispersive X-ray fluorescence spectroscopy (EDX). The obtained mapping image is shown in Figure 7b (white areas indicate the presence of Si element). From this mapping image, a microphase separation structure was observed on the surface of sample 3-3, and it was found that a silicone-based polymer constitutes the dispersed phase and a polyester-based polymer resin constitutes the continuous phase.

[0077] Each of the samples described above was implanted in rats at the location shown in Figure 6. After 28 days, the samples were removed from the rats, and observation specimens were prepared from the removed samples.

[0078] (Sample Preparation) Each specimen was cut from approximately the center of the sample in a head-to-tail direction (except for #3-3, which was cut in a left-to-right direction), and a cross-sectional image was taken (Figure 8). Since the specimens were not yet fixed, they were re-fixed with NBF, and then dehydrated using a dehydration and infiltration apparatus (Tissue-Tek V.I.P 6 AI; Sakura Finetek Japan CO. LTD, Tokyo, Japan) in an increasing concentration series of 70-100% ethanol, replaced with xylene, and then infiltrated with paraffin Parabet 60 GR (#43257; Muto pure chemicals CO. LTD, Tokyo, Japan) with a melting point of 58-60°C to create paraffin-embedded blocks. Using a sliding microtome (LS113; YAMATO-KOHKI industrial CO. LTD, Asaka, Saitama, Japan), thin sections were prepared at a set thickness of 4 mm from this block. These sections were then mounted on standard slide glass (#5116; Muto pure chemicals) coated with 3-aminopropyltrimethoxysilane (APS) as defined in Japanese Industrial Standards (JIS R 3703 / ISO 8255-1), dried overnight at 37°C, and then stained with hematoxylin-eosin (HE).

[0079] (Observation and Image Acquisition) A COOLPIX B600 (NIKON Corporation, Tokyo, Japan) was used to take cross-sectional images of the test specimens. The prepared specimens were observed with an optical microscope BX-43 (OLYMPUS Corporation, Tokyo, Japan), and the infiltration into the sample and signs of inflammation in the surrounding tissue were mainly evaluated. Infiltration into the sample was classified into tissue infiltration accompanied by tissue structures such as fibers and blood vessels, and cellular infiltration consisting only of free cells. In the display of the results, the word "fiber" was used for bio-derived fibers and "fiber" for artificial fibers (descriptions regarding other parts and directions are shown in Figure 9a (overall image of sample 1-1)). Microscope images were taken using an imaging device DP-22 (OLYMPUS). Whole images of the specimen were created by tiling and combining 20x magnified microscope images using the Multiple Image Alignment (MIA) function of CellSense (OLYMPUS). (Results) <Sample 1-1: ePTFE (control)> Sample 1-1 was a stretched PTFE (ePTFE) porous body implanted directly beneath the muscular layer of the skin and was used as a control sample. It was slightly curved toward the epidermis and the caudal end was somewhat rounded as if the corners had been trimmed, but there was no strong distortion or rupture in shape, and the rectangular cross-section of an almost disc-shaped sample was maintained (Figure 9a). The area around the sample was covered with a thin fibrous tissue with a relatively dense tissue density, mainly composed of fibroblasts, and capillary proliferation was observed on the outside of this, but no strong inflammatory cell infiltration, granulation tissue formation, hemorrhage, or calcification was observed in the surrounding tissue. Furthermore, on the cranial (left and right) short sides of the sample (hereinafter referred to as the "vertical ends"), a very small number of macrophages, including multinucleated giant cells, were observed lined up along the ePTFE fiber ends, and fibrous infiltration accompanied by fibrous components (presumably collagen fibers) was observed in some areas, eroding the sample (Figure 9b: arrow). In other areas, including the epidermal-deep (upper and lower) long sides of the sample (hereinafter referred to as the "horizontal ends"), cell infiltration (exudation) mainly consisting of lymphocytes was observed over a wide area, seeping from the surrounding tissue into the lattice-like spaces of the ePTFE fibers, but these cell infiltrations did not extend to the center of the sample (Figures 9b, 9c, 9d, and 9e).

[0080] <Sample 1-2: ePTFE-Polyester Thread Suturing> In Sample 1-2, similar to Sample 1-1, the implanted sample beneath the muscular layer of the skin was surrounded by fibrous tissue. The fibrous tissue appeared slightly thicker than in Sample 1-1 (especially in the area directly beneath the muscular layer), but no strong inflammatory cell infiltration, granulation tissue formation, or hemorrhage was observed in the surrounding tissue. The sample was slightly curved towards the epidermis and had three large ruptures at its horizontal end (Figure 10a). At two of these ruptures, closer to the craniocaudal (left and right) sides, bundles of polyester threads accompanied by macrophages were observed on both the upper and lower sides of the sample, and polyester fibers were also observed within the ePTFE fibers (Figure 10b: Polyester fibers are labeled "PET"). In addition, bundles of polyester fibers accompanied by macrophages were observed in several places, appearing to penetrate the ePTFE fibers from the epidermal-deep (upper and lower) side (Figure 10e: Polyester fibers are labeled "PET"). On the other hand, in the slightly larger fracture in the center of the horizontal end, no polyester fibers were observed. Instead, a somewhat edematous and sparse fibrous tissue accompanied by capillaries infiltrated and proliferated, almost bridging the ePTFE fibers from the epidermal-deep (upper and lower) sides. However, strong inflammatory cell infiltration and granulation tissue formation were not observed (Figures 10c and 10d). Furthermore, macrophages and lymphocytes, including a small number of multinucleated giant cells (Figure 10e: arrow), infiltrated somewhat broadly into the voids within the sample (Figure 10e). In other areas, cell infiltration from the surroundings into the ePTFE fibers was also observed, similar to sample 1-1, but deep infiltration into the central part of the sample, excluding the fractured area, was scarce.

[0081] <Sample 1-3: ePTFE-polyester thread sutured - low cleanliness> Sample 1-3 was located beneath the dermal muscle layer and surrounded by a thin fibrous tissue. It curved slightly toward the epidermis and was slightly compressed and indented near the center of the horizontal end (Figure 11a). This indented area appeared to be slightly compressed by the surrounding tissue, but there was no rupture of the ePTFE fibers due to deep fibrous tissue infiltration from the surrounding tissue, as seen in sample 1-2. Furthermore, there was no significant inflammatory cell infiltration, granulation tissue formation, or hemorrhage in the surrounding tissue. In sample 1-3, similar to sample 1-2, bundles of polyester thread were seen to be embedded in the ePTFE fibers from above and below (Figure 11b: polyester fibers are labeled "PET"), and polyester fibers were also seen within the ruptures (Figure 11c: polyester fibers are labeled "PET"), but the number and size (degree) of ruptures were fewer than in sample 1-2. Slight fibrous infiltration was observed at the cranial vertical end (Figure 11d: arrow), and mild cellular infiltration, mainly lymphocytes including some macrophages, was observed in the surrounding tissue on all sides, including between polyester fibers and ePTFE fibers, and in the voids within the ePTFE fibers (Figure 11e: polyester fibers are labeled "PET"), but the infiltration depth hardly extended to the center of the sample.

[0082] <Sample 1-4: ePTFE - Low porosity> In sample 1-4, the horizontal end (especially the epidermal side) of the ePTFE sample implanted beneath the dermal muscle layer was irregularly and significantly indented, as if partially compressed, resulting in an irregular shape (Figure 12a). Within the ePTFE fibers, layers with different porosity densities appeared to be stacked in a striped pattern in the direction of compression (Figures 12b, 12c). The outer periphery of the sample was covered with a thin fibrous tissue, and mild capillary proliferation was observed in the tissue outside of this, but no strong inflammatory cell infiltration, granulation tissue formation, or hemorrhage was observed in the surrounding tissue. Strong fibrous proliferation was not observed in the indented area on the epidermal side, and mild edema was observed in a narrow area (Figures 12b, 12c, 12d: arrows). Mild cellular infiltration, mainly lymphocytes, was observed from the surrounding tissues, seeping into the voids within the ePTFE fibers, but the infiltration did not extend far into the center of the sample (Figure 12e). Cellular infiltration from the epidermal tissues, in particular, was minimal.

[0083] <Sample 2-1: ePTFE - Low porosity - Different implantation location from Sample 1-4> Sample 2-1, implanted beneath the dermal muscle layer, showed two large depressions at the horizontal end on the epidermal side, similar to Sample 1-4 (Figure 13a). Within the ePTFE fibers, layers with different porosity densities appeared to be stacked in a striped pattern in the direction of compression (Figures 13b, 13c). The area surrounding the sample was covered with fibrous tissue that was slightly thicker than that of Sample 1-4, and capillary proliferation was observed at its outer edge, but no strong inflammatory cell infiltration, granulation tissue formation, or hemorrhage was observed in the surrounding tissue. Unlike Sample 1-4, Sample 2-1 did not show an edematous tendency in the tissue surrounding the depressions, and although there was some fibrous tissue proliferation, there was no strong fibrous tissue infiltration from the surrounding tissue. Furthermore, macrophages were observed in a thin line along the edge of the depression, but cellular infiltration into the ePTFE fibers was limited (Figure 13d). On the other hand, on the opposite side of the epidermis (i.e., the side without the depression), cellular infiltration, mainly lymphocytes, was observed, seeping from the surrounding tissue into the spaces within the ePTFE fibers (Figure 13e).

[0084] <Sample 2-2: ePTFE-polyester sutured - low cleanliness - different implantation location from Sample 1-3> Sample 2-2, implanted beneath the dermal muscle layer, was surrounded by relatively thin fibrous tissue. However, near the center of the horizontal end, it was fragmented (or, in other words, partially replaced by fibrous tissue) over a relatively wide area by loose fibrous connective tissue with capillaries. This area was the second largest among the samples in this study, after Sample 4-3 (Figures 14a and 14b). Within this fibrous connective tissue, fragmentary clumps of ePTFE fibers were observed, resembling a sandbar in a river mouth whose edges have been rounded by water flow. Some macrophages, lined up around these clumps, infiltrated the surrounding tissue along with the fibrous tissue containing capillaries, fragmenting the ePTFE fibers (Figure 14d: arrow). However, no necrosis, hemorrhage, strong granulation tissue formation, or inflammatory cell infiltration was observed. On the other hand, at the head and tail ends (left and right) of the sample, bundles of polyester fibers were observed sandwiching the ePTFE fibers from both above and below, and were embedded within the ePTFE fibers (Figure 14c). This was accompanied by infiltration of fibrous tissue that extended continuously from the tissue between the polyester fibers and the ePTFE fibers (Figure 14e). In addition, cell infiltration from the surrounding tissue was observed in other areas.

[0085] <Sample 2-3: ePTFE - polyester thread sutured - different polyester thread from Sample 1-2> In Sample 2-3, the ePTFE sample implanted beneath the dermal muscle layer was slightly curved and surrounded by fibrous tissue, but no bleeding, strong granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue (Figure 15a). Similar to Sample 1-3, bundles of polyester thread were observed at the horizontal ends, sandwiching the ePTFE fibers from both above and below, compressing the ePTFE fibers, and some of the threads were embedded within the ePTFE fibers (Figure 15b). However, the polyester fibers had a clearly thinner cross-sectional diameter compared to Sample 1-2 and Sample 2-2, and there were few findings indicating foreign body reactions such as macrophage infiltration around the fibers. In addition, infiltration of fibrous tissue accompanied by capillaries into the ePTFE fibers was observed from the surrounding tissue near the polyester fiber bundles (Figure 15c: polyester fibers are labeled "PET"). This fibrous tissue infiltration progressed deep into the ePTFE fibers by tearing them, and also infiltrated finely into the interior from the sides of the ePTFE fibers (Figure 15d: polyester fibers are labeled "PET", Figure 15e), and fragmented ePTFE fibers were also observed (Figure 15d: arrows).

[0086] <Sample 2-4: ePTFE - Different implantation location from Sample 1-1> Similar to Sample 1-1, Sample 2-4 was implanted beneath the dermal muscle layer and was surrounded by a thin layer of fibrous tissue. It curved slightly toward the epidermis, and one side of its vertical end was somewhat rounded, as if the corner had been shaved off. However, no bleeding, strong granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue (Figure 16a). However, near the center of the horizontal end of the sample on the epidermal side, it was depressed due to compression by the surrounding fibrous tissue, and striped density differences due to compression were observed within the ePTFE fibers, similar to Sample 1-4 (Figure 16b). At the vertical end of the sample, there were areas where a portion of the surrounding fibrous tissue had slightly infiltrated the ePTFE fibers in a notched or wedge-like manner (Figure 16c: arrow), and areas where a portion of the ePTFE fibers were finely divided by thin fibrous tissue along with cell infiltration into the fibers (Figure 16d: arrow). Furthermore, macrophages were observed along the ePTFE fibers in the compressed depressions at the horizontal ends, but no multinucleated giant cells were found, and cell infiltration into the ePTFE fibers was minimal (Figure 16e).

[0087] <Sample 3-1: ePTFE - same as sample 1-1 (control)> Sample 3-1, implanted beneath the dermal muscle layer, was surrounded by fibrous tissue and slightly curved toward the epidermis. However, it resembled sample 2-4 more than sample 1-1, exhibiting a depression due to compression near the center of the horizontal end on the epidermal side and a striped pattern due to differences in void density within the ePTFE fibers (Figure 17a). Microscopic findings were also similar to sample 2-4, with very slight incision-like fibrous tissue infiltration at the vertical end (Figure 17b: arrow) and rounded corners, and a small number of macrophages were observed along the depression at the horizontal end (Figure 17c). Cellular infiltration, mainly lymphocytes, from the surrounding tissue into the ePTFE voids was observed at both horizontal ends, but the degree was slightly stronger than in sample 2-1 and weaker than in sample 1-1 (Figures 17d, 17e). No bleeding, severe granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue, including the fibrous tissue enclosing the sample and the capillary proliferation area outside of it.

[0088] <Sample 3-2: Agglomeration of PE particles> Sample 3-2, implanted beneath the muscular layer of the skin, was completely encased in fibrous tissue with a relatively high tissue density. No warping or rupture was observed in the sample, and no significant deformation was seen at either the horizontal or vertical ends (Figure 18a). Furthermore, no bleeding, strong granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue. Unlike the ePTFE fibers in the groups of Samples 1-1 to 1-4 and Samples 2-1 to 2-4, which had a network of voids, the internal structure of Sample 3-2 consisted of small vacuoles, presumably traces of compressed powder, clustered in a foamy manner throughout the sample, but no obvious foreign matter remained. In the gaps between these foamy vacuoles, macrophages containing multinucleated giant cells, lymphocytes, and fibrous tissue containing capillaries were finely infiltrated throughout the sample from the surrounding tissue, and relatively large granulation tissue formation was observed near the center of the sample (Figures 18b, 18c, 18d, and 18e). Capillary congestion was observed, but no severe tissue bleeding or necrosis was found.

[0089] <Sample 3-3: Agglomeration of Silicone-Added Polyester Particles> Sample 3-3, implanted directly beneath the muscular layer of the skin, compressed the muscular layer at the implantation site, causing it to become slightly narrower. The sample was surrounded by fibrous tissue with a relatively high tissue density, and was divided almost exactly in the middle of the horizontal end by thin fibrous tissue continuous with this tissue, appearing as two rectangular cross-sections (Figures 19a and 19b). Furthermore, this sample appeared to have relatively poor flexibility, and a fairly wide gap was present between it and the fibrous tissue (Figure 19c). In particular, the surrounding tissue was sparse outside the fibrous tissue on the horizontal end side, and capillary proliferation was less pronounced compared to sample 3-2, but strong granulation tissue formation and inflammatory cell infiltration were not observed. The internal structure of the sample showed no obvious foreign matter, similar to sample 3-2. Vacuoles, presumably traces of compressed powder, were observed throughout in a foamy manner. Fibrous tissue, accompanied by macrophages, lymphocytes, and capillaries, infiltrated the gaps between the foamy vacuoles more continuously and finely than the surrounding tissue. However, compared to sample 3-2, the fused vacuolar portions were larger, resulting in a higher area ratio of vacuoles. Consequently, the formation of relatively large granulation tissue within the sample, as seen in sample 3-2, was poor (Figures 19d and 19e).

[0090] <Sample 3-4: ePTFE - Different sterilization method from sample 3-1> Sample 3-4, implanted beneath the dermal muscle layer, was slightly curved toward the epidermis, and a slight depression was observed at the center of the horizontal end, indicating compression from both the top and bottom (Figure 20a). In the depressed area, the voids within the ePTFE fibers appeared to be compressed, resulting in an irregularly high fiber density, but no tissue infiltration from the surrounding tissue was observed (Figures 20c, 20d). The sample was covered with fibrous tissue, and slight proliferation of capillaries was observed on the outside, but no bleeding, strong granulation tissue formation, or inflammatory cell infiltration was observed. Slight thickening of the surrounding tissue was observed at the vertical end of the sample, and very slight fibrous tissue infiltration was observed at the ends of the ePTFE fibers (Figure 20b). Mild cell infiltration, mainly lymphocytes, was observed from the horizontal end (Figure 20e).

[0091] <Sample 4-1: ePTFE - Different implantation location from Samples 3-4> Sample 4-1, implanted beneath the dermal muscle layer, was slightly curved toward the epidermis and its four corners were slightly rounded, but no compression of the outer edge, striped findings due to differences in void density, or ruptures were observed (Figure 21a). The sample was surrounded by fibrous tissue, and slight proliferation of capillaries was observed outside of this, but no hemorrhage, strong granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue. At the vertical end, the fibrous tissue was slightly thickened, and the ends of the ePTFE fibers were slightly jagged (Figure 21b). At the horizontal end, the surrounding fibrous tissue was thin, a small number of macrophages were observed along the ePTFE fibers, and cell infiltration mainly consisting of lymphocytes was observed in the voids within the ePTFE fibers (Figures 21c, 21d, and 21e).

[0092] <Sample 4-2: Agglomeration of PE particles - Different thickness from sample 3-2> Sample 4-2, implanted beneath the muscular layer of the skin, was completely encased in fibrous tissue with a slightly higher tissue density, similar to sample 3-2. No warping or rupture was observed in the sample, and no significant deformation was seen at either the horizontal or vertical ends (Figure 22a). Furthermore, no bleeding, strong granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue, including the fibrous tissue and the mildly proliferated capillaries outside it. The internal structure of the sample was also almost identical to that of sample 3-2, with small, foamy vacuoles, presumably traces of compressed powder, observed throughout, but no obvious foreign matter was found. In the spaces between these foamy vacuoles, fibrous tissue containing macrophages including multinucleated giant cells, lymphocytes, and capillaries was finely infiltrated throughout the sample, continuous with the surrounding tissue. However, compared to sample 3-2, large granulation tissue was scarce (Figures 22b, 22c, 22d, 22e). Mild congestion was observed in some capillaries, but no severe bleeding or necrosis was seen.

[0093] <Sample 4-3: Agglomeration of PE particles - Different implantation location from sample 3-2> Sample 4-3, implanted beneath the dermal muscle layer, was relatively widely separated at the center of its horizontal end by sparsely fibrous connective tissue with capillaries and multinucleated giant cells, resulting in a cross-section that was separated into two pieces, cranial and caudal (left and right) (Figure 23a). The entire sample was covered with a somewhat dense, capsular fibrous tissue, but this was absent in the broad, sparse fibrous connective tissue separating the two, which differed it from samples 2-2 and 3-3 (Figure 23b). The absence of this capsular fibrous tissue suggested that the sample was not originally a single piece that was split into left and right halves, but rather that it was originally separate and lacked continuity, meaning there was some degree of gap between the two and it was mobile. In any case, no bleeding, strong granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue (Figure 23c). The internal structure of both samples was almost identical to that of sample 3-2, and no obvious foreign matter remained. However, foamy vacuoles, which appeared to be traces of compressed powder, were observed throughout. In the spaces between the foamy vacuoles, macrophages containing multinucleated giant cells, lymphocytes, and fibrous tissue containing capillaries were finely infiltrated from the sparse fibrous connective tissue and the surrounding capsule-like fibrous tissue. However, there was no formation of relatively large granulation tissue as in sample 3-2, and tissue infiltration was less pronounced in the central part of the sample (Figures 23d and 23e).

[0094] <Sample 4-4: ePTFE - Different implantation position from sample 3-1> Sample 4-4 was closer to sample 3-1 than sample 1-1, and like the other ePTFE samples, the sample implanted beneath the dermal muscle layer was slightly curved towards the epidermis, with a depression observed near the center of the horizontal end on the epidermal side. The sample was covered with a membrane-like fibrous tissue, and the fibrous tissue was slightly thinner at the vertical end than in sample 3-1 (Figure 24b). However, no bleeding, strong granulation tissue formation, or inflammatory cell infiltration was observed in the surrounding tissue, including the fibrous tissue and the mild capillary proliferation area outside of it (Figure 24a). Similar to sample 3-1, the interior of the sample showed a striped pattern due to differences in void density in the depressed area (Figure 24c), but some macrophages observed along the surface of the depressed area had slightly infiltrated into the ePTFE (Figure 24d: arrow). Furthermore, while a slightly higher number of macrophages were observed along the horizontal edges of the ePTFE, cellular infiltration into the ePTFE fibers from there was mainly lymphocytes, which were relatively scattered and only found within the voids (Figure 24e).

[0095] As described above, while cell infiltration in ePTFE is limited to the surface, when polymer particle aggregates are implanted, cell infiltration is observed even within the tissue, demonstrating superior integration with biological tissue. Therefore, it can be understood that the present invention can provide a biocompatible material that easily integrates with biological tissue.

[0096] 1: Artificial blood vessel 1s1: Outer surface 1s2: Inner surface 2: Vascular access device 11: Main vessel 11L, 12L: Large diameter section 11S, 12S: Small diameter section 11a: One end 11b: Other end 12: Branch vessel 121: First branch vessel, 122: Second branch vessel, 123: Third branch vessel 21: Access port 21a: Housing space 21b: Nozzle section 22: Catheter 100: Porous material 211: Housing 211a: Inner housing 211b: Outer housing 211e: Opening edge 212: Septum 2111: Large diameter section BV: Blood vessel CX: Central axis D1: Depth direction D2: Planar direction LO: Biological tissue M1: Molding mold (outer mold) M2: Inner mold (core pin) M21, M22, M23, M24, M25: Split type MP: Molded surface MV: Cavity P: Polymer part SK: Skin V: Gap

Claims

1. A biocompatible member in which at least a portion of the surface is composed of a porous material, wherein the porous material is an aggregate of polymer particles.

2. The biocompatible member according to claim 1, wherein the porous body is a sintered body of polymer particles.

3. The biocompatible member according to claim 1 or 2, wherein the porous body comprises a first polymer which is a silicone polymer and a second polymer which is not a silicone polymer, and both the first polymer and the second polymer are exposed on the surface.

4. The biocompatible member according to claim 1 or 2, wherein the porous body comprises one or more polymer particles selected from the group consisting of polyethylene particles, polypropylene particles, polyester polymer particles, polystyrene polymer particles, polyether ether ketone particles, and polyurethane particles.

Images