An Improved Method for Preparing Histological Sections of Metallic Implants C

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An Improved Method for Preparing Histological Sections of Metallic Implants C. Maniatopoulos, D.D.S., M.Sc./A. Rodriguez/D.A. Deporter, D.D.S., Diplo. Perio., Ph.D./A.H. Melcher, M.D.S., H.D.D., Ph.D., D.Sc.

An improved method for preparing ground tissue-implant sections of good quality for light microscopy is presented. The specimen is embedded in methacrylate and cut using a diamond wafering saw. The cut surface is polished and stained with Stevenel's blue and a glass microscope slide is glued to the surface of the specimen using epoxy resin adhesive. A section is prepared, reduced to 30 to 50 µm using petrographic grinding techniques and protected by a glass coverslip. This method provides high quality sections in which fine tissue architecture, cellular detail, and the tissue-implant interface are preserved.

Histological preparations of metallic implants in situ present many technical difficulties, particularly if the implants have a complex surface geometry. Methods have been developed using diamond wafering saws1-22 that permit the obtaining of slices of such specimens. These slices are usually reduced in thickness to about 50 to 100 µm using petrographic grinding techniques.1-22 This thickness permits microscopic examination under transmitted light. Nevertheless, the relatively thick, unstained, or poorly stained ground sections so obtained generally lack cellular detail and do not permit recognition and differentiation of the types of cells and tissues present at the implant site. Moreover, problems related to embedding, grinding, and staining make it difficult to obtain histological sections of predictably good quality and artifacts are common. This report describes an improved method for preparing good quality ground tissue-implant sections. Materials and methods Fixation. The animal should be perfused while under a general anesthetic. The perfusate is a mixture of 37% formaldehyde:99.8% methanol: distilled water (1:1:1.5 v/v). Following perfusion, blocks of tissue are removed and fixed further by immersion in the same fixative. Other fixatives, such as buffered glutaraldehyde or Karnovsky's fixative, followed by the postfixation of tiny pieces in buffered osmium tetroxide, can also be used, particularly where tissues are to be processed for electron microscopy. Formulae for preparing these fixatives are readily available (for example, see Hayat23 and Glauert24). Following fixation, specimens are washed in running tap water overnight. Dehydration. The process of dehydration is accomplished by using an ascending series of ethanol (see below). Bulky specimens, particularly those containing substantial amounts of bone, should be defatted using a mixture of ether and acetone to facilitate infiltration and embedding. Steps in ethanol dehydration 1. 70% aqueous ethanol for four hours to two days (two changes) JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess…

2. 80% aqueous ethanol for four hours to four days (two to three changes) 3. 95% aqueous ethanol for eight hours to four days (two to three changes) 4. Absolute ethanol for eight hours to four days (two to three changes) 5. Defat in a mixture of ether: acetone (1:1 v/v) for sixteen hours to three days (two changes) 6. Clear in 100% acetone or 100% ethanol for eight hours to two days (two changes) Dehydration times given in the above schedule depend on specimen thickness and tissue type. Thus, the shorter dehydration time is adequate for processing of a small bone specimen (about 0.5 cm3), whereas the maximum time is required for processing a very large specimen, such as a hip joint replacement in a canine femur. For smaller specimens, and particularly for implants in soft tissues, dehydration times can be reduced. Also, for small soft tissue specimens, steps 5 and 6 may be omitted. Infiltration and embedding. After dehydration, the specimens are embedded in methylmethacrylate. This permits intact sections of both the implant and the adjacent tissues to be obtained. The method for infiltration and embedding is shown in Table 1. Method 1. Place the specimen in a mold (preferably a wide-mouth glass vial) containing solution A, and allow the solution to infiltrate for 8 to 24 hours at room temperature. 2. Replace the methacrylate solution with fresh solution A. 3. Evacuate the specimen in a dessicator or a vacuum oven to about 35 mm Hg at room temperature for 6 to 18 hours depending on the size of the specimen (see section 2 above). 4. Remove the specimen from the dessicator and replace solution A with solution B. 5. Evacuate the specimen again for 8 to 16 hours. 6. Polymerize the methacrylate in a water bath at 35¡C or at room temperature. Shrinkage of the methacrylate accompanies polymerization, so fresh solution B should be added periodically as required. Methacrylate polymerization time depends on the amount of methacrylate (specimen size), the type of initiator (benzoyl peroxide or Perkadox-16), and the polymerization temperature.28 For example, when 2.5-cm diameter containers are used (block size approximately 10 cm3), polymerization occurs in about two to three days at 35¡C when benzoyl peroxide is used. Or if Perkadox-16 is used polymerization occurs in 24 hours. For 5-cm diameter containers (block size approximately 80 cm3) polymerization time at 35¡C will be in three to four days when benzoyl peroxide is used, or about two days when Perkadox-16 is used. Containers 7.5 cm in diameter (300 cm3 blocks) require polymerization at room temperature for 25 to 30 days or 10 to 14 days for benzoyl peroxide and Perkadox-16, respectively. 7. Once polymerization has been completed, the specimen is removed from the mold and the block trimmed to convenient size using an electric band saw and 80-grit silicon JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess… carbide paper under water lubrication. Specimens processed in this way are well infiltrated, the blocks are hard and transparent, and can be sectioned relatively easily. Sectioning and grinding The implant is located and the block is cut so that the implant is sectioned into symmetrical halves. Sectioning is accomplished using a low-speed, low-deformation saw with a diamond wafering blade. A number of commercial apparatuses are available for this purpose. Implants larger than 1.5 mm in diameter are sectioned using a 12.7 cm diameter 0.3-mm thick diamond wafering blade. For sectioning implants smaller than 1.5 mm in diameter, thinner wafering blades (6.6 cm × 0.15 mm) are available. Cutting is usually performed at a blade speed of 100 to 500 rpm, using tap water as a lubricant, and with a force of 0.3 to 7 N acting on the specimen. It is often difficult to locate accurately the position in the tissues of small implants so that sections can be cut in the desired orientation. In such cases the following method can be used: 1. Drill four holes, each about 2 mm in diameter and 2 mm deep, one in each corner on a flat surface of the block from which the slices are to be cut. 2. Fill the holes with a radiopaque material (e.g., dental amalgam) to produce registration points on the block. 3. Place the block on a regular radiographic plate with the radiopaque markers facing the film, and take a contact radiograph. 4. After developing the radiograph, glue the film to the marked surface of the block with a cyanoacrylate adhesive so that the radiopaque marks on the film are accurately placed on the registration points on the block. 5. Bisect the radiograph and the specimen block using the radiograph as a reference to help obtain the desired orientation. The surface of the exposed specimen is now polished on 600 or 800 grit silicon carbide paper under water lubrication to remove cutting marks. To obtain a highly polished surface, 4,000 grit paper is used. Alumina lapping is not recommended, because it may result in formation of an uneven surface at the tissue-implant interface due to the difference in resistance to grinding of the implant and the tissue. The polished block surface is then stained with Stevenel's blue and Van Gieson or Stevenel's blue and alizarin red S (see the section on staining, below), and relatively thin sections are prepared as follows: 1. A standard microscope glass slide is glued to the stained block surface using a clear epoxy resin adhesive. Due to the high viscosity of these epoxy resin-catalyst systems, air bubbles are readily entrapped in the mixture. To avoid this, the epoxy adhesives are diluted in toluene. This is achieved by thoroughly mixing resin and catalyst in a small vial containing toluene (toluene:resin: catalyst 1:1:1 v/v). A small portion of the mixture is then applied to both the glass slide and the specimen surface, and left for JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess… five minutes before the two surfaces are joined. Full polymerization and bonding strength is achieved after 16 hours. When large specimens are sectioned, use of larger size (i.e., 38 × 75 mm or 50 × 75 mm) glass microslides may become necessary. 2. The block with the glass slide attached is returned to the sectioning machine and is sectioned proximal to the glass slide, so that a section about 0.1 mm in thickness that is attached to the glass slide is obtained. The block surface is then polished and stained again preparatory to cutting the next section. 3. The section, still glued to the microscope slide, is now ground using silicon-carbide paper (180 grit followed by 240 or 320 grit) under water lubrication. Polishing is achieved using 600 grit paper, followed by 800 and 4,000 grits. The sections can be reduced to a thickness of approximately 30 to 50 µm without disruption of the implant-tissue interface. A number of commercial apparatuses for metallographic analysis are available for grinding and polishing of the sections. To handle the slides during grinding, a custom-made holder (Fig. 1) can be used. The use of this holder results in sections of good quality and reproducible thickness. Commercial thin-section polishing holders are also available. 4. After completion of grinding, the section is rinsed in running tap water to remove grinding debris, dehydrated in 95% and then absolute ethanol (two changes each) and then cleared in xylene (three changes). The finished section is finally protected by a coverslip using a standard histological mounting medium. The method of preparation described above is typical for small implants. The glass slide is glued on the block surface to prevent curling and wrinkling of the section. When, however, large implants are processed, a relatively thick section (about 1- to 2-mm thick) should be cut from the block surface. This can then be stained and glued on the glass slide without any wrinkling. Staining The formulae for the preparation of Stevenel's blue, Van Gieson's picro-fuchsin, and alizarin red S, and the staining procedures, are as follows: Stevenel blue29 Solution Distilled water 75 ml A Methylene blue 1 g Solution Distilled water 75 ml B Potassium permanganate 1.5 g Preparation of the stains. Mix the two solutions and place in a boiling water bath until the precipitate has redissolved. This is a critical step in the procedure as failure to redissolve the precipitate completely gives a staining solution that tends to precipitate on the sections.30 Allow the stain to reach room temperature and then filter. The solution remains stable for several months and does not require periodic refiltering. Van Gieson picro-fuchsin31 Acid fuchsin, JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess… 1% aqueous solution 10 to 15 ml Saturated aqueous picric acid 100 ml Mix well and store at room temperature. The stain keeps for many months. Commercially prepared, ready to use, Van Gieson's solutions are available. However, commercial solutions tend to be low in acid fuchsin and stain bone (in undemineralized sections) less effectively than when the above formula is used. Alternatively, the Unna's32 variant can be used. This consists of acid fuchsin, 0.25 g; nitric acid, 0.5 ml; glycerin, 10 ml; distilled water, 90 ml; and picric acid to saturation. This solution gives deeper colors than Van Gieson's original stain. Alizarin red S31 Distilled water 100 ml Alizarin red S 2 g Dilute the alizarin red dye in warm (about 45¡C) distilled water stirring thoroughly. Allow the solution to reach room temperature and adjust the pH to 4.1 to 4.3 with dilute ammonium hydroxide. The solution should be a deep iodine color that keeps well for many months. Staining Procedures. Specimens are first stained en bloc with Stevenel's blue and then counterstained with Van Gieson or alizarin red S, as described below: 1. Place the block in a vessel containing Stevenel's blue preheated to, and maintained at, 60¡C in a water bath, taking care that the polished surface of the block does not touch the walls of the vessel. Stain for 15 minutes. 2. Rinse the block in distilled water at 60¡C and air dry. 3. Place the block in Van Gieson's stain at room temperature for five minutes. Blot the surface using absorbent paper to dry, or rinse briefly in absolute alcohol and air dry. 4. Alternatively, pipette a small amount of alizarin red S onto the block surface. Stain for 5 minutes at room temperature. Wash thoroughly in running distilled water to remove excess stain and air dry. Results Ground sections of tissue-implant interface prepared by the above method are illustrated in Figs. 2 to 5. Nuclear, cytoplasmic, and extracellular components of the tissues can be examined (Figs. 4 and 5). Only Stevenel's blue stains cells and extracellular structures in a subtle gradation of blue tones (except that mineralized tissues remain unstained). Counterstaining with Van Gieson's picro-fuchsin colors collagen fibers green or green-blue (Fig. 2); bone, orange or purple (Figs. 2, 3, and 4); osteoid, yellow-green (Figs. 3 and 4); and muscle fibers, blue to blue-green (Fig. 2). Alizarin red S specifically stains calcium so that only mineralized tissues are colored orange to purple. Other tissues remain blue (Fig. 5). Discussion JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess… The method of preparation described here is simple and gives well-stained sections of high quality in which cells, extracellular substance, and the tissue-implant interface can be preserved and demonstrated well using light microscopy. In previous studies, a variety of stains have been used to stain plastic-embedded tissue-implant sections. These include, acid fuchsin,1,3,9 toluidine blue and alizarin red S,12 toluidine blue and basic fuchsin,22 hematoxylin and eosin,4,5,10,18 Masson's trichrome,18,33 Giemsa,18,34 and Paragon-1301. 14,21,35-37 Kenner et al.38 have compared many of these stains, and have suggested that Paragon gives the best results, whereas the others were not satisfactory. Although Paragon stains tissues well, it did not always provide satisfactory cellular detail in this study. In common with many investigations,1-3,8,9,11,13-17,19,20,22,33-36 polymethylmethacrylate was used for infiltration and embedding. Low-viscosity epoxy resins have been used as an alternative4,6,7,10,12,18,21 to avoid foaming problems during polymerization and to reduce polymerization time.4 Use of bis (4-tert-butylcyclohexyl) peroxydicarbonate as the initiator in the methylmethacrylate solutions offers a reliable solution to both of these problems. According to Buijs and Dogterom,28 this initiator has advantages over benzoyl peroxide; it provides a better-controlled polymerization reaction and a significantly lower polymerization temperature so that there is less chance of reaching the boiling point of the methylmethacrylate. With this, foam formation is avoided, CO2 production during polymerization is significantly reduced, polymerization is rapid, and the material is not explosive and can be used without purification. The authors' experience is consistent with these findings. In addition, polymethylmethacrylate has two advantages over epoxy resins (1) an unmatched ability to completely infiltrate large pieces of properly prepared tissue and (2) the stainability of tissues embedded therein.38 Commercially available methylmethacrylate used for embedding biological specimens contains the inhibitor hydroquinone that provides a reasonable shelf life for the monomer. The most commonly used methods for preparing methylmethacrylate for embedding require either removal of the inhibitor2,8,35 or use of uninhibited monomer.36 Hydroquinone can be removed from the methacrylate monomer by washing with sodium hydroxide solution followed by distilled water,28,35,38 by the use of activated carbon,39 or by distillation.40 However, purification of the methacrylate is a tedious and laborious procedure and results in loss of as much as a third of the monomer.28 In recent studies35,38 it has been suggested that methylmethacrylate be used without removal of the inhibitor. Using inhibited monomers, the polymerization time is increased by about 30%28 Kenner et al.38 tested different concentrations of the initiator benzoyl peroxide and found that, for methylmethacrylate monomer inhibited with 25 ppm hydroquinone, an amount of 2 g/100 ml was optimal. Methylmethacrylate monomer inhibited with 10 ppm hydroquinone was used and it was found that 0.5 g of the initiator gives excellent polymerization results. Experience with 10 ppm hydroquinone-inhibited methylmethacrylate also suggests that this type of methacrylate monomer has a long shelf life and can be stored at room temperatures for periods longer than a year. The use of plexiglass slides has been suggested by some investigators as an alternative to microscope glass slides. Use of plexiglass slides is particularly helpful when microradiographs are prepared from the sections. However, plexiglass slides are usually thicker than glass and they bend or scratch more easily. Moreover, glass slides are readily JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess… available and are of a superior optical quality. The histological method described in this article was developed for the preparation of sections of metallic implant-containing tissues. It can also be used to good advantage for preparations of either ceramic or polymeric implants. However, it should be noted that methylmethacrylate embedding, acetone, and ether may adversely affect or remove some polymers (e.g., bone cement or polysulfone). Acknowledgment The authors are grateful to Dr. J.D. Bobyn and Mr. D. Wagner for helpful suggestions and discussion. JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess…

1. Galante, J., et al. Sintered fiber metal composites as a basis for attachment of implants to bone. J Bone Joint Surg 53A:101-114, 1971. 2. Hirschom, J.S., McBeath, A.A., and Dustoor, M.R. Porous titanium surgical implant materials. J Biomed Mater Res Symp 2:49-67, 1971. 3. Lembert, E., Galante, J. and Rostoker, W. Fixation of skeletal replacement by fiber metal composites. Clin Orthop 87:303-310, 1972. 4. Smith, L.G., and Karagianes, M.T. Histological preparation of bone to study ingrowth into implanted materials. Calc Tiss Int Res 14:333-337, 1974. 5. Nilles, I.L., Karagianes, M.T., and Wheeler, K.R. Porous titanium alloy for fixation of knee prosthesis. J Biomed Matr Res Symp 5:319-328, 1974. 6. Karagianes, M.T., et al. Development and evaluation of porous ceramic and titanium alloy dental anchors implanted in miniature swine. Biomed Mater Res Symp 5:391-399, 1974. 7. O'Keefe, P. A technique for preparing thin sections of porous-metal coated metallic implants to show bone ingrowth. In E.W. Filer, J.M. Hoegfeldt, and J. McCall (eds.) Microstructural Science. New York: Elsevier Science Publishing Co. Inc., 1976, vol. 4, pp. 165-177. 8. Page, K. M. Bone and the preparation of bone sections. In J.D. Bancroft and A. Stevens, (eds.) Theory and Practice of Histological Techniques. New York Churchill Livingstone Inc., 1977, chap 15, pp. 223-248. 9. Andersson, G.B.J., et al. Segmental replacement of long bones in baboons using a fiber titanium implant. J Bone Joint Surg 60A:31-40, 1978. 10. Babbush, C.A., Banks, B.A., and Weigand, A.J. Endosteal blade-vent implants modified by ion beam sputtering techniques. J Oral Implantol 8:509-533, 1979. 11. Pilliar R.M., et al. Bone ingrowth and stress shielding with a porous surface coated fracture fixation plate. J Biomed Makr Res 13:799-810, 1979. 12. Young, F.A., Spector, M., and Kresch, C.H. Porous titanium endosseous dental implants in rhesus monkeys: Microradiography and histological evaluation. J Biomed Matr Res 13:843-856, 1979. 13. Bobyn, J.D., et al. The optimum pore size for the fixation of porous-surfaced metal implants by the ingrowth of bone. Clin Orthop 150:263-270, 1980. 14. Peterson, L.J., et al. Clinical radiographic, and histological evaluation of porous rooted cobalt-chromium alloy dental implants. J Dent Res 59:99-108, 1980. 15. Szivek, J.A., et al. A study of bone remodelling using metal-polymer laminates. J Biomed Makr Res 15:853-865, 1981. 16. Bobyn, J.D., et al. Osteogenic phenomena across endosteal bone-implant spaces with porous surfaced intramedullary implants. Acta Orthop Scand 52:145-153, 1981. JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess…

17. Pilliar, R.M., et al. Radiographic and morphologic studies of load-bearing porous-surfaced structured implants. Clin Orthop 156:249-257, 1981. 18. Karagianes, M.T., et al. Investigation of long-term performance of porous-metal dental implants in nonhuman primates. J Oral Implantol 10:189-207, 1982. 19. Bobyn, J.D., et al: Biologic fixation and bone modeling with an unconstrained canine total knee prosthesis. Clin Orthop 166:301-312, 1982. 20. Ronningen, H., et al. Total surface hip arthroplasty in dogs using a fiber metal composite as a fixation method. J Biomed Mater Res 17:643-653, 1983. 21. Hirshorn, M.S., et al. Histological evaluation of porous titanium cardiac pacemaker electrode tips. J Biomed Makr Res 18:47-60, 1984. 22. Anderson, R.C., et al. An evaluation of skeletal attachment to LTI pyrolytic carbon, porous titanium, and carbon-coated porous titanium implants. Clin Orthop 182:242-257 1984. 23. Hayat, M.A. Principles and Techniques of Electron Microscopy. Biological Applications. 2nd ed. Baltimore: University Park Press, 1981, vol. 1, pp. 55-129. 24. Glauert, A.M. Practical methods in electron microscopy. Vol 3, Part I. In Fixation, Dehydration and Embedding. II. Ultramicrotomy, New York: Elsevier Science Publishing Co. Inc., 1974. 25. Anon. Embedding specimens in methacrylate resins. Pamphlet No. CM-102 L/ci, Philadelphia: Rohan and Haas. 26. Deichmann, W. Toxicity of methyl, ethyl and n-butyl methacrylate. J Industr Hyg Toxicol 23:343-351, 1941. 27. Spealman, C.R., et al. Monomeric methylmethacrylate—Studies on toxicity. Indiana Med 14:292-298, 1945. 28. Buijs, R., and Dogterom, A. An improved method for embedding hard tissue in polymethylmethacrylate. Stain Technol 58:135-141, 1983. 29. Gurr E. Biological staining methods. 8th ed. Buckinghamshire, England: Searle Diagnostic Gurr Products, 1973, pp 111. 30. del Cerro, M., Cogen, J, and del Cerro, C. Stevenel's blue, an excellent stain for optical microscopical study of plastic embedded tissues. Microscopica Acta 83:117-121, 1980. 31. Drury, R.A.B., and Wallington, E.A. Carleton's Histological Technique. 4th ed. New York: Oxford University Press, 1967, pp. 151, 166-167. 32. Lillie, R.D. Histopathological technic and practical histochemistry. New York: McCraw-Hill, 1965, chap 15, pp. 539-540. 33. Homsy, C.A., d al. Porous implant system for prosthesis stabilization. Clin Orthop 89:220-235, 1972. JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess…

34. Gross, U.M., and Strunz, V. Surface staining of sawed sections of undecalcified bone containing alloplastic implants. Stain Technol 52:217-219, 1977. 35. Klawitter, J.J., and Hulbert, S.F. Application of porous ceramics for the attachment of load bearing internal orthopedic applications. J Biomed Mater Res Symp 2:161-220, 1971. 36. Steflik, D.E., et al. Simultaneous histological preparation of bone, soft tissue and implanted biomaterials for light microscopic observations. Stain Technol 57:91-98, 1982. 37. Park, J.B., et al. Intramedullary fixation of artificial hip joints with bone cement-precoated implants. II. Density and histological study. J Biomed Mater Res 16:459-469, 1982. 38. Kenner, G.H., et al. Bone embedding technique with inhibited PPMA monomer. Stain Technol 57:121-126, 1982. 39. Franklin, R.M., and Martin, M.T. Staining and histochemistry of undecalcified bone embedded in a water-miscible plastic. Stain Technol 55:313-321, 1980. 40. Parsons, D.F., and Danden, F.J., Jr. Optimal conditions for methacrylate embedding of certain tissues and cells sensitive to polymerization damage. Exp Cell Res 24:466-483, 1961. JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess… JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess…

Fig. 1 The custom-made holder used for specimen grinding comprises an aluminum split polishing block (b, b') relieved (r) to accommodate a g/ass slide (25 × 75 mm). When the polishing block is not fully closed larger glass slides (38 × 75 mm or 50 × 75 mm) can be accommodated. Pieces of tungsten carbide (c) protrude 150 µm from the surface, to protect the section from overgrinding. The slide carrying the glued section is placed in the recess, secured in position using the screw(s) and ground to a thickness of 150 µm (glue plus section). The section is further reduced in thickness after raising the g/ass slide above the surface by placing metal strips beneath it. The process is repeated until the desired section thickness is obtained.

Fig. 2 Ground section of a porous-surfaced endosteal implant (im). The section has been stained with Stevenel's blue and Van Gieson's JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess… endosteal implant (im). The section has been stained with Stevenel's blue and Van Gieson's picro-fuchsin. A variety of tissues is observed, including bone (b), muscle (m), soft connective tissue (ct), blood vessels (bv), and nerve (n) (× 45).

Fig. 3 Ground section of a porous-surfaced implant (im). Deposition of bone (b) into the pores of the implant is evident, as is the osteoid (Od) lining on osteon. The morphology of the bone has been preserved and the bone-implant interface is intact. (Stevenel's blue and van Gieson's picro-fuchsin, × 90). JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess…

Fig. 4 A cutting cone, indicative of remodeling of compact bone, at the surface of a porous-surfaced implant (im). Osteoclasts (Oc), osteoblasts (Ob), osteoid (Od), bone (b), and osteocytes (Os) are demonstrated. (Stevenel's blue and Van Gieson's picro-fuchsin, × 360). JOMI on CD-ROM, 1986 Jan (31-37 ): An Improved Method for Preparing Histological Sectio… Copyrights © 1997 Quintess…

Fig. 5 Ground section of bone stained with Stevenel's blue and alizarin red S. Numerous osteoblasts (Ob) are demonstrated at the osteoid (Od) surface (× 280).