The purpose of this study was to evaluate the clinical success of an implant placed immediately after the explantation of a fractured blade implant. A healthy 58-year-old male nonsmoker presented with a fractured blade implant that had been subjected to biomechanical overload. A new blade implant was placed immediately after the removal of the fractured one. The new implant was placed with a composite graft of collagen gel and corticocancellous porcine bone and covered with a bioabsorbable membrane. Radiographic evaluation at 6 months postoperation showed complete bone healing. No residual bone defect was observed or probed during the uncovering phase; moreover, no mobility, pain, suppuration, or presence of peri-implant radiolucency were observed at the second-stage surgery.
Implant placement immediately or shortly after tooth extraction has proven to be a predictable treatment protocol regardless of implant shape.1,2 This technique has several advantages, such as the reduction of time between tooth extraction and placement of the definitive prosthetic restoration, and preservation of alveolar ridge height and width.3
Many clinical reports and experimental studies in animal models have demonstrated the favorable outcome of dental implants immediately placed in fresh extraction sockets, with or without the use of membranes or regenerative procedures.4 Schliephake and Hracht5 histologically and histometrically compared implants immediately placed into extraction sites with or without polylactic acid membranes and found no differences in bone-implant contact between implant sites treated with membranes and controls.
Celletti et al6 reported that titanium-reinforced ePTFE membranes achieved no significant gain in extraction socket bone height when compared with untreated control sites.
Over the years, implant systems have been introduced, and the indications for implant rehabilitation have gradually been expanded.9 Although high success rates have consistently been reported for many implant systems, complications leading to loss of the implant still occur.
Albrektsson and coworkers10 suggested clinical criteria to identify the success of dental implants, such as absence of infection, pain, mobility, and radiographic bone loss. The incidence of biological peri-implant and mechanical complications is usually due to bacterial infection and biomechanical overload.11 Biological complications include peri-implant radiolucency, bleeding on probing, increased probing depth, and radiographic signs of bone loss.12,13 Adverse occlusal forces on the implant-prosthetic complex have been reported to cause mechanical failure of the components14 and even loss of osteointegration.15,–17 Poor bone quality, such as that often found in the posterior jaw, could affect implant survival.
This article reports on the clinical and radiographic success of a blade implant placed immediately after removal of a preexisting fractured blade implant.
A healthy 58-year-old male nonsmoker was referred for treatment of a fractured blade implant that had been in function for 10 years. The prosthesis showed slight mobility, and it was decided to remove the fractured blade implant and replace it immediately with a new blade implant. A complete diagnostic workup was performed, which included fabrication of diagnostic casts for determining the intra-arch relationship, panoramic radiography, and computerized tomography. The treatment plan was thoroughly explained to the patient, who signed the informed consent.
Initial scaling and root planning were performed to provide an oral environment more favorable to wound healing.
Immediately before surgery, the patient rinsed for 1 minute with chlorhexidine mouthwash and was instructed to use it twice daily for 4 weeks after surgery.
Intrasulcular and vertical incisions that extended over the mucogingival junction were made to raise a mucoperiosteal flap. Examination of the fractured blade implant revealed that the intraossous portion was still osseointegrated (Figures 1 and 2). A thin bur was used at low speed under external irrigation to carefully isolate the residual apical part of the fractured implant Figure 3. A gentle explantation procedure was performed to minimize the bone trauma and to maintain the integrity of the walls of the alveolar housing (Figures 4 and 5). After blade implant removal, the surgical site was prepared using standard procedures, and a new implant with the same shape was placed (Figure 6). The peri-implant bone defect was grafted with a mixture of collagen gel and corticocancellous porcine bone (Osteobiol, Tecnos, Coazze, Italy) (Figures 7 and 8) and covered with a bioabsorbable membrane (Evolution, Tecnoss, Coazze, Italy). The mucoperiosteal flap was mobilized for the implant primary stability, then sutured to favor the wound (Figure 9). The patient was prescribed antibiotics, anti-inflammatories, and chlorhexidine mouthwashes.
Sutures were removed after 7 days, and a removable prosthesis was worn for the first 3 weeks. The patient reported light pain and swelling during the first week, but there were no other complications. During the follow-up period, the patient was seen monthly. The second-stage surgery was performed 6 months after implantation (Figure 11). A minimal incision was made at the crestal level to remove the cover screw of the implant and for placement of a healing abutment. Six months after implant placement, it was asymptomatic, immobile, and osseointegrated (Figure 10). No peri-implant bone defects were observed by probing. No signs of infection or bleeding on probing were detected (Figure 13).
Histological examination was performed on the removed blade implant according to the protocol described by Crespi and Grossi.21
The samples were immediately immersed and left overnight in a fixative consisting of an ice-cold mixture of 4% glutaraldehyde and 4% paraformaldehyde buffered to pH 7.2 with 0.2 M sodium cacodylate buffer. After fixation, the samples were placed in a 0.2-M sodium cacodylate buffer (pH 7.3) for several hours, postfixed for 2 hours in a 2% osmic acid solution in sodium cacodylate buffer, rinsed 3 times in this buffer, then dehydrated under agitation: 25% ethanol 10 minutes, 50% ethanol 10 minutes, 75% ethanol 15 minutes, 95% ethanol 50 minutes, 100% ethanol 1 hour, propylene oxide solvent 1 hour (Fluka Chimie AG, Buchs, Switzerland). The dehydrated samples were then infiltrated with resin in the following sequence: propylene-oxide + Epon-Araldite resin 2:1 (v/v), for 1 hour; propylene-oxide + Epon-Araldite resin 1:1 (v/v), for 1 hour; propylene-oxide + Epon-Araldite resin 1:2 (v/v), for 1 hour; Epon-Araldite resin overnight; polymerize at 80°C, 3 days.
The resin-embedded samples were sectioned using the Isomet Buelher microtome (Buelher, Lake Bluff, Ill). Sectioning time varied from 10 to 15 minutes per sample. The resulting plastic block was oriented in a predetermined position so that the coronal and apical surfaces of the section, as well as the distal, buccal, mesial, and lingual surfaces, were always in the same plane. The block was mounted on a chuck using Thermoplastic Cement (Buelher). A wafering blade 4 inches in diameter and with a thickness of 0.012 inches was used for the sectioning. The load applied was sufficient to allow cutting but not enough to cause glazing. Sample slabs of 200-μm thickness were produced as a result of the sectioning process. The resulting slabs were oriented as described above and affixed to a glass slide with Canadian balsam for subsequent grinding and polishing. Canadian balsam was selected over other cements for translucency and correct refractive index.
Root slabs attached to glass slides were ground using a Metaserv Buelher (Buelher) grinding machine, in which grinding is accomplished by rotation of a plate supporting different abrasive diamond disks at speeds varying from 50 to 500 rpm. Grinding speeds were maintained constant at 200 rpm.
The root slabs were ground in 2 steps using plates with abrasive diamond disks of 500 and 100 grain size. The slabs were held on the rotating abrasive surface with relatively mild finger pressure, resulting in an average force of 0.4 Å according to the pressure gauge of the grinding machine (with a range of 0 to 1.2 Å). The final thickness of the section was estimated from standard sections prepared prior to the study sections. These standard sections had been ground to thicknesses ranging from 8 to 50 μm. Optical resolution and fine-tissue detail were obtained in sections between 8- and 20-μm thick. Final section thickness was confirmed with a microscope calibrated in micrometers. Constant running water was used as a coolant during the grinding procedure and also served to remove debris from the section's surface. The sections were then allowed to air dry at room temperature. When thoroughly dried, a cover glass was placed on the section using resinous mounting medium.
Phase-Contrast Microscopy Technique
The sections postfixed with osmic acid were practically invisible by bright field microscopy. This technical disadvantage was overcome by viewing the osmic acid stained sections under phase-contrast microscopy using a Zeiss FOMI III microscope (Carl Zeiss, Oberkochen, Germany) equipped for phase-contrast illumination and for incident light.
At low magnification (×12), the implant surface revealed a perfect congruence with that of the bone surface. The tight adhesion of bone to the metal was demonstrated by examination of the interfacial bone. Direct bone apposition to the implant was indicated by the presence of corrugations on the bone surface that exactly matched the superficial rough irregularities of the metal (Figures 6 and 7).
High-power examination (×10 000) of the bone surface revealed the presence of small holes on the bone surface at the bone side of the interface. The diameter of these structures varied from 0.1 to 1 μ. Examination of the internal portion of these holes demonstrated that they were connected in a network-like system in a similar way to bone osteocyte canaliculi.
In the present case report, the implant immediately placed after removal of the fractured implant showed a good clinical outcome. There is a lack of data concerning implants placed immediately after the removal of a failed implant. When an implant fails, it must be immediately removed. The receptor site should then be examined to evaluate the presence of a soft-tissue lining, which is generally a combination of connective tissue and epithelium. This soft-tissue should be carefully curetted from the site. In some cases, when an implant is placed in a fresh extraction socket, it can achieve intimate contact with the alveolar walls of the receptor site, while in other sites, this close contact with the bone may be lacking. When the latter situation occurs, or a portion of the implant wall is exposed because of a dehiscence in the bone, guided tissue regeneration techniques can be employed using barrier membranes with or without bone graft materials.
The use of membranes with immediate postextraction implants has been recommended26 to prevent connective tissue down-growth during the healing phase between the socket walls and the implant surface in the most coronal portion of the bone-implant interface, which prevents osseointegration. Actually, several authors have reported a high rate of membrane dehiscence after immediate implant placement; Warrer et al27 demonstrated that predictable complete osseointegration of dental implants placed in fresh extraction sockets can occur when the membrane remains covered during the healing period.
Membrane exposure during healing in the oral cavity can lead to bacterial colonization and infection, which require membrane removal.
In 1988, Bowers and Donahue28 suggested that primary stability can be achieved over the immediate implant and membrane by using periosteal releasing incisions and vertical incisions to get sufficient mobility of the buccal flap.
The technique suggested by Bowers and Donahue28 has the disadvantage of moving any existing buccal attached gingiva coronally, resulting in a gap between the mucogingival junction of the treated site and adjacent site.
On the basis of these considerations, it has been recently observed that the use of a barrier membrane is not always necessary, especially in the presence of small bone defects not exceeding 2 mm, because they can heal spontaneously.
In the present case, a blade implant had to be removed because biomechanical complications caused it to fracture. A gentle explantation was performed, and a new implant of the same shape was immediately placed.
The histological analysis performed on the removed blade implant revealed the presence of high percentages of direct bone-implant contact, supporting the hypothesis that osseointegration can occur around a blade implant.
The histological data obtained from our samples agree with the findings of Michael et al.29 In their evaluation of consecutively placed unloaded root-form and plate-form implants, Michael and coworkers29 reported that unloaded plate-form or blade implants osseointegrated and achieved intimate bone-to-implant contact as well as reported on control implants in adult Macaca mulatta monkeys.
Piattelli and coworkers30 used light microscopy, scanning electron microscopy, and laser scanner microscopy to evaluate thin ground sections of blade implants retrieved after 7 to 20 years of clinical function and found in adult M mulatta monkeys that most of the implant surfaces had intimate contact with compact lamellar bone tissues.
The present case report demonstrated successful immediate replacement of a failed blade implant with a new implant of the same shape in the same location. Histological findings showed no fibrous tissue formation (encapsulation) and excellent bone-to-implant contact with the failed implant. Additional study with longer follow-up periods and a larger population are necessary before any definitive conclusions can be drawn.
Ugo Covani is the chair of oral pathology at Nanoword Institute, School of Dental Medicine, University of Genova, Italy.
Simone Marconcini is the chairman and Antonio Barone is a visiting professor of oral medicine and pathology at the School of Dental Medicine, University of Genoa, Italy. Address correspondence to Dr Marconcini at Piazza Diaz 10, 55041 Camaiore (Lu), Italy. (email@example.com)
Roberto Crespi is a research fellow at the University of Genoa, Italy.