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Year : 2012  |  Volume : 23  |  Issue : 6  |  Page : 753-757
The microbiology of the peri-implant sulcus following successful implantation of oral prosthetic treatments

1 Department of Prosthodontics, Dental Research Center, School of Dentistry, Mashhad University of Medical Sciences, Mashhad, Iran
2 Department of Microbiology, Mashhad University of Medical Sciences, Mashhad, Iran

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Date of Submission31-May-2011
Date of Decision27-Nov-2011
Date of Acceptance13-Apr-2012
Date of Web Publication3-May-2013


Background: Oral implants are widely used in partially and fully edentulous patients; however, the integration of an implant can be endangered by factors such as intraoral bacteria or inflammatory reactions. The purpose of this study was to evaluate the microbial flora present in the sulcus around dental implants and to assess the relationship between gingival health and microbial flora present.
Materials and Methods: Twenty patients who had received oral implants with no complications were followed for a period of 9 months. Assessment of probing depth, the presence of bleeding on probing and microbial sampling from the peri-implant sulcus were performed at three different time points- 4 weeks after surgery, 1 month and 6 months after loading. The samples were taken by paper points and transferred to the microbiology lab in thioglyocolate cultures. In order to do a colony count and isolate the aerobic capnophilic and anerobic bacteria the samples were cultured and incubated on laboratory media. The colonies were also identified using various diagnostic tests. Alterations in the presence of various bacterial species over time and gum health were tested using analysis of variance (ANOVA) with Tukey's test post hoc.
Results: The average pocket depth for each patient ranged from 1.37 ±0.39 mm to 2.55 ± 0.72 mm. The bacteria isolated from the cultured samples included aerobic, facultative anerobic, obligate anerobic and capnophilic bacteria.
Conclusion: The anerobic conditions created in the peri-implant sulcus might with time enhance the number of anerobic bacteria present following dental implant loading.

Keywords: Implants, loading, microbiology, prosthetic treatments

How to cite this article:
Asadzadeh N, Naderynasab M, Fard FG, Rohi A, Haghi HR. The microbiology of the peri-implant sulcus following successful implantation of oral prosthetic treatments. Indian J Dent Res 2012;23:753-7

How to cite this URL:
Asadzadeh N, Naderynasab M, Fard FG, Rohi A, Haghi HR. The microbiology of the peri-implant sulcus following successful implantation of oral prosthetic treatments. Indian J Dent Res [serial online] 2012 [cited 2021 Aug 5];23:753-7. Available from:
For more than 1500 years the replacement of lost teeth was an unattainable goal. However, recent advances in implant surgery, whereby titanium implants inserted into the jaw, integrate with the bone, was first described by Branemark in 1952. [1] The process of osseointegration of oral implants can be influenced by various parameters such as oral bacteria and inflammatory infection. The survival of a dental implant can also be influenced by factors such as occlusal over loading and peri-implantitis due to the presence of plaque which is dependent on the geometry and surface features of implants. Other patient-related parameters including smoking, bone quality, the presence of systemic disease, trauma or bacterial infection can also result in early implant failure. [2] Out of all these potential issues it has been suggested that the two most common causes of implant failure are occlusal overload, which is influenced by prosthetic reconstruction, and microbial attack, which is dependent on the microbial flora around implants and on bacterial plaque. [3]

In many cases overloading causes marginal bone loss and leads to a pocket surrounding the implant. After a while, the newly created anerobic environment may begin to host a variety of periodontal disease-causing flora. Therefore the presence of disease-causing sub-gingival flora after occlusal overload can reflect infection in the environment and this may result in more bone reduction. [4] Even without the presence of overloading Crawford stated that there is a clear difference between the microbiology of stable implants and diseased ones and that undoubtedly, Gram-negative bacteria are involved in this pathology. [5] In contrast, a study by Heydenrijk and colleagues of the bacteria present in peri-implantitis showed that the inflamed implants were colonized by a large number of Gram-positive anerobic bacteria such as Fusobacteria spirochetes,  Bacteroides forsythus Scientific Name Search  bacteria-containing black pigments such as Prevotella intermedia, Prevotella nigrescens and Prevotella gingivalis, Actinobacillus actinomycetemcomitans was also isolated from the samples taken, indicating that the microbial flora of inflammatory lesions around implants is very similar to that present in adult periodontitis including refractory type periodontitis. [6] It has also been shown that there is a relationship between probing depth and the presence of organisms such as Capnocytophaga spp. and Actinomyces odontolyticus around supportive implants in edentulous patients. [2] Another study by Puchads-Roman and colleagues indicated that increased probing depth was associated with a high percentage of Spirochetes around implants in comparison to natural teeth. [7]

However, there is some evidence that the presence of periodontal pathogens may not always end in a destructive process. [6]

In a long-term study by Leonhard and colleagues, all patients underwent advanced periodontal treatment. Before treatment and during the study, these patients were found to harbor bacteria such as P. gingivalis, P. intermedia, A. actinomycetemcomitans, Capnocytophaga spp. and C. rectus. which might indicate that the presence of periodontal pathogene is not related to the failure of implants and that these types of bacteria are a part of the normal periodontal flora. [8]

Similarly, Lee and colleagues showed that prosthetic rehabilitation has a limited effect on the microbial flora around implants. The results of the study suggested that periodontal disease history had more influence on the microbial flora around implants than implant loading interval and that the remaining teeth produced the greatest effect on the flora around implants. P. gingivalis, B. forsythus and red complex bacteria colonized some implants but all implants were successful. [9]

Given the amount of competing evidence regarding the microbial colonization of oral implants and oral implant failure this study aimed to evaluate the microbial flora present in the sulcus around dental implants and to assess the relationship between gingival health and the microbial flora present.

   Materials and Methods Top

Initially two patients were examined in a pilot study to survey bacterial colonization and identify relevant bacterial culture methods. On the basis of the statistical analysis from the pilot study and other similar studies 20 patients with 20 implants were selected and followed for 9 months. Smokers and patients taking antibiotics were excluded.

Bacterial samples were taken from the sulcus around the implants at the beginning of prosthetic treatment (4 weeks after implant surgery), and 1 month and 6 months after loading of the prosthetics. In addition, pocket depth was measured in 4 spots around the implants at each time point and the presence or absence of bleeding during probing was also recorded.

To take samples, the microbial plaque around the implant sulcus was removed using cotton roll, and the implant and its surrounding region were dried by a gentle pressure air. Following this a paper point was slowly placed inside the sulcus for 10 seconds at the buccal, mesial, distal and lingual sides. Then the samples were transferred to a thioglycolate environment in our microbial laboratory in order to isolate the aerobic from the anerobic bacteria present.

For each sample the test tube containing the thioglycolate medium was first mixed by vortex for 60 seconds. Then; in order to culture the bacteria and perform a colony count, a serial dilution (1:10, 1:100 and 1:1000) was prepared from the solution. In order grow the bacteria, blood taken from each patient during a blood test was added to the culture plates at a concentration of 5% v/v. Colonization of  Petri dish More Detailses was used to count the bacteria and isolated cultures in petri dishes were used to carry out Gram-positive coloring and diagnostic tests. The petri dishes were incubated in aerobic conditions for 48 hours at 37° C, and the number of colonies was counted. The number of colonies from the undiluted sample was considered as a positive control because the number of colonies in each petri dish for each serial dilution should be 10 times less than the previous dilution.

In addition, different culturing environments were used for specific bacteria. A tryptic soy environment was prepared and used to culture and raise A. actinomycetemcomitans along with 5% blood, 75 mg/ml bacitracin and 5 mg/ml vancomycin. The samples incubated in a CO 2 jar at 37°C for 5 days. [8] On the other hand, a basic blood agar environment was used to raise Prevotella spp. and Porphyromonas spp. along with 5% blood and 40 mg kanamycin. Peptostreptococcus spp. was also raised on this environment due to their resistance to kanamycin. These samples were incubated after planting in an anerobic colony jar at 37°C.

At time of closing of the door of the anerobic jar, a 2-pack gauze or gas-producing kit (N 2 85%, H 2 10%, CO 2 5% instead of oxygen) and an anerobic indicator were placed in the jar to maintain and monitor the precise anerobic conditions of the jar.

Once the samples had been cultured the diagnostic tests used to identify the bacteria included the scolin hydrolysis, nitrate reductase, indole production, catalase activity and DNase production tests. Petri dishes that were incubated in aerobic conditions were examined after 48 hours and slides were prepared from the various colonies that had been raised separately. After counting the aerobic colonies were stained using Gram's method.

Petri dishes incubated in CO 2 and media containing bacitracin, vancomycin, and/or kanamycin were removed from the jar and examined after 5 days. Similar to the aerobic bacteria, following counting slides were prepared and stained using Gram's method, and based on the accepted stain and the colony shape diagnostic tests were performed to biotype each colony.

Small colonies with a star-like shape under the microscope that had been soaked in agar and had a positive catalase test were recognized as A. actinomycetemcomitans.

Colonies of oral black-pigmented bacteria were recognized by the; production of indole and the fermentation of lactose. In order to distinguish between Porphyromonas spp. and Prevotella spp. sensitivity to vancomycin was used.

To recognize bacteria such as F. nucleatum, P. gingivalis, P. intermedia and A. actinomycetemcomitans; they were ATCC-positive control in the microbiology section that checked the results of the test.

Data regarding the different species of bacteria identified at the three time points as well as the probe depth and presence of bleeding was collected and analyzed using SPSS software, version 15.0(SPSS, Chicago, IL). Analysis of variance (ANOVA) with and Tukey's test post hoc was used to determine statistically significant results.

   Results Top

Absence or presence of bleeding during probing was recorded at the three times described above. There was a significant difference between the first and third visit (P = 0.031) in the percentage of bleeding during probing with the amount of bleeding increasing with time [Table 1]. There was also a significant difference in overall pocket depth between the first and second visit (P < 0.001) and between the first and third visit (P < 0.001). For each aspect of the implant specifically there was a significant difference in the pocket depth of the distal and buccal sample sites between the first and second visit (P = 0.04) and between the first and third visit (P < 0.001).
Table 1: Mean and standard deviation (SD) of bleeding (%) during probing

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Bacteria isolated from the samples included aerobic, facultative anerobe, capnophilic and obligate anerobe groups. Identified aerobic/facultative anerobic bacteria included Streptococcus spp., coagulase-positive Staphylococcus spp., coagulase-negative Staphylococcus spp., and a scant number of Klebsiella spp. and Enterobacter spp. Anerobic bacteria present included Peptostreptococcus spp., Porphyromonas gingivalis, Prevotella intermedia,  Neisseria More Details spp. and Veillonella spp. The most abundant anerobes were Fusobacterium nucleatum and the capnophilic Actinobacillus actinomycetemcomitans.

Bacterial counts did not have normal distribution so in order to normalize this distribution instead of analyzing direct bacterial counts we used logarithmic values to the base of 10. Using this method there was a significant difference between the number of Streptococcus bacteria detected in the first and second visit (P < 0.001), in the first and third visit (P < 0.001), and in the second and third visit (P = 0.027, [Table 2]).
Table 2: Mean and standard deviation (SD) of logarithmic bacterial counts for Streptococcus species

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There was also significant difference between the number of Peptostreptococcus colonies detected in the first and the second visit (P< 0.001), in the first and the third (P < 0.001), and in the second and the third visit (P = 0.027, [Table 3]).
Table 3: Mean and standard deviation (SD) of bacterial counts of Peptostreptococcus

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   Discussion Top

The present study surveyed the microbiology of the peri-implant sulcus and studied periodontal parameters such as bleeding during probing and the depth of the gingival pocket.

In a study by Shibli and colleagues which evaluated the on composition of the supra and sub-gingival biofilm, four bacterial groups showed the greatest abundance with Veillonella and Fusobacterium predominating. [10] Although this study examined the bacteria present around intact teeth, as most oral pathogens have been shown to disappear when all teeth are extracted it seems that each tooth holds bacteria, and that the bacteria around teeth and implants are similar to each other. This has been shown by several studies such as those by Apse and colleagues [11] and Quirynen and colleagues, [12] which indicate that the composition of the microbial flora around implants is dependent on the presence of teeth and, consequently, the bacteria present around implants and around teeth are similar. Another study by Buchmann and colleagues stated that among the bacteria cultured from around implants Peptostreptococcus and Streptococcus were the least abundant, Veillonella had a variable frequency, and of the Gram-negative bacteria Fusobacterium nucleatum, Porphyromonas gingivalis, and Peptostreptococcus micros were the most abundant. [14] In an analysis of the a biofilm that forms on the surface of oral implants Heuer and colleagues, showed using molecular methods that Porphyromonas gingivalis and A. actinomycetemcomitans and were more abundant than other bacteria. [14]

In the present study, Fusobacterium and Veillonella were cultured from samples taken at all three time points but while Fusobacterium was the predominant bacteria the amount of Veillonella varied as in Buchman's study [13] and was less abundant than the other anerobic bacteria detected.

Another one of our observations was that Actinomyces species were not isolated in this study although they have previously been isolated in other studies. [12],[13] Similarly, Quirynen and colleagues also did not isolate Actinomyces in their study and suggested that this might be due to differences in isolation methods (molecular vs. bacteria culturing), geographical differences and habits in different regions. [15] We also observed an absence of Spirochetes in these patients. However, a study by Nakou and colleagues suggested that the presence of Spirochetes is related to the presence of inflammation around implants and considering that the selected patients were healthy it is not surprising that we did not isolate any Spirochetes from the samples. [16] In other studies, Mombelli and colleagues stated that 80% of the bacteria cultured from around oral implants were facultative anerobic Gram-positive cocci species which are observed during the first 6 months following implant placement. They also did not detect Spirochetes but did detect an abundance of Fusobacterium and also anerobic black-pigmented Gram-negative bacteria. As mentioned above, in the present study there was no observation of Spirochetes as in the Mombelli study but the amount of facultative anerobic Gram-positive cocci we detected was not as large as in Mombelli study. [17]

In terms of gingival health, in the study by Shibli and colleagues the average depth of the pocket around implants was 3.41 ±1.18 mm, [10] while this parameter was more than 3 mm in 75% of cases in a study by Salvi's and colleagues, [18] 2.5 ± 0.67 mm in a study by Omlak and colleagues, [3] and 2.8 ± 0.8 mm in the study by Quirynen and colleagues. [14] In the present study, the average pocket depth over all time points had a minimum of 1.37 ± 0.39 mm and at maximum 2.55± 0.72 mm, which is very similar to the results observed by Olmak. [4]

Another parameter surveyed was the presence of bleeding during probing. Shibli and colleagues stated that after 2 years bleeding on probing was 4.9± 37.3 %. [10] Bleeding during probing in the study by Salvi was reported to be 2-11%. [18]

In the study by Olmak, this parameter was reported to be between 20 and 30 % [3] while Quirynen reported it as between 10 and 20%. [14] In the present study, 4 weeks after surgery, bleeding during probing was not observed and in the next 2 visits, bleeding during the probing was between 25 and 30% with a standard deviation of 44-47%. Lack of similarity between the results from this study and the others might be due to periodontal loading conditions and the difference in microbiology around implants.

Overall the results of this study indicate that the presence of bleeding in response to probing and the pocket depth around oral implants increases over time in the first 9 months following initiation of the implant procedure. Aerobic and anerobic bacteria are observed within a month following surgery and with the passage of time, especially after loading, the number of anerobic bacteria increases creating anerobic conditions in the peri-implant sulcus.

   References Top

1.Newman M, Takei H, Klokkevold P, Carranza F. Carranza's clinical periodontology. St Louis: Saunders, 2002. p. 882.  Back to cited text no. 1
2.Quirynen M, Soete MD, Van Steenberghe D. Infectious risks for oral inplants: a review of the literature. Clin Oral Imp Res 2002;13:1-19.  Back to cited text no. 2
3.Olmak U. Microbiologic findings with implant treated edentulous patients: a preliminary report. Dental Turk Uyeleri 2006;6:2368-73.  Back to cited text no. 3
4.Ellen RP. Microbial colonization of the peri-implant environment and its relevance to longterm success of osseointegrated implants. Int J Prosthodont 1998;11:433-41.  Back to cited text no. 4
5.Crawford JJ. Orofacial infections and antibiotic managemen. In: Newman MG, Goodman AD, editors. Guide to antibiotic use in dental practice. Chicago: Quintessence, 2006. p. 25-38.  Back to cited text no. 5
6.Heydenrijk K, Meijer HJ, Van der Reijden WA, Raghoebar GM, Vissink A, Stegenga B. Microbiota around root-from endosseous: a review of the literature. Int J Oral Maxillofac Implants 2002;17:829-38.  Back to cited text no. 6
7.Puchades-Roman L, Palmer RM, Palmer PJ, Howe LC, Ide M, Wilson RF. A clinical, radiographic and microbilogic comparison of Astr Tech and Branemark single tooth implants. Clin Implant Dent Relat Res 2000;2:78-84.  Back to cited text no. 7
8.Leonhardt A, Gröndahl K, Bergström C, Lekholm UL. Long-term folow- up of osseointegrated titanium implants using clinical, radiographic and microbilogical parmeters. Clin Oral Implant Res 2002;13:127-32.  Back to cited text no. 8
9.Lee KH, Maiden MF, Tanner AC, Weber HP. Microbiota of successful osseointegrated dental implants. J Periodontal 1999;70:131-8.  Back to cited text no. 9
10.Shibli JA, Melo L, Ferrari DS, Figueiredo LC, Faveri M, Feres M. Composition of supra and subgingival biofilm of subjects with healthy and diseased implants. Clin Oral Imp Res 2008;19:975-82.  Back to cited text no. 10
11.Apse P, Ellen RP, Overall CM, Zarb GA. MIcrobiota and crevicular fluid collagenase activity in the osseointegrated dental implant sulcus: A comparison of sites in edentulous and partially edentulous patients. J Periodont Res 1989;24:96-105.  Back to cited text no. 11
12.Quirynen M, Vogels R, Peeters W, Van Steenberghe D, Naert I, Haffajee A. Dynamics of initial subgingival colonization of "pristine" peri-implant pockets. Clin Oral Imp Res 2006;17:25-37.  Back to cited text no. 12
13.Buchmann R, Khoury F, Pingel D, Lange DE. The microflora recovered from the outer-surfaces of the Frialit-2 implantoprosthetic connector. Clin Oral Implants Res 2003;14:28-34.  Back to cited text no. 13
14.Heuer W, Elter C, Demling A, Neumann A, Suerbaum S, Hannig M, et al. Analysis of early biofilm formation on oral implants in man. J Oral Rehabilit 2007;34:377-82.  Back to cited text no. 14
15.Quirynen M, Vogels R, Pauwels M, Haffajee AD, Socransky SS, Uzel NG, et al. Initial subgingival colonization of "pristine" pockets. J Dent Res 2005;84:340-4.  Back to cited text no. 15
16.Nakou M, Mikx FH, Ooseterwaal PG, Kruusen JC. Early microbial colonization of permucosal implants in edentulous patients. J Dent Res 1987;66:1654-7.  Back to cited text no. 16
17.Mombelli A. Cimcobiology of the dental implant. Adv Dent Res 1993;7:202-6.  Back to cited text no. 17
18.Salvi GE, Fuörst MM, Lang KP, Persson GR. One-year bacterial colonization patterns of staphylococcus aureus and other bacteria at implants and adjacent teeth. Clin Oral Imp Res 2008;19:242-8.  Back to cited text no. 18

Correspondence Address:
Hamidreza Rajati Haghi
Department of Prosthodontics, Dental Research Center, School of Dentistry, Mashhad University of Medical Sciences, Mashhad
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Source of Support: None, Conflict of Interest: None

DOI: 10.4103/0970-9290.111253

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  [Table 1], [Table 2], [Table 3]


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