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Malcolm P. J. Young May Korachi Duncan H. Carter Helen V. Worthington James F. McCord David B. Drucker

Authors’ affiliations: Malcolm P. J. Young, Duncan H. Carter, Helen V. Worthington, James F. McCord, Department of Dental Surgery & Medicine, Turner Dental School, University of Manchester May Korachi, David B. Drucker, School of Biological Sciences, Turner Dental School, University of Manchester, Manchester, UK Correspondence to: Mr Malcolm P. J. Young Unit of Oral and Maxillofacial Surgery Department of Dental Medicine & Surgery University Dental Hospital of Manchester Higher Cambridge St, Manchester M15 6FH UK Tel: π44 161 274 6652 Fax: π44 161 275 6776 e-mail: D.Drucker/


Accepted 1 February 2001 To cite this article:

Young MPJ, Korachi M, Carter DH, Worthington HV, McCord JF, Drucker DB. The effects of an immediately pre-surgical chlorhexidine oral rinse on the bacterial contaminants of bone debris collected during dental implant surgery Clin. Oral Impl. Res. 13, 2002; 20–29 Copyright C Munksgaard 2002 ISSN 0905-7161


The effects of an immediately presurgical chlorhexidine oral rinse on the bacterial contaminants of bone debris collected during dental implant surgery

Key words: microbial analysis, chlorhexidine mouthrinse, collected bone debris, endosseous dental implants Abstract: Dental implant surgery produces bone debris that can be used in the ‘‘simultaneous augmentation’’ technique. Although this debris is contaminated with oral bacteria, a stringent aspiration protocol has been shown to reduce the levels of contamination. Chlorhexidine mouthrinse is a well-proven antibacterial rinse that has been shown to reduce infectious complications associated with dental implants. This study examined the effect of pre-operative rinsing with a 0.1% chlorhexidine digluconate mouthrinse on the bacterial contaminants present in collected bone debris bone (CBD). Twenty partially edentate patients were randomly allocated into equal groups and underwent bone collection using the Frios Bone CollectorA (FBC) during the insertion of two dental implants. In group T a pre-operative chlorhexidine rinse was used, whilst in group C sterile water was used. For both groups, a stringent bone collection protocol was used. Bone samples were immediately transported for microbial analysis. Colonial and microscopic morphology, gaseous requirements and identification kits were utilised for identification of the isolated microbes. Thirty-nine species were identified including a number associated with disease, in particular Actinomyces odontolyticus, Clostridium bifermentans, Prevotella intermedia, and Propionibacterium propionicum. Samples from group T (chlorhexidine mouthrinse) yielded significantly fewer organisms (P⬍0.001) than in group C (sterile water mouthrinse). Gram-positive cocci dominated the isolates from both groups. It is concluded that if bone debris is to be used for the purpose of immediate simultaneous augmentation, a preoperative chlorhexidine mouthrinse should be utilised in conjunction with a stringent aspiration protocol to reduce further the bacterial contamination of CBD.

It is accepted that a stringent approach to minimise bacterial contamination should be used for the insertion of endosseous oral implants if ‘‘osseointegration’’ is to be achieved with a minimum of infectious complications (Lambert et al. 1997; Adell et al. 1985). Furthermore, it is widely accepted that infection has a detrimental impact upon the success rates of endosseous dental implants, of guided tissue regeneration and guided bone regeneration (Dent et al. 1997; Yoshinari et al. 1998; De

Sanctis et al. 1996; Nowzari et al. 1995; Nowzari & Slots 1995). Bone debris produced during implant surgery has been collected and used for immediate implantation around implants with inadequate bone in the ‘‘simultaneous augmentation technique’’ (Lauer & Schilli 1994). The bacterial contamination of membranes during their insertion has already been established (Yoshinari et al. 1998; De Sanctis et al. 1996; Nowzari et al. 1995). In this context, the use of collected bone debris (CBD) from the oral

Young et al . Chlorhexidine: its effects on bacteria found in bone debris

cavity risks iatrogenic contamination of the inserted implants and associated barrier membranes. Recently, it has been established that CBD contains significant levels of bacteria but that the use of a stringent aspiration protocol can significantly reduce the levels of bacterial contamination (Young et al. 2001). Oral rinses containing chlorhexidine have long been recognised to be effective in reducing salivary bacterial levels (Altonen et al. 1976) and intra-alveolar dressings containing chlorhexidine have been shown to lower the incidence of alveolar osteitis associated with the removal of impacted third molars (Fotos et al. 1992). Furthermore, the use of a pre-operative chlorhexidine rinse has been shown to

be effective in reducing the incidence of oral infections associated with dental implants and to improve implant success rates (Lambert et al. 1997). The aim of this study was to establish whether an immediately pre-surgical, chlorhexidine mouthrinse reduces the levels of microbial contaminants of CBD further beyond that achieved by a stringent bone collection protocol alone.

Material and methods The Dental Ethics Committee at the University Dental Hospital of Manchester approved this study.


Twenty volunteers (10 men and 10 women) with a mean age of 46 years were recruited to participate in this study. Individuals were excluded from participating in this study if, following a detailed clinical examination, it was concluded that implant insertion would necessitate bone augmentation. Comprehensive medical histories were obtained and no subject presented a contraindication to minor oral surgery procedures. Furthermore, no patient was taking medication at the time of implant surgery. Patients were entered into this study if they were partially dentate and had no clinical or radiographic evidence of active oral disease. Group allocation

Twenty patients were randomly allocated into one of two equal groups on attending for implant surgery. All patients used an oral rinse for a timed duration of 2 min. Group T: Immediately prior to surgery, patients in this group carried out an oral rinse using 10 ml of chlorhexidine-containing oral rinse (CorsodylA, Smithkline-Beecham, Middlesex, United Kingdom). Group C: Immediately prior to surgery, patients in this group carried out an oral rinse using 10 ml of sterile water. A stringent bone collection protocol (restricted to within the surgical site only) was employed for bone collection in both

Fig. 1. Sample processing.

Fig. 2. Overview of sample processing for microbial analysis of bone debris collected during dental implant surgery.

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Young et al . Chlorhexidine: its effects on bacteria found in bone debris

groups. For both groups, salivary control was achieved utilising a separate, dedicated, sterile suction-tip and tubing. Clinical procedures

All surgeries were carried out in a dedicated, outpatient operating room. The operators and assistants scrubbed and wore sterile gowns, caps and gloves. Patients were covered with full-body, sterile disposable drapes and hats (Unitek 3M, Bradford, United Kingdom). The implant surgeries in these patients were carried out by one of the authors (MPJY) and experienced assistants carried out tissue-fluid control and bone aspiration. The Frialit-2A system (Friadent GmbH, Mannheim, Germany) was used according to the manufacturer’s instructions. During the preliminary soft-tissue surgery, a sterilised metal suction tip was used for tissue-fluid control (American pattern suction tipA, B.Braun-Aesculap, Essex, United Kingdom). Local anaesthesia was administered (ScandonestA, Deproco U.K. Ltd, Surrey, United Kingdom) with a minimum wait of 15 minutes allowed to ensure that adequate anaesthesia and vasoconstriction was achieved. A mucoperiosteal flap was raised, retracted and gently sutured to the adjacent mucosa. Immediately prior

to the preparation of the implant sites, a Frios bone trapA was connected to central suction by an aseptic technique. Bone collection for both groups was carried out using a stringent bone collection technique during the preparation of two maxillary endosseous dental implant sites. This aspiration protocol rigidly restricted the suction tip to collect bone, blood and irrigant only from directly within the surgical site. Salivary control was achieved utilising a separate suction (Young et al. 2001). On completion of implant insertion, and prior to wound closure, the bone sample was removed from the bone trap by means of a sterile curette. Samples were placed aseptically into a screw-capped, 7 ml, sterile, glass bottle containing 1 ml of freshly prepared, reduced transport fluid (RTF). Samples reached the laboratory within 3 to 5 min and the processing of samples commenced immediately. Laboratory procedures

An overview of processing procedures is shown in Figs 1 and 2. Each bone sample in RTF mixture was sonicated for 1 min in an ultra-sonic bath (ULT 300A, Wright Dental Company Ltd., Glamorgan, United Kingdom). Each sample was then agitated with a sterile 1 mm plastic

Pasteur pipette. Microbial analysis was undertaken of samples derived from the supernatant from uncrushed bone, from the supernatant fluid from crushed bone and from neat crushed bone. Bone sampling: supernatant

The bone-RTF mixture was allowed to stand and when it had settled an aliquot (200 ml) was removed from the supernatant solution and transferred to a sterile 7 ml, screw-capped, glass bottle. Phosphate buffer (pH 7.4, 0.01 m) in saline was used to prepare a ten-fold dilution series (10ª1 to 10ª6). Aliquots (10 ml) were removed from each ten-fold dilution and plated out separately on both Columbia-blood agar (CBA) and fastidious anaerobe agar (FAA) for aerobic and anaerobic incubation, respectively, at 37æC for 48 h. The remainder of the supernatant (800 ml) was divided into 2 equal aliquots. Each aliquot was transferred to a separate, sterile 7 ml screwcapped, glass bottle and passed through separate filter membranes (0.2 mm pore filter membrane) housed in a pre-sterilised unit (Whatman filtration unitA, Whatman International Ltd, Kent, England). A water-vacuum pump was used to effect the filtration. The filter units were opened using an aseptic technique

Fig. 3. Species detected in collected bone debris and number of samples from each group that contained these bacteria.

Species identified

Group T (chlorhexidine)

Group C (placebo)

(a) Species found only in group T Dermatococcus nishinomyaensis Fusobacteria sp. Gemella sp. Gemella morbillorum Staphylococcus sp. Staphylococcus aureus Wolinella sp.

3 1 2 3 2 1 1

0 0 0 0 0 0 0

(b) Species found only in group C Actinomyces odontolyticus Aerococcus viridans Clostridium bifermentans Clostridium tyrobutyricum Enterococcus faecalis Gemella haemolysans Micrococcus kristinae Micrococcus luteus Micrococcus lylae Micrococcus nishinomiyaensis Prevotella intermedia Staphylococcus capitis

0 0 0 0 0 0 0 0 0 0 0 0

1 2 1 1 1 1 1 1 1 1 1 3

* 0Ωspecies not detected

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Clin. Oral Impl. Res. 13, 2002 / 20–29

Species identified

Group T (chlorhexidine)

Group C (placebo)

Staphylococcus chromogenes Staphylococcus saccharolyticus Stomatococcus mucilaginosus Streptococcus anginosus Streptococcus oralis Streptococcus vestibularis Veillonella sp.

0 0 0 0 0 0 0

1 1 2 1 2 3 2

(c) Species found in both groups Fusobacterium nucleatum Leuconostoc sp. Micrococcus sp. Micromonas micros Propionibacterium propionicum Peptostreptococcus asaccharolyticus Peptostreptococcus prevotii Staphylococcus epidermidis Staphylococcus sciuri Streptococcus sp. Streptococcus mitis Streptococcus parasanguis Streptococcus salivarius ssp. salivarius

3 2 1 1 1 2 1 3 1 1 3 1 1

1 3 1 3 3 1 1 1 1 4 3 3 3

Young et al . Chlorhexidine: its effects on bacteria found in bone debris

and each membrane was removed using sterile tweezers. One filter was plated out onto CBA and one filter plated out onto FAA. They underwent aerobic and anaerobic incubation, respectively, at 37æC for 48 h.

Fig. 4. Species detected and the colony forming units produced by supernatant from uncrushed bone (SN), crushed bone (CB) and total (CBD) Colony-Forming Units (CFU*) Mouthrinse protocol Test group T

Species (chlorhexidine)

Sample T1

Gemella sp. Streptococcus mitis Micrococcus sp.

2 0.2 0.1 2.3

8 0.4 0.4 8.8

Sample T2

Fusobacterium sp. Fusobacterium nucleatum Staphylococcus epidermidis Wolinella sp.

0.02 0.01 0.2 0.02 0.25

0.08 0.04 0.8 0.04 0.96

0.1 0.05 1 0.06 1.21

Sample T3

Dermatococcus nishinomiyaensis Dermatococcus nishinomiyaensis Gemella morbillorum Staphylococcus sp. Staphylococcus sp. Staphylococcus epidermidis

0.1 0.2 0.3 0.1 0.1 0.1 0.9

0.8 0.8 1.6 0.4 0 0.4 4.0

0.9 1.0 1.9 0.5 0.1 0.5 4.9

Sample T4

Gemella morbillorum Leuconostoc sp. Peptostreptococcus asaccharolyticus Staphylococcus epidermidis

0.1 0.2 0.2 0.2 0.7

0.4 0.4 0.4 0.8 2.0

0.5 0.6 0.6 1.0 2.7

Sample T5

Dermatococcus nishinomiyaensis Fusonucleatum sp. Unidentified Gram-positive Unidentified Gram-negative

1 0.1 0.1 0.2 1.4

0 0 0.8 1.2 2.0

1 0.1 0.9 1.4 3.4

Sample T6

Streptococcus sp. Gemella sp. Leuconostoc sp

3 3 2 8

12 8 8 28

15 11 10 35

Sample T7

Propionibacterium propionicum Streptococcus mitis Streptococcus salivarius

3 4 3 10

8 12 12 32

11 16 15 42

Sample T8

Peptostreptococcus prevotii Staphylococcus parasanguis Staphylococcus sciuri Unidentified Gram-negative anaerobe

0.03 0.1 0.2 0.01 0.34

0.08 0.8 0.8 0.08 1.76

0.11 0.9 1.0 0.09 2.1

Sample T9

Gemella morbillorum Micromonas micros Peptostreptococcus asaccharolyticus

0.2 0.01 0.02 0.23

0.4 0.04 0.04 0.48

0.6 0.05 0.06 0.71

Sample T10

Fusobacterium nucleatum Staphylococcus aureus Streptococcus mitis

0.1 0.1 0.1 0.3

0.4 0.6 0.8 1.8

0.5 0.7 0.9 2.1

Ground bone debris: laboratory analysis

The sample of bone (minus the supernatant) was transferred to a sterile mortar and pestle and divided into half. One half was further divided into half again for plating out onto FAA and CBA, respectively. The remaining half was thoroughly ground for 2 minutes, then 2 ml of phosphate buffer (pH 7.4, 0.01 m in 0.8% w/v NaCl solution) was used to wash the mortar and pestle and the sample was transferred to a sterile 30 ml screwcapped plastic bottle. The crushed bone sample was sonicated for 15 min using an ultrasonic bath and then agitated using a sterile pipette. After settling, aliquots of 100 ml were removed and underwent serial dilution (10ª1 to 10ª6). The 10ª6 dilution (10 ml aliquots) were plated out for aerobic and anaerobic incubation (vide infra). The remaining bone suspension was divided equally into 2 plastic bottles and filtered. This stage was devised to ensure that bacteria present on the surface or within the surface irregularities were made available for isolation, cultivation and then identification. Bone sampling: neat bone debris

The remaining neat, crushed bone was divided into equal portions and plated out onto CBA and FAA plates and incubated as described previously, in case only very low numbers of bacteria were present. Microbial counts

Samples from all the serial dilutions, from both supernatant, crushed bone and from the neat bone were plated out and the number of colony-forming units (CFUs) was counted for each sample following incubation as described previously. The total CFU per sample of CBD was calculated by adding together the CFU for the supernatant from uncrushed bone (SN) to two times the CFU for half the crushed bone (CB), in order to yield a total CFU per ml of CBD. (The reason for doubling the CFU for crushed bone was to compensate for half already having been used).

Microbial culture and sub-culture

Samples for aerobic incubation were plated out onto CBA, then incubated aerobically at 37æC for 48 h. For anaerobic incubation, samples were plated out onto pre-reduced, FAA containing 5%



CBD 10 0.6 0.5 11.1

horse blood and placed into the anaerobic work-station (Modular Atmospheric Control SystemA or ‘‘MACS’’, Don Whitley Scientific, Shipley, United Kingdom). When microbial growth was evident, the isolates were examined

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Young et al . Chlorhexidine: its effects on bacteria found in bone debris

macroscopically for colour, texture and marginal integrity. Sub-culturing was carried out in order to obtain pure colonies for identification and to ensure propagation until identification was complete. Identification of microbes was achieved utilising colonial morphology, Gram stain, gaseous requirements, catalase test, and commercially available identification kits, specifically API StaphA, ID 32 StrepA, Rapid Ana IIA, ID 32AA, and Rapid NHA (bioMe´rieux, Hampshire, United Kingdom).

Fig. 4. Continued Colony-Forming Units Mouthrinse protocol Control group C

Species (sterile water)

Sample C1

Enterococcus faecalis Leuconostoc sp. Micromonas micros Propionibacterium propionicum Streptococcus sp.

Sample C2

Clostridium tyrobutyricum Fusobacterium nucleatum Micrococcus lylae Peptostreptococcus prevotii Streptococcus mitis Streptococcus parasanguis

Sample C3

Leuconostoc sp. Micrococcus sp. Micromonas micros Streptococcus mitis Streptococcus parasanguis Unidentified Gram-positive F. anaerobe

6 4 3 3 5 4 25

20 12 10 12 24 8 86

26 16 13 15 29 12 111

Sample C4

Staphylococcus saprophyticus Micrococcus lylae Streptococcus sp. Staphylococcus saccharolyticus Peptostreptococcus asaccharolyticus

2 6 4 6 10 28

0 8 6 10 14 38

2 14 10 16 24 66

Sample C5

Clostridium bifermentans Leuconostoc sp. Micromonas micros Staphylococcus capitis Staphylococcus sciuri Streptococcus anginosus Streptococcus oralis Streptococcus parasanguis Unidentified Gram-positive aerobe

4 14 3 2 4 3 6 4 3 43

8 28 4 12 12 12 16 12 8 112

12 42 7 14 16 15 22 16 11 155

Sample C6

Staphylococcus capitis Staphylococcus chromogenes Streptococcus sp. Streptococcus mitis Unidentified Gram-negative anaerobe

2 2 4 3 3 14

12 8 8 8 12 48

14 10 12 11 15 62

Statistical analysis

Sample C7

The bacterial levels (mean colony-forming units) between the two groups were compared using the Mann-Whitney (non-parametric) test for comparing two independent samples, at a 0.05 level of significance.

Aerococcus viridans Micrococcus nishinomiyaensis Streptococcus sp. Streptococcus vestibularis Stomatococcus mucilaginosus Veillonella

Sample C8

Micrococcus kristinae Staphylococcus epidermidis Stomatococcus mucilaginosus Streptococcus oralis


So that the same strains would not be repeatedly identified from individual samples, colonies were screened by means of Gram morphology and antibiograms. The latter would not be expected to yield identical zone sizes for all antibiotics if the strains being cultured were different. Isolates were lawninoculated onto CBA and FAA agar as appropriate and Mast-rings added (Mast, Liverpool, United Kingdom). The Mastrings used contained the following antibiotics: ampicillin, cephalothin, colistin sulphate, gentamycin, streptomycin, sulphathiad, tetracycline and cotrimoxazole, for Gram-negative bacteria. Chloramphenicol, erythromycin, fusidic acid, methicillin, novobiocin, penicillin G, streptomycin and tetracycline on a ring were used for Gram-positive bacteria. Catalase test

One loopful of the microbial isolates was placed onto a glass slide in a Petri dish. One drop of hydrogen peroxide (3% v/v) was added to the sample and effervescence sought (Cowan & Steel 1974).

Results All samples from the supernatant from uncrushed bone and from the crushed bone yielded viable microorganisms. The microorganisms identified in col-

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Clin. Oral Impl. Res. 13, 2002 / 20–29

SN 4 2 5 3 4 18 0.3 0.2 0.5 0.4 0.5 0.3 2.2

0.4 0.1 0.2 0.3 0.1 0.3 1.4 5 4 5 7 21

CB 12 8 24 4 16 54 0.8 0.8 1.6 1.2 1.2 1.6 7.2

1.2 0.8 1.2 0.4 0.4 0.5 4.5 16 16 20 28 80

CBD 16 10 29 7 20 82 1.1 1.0 2.1 1.6 1.7 1.9 9.4

1.6 0.9 1.4 0.7 0.5 0.8 5.9 21 20 25 35 101

Young et al . Chlorhexidine: its effects on bacteria found in bone debris

Fig. 4. Continued Colony-Forming Units Mouthrinse protocol Control group C

Species (sterile water)

Sample C9

Gemella haemolysans Peptostreptococcus asaccharolyticus Propionibacterium propionicum (a) Propionibacterium propionicum (b) Streptococcus sp. Streptococcus vestibularis

2 3 3 1 2 4 15

4 12 10 0 4 8 38

6 15 13 1 6 12 53

Sample C10

Actinomyces odontolyticus Leuconostoc sp. Staphylococcus capitis Streptococcus vestibularis Veilonella sp.

5 3 3 6 2 19

16 4 12 20 12 64

21 7 15 26 14 83




* CFU refers to colony-forming units calculated at 10ª3 dilution. The CFU per bone samples from this study were detected and counted at a wide range of dilutions. For the purpose of this study, the CFU are presented at a standardised dilution of 10ª3 standardised. This accounts for the wide variation in the CFU values seen. See text for experimental details.

lected bone debris and the number of samples that were positive for each species are presented for both groups (T and C) and listed in Fig. 3. Species isolated only from test Group T (chlorhexidine rinse) samples were: Dermatococcus nishinomyaensis, Fusobacterium sp., Gemella sp. (including Gemella morbillorum), Staphylococcus sp., (including Staphylococcus aureus) and Wolinella species. Species isolated only from control Group C samples (distilled water rinse) were: Actinomyces odontolyticus, Aerococcus viridans, Clostridium bifermentans, Clostridium tyrobutyricum, Enterococcus faecalis, Gemella haemolysans, Micrococcus kristinae, Micrococcus luteus, Micrococcus lylae, Micrococcus nishinomiyaensis, Prevotella intermedia, Staphylococcus sp. including Staphylococcus capitis, Staphylococcus chromogenes, Staphylococcus saccharolyticus, Stomatococcus mucilaginosus, Streptococcus sp. (Streptococcus anginosus, Streptococcus oralis, Streptococcus vestibularis) and Wolinella species. Finally, the species isolated from both groups were: Fusobacterium nucleatum, Leuconostoc sp., Micrococcus sp., Micromonas micros, Propionibacterium propionicum, Peptostreptococcus asaccharolyticus, Peptostreptococcus prevotii, Staphylococcus epidermidis and Staphylococcus sciuri. Streptococcus

species found in both groups included Streptococcus mitis, Streptococcus parasanguis and Streptococcus salivarius ssp. salivarius. Fig. 4 compares the microbial flora and levels of bacteria of samples from groups T and C, for each sample in turn. These data were used to calculate the total levels of bacterial contaminants present for each bone sample. With regard to collected bone debris (CBD), the mean CFU derived from test Group T samples (chlorhexidine rinse) was 0.72¿105 CFU per bone sample (SD 0.74) compared to 3.43¿105 CFU (SD 2.26) for control Group C (sterile water rinse), P⬍0.001.

Discussion In both this study and our previous microbiological study on collected bone debris (Young et al. 2001), bone collection was carried out using a separate dedicated, sterile suction line to minimise salivary and hence bacterial contamination. This study demonstrated that the adjunctive use of a 2-minute preoperative chlorhexidene mouthrinse further reduces the bacterial contaminants of collected bone debris beyond that achieved when a stringent bone collection protocol is used alone (Young et al. 2001). The data from these two independent studies revealed consistent

levels of bacterial contamination when bone is collected utilising a stringent aspiration protocol alone. Specifically, a mean CFU of 3.43¿105 was found per sample of CBD collected with a stringent collection protocol in this study, compared with mean CFU of 4.07¿105 in our previous study (Young et al. 2001). The slightly lower contamination levels for this latter study could be explained by the use of a 10 ml sterile water rinse prior to surgery and to the increased experience of the investigating team with this stringent bone collection technique. While it is widely recognised that preparatory techniques have a profound influence on the nature of bone mineral, the effects of chlorhexidine on the osteogenic potential of CBD are not yet known. Atropine sulphate is a wellknown anti-sialogogue (Sherman & Sherman 1999) and, in conjunction with the protocols we have already designed and tested, might further reduce the bacterial contamination of CBD. With regards to specific bacterial contaminants found in this study, their known clinical relevance is shown (Fig. 5) and several isolates are recognised pathogens that merit special consideration. In relation to bone samples isolated only from the test group T (chlorhexidine), Staphylococcus aureus was detected in one sample only. However, it has been shown to be a prime causative organism in infectious endocarditis (Ruiz Jr. et al. 2000) and sinusitis with spread to cause orbital cellulitis (Haddadin et al. 1999). Wolinella species was also found in one sample only from the test group but has been associated with refractory endodontic cases (Vigil et al. 1997). With regards to the control group C (sterile water rinse), the most unusual findings were the presence of Clostridum species, a rare oral isolate, in two samples from the control group. Clostridium bifermentans has been reported with serious but extremely rare complications such as metastatic osteomyelitis (Scanlan et al. 1994) and metastatic liver abscess (Nachman et al. 1989). Infectious complications might be linked to the colonisation of implants by bacteria and any associated ‘‘biofilm’’ formation. A biofilm has been described as ‘‘a functional consortium of micro-

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Clin. Oral Impl. Res. 13, 2002 / 20–29

Young et al . Chlorhexidine: its effects on bacteria found in bone debris

Fig. 5. Known clinical relevance of bacterial species isolated from CBD in this study Bacterial species

Known clinical relevance


Actinomyces odontolyticus Aerococcus viridans Clostridium bifermentans

Infections of teeth and bone Unknown Osteomyelitis Metastatic liver abscess Endocarditis Cranial abscess Unknown Unknown Dental root canal infection Colonisation of PTFE membranes Rapidly-progressing periodontitis Periodontal disease Papillon Lefevre syndrome (PLS) PLS PLS PLS PLS PLS Human periodontitis Human periodontitis

Summanen et al. 1993

Clostridium tyrobutyricum Dermatococcus nishinomyaensis Enterococcus faecalis Fusobacterium nucleatum

Leuconostoc sp. Micrococcus spp. M. kristinae M. luteus M. lylae M. nishinomiyaensis Micromonas micros Prevotella intermedia

Deep peri-implant bone defects Propionibacterium spp. Propionibacterium propionicum Peptostreptococcus sp. P. asaccharolyticus P. prevotii Staphylococcus spp. S. aureus S. capitis S. chromogenes S. epidermidis S. saccharolyticus S. sciuri Stomatococcus mucilaginosus Streptococcus sp. S. anginosus S. oralis S. parasanguis S. salivarius ssp. salivarius S. vestibularis Veillonella sp. Wolinella sp.

organisms organised within an extensive exopolymer matrix’’ (Elder et al. 1995). The adhesive property of the ‘‘glycocalyx’’ (exopolymer) plays a key role in the development of biofilms and its production is known to be species-dependent and strain-dependent (Wright et al. 1997). Biofilms have been shown to permit bacteria to survive antimicrobials in minimum inhibitory concentration (MIC) up to 1000 times greater than the MIC for the same strain in batch culture

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Clin. Oral Impl. Res. 13, 2002 / 20–29

Scanlan et al. 1994 Nachman et al. 1989 Kolander et al. 1989 Pencek & Burchiel 1986

Gomes et al. 1994 Sbordone et al. 1999 Fosse et al. 1999 Roberts 2000 Robertson 1999 Robertson 1999 Robertson 1999 Robertson 1999 Robertson 1999 Robertson 1999 Rams et al. 1992 Fosse et al. 1999 Clerehugh et al. 1997 Auguthun & Conrads 1997

Endocarditis Osteomyelitis Endodontic infection Ailing implants Endodontic bacteraemia Colonisation of catheters Colonisation of implants Endocarditis Orthopaedic prosthesis infection Endocarditis Not known Pin-track infections Not known Soft tissue infections Not known

Finegold et al. 1992 Summanen et al. 1993 Gomes et al. 1996 Rams et al. 1991 Debelian et al. 1995 Goldmann & Pier 1993 Barth et al. 1989 Chang 2000 Kalmeijer et al. 2000 al-Rashdan et al. 1998

Abscesses Dental caries Not known Dental caries Dental caries Dental and head & neck infections Refractory endodontic infection

Whiley et al. 1992 Wilcox et al. 1987

(Wright et al. 1997; Elder et al. 1995). Their presence can confuse the accurate diagnosis of causative organisms in clinical infection around implanted devices and present a most difficult diagnostic and treatment challenge (Gristina & Costerton 1985). Biofilms appear to permit bacteria to go undetected by the host for prolonged periods even in the presence of normal host defence mechanisms. While it is acknowledged that biofilms can safely co-exist with their

Elliott 1996 Marsou et al. 1999

Drucker et al. 1984 Wilcox et al. 1991 Summanen et al. 1993 Vigil et al. 1997

hosts for long periods, it is still not precisely understood why biofilms become ‘‘active’’ and contribute to clinical infection around implanted devices. It is quite feasible that biofilm ‘‘activation’’ plays a key role in late failures of endosseous dental implants. The importance of minimising the prevalence of infectious complications around implants is compounded by the fact that there is limited scientific evidence to support treatment strategies that are currently

Young et al . Chlorhexidine: its effects on bacteria found in bone debris

employed to stabilise failing implants (Esposito et al. 1999). With regards to oral implants and bone augmentation, infection is widely recognised to be associated with unfavourable clinical outcomes (Nowzari et al. 1996; Becker et al 1990; Mombelli et al. 1987). Whilst the precise origin of bacteria present in clinical infection around implanted devices has been debated, commensal organisms of the adjacent tissues have been cited as a prime source (Lewis & Sherertz 1997). As a consequence, skin preparatory techniques have been recommended to reduce levels of commensal organism prior to surgery and have a long and safe history of use (Sato et al. 1996). In a similar context, chlorhexidine is effective in the reduction of bacteria in the oral cavity (Altonen et al. 1976) and has recently been shown to reduce infectious complications associated with endosseous dental implants (Lambert et al 1997). Given this background, if CBD is collected for the purpose of immediate implantation, the use of a pre-surgical chlorhexidine oral rinse to minimise the contaminants found in CBD is strongly recommended.

Conclusions Our clinical studies have shown that stringent clinical protocols can exert a profound reduction in the levels and composition of the bacterial contaminants of CBD. This study strongly supports the use of a preoperative chlorhexidine oral rinse in conjunction with a stringent bone collection protocol if the CBD is to be implanted. Conversely, the implantation of heavily contaminated bone debris in the simultaneous augmentation technique is contra-indicated since it risks both minor infections and major (but extremely rare) infectious complications. For the reasons discussed in this paper, surgical implantation techniques should continue to be refined with the aim of further reducing the initial bacterial contamination of the implanted materials and devices (Pereira et al. 1999, Costerton et al. 1993). Acknowledgements: We would like to particularly thank Dr Igor Blum and Dr Shahram Tabibi for their assistance during

the clinical part of this research study. We are grateful to Mr R. Lloyd and Mr B. Musgrove (Unit of Oral & Maxillofacial Surgery), Mr Eldridge and Mr G. Smith (Unit of Prosthodontics) at the Turner Dental School, Manchester (UK) for permitting patients under their care to be included in this clinical research study. Research support was kindly provided by the Central Manchester Health Care NHS Trust (Manchester, England, Grant . G348), Friadent GmbH (Mannheim, Germany) and General Medical Equipment Co. Ltd., (Devizes, England).

protokolls das Ausmass der Kontamination reduziert werden kann. Die Chlorhexidinmundspu¨lung ist eine gut dokumentierte antibakterielle Spu¨lung, mit welcher die infektio¨sen Komplikationen, die bei dentalen Implantaten auftreten ko¨nnen, reduziert werden ko¨nnen. Diese Studie untersuchte den Einfluss einer pra¨opertiven Mundspu¨lung mit 0.1% Chorhexidinigluconat-Spu¨llo¨sung auf die bakterielle Kontamination von gesammeltem Knochenabrieb (CBD). Zwanzig teilbezahnte Patienten wurden zufa¨llig in zwei gleiche Gruppen aufgeteilt. Bei der Eingliederung von zwei dentalen Implantaten wurden mittels dem Frios KnochenkollektorA Knochenspa¨ne gesammelt. In der Gruppe T wurde pra¨operativ eine Chlorhexidinspu¨lung verwendet, wa¨hrend bei der Gruppe C mit sterilem Wasser gespu¨lt wurde. Bei beiden Gruppen wurde ein strenges Absaugprotokoll angewendet. Die Knochenproben wurden sofort zur mikrobiologischen Analyse u¨berfu¨hrt. Fu¨r die Identifizierung der isolierten Bakterien wurden die Morphogie der Kolonien und

Re´sume´ La chirurgie implantaire produit des de´bris osseux qui peuvent eˆtre utilise´s dans la technique d’augmentation osseuse simultane´e. Bien que ces de´bris soient contamine´s par les bacte´ries buccales, un protocole d’aspiration pre´cis a prouve´ qu’il pouvait re´duire les niveaux de contamination. Le bain de bouche a` la chlorhexidine est un antibacte´rien connu qui a prouve´ qu’il re´duisait les complications infectieuses associe´es aux implants dentaires. Cette e´tude a examine´ l’effet du rinc¸age pre´ope´ratoire avec 0.1% de digluconate de chlorhexidine sur des contaminants bacte´riens pre´sents dans les de´bris osseux collecte´s (CBD). Vingt patients partiellement e´dente´s ont e´te´ re´partis au hasard en groupe e´gaux et ont subi une collection osseuse utilisant le Frios Bone CollectorA durant l’insertion de deux implants dentaires. Dans le groupe T, un rinc¸age pre´ope´ratoire a` la chlorhexidine a e´te´ pratique´ tandis que dans le groupe C le rinc¸age a e´te´ effectue´ avec de l’eau ste´rile. Pour les deux groupes le protocole de collecte osseuse pre´cite´ a e´te´ utilise´. Les e´chantillons osseux ont ime´diatement e´te´ transporte´s pour une analyse microbiologique. La morphologie des colonies et microscopique, les ne´cessite´s en gaz et les kits d’identification ont e´te´ utilise´s pour l’identification des microbes isole´s. Trente-neuf espe`ces ont e´te´ identifie´es incluant un nombre d’espe`ces associe´es a` la maladie en particulier l’Actinomyces odontolyticus, le Clostridium bifermentans, le Prevotella intermedia et le Propionibacterium propionicum. Les e´chantillons du groupe T avaient significativement moins d’organismes (P∞0.001) que ceux du groupe C. Les coques Gram positif dominaient les isolats dans les deux groupes. Si des de´bris osseux doivent eˆtre utilise´s lors d’une technique chirurgicale d’augmentation osseuse simultane´e, un bain de bouche a` la chlorhexidine avant l’ope´ration devrait eˆtre utilise´ en association avec un protocole strict d’aspiration afin de re´duire la contamination bacte´rienne des CBD.

Zusammenfassung Wa¨hrend der Implantatchirurgie entsteht Knochenabrieb, welcher bei der ‘‘gleichzeitigen Augmentationstechnik’’ verwendet werden kann. Obwohl dieser Abrieb mit oralen Bakterien kontaminiert ist, konnte gezeigt werden, dass mittels eines strengen Absaug-

der Keime unter dem Mikroskop, der Gasbedarf und Identifikationskits verwendet. Es konnten 39 Arten identifiziert werden, eine gewisse Anzahl davon wird mit Erkrankungen assoziiert, na¨mlich Actinomyces odontolyticus, Clostridium bifermentans, Prevotella intermedia und Propionibacterium propionicum. Proben der Gruppe T zeigten (Chlorhexidinmundspu¨lung) signifikant weniger Organismen (P∞0.001) als Proben der Gruppe C (Spu¨lung mit sterilem Wasser). Grampositive Kokken dominierten die isolierten Keime in beiden Gruppen. Es wird die Schlussfolgerung gezogen, dass, falls Knochenabrieb fu¨r die gleichzeitige Knochenaugmentation verwendet wird, eine pra¨operative Chorhexidinmundspu¨lung zusammen mit einem strikten Absaugprotokoll verwendet werden sollte, um eine weitere bakterielle Kontamination des CBD zu reduzieren.

Resumen La cirugı´a de implantes dentales produce escombros de hueso que se pueden usar en la te´cnica de ‘‘aumento simultaneo’’. Aunque estos escombros esta´n contaminados con bacterias orales, un protocolo de aspiracio´n riguroso ha demostrado que reduce los niveles de contaminacio´n. El colutorio de clorhexidina es un enjuague antibacteriano bien probado que ha demostrado que reduce las complicaciones infecciosas asociadas con implantes dentales. Este estudio examino´ el efecto del enjuague preoperatorio con un colutorio de digluconato de clorhexidina al 0.1% en los contaminantes bacterianos presentes en los escombros de hueso recogidos (CBD). Se asignaron aleatoriamente a grupos iguales veinte pacientes parcialmente ede´ntulos y se sometieron a recoleccio´n de hueso usando el colector de hueso FriosA (FBC) durante la insercio´n de dos implantes dentales. En el grupo T se uso´ un colutorio de clorhexidina preoperativamente, mientras que en el grupo C se uso´ agua este´ril. Para ambos grupos se uso´ un protocolo de aspiracio´n riguroso. Las muestras de hueso se transportaron inmediatamente para ana´lisis microbiano. Para la identificacio´n de los microbios aislados se uso´ morfologı´a colonial y microsco´pica, requerimientos gaseosos y kits de identificacio´n. Se identificaron treinta y nueve especies incluyendo un nu´mero asociado a enfermedad, en particular Actinomyces odontolyticus, Clostridium bifermentans, Prevotella intermedia y Propionibacterium propionicum. Las muestras del grupo T (colutorio de clorhexi-

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Young et al . Chlorhexidine: its effects on bacteria found in bone debris

dina) produjeron significativamente menos organismos (P∞0.01) que las del grupo C (colutorio de agua este´ril). Los cocos gram positivos dominaron los aislamientos de ambos grupos. Se concluye que si los escombros de hueso se van a usar con el propo´sito inmediato de aumento simultaneo, se debe utilizar un colutorio de clorhexidina en conjunctio´n con un protocolo de aspiracio´n riguroso para reducir una posterior contaminacio´n bacteriana.

References Adell, R., Lekholm, U & Branemark, P.I. (1985) Tissue integrated prostheses. Surgical procedures. 1st ed, pp. 211–232. Chicago: Quintessence.

surgery: a study of 2,642 implants. Journal of Oral & Maxillofacial Surgery 55: 19–24. De Sanctis, M., Zucchelli, G. & Clauser, C. (1996) Bac-

al-Rashdan, A., Bashir, R., Khan, F.A. (1998) Staphylo-

terial colonisation of bioabsorbable barrier material

coccus capitis causing aortic valve endocarditis.

and periodontal regeneration. Journal of Periodonto-

Journal of Heart Valve Disease 7: 518–520.

logy 67: 1193–1200.

Altonen, M., Saxen, L, Kosunen, T. & Ainamo, J (1976)

Drucker, D.B., Shakespeare, A.P. & Green, R.M. (1984)

Effect of two antimicrobial rinses and oral prophy-

The production of dental plaque and caries by the

laxis on preoperative degerming of saliva. Int J Oral

bacterium Streptococcus salivarius in the gnotobi-

Surg 5: 276–284.

otic WAG/RIJ rat. Archives of Oral Biology 29: 437–

Auguthun, M. & Conrads, G. (1997) Microbial findings of deep peri-implant bone defects. Interna-

443. Elder, M.J., Stapleton, F., Evans, E. & Dart, J.K. (1995)

tional Journal of Oral and Maxillofacial Implants

Biofilm-related infections in ophthalmology. Eye 9:

12: 106–112.


Barth, E., Myrvik, Q.M., Wagner, W. & Gristina, A.G.

Elliott J. (1996) The microbiology of problem frac-

(1989) In vitro and in vivo comparative colonization

tures. Master of Science thesis, The University of

of Staphylococcus aureus and Staphylococcus epidermidis on orthopaedic implant materials. Biomaterials 10: 325–328.

Manchester. Esposito, M., Hirsch, J., Lekholm, U. & Thomsen, P. (1999) Differential diagnosis and treatment strat-

Becker, W., Becker, E.B., Newman, M.G., Nyman, S.

egies for biologic complications and failing oral im-

(1990) Clinical and microbiologic findings that may

plants: a review of the literature. International

contribute to dental implant failure. International

Journal of Oral and Maxillofacial Implants 14: 473–

Journal of Oral and Maxillofacial Implants 5: 31– 38.

490. Finegold, S.M., Baron, E.J. & Wexler, H.M. (1992) A

Chang, F.Y. (2000) Staphylococcus aureus bacteremia and endocarditis. Journal of Microbiological and Immunological Infection 33: 63–68. Clerehugh, V., Seymour, G.J., Bird, P.S., Cullinan, M., Drucker, D.B. & Worthington, H.V. (1997) The de-

clinical guide to anaerobic infection. Belmont, CA: Star Publishing Co., p. 34. Fosse, T., Madinier, I., Hitzig, C. & Charbit, Y. (1999) Prevalence




among 149 anaerobic gram-negative rods isolated

tection of Actinobacillus actinomycetemcomitans,

from periodontal pockets. Oral Microbiology and

Porphyromonas gingivalis and Prevotella interme-

Immunology 14: 352–357.

dia using an ELISA in an adolescent population

Fotos, P.G., Koorbusch, G.F., Sarasin, D.S. & Kist, R.J.

with early periodontitis. Journal of Clinical Period-

(1992) Evaluation of intra-alveolar chlorhexidine

ontology 24: 57–64.

dressings after removal of impacted mandibular

Costerton, J.W., Khoury, A.E., Ward, K.H. & Anwar, H. (1993) Practical measures to control device-related bacterial infections. International Journal of Artificial Organs 16: 765–770. Cowan, S.T. & Steel, K.J. (1974) Cowan and Steel’s

third molars. Oral Surgery, Oral Medicine and Oral Pathology 73: 383–388. Goldmann, D.A. & Pier, G.B. (1993) Pathogenesis of infections related to intravascular catheterization. Clinical Microbiology Reviews 6: 176–192.

manual for the identification of medically import-

Gomes, B.P., Lilley, J.D. & Drucker, D.B. (1994) Posi-

ant bacteria 2nd edition , pp. 27–28. London: Cam-

tive and negative associations between bacterial

bridge University Press.

species in dental root canals. Microbios 80: 231–

Debelian, G.J., Olsen, I. & Tronstad, L. (1995) Bacteremia in conjunction with endodontic therapy. Endodontics and Dental Traumatology 11: 142–149. Dent, C.D., Olson, J.W., Farish, S.E., Bellome, J., Casino, A.J., Morris, H.F. & Ochi, S. (1997) The influ-

243. Gomes, B.P., Lilley, J.D. & Drucker, D.B. (1996) Associations of endodontic symptoms and signs with particular combinations of specific bacteria. International Endodontic Journal 29: 69–75.

ence of preoperative antibiotics on success rates of

Gristina, A.G. & Costerton, J.W. (1985) Bacterial ad-

endosseous implants up to and including stage II

herence to biomaterials and tissue. The significance

28 |

Clin. Oral Impl. Res. 13, 2002 / 20–29

of its role in clinical sepsis. Journal of Bone & Joint Surgery of America 67: 264–273. Haddadin, A., Saca, E. & Husban, A. (1999) Sinusitis as a cause of orbital cellulitis. Eastern Mediterranean Health Journal 5: 556–559. Kalmeijer, M.D., van Nieuwland-Bollen, E., BogaersHofman, D. & de Baere, G.A. (2000) Nasal carriage of Staphylococcus aureus is a major risk factor for surgical-site infections in orthopedic surgery. Infectious Control and Hospital Epidemiology 21: 319– 323. Kolander, S.A., Cosgrove, E.M. & Molavi, A. (1989) Clostridial endocarditis. Report of a case caused by Clostridium bifermentans and review of the literature. Archives of Internal Medicine 149: 455–466. Lambert, P.M., Morris, H.F. & Ochi, S. (1997) The influence of 0.12% chlorhexidene digluconate rinses on infectious complications and implant success. Journal of Oral and Maxillofacial Surgery 55: 25– 30. Lauer, G. & Schilli, W. (1994) Collected, implant-cavity borings used as peri-implant osseous augmentation material. International Journal of Oral and Maxillofacial Implants 9: 437–443. Lewis, W.J. & Sherertz, R.J. (1997) Microbial interactions with catheter material. Nutrition 13: 5–9. Marsou, R., Bes, M., Boudouma, M., Brun, Y., Meugnier, H., Freney, J., Vandenesch, F. & Etienne, J. (1999) Distribution of Staphylococcus sciuri subspecies among human clinical specimens, and profile of antibiotic resistance. Research Microbiology 150: 531–541. Mombelli, A., von Osten, M.A.C., Schurch, E. & Lang, N.P. (1987) The microbiota associated with successful or failing osseointegrated implants. Oral Microbiology and Immmunology 2: 145–151. Nachman, S., Kaul, A., Li, K.I., Slim, M.S., San Filippo, J.A. & Van Horn, K. (1989) Liver abscess caused by Clostridium bifermentans following blunt abdominal trauma. Journal of Clinical Microbiology 27: 1137–1138. Nowzari, H. & Slots, J. (1995) Microbiologic and clinical study of polytetrafluoroethylene membranes for guided bone regeneration. International Journal of Oral & Maxillofacial Implants 10: 67–73. Nowzari, H., London, R. & Slots, J. (1995) The importance of periodontal pathogens in guided tissue regeneration and guided bone regeneration. Compendium of Continuing Education in Dentistry 16: 1042, 1044, 1046.

Young et al . Chlorhexidine: its effects on bacteria found in bone debris

Nowzari, H., MacDonald, E.S., Flynn, J., London,

A study of infectious endocarditis in Ribeirao Preto,

dius, Streptococcus coustellatus and Streptococcus

R.M., Morrison, J.L. & Slots, J. (1996) The dynamics of microbial colonisation of barrier membranes for

San Paulo, Brazil. Analysis of cases occurring between 1992 and 1997. Archives of Brazilian Cardi-

anginosus (the Streptococcus milleri group): association with different body sites and clinical infec-

guided tissue regeneration. Journal of Periodontology 67: 694–702.

Sato, S., Sakuragi, T. & Dan, K. (1996) Human skin

Wilcox, M.D.P., Drucker, D.B. & Green, R.M. (1987)

Pencek, T.L. & Burchiel, K.J. (1986) Delayed brain ab-

flora as a potential source of epidural abscess. Anes-

Relative cariogenicity and in vivo plaque forming

scess related to a retained foreign body with culture of Clostridium bifermentans. Case report. Journal of Neurosurgery 64: 813–815. Pereira, M.L., Carmo, L.S. do, Souki, M.Q., Santos, E.J. dos, Carvalho, M.A. de & Bergdoll, M.S. (1999) Staphylococci from dental personnel. Brazilian Dental Journal 10: 39–45. Rams, T.E., Roberts, T.W., Feik, D., Molzan, A.K. & Slots, J. (1991) Clinical and microbiological findings on newly inserted hydroxyapatite-coated and pure titanium human dental implants. Clinical Oral Implants Research 2: 121–127. Rams, T.E., Feik, D., Listgarten, M.A. & Slots, J. (1992) Peptostreptococcus micros in human periodontitis. Oral Microbiology & Immunology 7: 1–6. Roberts, G.L. (2000) Fusobacterial infections: an underestimated threat. British Journal of Biomedical Science 57: 156–162. Robertson K.L. (1999) Differentiation of Prevotella intermedia from P. nigrescens, including isolates from Papillon-Lefevre syndrome. PhD thesis, University of Manchester, Manchester, United Kingdom. Ruiz Jr., E., Schirmbeck, T. & Figueiredo, L.T. (2000)

ology 74: 217–231.

thesiology 85: 1276–1282.

tions. Journal of Clinical Microbiology 30: 243–244.

ability of the bacterium Streptococcus oralis in gno-

Sbordone, L., Barone, A., di Genio, M. & Ramaglia, L.

tobiotic WAG/RIJ rats. Archives of Oral Biology 32:

(1999) Bacterial colonisation during GTR treatment. A longitudinal analysis. Minerva Stomatology 48:

455–457. Wilcox, M.D.P., Knox, K.W., Green, R.M. & Drucker,


D.B. (1991) An examination of strains of the bac-

Scanlan, D.R., Smith, M.A., Isenberg, H.D., Engrassia,

terium Streptococcus vestibularis for relative cario-

S. & Hilton, E. (1994) Clostridium bifermentans

genicity in gnotobiotic rats and adhesion in rats.

bacteremia with metastatic osteomyelitis. Journal of Clinical Microbiology 32: 2867–2868. Sherman, C.R. & Sherman, B.R. (1999) Atropine sulfate: a current review of a useful agent for controlling salivation during dental procedures. General Dentistry 47: 56–60.

Archives of Oral Biology 36: 327–333. Wright, T.L., Ellen, R.P., Lacroix, J.M., Sinnadurai, S. & Mittelman, M.W. (1997) Effects of metronidazole on Porphyromonas gingivalis biofilms. Journal of Periodontal Research 32: 473–477. Yoshinari, N., Tohya, T., Mori, A., Koide, M., Kawase,

Summanen, P., Baron, E.J., Citron, D.M., Strong, C.A.,

H., Takada, T., Inagaki, K., & Noguchi, T. (1998) In-

Wexler, H.M., Finegold, S.M. (1993) Wadsworth Anaerobic Bacteriology Manual 5th Edition. Belmont, CA: Star Publishing Co.

flammatory cell population and bacterial contamination of membranes used for guided tissue regenerative procedures. Journal of Periodontology 69:

Vigil, G.V., Wayman, B.E., Dazey, S.E., Fowler, C.B. & Bradley Jr., D.V. (1997) Identification and antibiotic sensitivity of bacteria isolated from periapical lesions. Journal of Endodontics 23: 110–114. Whiley, R.A., Beighton, D., Winstanley, T.G., Fraser, H.Y. & Hardie, J.M. (1992) Streptococcus interme-

460–469. Young, M.P.J., Korachi, M., Carter, D.H., Worthington, H. & Drucker, D.B. (2001) Microbial analysis of bone collected during dental implant surgery: a clinical and laboratory study. Clinical Oral Implants Research 12: 95–103.

29 |

Clin. Oral Impl. Res. 13, 2002 / 20–29