Skip to main content

Sonication versus the conventional method for evaluation of the dental microbiome: a prospective pilot study



To investigate sonication as a new tool in microbiological probing of dental infections.


Comparison of a standard probing method: intraoperative swab, with sonication, and vortex of the removed tooth, was performed on 20 carious destructed teeth. Illumina high throughput sequencing of the 16S-rRNA-gene was used for assessing the microbial composition. Antibiotic susceptibility has been assigned based on known resistances of each detected species. Probing procedures were compared using Bland–Altmann-Test, and antibiotic susceptibility using the Friedmann-Test and alpha-adjusted post-hoc-analysis.


In total, 60 samples were analysed: 20 intraoperative swabs, 20 vortex fluids, and 20 sonication fluids. Sonication fluid yielded the highest number of bacterial sequencing reads in all three procedures. Comparing the operational taxonomic units (OTUs) of the identified bacteria, significantly more OTUs were found in sonication fluid samples. Phylum and order abundances varied between the three procedures. Significantly more Actinomycetales have been found in sonication fluid samples compared to swab samples. The assigned resistance rates for the identified bacteria (1.79–31.23%) showed no differences between the tested probing procedures. The lowest resistance rates were found for amoxicillin + clavulanate (3.95%) and levofloxacin (3.40%), with the highest in amoxicillin (30.21%) and clindamycin (21.88%).


By using sonication on extracted teeth, it is possible to get a more comprehensive image of the residing microbial flora compared to the standard procedure. If sonication is not available, vortexing is a potential alternative. In immunocompromised patients, especially when actinomycosis is suspected, sonication should be considered for a more detailed microbiological evaluation of the potential disease-causing microbiome. Due to the high rates of antibiotic resistance, a more targeted antibiotic therapy is favourable. Levofloxacin should be considered as a first-line alternative to amoxicillin + clavulanate in patients with an allergy to penicillin.

Peer Review reports


Worldwide, 10% of antibiotic prescriptions are due to dental infections [1]. The most commonly prescribed antibiotics in oral infections are penicillin, followed by lincosamides, macrolides, tetracyclines, and fluoroquinolones [2]. The overprescribing of antibiotics occurs at a rate of 55–80% by dentists [1], leading to an increasing level of bacterial resistance and changes in the composition of the microbiome in odontogenic infections [3,4,5]. Even though early studies focused on finding one specific microorganism causing dental diseases such as caries, gingivitis, and periodontitis, it is now generally accepted, that dental diseases are caused by a change in the specific surface microbiome of the affected oral tissue, driven forward by multispecies microbial interactions [6, 7].

Taking intraoperative swabs in severe odontogenic infections [8] to verify microbiological and antibiotic susceptibility of the present bacteria to ensure a targeted antibiotic regime is the usual procedure. Due to contamination or growth of too many different species or subspecies, which are hard to cultivate, the outcome of this method is poor and calls for better alternatives for microbiological sampling [9,10,11,12].

It is known that the composition and structure of the oral microbiota differ between the existing oral niches [13]. Testing of the saliva reveals microorganisms from various oral niches, but studies have demonstrated that it will not represent the entire oral microbiome [14]. However, tooth surfaces provide an ideal environment for bacterial growth and formation of dental plaque, so they will represent a higher microbial richness and diversity [15]. Besides preventing biofilm-related infections, the analysis of the sensitivity, specificity, and amount of the bacteria in the microbiome plays a decisive role in a targeted antibiotic regimen [10].

Bio-film-related infections do not only occur in the mouth but also in different regions of the human body. One such type of infection associated with biofilm formation is prosthetic joint infections (PJIs). PJIs are the second most common cause of prosthetic joint failure [16] and it is known that they are also caused by bacteria forming an organised biofilm on the implant surface, as oral bacteria do on teeth [17].

Orthopaedic studies have published data about the special procedure of sonication [16, 18]. Thus, the sensitivity and specificity of using the sonication fluid of explanted prostheses for detecting bacteria were shown to be increased compared to the normal procedure, such as synovial fluid cultures and tissue samples or swabs [18]. The number of bacteria identified was up to 10,000 times higher than in standard procedures [18].

Furthermore, sonication was used as an established method on breast, urinary, endovascular and cerebral implants [19,20,21,22]. Less is known about the use of sonication for microbiological testing in dentistry. It was mostly used to detach the biofilm from dentures, to evaluate the amount of Candida species [23], the microbiome of dentures in relation to denture stomatitis [24], and in vitro, to detach the formed biofilm from carbon or titanium surfaces [25] for analysis of the formed biofilm in relation to the implant surface. Regarding the orthopaedic and dental results, the sonication method promises improved detachment of the biofilm located on the teeth, resulting in a complete image of the potential disease-causing microbiome. In these previously published studies, only the sonication method was used to identify the bacterial colonisation process without comparable methods. Furthermore, Almaguer-Flores et al. could demonstrate the strong influence of chemical and physical properties of the substrate in the colonisation of oral bacteria (17).

To the best of our knowledge, no data are available to compare the sonication process to the conventional swab method for microbial investigation. Even in compromised patients, the safe and sufficient extraction of material for microbiological testing and therefore a better knowledge about the predominant microorganisms could lead to a more targeted approach in antibiotic treatment.

Therefore, the aim of the present study was to evaluate the outcome of bacterial DNA extraction and 16S-rRNA amplicon sequencing in the sonication fluid compared to vortex fluid and the standard method, intraoperative swab of the alveolus.


Inclusion criteria and surgical procedure

The research proposal for this prospective study was approved by the Ethics Committee of the Charité Universitätsmedizin Berlin, Germany (EA4/194/19) and complies with the STROBE guidelines. It conforms to the Declaration of Helsinki and the European Medicines Agency Guidelines for Good Clinical Practice. The study’s inclusion criteria were age of majority, at least one permanent tooth, which had to be removed due to carious destruction, no abscess, no systemic disease or drugs, and no nicotine abuses.

Twenty healthy patients, twelve men (mean age 54.2 years) and eight women (mean age 61.9 years) with at least one non-restorable premolar in the mandible were included in the study. All teeth were removed due to carious destruction.

All surgical procedures were performed under sterile conditions by one experienced surgeon under local anaesthesia. Extraction was performed atraumatically, using forceps and elevators. After the removal of the tooth, the extracted tooth was directly placed in a Falcon tube filled with 4 ml Urine Conditioning Buffer™ (UCB™, Zymo Research Corp, CA 92614, USA). Thereafter, a swab, using “DNA/RNA Shield Collection Tube w/ Swab” (Zymo Research Crop, CA 92614, USA), was taken from the extraction socket, representing the standard procedure of microbiological sampling.

All samples have been stored in the refrigerator after collection.

Microbiological preparation and assessment

All samples were transferred to the microbiological laboratory within 24 h, where the probes were processed on a laminar flow bench (Safety cabinet, Thermo Scientific, Langenselbold, Germany). The Falcon tube was vortexed for 30 s. Afterwards, 2 ml of the fluid was removed and placed in a second Falcon tube for further evaluation. Sonication was performed for 1 min at 40 kHz (BactoSonic, Bandelin electronic, Berlin, Germany), following vortexing for another 30 s, as already established in the sonication of endoprosthesis [26]. The swab and the two Falcon tubes per patient, containing the sonication fluid and vortex fluid, were transferred to the microbiology laboratory for analysis. For each procedure, one control sample (swab without probing, vortex, and sonication without tooth) was taken and analysed separately, to evaluate the kit-specific microbiome. Total processing of all samples was performed by the same person.

Microbial analysis

DNA was extracted and purified into 50 µl elution buffer using the DNeasy PowerSoil Pro-Kit (Qiagen). The 16S-V3-V4-PCR was performed using UCP-Multiplex-PCR Mastermix (Qiagen) according to the 16S Metagenomic Sequencing Library Preparation Protocol (Illumina, Primer: Fwd = CCTACGGGNGGCWGCAG, Rev = GACTACHVGGGTATCTAATCC) with 2 μl DNA [27]. In a subsequent PCR, index sequences were added to the purified PCR product. Samples were pooled in one sequencing library and sequenced on an Illumina MiSeq with v2-reagents in 2 × 250 bp paired-end reads with a mean sequencing depth of > 100,000 reads per sample. The number of reads per sample serves as a rough estimation of the amount of bacterial DNA.

After sequencing, paired reads of each amplicon were merged and clustered to Operational Taxonomic Units—OTUs, using the usearch package by clustering identical sequences with an identity of at least 97%. A representative consensus sequence was assigned to every OTU and OTUs were quantified by counting the number of reads mapped to each OTU consensus. The consensus sequences of each OTU were compared to the NCBI 16S-Microbial and NT Reference-Database, using NCBI BLAST (megablast). The OTUs were taxonomically classified based on the best database match (with a minimum identity of 97%) [28]. If multiple database hits matched the OTU sequence with the same identity the OTU was classified as the lowest common ancestor of the different database hits. If no match was found the respective OTU was labeled as unclassified. The reason for the alignment to more than one species relates to the fact that the V3-V4 region in different species can be identical.

Antibiotic susceptibility was evaluated by using “Antibiotics in Laboratory Medicine, 6th Edition” [29]. All found bacterial species were checked for existing enzymes or intrinsic resistances, which are able to inactivate the antibiotics used in clinical routine. Susceptibility was evaluated for amoxicillin, amoxicillin + clavulanate, clindamycin, doxycycline, and levofloxacin. For each sample, we calculated the percentage of bacterial species that have known resistances to any of the 4 antibiotics groups.


Statistical analysis was performed using “R” and MathCalc version 15.8, Graphs were created using “Phyton” and MathCalc version 15.8. Overall, 60 different probes, three for each patient, were analysed. Descriptive Statistics were performed for a number of reads, diversity, phyla distribution, and identified taxa for all samples. Bland–Altmann-test was used to check for significant differences over all three procedures. Friedmann-Test was used to check for differences in antibiotic resistance. Post-hoc analysis was alpha-adjusted and performed to check for significant differences between specific antibiotics. A P-value < 0.05 was considered statistically significant.


Of the 60 samples, the mean number of sequenced reads for each procedure, was 142,858 in the swab, 180,739 in the vortex, and 217,972 in the sonication. The diversity of all samples was evaluated by counting the number of OTUs. Each OTU symbolised one specific bacterial entity. 253 different species were detected in all samples.

In total OTUs, the swab compared to sonication showed a significant difference (p = 0.04; 95% CI − 22.19 to − 0.51). A negative confidence interval highlighted that sonication produced a significantly higher number of OTUs compared to the swab.

All other combinations showed no significant difference. The mean number of OTUs is shown in Fig. 1. Bland–Altman-Plot for comparison swab and sonication, regarding found OTUs is shown in Fig. 2.

Fig. 1
figure 1

Mean OTUs in swab, vortex, sonication

Fig. 2
figure 2

Bland–Altman-Plot for comparison swab and sonication, regarding found OTU

In Fig. 3, the distribution of relative phylum abundances is shown.

Fig. 3
figure 3

Distribution of relative phylum abundance in percentage

There is a high variance in relative phylum abundance in all three sampling procedures. In Actinobacteria (p < 0.01), Bacteroidetes (p < 0.01), and Tenericutes (p < 0.0396), significant differences between swabs and sonication could be found. The confidence interval in Actinobacteria (− 11.19 to − 2.57) showed a higher percentage in sonication in contrast to Bacteroidetes (0.86 to 5.18) and Tenericutes (0.01 to 0.56), where a higher percentage of appearance was found in the swab. Only slight tendencies towards a higher amount of Actinobacteria could be found in vortex, compared to swabs.

On Order-Level, as shown in the supplements, Actinomycetales (p < 0.01), Bacteroidales (p < 0.01), Corynebacteriales (p < 0.01), Flavobacteriales (p = 0.03), and Mycoplasmatales (p = 0.04) showed significant differences when comparing the swab to the sonication procedure (Additional file 1). In Actinomycetales (confidence-interval − 7.29 to − 2.39), Corynebacteriales (95% CI − 2.09 to − 0.34), and Flavobacteriales (95% CI − 2.32 to − 0.16), higher percentage abundances could be found in sonication, in contrast to Bacteroidales (95% CI 2.0777–6.4547) and Mycoplasmatales (95% CI 0.015–0.56) where a higher relative abundance was found in the swab compared to sonication (Additional file 2).

For each experimental group, the mean fraction of bacterial species with intrinsic resistance against certain antibiotics are shown in Table 1.

Table 1 Mean resistance among swab, vortex, and sonication in percentage point

In sonication fluid, vortex fluid, and swab, no significant differences were found in the resistance rate of the bacteria.

A significant difference (p < 0.01) could be found in bacterial resistance to different antibiotics using Friedmann-Test. Assigned ranks were 1.58, 1.73, 3.35, 3.60, and 4.75 for amoxicillin-clavulanate, levofloxacin, doxycycline, clindamycin, and amoxicillin, respectively, representing amoxicillin-clavulanate with the lowest and amoxicillin with the highest resistance percentage rate. The post-hoc test for differences in specific antibiotic resistance between each analysed individual showed significant differences in all combinations except for amoxicillin-clavulanate with levofloxacin and clindamycin with doxycycline. P-values for each comparison are shown in Table 2.

Table 2 Antibiotic resistance comparison

No relevant delay in microbiological results could be observed between the tested procedures.


To summarise the present results, by sonicating the tooth, significantly more bacteria could be detected compared to the swab, as shown by the higher number of OTUs in sonication samples. Furthermore, even the microbial composition of the analysed samples differed between the tested procedures. Moreover, it was possible to find some bacteria which could not be found in the standard procedure for microbiological testing. Focusing on antibiotic resistance rate, no significant difference between sonication fluid, vortex fluid, or swab could be found. A significant difference could be shown in the comparison of the resistance rate of the evaluated antibiotics.

The knowledge of the bacterial composition is crucial for a targeted and effective antibiotic regime. Next-generation sequencing using 16S rRNA gene has shown good results for identification of the oral microbiome [30,31,32,33,34,35]. The microbial composition reported is similar to earlier studies focusing on the oral bacterial composition [30, 32, 33, 36]. The most commonly used sample types to study the healthy oral microbiome and its changes in various diseases were saliva, oral rinse, or niche-specific samples, e.g. supra- or subgingival plaque or tongue swab [31]. Whole teeth have never been analysed before.

For evaluation of the purity of the collected specimen, one control sample for every procedure was taken and analysed separately, as recommended by Zaura et al. [31]. The results showed a low number of reads and diversity, confirming negligible contamination due to the kit-specific microbiome.

The number of reads is a semi-quantitative tool. It is highly affected by the number of PCR cycles performed, the taxa identified, and the sequencing runs itself [37, 38]. Therefore, no precise quantitative measurement is possible regarding how many times more DNA can be found in the sonication fluid compared to the swab or vortex.

In the present study, all patient samples were located on the same sequencing run, with the same number of cycles and same primer respectively, which provides comparability between the swab, vortex, and sonication fluid (Additional file 3).

Taking all of this into consideration, it is highly likely that there is a higher amount of DNA in the sonication fluid compared to the swab and vortex in each patient.

The procedural difference between sonication in contrast to the swab is that they are performed on extracted teeth, resulting in biofilm loosening on the whole surface, whereas the swab was only taken from the alveolus. Therefore, the quantity of the material gathered is probably higher and might have a distorting effect.

The composition of each oral microbiome is different, not only in the number of reads but also in the taxa found. A possible reason for this could be the diversity of the different microbiome surfaces and inter- and intra-individual variations [39]. The oral biofilm development over time is a complex interaction of different species which colonise oral surfaces to form an organised multispecies community with a specific composition. This is caused by the different prevailing physical and biological conditions in the oral habitat, such as surface texture, cell desquamation, or aerobic capacity in the specific niche [6]. Faust et al. demonstrated that the microbiome in different types of samples is similar but nevertheless different [40], highlighting that the source of sampling is crucial for proper microbial testing and especially so for antimicrobial susceptibility testing. This difference could be a reason for the differing microbiological results of the swab and vortex or sonication fluid. Especially regarding the aerophilic capacity of each bacterium, a more anaerobic bacterial composition should be expected in the alveolus or the periodontal pocket than on the tooth surface. By comparing the whole tooth surface, symbolised in the sonication fluid, the bacterial distribution is expected to be different from the bacterial distribution of the alveolus itself.

Apart from this, the contamination of sample extraction kits, during production, can have a potentially misleading impact on the microbiome analysis and consequent conclusion [41].

Even though the biofilm disruption on the whole surface of the tooth could be a potential bias, every bacterium located on the tooth could be able to cause further or could be the reason for the specific infection. Therefore, this setup resembles the reality of biofilm behaviour in the extraction setting in which the potential dissemination of parts of the biofilm can occur, resulting in severe consequences such as infectious endocarditis [42].

A potential criticism could be the amplification of the 16S rRNA gene. Using this procedure, there is no differentiation between bacteria, dead or alive.

Nonetheless, detection of dead bacteria is a potential benefit, because the procedure of swab taking negatively affects the viability of anaerobic bacteria [9]. In normal culture-based analysis, only living bacteria can be examined.

So, the procedure of sonication could be a further influencing factor and cause potential bias. During sonication, little air bubbles explode on the surface of the tooth, leading to the loosening of the biofilm. Bacteria, which are anaerobes or facultative anaerobes, will be highly affected by this excess oxygen [43], even though it could be shown that these bacteria are still alive after sonication of endoprosthesis [18]. Using the 16S rRNA gene, this correlation can be ignored, due to the stability of the genome, even when the bacteria are compromised.

Focusing on the study design, neither orthopaedic nor else studies, which are comparing direct 16S-rRNA-gene analysis and the difference in the microbiome distribution in different sampling modalities exist. Only a few studies have been investigating, whether using 16S-rRNA-gene analysis resulted in similar or even improved results, than normal microbiological testing [26, 44,45,46]. Due to this fact and the appropriate results of 16S-rRNA-gene analysis in microbiome analysis of the oral cavity, we assume, that this is a reliable tool for such investigation [30, 32, 34, 47]. Also in settings, where a dental infection is a potential causing of more severe disease such as medication-related osteonecrosis of the jaw, this sampling method is a potential tool to evaluate and identify the disease-causing bacteria, also in areas hard to reach or where contamination of the normal probing method, swab or tissue sample, is to be expected.

The present results revealed that the amount of Actinomycetales is underrepresented in the normal probing procedure. One potential life-threatening disease, which is hard to diagnose, is craniofacial actinomycosis [48]. In most cases, it is associated with odontogenic infections [49]. Therefore, it can be assumed, that by only taking a swab in combination with normal microbial culture, there is a general underestimation of this disease. Especially in patients undergoing or following radiotherapy due to head and neck cancer, this disease is a feared complication [48]. Also, in medication-related osteonecrosis of the jaw, Actinomyces spp. seem to play a major role in disease progression [50,51,52].

Focusing on Bacteroidales and Mycoplasmatales, which had a higher abundance in swabs, the read numbers of these orders were higher in sonication than in the swab. Showing that there were no bacteria missing but because of the higher amount of Actinomycetales, the relative abundance was lower than in the swab.

The antimicrobial susceptibility to specific antibiotics was not tested in bacterial culture or by molecular genetic analysis, which is a downside of this study. Nevertheless, it was not the primary objective of this investigation to precisely analyse antibiotic resistances, but rather to evaluate the procedure of sonication or vortex as a new tool in microbiological testing in oral surgery. Due to this limitation, no exact statement regarding proper antibiotic treatment can be made.

Heim et al. have already shown the increasing level of resistance found in cultures [53]. Literature-based resistances show the same percentage of resistance as that assigned by enzyme-based evaluation. In particular, the anaerobic species, which are hard to cultivate, show a higher resistance to clindamycin [54], which is still the antibiotic of choice in penicillin-allergic patients [2, 55] according to the German Guidelines [8]. By also evaluating doxycycline and levofloxacin, it could be shown that the resistance rate of amoxicillin with clavulanate was similar to levofloxacin and clindamycin to doxycycline.

Focusing on the analysed resistance rates, a change to the use of levofloxacin as a first-line alternative in severe cases of odontogenic infections should be considered in patients with an allergy to penicillin. Levofloxacin is similar to moxifloxacin, which is already in use in odontogenic infections [2]. In 2011, a comparison of clindamycin and moxifloxacin showed similar outcomes, but with a lower rate of adverse effects in moxifloxacin [56]. Taking this into consideration, levofloxacin or moxifloxacin should replace clindamycin as the first line alternative to amoxicillin with clavulanate in severe odontogenic infections. Due to the high rates of overprescribing of dental antibiotics [1], this change should be exclusively provided for hospitalised patients. The inpatient setting is also preferable for fluoroquinolones, due to their possible side effects like increasing the QT-interval [55]. Other possible side effects are tendinopathy, especially with long-term use [57], and drug-drug interactions. Because of the chondrotoxicity, it is not recommended to use this during pregnancy and in children.

In conclusion with the help of sonication, it was possible to find additional species which were not found in the standard procedure, swabs. The whole microbiome constitution differs, showing a potential incongruence between the standard method and our shown procedure. Still, the microbiomes found were similar in the swab, vortex, and sonication. Consequently, in high-risk patients requiring a more targeted antibiotic treatment (e.g., complex infections, former or ongoing radiotherapy, former or ongoing bisphosphonate medication, congenital heart disease, or immunosuppression) sonication of the tooth should be considered to gain a more complete image of the potential disease-causing microbiome. This sample provides the option to obtain as much information on the bacterial colonisation of the tooth as currently possible, meaning that it can therefore improve treatment as well as clinical outcomes. Early targeted treatment or the prevention of severe complications in high-risk patients can be necessary for their survival.


This evaluation was performed on healthy patients, where the hosts’ anti-infective capability is high and severe complications are rare. Due to the small cohort and the characteristic as a pilot study, further investigation should be performed not only focusing on the differences between sonication and the standard procedure for microbial testing in the treatment of infections of the maxillo-facial region. Additionally, the focus should be laid on changes in the oral microbiome in immune-compromised patients. No differentiation has been performed regarding the location of the teeth or the grade of carious destruction.

Yet, sonication could be a tool, especially for immune-incompetent patients, to improve the overview of the bacteria in the infected area, allowing for a more targeted antimicrobial therapy. Also, differences in conventional microbiological testing: bacterial culture, identification, and bacterial susceptibility are planned, to validate the found data, in a bigger cohort. If sonication is not accessible, we could show that vortex could also be considered for loosening of the biofilm on extracted teeth.

Availability of data and materials

The datasets used and/or analysed during the current study are available from the corresponding author upon reasonable request.


  1. Teoh L, Cheung MC, Dashper S, James R, McCullough MJ. Oral antibiotic for empirical management of acute dentoalveolar infections—a systematic review. Antibiotics. 2021;10(3):66.

    Article  Google Scholar 

  2. Ahmadi H, Ebrahimi A, Ahmadi F. Antibiotic therapy in dentistry. Int J Dent. 2021;2021:6667624.

    PubMed  PubMed Central  Google Scholar 

  3. Sobottka I, Wegscheider K, Balzer L, Boger RH, Hallier O, Giersdorf I, et al. Microbiological analysis of a prospective, randomized, double-blind trial comparing moxifloxacin and clindamycin in the treatment of odontogenic infiltrates and abscesses. Antimicrob Agents Chemother. 2012;56(5):2565–9.

    Article  PubMed  PubMed Central  Google Scholar 

  4. Robertson D, Smith AJ. The microbiology of the acute dental abscess. J Med Microbiol. 2009;58(Pt 2):155–62.

    Article  PubMed  Google Scholar 

  5. Olsen I, Tribble GD, Fiehn NE, Wang BY. Bacterial sex in dental plaque. J Oral Microbiol. 2013;5:66.

    Article  Google Scholar 

  6. Do T, Devine D, Marsh PD. Oral biofilms: molecular analysis, challenges, and future prospects in dental diagnostics. Clin Cosmet Investig Dent. 2013;5:11–9.

    PubMed  PubMed Central  Google Scholar 

  7. Lamont RJ, Hajishengallis G. Polymicrobial synergy and dysbiosis in inflammatory disease. Trends Mol Med. 2015;21(3):172–83.

    Article  PubMed  Google Scholar 

  8. Al-Nawas BK, Julia. Odontogene Infektionen. 2016;Version 1.0.

  9. Ogle OE. Odontogenic infections. Dent Clin N Am. 2017;61(2):235–52.

    Article  PubMed  Google Scholar 

  10. Böttger S, Zechel-Gran S, Schmermund D, Streckbein P, Wilbrand J-F, Knitschke M, et al. Clinical relevance of the microbiome in odontogenic abscesses. Biology. 2021;10(9):916.

    Article  PubMed  PubMed Central  Google Scholar 

  11. Siqueira JF Jr, Rôças IN. As-yet-uncultivated oral bacteria: breadth and association with oral and extra-oral diseases. J Oral Microbiol. 2013;5:66.

    Article  Google Scholar 

  12. Eckert AW, Maurer P, Wilhelms D, Schubert J. Soft tissue infections in oral, maxillofacial, and plastic surgery. Bacterial spectra and antibiotics. Mund Kiefer Gesichtschir. 2005;9(6):389–95.

    Article  PubMed  Google Scholar 

  13. Zhang Y, Wang X, Li H, Ni C, Du Z, Yan F. Human oral microbiota and its modulation for oral health. Biomed Pharmacother. 2018;99:883–93.

    Article  PubMed  Google Scholar 

  14. Ren W, Zhang Q, Liu X, Zheng S, Ma L, Chen F, et al. Exploring the oral microflora of preschool children. J Microbiol. 2017;55(7):531–7.

    Article  PubMed  Google Scholar 

  15. Li X, Liu Y, Yang X, Li C, Song Z. The oral microbiota: community composition, influencing factors, pathogenesis, and interventions. Front Microbiol. 2022;13: 895537.

    Article  PubMed  PubMed Central  Google Scholar 

  16. Trampuz A, Piper KE, Hanssen AD, Osmon DR, Cockerill FR, Steckelberg JM, et al. Sonication of explanted prosthetic components in bags for diagnosis of prosthetic joint infection is associated with risk of contamination. J Clin Microbiol. 2006;44(2):628–31.

    Article  PubMed  PubMed Central  Google Scholar 

  17. Temoin S, Chakaki A, Askari A, El-Halaby A, Fitzgerald S, Marcus RE, et al. Identification of oral bacterial DNA in synovial fluid of patients with arthritis with native and failed prosthetic joints. J Clin Rheumatol. 2012;18(3):117–21.

    Article  PubMed  PubMed Central  Google Scholar 

  18. Trampuz A, Piper KE, Jacobson MJ, Hanssen AD, Unni KK, Osmon DR, et al. Sonication of removed hip and knee prostheses for diagnosis of infection. N Engl J Med. 2007;357(7):654–63.

    Article  PubMed  Google Scholar 

  19. Oliva A, Pavone P, D’Abramo A, Iannetta M, Mastroianni CM, Vullo V. Role of sonication in the microbiological diagnosis of implant-associated infections: beyond the orthopedic prosthesis. Adv Exp Med Biol. 2016;897:85–102.

    Article  PubMed  Google Scholar 

  20. Prinz V, Bayerl S, Renz N, Trampuz A, Vajkoczy P, Finger T. Sonication improves pathogen detection in ventriculoperitoneal shunt-associated infections. Neurosurgery. 2019;85(4):516–23.

    Article  PubMed  Google Scholar 

  21. Bonkat G, Rieken M, Rentsch CA, Wyler S, Feike A, Schafer J, et al. Improved detection of microbial ureteral stent colonisation by sonication. World J Urol. 2011;29(1):133–8.

    Article  PubMed  Google Scholar 

  22. Rieger UM, Pierer G, Luscher NJ, Trampuz A. Sonication of removed breast implants for improved detection of subclinical infection. Aesthetic Plast Surg. 2009;33(3):404–8.

    Article  PubMed  Google Scholar 

  23. Kim E, Driscoll CF, Minah GE. The effect of a denture adhesive on the colonization of Candida species in vivo. J Prosthodont. 2003;12(3):187–91.

    Article  PubMed  Google Scholar 

  24. O’Donnell LE, Robertson D, Nile CJ, Cross LJ, Riggio M, Sherriff A, et al. The oral microbiome of denture wearers is influenced by levels of natural dentition. PLoS ONE. 2015;10(9): e0137717.

    Article  PubMed  PubMed Central  Google Scholar 

  25. Almaguer-Flores A, Ximenez-Fyvie LA, Rodil SE. Oral bacterial adhesion on amorphous carbon and titanium films: effect of surface roughness and culture media. J Biomed Mater Res B Appl Biomater. 2010;92(1):196–204.

    Article  PubMed  Google Scholar 

  26. Renz N, Cabric S, Janz V, Trampuz A. Sonication in the diagnosis of periprosthetic infections: significance and practical implementation. Orthopade. 2015;44(12):942–5.

    Article  PubMed  Google Scholar 

  27. Kurian SM, Gordon S, Barrick B, Dadlani MN, Fanelli B, Cornell JB, et al. Feasibility and comparison study of fecal sample collection methods in healthy volunteers and solid organ transplant recipients using 16S rRNA and metagenomics approaches. Biopreserv Biobank. 2020;18(5):425–40.

    Article  PubMed  Google Scholar 

  28. Olsson LM, Boulund F, Nilsson S, Khan MT, Gummesson A, Fagerberg L, et al. Dynamics of the normal gut microbiota: a longitudinal one-year population study in Sweden. Cell Host Microbe. 2022;30(5):726-39.e3.

    Article  PubMed  Google Scholar 

  29. Amsterdam D. Antibiotics in laboratory medicine, 6th edn. PA: Wolters Kluwer Health; 2014. p. 832.

  30. Keijser BJ, Zaura E, Huse SM, van der Vossen JM, Schuren FH, Montijn RC, et al. Pyrosequencing analysis of the oral microflora of healthy adults. J Dent Res. 2008;87(11):1016–20.

    Article  PubMed  Google Scholar 

  31. Zaura E, Pappalardo VY, Buijs MJ, Volgenant CMC, Brandt BW. Optimizing the quality of clinical studies on oral microbiome: a practical guide for planning, performing, and reporting. Periodontol. 2021;85(1):210–36.

    Article  Google Scholar 

  32. Lazarevic V, Whiteson K, Huse S, Hernandez D, Farinelli L, Osteras M, et al. Metagenomic study of the oral microbiota by Illumina high-throughput sequencing. J Microbiol Methods. 2009;79(3):266–71.

    Article  PubMed  PubMed Central  Google Scholar 

  33. Dewhirst FE, Chen T, Izard J, Paster BJ, Tanner AC, Yu WH, et al. The human oral microbiome. J Bacteriol. 2010;192(19):5002–17.

    Article  PubMed  PubMed Central  Google Scholar 

  34. Carda-Diéguez M, Bravo-González LA, Morata IM, Vicente A, Mira A. High-throughput DNA sequencing of microbiota at interproximal sites. J Oral Microbiol. 2020;12(1):1687397.

    Article  PubMed  Google Scholar 

  35. Oliveira SG, Nishiyama RR, Trigo CAC, Mattos-Guaraldi AL, Dávila AMR, Jardim R, et al. Core of the saliva microbiome: an analysis of the MG-RAST data. BMC Oral Health. 2021;21(1):351.

    Article  PubMed  PubMed Central  Google Scholar 

  36. Zaura E, Keijser BJ, Huse SM, Crielaard W. Defining the healthy “core microbiome” of oral microbial communities. BMC Microbiol. 2009;9(1):259.

    Article  PubMed  PubMed Central  Google Scholar 

  37. Drengenes C, Eagan TML, Haaland I, Wiker HG, Nielsen R. Exploring protocol bias in airway microbiome studies: one versus two PCR steps and 16S rRNA gene region V3 V4 versus V4. BMC Genomics. 2021;22(1):3.

    Article  PubMed  PubMed Central  Google Scholar 

  38. Abusleme L, Hong BY, Dupuy AK, Strausbaugh LD, Diaz PI. Influence of DNA extraction on oral microbial profiles obtained via 16S rRNA gene sequencing. J Oral Microbiol. 2014;6:66.

    Article  Google Scholar 

  39. Lazarevic V, Whiteson K, Hernandez D, Francois P, Schrenzel J. Study of inter- and intra-individual variations in the salivary microbiota. BMC Genomics. 2010;11(1):523.

    Article  PubMed  PubMed Central  Google Scholar 

  40. Faust K, Sathirapongsasuti JF, Izard J, Segata N, Gevers D, Raes J, et al. Microbial co-occurrence relationships in the human microbiome. PLoS Comput Biol. 2012;8(7): e1002606.

    Article  PubMed  PubMed Central  Google Scholar 

  41. Salter SJ, Turner C, Watthanaworawit W, de Goffau MC, Wagner J, Parkhill J, et al. A longitudinal study of the infant nasopharyngeal microbiota: the effects of age, illness and antibiotic use in a cohort of South East Asian children. PLoS Negl Trop Dis. 2017;11(10): e0005975.

    Article  PubMed  PubMed Central  Google Scholar 

  42. Han YW, Wang X. Mobile microbiome: oral bacteria in extra-oral infections and inflammation. J Dent Res. 2013;92(6):485–91.

    Article  PubMed  PubMed Central  Google Scholar 

  43. Joyce E, Phull SS, Lorimer JP, Mason TJ. The development and evaluation of ultrasound for the treatment of bacterial suspensions. A study of frequency, power and sonication time on cultured Bacillus species. Ultrason Sonochem. 2003;10(6):315–8.

    Article  PubMed  Google Scholar 

  44. Achermann Y, Vogt M, Leunig M, Wüst A Jr, Trampuz A. Improved diagnosis of periprosthetic joint infection by multiplex PCR of sonication fluid from removed implants. J Clin Microbiol. 2010;48(4):1208–14.

    Article  PubMed  PubMed Central  Google Scholar 

  45. Jacquier H, Fihman V, Amarsy R, Vicaut E, Bousson V, Cambau E, et al. Benefits of polymerase chain reaction combined with culture for the diagnosis of bone and joint infections: a prospective test performance study. Open Forum Infect Dis. 2019;6(12):ofz511.

    Article  PubMed  PubMed Central  Google Scholar 

  46. Renz N, Cabric S, Morgenstern C, Schuetz MA, Trampuz A. Value of PCR in sonication fluid for the diagnosis of orthopedic hardware-associated infections: Has the molecular era arrived? Injury. 2018;49(4):806–11.

    Article  PubMed  Google Scholar 

  47. Ng E, Tay JRH, Balan P, Ong MMA, Bostanci N, Belibasakis GN, et al. Metagenomic sequencing provides new insights into the subgingival bacteriome and aetiopathology of periodontitis. J Periodontal Res. 2021;56(2):205–18.

    Article  PubMed  Google Scholar 

  48. Karanfilian KM, Valentin MN, Kapila R, Bhate C, Fatahzadeh M, Micali G, et al. Cervicofacial actinomycosis. Int J Dermatol. 2020;59(10):1185–90.

    Article  PubMed  Google Scholar 

  49. Valour F, Senechal A, Dupieux C, Karsenty J, Lustig S, Breton P, et al. Actinomycosis: etiology, clinical features, diagnosis, treatment, and management. Infect Drug Resist. 2014;7:183–97.

    PubMed  PubMed Central  Google Scholar 

  50. Russmueller G, Seemann R, Weiss K, Stadler V, Speiss M, Perisanidis C, et al. The association of medication-related osteonecrosis of the jaw with Actinomyces spp. infection. Sci Rep. 2016;6(1):31–604.

    Article  Google Scholar 

  51. Arranz Caso JA, Flores Ballester E, Ngo Pombe S, Lopez Pizarro V, Dominguez-Mompello JL, Restoy LA. Bisphosphonate related osteonecrosis of the jaw and infection with Actinomyces. Med Clin. 2012;139(15):676–80.

    Article  Google Scholar 

  52. Raguse JD, Trampuz A, Boehm MS, Nahles S, Beck-Broichsitter B, Heiland M, et al. Replacing one evil with another: Is the fibula really a dispensable spare part available for transfer in patients with medication-related osteonecrosis of the jaws? Oral Surg Oral Med Oral Pathol Oral Radiol. 2020;129(6):e257–63.

    Article  PubMed  Google Scholar 

  53. Heim N, Jurgensen B, Kramer FJ, Wiedemeyer V. Mapping the microbiological diversity of odontogenic abscess: Are we using the right drugs? Clin Oral Investig. 2021;25(1):187–93.

    Article  PubMed  Google Scholar 

  54. Brook I. Spectrum and treatment of anaerobic infections. J Infect Chemother. 2016;22(1):1–13.

    Article  PubMed  Google Scholar 

  55. Flynn TR. Evidence-based principles of antibiotic therapy. In: Ferneini EM, Goupil MT, editors. Evidence-based oral surgery. Cham: Springer; 2019. p. 283–316.

    Google Scholar 

  56. Cachovan G, Boger RH, Giersdorf I, Hallier O, Streichert T, Haddad M, et al. Comparative efficacy and safety of moxifloxacin and clindamycin in the treatment of odontogenic abscesses and inflammatory infiltrates: a phase II, double-blind, randomized trial. Antimicrob Agents Chemother. 2011;55(3):1142–7.

    Article  PubMed  Google Scholar 

  57. Sendzik J, Shakibaei M, Schafer-Korting M, Lode H, Stahlmann R. Synergistic effects of dexamethasone and quinolones on human-derived tendon cells. Int J Antimicrob Agents. 2010;35(4):366–74.

    Article  PubMed  Google Scholar 

Download references


The authors thank PD Dr. Andrej Trampuz ( Charité Universitätsmedizin Berlin) for his support during the antimicrobial investigation and help by providing lab space.


Open Access funding enabled and organized by Projekt DEAL. This research received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.

Author information

Authors and Affiliations



OW: Contributed to conception and design, acquisition, analysis, and interpretation, performed the statistical analysis, drafted and critically revised the manuscript. PM: Contributed to analysis and interpretation, performed the statistical analysis, and critically revised the manuscript. RS: Contributed to analysis and interpretation and critically revised the manuscript. NN: Contributed to conception and design, acquisition, drafted and critically revised the manuscript. SP: Contributed to acquisition and critically revised the manuscript. MH: Contributed to interpretation and critically revised the manuscript. SN: Contributed to conception and design, interpretation, and critically revised the manuscript. All authors read and approved the final manuscript.

Corresponding authors

Correspondence to Oliver Wagendorf or Susanne Nahles.

Ethics declarations

Ethics approval and consent to participate

The research proposal for this prospective study was approved by the Ethics Committee of the Charité Universitätsmedizin Berlin, Germany (EA4/194/19) and complies with the STROBE guidelines. It conforms to the Declaration of Helsinki and the European Medicines Agency Guidelines for Good Clinical Practice. Informed consent was obtained from all subjects.

Consent for publication

Not applicable.

Competing interests

The authors have no conflict of interest to declare.

Additional information

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary Information

Additional file 1.

Distribution of relative order abundances in %.

Additional file 2.

Differences in percentage in relative order abundance in each procedure.

Additional file 3.

Rarefraction curves for every patient.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Wagendorf, O., Menzel, P., Schwarzer, R. et al. Sonication versus the conventional method for evaluation of the dental microbiome: a prospective pilot study. BMC Oral Health 22, 348 (2022).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: