Effect of different atmospheres on microcosm biofilm formation and tooth demineralization

doi:10.21203/rs.3.rs-2081258/v1

Download PDF

Research Article

Effect of different atmospheres on microcosm biofilm formation and tooth demineralization

https://doi.org/10.21203/rs.3.rs-2081258/v1

This work is licensed under a CC BY 4.0 License

Journal Publication

published 01 Jun, 2023

Read the published version in Journal of Applied Oral Science →

Version 1

posted

You are reading this latest preprint version

This study evaluated the effects of different atmospheres on the cariogenic potential of microcosm biofilms. Ninety bovine enamel and 90 dentin specimens were allocated into three atmospheres: 1) microaerophilia (5 days, 5% CO2); 2) anaerobiosis (5 days, jar); 3) mixed (2 days microaerophilia and 3 days anaerobiosis), which were subdivided into 0.12% chlorhexidine (positive control- CHX) and Phosphate-Buffered Saline (negative control- PBS) (n = 15). Biofilms were prepared using human saliva and McBain's saliva containing 0.2% sucrose. From the second day, the specimens were treated with CHX or PBS (1 x 1 min/day). After five days, colony-forming units (CFU) were counting and tooth demineralization was analyzed using transverse microradiography (TMR). Data were subjected to two-way ANOVA and Tukey–Sidak’s test (p < 0.05). Regarding CFU counting, most atmospheres were able to differentiate between CHX and PBS (differences of 0.3–1.48 log10 CFU/ml), except for anaerobiosis and microaerophilia for total microorganisms in enamel and dentin biofilm, respectively. In the case of dentin, no effect of CHX on Lactobacillus spp. was observed. All atmospheres were able to differentiate between CHX and PBS regarding enamel demineralization, showing lower mineral loss and lesion depth for CHX (78% and 22% reductions for enamel and dentin, respectively). The enamel mineral loss data did not differ between the models; however, the enamel lesion depth was greater under anaerobiosis. Dentin mineral loss was lower under anaerobiosis than under other atmospheres. Conclusion: The choice of atmosphere did not seem to interfere with the cariogenic potential of the microcosm biofilm.

dental caries

dentin

enamel

microorganisms

oral biofilm.

Dental caries is a multifactorial oral disease of great clinical relevance. It affects the tooth structure because of the presence of a biofilm in dysbiosis, which is rich in acidogenic, aciduric, and extracellular polysaccharide-producing microorganisms, which occur due to the frequent ingestion of sugar. These microorganisms can metabolize different types of sugars from the diet, mainly sucrose, forming acids that change the biofilm pH and cause tooth demineralization (Bowen 2002; Marsh et al. 2011; Walsh et al. 2015; Takahashi 2015; Pitts et al. 2017). The most common microorganisms involved in the development of dental caries are Streptococcus mutans and lactobacilli (Aas et al. 2008; Gross et al. 2010; Vieira et al. 2014; Henne et al. 2015, Mosaddad et al. 2019).

Because of the high degree of control of culture conditions and reproducibility, in vitro models have been well accepted to simulate cariogenic biofilms (Amaechi et al. 2019). In vitro models can be produced from monospecies, namely, S. mutans, or multispecies, namely, S. mutans + Lactobacillus casei biofilm models using microbial strains or from microorganisms derived from human saliva or dental biofilms, called microcosm biofilm (Filoche et al. 2007; Schwendicke et al. 2017; Amend et al. 2018).

Microcosm biofilms can accurately mimic the complexity of a dental biofilm (McBain 2009; Sim et al. 2016; Braga et al. 2021). However, within this model, different forms of cultivation have been used, comprising, 24-well microtiter plates, where specimens can be attached to the bottom of the wells (Zhang et al. 2015; Souza et al. 2018; Pires et al. 2019); meshes for the retention of microorganisms (Ayoub et al. 2020); or specimens suspended to prevent the formation of biofilm only by microorganism precipitation (Méndez et al. 2018). Some microcosm biofilm models use artificial mouths for programmed or continuous nutrient flow (Amend et al. 2018; Santos et al. 2019).

Studies differ concerning the sucrose concentrations and form of exposure, which can be either continuous or intermittent (Signori et al. 2018; Braga et al. 2020). Cultivation time can vary from 2–14 d depending on the selected response variables (Rudney et al. 2012; Souza et al. 2018). Incubation atmosphere is another important factor (Sim et al. 2016), which can vary between microaerophilia (5–10% CO₂) (Rudney et al. 2012; Zhang et al. 2015; Santos et al. 2019; Ayoub et al. 2020) and anaerobiosis with the use of jars, candles, or anaerobic cabins (O₂ < 0.1% and 5–10% CO₂; 10% CO₂, 10% H₂ and 80% N₂) (Maske et al. 2016; Signori et al. 2018).

Oral dental biofilms are constantly exposed to substantial fluxes of environmental conditions, namely, fast pH changes, availability of nutrients, nature of carbohydrates, and variations in redox potentials due to atmospheric conditions. These environmental parameters determine the composition of the microbial population in biofilms (Kolenbrander 2000; Dhaked et al. 2021).

Therefore, atmospheric conditions can modulate microbial changes and cariogenic potential during biofilm development (Sim et al. 2016; Maske et al. 2017). Environmental changes, physiological disturbances, or selective pressures that might occur during dental biofilm growth can stimulate the proliferation of certain species leading to the development of a potential cariogenic biofilm (Parisotto et al., 2010).

Despite this wide methodological variation between studies, the impact of the culture atmosphere on the development of microcosm biofilms and their potential to cause tooth demineralization has not yet been compared. Therefore, this study aimed to compare the three experimental cultivation models (microaerophile vs. anaerobiosis vs. experimental mixed) on the colony-forming units (CFU) of the cariogenic microorganisms and tooth demineralization. The null hypothesis is that atmospheric conditions do not influence the S. mutans and Lactobacillus spp. colonies and development of dental caries lesions (tooth demineralization).

Saliva Collection

This study was approved (CAAE: 35403320.1.0000.5417) by the local ethics committee of Bauru School of Dentistry-USP (Bauru-Brazil). The study was conducted according to the Declaration of Helsinki. After obtaining consent, saliva was collected from ten healthy participants (age: 23.8 ± 3; eight women and two men). The inclusion criteria for the saliva donors and the procedure for saliva collection (salivary flow > 1 mL/min for 10 min) followed previously reported protocols (Braga et al. 2019). Saliva samples were collected once in the morning. In total, 132 mL (pool) of saliva was collected and diluted in glycerol (70% saliva and 30% glycerol), and 1 mL of aliquots were stored at − 80° C (Pratten et al. 2003).

Tooth Specimen Preparation

Bovine specimens of cattle that were sacrificed in the food manufacturing industry (Frigol S. A., Lençóis Paulista, São Paulo, SP, Brazil) were obtained. The study was approved by the Ethics Committee on Animal Research (CEUA, Number 005/2020, Bauru School of Dentistry, University of São Paulo, Bauru, Brazil) following the guidelines provided by the National Council for Control of Animal Experimentation. Ninety bovine enamel and 90 bovine root dentin specimens (4 mm × 4 mm) were cut, polished, and evaluated for the average roughness (Ra; contact profilometer Mahr, Göttingen, Germany) (Braga et al. 2019) to standardize the tooth surfaces for biofilm growth. Two-thirds of the surface was covered with red nail polish (Estreia-Colorama, Rio de Janeiro, RJ, Brazil) to protect it from the biofilm and create two sound areas. This enabled appropriate analysis of tooth demineralization using transverse microradiography (TMR). Subsequently, the specimens were sterilized by exposure to ethylene oxide. Enamel and dentin specimens were randomly distributed into three groups and two subgroups (n = 15) according to their mean Ra values (Enamel Ra: 0.140 ± 0.029 µm and dentin Ra: 0.288 ± 0.056 µm), allowing the groups to have similar mean baseline Ra values.

Study conditions and sample size calculation

The experimental models differed regarding the atmosphere, namely, microaerophilia (greenhouse with 5% CO₂), anaerobiosis (greenhouse with jar and Microbiology Anaerocult A, Merck, Darmstadt, Germany, < 0.5% O₂), and mixed (two days in the microaerophilic model and three days in the anaerobic model). Under each atmospheric condition, half of the specimens were treated with Chlorhexidine (CHX), and the other half were treated with Phosphate buffered saline (PBS). The entire experiment was performed in biological triplicates (n = 5 each), with a final n = 15 (Table 1). The sample size was calculated using the website, http://powerandsamplesize.com, based on a previous study that indicated an integrated mineral loss (∆Z) of 3237.1 vol%. µm (SD: 781.0 µm) for CHX (Periogard) versus ∆Z 6151.3 vol%. µm (SD: 1084.5) for the PBS group (Braga et al. 2020), under a power of 80% and α error of 5% (n = 2/group).

Table 1

The tested atmospheres conditions and treatments in this experiment
Groups	Atmospheres conditions
Microaerophilia	Greenhouse with 5% CO₂
Anaerobiosis	Greenhouse with jar and Microbiology Anaerocult A (< 0.5% O₂)
Mixed	2 days in the microaerophilia and 3 days anaerobiosis
Subgroups	Company/City-Country	Composition/Concentration
PerioGard (positive control)	Colgate-Palmolive/São Paulo-Brazil	Active component: 0.12% chlorhexidine gluconate, sorbitol and cetylpyridinium chloride.
PBS (negative control)	-	-

Preparation of Artificial Saliva

McBain artificial saliva (McBain 2009), containing 2.5 g/L mucin (type II) from porcine stomach, 2.0 g/L tryptone, 2.0 g/L bacteriological peptone, 1.0 g/L yeast extract, 0.35 g/L NaCl, 0.2 g/L CaCl₂, 0.2 g/L KCl, 0.1 g/L cysteine hydrochloride, 0.001 g/L hemin, and 0.0002 g/L vitamin K1, was prepared. Some components were sterilized using an autoclave. Mucin was sterilized by pasteurization to protect its molecular structure and function and cysteine hydrochloride, hemin, and vitamin K1 were sterilized using a syringe filter.

Microcosm Biofilm Formation

In a 24-wells microtiter plate, each enamel or dentin specimen was exposed to 1.5 mL of inoculum (human saliva-glycerol + McBain saliva, 1:50) for 8 h. After the first 8 h, the inoculum was removed, the specimens were washed with PBS (5 s) and then received 1.5 mL fresh medium (McBain artificial saliva) with 0.2% sucrose for 16 h, for the first 24 h. From the second to the fifth day, the medium (McBain saliva) containing 0.2% sucrose was changed once a day. Before medium replacement, half of the specimens were treated daily with 0.12% CHX/PerioGard (positive control) and the other half with PBS (negative control) for 1 min (v = 1 mL/well). The experiment was performed at 37°C and each plate was stored under different atmospheric conditions.

Colony-forming units (CFU) counting

After the fifth day, the specimens were washed once with PBS solution to remove dead bacteria or those that did not adhere to the biofilm. Subsequently, the specimens were transferred to microtubes containing 1 mL of 0.89% NaCl solution and sonicated (Sonifier Cell Disruptor B-30, Branson, Danbury, USA) for 30 s at 20 W. For CFU counting, 100 µL of the microbial suspension from microcosm biofilm were diluted to 10^− 5 and spread on petri dishes (25 µL/dish) containing three types of agar: 1) Brain Heart Infusion agar (BHI, Kasvi, Curitiba, Brazil) for total microorganisms; 2) De Man, Rogosa and Sharpe (MRS- Kasvi, Curitiba, Brazil) supplemented with 0.13% glacial acetic acid to assess the number of total lactobacilli (Lima et al. 2009); and 3) SB-20M containing 15 g bacto-casitone (Difco, Detroit, USA), 5 g yeast extract (Kasvi, Curitiba, Brazil), 0.2 g L-cysteine hydro-chloride (Sigma, Steinheim, Germany), 0.1 g sodium sulfite (Sigma, Steinheim, Germany), 20.0 g sodium acetate (Synth, Diadema, Brazil), 200.0 g coarse granular cane sugar, 15.0 g agar (Kasvi, Curitiba, Brazil), and 1 L distilled water (autoclaved) plus 0.2 U/mL bacitracin (Sigma, Steinheim, Germany) for determination of mutans streptococci (S. mutans and S. sobrinus) (Silva et al., 2012; Li et al., 2014).

The plates were then incubated under the same conditions as the microcosm biofilm. In the mixed atmosphere, the plates were separately incubated under both microaerophilic and anaerobic conditions. After 48 h, CFU was computed and transformed to log₁₀ CFU/mL.

Transverse microradiography (TMR)

After removing the biofilm from the specimens, all enamel and dentin specimens were transversally sectioned and polished to obtain slices of 80–100 µm (enamel) and 100–120 µm (dentin) thickness (Braga et al. 2020; Vertuan et al. 2021). The slices were fixed in a sample holder together with an aluminum calibration step wedge with 14 steps. Ethylene glycol (Sigma-Aldrich, Steinheim, Germany) was applied on the dentin specimens for 24 h to avoid shrinkage due to desiccation during X-ray exposure desiccation (Buchalla et al. 2003). A microradiograph was obtained using an X-ray generator (Softex, Tokyo, Japan) on a glass plate at 20 kV and 20 mA (at a distance of 42 cm) for 13 min. The glass plates (high-precision photo plate, Konica Minolta Inc., Tokyo, Japan) were processed and analyzed using a transmitted light microscope fitted with a 20x objective (Zeiss, Oberkochen, Germany), a CCD- Charge-Coupled Device camera (Canon, Tokyo, Japan), and a computer. Two images per specimen were acquired using data acquisition (version 2012) and interpreted using the Inspektor Research System BV calculation software (version 2006) (Amsterdam, Netherlands). The mineral content was calculated based on the assumption of 87% volume of mineral content for sound enamel and 50% mineral content for sound dentin. Lesion depth (LD, µm) integrated mineral loss (∆Z, vol. µm), and the average mineral loss over the lesion depth (R, vol%) were obtained.

Statistical analysis

Data were statistically compared using the GraphPad Prism software (San Diego, CA, USA). Distribution and homogeneity were tested using the Kolmogorov–Smirnov and Bartlett’s tests, respectively. A two-way ANOVA (factors: atmospheres and treatments), followed by Tukey’s test or Sidak’s test, was applied. The level of significance was set at 5%.

CFU counting

Enamel

Table 2 presents the CFU values (log₁₀/mL) for the total microorganisms, Lactobacillus spp., and S. mutans/ S. sobrinus from the microcosm biofilm cultivated under various atmospheric growth conditions on the enamel specimens.

All atmospheres, except the anaerobic condition, were able to differentiate CHX from PBS, displaying the antimicrobial effect of the CHX on total microorganisms. A minor difference in the growth of total microorganisms was seen in microaerophilia compared to the other atmospheres, which did not differ from each other (anaerobic and mixed) regardless of the treatment.

Table 2

CFU counting (log₁₀ CFU/mL) of total microorganisms, *Lactobacillus* spp. and *Streptococcus mutans/Streptococcus sobrinus* from microcosm biofilm produced on enamel specimens.
Groups	Subgroups	Total microorganisms	Lactobacillus spp.	S. mutans/S. sobrinus
Microaerophilia	CHX	6.38 ± 0.52^Aa	5.87 ± 0.86^Aab	6.41 ± 0.36^Aa
Microaerophilia	PBS	7.37 ± 0.43^Ba	7.08 ± 0.11^Bab	8.05 ± 0.12^Ba
Anaerobiosis	CHX	7.35 ± 0.47^Ab	6.22 ± 0.39^Abc	6.53 ± 0.33^Aab
Anaerobiosis	PBS	7.68 ± 0.71^Aab	7.46 ± 0.21^Bbc	7.35 ± 0.43^Bb
Mixed-Anaerobic	CHX	7.15 ± 0.59^Ab	5.69 ± 0.47^Aa	6.88 ± 0.54^Ab
Mixed-Anaerobic	PBS	7.91 ± 0.25^Bb	7.17 ± 0.10^Ba	7.74 ± 0.11^Bab
Mixed-Microaerophilia	CHX	7.20 ± 0.57^Ab	6.31 ± 0.32^Ac	6.50 ± 0.81^Aab
Mixed-Microaerophilia	PBS	7.84 ± 0.18^Bb	7.70 ± 0.07^Bc	7.72 ± 0.09^Bab

Capital letters show significant difference between treatments, for each type of atmosphere. Lowercase letters show significant difference between atmospheres, for each type of treatment. For total microorganisms, two-way ANOVA was applied followed by Tukey's test (treatment p < 0.0001, atmosphere p < 0.0001 and interaction p = 0.0481). For Lactobacillus spp., two-way ANOVA was applied, followed by the Sidak’s test (treatment p < 0.0001, atmosphere p < 0.0001, no interaction, p = 0.7677). For S. mutans/ S. sobrinus, two-way ANOVA was applied, followed by the Tukey’s test (treatment p < 0.0001, atmosphere p = 0.0041 and interaction p = 0.0039).

All atmospheres were able to differentiate CHX from PBS, indicating the antimicrobial effect of CHX on Lactobacillus spp. and S. mutans/ S. sobrinus.

Lower growth of Lactobacillus spp. was observed under a mixed atmosphere (anaerobic CFU) than in the other conditions, except for the microaerophilic condition. The microaerophilic atmosphere was similar to the anaerobic atmosphere but displayed lower Lactobacillus spp. growth compared with the mixed atmosphere (microaerophilic CFU), which did not differ from the anaerobic and mixed-anaerobic CFU.

Regarding S. mutans/ S. sobrinus, lower microbial growth was observed in the microaerophilic atmosphere, which was similar to the mixed atmosphere (microaerophilic CFU). However, the microaerophilic atmosphere significantly differed from the anaerobic atmosphere when considering PBS and from the mixed atmosphere (anaerobic CFU) when considering CHX.

Dentin

Table 3 displays the CFU values (log₁₀/mL) for the total microorganisms, Lactobacillus spp., and S. mutans/ S. sobrinus from a microcosm biofilm cultivated under different atmospheric growth conditions on dentin specimens.

Table 3

CFU counting (log₁₀ CFU mL) of total microorganisms, *Lactobacillus* spp. and *Streptococcus mutans*/*Streptococcus sobrinus* from microcosm biofilm produced on dentin specimens.
Groups	Subgroups	Total microorganisms	Lactobacillus spp.	S. mutans/S. sobrinus
Microaerophilia	CHX	7.62 ± 0.22^Aa	6.95 ± 0.46^Aab	7.12 ± 0.39^Aa
Microaerophilia	PBS	7.74 ± 0.11^Aa	7.06 ± 0.38 ^Aab	7.63 ± 0.19^Ba
Anaerobiosis	CHX	7.45 ± 0.38^Aa	7.13 ± 0.46^Ab	7.24 ± 0.35^Aa
Anaerobiosis	PBS	7.94 ± 0.22^Ba	7.51 ± 0.20^Ab	7.60 ± 0.19^Ba
Mixed-Anaerobic	CHX	7.21 ± 0.53^Aa	7.25 ± 0.59^Ab	7.05 ± 0.35^Aa
Mixed-Anaerobic	PBS	7.84 ± 0.16^Ba	7.33 ± 0.51^Ab	7.69 ± 0.21^Ba
Mixed-Microaerophilia	CHX	7.49 ± 0.28^Aa	6.76 ± 0.36^Aa	7.21 ± 0.41^Aa
Mixed-Microaerophilia	PBS	7.80 ± 0.16^Ba	6.87 ± 0.56^Aa	7.77 ± 0.22^Ba

Capital letters show significant difference between treatments, for each type of atmosphere. Lowercase letters show significant difference between atmospheres, for each type of treatment. For total microorganisms, two-way ANOVA was applied, followed by Tukey's test (treatment p < 0.0001, atmosphere p < 0.1222 and interaction p = 0.0099). For Lactobacillus spp., two-way ANOVA was applied, followed by the Tukey’s test (treatment p = 0.0602, atmosphere p = 0.0002, no interaction, p = 0.5955). For S. mutans/ S. sobrinus, two-way ANOVA was applied, followed by the Tukey’s test (treatment p < 0.0001, atmosphere p = 0.4706 and interaction p = 0.4637).

All atmospheres, except microaerophilia, were able to differentiate between CHX and PBS, showing the antimicrobial effect of CHX on all microorganisms. When the atmospheres were compared, no differences were observed regardless of the treatment.

Regarding Lactobacillus spp. CFU, none of the atmospheres was able to differentiate between CHX and PBS. When the atmospheres were compared, a mixed atmosphere (microaerophilic CFU) presented lower growth of Lactobacillus spp. than the other atmospheres, except for microaerophilia.

In contrast, all atmospheres significantly differed in CHX and PBS regarding S. mutans/ S. sobrinus growth, whereas no differences were observed when the atmospheres were compared regardless of the treatment.

TMR analysis

Tables 4 and 5 present the TMR data for the enamel and dentin specimens, respectively. Figures 1 and 2 display representative TMR images of the enamel and dentin from each group, respectively.

Table 4

Mean ± SD of the integrated mineral loss (ΔZ, vol%. µm), the lesion depth (LD, µm) and the average mineral loss (R, vol%) of enamel specimens.
Groups	Subgroups	ΔZ (vol%.µm)	LD (µm)	R (vol%)
Microaerophilia	CHX	1035.62 ± 561.38^Aa	37.89 ± 15.46^Aa	27.92 ± 8.16^Aa
Microaerophilia	PBS	7223.33 ± 1206.52^Ba	119.84 ± 20.27^Ba	59.14 ± 3.11^Ba
Anaerobiosis	CHX	1702.22 ± 1056.60^Aa	61.19 ± 24.22^Ab	30.57 ± 10.52^Aa
Anaerobiosis	PBS	6555.00 ± 1681.07^Ba	134.22 ± 28.45^Bb	54.71 ± 4.33^Ba
Mixed	CHX	1436.11 ± 495.77^Aa	49.46 ± 13.01^Aab	23.94 ± 2.05^Aa
Mixed	PBS	7246.11 ± 350.14^Ba	119.54 ± 8.47^Bab	59.66 ± 2.06^Ba

Capital letters show significant difference between treatments, for each type of atmosphere. Lowercase letters show significant difference between atmospheres, for each type of treatment. For ΔZ, two-way ANOVA was applied, followed by Tukey’s test (treatment p < 0.0001; atmosphere p = 0.7627 and interaction p = 0.0168). For lesion depth, two-way ANOVA was applied, followed by Tukey’s test (treatment p < 0.0001, atmosphere p = 0.0148 and interaction p = 0.6412). For the average mineral loss, two-way ANOVA was applied, followed by Sidak’s test (treatment p < 0.0001, atmosphere p = 0.7709, interaction, p = 0.0284).

Table 5

Mean ± SD of the integrated mineral loss (ΔZ, vol%. µm), the lesion depth (LD, µm) and the average mineral loss (R, vol%) of dentin specimens.
Groups	Subgroups	ΔZ (vol%.µm)	LD (µm)	R (vol%)
Microaerophilia	CHX	4389.2 ± 603.51^Aa	159.2 ± 13.7^Aa	29.2 ± 3.4^Aa
Microaerophilia	PBS	5768.5 ± 523.95^Ba	184.2 ± 24.5^Ba	31.5 ± 2.9^Aa
Anaerobiosis	CHX	4242.3 ± 581.68^Aa	144.0 ± 14.0^Aa	27.8 ± 2.2^Ab
Anaerobiosis	PBS	5149.6 ± 551.40^Bb	194.3 ± 28.7^Ba	28.7 ± 3.1^Ab
Mixed	CHX	4744.3 ± 472.69^Aa	160.3 ± 18.4^Aa	29.0 ± 2.3^Aa
Mixed	PBS	5692.5 ± 243.04^Ba	183.1 ± 13.2^Ba	31.6 ± 2.5^Ba

Capital letters show significant difference between treatments, for each type of atmosphere. Lowercase letters show significant difference between atmospheres, for each type of treatment. For ΔZ, two-way ANOVA was applied, followed by Tukey’s test (treatment p < 0.0001; atmosphere p = 0.0034 and interaction p = 0.2346). For lesion depth, two-way ANOVA was applied, followed by Tukey’s test (treatment p < 0.0001, atmosphere p = 0.809 and interaction p = 0.0614). For average mineral loss, two-way ANOVA was applied followed by Tukey’s test (treatment p = 0.0039, atmosphere p = 0.0093, interaction, p = 0.4243).

Chlorhexidine reduced enamel demineralization in all the tested atmospheres. Similar carious lesions were produced in enamel in all atmospheres, except for microaerophilia, which induced a shallow lesion compared with the anaerobic atmosphere, both being similar to the mixed atmosphere (Table 4 and Fig. 1).

Chlorhexidine was also able to reduce dentin demineralization in all tested atmospheres, except for the average mineral loss in the case of microaerophilia and anaerobiosis. Microaerophilic and mixed atmospheres produced similar carious lesions in dentin with greater mineral loss (integrated and mean) than those produced under anaerobiosis (Table 5 and Fig. 2).

Although the microcosm biofilm model is a well-established practice, there are several differences between the protocols, as for example the growth atmosphere. Accordingly, the atmosphere that better represents oral conditions needs to be investigated. This study did not aim to validate a type of atmosphere based on the oral environment since no comparison was made between the tested protocol and in vivo conditions. This study intended to check the differences between the atmospheric models (with or without O₂), in vitro, and their relevance in the development of artificial dental caries.

Some minor differences were observed among the atmospheric conditions, but they were able to produce very similar carious lesions in both enamel and dentin. Furthermore, most atmospheres were able to differentiate between CHX and PBS, which is essential outcome, since the antimicrobial effect of CHX has been well established (Shapiro et al. 2002).

From a microbiological perspective, to analyze the effect of the atmosphere on the microorganisms from the microcosm biofilm, we selected CFU counting, which is a well-established, reproducible, and simple quantitative method (van Houte et al. 1993). However, it is not as precise and sensitive as PCR- Polymerase chain reaction method and does not provide an overview of the entire microbiota (Tanner et al. 2018). Based on the findings of the CFU count, there was a lesser number of Lactobacillus spp. and S. mutans observed under the microaerophilic than in the anaerobic environment, especially for enamel. The differences less than 0.2 log₁₀ CFU/mL might be clinically irrelevant and, in fact, they were important when TMR data were taken into account.

Both bacteria were chosen because they are highly established cariogenic species (Gao et al. 2016). A limitation of the study is that other species, which might be present in the microcosm biofilm, were not examined. Therefore, future studies using “omics” are desirable, especially to understand the reason that CHX did not reduce Lactobacillus spp. in microcosm biofilms produced on dentin. Interestingly, the number of total microorganisms was reduced under anaerobic conditions for CHX. This led to speculations regarding the contribution of other species. Herein, it is important to consider that the term total microorganism does not include species that require supplementation for their growth on BHI agar.

The association between lactobacilli and dental caries dates back to a century (Kligler 1915; Caufield et al. 2015). In another study, 0.2% chlorhexidine, utilized in an in situ model, was not able to reduce the CFU count for lactobacilli on dentin specimens when compared with the control (van Strijp et al., 1997). The author suggested that dentin can act as a "shelter” for this bacterium against the action of chlorhexidine (van Strijp et al., 1997).

The outside atmosphere had little impact on the microbiological analysis of the microcosm biofilms because the biofilm itself can create its atmosphere, with the deeper layers rich in strict anaerobic microorganisms and the superficial layers rich in facultative microorganisms (Diaz et al. 2006; Schoilew et al. 2019). Although dental biofilms are composed primarily of obligate anaerobic species with preferential growth in the presence of carbon dioxide (CO₂), these microorganisms may be protected from the toxic effects of oxygen, enabling their growth under microaerophilic conditions (Bradshaw et al. 1997; Sim et al. 2016). It should be noted that the findings of this study cannot be extrapolated to monospecies or other multispecies models.

Despite knowing the microbiological composition of the biofilm, it was clear by the tooth lesion analysis that even in the presence of different species, the set of metabolic products released into the biofilms under different atmospheres was able to induce similar artificial carious lesions. Metabolome analysis could be better used to discuss the results; however, the most important response variable is related to the tooth (TMR data).

Regarding enamel demineralization, the mineral loss (integrated and mean) was similar among different atmospheres; however, the anaerobic condition induced a deeper lesion, showing that in this case, the acids could have penetrated deeper into the environment, which may be related to the metabolic profile of the biofilm. While dentin lesion depth was similar among the atmospheres, dentin mineral loss (integrated and mean) was lower for anaerobic than for microaerophilic and mixed atmospheres, which behaved similarly. Therefore, as previously discussed, the metabolic products (primarily acids) induced by the anaerobic biofilm may have been neutralized by the organic content of dentin (Takahashi 2015; Wicaksono et al. 2020).

The mixed biofilm model was first performed by Braga et al. (2021), but with a different sequence: the first three days in an anaerobic and the last two days in a microaerophilic atmosphere. In their study, the biofilm model was able to differentiate CHX from PBS. The results of this study warrant further metabolome analysis to better explain the slight differences found in CFU counting and TMR analysis.

For both tissues, especially enamel, lesions induced in the absence of treatment (PBS group) were highly demineralized, and most of them were cavitated (about 85–100% for enamel and 31–55% for dentin), with no differences among the atmospheres. It is suggested that the higher cavitation of enamel is due to its lower organic content compared to dentin, which plays an important role in the modulation of de-mineralization progression. Also, the percentage of prevention fraction of CHX was much higher for enamel (~ 78%) than for dentin (~ 22%), which may be related to the high degree of de-mineralization of the enamel and the type of interaction of CHX with the biofilm, in agreement with previous reports (Santos et al. 2019; Braga et al. 2020; Pelá et al. 2021).

Our results provide new information about the effect of the atmosphere on microcosm biofilm growth and its potential to induce tooth demineralization. These different protocols should be analyzed in the future to identify their impact on the results and interpretation of metabolome. In addition, the biofilm should be analyzed by laser scanning microscopy or scanning electron microscopy using the 3D architecture of the biofilm to identify if the specific organization of microbial communities could be created by different atmospheric conditions.

In conclusion, this study identified some minor differences among the atmospheric conditions; however, in general, microcosm biofilms produced under microaerophilia, anaerobiosis, or mixed models produced very similar artificial carious lesions in both enamel and dentin.

Ethics approval and consent to participate: We collected human saliva from ten participants, who approved and signed the consent to participate of the present study. The study was first approved by by the local ethics committee of Bauru School of Dentistry-USP (Bauru-Brazil), number CAAE: 35403320.1.0000.5417.

Consent for publication: Not applicable.

Availability of data and material: Please contact the corresponding author for data requests.

Funding

This work was supported by the São Paulo Research Foundation (FAPESP Proc. 2019/21797-0; 2019/01730-9; 2017/17249-2). This funding source had no involvement in the study design; in the collection, analysis and interpretation of data; in the writing of the manuscript; and in the decision to submit the article for publication.

Authors' contributions

A.S.B.: Conceptualization, Methodology, Validation, Formal analysis, Investigation, Resources, Writing - Review & Editing, Visualization; R.R.K.: Methodology, Validation, Investigation, Resources, Funding acquisition; and A.C.M.: Conceptualization, Methodology, Validation, Formal analysis, Resources, Writing - Original Draft, Writing - Review & Editing, Visualization, Supervision, Project administration, Funding acquisition.

Authors' information: Not applicable.

Acknowledgments

We would like to thank the anonymous saliva donors and the São Paulo Research Foundation.

Disclosure Statement

The authors declare no potential conflict of interest.

Aas JA, Griffen AL, Dardis SR, Lee AM, Olsen I, Dewhirst FE, Leys EJ, Paster BJ (2008) Bacteria of dental caries in primary and permanent teeth in children and young adults. J Clin Microbiol 46(4):1407–1417. https://doi.org/10.1128/JCM.01410-07.
Amaechi BT, Tenuta L, Ricomini Filho AP, Cury JA (2019) Protocols to study dental caries in vitro: microbial caries models. Methods Mol biol 1922, 357–368. https://doi.org/10.1007/978-1-4939-9012-2_32.
Amend S, Frankenberger R, Lücker S, Domann E, Krämer N (2018) Secondary caries formation with a two-species biofilm artificial mouth. Dent Mater 34(5):786–796. https://doi.org/10.1016/j.dental.2018.02.002.
Ayoub HM, Gregory RL, Tang Q, Lippert F (2020) Comparison of human and bovine enamel in a microbial caries model at different biofilm maturations. J Dent 96:103328. https://doi.org/10.1016/j.jdent.2020.103328.
Bowen WH (2002) Do we need to be concerned about dental caries in the coming millennium? Crit Rev Oral Biol Me 13(2):126–131. https://doi.org/10.1177/154411130201300203.
Bradshaw DJ, Marsh PD, Watson GK (1997) Oral anaerobes cannot survive oxygen stress without interacting with facultative/aerobic species as a microbial community. Lett Appl Microbiol 25(6):385–387. https://doi.org/10.1111/j.1472-765X.1997.tb00001.x.
Braga AS, de Melo F, Saldanha LL, Dokkedal AL, Meissner T, Bemmann M, Schulz-Kornas E, Haak, R. Abdelbary M, Conrads G, Magalhães AC, Esteves-Oliveira M (2021) The effect of solutions containing extracts of Vochysia tucanorum Mart., Myrcia bella Cambess., Matricaria chamomilla L. and Malva sylvestris L. on cariogenic bacterial species and enamel caries development. Caries Res 55(3):193–204. https://doi.org/10.1159/000515234.
Braga AS, Girotti LD, de Melo Simas LL, Pires JG, Pelá, VT, Buzalaf M, Magalhães A C (2019) Effect of commercial herbal toothpastes and mouth rinses on the prevention of enamel demineralization using a microcosm biofilm model. Biofouling 35(7):796–804. https://doi.org/10.1080/08927014.2019.1662897.
Braga AS, Simas L, Pires JG, Souza BM, de Melo F, Saldanha LL, Dokkedal AL, Magalhães AC (2020) Antibiofilm and anti-caries effects of an experimental mouth rinse containing Matricaria chamomilla L. extract under microcosm biofilm on enamel. J Dent 99:103415. https://doi.org/10.1016/j.jdent.2020.103415.
Buchalla W, Attin T, Roth P, Hellwig E (2003) Influence of olive oil emulsions on dentin demineralization in vitro. Caries Res 37(2):100–107. https://doi.org/10.1159/000069017.
Caufield PW, Schön CN, Saraithong P, Li Y, Argimón S (2015) Oral Lactobacilli and Dental Caries: A Model for Niche Adaptation in Humans. J Dent Res 94(9 Suppl):110S–8S. https://doi.org/10.1177/0022034515576052.
Dhaked H, Cao L, Biswas I (2021) Redox Sensing Modulates the Activity of the ComE Response Regulator of Streptococcus mutans. J Bacteriol 203(23):e0033021. https://doi.org/10.1128/JB.00330-21.
Diaz PI, Chalmers NI, Rickard AH, Kong C, Milburn CL, Palmer Jr RJ, Kolenbrander PE (2006) Molecular characterization of subject-specific oral microflora during initial colonization of enamel. Appl Environ Microbiol 72(4):2837–2848. https://doi.org/10.1128/AEM.72.4.2837-2848.2006.
Filoche SK, Soma KJ, Sissons CH (2007) Caries-related plaque microcosm biofilms developed in microplates. Oral Microbiol Immunol 22(2):73–79. https://doi.org/10.1111/j.1399-302X.2007.00323.x.
Gao X, Jiang S, Koh D, Hsu CY (2016) Salivary biomarkers for dental caries. Periodontol 2000 70(1):128–141. https://doi.org/10.1111/prd.12100.
Gross EL, Leys EJ, Gasparovich SR, Firestone ND, Schwartzbaum JA, Janies DA, Asnani K, Griffen AL (2010) Bacterial 16S sequence analysis of severe caries in young permanent teeth. J Clin Microbiol 48(11):4121–4128. https://doi.org/10.1128/JCM.01232-10.
Henne K, Rheinberg A, Melzer-Krick B, Conrads G (2015) Aciduric microbial taxa including Scardovia wiggsiae and Bifidobacterium spp. in caries and caries free subjects. Anaerobe 35(Pt A):60–65. https://doi.org/10.1016/j.anaerobe.2015.04.011.
Kligler IJ, Gies WJ (1915) Chemical studies of the relations of oral microorganisms to dental caries. J Allied Dental Soc 10:141–166.
Kolenbrander PE (2000) Oral microbial communities: biofilms, interactions, and genetic systems. Annu Rev Microbiol 54:413–437. https://doi.org/10.1146/annurev.micro.54.1.413.
Li F, Weir M D, Fouad AF, Xu HH (2014) Effect of salivary pellicle on antibacterial activity of novel antibacterial dental adhesives using a dental plaque microcosm biofilm model. Dent Mater 30(2):182–191. https://doi.org/10.1016/j.dental.2013.11.004.
Lima JP, Sampaio de Melo MA, Borges FM, Teixeira AH, Steiner-Oliveira C, Nobre Dos Santos M, Rodrigues LK, Zanin IC (2009) Evaluation of the antimicrobial effect of photodynamic antimicrobial therapy in an in situ model of dentine caries. Eur J Oral Sci 117(5):568–574. https://doi.org/10.1111/j.1600-0722.2009.00662.x.
Marsh PD, Moter A, Devine DA (2011) Dental plaque biofilms: communities, conflict and control. Periodontol 2000 55(1):16–35. https://doi.org/10.1111/j.1600-0757.2009.00339.x.
Maske TT, Brauner KV, Nakanishi L, Arthur RA, van de Sande FH, Cenci MS (2016) An in vitro dynamic microcosm biofilm model for caries lesion development and antimicrobial dose-response studies. Biofouling 32(3):339–348. https://doi.org/10.1080/08927014.2015.1130824.
Maske TT, van de Sande FH, Arthur RA, Huysmans M, Cenci MS (2017) In vitro biofilm models to study dental caries: a systematic review. Biofouling 33(8):661–675. https://doi.org/10.1080/08927014.2017.1354248.
McBain A. J. (2009). Chapter 4: In vitro biofilm models: an overview. Adv Appl Microbiol 69:99–132. https://doi.org/10.1016/S0065-2164(09)69004-3.
Méndez DAC, Gutierres E, José Dionisio E, Afonso Rabelo Buzalaf M, Cardoso Oliveira R, Andrade Moreira Machado MA, Cruvinel T (2018) Curcumin-mediated antimicrobial photodynamic therapy reduces the viability and vitality of infected dentin caries microcosms. Photodiagnosis Photodyn Ther 24:102–108. https://doi.org/10.1016/j.pdpdt.2018.09.007.
Mosaddad SA, Tahmasebi E, Yazdanian A, Rezvani MB, Seifalian A, Yazdanian M, Tebyanian H (2019) Oral microbial biofilms: an update. Eur J Clin Microbiol Infect Dis 38(11):2005–2019. https://doi.org/10.1007/s10096-019-03641-9.
Parisotto TM, Steiner-Oliveira C, Duque C, Peres RC, Rodrigues LK, Nobre-dos-Santos M (2010) Relationship among microbiological composition and presence of dental plaque, sugar exposure, social factors and different stages of early childhood caries. Arch Oral Biol 55(5):365–373. https://doi.org/10.1016/j.archoralbio.2010.03.005.
Pelá VT, Braga AS, Camiloti GD, Lunardelli J, Pires JG, Toyama D, Santiago AC, Henrique-Silva F, Magalhães AC, Buzalaf M (2021) Antimicrobial and anti-caries effects of a novel cystatin from sugarcane on saliva-derived multi-species biofilms. Swiss Dent J 131(5):410–416.
Pires JG, Braga AS, Andrade FB, Saldanha LL, Dokkedal AL, Oliveira RC, Magalhães AC (2019) Effect of hydroalcoholic extract of Myracrodruon urundeuva All. and Qualea grandiflora Mart. leaves on the viability and activity of microcosm biofilm and on enamel demineralization. J Appl Oral Sci 27:e20180514. https://doi.org/10.1590/1678-7757-2018-0514.
Pitts NB, Zero DT, Marsh PD, Ekstrand K, Weintraub JA, Ramos-Gomez F, Tagami J, Twetman S, Tsakos G, Ismail A (2017) Dental caries. Nat Rev Dis Primers 3:17030. https://doi.org/10.1038/nrdp.2017.30.
Pratten J, Wilson M, Spratt DA (2003) Characterization of in vitro oral bacterial biofilms by traditional and molecular methods. Oral Microbiol Immunol 18(1):45–49. https://doi.org/10.1034/j.1399-302x.2003.180107.x.
Rudney JD, Chen R, Lenton P, Li J, Li Y, Jones RS, Reilly C, Fok AS, Aparicio C (2012) A reproducible oral microcosm biofilm model for testing dental materials. J Appl Microbiol 113(6):1540–1553. https://doi.org/10.1111/j.1365-2672.2012.05439.x.
Santos D, Pires JG, Braga AS, Salomão P, Magalhães AC (2019) Comparison between static and semi-dynamic models for microcosm biofilm formation on dentin. J Appl Oral Sci 27:e20180163. https://doi.org/10.1590/1678-7757-2018-0163.
Schoilew K, Ueffing H, Dalpke A, Wolff B, Frese C, Wolff D, Boutin S (2019) Bacterial biofilm composition in healthy subjects with and without caries experience. J Oral Microbiol 11(1): 1633194. https://doi.org/10.1080/20002297.2019.1633194.
Schwendicke F, Korte F, Dörfer CE, Kneist S, Fawzy El-Sayed K, Paris S (2017) Inhibition of Streptococcus mutans Growth and Biofilm Formation by Probiotics in vitro. Caries Res 51(2):87–95. https://doi.org/10.1159/000452960.
Shapiro S, Giertsen E, Guggenheim B (2002) An in vitro oral biofilm model for comparing the efficacy of antimicrobial mouthrinses. Caries Res 36(2):93–100. https://doi.org/10.1159/000057866.
Signori C, Hartwig AD, Silva-Júnior I, Correa MB, Azevedo MS, Cenci MS (2018) The role of human milk and sucrose on cariogenicity of microcosm biofilms. Braz Oral Res 32:e109. https://doi.org/10.1590/1807-3107bor-2018.vol32.0109.
Silva TC, Pereira AF, Exterkate RA, Bagnato VS, Buzalaf MA, Machado MA, Ten Cate JM, Crielaard W, Deng DM (2012) Application of an active attachment model as a high-throughput demineralization biofilm model. J Dent 40(1):41–47. https://doi.org/10.1016/j.jdent.2011.09.009.
Sim CP, Dashper SG, Reynolds EC (2016) Oral microbial biofilm models and their application to the testing of anticariogenic agents. J Dent 50:1–11. https://doi.org/10.1016/j.jdent.2016.04.010.
Souza BM, Santos D, Magalhães AC (2018) Antimicrobial and anti-caries effect of new glass ionomer cement on enamel under microcosm biofilm model. Braz Dent J 29(6):599–605. https://doi.org/10.1590/0103-6440201802163.
Takahashi N (2015) Oral microbiome metabolism: From "Who are they?" to "What are they doing?". J Dent Res. 94(12):1628–1637. https://doi.org/10.1177/0022034515606045.
Tanner A, Kressirer CA, Rothmiller S, Johansson I, Chalmers NI (2018) The Caries Microbiome: Implications for Reversing Dysbiosis. J Adv Dent Res 29(1):78–85. https://doi.org/10.1177/0022034517736496.
van Houte J (1993) Microbiological predictors of caries risk. J Adv Dent Res 7(2):87–96. https://doi.org/10.1177/08959374930070022001.
van Strijp AJ, van Steenbergen TJ, ten Cate JM (1997) Effects of chlorhexidine on the bacterial colonization and degradation of dentin and completely demineralized dentin in situ. Eur Oral Sci 105(1):27–35. https://doi.org/10.1111/j.1600-0722.1997.tb00177.x.
Vertuan M, da Silva JF, Braga AS, de Souza BM, Magalhães AC (2021) Effect of TiF₄/NaF and chitosan solutions on biofilm formation and prevention of dentin demineralization. Arch Oral Biol 132:105275. https://doi.org/10.1016/j.archoralbio.2021.105275.
Vieira DR, Amaral FM, Maciel MC, Nascimento FR, Libério SA, Rodrigues VP (2014) Plant species used in dental diseases: ethnopharmacology aspects and antimicrobial activity evaluation. J Ethnopharmacol 155(3):1441–1449. https://doi.org/10.1016/j.jep.2014.07.021.
Walsh T, Oliveira-Neto JM, Moore D (2015) Chlorhexidine treatment for the prevention of dental caries in children and adolescents. Cochrane Database Syst Rev (4):CD008457. https://doi.org/10.1002/14651858.CD008457.pub2.
Wicaksono DP, Washio J, Abiko Y, Domon H, Takahashi N (2020) Nitrite Production from Nitrate and Its Link with Lactate Metabolism in Oral Veillonella spp. Appl Environ Microbiol 86(20):e01255-20. https://doi.org/10.1128/AEM.01255-20.
Zhang N, Ma J, Melo MA, Weir MD, Bai Y, Xu HH (2015) Protein-repellent and antibacterial dental composite to inhibit biofilms and caries. J Dent 43(2):225–234. https://doi.org/10.1016/j.jdent.2014.11.008.

No competing interests reported.

Download PDF

Journal Publication

published 01 Jun, 2023

Read the published version in Journal of Applied Oral Science →

Version 1

posted

You are reading this latest preprint version

Effect of different atmospheres on microcosm biofilm formation and tooth demineralization

Status:

Journal Publication

Version 1

Abstract

Figures

Introduction

Materials And Methods

Saliva Collection

Tooth Specimen Preparation

Study conditions and sample size calculation

Preparation of Artificial Saliva

Microcosm Biofilm Formation

Colony-forming units (CFU) counting

Transverse microradiography (TMR)

Statistical analysis

Results

CFU counting

Enamel

Dentin

TMR analysis

Discussion

Conclusion

Declarations

References

Additional Declarations

Status:

Journal Publication

Version 1