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Brazilian Journal of Microbiology logoLink to Brazilian Journal of Microbiology
. 2024 Oct 31;55(4):3921–3932. doi: 10.1007/s42770-024-01550-4

High release of Candida albicans eDNA as protection for the scaffolding of polymicrobial biofilm formed with Staphylococcus aureus and Streptococcus mutans against the enzymatic activity of DNase I

Santiago M Lattar 1,✉, Rene Peter Schneider 2, Vidal Jorge Eugenio 3, Gabriel Padilla 4,✉
PMCID: PMC11711860  PMID: 39480631

Abstract

This study aimed to determine the protective role of the high release of C. albicans extracellular DNA (eDNA) in a polymicrobial biofilm formed by S. aureus and S. mutans in the course of DNase I treatment. A tube-flow biofilm bioreactor was developed to mimic biofilm formation in the oral cavity. eDNA release was quantified by real-time PCR (qPCR) and confocal microscopy analysis were used to determine the concentration and distribution of eDNA and intracellular DNA (iDNA). The mean amount of eDNA released by each species in the polymicrobial was higher than that in monospecies biofilms (S. aureus: 3.1 × 10−2 ng/μl polymicrobial versus 5.1 × 10−4 ng/μl monospecies; S. mutans: 3 × 10−1 ng/μl polymicrobial versus 2.97 × 10−2 ng/μl monospecies; C. albicans: 8.35 ng/μl polymicrobial versus 4.85 ng/μl monospecies). The large amounts of eDNA released by C. albicans (96%) in polymicrobial biofilms protects the S. aureus and S. mutans cells against the degradation by DNase I and dampens the effect of clindamycin.

Keywords: C. albicans, Biofilms, Staphylococcus aureus, Streptococcus mutans, Antibiotics

Introduction

Oral biofilms can be defined as a cluster of bacterial cells embedded in a matrix [1]. The biofilm matrix is mainly composed of polysaccharides, proteins, and extracellular DNA (eDNA) [2]. Moreover, eDNA has been revealed as an important component of the extracellular matrix of multicellular communities, such as biofilms formed by bacteria, archaea, and fungi [3]. The proposed functions for eDNA in the biofilms are (i) structural component within the biofilm that provides stability [4]; (ii) a signal factor that promotes biofilm formation and extracellular matrix production as evidenced by the fact that treatment with DNase I inhibits biofilm formation by Pseudomonas aeruginosa [5, 6]; and (iii) stimulates gene transfer by transformation of competent sister bacteria [7]. Additionally, eDNA has been proven to play a role in maintaining cell-to-cell interactions within biofilms and to contribute to antibiotic resistance [8]. Clindamycin (CLi), a protein synthesis inhibitor used in the treatment of Gram-positive bacterial infections, poorly penetrates microbial biofilms, but its combination with DNase I increases it action on biofilm cells [9].

Release of eDNA from Staphylococcus epidermidis and Candida albicans is increased in mixed-species biofilms [10]. Staphylococcus aureus, Streptococcus mutans, and Candida albicans are pathogenic microorganisms that can be isolated from the oral cavity [11]. S. mutans is one of the most important cariogenic bacteria [12]. S. mutans eDNA is released not only by cell lysis, but also inside membrane vesicles during different phases of biofilm development [16]. S. aureus is a Gram-positive opportunistic bacterium that produces biofilms under stress circumstances with eDNA a major structural component in the biofilm matrix [13–15, 17]. C. albicans is a dimorphic fungal pathogen that causes systemic candidiasis [18]. eDNA plays a crucial role in C. albicans biofilm formation and scaffolding integrity [19]. In the context of human health, a crucial biofilm property is enhancement of protection during antimicrobial compound exposure [20].

The study of eDNA from C. albicans, S. mutans, and S. aureus biofilms is important because antibiotic resistance is enhanced by biofilms, and eDNA plays a role in maintaining the biofilm scaffolding and should be considered a possible target for modifying biofilms; eDNA release is essential for biofilm formation and structural integrity and bacteria in biofilms have stronger resilience compared to planktonic bacteria because the biofilm serves as a physical shield against potential antibiotics such as clindamycin. There is also evidence that eDNA removal can increase the susceptibility of microbial cells in biofilms to various antimicrobial agents [16, 18]. Many oral bacteria potentially utilize eDNA as nutrient storage for phosphate, fixed nitrogen, and carbon [20]. Furthermore, eDNA is important for establishing the transfer of genetic information within oral biofilms [20].

In this study, a modified biofilm bioreactor [21] was designed to investigate eDNA liberation from biofilms by three clinically important microorganisms, S. aureus, S. mutans, and C. albicans associated with oral human infections. Molecular studies included the quantification of eDNA by quantitative real-time PCR (qPCR) using qPCR oligonucleotides specific for each strain. Much is known about the role of eDNA in biofilm, but we do not know the role of eDNA in the protection of microorganisms in the polymicrobial biofilm. We check the role of C. albicans eDNA in the protection of the polymicrobial biofilm from the action of the DNase I enzyme and the CLi. In this paper we demonstrate for the first time how the high eDNA release of one species (C. albicans) protects other species (S. mutans and S. aureus) in the polymicrobial biofilm.

Materials and methods

Bacterial strains and growth conditions

S. aureus ATCC 27853, S, mutans ATCC 25175 and C. albicans ATCC 10231 were routinely grown on salt mannitol agar (Difco Laboratories), Mitis Salivarius agar (Difco Laboratories) + bacitracin + potassium tellurite, and Sabouraud dextrose agar (Difco Laboratories) + erythromycin, respectively.

Natural not stimulated saliva collection

A total of 70 ml of not stimulated natural saliva was collected purified and sterilized as previously described [22].

Biofilm assays into the tube-flow biofilm bioreactor

The tube-flow biofilm bioreactor system (Fig. 1) was a modification of the bioreactor system previously described [21, 23]. The medium reservoir was a sterilizable glass balloon that contained 250 ml BHI that flowed from top to bottom by gravity through a 7.0 × 0.3 cm autoclavable silicone tube. Flow was regulated with a flow valve (0.25 ml/min for 16 h). The reactor was operated at room temperature in sterile conditions. During the experiments, 500 μl of sterile not stimulated natural saliva was added into the autoclaved silicone tube and incubated for 3 h at 37 °C to allow salivary pellicle formation. The saliva was then discarded and substituted by an inoculation mix containing 500 μl. of BHI, saliva, and 2 × 106 CFU (colony forming unit) of each microorganism for monospecies biofilm, or 2 × 106 CFU microbial counts for each strain for the polymicrobial biofilms. Colonization of the internal surfaces of the tube-flow biofilm bioreactor was achieved by contacting them for 3 h at 37 °C in microaerophilic conditions with the microorganism inoculum. The colonized tube was then connected to the medium reservoir and percolated for 16 h. Cells eroded from the biofilm were collected at the reactor outlet in a collector flask.

Fig. 1.

Fig. 1

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Tube-flow biofilm bioreactor

Viable cells in the samples (CFU/ml) were quantified by dilution plating. The biofilms were removed from the flow tubes with 1 ml of PBS (2.7 mmol /l KCl, 138 mmol /l NaCl, 1.5 mmol/ l KH2PO4, and 20.4 mmol/l NaH2PO4 [pH 7.2]) then disintegrated in a vortex mixer for 60 s, followed by serial decimal dilutions in PBS. Subsequently, 100 μl aliquots from the dilutions (10−4, 10−5, and 10−6) were inoculated on Petri dishes prepared with culture media specific for each microorganism, previously described above and incubated for 48 h at 37 °C.

Purifications and quantifications of extracellular DNA by qPCR

The biofilm was removed from the drained flow-tube surface by vortexing three times in 1 ml PBS at 25 °C for 5 min, followed by separation of cells from matrix material by centrifugation at 12.000 g for 15 min at 4 °C. Cells were removed from the supernatant containing eDNA by filtration (0.22 µm) and the supernatant was subsequently stored at − 20 °C [24]. eDNA was quantified using qPCR. Reactions were performed with SYBR green super mix (Bio-Rad), using 6 µl of mix, 2 µl of 2 qPCR oligonucleotides (20 µmol/l), and 2 µl of eDNA sample. The final volume of 10 µl (mix, primers and samples for qPCR), was dispensed into 0.2 ml flat optic cap tubes (4ti-0790-flat optical cap, SINAPSE). The following qPCR primers were used: 16sRNA F 5-GCAAGCGTTATCCGGATTT-3, 16sRNA R 5-CTTAATGATG GCAACTAAGC-3 [25], gtfB F 5-GCCTACAGCTCAGAGATGCTAT TCT-3, gtfB R 5 GCCATACACCACTCATGAATTGA-3 [26], cat1 F 5-GATTCTCTACT GTTGGTGGTG-3, and CAT1 R 5-GTGAGTTTCTGGGTTTCTCTT-3 [27]. Reactions were run in duplicate using a Rotor Gene 6000 qPCR detection system (Corbett). The qPCR protocol for S. aureus (16S rRNA reference gene) and S. mutans (gtfB reference gene) eDNA analyses started with an initial denaturation (Cq) at 95 °C for 10 min, followed by 45 cycles with denaturation at 95 °C for 20 s, reannealing at 58 °C for 20 s, and extension at 72 °C for 20 s. eDNA quantification of C. albicans (CAT1 reference gene) started with an initial denaturation at 95 °C for 10 min, followed by 40 cycles of denaturation at 95 °C for 15 s, reannealing at 57 °C for 15 s, and amplification at 72 °C for 20 s. A standard curve with different concentrations of genomic DNA (0.0001 ng/µl, 0.001 ng / µl, 0.01 ng/μl, 0.1 ng/μl, 1 ng/μl, and 10 ng/μl per reaction) from either S. aureus, S. mutans, or C. albicans per reaction was established in parallel during qPCR assays. The amounts of eDNA in the samples were calculated in ng/μl using Bio-Rad CFX Manager (Bio-Rad) software. This study complies with the MIQE guidelines [28] (where applicable).

Biofilm treatment with DNase I, CLi or DNase I and CLi

Biofilms were pretreated with DNase I 500 µg/ml (Roche) for 3 h at 37 °C in the flow-tube bioreactor. Subsequently, the tubes were percolated with 250 ml of BHI containing DNase I 20 µg/ml for 16 h. For testing the effect of CLi on biofilm development, the pretreated DNase I tubes were exposed to 250 ml of BHI with CLi 1 µg/ml (Sigma) at flow rate for 16 h in the flow-tube bioreactor. Finally, the combined effect of DNase I and CLi on the development of biofilms pre-treatment with 500 µg/ml DNase I was investigated by percolating the tubes with 250 ml of BHI with DNase I 20 µg/ml and CLi 1 µg/ml (Sigma) for 16 h.

Intra- and extracellular DNA labeling and confocal microscopy (CM) Zeiss LSM 780-NLO analysis

Intracellular DNA (iDNA) and eDNA in biofilms were stained with Hoechst (bisBenzimide 33,342, Sigma Aldrich, St. Louis, Missouri, USA) and propidium iodide (SigmaAldrich, St. Louis, Missouri, USA), respectively, following the procedure described in the article [29]. The mean fluorescence intensity of eDNA and iDNA was obtained using the Fiji program [30]. For analysis using CM Zeiss LSM 780-NLO analysis, the following patterns were used: the step size: 1.84 μm, objective: EC Plan-Neofluar 20x/0.50 M27, resolution: 2.40 pixels/μm, laser power: 30% (Diode 405) and 25% (Argon 488) gain: 668 (Diode 405) and 897 (Argon 488), imaging software: Zen 2011 SP7.

Statistical methods

The experiments were performed with at least three independent biological replicates. The dependent variables are eDNA concentration and CFU. The independent variables are the controls, treatments (DNase I, CLi and DNase I + CLi), and strains (S. aureus, S. mutans, and C. albicans). A nonparametric methodology was used for multiple comparison implemented in the nparcomp [31] software R package. This methodology considers that there is more than one group (characterized by the categories of independent variables), so that the comparison between the groups is made jointly, that is, the p-values ​​are adjusted to express the fact that there are more than two groups and more than one comparison. The non-parametric test, proposed in [40], determines which of two groups tends to have a higher value in the variable, that is, if we randomly select an individual from each group, which of the two has the greater probability of having a higher value in the variable. Statistical significance is determined at p = 0.0001.

Results

Release of species-specific eDNA from Biofilms grown in the tube-flow biofilms bioreactor with surfaces conditioned with human saliva

The tube-flow biofilm bioreactor covered with human saliva developed in this study (Fig. 1) allowed the investigation of biofilms produced by either pure or mixed cultures of potential pathogens on surfaces conditioned with a salivary film, such as it would occur in the human oral cavity. eDNA is an important component of the biofilm matrix. The concentration of species-specific eDNA in polymicrobial biofilms was significantly higher than that of monospecies biofilms. Polymicrobial biofilms contained much higher amounts of eDNA, than the monospecies biofilms, S. aureus polymicrobial 3.1.10–2 ng/µl and monospecies 5.10–4 ng/µl (p = 0.0001); S. mutans polymicrobial 3.10–1 ng/µl and monospecies 2.97.10–2 ng/µl (p = 0.0001) and C. albicans polymicrobial 8.35 ng/µl and monospecies 4.85 ng/µl (p = 0.0001) (Fig. 2).

Fig. 2.

Fig. 2

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eDNA from monospecies and polymicrobial biofilm

Relative to values measured in pure culture biofilms, the eDNA in polymicrobial biofilms increased ~ 100-fold for S. aureus (p = 0.0001), ~ ninefold for S. mutans (p = 0.0001) and about twofold for C. albicans (p = 0.0001) (Fig. 2). C. albicans released more eDNA than the other two species, in both monospecies and polymicrobial biofilms. Amongst the bacterial species, eDNA release by S. mutants was 40-fold (p = 0.0001) and tenfold (p = 0.0001) higher than the eDNA released by S. aureus in monospecies and polymicrobial biofilms, respectively (Fig. 2). S. aureus, S. mutans and C. albicans released significantly more eDNA when growing in polymicrobial biofilms than in monospecies.

The high amount of eDNA in the biofilm should render the biofilm matrix susceptible to disaggregation by DNase I treatment

DNase I significantly reduced eDNA in polymicrobial biofilms in 21-fold of S. aureus; 12-fold from S. mutans and 7.3-fold from C. albicans (Fig. 3A), eDNA concentration in S. aureus before DNase I treatment was 3.1.10–2 ng/µl and after DNase I treatment was 1.47.10–3 ng/µl (p = 0.0001); eDNA concentration in S. mutans before DNase I treatment was 3.10–1 ng/µl and after DNase I treatment was 2.47 0.10–2 ng/µl (p = 0.0001) and eDNA concentration in C. albicans before DNase I treatment was 8.35 ng/µl and after DNase I treatment was 1.15 ng/µl (p = 0.0001) (Fig. 3A).

Fig. 3.

Fig. 3

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eDNA and colony forming unit (CFU) from polymicrobial and monospecies biofilm trated with DNase I

Bacterial and fungal cellular density was only slightly affected by the reduction of eDNA, with only a 1.4-fold; 1.33-fold and 1.97-fold decrease in density of S. aureus, S. mutans and C. albicans, respectively (Fig. 3B); cell density of S. aureus before DNase I treatment was 4.63. 106 UFC/ml and after DNase I treatment was 3.3. 106 UFC/ml (ns); cell density of S. mutans before DNase I treatment was 1.33. 105 UFC/ml and after DNase I treatment was 9.95. 104 UFC/ml (ns) and C. albicans before DNase I treatment was 5.34. 105 UFC/ml and after DNase I treatment was 2.7. 105 UFC/ml (ns) (Fig. 3B).

A high amount of eDNA in the matrix of biofilms should turn the matrix unstable in the presence of a DNA degrading enzymes such as DNase I. Exposure of the biofilms to the enzyme in the growth phase should result in lower eDNA contents in their matrices as well lower cell densities on the surfaces. Treatment of monospecies biofilms with DNase I led to a 102-fold, 7.61-fold, and fourfold reduction of S. aureus, S. mutans, and C. albicans eDNA in the biofilm, respectively (Fig. 3C); eDNA concentration in S. aureus before DNase I treatment was 5.1. 10–4 ng/µl and after DNase I treatment was 5.10–6 ng/µl (p = 0.0001); eDNA concentration in S. mutans before DNase I treatment was 2.97.10–2 ng/µl and after DNase I treatment was 3.9.10 −3 ng/µl (p = 0.0001) and eDNA concentration in C. albicans before DNase I treatment was 4.87 ng/µl and after DNase I treatment was 1.1 ng/µl (p = 0.0001) (Fig. 3C). The corresponding decrease of cell density was 12.3-fold, 5.6-fold, and 3.4-fold for S. aureus, S. mutans, and C. albicans biofilms, respectively (Fig. 3D); cell density of S. aureus before DNase I treatment was 1.89. 107 UFC/ml and after DNase I treatment was 1.54. 106 UFC/ml (p = 0.0001); S. mutans before DNase I treatment was 2.14 0.105 UFC/ml and after DNase I treatment was 3.8.104 UFC/ml (p = 0.0001) and C. albicans before DNase I treatment was 2.9 0.105 UFC/ml and after DNase I treatment was 8.5.104 UFC/ml (ns) (Fig. 3D).

CLi drastically reduced cell density and eDNA of prokaryotes in biofilms

CLi treatment of polymicrobial biofilms induced a dramatic decrease of eDNA in the supernatant of S. aureus by 12.000-fold, S. mutans by 19-fold and C. albicans by 12.5-fold; eDNA from S. aureus before CLi treatment was 3.1. 10–2 ng/µl and after CLi treatment was 2.5.10–6 ng/µl (p = 0.0001); S. mutans before CLi treatment was 3 0.10–1 ng/µl and after CLi treatment was 3.17.10–2 ng/µl (p = 0.0001) and C. albicans before CLi treatment was 8.35 ng/µl and after CLi treatment was 6.67.10–1 ng/µl (p = 0.0001) (Fig. 4A).

Fig. 4.

Fig. 4

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eDNA and colony forming unit (CFU) from polymicrobial and monospecies biofilm trated with Clindamycin

In polymicrobial biofilms the decrease of cell density after treatment with Clindamycin was 10.45-fold, 19-fold, and 1.4-fold for S. aureus, S. mutans, and C. albicans respectively; cell density of S. aureus before CLi treatment was 4.63. 106 UFC/ml and after CLi treatment was 4.43. 105 UFC/ml (p = 0.0001); S. mutans before CLi treatment was 1.33. 105 UFC/ml and after CLi treatment was 7.103 UFC/ml (p = 0.0001) and C. albicans before CLi treatment was 5.34 0.105 UFC/ml and after CLi treatment was 3.79.105 UFC/ml (ns) (Fig. 4B).

CLi treatment of monospecies biofilms induced a decrease of eDNA in the supernatant of S. aureus 214.6-fold, S. mutans 9.9-fold and C. albicans 0.8-fold respectively; eDNA from S. aureus before CLi treatment was 5. 10–4 ng/µl and after CLi treatment was 2.33. 10–6 ng/µl (p = 0.0001); S. mutans before CLi treatment was 2.97 0.10–2 ng/µl and after CLi treatment was 3.47.10–3 ng/µl (p = 0.0001) and C. albicans before CLi treatment was 4.85 0.10–1 ng/µl and after CLi treatment was 6.10–1 ng/µl (p = 0.0001) (Fig. 4C).

Treatment with CLi (1 µg/ml) drastically decreased the density of S. aureus and S. mutans in both polymicrobial and monospecies biofilms, but as expected, the population of C. albicans was not affected by the antibiotic (Fig. 4B and D). The corresponding decrease of cell density was 10.16-fold, 5.6-fold, and twofold for S. aureus, S. mutans, and C. albicans monospecies biofilms, respectively; cell density of S. aureus before CLi treatment was 1.89. 107 UFC/ml and after CLi treatment was 1.86. 106 UFC/ml (p = 0.0001); S. mutans before CLi treatment was 2.14 0.105 UFC/ml and after CLi treatment was 2.65.104 UFC/ml (p = 0.0001) and C. albicans before CLi treatment was 2.9 0.105 UFC/ml and after CLi treatment was 1.4.105 UFC/ml (ns) (Fig. 4D).

Combined exposure of biofilms to DNase I and CLi reduced the cellular density and eDNA of monospecies and polymicrobial biofilms

CLi and DNase I treatment from polymicrobial biofilms induced a dramatic decrease of eDNA in the supernatant of S. aureus by 1.161-fold, S. mutans by 39-fold and C. albicans by 11-fold respectively; eDNA from S. aureus before CLi and DNase I treatment was 3.1. 10–2 ng/µl and after Cli and DNase I treatment was 2.67.10–5 ng/µl (p = 0.0001); S. mutans before CLi and DNase I treatment was 3 0.10–1 ng/µl and after CLi and DNase I treatment was 7.7.10–3 ng/µl (p = 0.0001) and C. albicans before CLi and DNase I treatment was 8.35 ng/µl and after CLi and DNase I treatment was 7.67.10–1 ng/µl (p = 0.0001) (Fig. 5A).

Fig. 5.

Fig. 5

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eDNA and colony forming unit (CFU) from polymicrobial and monospecies biofilm trated with DNase I and clindamycin

In polymicrobial biofilm cell density of S. aureus before CLi and DNase I treatment was 4.63. 106 UFC/ml and after CLi and DNase I treatment was 1.72. 106 UFC/ml (p = 0.0001); S. mutans before Cli and DNase I treatment was 1.33. 105 UFC/ml and after CLi and DNase I treatment was 2.72. 103 UFC/ml (p = 0.0001) and C. albicans before CLi and DNase I treatment was 5.34 0.105 UFC/ml and after CLi and DNase I treatment was 2.76. 105 UFC/ml (ns) (Fig. 5B).

CLi and DNase I treatment of monospecie biofilms induced a decrease of eDNA in the supernatant of S. aureus by 18.7-fold, S. mutans by 14.85-fold and C. albicans by 32.33-fold; eDNA from S. aureus before CLi and DNase I treatment was 5. 10–4 ng/µl and after CLi and DNase I treatment was 2.67. 10–5 ng/µl (p = 0.0001); S. mutans before Cli and DNase I treatment was 2.97. 10–2 ng/µl and after CLi and DNase I treatment was 2.033. 10–3 ng/µl (p = 0.0001) and C. albicans before CLi and DNase I treatment was 4.85 ng/µl and after CLi and DNase I treatment was 1.5. 10–1 ng/µl (p = 0.0001) (Fig. 5C).

Treatment with CLi (1 µg/ml) and DNase I drastically decreased the cell density of S. aureus, S. mutans and C. albicans in both polymicrobial and monospecies biofilms. In monospecies biofilm cell density of S. aureus before CLi and DNase I treatment was 1.89. 107 UFC/ml and after Cli and DNase I treatment was 9.52. 105 UFC/ml (p = 0.0001); S. mutans before CLi and DNase I treatment was 2.14 0.105 UFC/ml and after CLi and DNase I treatment was 2.103 UFC/ml (p = 0.0001) and C. albicans before CLi treatment 2.9 0.105 UFC/ml and after CLi and DNase I treatment was 2.5.104 UFC/ml (ns) (Fig. 5D).

Ultrastructural analysis of S. aureus biofilms and post-treatment with DNase I and CLi

S. aureus cells forming the biofilm were stained with a membrane-permeable fluorophore (i.e., Hoechst) and a non-membrane-permeable fluorescence reporter. The confocal images reveal clear patches of dense biofilm populations whilst the border areas contained cells distributed in a more diffuse pattern (Fig. 6a1). The eDNA marker revealed some localized small clusters with intensive color, all of which were located in zones with strong iDNA marker emission (Fig. 6a2). eDNA was also present in a rather diffuse zone which coincided with regions of highest iDNA density (Fig. 6a1 and a2). The mean fluorescence intensity for iDNA was about 3.5 times that of eDNA (Table 1). The pattern of distribution of iDNA and eDNA fluorescence remained similar after DNase I treatment (Fig. 6b1 and b2), but the mean fluorescence intensity of both iDNA and eDNA decreased by about half. The proportion of iDNA to eDNA fluorescence remained similar to that of the untreated control (Table 1). Exposure of biofilms to clindamycin during growth resulted in a significantly lower surface colonization (Fig. 6c1 and c2). The biofilm consisted mostly of diffuse low cell density biofilm interconnected biofilm strings interspersed with only a few small high cell density clusters. Mean fluorescene intensity for both iDNA and eDNA as well as their proportion were similar to that observed after for DNase I exposed biofilms (Table 1). Growth of the biofilms exposed to both clindamycin and DNase I resulted in a stringly biofilm morphology similar to that observed for the individual treatments, with less dense iDNA cell clusters and almost no eDNA dense cell clusters (Fig. 6d1 and d2). The combination treatment resulted in a strong reduction of iDNA mean fluorescence to about 23% of the control value, whilst eDNA mean fluorescence remained at an intensity similar to that observed for the individual treatments. The strong reduction of iDNA fluoresce shifted the proportion of iDNA/eDNA fluorescence to about twofold (Table 1).

Fig. 6.

Fig. 6

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Confocal microscopy analysis from monospecies and polymicrobial biofilm

Table 1.

Mean fluorescence intensity of eDNA and iDNAfrom monospeciesand polymicrobial biofilms

Treatment DNA Biofilm type
S. aureus S. mutans C. albicans Polymicrobial

Control

DNaseI

 iDNA 27.022/μm2 20.982/μm2 14.743/μm2 14.143/μm2
eDNA 7.629/μm2 8.591/μm2 9.015/μm2 11.045/μm2
iDNA + eDNA 33.069/μm2 27.015/μm2 19.257/μm2 19.100/μm2
DNaseIiDNA 13.091/μm2 5.600/μm2 4.094/μm2 7.732/μm2
eDNA 3.419/μm2 3.190/μm2 3.630/μm2 3.954/μm2
CLi iDNA + eDNA 13.642/μm2 6.951/μm2 4.682/μm2 7.727/μm2
iDNA 13.291/μm2 3.270/μm2 10.464/μm2 5.199/μm2
eDNA 3.145/μm2 2.045/μm2 7.479/μm2 2.982/μm2
DNaseI + CLi iDNA + eDNA 13.352/μm2 3.745/μm2 15.245/μm2 5.678/μm2
iDNA 6.288/μm2 1.771/μm2 4.033/μm2 3.340/μm2
eDNA 3.241/μm2 1.909/μm2 3.568/μm2 3.727/μm2
iDNA + eDNA 6.930/μm2 2.187/μm2 5.243/μm2 5.338/μm2
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eDNA extracellular, DNA, iDNA intracellular DNA, DNase I Desoxyribonuclease I, CLi Clindamycin

Ultrastructural analysis of S. mutans biofilms and post-treatment with DNase I and CLi

S. mutans forming a biofilm with high cell density was stained with a membrane-permeable fluorophore. The eDNA marker revealed small clusters with intensive color, all of which were located in zones with strong iDNA marker emission (Fig. 6e1 and e2). The mean fluorescence intensity for iDNA was about 2.44 times that of eDNA (Table 1). Treatment of eDNA biofilm with DNase I produces a notable reduction in biofilm structure (Fig. 6f1 and f2). The distribution of iDNA and eDNA was different after DNase I treatment (Fig. 6f1 and f2). The proportion of iDNA to eDNA fluorescence was reduced 4 and 3 times respectively (Table 1). Treatment with Clindamycin during biofilm growth results in low colonization of the surface, enabling the formation of a biofilm with low cell density (Fig. 6g1 and g2). Mean fluorescence intensity for both iDNA and eDNA was reduced 6 and 4 times respectively (Table 1). Biofilm treatment with Clindamycin and DNase I produces a very strong reduction in cell density (Fig. 6h1 and h2). The strong reduction of iDNA fluoresce shifted the proportion of iDNA/eDNA fluorescence to about onefold (Table 1).

Ultrastructural analysis of C. albicans biofilms and post-treatment with DNase I and CLi

The confocal images reveal patches of dense biofilm populations whilst the border areas contained cells distributed in a diffuse pattern (Fig. 6i1). The eDNA marker revealed some localized small clusters with intensive color, all of which were located in zones with strong iDNA marker emission (Fig. 6i2).

The mean fluorescence intensity for iDNA was about 1.64 times that of eDNA (Table 1). The pattern of distribution of iDNA and eDNA fluorescence was different after DNase I treatment (Fig. 6j1 and j2), the proportion of iDNA to eDNA fluorescence was reduced 3.6 and 2.5 times respectively (Table 1). Treatment with Clindamycin during biofilm growth does not produce obvious changes in the biofilm (Fig. 6k1 and k2). The proportion of iDNA to eDNA fluorescence remained similar to that of the untreated control (Table 1). Biofilm treatment with Clindamycin and DNase I produces a reduction in cell density (Fig. 6l1 and l2). The strong reduction of iDNA fluoresce shifted the proportion of iDNA/eDNA fluorescence to about onefold (Table 1).

Ultrastructural analysis of polymicrobial biofilms and post-treatment with DNase I and CLi

Polymicrobial biofilm has a high cell density, the confocal images reveal clear patches of dense biofilm populations (Fig. 6m1). The eDNA marker revealed small clusters with intensive color, all of which were located in zones with strong iDNA marker emission (Fig. 6m2). The mean fluorescence intensity for iDNA was about 1.28 times that of eDNA (Table 1). Treatment of eDNA biofilm with DNase I produces a slight reduction in biofilm structure (Fig. 6n1 and n2); the proportion of iDNA to eDNA fluorescence was reduced 2 and 3 times respectively (Table 1). Treatment with Clindamycin during biofilm growth enabling the formation of a biofilm with low cell density (Fig. 6o1 and o2). Mean fluorescence intensity for both iDNA and eDNA as well as their proportion were similar to that observed after for DNase I exposed biofilms (Table 1). Biofilm treatment with Clindamycin and DNase I produces a very strong reduction in cell density (Fig. 6p1 and p2). The strong reduction of iDNA fluorescence shifted the proportion of iDNA/eDNA fluorescence to about onefold (Table 1).

Discussion

The tube-flow biofilms bioreactor covered with saliva is an excellent system for eDNA study in an experimental model mimicking oral biofilms (Fig. 1). To the best of our knowledge, this is the first study to describe the production and distributions of eDNA from viable polymicrobial and monospecies biofilms formed by S. aureus, S. mutans, and C. albicans into the tube-flow biofilm reactor. The bioreactor allowed us to efficiently test the effect of DNase I and CLi on the content of eDNA and viable cell counts in the biofilm.

Release of species-specific eDNA increases in polymicrobial biofilms. S. aureus, S. mutans, and C. albicans release more eDNA in polymicrobial biofilms. An enhanced release of the microbial eDNA was observed in multispecies biofilms, which may be caused by autolysis or cell death [5]. Our results showed that C. albicans, S. aureus, and S. mutans enhanced the release of eDNA, which may be caused by autolysis or cell death in the polymicrobial biofilm, and it caused a high accumulation of eDNA. C. albicans eDNA release is associated with biofilm formation, and more eDNA concentration leads to a higher biofilm formation [32]. In the polymicrobial biofilm, the large amount of eDNA released by S. aureus, S. mutans, and C. albicans allows the construction of a safe scaffolding for the development of a robust biofilm (Figs. 2 and 3).

C. albicans eDNA protects the scaffolding of polymicrobial biofilm against the action of DNase I. The high release of C. albicans eDNA in polymicrobial biofilm protects the CFU of the scaffolding of biofilm of S. aureus and S. mutans against the enzymatic activity of DNase I. During polymicrobial biofilm formation, C. albicans, S. mutans and S. aureus contribute 96%, 3.5%, and 0.5% of eDNA respectively (Fig. 2). In a previous study, eDNA and biofilm matrix proteins are vital constituents of biofilms and may carry significant risk when coupled with antibiotic resistance [33]. This is the first investigation demonstrating by qPCR that the great release of eDNA by C. albicans protects the CFU formed by S. mutans and S. aureus against disintegration in the polymicrobial biofilm. C. albicans eDNA (eukaryotic) enables the polymicrobial biofilm formed by S. aureus and S. mutans (prokaryotes) to be resistant to the action of DNase I, C. albicans contributes 96% of eDNA and cushioning the effect of DNase I.

CLi drastically reduces the bacterial density from prokaryote biofilms. The action of CLi decreases the cellular density of S. aureus and S. mutans but not C. albicans in polymicrobial and monospecies biofilms and causing a decrease in the release of eDNA only in polymicrobial biofilm. CLi, a protein synthesis inhibitor used in the treatment of Gram-positive bacterial and fungi infections in combination with an antifungal, modulates the production of several toxins and virulence factors [34, 35]. The eDNA was detected to increase resistance against antibiotics in many pathogenic bacteria [36]. Our results indicate that inhibitory concentrations of CLi (1 µg/µl) reduce cell density, causing the decrease of eDNA release from polymicrobial biofilm with a minimal effect on C. albicans (Fig. 4).

A combined therapy with DNase I and CLi reduced the cellular density of S. aureus and S. mutans but not C. albicans in polymicrobial and monospecies biofilms. DNase I degrades eDNA from biofilms, but the effect on cell density is different. The polymicrobial biofilm shows a slight decrease in the cell density, which is non-significant when compared from those not treated with DNase I; however, the monospecies has a strong decrease in cell density (Fig. 5). The action of CLi decreases the cellular density of S. aureus and S. mutans but not C. albicans polymicrobial and monospecies biofilms and decreases the release of eDNA. DNase I degrades the eDNA and disaggregates the biofilm, enhancing the action of CLi in the polymicrobial and monospecies biofilm. Agents that disrupt the biofilm matrix, such as DNase I, enhance the antibiofilm activity of antibiotics, so that the biofilm cells become susceptible to antibiotic treatment [9].

Biofilm eDNA is secreted in various ways, and different microorganisms have different modes of releasing eDNA, eDNA aids in the structural integrity of the biofilm matrix, as wells as bacterial adhesion, metal chelation, metabolic vitality, antibiotic resistance, and horizontal gene transfer. eDNA may serve as an effective anti-biofilm target. Research has focused on DNase treatments ever since DNase I was used to degrade eDNA and disrupt biofilms [3, 5]. By applying nucleases directly into biofilms, eDNA is degraded and the biofilm matrix is weakened; DNase I could inhibit formation and disrupt the bioflm matrix, allowing the antimicrobial substance to target the detached cells. These findings suggest that DNase I in combination with CLi may be a better alternative approach against biofilm-associated pathogens to disperse biofilm and enhance bactericidal efficiency.

Ultrastructural analysis of biofilms and post-treatment with DNase I and CLi. S. aureus, S. mutans, and C. albicans develop a robust oral biofilm stained with Hoechst that stains intracellular DNA (iDNA) present in the living cells that constitute the biofilm. eDNA is present in oral biofilms and represents a potential target for biofilm control and exclusion of potential pathogens [37]. In polymicrobial biofilm, the eDNA concentration stained with propidium iodide is higher than the monospecies; the mean fluorescence intensity for iDNA was about 3.5 times that of eDNA from S. aureus monospecies biofilm, 2.44 times from S. mutans monospecies biofilm, 1.64 times from C. albicans monospecies biofilm and 1.28 times from polymicrobial biofilm; C. albicans monospecies biofilm and polymicrobial biofilm release a lot of eDNA (Table 1). C. albicans release more eDNA essential for the scaffolding of bacteria in the polymicrobial biofilm [38].

Destruction of eDNA with DNase I causes the rupture of the monospecies biofilm ultrastructure but in the polymicrobial biofilm the break is slight (Fig. 6). In monomicrobial biofilms the proportion of iDNA that degrades is equal or greater than eDNA and in polymicrobial biofilms the proportion of iDNA that degrades is lower than eDNA (Table 1). iDNA from polymicrobial biofilm is more resistant to action of DNase I than monospecies biofilm because the high concentration of eDNA from polymicrobial maintaining the biofilm scaffolding and dumpins the action of DNase I (Fig. 6n1 and n2).

CLi destroys the cell density of S. aureus and S. mutans but not C. albicans from monospecies and polymicrobial biofilm. CLi caused significant disruption to the bacterial monospecies biofilm structure, resulting in a more dispersed and accesible structure (Fig. 6c1 and g1); but Cli does not disperse the polymicrobial biofilm (Fig. 6o1); in polymicrobial biofilm C. albicans mediated enhanced S. aureus and S. mutans biofilm structure. Previous work determined that C. albicans increased the tolerance of Streptococcus gordonii to clindamycin [39].

Treatment with DNase I plus CLi reduced the fluorescence intensity of the eDNA and iDNA of the monospecies and polymicrobial biofilms (Fig. 6), but the polymicrobial and C. albicans biofilms were more refractory to DNase I and CLi treatment compared with than S. aureus and S. mutans monospecies biofilms (Table 1).

In conclusions, C. albicans releases a high amount of eDNA in polymicrobial biofilm, which protects the scaffolding of S. aureus and S. mutans against the enzymatic activity of DNase I and dampens the effect of DNaseI and CLi on prokaryotic cells. The eDNA forming the biofilm matrix is degraded by DNase I, and this enhances the action of CLi that causes a reduction of the biofilm ultrastructure of S. aureus and S. mutans but not C. albicans. The eDNA from C. albicans should be considered a possible target for modifying robust polymicrobial biofilms for successful treatment of biofilm-associated prokaryote infections with antibiotics.

Acknowledgements

The authors thank Mario Costa Cruz from the Research Facility Center (CEFAP) and Marcia Harumi for the excellent technical support.

Funding

Sao Paulo State Research Foundation Grant Number: 2017/07339–4. UFMG Support Foundation (FUNDEP) Grant Number: 30201*52.

Declarations

Conflict of Interest

The authors declare no potential conflicts of interest with respect to the authorship and/or publication of this article.

Footnotes

Publisher's Note

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

Contributor Information

Santiago M. Lattar, Email: santiagolattar@gmail.com, Email: slattar@ufmg.br

Gabriel Padilla, Email: gpadilla@icb.usp.br.

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