1 Volume 23 2024 e246163 Original Research Braz J Oral Sci. 2024;23:e246163http://dx.doi.org/10.20396/bjos.v23i00.8676163 1 Faculdade de Odontologia de Piracicaba (FOP), Universidade Estadual de Campinas (UNICAMP), Piracicaba, SP, Brazil. 2 Institute of Oral Biology, Faculty of Dentistry, University of Oslo, Oslo, Norway. 3 São Carlos Institute of Physics, University of São Paulo, São Carlos, SP, Brazil. Corresponding author: Antonio Pedro Ricomini Filho Department of Biosciences, Faculdade de Odontologia de Piracicaba (FOP), Universidade Estadual de Campinas (UNICAMP) Avenida Limeira, nº 901, Areião, CEP 13414-903, Piracicaba, SP, Brazil. e-mail: ricomini@unicamp.br Editor: Dr. Altair A. Del Bel Cury Received: April 02, 2024 Accepted: June 04, 2024 Absence of inhibitory effects of two new glucanases on streptococcus mutans growth Mateus Xavier-Queiroz1,2 , Pedro Ricardo Vieira Hamann3 , Heidi Aarø Åmdal2, Igor Polikarpov3 , Fernanda Cristina Petersen2 , Antônio Pedro Ricomini-Filho1* Glucanohydrolases have shown promise in degrading exopolysaccharides in cariogenic biofilms, making them a potential strategy for biofilm control without disrupting the oral microbiota. However, their direct antimicrobial effects remain unclear. Aim: To determine the antimicrobial activity on S. mutans of two newly discovered glucanases characterized by our group, PmGH87 (mutanase) from Prevotella melaninogenica and CoGH66 (dextranase) from Capnocytophaga ochracea, using a commercial dextranase from Penicillum sp. as a control. Methods: Their effects on growth were assessed using a luciferase reporter system coupled with the promoter of the ldh gene in Streptococcus mutans. Results: Quantification of optical density and luminescence over a 10-hour growth period revealed that the commercial dextranase exhibited inhibitory effects on S. mutans growth. However, these effects were neutralized by heat treatment, suggesting the presence of a heat-sensitive contaminant or an additional antimicrobial property associated with the commercial dextranase from Penicillum sp. On the other hand, the purified mutanase and dextranase enzymes had no inhibitory effect on S. mutans growth. Conclusion: In conclusion, the absence of inhibitory effects on S. mutans growth by the newly discovered enzymes emphasizes their potential for biofilm control while preserving the delicate balance of the oral microbiota and preventing the emergence of resistance. Keywords: Streptococcus mutans. Dextranase. Anti-Infective agent. https://orcid.org/0000-0001-8292-1230 https://orcid.org/0000-0002-6654-0646 https://orcid.org/0000-0001-9496-4174 https://orcid.org/0000-0003-4649-247X https://orcid.org/0000-0002-6593-6040 2 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 Introduction Glucanohydrolases have demonstrated potential in degrading biofilms by specifi- cally targeting the polysaccharides within the extracellular matrix1. In the oral cavity, biofilms are constantly formed on tooth surfaces, and due to the frequent consump- tion of sucrose, large amounts of extracellular polysaccharides (EPS) are synthe- sized, mainly glucans, such as α(1→3)-linked (mutans) and α(1→6)-linked (dextrans) glucans2. EPS favor the adhesion and accumulation of biofilm, and if not removed, contribute to the development of dental caries. Therefore, the degradation of dex- trans and mutans by the enzymes α(1→3)- and α(1→6)-glucanases (mutanase and dextranase, respectively) have shown promise in degrading exopolysaccharides in oral biofilms3. Furthermore, these enzymes exhibit substrate-specificity, making them a poten- tial strategy for controlling biofilms without disrupting the oral microbiota4. This differentiates them from traditional antimicrobial therapies, which may harm the commensal microbiota and contribute to antimicrobial resistance. As an alternative approach, they have been explored in combination with conventional antimicrobials to investigate potential synergistic anti-biofilm effects4-6. By breaking down the EPS barrier, glucanases facilitate the penetration of antimicrobials deeper into biofilms to prevent bacterial regrowth. However, it is crucial to consider the possible impact of these enzymes on bacterial growth, as they may carry antimicrobial compounds from the microorganisms used for their isolation or other impurities that could inhibit bacterial growth, and thus result in anti-biofilm effects not solely related to the direct activity of glucanases in degrading EPS. Of note, this aspect of glucanase activity is often overlooked in studies investigating their efficacy as anti-biofilm agents4-9. Therefore, we investigated the antimicrobial activity of two newly discovered gluca- nases characterized by our group, PmGH87 (mutanase) from Prevotella melaninogen- ica and CoGH66 (dextranase) from Capnocytophaga ochracea, using a commercial dextranase from Penicillum sp. as a control. Previously, our group demonstrated the potential of such enzymes on S. mutans biofilm inhibition and degradation10-11. Here, we assessed the impact of the enzymes on the growth and metabolism of Strepto- coccus mutans cultures in real-time using both optical density and a luciferase ldh gene reporter, respectively. Material and Methods Expression and purification of cloned mutanase (PmGH87) and dextranase (CoGH66) The nucleotide sequences encoding the GH87 mutanase from Prevotella melanino- genica (PmGH87) (GenBank ID: WP_004358976.1) and the GH66 dextranase from Capnocytophaga ochracea (CoGH66) (GenBank ID: WP_128091406.1) were ampli- fied from the respective genomic DNA and cloned, as previously described by Cor- tez et al.10 (2023). Heterologous protein expression was performed in Luria-Bertani (LB) broth containing 50 μg/mL of kanamycin at 37°C and 180 rpm until OD 0.6 3 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 (600 nm) followed by induction with 1 mM IPTG at 18°C for approximately 16 h. Then, the cells were harvested by centrifugation and resuspended in 50 mM Tris-HCl (pH 8.0), 200 mM NaCl, 1 mM PMSF buffer and stored at 4 °C before purification. Purification of expressed mutanase (PmGH87) and dextranase (CoGH66) Enzyme purification was carried out following the protocol described by Camilo and Polikarpov12 (2014). Briefly, the resuspended cells were disrupted performing six cycles of sonication on ice bath using a Sonic Dismembrator Sonifier (Fisher Sci- entific, Hampton, USA) at 40% amplitude. The resulting lysate was then clarified by centrifugation at 20,200 x g at 4°C for 30 min to remove cell debris, and the result- ing supernatant was used for mutanase or dextranase purification. The superna- tant from each sample was loaded in a column containing Ni-NTA Superflow resin (Qiagen, Hilden, Germany), previously equilibrated with 50 mM Tris-HCl (pH 8.0) and 200 mM NaCl buffer. Elution was performed using an imidazole gradient. The result- ing eluted protein fractions were combined and concentrated using a 50 kDa molec- ular cut-off concentrator to reduce imidazole concentration to less than 20 mM. The resultant sample was incubated with recombinant TEV protease at 4°C for 16 h for 6xHis-thioredoxin tag removal from the target enzyme. Finally, a second Ni²+ affinity chromatographic step was used for separation of tag-free mutanase or dextranase from contaminants and TEV protease. The protein sample’s purity was confirmed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) under denaturing conditions. The purified enzymes were freeze-dried and, previously to start the biofilm assay, enzymes were resuspended Phosphate-Buffer Salive (PBS) solution 1 X, pH 8.0. The concentration was determined using Nanodrop 1000 spec- trophotometer (Thermo Scientific, Waltham, EUA), measuring absorbance at 280 nm and applying the calculated extinction coefficient (Ɛ = 165180 M−1 cm−1 for muta- nase and Ɛ = 122000 M−1 cm−1 for dextranase)12. Experimental treatments In addition to mutanase (PmGH87) from Prevotella melaninogenica and dextranase (CoGH66) from Capnocytophaga ochracea, both of which were expressed and puri- fied, a commercially available dextranase (α-(1→6) glucanase, EC3.2.1.11, from Peni- cillium sp.; Product number: D4668-1KU; Batch number: 0000136817) was evaluated as control. The three enzymes were also evaluated in their denatured form by heating them at 100°C for 15 min10. The antimicrobial chlorhexidine digluconate (CHX), com- monly used in mouthwash, was used as the positive control, while only pure CDM medium as the negative control. Bacterial growth and luciferase reporter bioassay Lactate dehydrogenase (ldh) gene is constitutive in streptococci and plays a cru- cial role in the ATP-generating pathway, providing information into both viability and metabolic status13. In this study, an isogenic mutant strain of Streptococcus mutans UA159 containing a luciferase reporter for ldh gene (designated as SM120) as described by Dornelas-Figueira et al.14 (2023), was used in the bioassay. Briefly, bacterial inoculum of S. mutans SM120 was prepared in Chemical Defined Medium 4 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 (CDM) containing 1% glucose15. Overnight culture of S. mutans SM120 reporter strain was adjusted to an OD 0.7 (600 nm). The enzymes, active or denatured, were prepared at a concentration of 0.125 mg/mL (w/v) in the bacterial inoculum. This enzyme concentration was chosen based on our previous study3, where the lower dosage to cause a significant S. mutans biofilm degradation was 0.125 mg/mL (w/v). The positive control was prepared with 0.12% chlorhexidine digluconate in bacterial inoculum, as well as a group with 0.12% chlorhexidine digluconate in pure medium. A negative control group consisting of the inoculum without any treatment and a blank containing only pure medium were also prepared. A group containing inoculum without added luciferin was also included as a control in the luminescence assay. Aliquots of 200 µL of the mixtures were added in 96-well flat bottom plates (Nunc Thermo Scientific), followed by the addition in each well of 10 μL of 1.0 mM luciferin solution (Synchem, Felsberg-Altenberg, Germany) at room temperature. The plates were incubated at 37°C, 10% CO2 for 10 h. The optical density (OD), as an indicative of bacterial growth, and relative light unit (RLU), as an indicative of bacterial metabolism, were measured every 30 min after a 10-seconds elliptical agitation in a microplate reader (Synergy HT; BioTek, Winooski, VT, USA). The data of CHX in inoculum was subtracted from that in the pure medium, as the antimicrobial produced a whitish color in the medium, which affected OD measurement. RLU and OD data were log transformed and the area under the curve (AUC) for OD or RLU x h were calculated. Three independent experiments were conducted in triplicate, with each experiment carried out on different days. Statistical Analyses The data were analyzed by comparing treatment groups to the positive control group (CDM + S. mutans for OD data, or CDM + S. mutans + luciferin for RLU data). Each time point for OD and RLU curves was analyzed by two-way Anova followed by Dunnett’s Test (α=5%), considering as factors treatments and time. The AUC (OD or RLU x h) was analyzed by one-way Anova, Dunnett’s Test (α=5%). Statistical analysis was conducted using GraphPad Prism 9, and a p-value < 0.05 was consid- ered statistically significant. Results Quantification of optical density (OD) and luminescence (RLU) over the 10-h incuba- tion period revealed that the commercial dextranase exhibited inhibitory effect on the growth of S. mutans. The commercial dextranase, chlorhexidine and negative control groups presented no growth quantified by OD or ldh activity by RLU when compared to the respective positive control groups after 5 h-growth (Fig 1.A) and 1 h-growth (Fig 2.A), respectively. However, this effect was not observed for the commercial dex- tranase submitted to the heat-denaturation procedure (Fig 1.A and 2.A). Additionally, cultures in the presence of the other enzymes (whether denatured or not) exhibited similar growth and metabolic patterns to their respective positive control groups over the incubation period (Fig 1.A and 2.A). 5 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 A 0.5 0.0 -0.5 -1.0 -1.5 Time (h) Lo g ab so rb an ce (O D 60 0 nm ) Mutanase Mutanase denatured Dextranase sig: CDM and Dextranase Sigma; ns: all other groups. Commercial dextranase Commercial dextranase denaturedDextranase denatured CDM CDM + S. mutans CDM + S. mutans CDM Mutanase Mutanase denatured Dextranase Dextranase denatured Commercial dextranase Commercial dextranase denatured Chlorhexidine 0.12% Chlorhexidine 0.12% B 0 2 4 6 8 10 AUC Log abs (600 nm)xh Figure 1. Optical density (OD) quantification (Log). A. S. mutans 10 h-growth curve (n=3; mean ± SD) according to the treatments. B. Area under the curve for OD x h. * means significant differences against CDM + S. mutans control group. The AUC data for both OD and RLU compile the results observed at each time point throughout the entire period of the experiment. The active commercial dextranase exhibited lower values for both growth (AUC-OD; p<0.05; Fig. 1B) and metabolism (AUC-RLU; p<0.05; Fig. 2B) of S. mutans compared to the positive control. However, the commercial heat-denatured dextranase did not differ significantly from the posi- tive control for both analyses (p > 0.05; Fig. 1.B and 2.B). Mutanase and dextranase, whether denatured or not, showed higher AUC values for OD (p<0.05) and RLU (p<0.05) compared to the positive control. 6 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 A 5 4 3 2 1 0 Time (h) Lo g RL U (R el at iv e Li gh t U ni t) Mutanase Mutanase denatured Dextranase sig: CDM, CDM + S. mutans, Commercial dextranase and Chlorhexidine 0.12%; ns: other groups. Commercial dextranase Commercial dextranase denaturedDextranase denatured CDM CDM + S. mutans CDM + S. mutans + Luciferin CDM + S. mutans + Luciferin CDM + S. mutans CDM Mutanase Mutanase denatured Dextranase Dextranase denatured Commercial dextranase Commercial dextranase denatured Chlorhexidine 0.12% Chlorhexidine 0.12% B 0 10 20 30 40 AUC Log RLU (Relative Light Unit)xh Figure 2. Relative light unit (RLU) quantification (Log). A. S. mutans 10 h-growth curve (n=3; mean ± SD) according to the treatments. B. Area under the curve for RLU x h. * means significant differences against CDM + S. mutans + luciferin control group. Discussion The great advantage of employing enzymes for biofilm control lies in their ability to target specific molecules, thereby preventing the formation or disruption of the biofilm structure, without directly interfering with microbial viability. In the context of oral bio- films, it is expected that mutanases target mutans, while dextranases target dextrans. Therefore, the use of such enzymes to control oral biofilms could offer a beneficial therapy without disrupting the biological balance in the oral microbiota through anti- microbial effects. The newly discovered and characterized mutanase from Prevotella melaninogenica and the dextranase from Capnocytophaga ochracea have been used 7 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 by our group to degrade S. mutans biofilms10-11. In this study, we confirmed that these enzymes do not exhibit antimicrobial effects on S. mutans cultures, highlighting their potential to degrade oral biofilms solely through specific hydrolytic action which was verified previously10-11. The lack of antimicrobial effect for the recombinant enzymes aligns with findings by Rikvold et al.16 (2024) that also evaluated the antimicrobial effects of mutanase, DNAse, and beta-glucanase. In this context, the observed reduction of viability in enzymatically treated biofilms4,6 may be attributed to the degradation of extracel- lular polymeric substances, which impairs bacterial aggregation during biofilm for- mation or detaches bacterial cells from already formed biofilms. Our study supports this hypothesis, as the recombinant enzymes did not affect the planktonic growth of S. mutans (Fig. 1 and 2), which has been considered one thousand more suscepti- ble to antimicrobials in its planktonic than in biofilms17,18. Differently, commercial dextranase exhibited a similar inhibition of bacterial growth as digluconate chlorhexidine (Fig. 1 and 2). However, when the commercial dextranase was heated-denatured, bacterial growth occurred similarly to the untreated group. Thus, it could be speculated a possible antimicrobial effect of commercial dextra- nase. However, the substrate-specificity action of such enzymes for the recombinant enzymes, as observed in our study and the reported by Rikvold et al.16 (2024), do not support this hypothesis. Additionally, S. mutans cultures were grown in a chemically defined medium (CDM) supplemented with glucose instead of sucrose, which cannot lead to the synthesis of exopolysaccharides such as dextran and mutan that com- prise the biofilm matrix. As a result, the antimicrobial effect could also not be due to EPS protection. Thus, the absence of enzymatic activity observed in this experiment could be attributed both to the lack of available substrate and the enzymes’ inability to exert antibacterial effects. A more plausible explanation would be contaminants in the lyophilized enzyme product that could affect the bacterial viability. Indeed, the certificate of analyses of the commercial dextranase does not offer details regarding the enzyme’s production or purification process. In fact, it states that the protein content is 20.85% of the whole product. However, it is unclear whether this percentage specifically refers to the dextranase, as the total protein content was quantified using Lowry’s method19. Additionally, fungal dextranases are known to be produced by induction methods3 where the enzyme is secreted extracellularly along with many others fungal products that may not be removed if enzymes are not prop- erly purified. Considering that the commercial dextranase was produced by Penicil- lium spp., it is possible to speculate that the presence of penicillin as a contaminant could have contributed to the observed antimicrobial effect. In contrast, in our study, we used a well standardized protocol for intracellular heterologous expression of the mutanase and dextranase, as well as for protein purification10-12. It is worth noting that the commercial dextranase used in our study has been widely utilized in studies as a test or control enzyme in biofilm assays4-5,8. Upon reviewing the literature, we found one study7 in which another commercial enzyme was employed to specifically test its effect on the growth of S. mutans in culture, with notable inhibitory results. While this may be a true effect, the purification grade of such commercial enzyme is not informed and may have an impact on its observed effects. 8 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 Overall, our study highlights the necessity of implementing a purification step after enzyme production to ensure the removal of any contaminants that may not only affect microbial viability but also pose toxicity risks to humans, particularly in the context of enzyme appli- cation in oral care products. Enzymes can also be a desirable strategy when antimicrobi- als are needed. Despite biofilms being resistant to antimicrobials17, enzymes may play a crucial role in degrading EPS, facilitating antimicrobial diffusion, and enhancing bacterial killing4-6. Thus, enzymes contaminated with antimicrobials may lead to an overestimated reduction in bacterial viability. Besides, the presence of antimicrobials in biofilms can result in secondary effects18 that may also impact other evaluated outcomes. Some limitations of our study should be highlighted. While the oral cavity hosts hun- dreds of different species20, our study focused only on S. mutans. However, S. mutans is a relevant species due to its ability to produce an EPS-rich matrix and its association with biofilms, which contribute to dental caries21-23. Since EPS is the target substrate for dextranases and mutanases, the use of this microorganism was appropriate. Regard- ing the impact on other bacterial species, Rikvold et al.16 (2024) did not observe any antimicrobial effects. Therefore, studies evaluating commercially available enzymes aimed at degrading components of the biofilm matrix must include bacterial viability controls, as contaminants within the contents may interfere with bacterial growth or metabolism. In addition, the inhibitory effects of the commercial dextranase are not clear and require further investigation. In conclusion, the absence of inhibitory effects on S. mutans growth by the newly discovered and characterized enzymes emphasizes their potential for biofilm control while preserving the delicate balance of the oral microbiota and mitigating the emer- gence of resistance. In addition, this study highlights the potential of biofilm reduction previously demonstrated by our group to be solely attributable to the enzymatic spe- cific action on the EPS substrate. Furthermore, it is crucial to further investigate the underlying mechanisms responsible for the observed inhibitory effects of the com- mercial dextranase, as it is commonly employed in research to evaluate anti-biofilm properties. Additionally, future research should explore the broader applications of the newly discovered enzymes to harness their full potential in oral health interventions. Declarations Acknowledgements The authors are thankful to the Institute of Oral Biology at the University of Oslo, the Institute of Physics of São Carlos at USP, and the Laboratory of Biochemistry at FOP-UNICAMP for providing the necessary facilities to conduct the analyses performed in this study. Mateus Xavier de Queiroz is a recipient of PhD and PrInt scholarships from Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES), grants 88887.800941/2023-00 and 88887.682625/2022-00. This study was supported by CAPES, grant 001; by Fundo de apoio ao ensino, pesquisa e extensão (FAEPEX), grant 2501/23; by Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP), grants 2020/15092-1, 2021/08780-1, and 2023/07897-8; by Conselho Nacional de Desenvolvi- mento Científico e Tecnológico (CNPq), grants 306852/2021-7 and 440180/2022-8; and by Research Council of Norway (RCN), grants 274867 and 322375. 9 Queiroz et al. Braz J Oral Sci. 2024;23:e246163 Conflict of Interest The authors have no conflict of interest to disclose. Data availability Datasets related to this article will be available to the corresponding author upon request. Author Contribution Mateus Xavier de Queiroz: study conception and design, experimental work, data acquisition, data acquisition, data analyses, writing and manuscript review. Pedro Ricardo Viera Hamann: experimental work, data acquisition. Heidi Aarø Åmdal: experimental work, data acquisition, data analyses. Igor Polikarpov: study conception and design, data analyses, writing and manuscript review, funding acquisition. Fernanda Cristina Petersen: study conception and design, data analyses, writing and manuscript review, funding acquisition. Antonio Pedro Ricomini Filho: study conception and design, data analyses, writing and manuscript review, funding acquisition. All authors reviewed the final version of the manuscript. References 1. Thallinger B, Prasetyo EN, Nyanhongo GS, Guebitz GM. Antimicrobial enzymes: an emerging strategy to fight microbes and microbial biofilms. Biotechnol J. 2013 Jan;8(1):97-109. doi: 10.1002/biot.201200313.  2. Bowen WH, Koo H. Biology of Streptococcus mutans-derived glucosyltransferases: role in extracellular matrix formation of cariogenic biofilms. Caries Res. 2011;45(1):69-86. doi: 10.1159/000324598.  3. Pleszczyńska M, Wiater A, Janczarek M, Szczodrak J. (1→3)-α-D-Glucan hydrolases in dental biofilm prevention and control: a review. 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