A subscription to JoVE is required to view this content. Sign in or start your free trial.

Method Article

Oral Biofilm Formation on Different Materials for Dental Implants

11.4K views

⸱

DOI:

10.3791/57756

⸱

June 24th, 2018

 ,  ,  ,  ,  , 
1Department of Dental Materials and Prosthodontics, School of Dentistry of Ribeirão Preto, University of São Paulo, 2Department of Restorative Dentistry, School of Dentistry of Ribeirão Preto, University of São Paulo

In This Article

Summary

Here, we present a protocol to evaluate oral biofilm formation on titanium and zirconia materials for dental prosthesis abutments, including the analysis of bacterial cells viability and morphological characteristics. An in situ model associated with powerful microscopy techniques is used for the oral biofilm analysis.

Abstract

Dental implants and their prosthetic components are prone to bacterial colonization and biofilm formation. The use of materials that provides low microbial adhesion may reduce the prevalence and progression of peri-implant diseases. In view of the oral environment complexity and oral biofilm heterogeneity, microscopy techniques are needed that can enable a biofilm analysis of the surfaces of teeth and dental materials. This article describes a series of protocols implemented for comparing oral biofilm formation on titanium and ceramic materials for prosthetic abutments, as well as the methods involved in oral biofilms analyses at the morphological and cellular levels. The in situ model to evaluate oral biofilm formation on titanium and zirconia materials for dental prosthesis abutments as described in this study provides a satisfactory preservation of the 48 h biofilm, thereby demonstrating methodological adequacy. Multiphoton microscopy allows the analysis of an area representative of the biofilm formed on the test materials. In addition, the use of fluorophores and the processing of the images using multiphoton microscopy allows the analysis of the bacterial viability in a very heterogeneous population of microorganisms. The preparation of biological specimens for electron microscopy promotes the structural preservation of biofilm, images with good resolution, and no artifacts.

Introduction

Bacterial biofilms are complex, functionally and structurally organized microbial communities, characterized by a diversity of microbial species that synthesize an extracellular, biologically active polymer matrix1,2. The bacterial adhesion to biotic or abiotic surfaces is preceded by a formation of the acquired pellicle, mainly consisting of salivary glycoproteins1,3,4. Weak physicochemical interactions between the microorganisms and the pellicle are initially established and followed by stronger interactions between bacterial adhesins and glycoprotein receptors of the acquired pellicle. Microbial diversity gradually increases through the coaggregation of secondary colonizers to the receptors of the already attached bacteria, forming a multispecies community1,3,4,5.

Homeostasis of the oral microbiota and its symbiotic relationship with the host is important in maintaining oral health. The dysbiosis within oral biofilms may increase the risk for the development of caries and periodontal disease2,5. Clinical studies demonstrate a cause-and-effect relationship between the accumulation of biofilm on teeth or dental implants and the development of gingivitis or peri-implant mucositis6,7. The progression of the inflammatory process leads to peri-implantitis and the consequent loss of the implant8.

Dental implants and their prosthetic components are prone to bacterial colonization and biofilm formation9. The use of materials with a chemical composition and surface topography that provides low microbial adhesion may reduce the prevalence and progression of peri-implant diseases9,10. Titanium is the most-used material for the manufacture of prosthetic abutments for implants; however, ceramic materials were recently introduced and are gaining popularity as an alternative to titanium because of their aesthetic properties and biocompatibility11,12. Also importantly, ceramic materials have been associated with a supposedly reduced potential to adhere to microorganisms, mainly due to their surface roughness, wettability, and surface free energy10,13.

In vitro studies have contributed to significant advances in the understanding of microbial adhesion to prosthetic abutment surfaces9,14,15,16,17. However, the dynamic environment of the oral cavity, characterized by its varying temperature and pH and nutrient availability, as well as by the presence of shear forces, is not reproducible in in vitro experimental protocols18,19. To overcome this problem, an alternative is the use of in situ models of biofilm formation, which advantageously preserves its three-dimensional structure for ex vivo analysis10,20,21,22,23,24.

The analysis of the complex structure of the biofilm formed on oral substrates requires the use of microscopy techniques capable of displaying optically dense matter25. Multiphoton laser scanning microscopy is a modern option for biofilm structural analysis26. It is characterized by the use of nonlinear optics with an illumination source close to the infrared wavelength, pulsed to femtoseconds27. This method is indicated for the image acquisition of autofluorescence materials or materials marked by fluorophores, in addition to images generated by non-linear optical signals derived from a phenomenon known as Second Harmonic Generation. Among the advantages of multiphoton microscopy is the great image depth obtained with minimum cell damage caused by the intensity of the excitation light27.

For a viability analysis of biofilm on abiotic surfaces by multiphoton microscopy, the use of fluorescent nucleic acid dyes with different spectral characteristics and a penetration capacity in bacterial cells is required28. Fluorophores SYTO9 (green-fluorescent) and propidium iodide (red-fluorescent) can be used for a visual differentiation between live and dead bacteria28,29,30. Propidium iodide penetrates only bacteria with damaged membranes, while SYTO9 enters bacterial cells with an intact and compromised membrane. When both dyes are present inside a cell, propidium iodide has a greater affinity for nucleic acids and displaces SYTO9, marking it red28,30.

In view of the oral environment complexity and oral biofilm heterogeneity, microscopy techniques are needed that can enable the biofilm analysis of the surfaces of teeth and dental materials. This article describes a series of protocols implemented for comparing oral biofilm formation on titanium and ceramic materials for prosthetic abutments, as well as the methods involved in oral biofilms analyses at the morphological and cellular levels.

Access restricted. Please log in or start a trial to view this content.

Protocol

This study was approved by the Institutional Review Board of the School of Dentistry of Ribeirão Preto, and the volunteer participant signed the written consent (Process 2011.1.371.583).

1. Biofilm Formation in Situ

  1. Selection of participants
    1. Select patients based on the following inclusion criteria: a healthy individual with a complete dentition and no clinical signs of oral diseases.
    2. Exclude patients based on the following exclusion criteria: pregnancy, lactation, dental caries, a periodontal disease or antibiotic treatment in the last 3 months, smoker, or any systemic disease that could influence the periodontal status.
  2. Preparation of intraoral device
    1. Record the maxillary arch by means of an alginate impression.
    2. Prepare type IV stone by adding 19 mL of water to 100 g of stone powder. Pour the stone into the alginate mold to make a model of the maxillary arch.
    3. Design and fabricate an acrylic intraoral device to support the test specimens (titanium and ceramic disks).
      1. Fabricate retentive clasps from NiCr wire (0.7 mm in diameter) using orthodontic pliers #139 and position them on the model. To position the clamp between the upper premolars, bend one end of each wire so that they form a loop.
        1. Adapt the loop in the region of the interdental papilla. In the occlusal, insert a gentle curvature so as not to interfere with the points of contact and with the occlusion. Make a 90° fold at the end of the clamp for a retention in the acrylic.
        2. To fit the clamp in the upper second molar, adapt the curvature of the wire in the cervical third (vestibular) to contour the tooth (distal) and make a 90° fold at the end of the clamp for a retention in the acrylic.
          NOTE: In order to provide adequate retention/stability to the intraoral device, the retentive clasps were positioned between the upper premolars and distal aspect of the second molar on both sides of the dental arch.
      2. Manipulate the self-curing acrylic resin according to the manufacturer's instructions and press the acrylic resin between 2 glass plates (with a 3 mm thick spacer interposed) during the plastic phase, to make a 3 mm thick sheet.
      3. Lay the acrylic sheet on the palatal region of the model, according to the device's design, and trim off the surplus acrylic resin with a carver, before polymerization.
      4. Fabricate wax disks using a metallic matrix [10 mm in diameter, 2 mm in thickness (surface area of 78.5 mm2)] by placing molten wax in the matrix and waiting for the wax to solidify. Manually embed 4 wax disks into the acrylic resin, 2 of them between the premolars and the other 2 next to the second molars.
      5. Let the acrylic resin polymerize in a pressure pot under 20 psi of compressed air for 20 min.
      6. Finish the intraoral device with an electric dental lab handpiece and cutter and polish it with abrasive rubber.
      7. Adjust the device for a proper fit in the mouth using the equipment described above.
  3. Preparation of the titanium and zirconia specimens
    Note: Specimens (n = 14) are 10 mm in diameter and 2 mm in thickness and have a surface area of 78.5 mm2.
    1. Polish the specimens' surfaces with water-cooled sandpapers of decreasing abrasiveness (600, 1200, and 2000 grit), for approximately 20 min.
      NOTE: The polishing of the specimens was performed to standardize the surface roughness at 0.2 µm.
    2. Clean and disinfect the specimens and intraoral device with liquid detergent and tap water, and then use an isopropyl alcohol ultrasonic bath for 15 min. Dry them with absorbent paper towels.
    3. Fix the specimens in the intraoral oral device with about 0.1 mL of non-toxic hot melt adhesive (Figure 1).
    4. Install the device containing the specimens in the oral cavity.
      NOTE: The intraoral device containing the specimens should be worn for 48 h. The device was removed and stored in phosphate buffered saline (PBS) while the patient was eating and performing oral cleaning.

2. Assessment of Bacterial Viability

Note: The sample size n = 10.

  1. Prepare a dyeing solution by adding 3 µL of SYTO9 (green fluorescent nucleic acid dye) and 3 µL of propidium iodide (red fluorescent nucleic acid dye) to 1 mL of sterile distilled water.
    NOTE: Prepare solutions protected from light.
  2. Transfer the specimens to a 24-well plate and wash them thoroughly with PBS to remove any non-adherent cells.
  3. Add the appropriate volume (1 mL) of fluorescent dye solution to cover the biofilm-containing specimen. Add the dye very carefully so as not to disorganize the biofilm.
  4. Incubate the specimen for 20 - 30 min at room temperature, protected from light.
  5. Gently wash the biofilm sample with sterile distilled water to remove any excess dye.
  6. Place the specimen in a glass bottom dish and perform multiphoton laser scanning microscopy for the biofilm analysis.
    NOTE: The analysis of the bacterial cells viability was carried out using a multiphoton microscopy system. The fluorescence of propidium iodide was detected using a filter with an excitation/emission wavelength of 546/680 nm and 477/600 nm for SYTO9. The size of the images obtained was 5.16188 x 5.16188 mm, which corresponds to 26.64 mm2 of the total area of each specimen (78.5 mm2) or 33.94% of the total area. The images were made from the most central portion of the sample, with a resolution of 1,024 x 1,024 pixels. The red channel and green channel images were analyzed separately using Fiji software31. Each cell was selected using a selection tool and the intensity of the fluorescence was measured through the integrated density of the pixels, subtracting the image background.

3. Analysis of the Specimens' Chemical Composition by Energy Dispersive Spectroscopy (EDS)

Note: The sample size n = 3.

  1. Select 3 specimens of each biofilm-free test material, and assess the chemical composition in 2 different areas of each specimen using a scanning electron microscope coupled to a dispersive energy spectrometer, with an electron beam voltage of 10 kV32.

4. Morphological Analysis of the Bacterial Biofilm by Scanning Electron Microscopy

Note: The sample size n = 1.

  1. Fix the biofilm by immersing the specimens in 2.5% glutaraldehyde diluted in 0.1 M sodium cacodylate buffer, pH 7.0 - 7.3, for 24 h.
  2. Wash the sample in PBS buffer (pH = 7.6).
  3. Postfix the biofilm with 1% osmium tetroxide for 1 h.
  4. Wash the sample in PBS buffer (pH = 7.6).
  5. Dehydrate the biofilm samples carefully, keeping them immersed in increasing concentrations of ethanol solutions (50%, 70%, 90%, 95%, and 100%).
    NOTE: Perform 3 exchanges of ethanol at a 100% concentration. The total dehydration step takes about 2 h.
  6. Transfer the specimens to the critical point dryer and make several substitutions with carbon dioxide (CO2) until the specimens are dry.
  7. Remove the dried specimens from the apparatus and mount them onto the scanning electron microscope holders.
  8. Sputter-coat a 20 nm layer of gold onto the specimen's surface for 120 s.
  9. Insert the gold-coated discs in the scanning electron microscope chamber. Obtain images from 5 different areas of each specimen, at 20 - 30 kV under variable pressure and at 650X magnification12.

Access restricted. Please log in or start a trial to view this content.

Results

The colonization density of the biofilm after 48 h of in situ growth was represented in this study by the proportion of the colonized area on the titanium and zirconia disks in relation to the total scanned area of the specimen using multiphoton microscopy (26.64 mm2). Figure 2 represents the bacterial colonization density on the surface of the 3 tested materials. A higher density of biofilm was observed on the surfaces of the cast and on ...

Access restricted. Please log in or start a trial to view this content.

Discussion

The protocol described in this study was developed to evaluate the biofilm formation on titanium and zirconia materials for prosthetic abutments, including the analysis of bacterial cell viability and morphological characteristics. In order to accomplish this, an in situ model of biofilm formation was designed, consisting of an intraoral device capable to accommodate samples of the test materials and keep them exposed to the dynamic oral environment for 48 h. The device was considered comfortable and easy to ins...

Access restricted. Please log in or start a trial to view this content.

Disclosures

The authors have nothing to disclose.

Acknowledgements

The authors thank José Augusto Maulin from Microscopy Multiuser Laboratory (School of Medicine of Ribeirão Preto) for his generous assistance with the EDS and SEM analyses and Hermano Teixeira Machado for his generous technical assistance in the video edition.

Access restricted. Please log in or start a trial to view this content.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Hydrogum 5Zhermack DentalC302070
Durone IVDentsply17130500002
NiCr wire Morelli55.01.070
JET auto polymerizing acrylicClássico
Dental wax Clássico
Pressure pot Essencedental
Sandpapers 600 gritNORTONT216
Sandpapers 1200 gritNORTONT401
Sandpapers 2000 gritNORTONT402
Metallographic Polishing MachineArotec
Isopropyl alcoholSIGMA-ALDRICHW292907
Hot melt adhesiveTECSILPAH M20017
Filmtracer LIVE/DEAD Biofilm Viability KitInvitrogenL10316
Pipette Tips, 10 µLKASVIK8-10  
Pipette Tips, 1,000 µLKASVIK8-1000B  
24-well plate KASVIK12-024
Glass Bottom DishThermo Scientific150680
AxioObserver inverted microscope ZEISS
Chameleon vision ii laserCoherent
Objective EC Plan-Neofluar 40x/1.30 Oil DICZEISS440452-9903-000
SDD sensors - X-Max 20mm²Oxford Instruments
Glutaraldehyde solutionSIGMA-ALDRICHG5882
Sodium cacodylate Buffer SIGMA-ALDRICH97068 
Osmium tetroxideSIGMA-ALDRICH201030
Na2HPO4SIGMA-ALDRICHS9638Used for preparation of phosphate buffered saline
KH2PO4SIGMA-ALDRICHP9791 
NaClMERK1.06404
KclSIGMA-ALDRICHP9333 
Ethanol absolute for analysis EMSUREMERK1.00983
CPD 030 Critical Point DryerBAL-TEC
JSM-6610 Series Scanning Electron MicroscopeJEOL
SCD 050 Sputter CoaterBAL-TEC

References

  1. Do, T., Devine, D., Marsh, P. D. Oral biofilms: molecular analysis, challenges, and future prospects in dental diagnostics. Clinical, Cosmetic and Investigational Dentistry. 5, 11-19 (2013).
  2. Samaranayake, L., Matsubara, V. H. Normal Oral Flora and the Oral Ecosystem. Dental Clinics of North America. 61 (2), 199-215 (2017).
  3. Larsen, T., Fiehn, N. E. Dental biofilm infections - an update. Acta Pathologica, Microbiologica, et Immunologica Scandinavica. 125 (4), 376-384 (2017).
  4. Marsh, P. D., Do, T., Beighton, D., Devine, D. A. Influence of saliva on the oral microbiota. Periodontology 2000. 70 (1), 80-92 (2016).
  5. Marsh, P. D., Zaura, E. Dental biofilm: ecological interactions in health and disease. Journal of Clinical Periodontology. 44 Suppl 18, S12-S22 (2017).
  6. Zitzmann, N. U., Berglundh, T., Marinello, C. P., Lindhe, J. Experimental peri-implant mucositis in man. Journal of Clinical Periodontology. 28 (6), 517-523 (2001).
  7. Meyer, S., et al. Experimental mucositis and experimental gingivitis in persons aged 70 or over. Clinical and biological responses. Clinical Oral Implants Research. 28 (8), 1005-1012 (2017).
  8. Salvi, G. E., Cosgarea, R., Sculean, A. Prevalence and Mechanisms of Peri-implant Diseases. Journal of Dental Research. 96 (1), 31-37 (2017).
  9. Hahnel, S., Wieser, A., Lang, R., Rosentritt, M. Biofilm formation on the surface of modern implant abutment materials. Clinical Oral Implants Research. 26 (11), 1297-1301 (2015).
  10. Nascimento, C., et al. Bacterial adhesion on the titanium and zirconia abutment surfaces. Clinical Oral Implants Research. 25 (3), 337-343 (2014).
  11. Nakamura, K., Kanno, T., Milleding, P., Ortengren, U. Zirconia as a dental implant abutment material: a systematic review. The International Journal of Prosthodontics. 23 (4), 299-309 (2010).
  12. Scarano, A., Piattelli, M., Caputi, S., Favero, G. A., Piattelli, A. Bacterial adhesion on commercially pure titanium and zirconium oxide disks: an in vivo human study. Journal of Periodontology. 75 (2), 292-296 (2004).
  13. Nascimento, C., et al. Microbiome of titanium and zirconia dental implants abutments. Dental Materials. 32 (1), 93-101 (2016).
  14. Rimondini, L., Cerroni, L., Carrassi, A., Torricelli, P. Bacterial colonization of zirconia ceramic surfaces: an in vitro and in vivo study. The International Journal of Oral & Maxillofacial Implants. 17 (6), 793-798 (2002).
  15. de Avila, E. D., Avila-Campos, M. J., Vergani, C. E., Spolidorio, D. M., Mollo Fde, A. Jr Structural and quantitative analysis of a mature anaerobic biofilm on different implant abutment surfaces. Journal of Prosthetic Dentistry. 115 (4), 428-436 (2016).
  16. de Avila, E. D., et al. Impact of Physical Chemical Characteristics of Abutment Implant Surfaces on Bacteria Adhesion. Journal of Oral Implantology. 42 (2), 153-158 (2016).
  17. de Avila, E. D., et al. Effect of titanium and zirconia dental implant abutments on a cultivable polymicrobial saliva community. Journal of Prosthetic Dentistry. 118 (4), 481-487 (2017).
  18. Lin, N. J. Biofilm over teeth and restorations: What do we need to know? Dental Materials. 33 (6), 667-680 (2017).
  19. Prada-Lopez, I., Quintas, V., Tomas, I. The intraoral device of overlaid disk-holding splints as a new in situ oral biofilm model. Journal of Clinical and Experimental Dentistry. 7 (1), e126-e132 (2015).
  20. Prada-Lopez, I., Quintas, V., Vilaboa, C., Suarez-Quintanilla, D., Tomas, I. Devices for in situ Development of Non-disturbed Oral Biofilm. A Systematic Review. Frontiers in Microbiology. 7, 1055(2016).
  21. Burgers, R., et al. In vivo and in vitro biofilm formation on two different titanium implant surfaces. Clinical Oral Implants Research. 21 (2), 156-164 (2010).
  22. do Nascimento, C., et al. Oral biofilm formation on the titanium and zirconia substrates. Microscopy Research and Technique. 76 (2), 126-132 (2013).
  23. Al-Ahmad, A., et al. In vivo study of the initial bacterial adhesion on different implant materials. Archives of Oral Biology. 58 (9), 1139-1147 (2013).
  24. Al-Ahmad, A., et al. Bacterial adhesion and biofilm formation on yttria-stabilized, tetragonal zirconia and titanium oral implant materials with low surface roughness - an in situ study. Journal of Medical Microbiology. 65 (7), 596-604 (2016).
  25. Thomsen, H., et al. Delivery of cyclodextrin polymers to bacterial biofilms - An exploratory study using rhodamine labelled cyclodextrins and multiphoton microscopy. International Journal of Pharmaceutics. 531 (2), 650-657 (2017).
  26. Lakins, M. A., Marrison, J. L., O'Toole, P. J., van der Woude, M. W. Exploiting advances in imaging technology to study biofilms by applying multiphoton laser scanning microscopy as an imaging and manipulation tool. Journal of Microscopy. 235 (2), 128-137 (2009).
  27. Zipfel, W. R., Williams, R. M., Webb, W. W. Nonlinear magic: multiphoton microscopy in the biosciences. Nature Biotechnology. 21 (11), 1369-1377 (2003).
  28. Stocks, S. M. Mechanism and use of the commercially available viability stain, BacLight. Cytometry Part A. 61 (2), 189-195 (2004).
  29. Johnson, M. B., Criss, A. K. Fluorescence microscopy methods for determining the viability of bacteria in association with mammalian cells. Journal of Visualized Experiments. (79), e50729(2013).
  30. Stiefel, P., Schmidt-Emrich, S., Maniura-Weber, K., Ren, Q. Critical aspects of using bacterial cell viability assays with the fluorophores SYTO9 and propidium iodide. BMC Microbiology. 15, 36(2015).
  31. Schindelin, J., et al. Fiji: an open-source platform for biological-image analysis. Nature Methods. 9 (7), 676-682 (2012).
  32. Placko, H. E., Mishra, S., Weimer, J. J., Lucas, L. C. Surface characterization of titanium-based implant materials. The International Journal of Oral & Maxillofacial Implants. 15 (3), 355-363 (2000).
  33. So, P. T., Dong, C. Y., Masters, B. R., Berland, K. M. Two-photon excitation fluorescence microscopy. Annual Review of Biomedical Engineering. 2, 399-429 (2000).
  34. Benninger, R. K., Piston, D. W. Two-photon excitation microscopy for the study of living cells and tissues. Current Protocols in Cell Biology. , Chapter 4, Unit 4.11 11-24 (2013).
  35. Gardi, J. E., Nyengaard, J. R., Gundersen, H. J. The proportionator: unbiased stereological estimation using biased automatic image analysis and non-uniform probability proportional to size sampling. Computers in Biology and Medicine. 38 (3), 313-328 (2008).
  36. Melvin, N. R., Poda, D., Sutherland, R. J. A simple and efficient alternative to implementing systematic random sampling in stereological designs without a motorized microscope stage. Journal of Microscopy. 228 (Pt 1), 103-106 (2007).
  37. Neu, T. R., Kuhlicke, U., Lawrence, J. R. Assessment of fluorochromes for two-photon laser scanning microscopy of biofilms. Applied and Environmental Microbiology. 68 (2), 901-909 (2002).
  38. Neu, T. R., Woelfl, S., Lawrence, J. R. Three-dimensional differentiation of photo-autotrophic biofilm constituents by multi-channel laser scanning microscopy (single-photon and two-photon excitation). Journal of Microbiological Methods. 56 (2), 161-172 (2004).
  39. Neu, T. R., Lawrence, J. R. Innovative techniques, sensors, and approaches for imaging biofilms at different scales. Trends in Microbiology. 23 (4), 233-242 (2015).
  40. Lacroix-Gueu, P., Briandet, R., Leveque-Fort, S., Bellon-Fontaine, M. N., Fontaine-Aupart, M. P. In situ measurements of viral particles diffusion inside mucoid biofilms. Comptes Rendus Biologies. 328 (12), 1065-1072 (2005).
  41. Briandet, R., et al. Fluorescence correlation spectroscopy to study diffusion and reaction of bacteriophages inside biofilms. Applied and Environmental Microbiology. 74 (7), 2135-2143 (2008).
  42. Berney, M., Hammes, F., Bosshard, F., Weilenmann, H. U., Egli, T. Assessment and interpretation of bacterial viability by using the LIVE/DEAD BacLight Kit in combination with flow cytometry. Applied and Environmental Microbiology. 73 (10), 3283-3290 (2007).
  43. Bergmans, L., Moisiadis, P., Van Meerbeek, B., Quirynen, M., Lambrechts, P. Microscopic observation of bacteria: review highlighting the use of environmental SEM. International Endodontic Journal. 38 (11), 775-788 (2005).
  44. Hannig, C., Follo, M., Hellwig, E., Al-Ahmad, A. Visualization of adherent micro-organisms using different techniques. Journal of Medical Microbiology. 59 (Pt 1), 1-7 (2010).
  45. Knutton, S. Electron microscopical methods in adhesion. Methods in Enzymology. 253, 145-158 (1995).
  46. Fischer, E. R., Hansen, B. T., Nair, V., Hoyt, F. H., Dorward, D. W. Scanning electron microscopy. Current Protocols in Microbiology. , Chapter 2, Unit 2B.2 (2012).

Access restricted. Please log in or start a trial to view this content.

Reprints and Permissions

Tags

Titanium MaterialsZirconia MaterialsMultiphoton MicroscopyScanning Electron MicroscopyBiofilm AnalysisBacterial ViabilitySpecimen PreparationIn Situ Model
Oral Biofilm Formation on Different Materials for Dental Implants
Contact Us
Recommend to Library

Privacy

Terms of Use

Policies

Copyright © 2026 MyJoVE Corporation. All rights reserved.