Method Article

Multimodal Approach to Assess Bone Regeneration and Scaffold Performance

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DOI:

10.3791/69483

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February 13th, 2026

 ,  ,  ,  ,  ,  , 
1Institute of Biomedical Sciences, Federal University of Rio de Janeiro, 2Department of Condensed Matter, Applied Physics and Nanoscience, Brazilian Center for Research in Physics, 3Alberto Luiz Coimbra Institute for Graduate Studies and Research in Engineering (COPPE), Federal University of Rio de Janeiro, 4Brazilian Synchrotron Light Laboratory (LNLS), Brazilian Center for Research in Energy and Materials (CNPEM)

In This Article

Summary

We propose a protocol for assessing bone regeneration at various hierarchical levels. This multimodal approach allows for structural analysis across different length scales, effectively addressing challenges in sample preparation. It also provides a reliable workflow for evaluating the effectiveness of biomaterials in promoting bone regeneration.

Abstract

The clinical success of an implant largely depends on its ability to promote bone repair and facilitate osteointegration. While significant advancements have been made in designing new scaffolds, a thorough evaluation of implant success at various bone length scales has not been sufficiently investigated. Bone is a hierarchically organized tissue, with an intricate structure that ranges from the organization of collagen fibrils and their mineralization at the nanometer and micrometer scales to the macroscopic arrangement of compact and cancellous bone. This hierarchical organization is crucial for ensuring bone function and mechanical performance. In this context, a comprehensive assessment of the effects of biomaterials on bone repair at multiple length scales is essential for evaluating the effectiveness and safety of implanted materials. However, preparing samples for integrated multimodal analyses poses a technical challenge. In this work, we present a protocol designed to evaluate bone regeneration across various hierarchical levels and analyze the interface between newly formed bone and the implanted scaffold. This approach provides a robust method for assessing the effectiveness of scaffolds in inducing bone repair. The combination of selected 3D and 2D imaging techniques is crucial for a proper evaluation of the complex mineralized tissue that forms during the bone healing process

Introduction

Bone is a complex, hierarchically organized tissue1,2. Understanding its structural organization is essential for addressing fundamental questions related to how cells contribute to tissue architecture and how structural changes impact mechanical function3. In this context, bone remodeling after injury remains a central topic of investigation. Fracture healing is a complex multistage process, beginning with an initial inflammatory response, followed by angiogenesis and the differentiation of progenitor cells into chondrocytes and osteoblasts4,5. Much has been debated regarding the role of biomaterials to stimulate osteointegration, particularly about biocompatibility and their ability to prevent fibrous encapsulation6.

A commonly adopted method to assess biomaterial-stimulated osteointegration is optical microscopy, which enables both quantitative and qualitative histological analyses. This approach has been shown to be a reliable and effective way to evaluate the success of bone implants7,8. Nevertheless, several issues related to bone hierarchical organization cannot be explored by using a single imaging approach3, particularly the histological one, which is limited at least in part by the magnification restriction of the method.

Advances in three-dimensional imaging have proven essential for understanding bone hierarchical organization. Techniques such as microcomputed tomography (microCT; either synchrotron radiation or conventional laboratory-based X-ray sources)9,10,11, dual-beam scanning electron microscopy (FIB-SEM) associated with slice-and-view tomography12,13,14 and transmission electron tomography15,16 provided new insights into bone architecture at multiple scales.

Based on different 3D imaging approaches, a model of bone hierarchical organization has been proposed, which initially identified nine distinct levels of bone structural organization1. This scheme has recently been revised and expanded to encompass approximately 12 hierarchical levels, serving as a valuable reference for exploring the relationship between structure and function in bone tissue2,17. Importantly, this proposed hierarchical model provides a foundation for evaluating the success of bone repair following injury. Evaluating whether the regenerated bone tissue reproduces the native hierarchical structure is a key standard in assessing the quality of bone healing, particularly when biomaterials are used within the damaged site.

Although the use of multiple imaging techniques to investigate bone organization has been discussed6,7,18,19,20, little has been explored regarding a workflow that integrates different imaging modalities to assess bone mineral organization across multiple spatial scales. As mentioned, bone is a complex composite tissue, consisting of an organic phase (cells, collagen, and macromolecules) and a mineralized matrix. From a sample preparation perspective, this complexity presents technical challenges regarding solvent permeation through the sample, sectioning of a hard specimen, and the integration of different imaging strategies on the same sample.

The key innovation in this work is the establishment of a reproducible, correlative, and multiscale imaging workflow using three-dimensional imaging and complementary techniques. As illustrated in Figure 1A, the workflow is designed so that the same sample can be sequentially examined using different imaging modalities. Because some of these techniques are destructive or require sectioning, the workflow follows an order that begins with non-destructive methods and progresses to approaches that involve cutting or material removal.

This sample preparation and analysis workflow is broadly applicable to hard or mineralized specimens, including cortical and trabecular bone, pre-implantation scaffolds, and the evaluation of the bone/biomaterials interface after implantation. By combining imaging modalities that span different fields of view and resolution ranges (Figure 1B), the workflow enables the analysis of samples from the centimeter scale down to the micrometer and nanometer scales. While the sequence is optimized to support correlative analysis of a single specimen, each technique can also be applied independently according to the experimental goal. This approach enables a detailed assessment of biomaterials' osteointegration and the evaluation of the regenerated bone growth in the healing area.

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Protocol

The results presented in this protocol were obtained from an experimental procedure conducted in a previous study21. All procedures were carried out in accordance with the ethical guidelines of the Ethical Committee on the Use and Care of Experimental Animals at Fluminense Federal University, Brazil (CEUA/UFF: 779). Briefly, 20 male Wistar rats weighing between 300 and 400 g were randomly assigned to two groups: Sham and cHA-microsphere implantation. All animals underwent surgery in which a 2 mm defect was created in the tibial diaphysis. The Sham group served as a clot control to evaluate bone regeneration in a non-critical defect, whereas the experimental group received CHA37 microspheres implanted into the defect site. After 7 and 21 days, animals were euthanized with an overdose of general anesthesia, and tibial bone blocks from each group and time point (7 and 21 days) were collected. Bone quality and the extent of newly formed bone were assessed using optical microscopy and microCT.

1. Sample fixation and resin embedding

  1. Immediately after euthanasia, remove the tibia containing the implanted biomaterial and transfer it to a vial containing Karnovsky fixative solution (2.5% glutaraldehyde, 4% formaldehyde in 0.1 M cacodylate buffer). Store the sample at -4 °C for at least 1 day.
    NOTE: Prepare and handle the fixative under a fume hood to prevent exposure to volatile substances. All chemical handling must be performed using appropriate personal protective equipment (PPE), and additional technical safety measures should be implemented whenever required.
  2. Transfer the sample from the fixative to a vial containing 0.1 M cacodylate buffer. Keep the sample under gentle rotation, changing the buffer 2x at 10-minute intervals. At this stage, trim the sample edges using a small saw to facilitate resin infiltration. When cutting the sample, keep it wet in the 0.1 M cacodylate buffer to prevent it from drying.
    NOTE: Always keep the sample in a buffer volume that is at least 2x the sample volume. This should be followed for all the solutions and resin used in this protocol. When handling the chemical components mentioned above, work inside a fume hood and wear appropriate PPE. Collect the fixative solution and cacodylate buffer waste in clearly labeled, chemically compatible containers with properly fitted caps. Follow the institutional Standard Operating Procedures (SOPs) for the safe storage and disposal of hazardous chemical waste.
  3. Begin the dehydration process using increasing concentrations of acetone (10%, 30%, 50%, 70%, 90% and 100%). Keep the sample under rotation for 30 min at each concentration. Repeat the 100% anhydrous acetone step 3x.
    NOTE: When handling acetone, work inside a fume hood and wear appropriate PPE. After using it, collect any residual solvent in clearly labeled, chemically compatible waste containers with properly fitted caps. Follow the institutional SOPs for the safe storage and disposal of hazardous chemical waste.
  4. Infiltrate with spur resin by transferring the sample through a series of graded acetone-to-resin ratios: 2:1, 1:1, 1:2, and ultimately pure resin. Each step should last 12 h and be conducted under rotation at room temperature. After graded acetone-to-resin infiltration, change the pure resin 2x. Finally, transfer the sample to a mold containing pure resin and polymerize the sample at 70 °C for 24 h.
    NOTE: The resin was prepared according to the data sheet to achieve medium hardness. When handling spur resin, work inside a fume hood and wear appropriate PPE. After use, collect any residual in clearly labeled, chemically compatible waste containers with properly fitted caps. Follow the institutional SOPs for the safe storage and disposal of hazardous chemical waste.

2. Sample visualization using microcomputed X-ray tomography (microCT)

NOTE: microCT is used to assess implant positioning and bone regeneration. It also helps identify potential regions of interest for further analysis. This step should be performed with the sample embedded in resin.

  1. Utilize a sample holder provided by the manufacturer as a base, ensuring that the sample is positioned securely to prevent movement during the scan. Use low-density materials for support, such as polystyrene, gauze, or microcentrifuge tubes.
  2. Visualize using the following optimal scanning parameters: tube voltage/current = 60 kV/133 mA; angular step = 0.33°; averaging frames = 5; voxel size = 5 µm; rotation mode = 180°. For samples containing metallic implants, use a 360 ° rotation and a 0.5 mm aluminum filter.
    NOTE: During bone regeneration, the thickness of trabecular-like woven bone is typically within the resolution limits of conventional microCT imaging. To accurately quantify conventional morphometric parameters of trabecular bone, such as trabecular thickness, separation, and number, it is recommended to use the smallest voxel size (5 µm as specified). If the Region of Interest (ROI) exceeds the Field of View (FOV) at this nominal resolution, first use the multiscan feature to tile the ROI. Consider increasing the voxel size only as a last resort if the ROI cannot be captured via multiscan.
  3. After scanning, the raw acquisition data consists of a series of numbered image files. Perform reconstruction using the software provided by the manufacturer, with a focus on minimizing artifacts in the region of interest of the sample.
    NOTE: It is common to encounter image artifacts along the borders of the images, particularly in areas that will not be analyzed. The final volume can be cropped using the manufacturer's software to eliminate these regions.
  4. Analyze the reconstructed volume using various 3D visualization software options, such as Avizo, VGStudio Max, CTVox, and Dragonfly22. Apply digital image filters to remove artifacts. In this case, use the Non-local Means Denoising method for this purpose19.
    NOTE: Non-local Means Denoising filter smoothing values should be kept as low as possible, not to hinder the interface's important details.

3. Sample preparation for optical microscopy

  1. Select the sample sectioning direction according to the experimental objective. The tibia can be sectioned either transversely or longitudinally along the long axis of the bone. Use transverse sectioning to produce serial sections that cover the entire defect volume. Longitudinal sectioning is appropriate to evaluate bone regeneration along the long-axis direction. However, this approach provides only a single section of the defect.
    1. Use a low-speed diamond saw disk to section the sample into 1 mm slices, following the experimental strategy.
      NOTE: It is recommended to use a diamond disk with a thickness of 0.006 to 0.012 inches. Thinner blades allow for highly accurate cuts, with minimal material loss due to blade thickness.
    2. Fill the liquid reservoir until the fluid level covers the bottom edge of the blade.
      ​NOTE: Distilled water is suitable for bone samples. However, if excessive heating is anticipated, such as with samples containing metal implants, use the oil provided by the equipment manufacturer to fill the reservoir.
    3. Secure the sample to the specimen holder and attach it to the instrument arm. Ensure that the sample is stable and horizontal to the surface of the disk. Use the positioning knobs to adjust the specimen cut region to the disk.
    4. Begin cutting the sample using higher speed settings (210 - 250 RPM) and apply low pressure until an indentation forms on the sample. Then, reduce the speed (to 93 - 124 RPM) to complete the sectioning smoothly and prevent excessive vibration.
    5. Stop the saw immediately after the cut is complete. Adjust the system properly to ensure cutting ceases as soon as the sample is detached from its original block.
  2. Examine the sample using a stereomicroscope. Begin at low magnification (e.g., 5x-10x) to inspect the entire specimen, then increase the magnification (e.g., 20x-40x) to select the region of interest according to the scientific question. The exact magnification range may vary depending on the instrument used. In this work, regions exhibiting a higher density of interfaces between newly formed bone and the biomaterial were selected for further analysis.
  3. Sample mounting for thinning
    1. Mount the sample on a support to facilitate proper handling during polishing. If the sample is intended solely for optical microscopy analysis, attach it directly to a plastic coverslip using superglue on the previously polished side.
      ​NOTE: Plastic slides are recommended due to their improved adhesion. Glass slides may also be used (after extensive washing with neutral soap). However, glass slides increase the risk of sample detachment during polishing.
    2. For samples that will be further analyzed using a correlative approach, mount on a two-part aluminum tube assembly. In this protocol, a homemade aluminum tube assembly was developed, consisting of an inner solid cylinder with a diameter of 2.5 cm and an outer hollow cylinder that provides mechanical support during polishing. The sample is attached to the inner solid cylinder.
    3. Place the inner aluminum cylinder on a hot plate and heat it to a temperature suitable for melting the wax (typically around 60-70 °C, depending on the type of wax used).
      ​NOTE: Beeswax is also suitable. When handling mounting wax, work inside a fume hood and wear appropriate PPE. After use, collect the residual in clearly labeled, chemically compatible waste containers with properly fitted caps. Follow the institutional SOPs for the safe storage and disposal of hazardous chemical waste.
    4. Apply a small amount of wax onto the surface of the cylinder and allow it to melt completely.
    5. Once melted, place the sample with the surface of interest down onto the wax-coated surface and gently press it to ensure complete contact.
      ​NOTE: The surface of interest should face the cylinder surface so that thinning is performed on the opposite side. Avoid using excessive wax, as this may lead to uneven mounting and affect polishing quality.
    6. Remove the cylinder from the hot plate and allow the wax to solidify at room temperature.
  4. Monitoring sample thickness during polishing
    1. To monitor the sample thickness during the polishing process, use reflection microscopy at low to intermediate magnification (e.g., 10x-20x). A diagram illustrating the microscope calibration and the sample thickness measurement is shown in Figure 2.
    2. Calibrate the microscope Z-axis with a standard coverslip, which is typically around 150 µm thick. Mark one side of the coverslip with a red permanent marker and an adjacent area on the opposite side with a blue marker. Mount the coverslip on a glass slide.
    3. Using the fine focus knob and the micrometer scale, focus first on the bottom surface of the coverslip and then on the top surface.
    4. Record the Z-displacement value. This value corresponds to the coverslip thickness (approximately 150 µm) and serves as a reference.
    5. While polishing the sample, periodically use this Z-axis calibration to estimate the sample thickness without removing it from the aluminum cylinder.
      ​NOTE: This technique allows for quick and non-destructive thickness estimation during polishing. Alternatively, the sample can be removed from the mount and measured with a digital caliper or micrometer.
  5. Thinning and polishing
    1. Begin the coarse and intermediate polishing using sandpaper with grit sizes of 1200, 2500, and 4100. The objective of this step is to reduce the sample thickness to approximately 0.15 mm while removing significant surface irregularities.
    2. Start with 1200 grit sandpaper, then progress to 2500 grit. Apply light pressure during this process. Periodically monitor the sample thickness using reflection microscopy while polishing.
    3. Once the desired thickness is achieved, polish both sides of the sample using 4100-grit sandpaper. Polishing can be performed manually or with a semi-automatic polishing machine set to low rotation speed. Clean the sample thoroughly with distilled water continuously during each polishing step.
      ​NOTE: Keeping the samples and sandpapers wet during polishing helps reduce heat and prevents surface contamination from the rougher sandpapers.
    4. For final polishing, apply a small amount of diamond paste (e.g., 6 µm or 3 µm) onto a felt polishing cloth mounted on a polishing disc. Select the diamond paste particle size according to the surface roughness of the sample.
      ​NOTE: A 6 µm paste is suitable for preliminary fine polishing, whereas a 3 µm paste is recommended for final mirror polishing. When possible, using both pastes sequentially yields the best results. Nevertheless, satisfactory polishing can still be achieved using only the 6 µm paste.
    5. Gently polish both sides of the sample with minimal pressure. Use compressed air or cotton to dry the sample's polished surface before microscopy investigations.
    6. To detach the sample from the cylinder, warm the wax using a hot plate and clean any residual wax off the sample using anhydrous acetone.
      ​NOTE: When handling mounting wax and anhydrous acetone, work inside a fume hood and wear appropriate PPE. After use, collect the residual in clearly labeled, chemically compatible waste containers with properly fitted caps. Follow the institutional SOPs for the safe storage and disposal of hazardous chemical waste.
    7. Mount the sample between a glass slide and coverslip using immersion oil as the mounting medium and evaluate collagen fiber orientation using polarized light microscopy. Start at low magnification (e.g., 5x-10x) to survey the sample and increase to intermediate magnification (e.g., 20x-40x) to identify potential regions for further exploration using electron microscopy.
      NOTE: To proceed with the correlative workflow and subsequent analysis of the sample by scanning electron microscopy, it is advisable not to seal the sample.

4. Sample preparation for scanning electron microscopy

  1. Carefully remove the sample from the glass slide. Gently slide the coverslip off the glass slide using a pipette tip. Use filter paper to remove the excess immersion oil. Wash the sample with neutral soap. Perform the washing inside a small container to avoid accidental sample loss.
    ​NOTE: If residual immersion oil is still present on the sample, gently clean the surface using a cotton swab lightly moistened with 70% ethanol.
  2. Leave the sample to air dry on a filter paper. Compressed air or cotton can also be used to dry the sample. Once fully dry, mount the sample on a stub using carbon tape. Sputter-coat the sample with a 10 nm layer of gold to prevent charging during imaging.
  3. Utilize the dual beam FIB-SEM microscope to explore the sample through various methods.
    1. In the SEM (scanning electron microscopy) mode, using either secondary electrons or backscattered electrons, evaluate the sample morphology, biomaterial, and the newly formed bone interface in regions previously selected by optical microscopy.
    2. In focused ion beam-scanning electron microscopy (FIB-SEM) mode, perform slice-and-view tomography.
  4. In SEM mode with the secondary electron detector, select areas that were previously analyzed using polarized light microscopy and SEM to proceed with slice-and-view tomography acquisition.
  5. Adjust the parameters for trench volume, pixel size, and slice thickness based on the features of interest. In our setup, slice-and-view tomography was conducted with a trench volume of 15 µm x 10 µm x 10 µm, a pixel size of 20 nm, and a slice thickness of 60 nm. It is recommended to use an isometric voxel size whenever possible for enhanced imaging quality across all three dimensions (XYZ).
    NOTE: These parameters may vary depending on the specific FIB-SEM system. Consult the equipment operator to determine the optimal conditions based on your analytical requirements and the capabilities of the system.

5. Alignment, filter processing, and segmentation of FIB-SEM slice-and-view tomography

  1. Slice-and-view tomography generates a series of images that are numbered sequentially. To proceed with alignment, convert the series into an MRC file, commonly referred to as a stack.
  2. Perform MRC file generation and image stack alignment using IMOD software.
    NOTE: Other software, such as Avizo and FIJI/ImageJ, also offer stack alignment plugins. For detailed guidance, refer to each software-specific documentation. Additionally, the work of Peddie et al23. offers an overview of the approaches adopted during stack alignment.
  3. To generate the MRC file, open the terminal or command prompt, navigate to the folder containing the image sequence, and execute the following command: tif2mrc imagerootfile*.tif stackname.mrc
    1. For images that are not in TIFF format, use raw2mrc instead. Add -g (e.g., tif2mrc -g) to convert 24-bit RGB images to 8-bit grayscale. This will substantially reduce the computation time.
  4. Launch Etomo from the terminal and select the option Align Serial Sections. Follow the sequence to align the stack: (1) Load the unaligned stack into a single frame. (2) Algin tab using Initial Auto Alignment. (3) Make a Stack tab using Make Aligned Stack, then open the aligned stack.
    NOTE: This process may take several minutes, depending on the stack size and the performance of the system.
  5. Once completed, review the result. The aligned stack file will be named stackname_ali.mrc.
    NOTE: This procedure uses the system default settings for stack alignment. However, parameters related to the initial alignment and final stack generation can be modified according to user preferences. For detailed guidance, refer to the IMOD documentation: https://bio3d.colorado.edu/imod/doc/serialalign.html.https://bio3d.colorado.edu/imod/doc/serialalign.html.
  6. Perform additional image filtering using the Avizo software. Open the aligned stack and sequentially apply the following filters: Fast Fourier transform (FFT) filter, available in the Sandbox Filter plugin (select the option of vertical stripe orientation, tolerance value: 2), Shading correction wizard (adjust mask selection to cover all the shadow area of the image, use normalizing factor of 100), Histogram equalization (default configuration), and Non-local means (adjust values to 5; 0.1; 10; 3).
    NOTE: Equivalent filters are available in other image analysis software. In FIJI/ImageJ, we recommend the following sequence: FFT filter, Subtract Background, Enhance Local Contrast (CLAHE), and Non-Local Means filter.
  7. Proceed to image segmentation and 3D rendering using the software. Different segmentation strategies are available, including manual segmentation and threshold-based segmentation. Additional tools, such as the Watershed algorithm, can help automate the process and reduce the time required for segmentation.

6. Sample preparation for transmission electron microscopy

  1. Prepare lamellae using focused ion beam scanning electron microscopy (FIB-SEM) for assessing the bone/biomaterial interface.
    NOTE: If the implanted biomaterial is not ceramic or metallic, as in this study, ultrathin sections suitable for transmission electron microscopy (TEM) can be prepared using an ultramicrotome. However, it is important to note that sectioning mineralized materials can significantly reduce the lifespan of the diamond knife.
  2. Post-stain the ultrathin sections for 20 min in an aqueous solution of 5% uranyl acetate.
    NOTE: When handling uranyl acetate, work inside a fume hood and wear appropriate PPE. After use, collect any residual solvent in clearly labeled, chemically compatible waste containers with properly fitted caps. Follow the institutional SOPs for the safe storage and disposal of hazardous chemical waste.
  3. Analyze the sections using TEM. For a comprehensive analysis, combine imaging with selected area electron diffraction (SAED), energy-dispersive X-ray spectroscopy (EDS), and electron tomography.

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Results

The initial analysis of resin-embedded samples is performed using microcomputed tomography (microCT). This non-destructive technique facilitates 3D visualization of the entire defect area, enabling both quantitative and qualitative analyses of the sample. Contrast in microCT imaging derives mostly from the density differences of the materials present in the sample. Figure 1 illustrates the microCT analysis of a non-critical defect in a rat tibia that was implanted with the CHA37 biomaterial....

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Discussion

Different imaging techniques have been employed to investigate bone architecture and assess the effectiveness of biomaterials in inducing osteointegration7,21,22,23,24. In this protocol, we established a workflow that incorporates X-ray microtomography, optical microscopy, and electron microscopy to analyze the mineralized bone tissue formed in a rat tibia def...

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Disclosures

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

This work was supported by grants to the authors from the following Brazilian agencies: Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq/Brazil) and Fundação Carlos Chagas Filho de Amparo à Pesquisa do Estado do Rio de Janeiro (FAPERJ). We thank LABNANO/CBPF and CENABIO/UFRJ for the electron microscopy facilities. Language refinement and grammar revision were performed using ChatGPT (OpenAI, GPT-5.1, 2025). All edits were reviewed and verified by the authors.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Acetone Merck67-64-1
Aluminium cylinderhome-maden/a
Avizo version 2022.1ThermoFishern/asoftware for image filtering, segmentation and 3D visualization
Cacodylic Acid, Sodium Salt, trihydrateTed Pella18851powder for Cacodylate Buffer. Prepare a stock solution of 0.2 M
Diamond Polishing Compound 3 µm sizeTed Pella895-8
Diamond Polishing Compound 6 µm sizeTed Pella895-9
Diamond Wheel x.006South Bay TechnologyDWH 3063
Diamond Wheel x.012South Bay TechnologyDWH 4122
Felt polishing clothTed Pella816-20
Flat Embedding CapsulesEMS70021
Glass cover slipKnitteln/a
Glass slideKnitteln/a
Glutaraldehyde, 25% EM gradeTed Pella18426/18427
Gold sputter K550XEMITECHn/a
ImageJNational Institutes of Health (NIH)software for image aligment, filtering, segmentation and 3D visualization
IMOD version 5.1University of Coloradosoftware for image aligment, segmentation and 3D visualization
Lapping e Polishing MachineSouth Bay Technologymodel 910
Low-Speed Diamond Wheel Saw South Bay Technologymodel 650
Low Viscosity Embedding KitEMS14300kit for SPURR resin. Prepare according to the Data sheet specifications 
Micro-CT Skyscan 1273 Brukern/a
Microscope Jeol 2100F 200kV Jeoln/a
Microscope Olympus BX51Olympusn/a
Microscope Tescan Lyra3Tescann/a
Microscope Zeiss AxioplanZeissn/aequipped with a first order lambda plate
Paraformaldehyde, EM GradeSigmaP6148powder for formaldehyde solution. Prepare a stock solution of 16%
Plastic slideExakt41500
QuickStick 135 Mounting WaxSouth Bay Technologyn/a
Sandpapers  grit sizes 1200n/an/a
Sandpapers  grit sizes 2500n/an/a
Sandpapers  grit sizes 4100n/an/a
Super GlueLoctiten/a
Water Soluble Coolant South Bay Technologyn/a
Whatmann grade 1 filter papersigmaWHA1001150

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Multimodal ImagingBone RepairBiomaterial EvaluationBone Biomaterial InterfacePolarized Light MicroscopyScanning Electron MicroscopyTransmission Electron MicroscopyEnergy Dispersive X Ray

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