Method Article

Animal Model of Implant-Associated Infections in Mice

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DOI:

10.3791/68041

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June 27th, 2025

 ,  ,  ,  ,  ,  ,  , 
1Department of Orthopedics, Centre for Leading Medicine and Advanced Technologies of IHM, The First Affiliated Hospital of USTC, Division of Life Sciences and Medicine, University of Science and Technology of China

* These authors contributed equally

In This Article

Summary

The present protocol describes a mouse model employing subcutaneous dorsal implantation to investigate implant-associated infections, enabling comprehensive investigation of pathophysiological mechanisms and supporting the development of diagnostic criteria with targeted therapeutic strategies.

Abstract

Implant-associated infections represent a significant clinical challenge, as they arise when pathogenic microorganisms infiltrate the site of medical implants, such as joint replacements, catheters, or pacemakers. These infections can lead to severe complications, including implant failure, prolonged hospitalization, and the need for additional surgical interventions. To investigate these critical issues, murine models provide an invaluable platform for studying implant-associated infections and evaluating therapeutic strategies against bacterial biofilms, yielding insights that may ultimately improve clinical outcomes for patients afflicted by infected implants. The commonly employed materials in these studies include titanium, polyethylene, and stainless steel, which closely mimic those used in clinical practice. The experimental design generally involves a surgical procedure to implant the device, which is often performed subcutaneously to simulate joint or skin-related infections. Following implantation, a biofilm-forming bacterial strain, such as Staphylococcus aureus or Escherichia coli, is inoculated at the site to initiate infection, thereby establishing a model that mirrors the pathophysiology of human infections. Researchers have rigorously assessed the effectiveness of various antibiotic regimens against established biofilm infections, employing both systemic and localized delivery methods. Investigations have focused on novel antimicrobial agents, biofilm-disrupting compounds, and innovative therapeutic approaches aimed at augmenting the host immune response. Efficacy evaluation encompasses quantifying bacterial loads on the implant and surrounding tissue, alongside assessing reductions in biofilm thickness and density posttreatment. Clinical symptomatology, including changes in swelling and pain, serves as an additional indicator of therapeutic success. Furthermore, overall survival and morbidity rates among murine subjects provide critical insights into the safety and effectiveness of the evaluated treatments. It is imperative that all studies adhere to rigorous ethical guidelines and secure necessary institutional approvals, ensuring that findings can be reliably replicated across different laboratories. This methodological rigor is essential for validating results and advancing our understanding of effective treatments for implant-associated infections.

Introduction

Animal models of implant-associated infections are essential tools that bridge basic research and clinical applications1,2. These findings provide practical support for scientific research and facilitate the development and refinement of novel therapeutic and diagnostic approaches, ultimately improving patient outcomes and quality of life3,4,5,6. These models offer a platform for long-term observation, which aids in understanding the pathophysiological changes associated with implant-associated infections7,8. For example, by observing the effects of infection on surrounding tissues and bones, researchers can elucidate how infections lead to tissue damage, osteolysis, and other complications, thereby guiding improvements in clinical treatment strategies.

Successful animal model studies provide foundational data for clinical trials and help optimize therapeutic strategies9,10. Through these experiments, researchers can determine the most effective dosage, treatment duration, and administration route, which can inform clinical research11,12. Additionally, animal models enable the assessment of treatment limitations, allowing for necessary adjustments before transitioning to human trials. Animal models of implant-associated infections, also referred to as device-associated infections, are invaluable in studying the mechanisms, detection methods, and treatment approaches for these infections13,14. Implant infections often occur in fields such as orthopedics and dentistry, where devices such as joint replacements or fracture fixations are commonly used15,16,17,18. Owing to the presence of implants, infections are challenging to eradicate and often lead to chronic inflammation and tissue damage, potentially hindering patient recovery and necessitating implant removal for further treatment.

The purpose of establishing animal models is to simulate the occurrence and progression of implant-associated infections under controlled conditions, enabling in-depth exploration of pathophysiological processes, the development of novel diagnostic methods, and the evaluation of potential therapeutic strategies19,20,21,22,23. An ideal animal model should closely mimic the pathological features and immune responses observed in human infections, with consistent reproducibility in experiments24,25,26. Commonly used animals include rats, mice, and rabbits, each model being tailored to specific research goals, such as the choice of implant type, bacterial strains, and infection methods. With these models, researchers can monitor infection spread, immune responses, and pathogen colonization behaviors, including biofilm formation on the implant11,23,27. Furthermore, these models are instrumental in testing the efficacy of antimicrobial therapies, such as antibiotics, antimicrobial coatings, and photodynamic therapy, in controlling infection4,12,28. The application of new diagnostic methods and imaging technologies within animal models also enhances the potential for early diagnosis and effective management of implant-associated infections in clinical settings.

The long-term goal is to establish an animal model that accurately replicates the physical and physiological microenvironments of implant-associated infections in humans. Although this endeavor involves high complexity, substantial costs, and ongoing challenges, the development of a highly realistic animal model for implant-associated infections provides a critical early-stage testing platform for therapeutic agents and strategies. In this study, a mouse implant-associated infection model and a mouse subcutaneous infection model were constructed. By comparing relevant indicators, the degree of matching between the animal model and clinical periprosthetic infections was explored.

Protocol

All animal experimental procedures were approved by the Institutional Animal Care and Use Committee of the Institute of Health and Medicine, Hefei Comprehensive National Science Center (approval number: IHM-AP-2024-080). All mice used were male C57BL/6J mice of specific pathogen-free (SPF) grade, aged 6 weeks, with body weights of 20-30 g.The animals were housed at five mice per cage and were allowed free access to food and water. The environment was maintained at 22-26 °C with 50-60% relative humidity under a 12-h light/dark cycle (Light: 7:00 am-7:00 pm; Dark: 7:00 pm-7:00 am). Details of the reagents and equipment used in this study are listed in the Table of Materials.

1. Bacterial preparation for subcutaneous implant-associated infections

  1. Remove a loop (10 µL) of Staphylococcus aureus (S. aureus, ATCC 43300) culture from the -80 °C stock tube. Streak the suspension onto a blood agar plate using the streak plate technique. Incubate at 37 °C for 24 h under aerobic conditions without shaking.
    NOTE: Either three-sector or four-sector streak plate methods are permissible, provided the initial and terminal sectors remain non-confluent. Inter-sector streaking requires the use of a new inoculation loop. Parallel, equidistant streaks should be executed to facilitate uniform microbial distribution. Strict adherence to aseptic protocols is mandatory to prevent cross-contamination.
  2. Examine the morphology of typical S. aureus colonies and the presence of clear hemolytic zones.
    NOTE: Typical colony morphology includes golden-yellow pigmentation, a round shape, and a smooth surface. A clear zone of at least 2 mm in diameter is considered positive for β-hemolysis.
  3. Perform Gram staining and microbial mass spectrometry analysis to confirm S. aureus identity.
    NOTE: Strain identification should be conducted by investigators using established selection criteria.
  4. Select an independent typical colony (as described in step 1.2). Inoculate it into a 15 mL sterile centrifuge tube containing 5 mL of sterile tryptic soy broth (TSB). Incubate at 37 °C for 12 h with shaking at 200 rpm.
  5. Dilute the bacterial suspension from step 1.4 with TSB at a 1:50 ratio. Incubate at 37 °C with shaking at 200 rpm.
  6. Monitor and record the optical density (OD) at 600 nm using a spectrophotometer to identify the logarithmic growth phase.
  7. Establish a correlation between OD600 values and Colony Forming Units per milliliter (CFU/mL) using the spread plate method (refer to step 4.1.7-4.1.10).
    NOTE: In this experiment, the ideal OD600 is 0.700-0.750, corresponding to approximately 4 × 105 CFU/mL. Determining the relationship between OD600 and CFU/mL ensures experimental accuracy.
  8. Dilute 3 mL of bacterial suspension (from step 1.7, 4 × 105 CFU/mL) with 3 mL of cold sterile phosphate-buffered saline (PBS). Centrifuge at 3,000 × g for 10 min at 4 °C and resuspend the pellet. Final bacterial concentration: 2 × 105 CFU/mL.
    NOTE: The 1:1 ratio balances the effective dilution of culture medium impurities with the maintenance of cell integrity and osmotic stability. It ensures efficient washing while minimizing cell damage due to osmotic stress and promotes uniform cell pellet formation during centrifugation, which is critical for downstream processing.
  9. Repeat step 1.8 in duplicate.
  10. Resuspend the bacterial pellet in PBS for further experimentation.

2. Mouse and implant preparation

  1. Randomly divide 20 C57BL/6J wild-type mice into two groups (10 mice/group): the Implant-associated Infection group and the Subcutaneous Abscess group29.
    NOTE: Each group should be housed in two individually ventilated cages (IVCs), with five mice per cage.
  2. Apply tail markings to each mouse for identification.
    NOTE: Use ear tags for mice identification to prevent tail markings from fading.
  3. Sterilize circular titanium plates (TA1, 1 cm in diameter, 0.5 mm in thickness) by autoclaving at 121 °C, 0.1 MPa for 30 min.

3. Animal surgery

  1. Place the mice in an isoflurane (2%) gas induction chamber with general animal gas anesthetics for approximately 5 min to induce anesthesia (following institutionally approved protocols).
  2. Monitor respiration to ensure appropriate anesthetic depth.
    NOTE: Respiration should be regular and slower than when awake and should not change in response to noxious stimuli, such as surgical procedures or toe pinching.
  3. Maintain anesthesia with the nasal cone delivery system using continuous inhalation of 2% isoflurane and 2 L/min oxygen flow (Figure 1A).
    NOTE: Perform all procedures under anesthesia to prevent stress-induced struggling and ensure compliance with animal welfare regulations.
  4. Transfer the anesthetized mice to the preparation zone of the biosafety cabinet.
    NOTE: Divide the biosafety cabinet into three compartments: preparation, disinfection, and operation zone. Place sterile disposable mats within each compartment. Perform all procedures within the cabinet to maintain aseptic conditions.
  5. Shave the hair on the back (3 cm × 4 cm area) and apply depilatory cream for approximately 1 min.
  6. Disinfect the skin with iodine solution and 75% ethanol three times (Figure 1B).
  7. Make an approximately 1 cm incision in the dorsal region using sterile surgical blades. Insert the sterile titanium implants (prepared in step 2.3) into the subcutaneous pocket (Figure 1D).
    NOTE: Elevate the distal skin of the incision using sterile forceps to form the subcutaneous pocket.
  8. Inject 100 µL of S. aureus bacterial suspension (2 × 105 CFU/mL) directly onto the titanium surface using a 1 mL sterile syringe (Figure 1E).
  9. Disinfect the skin with 75% ethanol and apply 3-0 non-absorbable sutures to close the incision (Figure 1F).
    NOTE: Suture size selection (3-0, 4-0, 5-0, or 6-0) should correspond to the mice's body size.
  10. For the subcutaneous abscess group, create, disinfect, and suture the incision directly without inserting any implant.
  11. Inject 100 µL of S. aureus suspension (2 × 105 CFU/mL) via the incision using a sterile 1 mL syringe.
    NOTE: Continuous slow-rate injection was maintained throughout the administration to mitigate rapid fluid accumulation and prevent bacterial suspension leakage.
  12. Monitor respiratory function until spontaneous breathing resumes before transferring the animals to new IVCs.
  13. Administer meloxicam (0.02 mL/10 g/day) subcutaneously for three consecutive days to manage post-surgical pain.
  14. Ensure all mice can bear full weight on their limbs, move without restriction, and access food and water freely.
    NOTE: House all treated mice in new IVCs to prevent cross-contamination with untreated mice or old IVCs.

4. Postoperative observations and evaluation

  1. Evaluation of infections in peripheral tissue
    1. Observe the condition of the dorsal incision on each mouse daily and record observations on days 1, 3, 5, 7, 10, and 14.
      NOTE: Monitor for implant dislodgement; if detected, retrieve the implant promptly for subsequent scanning electron microscopy (SEM) analysis.
    2. Euthanize the mice via CO2 asphyxiation followed by cervical dislocation on days 3, 7, and 14. Collect the implants and peripheral tissue for further analysis (Figure 2).
    3. Shave the dorsal hair and disinfect the skin using iodophor and 75% ethanol three times before tissue collection.
      NOTE: Shaving the dorsal hair and disinfecting the wound surface minimizes contamination from surface bacteria.
    4. Excise the skin, subcutaneous tissue, and muscle within a 0.5 cm margin around the implant or wound using sterile surgical scissors.
      NOTE: Collect tissue samples without size restrictions to maximize yield. Weigh and homogenize each sample with an equal mass of sterile PBS to maintain consistent g/mL units. Divide each tissue sample longitudinally and equally into proximal and distal segments for bacterial quantification (proximal) and histopathological evaluation (distal).
    5. Place the harvested tissue into sterile tubes. Add an equal mass of sterile PBS and three sterile steel grinding beads to each tube.
    6. Homogenize the tissues in three cycles (70 Hz, 60 s grinding time, 20 s pause time), then vortex for 5 min.
    7. Perform serial dilutions of the homogenate (from step 4.1.6) with PBS from 101 to 105.
    8. Drop 15 µL of each diluted homogenate onto designated areas of blood agar plates.
    9. Incubate the plates at 37 °C without shaking for 24 h.
      NOTE: Divide each blood agar plate into six equal sectors and apply serial dilutions to their respective areas.
    10. Record bacterial colonies using a digital camera and count them manually.
  2. Observation of S. aureus on the implants
    1. On days 3, 7, and 14, harvest the implants using sterile scissors and tweezers.
    2. Immerse the implants in 2.5% glutaraldehyde at 4 °C for 48 h for fixation.
    3. Rinse the implants with sterile PBS three times, then dehydrate them using ethanol solutions (50%, 70%, 80%, 90%, and 100%) for 10 min each.
    4. Freeze-dry the implants in a freeze-dryer at -85 °C with vacuum pressure maintained under 20 Pa.
    5. Mount the implants on a sample stage using conductive tape and coat with a 10 nm layer of gold.
    6. Observe the samples under SEM at high vacuum and 1.5 kV. Adjust magnification to at least 2500×, as needed.
  3. Pathology analysis of the peripheral tissue
    1. On days 3, 7, and 14, collect peripheral tissue or major organs using sterile scissors after disinfecting the skin.
    2. Immerse the collected tissue in 4% paraformaldehyde at 4 °C for 48 h for fixation.
    3. Dehydrate the tissue by immersing it in graded ethanol solutions (50%, 70%, 80%, 90%, 100%) for 15 min each.
    4. Embed the dehydrated tissue in paraffin and section it into 4 µm slices using a microtome.
    5. Stain the sections with hematoxylin and eosin (H&E) or Giemsa using standard protocols30.
    6. Observe the stained slices using the tissue cell quantitative analysis system.

Results

A mouse subcutaneous implant-associated infection model was established via the methodology described in this study (Figure 1). All the mice were in good condition within 3 h after the surgical operation, while their diet, sleep, and activity returned to normal on the same day.

Postoperatively, we continued to observe the wound areas on the back of the two groups and found that the range of subcutaneous abscesses in the implant-associated infection group was greater than that in the subcutaneous abscess group until the end of the experiment (Figure 3). Compared with the subcutaneous abscess group, the implant-associated infection group developed rupture on day 3, which occurred 4 days earlier. On day 10, both groups of mice showed signs of wound recovery, with the subcutaneous abscess group showing more pronounced recovery on day 14. In contrast, the wound has not yet healed in the implant-associated infection group, in which some mice had shed titanium sheets within 14 days, indicating a more severe degree of infection.

To distinguish more visually the difference in the extent of infection between the two groups, we collected tissue from the infection site in each mouse. Tissue homogenate dilutions of the infection site from both groups were analyzed for bacterial load on blood agar plates (Figure 4A). Strikingly, the amount of infected tissue in the implant-associated infection group had no tendency to decrease on days 7 and 14, whereas the amount of infected tissue bacteria in the subcutaneous abscess group decreased successively on days 3, 7, and 14 (Figure 4B). SEM observations of titanium sheets removed from mice in the implant-associated infection group further confirmed the above findings, namely that there was a gradual increase in the bacterial counts on the titanium sheets in the mice over a period of 14 days (Figure 5). Giemsa staining also revealed that the bacterial counts of infected tissues in the subcutaneous abscess group were significantly lower on day 7 when compared with that on day 3 and further decreased on day 14, during which time there was no sign of a decrease in bacterial counts in the implant-associated infection group (Figure 6). Thus, the implant-associated infection model showed more severe and persistent infection than the subcutaneous abscess model under an equivalent infection dose.

Further analysis of the intensity of inflammatory cell infiltration in the infected tissues of the mice via H&E staining revealed that the subcutaneous abscess group exhibited a lower degree of inflammatory cellular infiltration on day 7 when compared with that on day 3 and a remarkable decrease on day 14 (Figure 7). Inflammatory cell infiltration in the implant-associated infection group continued to be high throughout the 14-day period. Finally, the histopathological analysis of the major organs in the implant-associated infection group revealed that all the major organs were essentially normal using our modeling method and that there was no obvious manifestation of lesions in the outcome compared with normal mice (Figure 8). These results suggested that our modeling method is highly safe and reliable and is suitable for the study of implant-associated infections.

Mouse surgical procedure: anesthesia, skin prep, incision, placement, infection, suture steps.
Figure 1: Protocol of the implant-associated infection model. (A) Anesthesia was maintained using an inhalation anesthesia system with nasal cone delivery; inset: induction anesthesia via a gas induction chamber. (B) Skin preparation was performed by shaving and disinfecting the surgical site. (C) A transverse dorsal incision was made using a sterile blade. (D) A sterile titanium implant was inserted subcutaneously through the incision. (E) Bacterial suspension (2 × 105 CFU/mL) was applied to the titanium surface to induce infection. (F) The incision site was sterilized and sutured. Please click here to view a larger version of this figure.

Implant-associated infections diagram; SEM analysis, bacterial quantification, histopathology.
Figure 2: Schematic of experimental grouping, sample collection, and analysis procedures. Please click here to view a larger version of this figure.

Mice healing study, subcutaneous vs. intraperitoneal injection, time-lapse results over 14 days.
Figure 3: Representative images of the incision sites in both models on days 1, 3, 5, 7, 10, and 14. Scale bar: 1 cm. Please click here to view a larger version of this figure.

Petri dish cultures, infection growth; chart of bacterial count over time; gradient dilution diagram.
Figure 4: Bacterial load analysis from peripheral tissue. (A) Representative images of bacterial colonies from spread plate cultures on days 3, 7, and 14. Top row: subcutaneous abscess model; bottom row: implant-associated infection model. (B) Quantitative changes in bacterial load over time in both groups. (C) Schematic of the serial dilution method used for bacterial quantification. Please click here to view a larger version of this figure.

Bacterial growth over time, implant infections, microscopy image, Days 3, 7, 14, medical research.
Figure 5: Scanning electron microscopy (SEM) images of the titanium implant surface on days 3, 7, and 14. Yellow spheres indicate S. aureus; the blue background represents the titanium surface. Scale bar: 20 µm. Please click here to view a larger version of this figure.

Histology of abscess and infection at days 3, 7, 14; tissue response to bacteria; microscopy analysis.
Figure 6: Giemsa staining of peripheral tissue on days 3, 7, and 14 in both models. Red arrows indicate bacterial colonies. Scale bar: 50 µm. Please click here to view a larger version of this figure.

Histological analysis of infection progression in subcutaneous abscess and implant sites over 14 days.
Figure 7: Hematoxylin and eosin (H&E) staining of peripheral tissue on days 3, 7, and 14 in both models. Scale bar: 50 µm. Please click here to view a larger version of this figure.

Histology comparison diagram; control vs. implant infection in heart, liver, spleen, lung, kidney.
Figure 8: H&E staining of major organs on day 14 in the implant-associated infection model and healthy control mice. Scale bar: 50 µm. Please click here to view a larger version of this figure.

Discussion

As with all animal models, intrinsic immunological divergences between murine and human responses31 necessitate cautious interpretation of findings regarding clinical translation. The implant infection model represents a controlled simulation that does not fully recapitulate the complexity of clinical scenarios. Furthermore, clinical biomaterial heterogeneity (e.g., composition variance) may significantly modulate bacterial pathogenesis, a parameter standardized in this experimental setting32.

While the murine model provides practical advantages for preliminary investigations, its translational limitations must be acknowledged. The incorporation of aged (18-month-old) or genetically obese animals would better recapitulate the metabolic and immunological conditions of patients most vulnerable to implant-associated infections33,34. Additionally, controlled mechanical loading paradigms could bridge the gap between static implant models and the dynamic joint environment35.

The current subcutaneous implant-associated infections model, while demonstrating utility in studying early biofilm formation, presents several technical challenges that warrant discussion. Regarding infection rate variability, the observed spectrum from spontaneous clearance to uncontrolled dissemination likely reflects heterogeneity in host immune responses. To address this, immunosuppressive pretreatment (e.g., dexamethasone or cyclophosphamide) could better simulate the compromised immunity characteristic of typical implant-associated infection patients36, particularly those with diabetes or advanced age. Furthermore, the implementation of local ischemic conditions during surgery through brief vascular occlusion may promote more consistent bacterial colonization by creating a transient hypoxic microenvironment favorable for microbial adherence.

Technical challenges in biofilm visualization, particularly the disruption of delicate extracellular matrices during conventional SEM processing, necessitate methodological refinements. The adoption of critical point drying techniques represents a superior alternative to standard freeze-drying for structural preservation. Complementary approaches such as confocal microscopy with fluorescent lectin staining enable three-dimensional biofilm characterization without the artifacts introduced by dehydration protocols37.

The advantages of assessing implant-associated infection are the simplicity of the procedure, the high success rate, and the broad applicability. Comparatively speaking, the method of subcutaneous MRSA infection constructed by Ching-Wen Tseng et al. is simpler but not applicable to implant-associated infection models38. The mouse periprosthetic joint infection models constructed by39, and the spinal implant-associated infection model constructed by Benjamin V. Kelley et al. are closer to real clinical situations40. However, their implant insertion procedure requires more practice for a novice researcher. This modeling method is highly safe and reliable, enables comprehensive investigation of pathophysiological mechanisms, and supports the development of diagnostic criteria with targeted therapeutic strategies.

Looking forward, this model system offers exciting opportunities for technological innovation. The integration of bioluminescent bacterial tracking (via lux-tagged strains) would enable longitudinal monitoring of infection progression in living animals41. The platform is particularly well-suited for evaluating novel antimicrobial strategies, including antibiotic-functionalized implant coatings. Genetic manipulation approaches, such as targeted knockout of immune signaling pathways (e.g., MyD88 deficiency), could yield important insights into host determinants of biofilm persistence. These methodological advancements promise to enhance both the reliability of preclinical implant-associated infection models and their predictive value for clinical translation.

Disclosures

The authors declare that they have no competing financial interests.

Acknowledgements

The authors would like to acknowledge the financial support from the National Natural Science Foundation of China (82572779 to Z.S., 82102586 to Z.S., 82272512 to C.Z., 82472463 to C.Z.), Anhui Provincial Health Research Program (AHWJ2024Aa20445 to Z.S., AHWJ2024Aa10144 to C.Z.), the Natural Science Foundation of Anhui Province, Distinguishing Youth Project (2108085J40 to C.Z.), the Anhui Provincial Key Research and Development Program-Clinical Medical Research Translation Specialization (202304295107020020 to C.Z.), the China Postdoctoral Science Foundation (2020M671893 to Z.S.), the Fundamental Research Funds for the Central Universities of China (WK9110000155 to Z.S., YD9110002053 to C.Z.), the Anhui Provincial Scientific Research Compilation Project (2022AH051251 to Z.S., 2022AH010076 to C.Z.), and the Research Funds of Centre for Leading Medicine and Advanced Technologies of IHM (2023IHM01070 to C.Z., 2023IHM01074 to C.Z.). We are also thankful for support from the Core Facility Center for Medical Sciences, The First Affiliated Hospital of USTC (Anhui Provincial Hospital). This work was partially carried out at the Instruments Center for Physical Science, University of Science and Technology of China. We are also thankful for the support of the Biosafety Level-2 Laboratory from the Core Facility, Institute of Health and Medicine, Hefei Comprehensive National Science Center.

Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1% osmium tetroxide solutionAny make
2.5% glutaraldehyde solutionAny make
4% paraformaldehyde solutionAny make
75% ethanolAny make
Automated Gram Staining SystemBIOMERIEUXPrevi Color Gram
Biosafety CabinetThermo Fisher Scientific1500
Blood agar plateSHIGOUYIKLCP002
Depilatory creamVeet
Freeze drierEYELAFDU-2110
IsofluraneRingpuIsoflurane
Low-Temperature High-Speed CentrifugeSCILOGEXCF1524R
Microbial Mass Spectrometry Analysis BIOMERIEUXVITEK MS
Non-Absorbable Sutures with needlesShanghai JinhuanF308
Paraffi sectioning systemLeica
Phosphate buffered saline (PBS)BiosharpBL302A
Scanning electron microscopeHITACHISU8010
Small Animal Anesthesia MachineRWDR500
SpectrophotometerJINGHUA721
Tert-butanolAny make
TG Panoramic Tissue Cell Quantitative Analysis SystemTissueGnosticsTISSUEFAXS PLUS
Tissue homogenizerIKAT10 basic ULTRA-TURRAX
Tryptic soy broth (TSB)SolarbioLA0110
Vertical single-door oscillation incubatorSakeziEsci-1020

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