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Method Article

Surgical Bone Implantation Technique for Rat Tibia Models of Diabetes and Osteoporosis

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DOI:

10.3791/66591

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July 5th, 2024

 ,  ,  ,  ,  ,  ,  , 
1Faculty of Medicine and Health Sciences, University of Barcelona, 2Department of Oral Surgery, Davallmed Clinic. Porto, 3Biomaterials, Biomechanics and Tissue Engineering Group, Department of Materials Science and Engineering, Universitat Politècnica de Catalunya, 4Barcelona Research Centre in Multiscale Science and Engineering, Universitat Politècnica de Catalunya, 5Department of Periodontology, Davallmed Clinic. Porto, 6Department of Odontostomatology, University of Barcelona, 7Oral Health and Masticatory System Research Group at the Bellvitge Biomedical Research Institute (IDIBELL), 8Preclinical imaging platform, Vall d'Hebron Research Institute, Barcelona, 9Preclinical Therapeutics Core, Helen Diller Family Comprehensive Cancer Center, University of California, San Francisco, 10Animal facility, Vall d'Hebron Research Institute, Barcelona, 11UCSF Gnotobiotics Core Facility, University of California, San Francisco, 12Human Anatomy and Embryology Unit, Experimental Pathology and Therapeutics Department, Faculty of Medicine and Health Sciences, University of Barcelona

In This Article

Summary

The placement of implants in a rat model is an essential experimental procedure for clinical research. This study presents a comprehensive surgical protocol for implanting titanium implants into the tibia of rat models with diabetes and osteoporosis.

Abstract

The rat has long served as a valuable animal model in implant dentistry and orthopedics, particularly in studying the interactions between biomaterials and bone tissue. The rat's tibia is frequently chosen due to its easy surgical access through thin tissue layers (skin and muscle) and the flattened shape of its medial face, facilitating the surgical insertion of intraosseous devices. Additionally, this model enables the induction of specific diseases, mimicking various clinical conditions to assess biological responses to different implant conditions like geometry, surface texture, or biological cues. However, despite its robust cortical structure, certain intraosseous devices may require adaptations in design and size for successful implantation. Therefore, establishing standardized surgical methods for manipulating both soft and hard tissues in the implantation region is essential for ensuring proper implant or screw device placement, particularly in fields like implant dentistry and orthopedics. This study included eighty Sprague Dawley rats divided into two groups based on their respective diseases: Group 1 with osteoporosis and Group 2 with Type 2 Diabetes. Implantations were performed at 4 weeks and 12 weeks, with the same surgeon following a consistent surgical technique. A positive biological response was observed, indicating complete osseointegration of all implants placed. These results validate the success of the surgical protocol, which can be replicated for other studies and serve as a benchmark for the biomaterials community. Notably, osseointegration values remained stable at both 4 weeks and 12 weeks for both disease models, demonstrating a durable integration of the implant over time and emphasizing the establishment of an intimate bone connection as early as 4 weeks.

Introduction

The common choice of rats as experimental subjects is due to the fact that they are easy to breed and relatively inexpensive compared to larger animal models. The emergence of new procedures, such as the reliable reproduction of a disorder, e.g., osteoporosis or diabetes, makes this model especially useful for analyzing the potential use of treatments and/or the influence of the disease in the biological response to drugs and surgical devices or procedures1,2.

The rat's bone mass gain occurs mostly during the first 6 months of life, although some researchers believe that the long bone grows constantly for at least a year with a progressive increase in length1. With aging, there is a transition from modeling to remodeling, which does not occur in all cases equally throughout the bones2. Female Sprague Dawley rats grow more slowly than male rats and achieve a lower peak in weight than male rats1. Continuous bone elongation and varied bone remodeling dynamics in rats are factors that have to be taken into account when addressing human health issues; however, it has not yet been possible to find any experimental research that shows either lifelong rat bone development or the species' inability to remodel bone1. If the experimentation starts around 10 months of age, a margin of at least 1 mm from the growth plate of the tibia should be left intact due to this longitudinal bone growth, an issue to be considered in dental implant studies2. Hormones are also a key parameter in bone research since at 8 months of age, male rats were found to have 22% greater bone width and 33% greater breaking strength than females in the tibia3.

The reliable reproduction of a disorder is thus very important in orthopedics and implant dentistry since osseointegration of an orthopedic screw or a dental implant is a complex process that depends on numerous factors influencing the systemic response to the device implantation into the bone. Systemic disorders like osteoporosis and diabetes are known to affect the success rate in orthopedics and implant dentistry, so the reliable reproduction of those disorders in rat models can be applied to explore ways to overcome these limitations.

The rat tibia, due to the easy surgical access, moderate bone volume, and the flat shape on the medial plate, makes it suitable for surgical bone implantation experiments4,5, and it has been used in numerous research studies exploring the effects of implant surface on osseointegration4,5,6. A growing number of studies assess the effects on osseointegration of coatings and substances added to the implant surface in both healthy animals7 and in compromised animals affected by diabetes or osteoporosis8,9,10,11,12,13,14.

The number of implant devices placed in one rat's tibia is limited and can differ depending on the type of study. Depending on the number of implants or study conditions, the dimensions of the devices must be adapted to minimize surgical trauma. In studies with one implant, a nearly human-size implant can be placed (2.0 mm in diameter and 4 to 5 mm in length), and bi-cortical anchorage can be achieved6,7,15,16. The dimensions of the implants in multi-implant protocols should adopt an appropriate implant size (1.5 mm in diameter and 2.5 mm in length)4,17.

The present study aims to describe a standardized surgical protocol for titanium implant placement on the tibia of two rat models: the osteoporosis and the diabetes rat model. Moreover, this study permits testing the surgical protocol to assess different types of implant surface biofunctionalization and its effect on osseointegration.

A sample of 80 rats was divided into two groups. In group 1, 40 ovariectomized Sprague Dawley females and 5 sham animals were selected, with a mean weight of 484 g and a mean age of 12 weeks. Based on vendor recommendations (see Table of Materials), three months after neutering, the experiment started. This waiting period ensured the disappearance of sex hormones. Osteoporosis was confirmed at the time of surgery based on micro-computed tomography (micro-CT) bone analysis, which reflected an average of 20% bone loss compared to the sham group. Group 2 consisted of 40 BBDR (Bio Breeding Diabetes Resistant) genetically modified Sprague Dawley rats with type II diabetes. The mean weight was 730 g, and the average age was 12 weeks. Prior to surgery, the diabetic status was confirmed with three consecutive days of glucose measurements with results higher than 200 mg/dL. Glucose was measured with a glucometer in 6 h of fasting, and a blood drop was collected by tail puncture.

Grade 3 titanium implants measuring 2 mm in length and 1.8 mm in diameter were used. All implants were sterilized in clean room conditions, by being ultrasonically cleaned in cyclohexane (3 times for 2 min), acetone (once for 1 min), deionized water (3 times for 2 min), ethanol (3 times for 2 min), and acetone (3 times for 2 min) using an ultrasound bath (230 VAC, 50/60 Hz, 360 W). Then, the samples were dried with nitrogen gas, and a nitrogen beam at 0.5 bar was applied directly onto the samples. Prior to implantation, the implants were first soaked in deionized water and then immersed in 70% ethanol (v/v) for 10 min. After this, the implants were transferred to sterile microcentrifuge tubes, and kept under sterile conditions until the surgery.

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Protocol

All experimental procedures were conducted in accordance with the European Community Guidelines for the protection of animals used for scientific purposes (Directive 2010/63/EU) as implemented in Spanish law (Royal Decree 53/2013) and Generalitat de Catalunya regulations (Decree 214/97). Ethics approval for all animal procedures and handling was obtained from the Ethics Committee for Animal Experimentation of the Vall D'Hebron Institut de Recerca (registration number 72/18 CEEA). For the osteoporotic model, female Sprague Dawley rats with an average weight of 484 g and an average age of 12 weeks were utilized. As for the diabetic model, genetically modified female BBDR (Bio Breeding Diabetes Resistant) rats with an average weight of 730 g and an average age of 12 weeks were employed. All animals were sourced from a commercial supplier. The specific details of the animals, reagents, and equipment utilized in the study are listed in the Table of Materials.

1. Anaesthesia/pharmacology and preparation of animals

  1. Administer presurgical analgesia using buprenorphine at 0.05 mg/kg and meloxicam at 2 mg/kg through subcutaneous injections 10-15 min before starting the surgical procedure.
  2. Perform intra-surgical anesthesia with inhaled isoflurane: initiate at 5% in fresh air during induction and maintain at 3%. Induce anesthesia in a rat chamber and maintain the isoflurane supply with a conical nose adaptation during surgery.

2. Preparation for the surgery

  1. Measure the anesthetized animal's body temperature with a rectal probe and use an electronically controlled heating pad for thermal support throughout the surgical procedure. Eye ointment must be applied prior to surgery beginning to avoid corneal dryness.
    NOTE: If necessary, the ointment must be reapplied after checking eye dryness.
  2. Trim hair with an electric shaver and apply hair removal cream to eliminate any remaining fur.
  3. Obtain an aseptic surgical field by cleaning the knee skin in a pattern of iodine and 70% (v/v) ethanol using sterile swabs, starting from inside and moving outward the incision line without retracing. Perform a minimum of three consecutive cleaning rotations (iodine-ethanol-iodine).
  4. Isolate the operating field by positioning a sterile fenestrated surgical drape over the animal, exposing the leg through the central opening (Figure 1).

3. Surgery

  1. Surgical exposure
    1. Make a full-thickness skin incision of approximately 1 cm in length vertically along the proximal border of the anteromedial face of the tibia, in the metaphysis region to expose the bone (Figure 2).
    2. Stabilize the leg and pull the skin taut against the underlying bone while making the incision, ensuring a clean incision stays in the correct location. Manage by cleaning the expected light bleeding with a compress soaked in saline solution (Figure 2).
      NOTE: The rat skin is thin and sagging or loose. Stabilization of the skin is crucial and required.
    3. Fully detach the tissue from the bone using small periosteal elevators (Figure 3).
    4. Expose the bone until identification of the insertion of the tibialis cranialis muscle, the gracilis, and the gastrocnemius lateral head muscle in the posterior border of the medial aspect of the tibia, as a fibrous white tissue firmly adhered to the bone (Figure 3).
      NOTE: It is important to identify this group of muscular insertions to enable the implant to be placed in a bone region with similar characteristics and stimuli across the entire sample, regardless of the rat's size.
  2. Drilling process
    1. Begin the drilling process at the correct region between the proximal tibial crest and the posterior limit of the medial face of the tibia bone, contiguous to the insertion of the tibialis cranialis muscle, the gracilis, and the gastrocnemius lateral head muscle, avoiding any muscle injury.1
      NOTE: The correct location should be 5 mm ± 2 mm from the tibial plateau.
    2. Drill with a maximum of 150 rpm (rotations/min) under saline solution irrigation at a temperature close to 20 °C with a surgical electric motor with a 20:1 reduction contra-angle.
      NOTE: Only two drills were required.
    3. Start with a lance pilot drill (Figure 4) at a depth of 2.4 mm under saline solution irrigation.
      NOTE: Each drill had a maximum of 10 uses.
    4. As a second drill, use (Figure 4) a twist design drill at a depth of 2.4 mm with a diameter of 1.6 mm under saline solution.
      NOTE: Each drill had a maximum of 10 uses.
  3. Implant placement
    1. Insert the implant with an intermediary piece (Figure 5) attached to the 20:1 reduction contra-angle.
    2. Before placing the implant, clean the implant from any residual chemical sterilant by rotating it in the contra-angle with simultaneous saline irrigation for 10 s (Figure 5).
    3. Place the titanium implant (2 mm lengthwise and 1.8 mm in diameter) using the intermediate piece at 20 rpm while monitoring real-time torque value, registering the maximum insertion torque.
      NOTE: An initial difficulty in inserting the implant is expected due to the difference between the final drill and the implant, as well as the cylindrical profile of the implant; however, this initial difficulty is quickly normalized as soon as the implant penetrates the initial cortical bone.
    4. Finish implant insertion before it completely passes the cortical bone where it is inserted, that is, the flat medial face of the tibia.
      NOTE: At the final moment of implant insertion, it is important to leave the implant slightly outside the cortical bone or level with the cortical bone where it is inserted, that is, the flat medial face of the tibia, to ensure primary stability (Figure 6).
  4. Wound closure
    1. Suture the muscle tissue borders with simple internal sutures using a 4/0 monofilament synthetic resorbable suture (Glyconate) (Figure 7).
    2. Perform skin closure with an intradermal suture using a 4/0 monofilament synthetic resorbable suture (Glyconate) (Figure 7).

4. Micro-CT scan

  1. After completing the surgery and still under general anesthesia, conduct a micro-CT scan to confirm proper implant placement.
  2. Remove the rat from the surgery bed and place it on the scanning bed. Locate the operated leg using the micro-CT live scan mode and center the Field of View on the implant.
    NOTE: Recommended acquisition parameters are Field of view of 5 mm, spatial resolution of 0.0001 mm3, 50 kV, 200 µA, and acquisition time of 3 min.
  3. Once the scan is acquired, confirm the correct distance between the proximal aspect of the implant and the surface of the tibial plateau, according to step 5.2.1.
    NOTE: This value will be helpful for the standardization of the technique (Figure 8).

5. Postoperative care

  1. Following imaging acquisition, return the rat to its cage and monitor until full recovery.
    NOTE: This takes approximately 5-10 min, depending on the animal model. Diabetic rats are expected to remain anesthetized longer and have a longer recovery time due to the metabolic changes associated with diabetes.
  2. Administer buprenorphine (0.1 mg/kg) every 6-8 h and meloxicam (5 mg/kg) every 24 h, subcutaneously, up to 72 h.
  3. Suture removal time
    1. Examine the surgical wound daily for infection, suture integrity, or other issues, and remove suture remnants, if necessary, 15 days post-surgery.

6. Euthanasia

  1. Euthanize the animals using a CO2 chamber, according to the National Institutes of Health (NIH) guidelines, with a CO2 fill rate of 30%-70% of the chamber volume/min after the implantation time (4 weeks or 12 weeks).

7. Postsurgical analysis

  1. For both models (osteoporosis and diabetes), remove the tibia by disarticulation after euthanasia for further analysis.
    NOTE: Regarding the micro-CT analysis, data allowed for calculating bone-implant contact (BIC). The CT acquisition was conducted using the previously described parameters, and the BIC analysis was performed by dividing the bone area (mm2) by the implant area (mm2) calculated at the cortical region, adapting a protocol already described in the literature18. Representative images are shown for each model, osteoporosis (Figure 9) and diabetes (Figure 10).

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Results

Surgical phase
It is important to mention that both animal models used in this study present certain constraints due to the induced diseases. These constraints regarding the manipulation of hard and soft tissues are reflected during the surgical procedure.

In the diabetic model, the rat is larger, making it difficult to stabilize the legs during surgical procedures. This increases the surgical time and, consequently, the anesthesia time, which requires a longer recovery ti...

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Discussion

Although the rat is a widely used model for studying osseointegration, it is important to define and describe a reproducible surgical technique for adequately placing implants. Such a technique could serve as a guide for the scientific community. Moreover, the fact that certain diseases, such as osteoporosis and diabetes, alter bone metabolism implies stronger demands for correctly designing surgical procedures. The rat compares favorably with other animal models since it presents the main features of both osteoporosis (...

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Disclosures

We hereby declare that there is no conflict of interest regarding this scientific article.

Acknowledgements

The authors thank the Spanish State Research Agency for financial support through projects PID2020-114019RBI00 and PID2021-125150OB-I00.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
22 G needles+A2:C30TerumoNN-2238R
4/0 monofilament synthetic resorbable sutureBraun ( MonoSyn)
5 mL, 10 mL syringesBraun4617100V-02 4606051V
Adson forcepsAntão MedicalRef: A586
BBDR ( Biobreeding Diabetes Resistant ) Sprague Dawley RatsJanvier Labs
BetadineMylan
BuprecareAnimalcare (UK)
Castroviejo Caliper 0-40 mm 15 cm angledUL AMIN Industries
Castroviejo Needle HolderAntão MedicalRef: AM1702
Dental surgery scissors curved and straightAntão MedicalAMA603 / AMA600
Electric shaverOster Pro 3000i34264482227
Extra Fine Graefe ForcepsF.S.TRef: 11150-10
Gauze padsCOVIDIEN441001
GlucometerMenarini (Italy)
Helicoidal Drill / OSTEO-PIN DRILL Ø1.6 mmsoadcoRef. OS-8001
Implants / SCREW OSTEO-PIN Ø1.8 x 2.0 mmsoadcoRef. OS-3
IsofloLe Vet Pharma (Netherlands)
Lance pilot drill / Lanceolate Drill (DS)soadcoRef. 10 02 01 T
Latex gloves - Surgical gloves sterileHartmannRef: 9426495
Lucas Surgical CuretteAntão MedicalRef: AMA940-3
MetacamBoehringer Ingelheim(Germany)
Micro forceps straightnopaRef: AB 542/12
Micro-CT scan( Quantum Fx microCT )Perkin Elmer (US)
Osteoporotic Sprague Dawley females RatsJanvier Labs
Periosteal elevator -  Molt 2-4Antão MedicalRef: A1564
Physiologic solution for IrrigationHygitechRef:10238
Scalpel Blade Carbon Steel 15CRazor MedRef: 02846
Sterile Gauze SwabsAlledentalRef: 270712
Sterile Irrigation systemHygitechRef:HY1-110001D
Sterile towels (1 piece per animal)Dinarex4410
Surgical contra-angle handpieceW&HRef: WS-75 LED G
Surgical contra-angle handpieceW&HSN 08877
Surgical contra-angle handpieceW&HSN 01309
Surgical Electric MotorWH Implantmed Type: SI-1023 Ref: 30288000
Surgical scalpel handleAsaDentalRef: 0350-3
Towel clampsXelpov surgicalAF-773-11
Ultrasonic deviceJ.P. Selecta, Abrera, Spain

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