Implantation
1. Obtain Wistar rats (12-week-old is here exemplified)
2. Perform a unique animal codification (1, 2, 3…)
3. Mirror the codification in a spreadsheet column and highlight the relevant experimental group division by material(s) to be implanted and time point(s) (to be printed and affixed in the surgical room for guidance)
4. Acquire the selected material(s) for the in vivo implantation after the in vitro validation
5. Chemically characterise the material before implantation using preferably different analytical techniques14,36 to double-check the material chemical composition stated by the manufacturer. It is utterly encouraged to report the material(s) batch(es)
6. Stablish in vitro replicable material handling and manipulation (including material flow and setting time) in advance (in a sufficient number of samples that allow replicability) by either following the manufacturer's instructions or a relevant ISO standard37 for the material testing
7. Polyethylene tubes of 1.0 mm internal diameter, 1.6 mm external diameter and 10.0 mm length can be used36 – this tube configuration can be adjusted according to the study design following the instructions of the relevant ISO standard38 that suggest: ‘Non-solid materials (including powders) may be contained in open-ended cylindrical tubes for the purpose of testing for local effects after implantation’ and ‘test samples shall be 1,5 mm to 2 mm in diameter, 5 mm to 10 mm in length and have rounded ends’
8. Sterilise the tubes using ethylene oxide according to the relevant ISO standard (ISO 10993-7:2008 Biological evaluation of medical devices Part 7: Ethylene oxide sterilization residuals)
9. Run all the following steps in a dedicated private room with low light, clean floor, with disinfected surfaces/covered by a surgical sterile cloth and isolated from noise. It is crucial to perform the experiment at one animal at a time, to avoid stress
10. Veterinary scale using the 'live animal' mode (i.e., Mars AD500S digital scale, São Paulo, SP) for the weight assessments
11. Run normality test (i.e., Shapiro Wilk) to evaluate the data distribution (a normal distribution is desired)
12. Separate one animal at a time to perform intramuscular anaesthesia with a combination of ketamine (Dopalen®) (80 mg/kg) and xylazine (Anasedan®) (10 mg/kg). Immobilisation should be performed using a clean cloth covering the animal vision (Figure 1)
13. After approximately five minutes of anaesthesia, test the animal's responsiveness by gentle digital compression of the paw. If the animal is still responsive, the anaesthesia can be supplemented with 1/4 of the initial dose, followed by three minutes wait prior to the second responsiveness test.
14. Perform trichotomy of the dorsal region gently in a squared format (~5 x 5 cm) using blades or electric shaving machines (Figure 2)
15. Perform antisepsis of the trichotomized area with povidone-iodine (PVPI) (Figure 2)
16. Hold the tissue up to avoid a deep blade perforation. This step can be performed using either the opposite hand or with a delicate Lane tissue forceps
17. Make a continuous longitudinal incision in the mid-dorsal region using a #15 scalpel (Figure 2)
18. Perform the divulsion of the subcutaneous tissue with blunt-tip scissors bilaterally along the animal flanks for the insertion of the filled and control tubes (Figure 2)
19. When 4 tubes are to be implanted, we strongly recommend that two separate smaller incisions are performed, and one tube can be placed in each side (Figure 3)
20. Hydrate the calcium silicate-based materials according to the manufacturer's instructions or to prior in vitro validation observing steps 4, 5 and 6
21. Considering that 4 tubes will be implanted in each animal, fill 3 sterile tubes with the freshly mixed material and leave 1 tube empty, for control
22. Place one filled tube in positions ‘a’, ‘b’ and ‘c’ and the empty tube in position ‘d’ for control (Figure 3)
23. Make sure that the tubes are in direct contact with the exposed connective tissue and sufficiently far from the incision – into the animal flank – to prevent their expulsion during the animal's mobility
24. Perform suture with 4.0 silk thread (i.e., Ethilon, Johnson & Johnson, USA) with four or five suture points for the continuous incision or two to three suture points for the smaller incision
25. Identify animals grouping by material in identified cages. Never mix animals with different implanted materials to avoid any contamination through contact, saliva, urine, faeces and/or cross-contamination by the food provided
26. Administer dipyrone at a dosage of 100 mg/kg on the opposite paw used for the anaesthesia (Figure 4)
27. Evaluate the animal recover daily considering any signs of distress. If animals present extended signs of discomfort, consider anticipated euthanasia (and exclusion from the study) with the consultation of animal research facility veterinarian
28. Keep, for five to seven days, the animals in individual cages for recovery
29. An initial hair growth can be observed in the trichotomy region after this period
30. Group the animals according to the implanted material
31. Provide a controlled room temperature (20 ± 1°C) during the experimental period with an automatic 12/12-hour light cycle (lights on from 8 am to 8 pm) and a relative humidity of ~55%. The normal temperature range in rats is between 35.9 and 37.5°C39; this physiological parameter can be used during postoperative care after the tube implantations
32. Provide a balanced commercial solid food ad libitum
33. Provide filtered water ad libitum
34. Sanitize the cages by replacing the animals into a new cage containing ‘fresh’ bedding (wood shavings) periodically (every 2 to 3 days) throughout the experimental period
Euthanasia
35. The animal needs to be deeply anesthetized (assessing again its weight) with a tripled dose, also intramuscularly
36. At this point, intracardiac blood sampling (3 to 5 ml for each animal) can be performed using a 25.06 needle and a 5 ml syringe after ventral full access, if planned40. Store the blood sample in a vial with an anticoagulant for erythrograms and leukograms (purple lidded vial) or biochemical analysis light-protected (red lidded vial) (Figure 5)
37. The euthanasia can be confirmed by observing the absence of respiratory movement (apnoea) and the loss of the corneal reflex
38. Locate the implanted tube area by palpation and incise and dissect the tissue. Make sure to cover sufficient surrounding normal tissue of at least 5 mm to each tube side
39. Immerse the tissue segments containing the tube for fixation in a 10% buffered formalin at neutral pH, in individual identified (using the unique code described in steps 2 and 3). It is highly recommended to have all the necessary individual flasks identified and filled with the 10% buffered formalin to at least ten-fold the sample volume (5 to 8 ml, for these shown samples) previously to the proceedings. Do not perform the euthanasia without the flasks ready to be used
40. The tissues in the flasks containing 10% buffered formalin can be stored at room temperature. A minimum of 24 to 48 hours for tissue fixation is required
Histological processing
(Briefly described here, and adjustable to the availability of equipment)
41. For the histological processing the samples must be washed in running water for 12 hours
42. The excess adjacent tissue can be eliminated manually using microtome blades, leaving the sample with a rectangular shape and tissue surrounding the implant, sufficient to carry out the processing
43. The tubes can be now carefully separated from the fixated tissue and the tissue allocated separately in histological cassettes for a brief second fixation (Figure 6)
44. Washed in running water overnight
45. Sample dehydration is accomplished by immersion in graded steps of ethanol (50%, 70% and 100%) for dehydration for 1.5 hours for each concentration. CRUCIAL STEP: Additionally, two extra 1.5-hour immersion cycles must be performed using the 100% ethanol
46. Immerse samples in 1:1 proportion of 100% ethanol and xylene P.A. for 12 hours
47. Immerse samples in fresh xylene P.A. for three cycles of 1.5 hours
48. Paraffin embedding (i.e., Histosec® or Paraplast®) – suggested to liquefy at 62°C and immerse samples twice for 1 hour
49. Place the sample in aluminium moulds for cooling
50. The paraffin blocks can now be subjected to semi-serial microtomy using a microtome in 5 µm sections and its placement over glass slides. CRUCIAL STEP: make sure that the selected slices are obtained in longitudinal direction of the tubular tissue
51. Perform routine haematoxylin and eosin staining (H&E)
a. Haematoxylin (Gills III)
i. Reagents (for ~1 L of solution)
1. Ethylene glycol – 250 ml
2. Distilled water – 750 ml
3. Haematoxylin – 6 g
4. Sodium Iodate – 0.6 g
5. Aluminium sulphate – 80 g
6. Glacial acetic acid – 20 ml
ii. Dissolve ethylene glycol in distilled water
iii. Add haematoxylin crystals
iv. Add sodium iodate
v. Add acetic acid
vi. Add chloral hydrate – agitate gently until fully homogeneous (~15 min). If prepared correctly, the solution will present a strong red colour and in contact with tap water will become blue
b. Eosin Y stock solution:
i. Reagents (for~ 200 ml of stock solution)
1. Eosin Y – 2 g
2. Distilled water – 40 ml
3. 95% ethanol – 160 ml
ii. Dissolve Eosin Y in water, mix until dissolved
iii. Add ethanol and mix
c. Eosin Y working solution:
i. Reagents (for ~800 ml of working solution)
1. Eosin stock solution – 200ml
2. 70% ethanol – 200 ml
3. Glacial acetic acid – 4 ml
ii. Dilute eosin stock solution in 70% ethanol
d. Other required solutions:
i. 0.3% acetic acid in distilled water
ii. 0.3% hydrochloric acid in 70% ethanol
iii. Scott’s tap water substitute
e. H&E staining41, 42 - place slides in a suitable holder, adding enough of the following solutions to ensure slides are covered:
i. Dewaxing:
1. Xylene – 5 min (twice)
ii. Sample rehydration:
1. 100% ethanol – 1 minute (twice)
2. 95% ethanol – 1 minute
3. 70% ethanol – 1 minute
4. Tap water – 2 minutes
iii. Staining:
1. Haematoxylin (Gills) – 4 mins
2. Running tap water – 2 minutes (until water runs clear)
3. Differentiate with 0.3% acetic acid – 30 seconds
4. Drain
5. Treat with 0.3% hydrochloric acid in 70% ethanol – 30 seconds
6. Running tap water – 2 minutes
7. Treat with Scott’s tap water substitute – 2 mins
8. Running tap water – 2 mins
9. Eosin – 1 min
10. Running tap water – 2 minutes (until water runs clear)
11. Drain well
iv. Dehydration:
1. 100% ethanol – 1 min (twice)
2. Xylene – 1 min (twice)
f. Canada balsam or DPX for permanent mounting of slides for microscopy, including a coverslip.
52. Immunohistochemistry techniques can also be performed after these described steps according to different protocols43 and markers of interest can be selected to stain the obtained sections
53. The number of inflammatory cells can be counted for each specimen image (i.e., n = 35 slides for each material/time point or control/time at 40× magnification, as previously reported by our research group36) by a calibrated operator ideally blinded to the image condition, using a light microscopy (i.e., DM 4000 B; Leica Microsystem, Wetzlar, Germany)
54. Acquire and export images in RAW, TIFF or other uncompressed format (compressed image formats are not suitable)
55. Scale bars can be positioned within the images using ImageJ software (https://imagej.net/ij/download.html) after calibration (adjustable for each microscopy and image acquisition system)