Effect of Different Storage Media on the Regenerative Potential of Autogenous Bone Grafts: A Histomorphometrical Analysis in Rabbits

Bone grafts are frequently used for treatment of severe bone loss.^1–3  Autogenous bone grafts are more predictable when compared with other biomaterials because of their biological properties,⁴  and they are considered the gold standard for bone reconstruction.^1,2,4  Graft volume maintenance and early bone remodeling are important factors in the success of these surgical procedures.⁵ 

During a bone-grafting procedure, there is usually a delay between removal of the graft and its stabilization. Preparation of the recipient sites may sometimes begin after the donor site has been surgically closed.⁶  This raises several questions concerning the harmful effects of this delay over osteogenesis and the optimal environment for the graft.^7,8 

Experimental evidence demonstrates that the living cells within a fresh bone graft contribute to osteogenesis within and around the graft.²  If the graft cells are not viable or lost before implantation, little or no bone is produced, whereas in an intact living graft, abundant new bone appears.^7–9  Even with the occurrence of deleterious effects due to inappropriate graft handling during the transsurgical period, osteogenic cells may survive and contribute to the synthesis of new bone after transplantation under favorable conditions. Therefore, a proper storage medium becomes important to preserve as many cells as possible and improve bone repair^10,11  when immediate graft placement at the recipient site is not possible. Despite several solutions tested as storage media for bone grafts such as saline solution,^7,9,11–14  distilled water,^14,15  blood,^9,13  platelet-poor plasma (PPP),^6,13  lactate solution of dextrose 5%,^13–15  air exposure,^11,13,16  and different culture media,^9,14  only little is known about the in vivo influence of autogenous bone grafts storage on the outcome. Furthermore, the results of these studies remain controversial, and no conclusions could be drawn regarding the most adequate storage media.

Bone graft maintenance in air exposure or in saline solution is often seen in clinical practice. Air exposure is considered the most detrimental media²  because it leads to absence of nutrients and oxygen and consequently causes cell balance disturbance. Saline solution, easily obtained, can also be used for bone graft storage with favorable in vitro results.²  Currently, no studies have attempted to evaluate the in vivo effect of PPP as an alternative storage media, although there is evidence of the positive effect of PPP when compared with saline solution in an in vitro model.⁶  Therefore, the aim of this study was to evaluate in vivo the effect of storage autogenous bone grafts in dry, saline solution, and PPP media during the transsurgical time.

Eighteen healthy New Zealand white female rabbits (Oryctolagus cuniculus) weighting 2.5 to 3.5 kg were randomly divided into 3 groups, as described in Table 1. This study was performed in agreement with the rules of the Brazilian College of Animal Experiments.

For the surgical procedures, the animals were anesthetized intramuscularly with ketamine (25 mg/kg)/xylazine (10 mg/kg)/acepran (0.2 mg/kg)/midazolam (0.2 mg/kg) and local anesthesia with 0.9 mL of mepivacaine with epinephrine. To avoid infection, a single prophylactic dose of antibiotic therapy with cephalosporin (30 mg/kg) was administered intravenously at the same time as the administration of anesthesia. Five milliliters of autologous blood was drawn from each rabbit from the auricular vein several minutes before administration of anesthesia. The 5 mL of autologous blood was combined with 0.5 mL of 3.8% sodium citrate to prevent coagulation. The blood was centrifuged (206-BL-Fanem; Datamed, São Paulo, Brazil) according to the Sonnleitner modified method¹⁷  at 1000 rpm (160g) for 20 min to separate the plasma containing the platelets from the red cells. The supernatant and 2 mm below the “mist” between the phases was pipetted and transferred to a tube without anticoagulant. An additional centrifugation for 15 minutes at 1600 rpm (400g) was done to separate the platelets. The precipitate formed in the tube by this second centrifugation was the platelet-rich plasma (PRP), used for another experiment, and the superficial part was the PPP used in this study. The concomitant use of PPP, already prepared, could be an interesting storage media for autogenous bone grafts.

Surgery was performed under aseptic conditions. With the rabbits in the ventral position, trichotomy and antisepsis with aqueous solution of povidone-iodine (PVP iodine) was performed. Midsagittal incisions were made along the frontal bone to the occipital bone (Figure 1a), the periosteum was raised laterally (Figure 1b), and 2 bicortical standardized bony fragments were harvested from parietal bone with a 8.0-mm trephine drill (Figure 1c and d) under copious saline solution irrigation.⁵ 

Recipient sites were prepared through bilateral incisions in the submandibular area (Figure 2a) and elevation of muscle and periosteum (Figure 2b and c). The center of the fragment was perforated with a 1.0-mm drill under saline solution irrigation. The first fragment removed was immediately stabilized with a 1.2-mm × 6-mm titanium screw in the right mandibular angle (control group). The second fragment was maintained in air exposure (dry group), 0.9% NaCl solution (saline group), or PPP (PPP group) for 30 minutes and then fixed in left mandibular angle, in a symmetrical location of the control graft, in the same animal (Figure 2d). The periosteum, muscle fascia, and skin were sutured with nylon 5-0, and the skin sutures were removed after 7 days. Attending veterinary recommendations, all animals received intramuscular antibiotics Flotril (Enrofloxacina 2.5%–2.5 mL/kg) during 5 days after surgery. The animals received a normal diet consisting of granular food and water ad libitum. After a 4-week survival period, they were anesthetized with pentothal sodium 2.5% and euthanized with an overdose of potassium chloride 19.1%.

Bone fragments containing the grafted bone and surrounding tissue were removed and immediately immersed in 10% phosphate-buffered formaldehyde solution during 48 hours. All specimens were decalcified in EDTA 4.13%, dehydrated with graded ethanol, and embedded in paraffin. From the central perforation of the graft (screw region), 5-μm-thickness semiserial sections were obtained and stained in hematoxylin and eosin (HE) and Mallory trichrome (MT). Histological evaluation was performed under light microscope.

Empty lacunae (represents the loss of living cells) and bone graft area (mm² ; represents the amount of bone maintained) were quantified for each section by the same examiner in a blind way. Empty lacunae analysis was performed in 4 randomly selected histological fields per section using ×100 magnification (Instrutherm Mod MBB-200) in sections stained in HE.

The histological images of the graft area were captured at ×4 magnification in sections stained in MT, using an Olympus BX 40 binocular microscope (Shinjuku-ku, Tokyo, Japan) coupled with an Olympus OLY 200 camera (Center Valley, Penn) linked to a PC computer through a 3153 Data Translation digitizer plate (Marlboro, Mass).

All graft area images were captured from each histological section (Figure 3a) and then edited using Photoshop CS2 software to maintain only the bone graft area without the recipient bed and soft tissues, which were identified in histological images (Figure 3b). The bone graft area was converted into binary images and quantified with the HL Image program (Western Vision Software, Salt Lake City, Utah; Figure 3c).¹⁷  The editing process was performed twice in each section by the same examiner. The results obtained were submitted to normality test and paired Student t test, analysis of variance, and Tukey test. Differences were considered statistically significant if P ≤ .05.

Histological analysis in all experimental groups showed bone tissue with normal morphology (Figure 4). In all groups, we observed bone graft integration to recipient bed with bone formation areas. There was a progressive decrease in the empty lacunae next to the periosteum in all groups.

The median number of empty lacunae was significantly greater in the dry group (13.80 ± 1.74) compared with its control (11.08 ± 2.38; P = .05). No significant differences in the number of empty lacunae was found in saline group (12.10 ± 2.56) when compared with its control (11.10 ± 2.47; P = .22) and the PPP group (8.30 ± 1.64) when compared with its control (9.03 ± 3.03; P = .31). The dry group showed more empty lacunae than the saline group (P < .05) or PPP group (P < .01; Figure 5).

The analysis of bone graft area (mm²) revealed no significant differences between the dry group (1.30 ± 0.26) and its control (1.84 ± 0.30; P = .06), between the saline group (1.55 ± 0.27) and its control (1.63 ± 0.23; P = .14), or between the PPP group (2.17 ± 0.35) and its control (2.16 ± 0.56; P = .25). However, specimens stored in saline and PPP had a significantly higher bone graft area when compared with the dry group (P < .01; Figure 6).

In the incorporation of autogenous bone grafts, an important, but less often studied, matter concerns the effect of temporary methods for bone storage on osteocyte viability and bone volume maintenance. In the current in vivo study, osteocyte survival and bone graft area were affected by different handling techniques, resembling available options in clinical practice.

The maintenance in a solution allows the graft to become impregnated with the nutritious media that could sustain it after implantation⁷  until the blood supply is established. Viable cells are required to achieve bone formation instead of only having the presence of a potentially osteoinductive extracellular matrix.^18,19  The induction of osteocyte apoptosis suggests a possible causative pathway linking cell death and regulation of bone remodeling.¹¹  Resorption of bone containing living osteocytes is smaller than that occurring in bone where the osteocytes are dead.²⁰  When graft living cells are lost at the implantation time, due to inappropriate handling, bone-forming cells will be derived from host tissues; likewise, new bone formation is delayed. It seems feasible to hypothesize that various storage techniques might affect osteogenic potential.¹¹ 

Bone tissue should not stay for too long in completely hypoxic conditions because air exposure of a graft for a prolonged time reduces the proliferation of osteoblasts in vitro¹¹  and bone neoformation in vivo.²¹  We found an increased number of empty lacunae and decreased bone graft area after air exposure (dry group) when compared with the saline and PPP groups, suggesting loss of viable cells and damage to the bone graft volume maintenance. Although the absence of cells in the graft probably does not directly interfere with osteoconduction,²⁰  it compromises osteoinduction and increases resorption.^20,22  As long as osteocytes stay active, they may inhibit osteoclasts, preventing bone resorption. As soon as its network is damaged, this inhibition process is abolished and resorption would begin.²²  However, although osteocytes are important cells in molecular signaling and maintenance of bone vitality, the induction of bone resorption seems to be a more complex process in which several other factors participate.

In vitro studies have shown that graft storage in saline solution reduces the number of viable cells^7,9,12,13,15  and cellular metabolism.¹⁰  However, studies evaluating the bone neoformation in vivo²¹  and the proliferation of osteoblasts in vitro¹⁴  did not observe any differences between the control graft and the graft maintained in saline solution. In the current study, saline solution demonstrated satisfactory results as a storage solution due to osteocyte preservation and graft area. As known, osteocytes maintained in saline solution show morphological changes as time goes by¹²  and could be metabolically altered, which was not evaluated. Therefore, the present analysis could have been influenced by the absence of longer storage periods. Divergent results may be due to different methods of study, including the storage period, study in vivo or in vitro, animal model, and others.

Platelet-poor plasma has been suggested as an alternative storage medium.⁹  Grafts stored in PPP are found to have similar results to control grafts and to present more osteocyte preservation than saline solution.^9,13  Although the results were not significant, in the present study the PPP group showed greater results than the saline group. This fact could be related to the presence of growth factors in PPP, which might be favorable for cell balance.²³  Platelet-poor plasma contains the same growth factors found in PRP at lower concentrations.²⁴  It has been already reported that the growth factors from platelet concentrates could have a profound effect on bone graft physiology, participating in the production of cell survival factors,²⁵  which would increase the cell survival time. Moreover, the preparation of PPP does not demand complex techniques⁹ ; thus, in procedures in which PRP will be used, the concomitant use of PPP could be an interesting storage medium for autogenous bone grafts.

Further studies are necessary to clarify the complex cellular and molecular interactions between the bone graft and host bed and will provide the basis for new approaches to enhancement of bone graft incorporation and perhaps enable the clinician to ensure successful bone grafts even when adverse conditions exist.

The dry group showed more empty lacunae and less bone graft area maintenance than the PPP and saline solution groups, which was statistically significant. However, bone graft area maintenance revealed no significant differences between the dry, saline, and PPP groups when compared with controls. Therefore, saline and PPP might be considered optimal storage media for autogenous bone grafts during the transsurgical period in comparison with dry storage in this preclinical model.

Abbreviations

HE

hematoxylin and eosin

MT

Mallory trichrome

PPP

platelet-poor plasma

PRP

platelet-rich plasma