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While in vitro studies have shown that nanoscale surface modifications influence cell fate and activity, there is little information on how they modulate healing at the bone–implant interface.
This study aims to investigate the effect of nanotopography at early time intervals when critical events for implant integration occur.
Untreated and sulfuric acid/hydrogen peroxide-treated machined-surface titanium alloy implants were placed in rat tibiae. Samples were processed for DNA microarray analysis and histomorphometry.
At both 3 and 5 days, the gene expression profile of the healing tissue around nanotextured implants differed from that around machined-surface implants or control empty holes, and were accompanied by an increase in bone–implant contact on day 5. While some standard pathways such as the immune response predominated, a number of unclassified genes were also implicated.
Nanotexture elicits an initial gene response that is more complex than suspected so far and favors healing at the bone–implant interface.
The current trend in implant research is the development of biomaterials with surfaces that will improve tissue integration and stability, and ensure clinical performance [1,2]. Surface characteristics such as chemical composition, morphology and energy can generate conditions at the bone–implant interface that significantly influence cell and tissue responses . While all scales of implant topography can potentially influence tissue healing events, it is increasingly recognized that interactions between biomaterials and host tissues occur on the nanoscale, the scale at which cells normally carry out their activity [4,5]. Several studies have now demonstrated that nanoscale surface cues can elicit change in cell attachment [6,7], proliferation and differentiation [8–10], organization of cell shape and cytoskeleton [10,11], and enhance the precipitation of apatite .
While there have been several in vitro studies on cell behavior and surface texture, very few have investigated the effect of nanoscale surface modifications on implant osseointegration in the complex and multifactorial environment of the body . Results from these few in vivo studies have suggested that nanoscale surface topography has a major impact on contact osteogenesis [13,14] and promotes interfacial strength . Despite these reports, little is known on how nanotopography influences the early healing response and the cellular and molecular events that are critical for determining the outcome of an implant. The objective of this study was, therefore, to determine the gene expression and histological profile of tissue healing at early time points around nanostructured titanium (Ti) alloy screw-shaped miniature implants placed in rat tibiae, as compared with ones with a machined surface and the empty implantation hole. We selected early time points that reflect initial healing events in noncritical size defects in rat . We used a chemical oxidative patterning approach with a mixture of sulfuric acid (H2SO4) and hydrogen peroxide (H2O2), which has proved efficient for various medically relevant metals [16–19], to nanostructure the implants. Such controlled oxidation predictably produces networks of nanosized pits of approximately 20 nm in diameter and with a root mean square roughness of approximately 15 nm [16–18,20]. This treatment has been shown to enhance osteogenic activity in vitro, to directly and selectively signal to cells, and to influence stem cell division and differentiation [17,19,21].
Miniature machined screw-shaped implants made of Ti-6Al-4V (Micro Med Machining Inc, FL, USA) were used. The screws had the following dimensions: 1.0 mm inner diameter; 1.5 mm outer diameter; 2.4 mm head diameter; 0.4 mm head thickness; and 1.6 mm implant length. Implants were washed with toluene and then nanotextured by treatment with a solution consisting of equal volumes of concentrated H2SO4 and H2O2 for 2 h, at room temperature (Figure 1) . Samples were rinsed with nanopure distilled water, washed with 70% ethanol and air-dried. Machined screw implants were washed with 70% ethanol, rinsed with water and air-dried. The surface characteristics of the machined and nanotextured screws were confirmed using a JEOL JSM-7400F (Tokyo, Japan) field emission scanning electron microscope (SEM) operated at 1–2 kV.
Forty male Wistar rats weighting 200–225 g (Charles Rivers Canada, QC, Canada) were anesthetized with an intraperitoneal injection of a mixture of Ketalean® (ketamine hydrochloride; Biomeda-MTC, ON, Canada), Rompun® (xylazine; Bayer Inc., ON, Canada) and Acevet® (acepromazine maleate; Vetoquinol Inc., QC, Canada). The anteromedial side of each hind limb was shaved and cleaned with Baxedin® (chlorhexidine gluconate; Omega Laboratories, QC, Canada). A 1-cm incision was made through the skin, and the skin and muscle were gently pried apart to expose the periosteum. Two holes were drilled through the periosteum and across the bone cortex at 5 mm from the knee joint, using a 1.4-mm round carbide burr (Brasseler, QC, Canada). The drilling site was irrigated with physiological saline and implants were placed in both tibiae. In a randomized manner, one tibia received two machined implants, while the other received two nanotextured implants. Control groups consisted of animals with holes without placement of a screw (empty hole). The muscle was sutured with 4-0 chromic gut sutures and the skin was closed with 4-0 sofsilk sutures (distributed by Patterson Dental Supply Inc., MN, USA). The surgical site was cleaned and disinfected with Baxedin. The animals received an injection of Temgesic® (buprenorphine hydrochloride; Reckitt and Colman, Hull, UK) after surgery and were fed with soft food containing Temgesic. All experimental protocols and animal handling described above were approved by the Comité de déontologie de l’expérimentation sur les animaux of Université de Montréal, Canada.
After 3 (n = 14) or 5 (n = 14) days, some animals were anesthetized, the wound was cleaned with 70% ethanol and opened with a scalpel blade and the screws were gently removed with a miniature jewelry screwdriver. Under physiological saline irrigation, the bone at the surgical site was harvested using a trephine drill (2.4-mm diameter, ACE Dental Implant System, MA, USA) fitted on a slow-speed hand-piece (Physiodispenser 3000, Henry Schein Inc., ON, Canada), placed in Trizol® (Invitrogen, ON, Canada) and immediately homogenized with the Polytron® (Kinematic Inc., NY, USA) at full speed for 1 min. Total RNA was extracted from these samples as recommended by the manufacturer and purified with the RNeasy® MiniElute® Cleanup kit (Qiagen, ON, Canada). RNA concentration, integrity and quality were analyzed using the Agilent Bioanalyzer 2100 at McGill University and Génome Québec Innovation Centre (QC, Canada). Only RNA samples showing a sharp distinction of clear 18s and 28s ribosomal RNA peaks and a high RNA integrity number were considered of ‘good quality’ and were used for DNA microarray analyses. The number of samples for day 3 (n = 14) and 5 (n = 14) was calculated after eliminating samples with poor-quality RNA.
Microarray analysis using an Illumina® RatRef-12 Expression BeadChip (22106 probe sets; Illumina, S CA, USA) was performed at McGill University and Génome Québec Innovation Centre. Expression analyses of microarray data were performed by Genexanalysis (QC, Canada ) using the method of consecutive sampling and coincidence testing [22–24] and statistical significance was estimated based on the method of stabilized variance .
Using the PANTHER Classification System , a ‘gene ontology’ classification of the genes/expressed sequence tags (ESTs) that were significantly (p < 0.05) up- or down-regulated was performed for both implants and time points (nanotextured implants vs empty hole, machined implants vs empty hole, and nano-textured vs machined implants at both 3 and 5 days post-surgery). Pathway analyses of differentially expressed genes (probability interval: 0.9) were carried out using the GO-Elite software (Gene Map Annotator and Pathway Profiler [GenMAPP], Gladstone Institute at the University of California at San Francisco [CA, USA]) .
After 3 (n = 6) or 5 days (n = 6), animals were anesthetized and perfusion-fixed . Tibia were dissected out and immersed in a fixative solution consisting of 4% paraformaldehyde (Fisher Scientific, ON, Canada) and 0.1% glutaraldehyde (Electron Microscopy Sciences, PA, USA) in 0.08 M sodium cacodylate (Electron Microscopy Sciences) buffer containing 0.05% calcium chloride (Sigma-Aldrich Canada Ltd, ON, Canada), pH 7.2, for 3 h at 4°C. They were then washed with 0.1 M sodium cacodylate buffer, pH 7.2. Prior to processing for embedding in methylmethacrylate resin (Technovit® 9100 new, Electron Microscopy Sciences) , x-rays (7 mA, 60 kv, 0.500 s; Progeny Dental, IL, USA) of the dissected tibiae were taken to verify the positioning of the implant (Figure 2). Longitudinal sections that were 5- to 8-μm thick including the implant and surrounding tissue, were cut with a Reichert-Jung Polycut E microtome (Leica, Heidelberg, Germany), collected on superfrost-plus slides for histology, deplasticized with 2-methoxyethyl acetate (Sigma-Aldrich Canada Ltd, ON, Canada), rehydrated with decreasing concentrations of ethanol and stained according to the Goldner’s trichrome protocol. Goldner’s trichrome stains mineralized bone in green and unmineralized bone (osteoid) in orange/red color. Histological observations were carried out with an Axiophot light microscope (Carl Zeiss, Oberkochen, Germany).
Goldner’s trichrome stained sections were photographed and 12 bone areas in each group were analyzed. A 250-μm square was drawn around the implant or in the empty hole; green pixels were picked up as a bone tissue using imaging software (Photoshop CS4; Adobe, CA, USA). These pixels were translated into the bone area and expressed by the ratio of total tissue area in the 250-μm square. The length of bone–implant contact was expressed by the total length of green pixels on the implant–tissue surface, and demonstrated by the ratio of total implant–tissue surface length in the 250-μm square. Statistical differences of bone area around two different implants and in the empty hole were examined using the Kruskal–Wallis one-way analysis of variance. A Mann–Whitney U test was also used to statistically compare the bone–implant contact between nanotextured and machined-surface implant groups; p < 0.01 was considered significant.
As previously reported , SEM examination confirmed that surface treatment of Ti-6Al-4V screw-shaped implants with the H2SO4/H2O2 oxidative solution created a nanopatterned surface with uniformly distributed pores of approximately 20 nm in size (Figure 1).
For all animals, healing was uneventful following implant installation. No sign of infection or inflammation was noticed at the surgical site and all implants were stable. Histological sections stained with Goldner’s trichrome revealed that as early as 3 days post-surgery, in the empty hole group, newly formed bone was noted within the bone marrow cavity (Figure 3A & B) and almost completely filled the surgical site by day 5 (Figure 3C). The presence of residual bone debris produced during drilling and implantation of machined (Figure 3D & e) and nanotextured screws (Figure 3g & H) was also noted in the bone marrow cavity. However, new bone formation was not evident at this time point for both types of implants. By day 5, there was an increase in the amount of bone forming around both machined (Figure 3F) and nanotextured screw-shaped implants (Figure 3i). Direct bone contact with the implant surface was clearly observed in nanotextured implants (Figure 3i), while it was only occasionally noted in machined implants (Figure 3F).
Histomorphometric analysis did not reveal any statistical difference in the bone formation area in empty hole, machined and nanotextured implants at days 3 and 5 (Figure 4). At day 3, no difference of bone formation and bone–implant contact was measured between the machined and nanotextured implant. However, by day 5, the bone–implant contact was significantly increased in nanotextured implants. The high standard deviations measured are probably related to the variability in bone formation in the various regions along the implant. However, statistical analyses validated the differences in bone–implant contact observed.
We examined the global gene expression profiles in RNA pools from the healing tissue harvested in an empty hole and around machined and nanotextured screw-shaped implants, at both days 3 and 5 post-surgery. On both days, stable nanotextured screws elicited a different gene expression profile in the healing tissue compared with machined-surface screws and the empty hole. A detailed list of differentially expressed genes for both surfaces and time points is presented in Supplementary TAbles 1–6 (see online, www.futuremedicine.com/doi/suppl/10.2217/NNM.12.167).
Since the identification of individual genes conveys limited information on how surface roughness affects interfacial tissue healing, differentially expressed genes were classified into various biological processes in order to assess their functional significance (Figures 5–7). The proportional distribution of up-(Figures 5A & B, 6A & B, 7A & B) and down-regulated (Figures 5C & D, 6C & D, 7C & D) genes in each process is illustrated for day 3 (Figure 5) and 5 (Figure 6) post-surgery.
To identify potential local signaling pathways associated with tissue healing response around nanotextured and machined implants and in an empty hole, we analyzed our microarray expression data using GenMAPP and results are listed in Tables 1–5. These results indicate that different pathways have been activated with the placement of nanotextured and machined implants.
Biochemical signals and physicochemical cues both influence the cellular response to a biomaterial at the site of implantation [28,29]. Learning how to best integrate these two contributions is, therefore, essential to optimize tissue repair and even achieve regeneration. Although several in vitro studies have been carried out on the benefits of nanostructured surfaces on the biological response to implants , investigating the relevance of such surfaces in the complex biological environment of the body, where implants are exposed to multiple cell types that may not all respond in the same way to a given surface cue needs to be defined. Moreover, while whole-genome expression profiles of healing bone around microtextured implants have just recently been reported , that of next-generation nanotextured surfaces remains to be addressed. Data from this combined histomorphometric and gene expression study show that nanotextured surfaces lead to changes in expression of gene sets at the bone–implant interface that result in differences in bone healing that favor implant stability. Moreover, they provide insight into the molecular mechanisms involved in osseointegration of implants and on the response to surface topography. Importantly, all of our analyses were carried out on calcified tissues from which the implants were not removed. This avoided tissue damage, which normally occurs when an implant is pulled out, and therefore, to characterization of the healing process at the real tissue–implant interface.
A major distinction between our study and the recently reported microarray analyses of bone healing around microtextured dental implants placed in human patients  is the nature of the bone–implant contact. In the case of dental implants, they are usually placed in compact bone and are largely in contact, throughout their length, with old bone. Changes in gene expression around such implants would, therefore, probably include a strong bone remodelling component. In our case, most of the implant surface extended in the medullary cavity of the tibia, and hence the resulting gene expression profile most likely reflected bone induction events around the implant. In addition, because our sampling region was very close to the implant (0.45 mm), the changes observed are likely to relate to events at the tissue–implant interface where cell surface interactions actually take place.
One striking aspect of gene profiling studies following marrow ablation , mechanical loading of bone , and of our study, is that initially the genes that are up- or down-regulated are related to general biological processes such as protein metabolism and modification, signal transduction and immune response. In the case of the marrow ablation experiment  and with our nanostructured surfaces, note also the participation of several miscellaneous genes and/or those with undefined name/function. This suggests that the early bone healing response, in particular to implantable metals, may be more complex than so far suspected and probably initially involves general healing genes that will eventually be supplanted by osteogenic ones as bone builds up. Altogether, this suggests that focusing on standard pathways may only provide a limited picture, particularly at early time intervals when the fate of a biomaterial is determined.
Omar et al. compared the expression of selected proinflammatory cytokines, chemokines, adhesion molecules and bone formation and resorption markers at various time points following the implantation in rat tibiae of machined-surfaced and anodically oxidized Ti implants [34–36]. They concluded that the microtexture generated by their anodization process stimulates the recruitment and adhesion of cells that are crucial for regenerative processes at implant surfaces. In particular, they note that microtexture elicits a reduced inflammatory response. In the case of our nanotextured implants, there is a major modulation of the immunoglobulin and immune response gene category, with some genes undergoing downregulation while others are upregulated. The screening and identification of these up- and down-regulated genes will be validated using quantitative PCR in future studies. Thus, a better understanding of the temporally adapted control of the various aspects of the inflammatory process may lead to the rational design of surfaces that will optimize healing and new bone formation around implants.
In conclusion, our study provides the first integrated histomorphometric and molecular profile of the early bone healing events around implants with machined as compared with nanostructured surfaces. The data show that a nanoporous Ti alloy surface unquestionably has a major effect on how the body responds to the implant. The data also point to a number of potential candidate genes, so far unsuspected, that may help us understand how nanotexture influences local and systemic aspects of the tissue healing response. Recent literature from various laboratories and using different models indicates that altering the physicochemical properties of the surface of various types of biomaterials influences the activity of several cell types [5,37,38]. Most of these studies [37,38] are in vitro and will need to be validated in the complex environment of the body.
In the years to come, one can expect that novel generations of implants will be available that will have rationally designed surfaces capable of providing signaling cues to the surrounding environment in order to modulate cell activity at tissue–implant interface. These surfaces will be capable of promoting bone healing and regeneration, to improve implant integration. These innovations are needed to address the challenges of a demographically ageing population with compromised bone status.
The authors acknowledge the services from Génome Québec Innovation Center at McGill University for performing the DNA microarray analysis and Genexanalysis for the service with quantitative analysis of data.
Ethical conduct of research
The authors state that they have obtained appropriate institutional review board approval or have followed the principles outlined in the Declaration of Helsinki for all human or animal experimental investigations. In addition, for investigations involving human subjects, informed consent has been obtained from the participants involved.
Financial & competing interests disclosure
This work was supported by Canadian Institutes of Health Research, The Network for Oral and Bone Health Research and NIH EB000504-04 grants. The authors have no other relevant affiliations or financial involvement with any organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript apart from those disclosed.
No writing assistance was utilized in the production of this manuscript.
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