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Published on 04 October 2018. Downloaded by Xian Jiaotong University on 11/9/2018 1:45:39 AM.

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Cite this: DOI: 10.1039/c8bm00901e

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Osteoimmunomodulation, osseointegration, and in vivo mechanical integrity of pure Mg coated with HA nanorod/pore-sealed MgO bilayer† Bo Li,a,d Peng Gao,b Haoqiang Zhang,b Zheng Guo,b Yufeng Zhengc and Yong Han *a Fast degradation of Mg-based implants results in the loss of mechanical integrity and poor osseointegration. Herein, a bilayer-structured coating (termed as HAT), comprising an outer layer of hydroxyapatite (HA) nanorods and an inner layer of pores-sealed MgO with HA/Mg(OH)2, was formed on Mg using plasma electrolytic oxidation and hydrothermal treatment. Osteoimmunomodulation, osseointegration, mechanical integrity, and bone–implant interfacial structure evolution of the HAT-coated Mg were investigated by implantation in rabbit femora, together with Mg coated with plasma electrolytic oxidized porous MgO (termed as PEO0) and bare Mg. As compared to PEO0-coated and bare Mg, HAT-coated Mg greatly downregulated pro-inflammatory TNF-α and IL-1β, upregulated anti-inflammatory IL-10, and suppressed osteoclastogenesis, modulating the surrounding microenvironment toward favoring the recruitment of osteogenetic cells. Moreover, HAT-coated Mg accelerated bone sialoprotein and osteopontin secretion of osteogenetic cells and their mineralization to form a cement line matrix. It also promoted the differentiation of osteogenetic cells, secretion of collagen overlying on the cement line matrix, inducing an earlier and more pronounced bone matrix formation. The cement line matrix wrapped the HA nanorods and filled the interrod spaces of the HAT coating, forming strong interdigitation at the bone–coating interface, and therefore, yielding enhanced osseointegration by means of contact osteogenesis. Due to

Received 31st July 2018, Accepted 3rd October 2018 DOI: 10.1039/c8bm00901e rsc.li/biomaterials-science

the considerably reduced corrosion of Mg by the pores-sealed bilayer structure of HAT coating, HATcoated Mg maintained the mechanical integrity for a longer duration than PEO0-coated and bare Mg. It is clarified that the degradation of MgO and HA, rather than delamination, was the vanishing mode of PEO0 and HAT coatings during long-term implantation, avoiding osteolysis induced by the delamination-generated particles.

1. a State Key Laboratory for Mechanical Behavior of Materials, Xi’an Jiaotong University, Xi’an 710049, China. E-mail: [email protected]; Fax: +86-02982663453; Tel: +86-02982665580 b Department of Orthopedics, Xijing Hospital, Fourth Military Medical University, Xi’an, 710032, China c Center for Biomedical Materials and Tissue Engineering, Academy for Advanced Interdisciplinary Studies, Peking University, Beijing 100871, China d School of Life Science and Technology, Xi’an Jiaotong University, Xi’an 710049, China † Electronic supplementary information (ESI) available: Position of the implant vertically inserted in the rabbit femoral shaft, the fluorescent and histological analyses performed in the region marked by the dotted squares, and surgery procedure schematic of the pillars parallelly implanted in the rBMC; immunofluorescence images of TNF-α, IL-10, and OPN within the bone tissue surrounding the implanted pillars for different times; panoramic histological stained images of bare and coated Mg pillars at implantation time of 12 weeks; the residual volumes and micro-CT images of bare Mg and PEO0- and HAT-coated Mg vertically implanted in the rabbit femora for different times. See DOI: 10.1039/ c8bm00901e

This journal is © The Royal Society of Chemistry 2018

Introduction

Magnesium (Mg) and its alloys provide an attractive alternative for use as temporary orthopedic implants due to their degradability in the physiological environment without requiring additional surgeries for removal after healing, which is the case with nondegradable implants. However, the rapid degradation and consequent loss of mechanical integrity restricts their clinical usage.1 Furthermore, Mg-degradation-induced hydrogen accumulation and strong alkalization of the surrounding body fluids would impair osteogenesis and induce the failure of osseointegration.2 To overcome these demerits, various improved strategies have been proposed,2–4 where surface-modification-derived coatings have shown promising potential by means of slowing down the degradation of the underlying Mg alloys, as well as simultaneously promoting osteogenesis, and therefore, osseointegration.5–9 To be more effective in orthopedic appli-

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cations, besides the abilities of substrate protection and osteogenesis, the coatings are required to be degradable as nonbiodegradable coatings can remain and peel off when the underlying Mg alloys partially or completely degrade in vivo, posing the risk of inducing inflammation.10 Typically, the currently developed coatings on Mg alloys playing the aforementioned three kinds of roles are MgO/poly (L-lactic acid) (PLA) composite coating,5 Ca–P layer,6 zoledronic-acid-loaded PLA/brushite coating,7 hydroxyapatite (HA) and bone morphogenetic protein-2 containing polydopamine coating,8 and Mg–Allayered double hydroxide coating.9 Among these coatings, micro-arc oxidation (MAO)-derived MgO coating on Mg alloy showed significantly higher protective efficacy11 and firmly adhered to the substrates, avoiding peeling off. However, MAOderived coatings are inherently porous, owing to the inherent spark-discharging feature of MAO,12 which seriously weakens their resistance against corrosion.13,14 Although TiO2 and SiO2 nanoparticles were used to reduce the porosity,15,16 the sealing agents are nonbiodegradable; moreover, biodegradable polymeric molecules used as sealants also failed to provide effective protection due to their insufficient thicknesses to fill the pores in the MAO coating.17 The main inorganic mineral in bone, hydroxyapatite (HA), particularly with 3-dimensional (3D) nanorod-patterned topography, has shown an enhanced role in promoting osteogenesis-related cell (osteoblasts and mesenchymal stem cells (MSCs)) functions in vitro and osseointegration in vivo as compared to 2D-patterned HA and other inorganic components.18–20 Moreover, the slow in vivo degradability is also the reason for choosing HA as the sealant in MAO-derived porous MgO coating.21 Actually, bone formation on implants can be divided into distance osteogenesis and contact osteogenesis; the latter is crucial for osseointegration.22 Contact osteogenesis begins with macrophage-mediated inflammatory response. Following the in vivo implantation of a material, macrophages as a kind of immune cells, which usually reveals two polarized phenotypes, M1 and M2, were recruited to the implant surface from the surrounding tissue to initiate inflammation.23 M1 macrophages secrete cytokines such as interleukin (IL)-1β, IL-6, IL-8, and tumor necrosis factor (TNF)-α to cause inflammation, leading to the destruction of adjacent bone and the formation of fibrous tissues. M2 macrophages produce antiinflammatory cytokines such as IL-1Ra, IL-4, and IL-10 to resolve inflammation and promote bone regeneration24 and can also secrete growth factors to support migration, homing, and osteoblastic differentiation of MSCs.23 In addition, macrophages are able to differentiate into multinucleate osteoclasts, inducing bone resorption. Following the inflammatory response, the recruitment of osteogenesis-related cells to the implant surface, differentiation, and extracellular matrix mineralization of the cells occur sequentially to induce contact osteogenesis.25,26 Recently, some works involving surfacemodified Ti alloys and bioactive ceramics clearly demonstrated that tuning the surface architecture23 and chemical compositions27,28 of materials can modulate the phenotypes of macrophages (revealing M1 or M2) and osteoclastogenesis, as

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well as the inflammatory response, i.e., the immunomodulatory effect, while the osteoimmunomodulatory effect that favors in vivo osteogenesis and osseointegration are expected.23,27,28 Focusing on bare and coated Mg alloys, unfortunately, the in vitro investigations regarding their osteoimmunomodulatory effects are fairly limited, although a recent work showed that coating β-Ca3(PO4)2 on a Mg scaffold could more effectively switch the macrophages to the M2 phenotype, inhibiting the pro-inflammatory response and promoting osteoblastic differentiation of MSCs.29 Moreover, current in vivo reports on coated Mg-alloy-induced inflammatory response are confined to the appearance of inflammatory cells or fibrous capsules at the bone–implants interfaces8,9,30 and the lack of a careful evaluation of the coated-Mg-alloy-modulated switch of the macrophage phenotype. Notably, current studies on the osseointegration of coated Mg alloys have mainly focused on new bone formation,6–9 while the interfacial structure and mechanism by which the coated Mg alloys bond to the bone, which is vital for osseointegration, have been rarely studied. On the other hand, the in vivo structural integrity of the coatings on Mg alloys over time is a major concern. An earlier study has shown that in vivo Mg2F coatings vanished within 4 weeks.31 The bilayer coating on the Mg alloy, comprising electrodeposited HA and MAO MgO disappeared after in vivo implantation for 18 weeks, in which HA vanished within 12 weeks after surgery.32 Actually, the vanishing of the coatings on Mg alloys in vivo can be attributed to either their degradation or delamination from the substrates into nanoparticles or fragments. The latter would simulate the release of inflammatory cytokines from the macrophages, resulting in inflammation and osteolysis.10 Previous works have mainly reported the protective efficacy of the coatings to Mg-based implants;5–9 however, the vanishing of these coatings is not clearly explored. In the present work, we fabricated a bilayer-structured coating on Mg, consisting of an outer layer of nanorodpatterned HA and an inner layer of MgO with Mg(OH)2 nanoplates/HA nanorods sealing the pores. The osteoimmunomodulation of the coated Mg and contact osteogenesis process on the coated Mg within the early stage of implantation, as well as the bone–implant interfacial structure evolution of the coated Mg during implantation period, were evaluated in the rabbit femora together with the pore-unsealed MgO coating and bare Mg. The evolution of mechanical integrities of the coated and bare pillars in rabbit femoral bone marrow cavities (rBMCs) was also investigated.

2. Materials and methods 2.1. Plasma electrolytic oxidation and hydrothermal treatment ∅ 14 × 4 mm pure Mg discs, as well as ∅ 2.5 × 10 mm and ∅ 3.2 × 12 mm pure Mg pillars, were employed as the anode and stainless steel plate was used as the cathode in an aqueous electrolyte containing β-glycerophosphate disodium, Ca(OH)2,


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and NaOH (0.02 M, 0.135 M, and 0.125 M, respectively) to receive sequential plasma electrolytic oxidation (PEO). The PEO process lasts for 10 min at 450 V positive pulse voltage, 100 Hz pulse frequency, and 26% duty ratio. The obtained coating was assigned as PEO0 coating and the substrates with PEO0 coatings were termed as PEO0-coated Mg. Subsequently, PEO0-coated Mg was subjected to hydrothermal treatment (HT) in a 52 mL volumetric Teflon-lined autoclave. An aqueous solution containing 0.1 M C10H12CaN2Na2O8 (Ca-EDTA) and 0.5 M NaOH with a volume of 7.8 mL was added into the autoclave to immerse the PEO0coated Mg and HT at 90 °C for 24 h. The obtained coating was assigned as HAT coating, and the substrate with the coating was named as HAT-coated Mg. 2.2.

Characterizing microstructures of PEO0 and HAT coatings

A field-emission scanning electron microscope (FE-SEM; FEI QUANTA 600F) with an energy-dispersive X-ray spectrometer (EDX) was used to examine the morphologies and chemical compositions of the coatings. An X-ray diffractometer (XRD; X’Pert PRO, Netherlands) was carried out to identify the phase compositions using Cu-Kα (λ = 0.15406 nm) radiation over a 2θ angle of 20–63° at a step of 0.02°. A nanoplate and nanorod were scratched from the HAT coating and investigated using a transmission electron microscope (TEM; JEOL JEM-2000FX, Japan) with EDX. 2.3. Cell culture and in vitro assessment of mineralizationinduced proteins MSCs were harvested from 1-week-old New Zealand rabbits. Briefly, the bone marrow was aspirated from the femora and tibias, from which the mononucleated cells were isolated via density gradient centrifugation. The obtained MSCs were plated in the culture medium that consisted of Dulbecco’s modified Eagle medium (DMEM; HyClone) with 10% fetal bovine serum (FBS; HyClone) and 1% antibiotics and cultured at 37 °C in a humidified atmosphere of 5% CO2 and 95% air. Non-adherent MSCs were removed and the adherent MSCs were collected for further expansion. In parallel, the human fetal osteoblastic cell line, hFOB 1.19, was purchased and injected into the culture medium (DMEM with 10% FBS, 0.3 mg mL−1 Geneticine-418 (Sigma), 0.5 mM sodium pyruvate (Sigma), and 1.2 mg L−1 Na2CO3) and cultured in a humidified atmosphere incubator with 5% CO2 at 37 °C. The medium used for the culture of the MSCs and hFOB 1.19 cells were refreshed every 2 days throughout the culture duration. Fluorescence staining assay of the mineralization-induced proteins of bone sialoprotein (BSP) and osteopontin (OPN), early secreted by rabbit MSCs and hFOB 1.19 cells onto PEO0and HAT-coated Mg discs, was performed as follows. The discs were centrally placed in 24-well plates and then seeded with hFOB 1.19 cells and rabbit MSCs at densities of 8 × 104 and 2 × 104 cells per well, respectively. After 4, 24, and 72 h of culture, the cell-seeded-coated Mg was fixed in 4% paraformaldehyde for 10 min and then incubated in a blocking medium containing 1% bovine serum albumin (Sigma) for 30 min at 37 °C.


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Rabbit polyclonal anti-human BSP (Abcam)/mouse monoclonal (MM) anti-rabbit BSP (Immundiagnostik AG) and MM anti-human OPN (Abcam)/MM anti-rabbit OPN (Novusbio) were correspondingly added into the hFOB 1.19 cells/rabbit MSCs-seeded coatings and incubated at 37 °C for 2 h; then, 10 μg mL−1 second antibodies, rhodamine-conjugated goat anti-rabbit IgG (Chemicon International), or FITC-conjugated goat anti-mouse IgG (Chemicon International) were correspondingly added to the coatings seeded by the abovementioned cells and incubated at 37 °C for 1 h. Following every staining event, the discs were washed five times with PBS. The stained BSP and OPN secreted by the cells were pictured using an OLYMPUS laser confocal microscope (FV1000). For convenient observation and comparison, a pseudo-color of red was added to the captured images of BSP secreted by rabbit MSCs using the FV10-ASW (viewer 3.1) software. 2.4.

Animal experiments

2.4.1 Animals and surgery. All the animal experiments were conducted according to the ISO 10993-2: 2006 animal welfare requirements, and all the procedures in the animal experiments were approved by the animal research committee of Xi’an Jiaotong University. Adult male New Zealand rabbits (male, ∼2.5 kg) were employed to evaluate the osseointegration and mechanical integrities of the coated and bare Mg pillars. The rabbits were anesthetized by an intramuscular injection of 2.5 wt% pentobarbital sodium solution. For osseointegration evaluation, pillars with a size of ∅ 2.5 × 10 mm were employed, and each rabbit was implanted one kind of pillar, as schematically shown in Fig. S1a.† Briefly, two holes (∅ 2.5 mm) were drilled in the right and left femoral shafts of each rabbit; then, the pillars were inserted, keeping the pillars vertical to the long axis of the femur, followed by suturing the muscle, subcutaneous tissue, and skin. For determining the mechanical integrity during the implantation period, pillars with a size of ∅ 3.2 × 12 mm were employed, and each of them was parallelly inserted into the bone marrow cavity of the femur through a pre-drilled hole (∅ 3.2 mm) on the femur, as schematically shown in Fig. S1b,† followed by suturing of the muscle, subcutaneous tissue, and skin. In both the femoral shafts of the rabbit, the implanted pillars were of the same kind. Postoperatively, the rabbits received brizolina (antibiotics) intramuscular injection for three days and were allowed to move freely in their cages. 2.4.2 Histological examinations and sequential fluorescent labeling assays. After 4, 8, and 12 weeks of surgery, the rabbits were sacrificed. The femora containing the vertically implanted pillars (n = 4 for each group) were fixed in formalin, dehydrated by ethanol with ascending concentrations, and polymethyl methacrylate (PMMA) were used for embedding. The embedded samples were cut into sections (∼150 μm thickness) using a microtome (Leitz 1600); the sections were ground, polished to foils with a thickness of 20 μm, and subsequently stained with Van Gieson’s picric–fuchsine for histological analysis. The sections were observed by a Olympus IX 71 microscope and analyzed using Image-Pro Plus software. The ratio

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of the bone–implant contact was evaluated on four sections of each implant. The bone–implant interfaces were examined using FE-SEM equipped with EDX. In addition, a polychrome sequential fluorescent method was employed to label new bone formation over time. At weeks 2, 4, and 6 of the implantation, three types of fluorochromes were intramuscularly injected into the pillar-implanted rabbits in a sequence of 25 mg kg−1 tetracycline hydrochloride (TE; Sigma), 30 mg kg−1 Alizarin Red S (AL; Sigma), and 20 mg kg−1 calcein (CA; Sigma), respectively. The rabbits were sacrificed at week 8; the fluorescent foils were prepared using the same procedure as that for the aforementioned histological examination, but without using Van Gieson’s picric–fuchsine staining. Then, the foils were observed under confocal laser scanning microscope (CLSM, OLYMPUS FV1000) with excitation wavelengths of 405/560–590 nm for TE (yellow), 543/580–670 nm for AL (red), and 488/500–550 nm for CA (green).33 2.4.3 Push-out tests and observation of the pushed-out surface morphologies. Push-out tests were used to evaluate the bone–implant integration strength at days 3, 7, and 14, as well as weeks 4, 8, and 12 post-implantation (n = 8 for each group at each time point). Each of the vertically implanted pillars in the femora was mounted on a device. The device was accurately aligned with the pillar having a perpendicular load transfer. Thereafter, the testing appliance pushed the pillar vertically out at 1 mm min−1. The load vs. displacement curve for each pillar was recorded, from which the push-out force was obtained. In parallel, the pushed-out PEO0- and HATcoated Mg pillars, which were implanted for 3, 7, and 14 days, as well as the pushed-out bare Mg pillars implanted for 14 days, were fixed in 4% paraformaldehyde at room temperature, followed by immersion in a solution of 3% sodium hypochlorite for 15 min. Subsequently, the pillars were washed with running water, dehydrated in serial concentrations of ethanol (50, 70, 95, 100, 100% v/v) for 20 min for each instance and vacuum-dried. Thereafter, the pushed-out surfaces on the sides of the pillars were examined using Raman-SEM (TESCAN-RISE, Czech Republic) with an approximately 1 μmdiameter electron beam spot to reveal the initial contact osteogenesis procedure and bone–implant bonding mechanism. 2.4.4 Real-time polymerase chain reaction (RT-PCR) analysis of in vivo inflammation-, osteoclastogenesis-, and osteogenesis-related gene expressions. The evaluations of inflammation-, osteogenesis-, and osteoclastogenesis-related gene expressions within the bone surrounding bare and coated Mg pillars vertically implanted in the rabbit femora for 3, 7, and 14 days (n = 8 per group at each time point) were performed according to the method described elsewhere.34 Briefly, the rabbits were anesthetized, followed by an operation to expose the implants. Following the pulling-out of the implants, their surrounding bone tissues were carefully retrieved using a stainless steel trephine bur (inner diameter: 3 mm); then, they were immediately frozen in liquid nitrogen and maintained at −80 °C. Prior to the RT-PCR analysis, the retrieved bone cores were placed in liquid nitrogen and grounded into a powder with a pestle. The total RNA from each of the pulverized bone

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powders was extracted using TRIZOL reagent (Invitrogen, Carlsbad, CA) according to the manufacturer’s protocol and then quantified. The total RNA from each of the peri-implant bone tissues was increased by 5 μg and reverse-transcribed into complementary DNA using a Prime-Scrip RT reagent kit (Takara, Japan). The expressions of the inflammation-related genes (such as IL-1β, TNF-α, and IL-10), osteoclastogenesis-related gene (such as tartrate-resistant acid phosphatase (TRAP)), and osteogenesis-related genes (such as alkaline phosphatase (ALP), BSP, OPN, and osteocalcin (OCN)) were quantified using a RT-PCR detection system (Roche Lightcycler 96) with SYBRsPremix ExtTaqII (Takara, Japan). Data analysis was carried out using the Lightcycler 96 instrument software 1.1 (Roche, Switzerland). The housekeeping gene, glyceraldehyde3-phosphate dehydrogenase (GAPDH), was used as an endogenous reference to normalize the calculation through the comparative Ct value method. The sequences of the specific primer sets were listed as follows: IL-1β (5′CCACAGTGGCAATGAAAATGA3′ and 5′ACCTGCCGGAAGCTCTTGTT3′), TNF-α (5′TGCTGCACTTCAGGGTGATC3′ and 5′CTTGCGGGTTTGCTACTACGT3′), IL-10 (5′AGAACCACAGTCCAGCCATCA3′ and 5′GCTTGCTGAAGGCGCTCTT3′), TRAP (5′GTCTCAGCCCAGATCGCCTAT3′ and 5′GTCGTTTGAGTTGCCACACAA3′), ALP (5′ CAGCAGCACACCAGCTGTGT3′ and 5′TGTTGTGCGCGGTACCAAT3′), OPN (5′TCACGAGCAGGCCAGACA3′ and 5′TCGGCTCGATGGCTAGCTT3′), OCN (5′ACGCACAGAGCGACAGCAT3′ and 5′ CTCTTGGACACGAAGGCTGA3′), BSP (5′GGGAGTACGAACAAACAGGTAACC3′ and 5′GACCCTCGTAGCCCTCATAGC3′), and GAPDH (5′ATCAAGTGGGGTGATGCTGG3′ and 5′TATTCTCGTGGTTCACGCC3′). 2.4.5 Micro-CT assays and mechanical integrities of the coated and bare Mg pillars implanted in rBMCs. Micro-CT assays were used to evaluate the corrosion rates of the coated and bare Mg pillars parallelly implanted in the rBMCs for different times. The rabbits with the pillars were sacrificed at weeks 4, 8, and 12 post-implantation; then, the femora containing the pillars were collected and scanned by a Micro-CT device (eXplore Locus SP, GE company, USA) for 3D reconstruction (working voltage: 80 kV; current: 80 μA). The reconstructed images and residual volumes of the implants at each time point were obtained using the rebuilding software in the Micro-CT system (Micview, USA). The corrosion rates (CR) of the pillars in the bone marrow were calculated according to the following equation: CR ¼ ðV o V r Þ=ti

ð1Þ

where Vo is the original volume of the pillars before implantation, Vr is the residual volume after implantation obtained from Micro-CT reconstruction, and ti is the implantation time. After Micro-CT examination, the pillars were pushed-out from the bone marrow cavities; thereafter, the following threepoint bending tests were carried out to identify their mechanical integrities. By employing mechanical test equipment (Instron 5500, USA), each sample was centrally located on two


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separated supports with a distance of 6 mm from the testing machine. A crosshead in contact with the surface center of the pillars moves downward at 0.5 mm min−1, bending it to failure at an applied load. The load was designed as the maximum bending force and the obtained values were averaged (n = 4 for each group at each time point).


2.5.

Statistical analysis

The data were expressed as mean ± standard deviation (SD) from repeated independent experiments. The data were analyzed using SPSS 16.0 software (SPSS, USA). A one-way ANOVA followed by a least-significant-difference (LSD) post hoc test was used to determine the level of significance. Here, p < 0.05 and 0.01 were considered to be significant and highly significant, respectively.

3. Results 3.1.

Microstructures of PEO0 and HAT coatings

Fig. 1 shows the microstructural features of PEO0 and HAT coatings on Mg. The PEO0 coating was microporous and the

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2–3 μm-diameter pores were homogeneously distributed on its surface (left image in Fig. 1a). The thickness of the coating was about 15 μm, as identified by the Mg, Ca, and P distributions detected by EDX on its cross-section; some micro-arc-discharge-derived micropores were distributed throughout the cross-section (right image in Fig. 1a), which may act as channels for body fluid permeation, and consequently, weaken the ability of the coating to protect Mg from corrosion. In addition, both surface and cross-sectional views exhibited a denser structure of the coating matrix, which consisted of MgO (Fig. 1c). Moreover, no discontinuity was observed at the coating–Mg substrate interface. With regard to HAT coating (i.e., the one resulting from HT of PEO0 for 24 h), its surface was fully covered with nanorods, which appeared to directly grow on the MgO matrix with a mean diameter of 90 ± 5.6 nm and interrod spacing of 65.2 ± 5.3 nm (Fig. 1b). Notably, the nanorods also grew on the pores’ walls to seal them, as proven by the ellipse-marked region in the enlarged inset in the right image of Fig. 1b. In addition, nanoplates appeared among the nanorods on the dense surface and pores’ walls of MgO, further sealing the pores. As identified by XRD (Fig. 1c) and TEM (Fig. 1d and e), the nanoplates and nanorods comprised

Fig. 1 FE-SEM surface and cross-sectional morphologies of (a) PEO0 and (b) HAT coatings; (c) XRD patterns detected on PEO0 and HAT coatings; TEM bright-field images of the scratched (d) nanoplate and (e) nanorod from the HAT coating; the insets show the SAED patterns and EDX spectra from the dotted-circle marked areas.


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well-crystallized Mg(OH)2 and HA, respectively. From the EDXdetected distribution of Ca, P, and Mg, it can be concluded that the bilayer-structured HAT coating comprised a HA nanorod-topographical (shortened as HAT) outer layer and an inner layer of MgO with HA nanorods/Mg(OH)2 nanoplates, sealing the pores; moreover, the HA nanorods were embedded in the MgO matrix rather than the Mg(OH)2 nanoplates, ensuring the fixation of the HA nanorods. 3.2. Early secretion of non-collagenous proteins by MSCs and osteoblasts on coated Mg A hypothesis states that contact osteogenesis on a Ti-based implant in vivo originates from the secretion of non-collagenous proteins such as BSP and OPN by osteogenesis-related cells onto the implant, which further induces mineralization to form a cement line matrix, bonding the implant to the new bone.26 To test the availability of the hypothesis for Mg-based implants, fluorescence staining of the non-collagenous proteins secreted by MSCs and osteoblasts in vitro onto HAT- and PEO0-coated Mg was performed at the culture times of 4, 24, and 72 h, as shown in Fig. 2. The MSCs-secreted BSP and OPN tended to increase in amount with incubation time on both the coated Mg. Notably, as early as 4 h, a heap of BSP and a small amount of OPN appeared only on HAT-coated Mg and not on PEO0-coated Mg; HAT-coated Mg promoted the secretion of BSP and OPN at the time points of 24 and 72 h each as compared to PEO0-coated Mg (Fig. 2a). For osteoblasts, their secreted non-collagenous proteins on both the coated Mg displayed similar variation trends to those of MSCs (Fig. 2b), and they were slightly more in amount than those in MSCs at each time point. 3.3.

Osteoimmunomodulation of the coated and bare Mg pillars

Following the endosseous implantation of a material, the immune response plays a vital role in osteogenesis and osseointegration.25 To investigate the immunomodulatory effects of coated and bare Mg pillars, the inflammatory and osteoclastogenesis- and osteogenesis-related genes of the pillars surrounding the bone during the early period of 14 days were evaluated (Fig. 3). At each time point of 1, 3, 7, and 14 days, the macrophage-secreted pro-inflammatory cytokines TNF-α and IL-1β were downregulated in gene expression by HAT-coated Mg as compared to PEO0-coated Mg and, more significantly, as compared to bare Mg (Fig. 3a); simultaneously, macrophage-secreted anti-inflammatory cytokine IL-10 was highly expressed in the gene level for HAT-coated Mg as compared to PEO0-coated Mg and bare Mg (Fig. 3b). Moreover, the variation trend of TNF-α and IL-10 expressions at the protein level in the bone tissues surrounding bare, PEO0-, and HAT-coated Mg is similar to that of their corresponding genes according to immunofluorescence staining results at 3 and 7 implantation days (Fig. S2†). Further, based on the variations in the expressions of TNF-α and IL-10 both at the gene and protein levels with time, particularly the highest levels of TNF-α for bare Mg and IL-10 for HAT-coated Mg at day 7, it is indicated that the lasting duration of inflammation

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was shortened in the case of HAT-coated Mg, and the macrophages around HAT-coated Mg switched quickly from proinflammatory phenotype (M1) to anti-inflammatory phenotype (M2). An earlier study has reported that nanosized HA nanoparticles would induce a high level of pro-inflammatory cytokine secretion of macrophages and impair tissue remolding.35 However, our previous and present works have shown that both nanograined and HA nanorod-patterned surfaces can enhance the osteogenetic differentiation of osteoblasts18,36 and quickly switch the macrophages into the M2 phenotype. The conflicting results may be attributed to the adhesion of the macrophages on the nanograined and nanorod-patterned surfaces rather than the uptaken nanoparticles. TRAP is a marker of the osteoclastic differentiation of macrophages and highly expressed by osteoclasts.37 The gene expression of TRAP was significantly downregulated by HAT-coated Mg as compared to bare and PEO0-coated Mg at each implantation time point (Fig. 3c), implying the suppression of osteoclastogenesis and subsequent bone resorption.38 For the osteogenesisrelated markers detected as ALP, BSP, OPN, and OCN (Fig. 3d), coated Mg was observed to upregulate its mRNA levels with implantation from 1 to 14 days; however, no obvious changes in their expressions were observed around bare Mg. At each time point, the gene expressions of ALP, BSP, OPN, and OCN around bare and coated Mg pillars followed the rank of HATcoated Mg > PEO0-coated Mg > bare Mg, suggesting that HATcoated Mg could greatly promote in vivo osteogenesis as compared to bare and PEO0-coated Mg during the early period of implantation. 3.4. In vivo contact osteogenesis and osseointegration of coated and bare Mg 3.4.1 Morphological observation. Following the implantation of coated and bare Mg pillars in the rabbit femora for 4 to 12 weeks, the rectangle-marked region on each of the pillars (Fig. S1†) was imaged to evaluate new bone formation. This formation includes distance osteogenesis and contact osteogenesis. In distance osteogenesis, new bone is formed on the surface of the old bone in the peri-implant site to encroach upon the implant, leading to the bone approximating the implant surface; however, contact osteogenesis includes the recruitment and migration of osteogenic cells and bone formation by those cells, resulting in bone apposition to the implant surface.39 Based on the definition and the fact that the aforementioned rectangle-marked region is adjacent to the cancellous bone (but still in the bone marrow cavity), it can be deduced that in this region, the image characterizes contact osteogenesis rather than distance osteogenesis. As shown in Fig. 4a (which is pictured following the polychrome sequential fluorescent labeling and statistical result (Fig. 4b)), the amounts of newly mineralized bone matrix labeled with TE at week 2 (yellow area), AL at week 4 (red area), and CA at week 6 (green area) were more on HAT-coated Mg than PEO0-coated Mg. Notably, almost no newly mineralized bone appeared on PEO0-coated and bare Mg within 2 weeks. It is indicated that HAT-coated Mg significantly promoted new bone formation as


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Fig. 2 Fluorescent staining images of mineralization-induced non-collagenous proteins secreted by (a) MSCs and (b) osteoblasts cultured on PEO0- and HAT-coated Mg for 4, 24, and 72 h.

compared to bare and PEO0-coated Mg at various stages within 8 weeks. The histological stained images (Fig. 4c) showed new bone to be formed and increased on all the pillars without any intervening fibrous layers along with prolonging the implantation time at least up to 8 weeks. However, the new bone amounts and bone–implant contact ratios (Fig. 4d) are fairly different, showing the highest values on HAT-coated Mg and the lowest values on bare Mg. Meanwhile, the compactness of the new bone formed on bare and coated Mg increases


with implantation time within 8 weeks, but the bone formed on HAT-coated Mg shows the densest structure as compared to that on bare and PEO0-coated Mg. Noticeably, at week 12, the pillars’ degradation-derived gaps appear between the newly formed bone and the pillars, leading to a decrease in the bone–implant contacts for the pillars as compared to those at week 8. Further, there was no connection between the new bone and bare Mg, whereas HAT-coated Mg maintained the highest bone–implant contact ratio. Moreover, due to the fast degradation of HAT-coated Mg from week 8 to

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Fig. 3 Gene expressions of the markers in bone tissues around bare and coated Mg: (a) pro-inflammatory cytokines TNF-α and IL-1β; (b) antiinflammatory cytokine IL-10; (c) osteoclastogenic marker TRAP; and (d) osteogenic markers ALP, BSP, OPN, and OCN. Data are presented as mean ± SD, n = 8. (*) p < 0.05 and (**) p < 0.01 when compared with bare Mg and (#) p < 0.05 and (##) p < 0.01 when compared with PEO0-coated Mg.

week 12, the new bone surrounding the implant would not form as compact as that at week 8, but it was still much denser than that on PEO0-coated Mg. In addition, the original cortical bones around the bare and coated pillars exhibit no destruction or osteolysis even at week 12 post-surgery (Fig. S3†), indicating that the corrosion products of bare and

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coated Mg do not induce any adverse effects on the localized tissues. Fig. 5 shows the cross-sectional FE-SEM morphologies at the interfaces of new bone and bare and coated Mg vertically implanted in the rabbit femora for 4 and 12 weeks, respectively. Here, the new bone, coating, and Mg substrate on each


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Fig. 4 New bone formation around bare and PEO0- and HAT-coated Mg pillars implanted in rabbit femora over time: (a) polychrome sequential fluorescent labeling images; (b) corresponding fluorochrome areas; (c) histological stained images at implantation times of 4, 8, and 12 weeks (NB: new bone; BM: bone marrow); and (d) corresponding bone–implant contacts. Data are presented as mean ± SD, n = 4. (*) p < 0.05 and (**) p < 0.01 when compared with bare Mg and (#) p < 0.05 and (##) p < 0.01 when compared with PEO0-coated Mg.

bone–implant cross-section were distinguished by EDXdetected elemental distribution. At week 4, even at high magnifications, HAT coating still exhibited a tight connection with the new bone at a major area of the coating–bone interface (Fig. 5a3). However, a few intervals were observed at the PEO0 coating–new bone interface (Fig. 5a2), indicating a weaker osseointegration ability of PEO0 coating. Even so, the PEO0coated pillar still exhibited enhanced osseointegration than bare Mg, which induced a large area of intervals at the implant–new bone interface (Fig. 5a1). At week 12, bare Mg pillars exhibited severe pitting corrosion, exhibiting an irregular outline, which was closely surrounded by the corrosion product, Mg(OH)2, sequentially followed by bone marrow and bone trabecula, as identified by the EDX-detected distribution maps of Mg, O, Ca, and P (Fig. 5b1). PEO0-coated Mg showed a more regular shape than bare Mg, but Mg(OH)2 was also observed around Mg, as identified by EDX-detected distribution maps of Mg and O. Although degradation-derived gaps appeared at the bone–implant interface, a few connective regions still existed. The details of the connective regions are shown in the magnified image of site A, where Mg(OH)2 overlying on Mg directly connected with the bone trabecula and no osteolysis-derived gaps could be observed at the interface. For HAT-coated Mg (Fig. 5b3), most of the bone– coating interfacial area displayed a tight contact without any trace of osteolysis, as displayed in the magnified image of site C with the aid of EDX profiles. However, a few Mg(OH)2 regions (e.g., site B) appeared between Mg and the newly formed bone, and Mg(OH)2 tightly connected with the bone without any trace of osteolysis at the Mg(OH)2–bone interface, as identified by the distribution maps of Mg and O (Fig. 5b3)


and EDX profiles shown in the magnified image of site B. Notably, the Mg(OH)2 overlying on Mg in sites A and B could be derived from the corrosion of Mg and/or the conversion of MgO in PEO0 and HAT coatings. 3.4.2 Push-out forces and components of push-out disrupted surfaces. As shown in Fig. 6a, the push-out forces that characterize the fixations of bare and coated pillars with bone, as measured by the push-out tests, increased with the implantation time from 3 days to 8 weeks, following the rank of HAT > PEO0 > bare Mg. Then, the push-out forces of bare and coated pillars decreased with time to 12 weeks due to the formation of degradation-derived gaps between the bone and pillars (Fig. 4c and 5b), but the HATcoated Mg still exhibited significantly higher push-out force than the other two implants. This result reveals that HATcoated Mg could enhance the fixation with bone, most notably at an earlier stage after surgery. Meanwhile, it is noteworthy that due to the yet decreased push-out forces of bare and coated Mg at week 12, the bone–implant interfacial failure, particularly for HAT-coated Mg, would be investigated in our future work for a longer implantation period (more than 12 weeks). The pushed-out surface on the side of the bare Mg pillar implanted for 14 days were observed (Fig. 6b), revealing that the plain regions (Fig. 6b2 as an example) of the surface were covered with the corrosion product Mg(OH)2 without any bone matrix, as identified by the EDX-detected high contents of Mg and O, but the absence of Ca and P (Fig. 6b4). Moreover, some corrosion pits (Fig. 6b3 as an example) were observed on the surface, surrounded by Mg(OH)2, as identified by EDX. It is indicated that even after 14 days of implantation, no new bone

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Fig. 5 Cross-sectional FE-SEM morphologies at the bone–implant interfaces vertically implanted in the rabbit femora for (a) 4 and (b) 12 weeks (a1 and b1 for bare Mg, a2 and b2 for PEO0-coated Mg, and a3 and b3 for HAT-coated Mg); the distribution maps of Mg, O, P, and Ca elements detected with EDX over (b1)–(b3), and the magnified images of the dotted-squares-marked sites A, B, and C in corresponding (b2) and (b3). (NB: new bone; BM: bone marrow).

Fig. 6 (a) Push-out forces of bare Mg and PEO0- and HAT-coated Mg pillars vertically implanted in the rabbit femora at 3, 7, and 14 days, as well as 4, 8, and 12 weeks; (b) FE-SEM analysis of the pushed-out surface on the side of the bare Mg pillar implanted for 14 days, where (b1) is the panoramic view and (b2) and (b3) are the magnified views of the dotted squares in (b1); (b4) lists the elemental contents detected within the Arabic numbered micro-regions ① and ② in images (b2) and (b3). Data are presented as mean ± SD, n = 8. (*) p < 0.05 and (**) p < 0.01 when compared with bare Mg and (#) p < 0.05 and (##) p < 0.01 when compared with PEO0-coated Mg.

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matrix formed on the bare Mg pillar, which is in good agreement with the sequential fluorescent labeling results (Fig. 4a). To clarify the mechanism underlying the bonding of coated Mg onto the bone, the pushed-out surfaces on the sides of PEO0- and HAT-coated Mg pillars implanted for 3 to 28 days, obtained from the push-out tests, were carefully examined with FE-SEM, EDX, and in situ Raman analyses. For the implanted PEO0-coated Mg, no changes in the morphology could be observed on the coating at day 3 as compared to the as-received one (Fig. 7a2 vs. 1a), and no other Raman peaks were detected besides the peaks at 3220 cm−1 assigned to O–H in water40 and at 2160 cm−1 assigned to absorbed C–O in CO2.41 This is in good agreement with the immunofluorescence staining results in Fig. S4,† which shows that no mineralization-induced OPN was observed surrounding the PEO0coated Mg. At day 7, a few globules appeared on the microporous MgO surface (Fig. 7b). They were richened with Ca and P as identified by EDX and contained OPN and phosphate (PO43−) as identified by the Raman spectrum, which reveals the peak at 1464 cm−1 (caused by CH2 scissoring or CH3 asymmetric stretching) intrinsically attributed to OPN42 and the peaks at 441 and 608 cm−1 assigned to PO43−.40 Meanwhile, the appearance of OPN on the PEO0-coated Mg was further confirmed by the result in which a spot of OPN distributes in close proximity to the pillar in the immunofluorescence staining image at day 7 (Fig. S4a†). It is indicated that the globules consisted of OPN and mineralized apatite, with chemical components fairly similar to the cement line matrix (comprising non-collagenous proteins and their induced mineralized apatite) between the new and old bones at the site of bone remodeling,22,43 suggesting the initial formation of a cement line matrix on the microporous MgO surface at day 7. Notably, the cement line matrix intimately juxtaposed to the walls of the micropores (Fig. 7b2). At day 14, fibrous clusters appeared to overlay on the collected globules (i.e., cement line matrix) (Fig. 7c), which consisted of collagen (identified from the Raman spectrum) that exhibits the peaks at 2940 cm−1 (caused by CH2 and CH3 amino acid side chains stretching) and 1670 cm−1 (caused by amide I) intrinsically attributed to collagen.40 At day 28, the disrupted surface reveals a construction of fibrous clusters wrapped and filled with minerals (Fig. 7d) composed of collagen and apatite, as identified from the Raman spectrum, which reveals a peak at 2940 cm−1 ascribed to collagen and peaks at 448, 598, and 958 cm−1 assigned to PO43−,40 suggesting that the collagen fibers induced the mineralization of apatite. Moreover, the increase in the Ca and P contents of the corresponding constructions (Fig. 7e) further proves the progress in apatite mineralization over time. It is highlighted that the collagen fibers further induced apatite mineralization to form the bone matrix over time. For HAT-coated Mg, in which the outer layer of the coating was patterned with HA nanorods, at day 3 of the implantation, the HA nanorods were wrapped and filled, even partially overlaid with the newly formed matrix (Fig. 8a). The matrix contained OPN, as identified by the Raman spectrum that reveals the peak at 1464 cm−1 attributed to OPN,42 and the immuno-


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fluorescence staining results (Fig. S4b†). It also consists of mineralized apatite, as confirmed by the Raman peak at 958 cm−1 assigned to PO43−,40 which may be partially caused by the underlying HA nanorods. Moreover, the amount of secreted OPN on the HAT-coated Mg was more than that on PEO0-coated Mg (Fig. S4†), which was in line with the in vitro result (Fig. 2). It is indicated that the formation of a cementline-like matrix could be promoted by the nanorod-patterned HA as compared to Fig. 7a (microporous MgO). At day 7, the layered and overlapped bone matrix appeared on the pushedout surface of HAT-coated Mg (Fig. 8b1); Fig. 8b2 shows the overlapped sequence. The marked top layer revealed fibrous clusters embedded or wrapped with some particles (Fig. 8b3), which consisted of collagen and its induced mineralized apatite, respectively, as identified by the Raman spectrum that reveals the peak at 2940 cm−1 attributed to collagen and the peak of 958 cm−1 ascribed to PO43−.40 Notably, the collagen fibers are more organized and compact as compared to that formed on PEO0 coating (Fig. 7c2), which would lead to a denser structure and better mechanical resistance of the newly formed bone to promote bone–implant interfacial stability,44 which is in good agreement with the histological staining and push-out tests results (Fig. 4c and 6a). Underlying the top layer, the collagen fibrous clusters were almost fully filled with the mineralized apatite (Fig. 8b4), as further supported by the fact that the Ca and P contents detected within the ③-marked region is significantly higher than those detected within the ②-marked region (Fig. 8d). Underlying the layer constructed with the apatite-filled collagen fibrous clusters is the marked bottom layer, which consisted of a cement line matrix (Fig. 8b5), as identified by the Raman spectrum that reveals the peak at 1464 cm−1 attributed to OPN42 and an increased amount of OPN appeared surrounding the HAT-coated Mg (Fig. S4†). Further, the peak at 958 cm−1 is assigned to PO43−.40 Fig. 8b along with Fig. 8a reveal that the newly formed bone matrix is directly bound to the cement line matrix, which further formed firm interdigitation with the HA nanorods. At day 14, the pushed-out surface on the side of the HAT-coated Mg exhibited a more mature bone matrix composed of collagen (Fig. 8c2) and more mineralized apatite (Fig. 8c3), as identified by the Raman spectrum, which reveals the characteristic peaks of PO43− at 448, 598, and 958 cm−1,40 but without the peaks assigned to collagen, as well as the considerably high Ca and P contents in the apatite (Fig. 8c and d). 3.5. Degradation behavior and mechanical integrities of bare and coated Mg in rBMCs Fig. 9a shows the Micro-CT images, reconstructed with and without the new bone, of the coated and bare pillars parallelly implanted in the rBMCs for 4, 8, and 12 weeks. At each time point, HAT-coated Mg was observed to induce new bone more than PEO0-coated Mg and considerably more than bare Mg. Obviously, new bone on the pillars implanted in this case (far away from cancellous bone) was less than that shown in Fig. 4c, in which the observed rectangle region (Fig. S1a†) is adjacent to the cancellous bone, which can be attributed to

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Fig. 7 Structural analyses of the pushed-out surfaces on the sides of PEO0-coated Mg pillars vertically implanted in the rabbit femora over time. FE-SEM surface morphologies after implantation for (a) 3, (b) 7, (c) 14, and (d) 28 days, where (a1)–(d1) are the panoramic views and (a2)–(d2) are the magnified views of the dotted squares in (a1)–(d1), respectively; Raman spectra detected within the entire (a2) area and Arabic ①, ②, and ③ microregions in (b2)–(d2); (e) listing the EDX-detected elemental contents within the marked regions; (f ) Raman peaks assignment for OPN, collagen and, apatite.

the rapid revascularization in the region adjacent to the cancellous bone.45 Notably, all the coated and bare pillars underwent, at different levels, volume reduction. At each time point, the residual volumes of the pillars followed the order: HATcoated Mg > PEO0-coated Mg > bare Mg (Fig. 9a and b). Meanwhile, the corrosion pits were less on PEO0-coated Mg and far less on HAT-coated Mg as compared to those on bare Mg at week 4, revealing effective corrosion protection of the Mg coatings within the initial implantation period. However, corrosion pits appeared on the coated pillars after 8 weeks of

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implantation and were more severe on PEO0-coated Mg as compared to HAT-coated Mg (Fig. 9a), suggesting higher protective efficacy of HAT coating than PEO0 coating. Consequently, the average corrosion rates of both the coated Mg measured within the period of 8 to 12 weeks dramatically increased as compared to those measured within the period of 0 to 8 weeks (Fig. 9c). Furthermore, the volume reduction of bare and coated Mg pillars vertically implanted in the rabbit femora show a similar variation trend as compared to that parallelly implanted in the rBMCs (Fig. S5†). The results obtained


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Fig. 8 Structural analyses of the pushed-out surfaces on the sides of the HAT-coated Mg pillars vertically implanted in the rabbit femora over time. FE-SEM surface morphologies after implantation for (a) 3, (b) 7, and (c) 14 days, where (a1)–(c1) are the panoramic views and magnified views of the dotted squares in (a1)–(c1) are presented, while (b3)–(b5) are the magnified images of the corresponding dotted squares in (b2); Raman spectra detected within the Arabic ①, ②, ④, and ⑤ micro-regions in images (a)–(c); (d) listing the elemental contents (at%) detected within the ①–⑤ regions.

from different implantation cases confirm the effective protection of HAT coating to the underlying Mg, particularly within the initial implantation period. As previously demonstrated, the pitting corrosion of Mg and its alloys could lead to a premature loss of their mechani-


cal integrities.14 Hence, three-point bending tests were used to evaluate the mechanical integrities of bare and coated Mg pillars parallelly implanted in rBMCs. Fig. 9d shows the variations in the maximum bending forces (BFmax) of the pillars with implantation time. The values were obtained from the

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Fig. 9 (a) Micro-CT images of bare Mg, PEO0-, and HAT-coated Mg (marked in white color) parallelly implanted in rBMCs with or without the periimplant bone (marked in green color) for different times; (b) residual volumes of the pillars; (c) corrosion rates of the pillars implanted for different times; (d) variations in BFmax of bare and coated pillars with time of implantation in rBMCs; (e) representative bending loads–displacement curves; (f ) in vivo percentage reduction rate in the strength or BFmax of the coatings on Mg alloys listed in the literature and our HAT-coated Mg. Data are presented as mean ± SD, n = 4. (*) p < 0.05 and (**) p < 0.01 when compared with bare Mg and (#) p < 0.05 and (##) p < 0.01 when compared with PEO0-coated Mg.

bending loads–displacements curves, as shown in Fig. 9e. Before implantation, the BFmax values of bare and coated Mg were similar, indicating that there was no obvious influence of the coatings on the mechanical strength. Following implantation, the BFmax values of all the pillars decreased, but with different amplitudes. HAT-coated Mg exhibited the lowest decrease in BFmax, indicating that HAT-coated Mg can maintain its mechanical integrity for a longer duration. Some works have investigated the mechanical integrities of coated Mg alloys after in vivo implantation. For instance, it was reported that the MgF2-coated Mg alloy decreased to about 40% in BFmax after implantation in rBMCs for 3 months;46 HA-coated Mg demonstrated a 30% decrease in the tensile strength after 12 weeks of implantation in rat pericranium pouches that have relatively moderate corrosive environments as compared to the bone marrow cavity.47,48 For comparison of the protective effects of HAT coatings with the coatings in the aforementioned literature, Fig. 9f shows the percentage decrease rates in BFmax or strength (i.e., the ratio of the percentage decrease in BFmax or strength and implantation time), indicating that our HAT-coated Mg exhibits higher in vivo mechanical integrity as compared to the aforementioned implants, revealing enhanced corrosion protective efficacy of the bilayer-structured coating for Mg-based implants.

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4.

Discussion

4.1. Contact osteogenesis and osseointegration mechanism of coated Mg J. E. Davies proposed the hypothesis that contact osteogenesis on a Ti-based implant originates from the secretion of non-collagenous proteins by osteogenetic cells onto the implant, which further induces mineralization to form a cement line matrix, bonding the implant to the new bone.26,49 However, when a material was implanted, a macrophage-mediated inflammatory response is immediately initiated.23 Moreover, in the hypothesis, the endosseous peri-implant proteins were not investigated and the identification of the so-called cement line was only based on morphology without any confirmation of the chemical composition.26,49 Therefore, it is needed to elucidate contact osteogenesis and osseointegration mechanism of the coated Mg as an alternative orthopedic implant. It is well known that the osseointegration of implants depends on contact osteogenesis, which originates from the macrophages-mediated inflammatory response; a quick phenotype switch of the macrophages from M1 to M2 at an early stage of implantation would enhance the recruitment of MSCs on the implants and their osteoblastic differentiation.23,24 Herein, following endosseous implantation, HAT-coated Mg


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was shown to considerably downregulate pro-inflammatory TNF-α and IL-1β, upregulate anti-inflammatory IL-10, and suppress osteoclastogenesis as compared to PEO0-coated and bare Mg (Fig. 3a–c and S2†), inducing a quick M1 to M2 phenotype switch of the macrophages around HAT-coated Mg. It can be expected that HAT-coated Mg could modulate the surrounding immune microenvironment toward favoring the recruitment of osteogenetic cells. Our in vitro result confirmed that osteogenetic cells, MSCs, and osteoblasts, could secrete BSP and OPN onto both the coated Mg within a culture time of 3 days (Fig. 2). Given that BSP and OPN initiate the nucleation of apatite,22,43,50,51 it is expected that their continued mineralization forms a cement line matrix. This is supported by our result (as shown in Fig. 7b, 8a, and S4†), where OPN and its induced apatite appeared on both the coated Mg. It was also reported that the non-collagenous proteins in the cement line matrix are rich in OPN and sometimes contain BSP;51 the absence of BSP peaks in the Raman spectra of Fig. 7b and 8a is acceptable and seems reasonable. Given that the half-life is about 4–5 h for BSP and 6 h for OPN,52 this would provide sufficient time to allow the diffusion of the secreted proteins into the space among the HA nanorods of the HAT coating and the micropores of the PEO0 coating, inducing mineralization to form cement line matrix wrapping and filling them. Due to the earlier and enhanced secretion of BSP and OPN on HAT-coated Mg as compared to PEO0-coated Mg (Fig. 2 and S4†), their mineralization formed a cement line matrix that appeared on HAT-coated Mg within the endosseous implantation time of 3 days (Fig. 8a), which was earlier than that for PEO0-coated Mg (within 7 days, but out of 3 days, Fig. 7b and a), while no cement line matrix appeared on bare Mg even on day 14 of implantation (Fig. 6b). Besides the recruitment of osteogenetic cells, HAT-coated Mg indeed accelerated their differentiation as the osteogenesis-related markers within its surrounding bone (e.g., ALP, BSP, OPN, and OCN) were dramatically upregulated at the gene level as compared to PEO0-coated and bare Mg (Fig. 3d). Subsequently, the differentiating cells secreted collagen, assembling into fibers to overlay on the cement line matrix, which would further induce the mineralized apatite to form a bone matrix (Fig. 7c, d and 8b, c). Owing to the cement line matrix wrapping each of the HA nanorod and filling its space, stronger mechanical interdigitation could be formed as compared to the case presented on microporous MgO/PEO0-coated Mg. Moreover, the cement line matrix overlying the bone matrix could be more mature on HAT-coated Mg as compared to PEO0-coated and bare Mg at the same implantation time due to the earlier secretion of collagen fibers and their mineralization on the former. Collectively, these can provide resistance to the removal of the bone matrix in three orthogonal planes to enhance the mechanical stability at the bone– implant interface (Fig. 6a). In total, HA-nanorod-patternedcoated HAT capably modulated the endosseous environment favoring contact osteogenesis and formed mechanical interdigitation with the new bone to achieve the fixation of implants with the bone.


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4.2. In vivo mechanical integrities of coated Mg and structural evolution of coatings During the initial period following a surgery, implants should provide sufficient mechanical support for the fractured bone. However, the un-uniform degradation of Mgbased implants in the physiological environment could induce a premature loss of mechanical integrity,14 suggesting that coatings conducted on Mg-based implants should inhibit the severe pitting corrosion of the substrate. In both the cases of implantation in the bone marrow cavity and femur, PEO0 coating can, to some extent, protect the Mg substrate from corrosion within the initial 4 weeks; thereafter, however, there is a dramatic increase in the corrosion rate and rapid decrease in the mechanical integrity on PEO0coated Mg (Fig. 9b–d and S5†). This may be due to the permeation of body fluids through, on one hand, the original micropores of the PEO0 coating; on the other hand, the conversion of MgO to Mg(OH)2 resulted in pathways and even loss of structural integrity of the coating (similar to the case shown in Fig. 5b2), onto the Mg substrate, leading to the formation of corrosion pits (Fig. 9a). Alternatively, HAT coating maintains the in vivo mechanical integrity of the substrate for a longer duration as compared to PEO0 coating (Fig. 9d). This is attributed to the unique HA-nanorod-patterned bilayer structure of HAT coatings. On one hand, the pores in the MgO layer were sealed with HA and Mg(OH)2, effectively avoiding the penetration of body fluids through the coating on the Mg substrate. On the other hand, the HA nanorods overlying on the MgO layer could partially inhibit the contact of MgO to body fluids, delaying the transition of MgO to Mg (OH)2, and therefore, maintaining the structural integrity of the coating for a longer duration (as the case shown in Fig. 5b3). In addition, the HA nanorods induced a new bone by means of contact osteogenesis, which is more pronounced as compared to the MgO-induced one. Further, this can effectively act as a barrier to impair the penetration of body fluids into the coating, and therefore, onto the substrate. Noticeably, the increase in the corrosion rate and decrease in the mechanical integrity of HAT-coated Mg are more pronounced after 8 weeks of implantation, implying a gradual loss in the protective efficacy of the coating with further prolonging implantation. However, HAT-coated Mg retains the in vivo mechanical integrity to a large extent for a longer duration as compared to the currently developed surfacemodified Mg implants (Fig. 9f ), satisfying the clinical requirement for Mg-based implants involving increased corrosion resistance during the initial implantation and relatively fast degradation after the consolidation of the fractured bone.13 Following endosseous implantation, PEO0 and HAT coatings underwent, at different levels, structural evolution with time. At week 12, PEO0 coating overlying on Mg could not be observed; instead, Mg(OH)2 appeared around Mg (Fig. 5b2). This phenomenon also occurred in some regions of the HAT coating, although it still persisted elsewhere and seemed rela-

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tively integrated (Fig. 5b3). In fact, the vanishing of the coatings on Mg alloys in vivo can be ascribed to either the degradation or delamination from the substrates resulting in fragments or nanoparticles; the latter would result in osteolysis.10 Based on the fact that no osteolysis could be observed at the interfaces of Mg(OH)2–bone (the magnified images of site A in Fig. 5b2 and site B in Fig. 5b3) and HAT coating–bone (the magnified image of site C in Fig. 5b3), it is suggested that the entire vanishing of the PEO0 coating and partial vanishing of the HAT coating at week 12 can be attributed to the degradation of MgO and HA, rather than the delamination of the coatings from Mg to generate fragments or nanoparticles. Our results clarified that the MgO in PEO0 and HAT coatings would convert to Mg(OH)2 and then react with Cl− in the body fluid to transform into soluble MgCl2.

5. Conclusions A bilayer-structured coating (HAT) has been formed by the hydrothermal treatment of plasma electrolytic oxidized porous MgO (PEO0) on Mg. The HAT coating comprised an inner layer of MgO with HA nanorods/Mg(OH)2 nanorods sealing the pores and an outer layer of HA nanorods. As compared to bare and PEO0-coated Mg, HAT-coated Mg induced a quicker phenotype switch of the macrophages from M1 to M2, significantly downregulated TNF-α and IL-1β, upregulated IL-10, and suppressed osteoclastogenesis, modulating the surrounding environment to facilitate the recruitment of osteogenetic cells. Moreover, HAT-coated Mg accelerated the secretion of BSP and OPN and their mineralization to form a cement line matrix. It also promoted the differentiation of osteogenetic cells and secretion of collagen fibers overlying on the cement line matrix, inducing earlier and more pronounced bone matrix formation. The cement line matrix wrapped each of the HA nanorods and filled the interrod space of the HAT coating to form strong interdigitation at the bone–coating interface, achieving enhanced osseointegration by means of contact osteogenesis. Owing to the enhanced inhibition of the body fluids to penetrate onto the Mg substrate because of the HAT coating, which was attributed to the sealed pores in the MgO layer by HA and Mg(OH)2. Therefore, the conversion of HA-nanorod MgO into Mg(OH)2 was delayed and the new bone acted as a more pronounced barrier. HAT-coated Mg maintained its mechanical integrity for a longer duration as compared to PEO0-coated and bare Mg. It is also clarified that the degradation of MgO and HA, rather than delamination, was the vanishing mode of PEO0 and HAT coatings during long-term implantations, avoiding osteolysis induced by the delamination-generated particles.

Conflicts of interest The authors declare no conflict of interest.

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Acknowledgements We appreciate Research Fund for the National Natural Science Foundation of China (Grant number 51631007, 51371137 and 31700860), China Postdoctoral Science Foundation (2017M613128) and the Fundamental Research Funds for the Central Universities (xjj2018230) for financially supporting this work.

References 1 M. P. Staiger, A. M. Pietak, J. Huadmai and G. Dias, Biomaterials, 2006, 27, 1728–1734. 2 Y. F. Zheng, X. N. Gu and F. Witte, Mater. Sci. Eng., R, 2014, 77, 1–34. 3 H. Hornberger, S. Virtanen and A. R. Boccaccini, Acta Biomater., 2012, 8, 2442–2455. 4 T. S. N. S. Narayanan, I. S. Park and M. H. Lee, Prog. Mater. Sci., 2014, 60, 1–71. 5 R. C. Zeng, L. Y. Cui, K. Jiang, R. Liu, B. D. Zhao and Y. F. Zheng, ACS Appl. Mater. Interfaces, 2016, 8, 10014– 10028. 6 Y. K. Kim, Y. S. Jang, Y. H. Lee, H. K. Yi, T. S. Bae and M. H. Lee, Sci. Rep., 2017, 7, 712. 7 G. Y. Li, L. Zhang, L. Wang, G. Y. Yuan, K. R. Dai, J. Pei and Y. Q. Hao, Acta Biomater., 2018, 65, 486–500. 8 Y. N. Jiang, B. Wang, Z. R. Jia, X. Lu, L. M. Fang, K. F. Wang and F. Z. Ren, J. Biomed. Mater. Res., Part A, 2017, 105, 2750–2761. 9 F. Peng, H. Li, D. H. Wang, P. Tian, Y. X. Tian, G. Y. Yuan, D. M. Xu and X. Y. Liu, ACS Appl. Mater. Interfaces, 2016, 8, 35033–35044. 10 J. Pajarinen, V. P. Kouri, E. Jamsen, T. F. Li, J. Mandelin and Y. T. Konttinen, Acta Biomater., 2013, 9, 9229–9240. 11 X. N. Gu, N. Li, W. R. Zhou, Y. F. Zheng, X. Zhao, Q. Z. Cai and L. Q. Ruan, Acta Biomater., 2011, 7, 1880–1889. 12 H. H. Luo, Q. H. Cai, B. K. Wei, B. Yu, D. J. Li, H. He and Z. Liu, J. Alloys Compd., 2008, 464, 537–543. 13 S. F. Fischerauer, T. Kraus, X. Wu, S. Tangl, E. Sorantin, A. C. Hänzi, J. F. Löffler, P. J. Uggowitzer and A. M. Weinberg, Acta Biomater., 2013, 9, 5411–5420. 14 X. Lin, L. L. Tan, Q. Zhang, K. Yang, Z. Hu, J. Qiu and Y. Cai, Acta Biomater., 2013, 9, 8631–8642. 15 H. P. Duan, K. Q. Du, C. W. Yan and F. H. Wang, Electrochim. Acta, 2006, 51, 2898–2908. 16 W. Shang, B. Z. Chen, X. C. Shi, Y. Chen and X. Xiao, J. Alloys Compd., 2009, 474, 541–545. 17 P. Lu, L. Cao, Y. Lin, X. Xu and X. Wu, J. Biomed. Mater. Res., Part B, 2011, 96, 101–109. 18 J. H. Zhou, Y. Han and S. M. Lu, Int. J. Nanomed., 2014, 9, 1243–1260. 19 J. H. Zhou, B. Li, S. M. Lu, L. Zhang and Y. Han, ACS Appl. Mater. Interfaces, 2013, 5, 5358–5365. 20 J. H. Zhou, B. Li, Y. Han and L. Z. Zhao, Nanomedicine, 2016, 12, 1161–1173.


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21 S. Wenisch, J. P. Stahl, U. Horas, C. Heiss, O. Kilian, K. Trinkaus and R. Schnettler, J. Biomed. Mater. Res., Part A, 2003, 67, 713–718. 22 J. E. Davies, Biomaterials, 2007, 28, 5058–5067. 23 Q. L. Ma, L. Z. Zhao, R. R. Liu, B. Q. Jin, W. Song, Y. Wang, Y. S. Zhang, L. H. Chen and Y. M. Zhang, Biomaterials, 2014, 35, 9853–9867. 24 J. A. Jones, D. T. Chang, H. Meyerson, E. Colton, I. K. Kwon, T. Matsuda and J. M. Anderson, J. Biomed. Mater. Res., Part A, 2007, 83, 585–596. 25 Z. T. Chen, C. T. Wu, W. Y. Gu, T. Klein, R. Crawford and Y. Xiao, Biomaterials, 2014, 35, 1507–1518. 26 J. E. Davies, V. C. Mendes, J. C. H. Ko and E. Ajami, Biomaterials, 2014, 35, 25–35. 27 Z. T. Chen, J. Yuen, R. Crawford, J. Chang, C. T. Wu and Y. Xiao, Biomaterials, 2015, 61, 126–138. 28 C. T. Wu, Z. T. Chen, Q. J. Wu, D. L. Yi, T. Friis, X. B. Zheng, J. Chang, X. Q. Jiang and Y. Xiao, Biomaterials, 2015, 71, 35–47. 29 Z. T. Chen, X. L. Mao, L. L. Tan, T. Friis, C. T. Wu, R. Crawford and Y. Xiao, Biomaterials, 2014, 35, 8553–8565. 30 J. D. Sun, Y. Zhu, L. Meng, P. Chen, T. T. Shi, X. Y. Liu and Y. F. Zheng, Acta Biomater., 2016, 45, 387–398. 31 F. Witte, J. Fischer, J. Nellesen, C. Vogt, J. Vogt, T. Donath and F. Beckmann, Acta Biomater., 2010, 6, 1792–1799. 32 S. Chen, S. K. Guan, W. Li, H. Wang, J. Chen, Y. Wang and H. Wang, J. Biomed. Mater. Res., Part B, 2012, 100, 533– 543. 33 J. Zhao, G. Shen, C. Liu, S. Y. Wang, W. J. Zhang, X. C. Zhang, X. L. Zhang, D. Ye, X. J. Wei, Z. Y. Zhang and X. Q. Jiang, Tissue Eng., Part A, 2012, 18, 185–197. 34 J. W. Park, Y. J. Kim, J. H. Jang and H. Song, J. Biomed. Mater. Res., Part A, 2013, 101, 298–306. 35 F. Lebre, R. Sridharan, M. J. Sawkins, D. J. Kelly, F. J. O’Brien and Ed. C. Lavelle, Sci. Rep., 2017, 7, 2922. 36 R. Huang, Y. Han and S. M. Lu, J. Mater. Chem., 2014, 2, 4531–4543. 37 C. Minkin, Calcif. Tissue Int., 1982, 34, 285–290.


Paper

38 Z. J. Zhai, X. H. Qu, H. W. Li, K. Yang, P. Wan, L. Tan, Z. Ouyang, X. Liu, B. Tian, F. Xiao, W. Wang, C. Jiang, T. Tang, Q. Fan, A. Qin and K. Dai, Biomaterials, 2014, 35, 6299–6310. 39 J. E. Davies, J. Dent. Educ., 2003, 67, 932–949. 40 K. Chatzipanagis, C. G. Baumann, M. Sandri, S. Sprio, A. Tampieri and R. Kröger, Acta Biomater., 2016, 46, 278– 285. 41 M. A. Ulla, R. Spretz, E. Lombardo, W. Daniell and H. Knözinger, Appl. Catal., B, 2001, 29, 217–229. 42 K. Z. Nastaran, H. Chen, H. A. Goldberg, G. K. Hunter, B. Grohe and F. Lagugne-Labarthet, J. Am. Chem. Soc., 2012, 134, 17076–17082. 43 D. A. Puleo and A. Nanci, Biomaterials, 1999, 20, 2311– 2321. 44 A. Levillain, C. Boulocher, S. Kaderli, E. Viguier, D. Hannouche, T. Hoc and H. Magoariec, Osteoarthr. Cartil., 2015, 23, 1186–1193. 45 D. W. Lee, K. T. Koo, Y. J. Seol, Y. M. Lee, Y. Ku, I. C. Rhyu, C. P. Chung and T. Kim, J. Periodontal Implant Sci., 2010, 40, 132–138. 46 M. Thomann, C. Krause, N. Angrisani, D. Bormann, T. Hassel, H. Windhagen and A. Meyer-Lindenberg, J. Biomed. Mater. Res., Part A, 2010, 93, 1609–1619. 47 S. M. Kim, J. H. Jo, S. M. Lee, M. Kang, H. Kim, Y. Estrin, J. H. Lee, J. Lee and Y. Koh, J. Biomed. Mater. Res., Part A, 2014, 102, 429–441. 48 J. Draxler, E. Martinelli, A. M. Weinberg, A. Zitek, J. Irrgeher, M. Meischel, S. E. Stanzl-Tschegg, B. Mingler and T. Prohaska, Acta Biomater., 2017, 51, 526–536. 49 J. E. Davies, E. Ajami, R. Moineddin and V. C. Mendes, Biomaterials, 2013, 34, 3535–3546. 50 G. S. Baht, G. K. Hunter and H. A. Goldberg, Matrix Biol., 2008, 27, 600–608. 51 J. Sodek, B. Ganss and M. D. McKee, Crit. Rev. Oral Biol. Med., 2000, 11, 279–303. 52 A. Oldberg, B. Jirskog-Hed, S. Axelsson and D. Heinegård, J. Cell Biol., 1989, 109, 3183–3186.

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