In vivo biodegradation and biological properties of a Mg-Zn-Ca amorphous alloy for bone defect repair

ABSTRACT

Amorphous magnesium alloys, characterised by their unique disordered structure, exhibit exceptional mechanical properties and superior corrosion resistance. This study introduced a novel biodegradable Mg-Zn-Ca amorphous alloy. Immersion and electrochemical tests revealed uniform degradation in a simulated body fluid environment. The degradation products predominantly contained zinc and oxygen. After 28 days of immersion, the alloy’s structure largely remained intact, and it exhibited a minimal weight loss rate of 11.64 ± 1.85%, indicating its excellent corrosion resistance. In addition, the homogeneous and rapid degradation of the Mg-Zn-Ca amorphous alloy in rats (the residual volume at 12 weeks was 14.94 ± 5.05%, at least 2–3 times of the corrosion rate of immersion in vitro), thus increasing the bone volume/total volume and trabecular number. The new bone and the residual material surface showed good bone integration. Blood and organ tissue tests confirmed the in vivo biological safety. Therefore, Mg-Zn-Ca amorphous alloys have excellent potential as bone-repair materials, especially in load-bearing areas.

KEYWORDS:

• Mg-Zn-Ca amorphous alloy
• bone defects
• biodegradation
• osteogenesis

Introduction

The incidence of bone defects caused by high-energy trauma, infections, bone tumours, and other diseases is increasing. Almost all segmental bone defects cannot heal on their own and must be filled with implants to complete the replacement and fusion of the bone tissue [1]. Autologous bone has excellent biocompatibility, osteoconduction, and osteoinduction properties; however, its inherent disadvantages and potential complications, such as insufficient bone volume, size mismatch, and donor site injury, limit its application [2,3]. Allografted bone is an effective substitute for autologous bone. Homologous allograft bone originating from bone banks needs special treatment, such as freeze-drying and irradiation, to be applied in the clinic, and the cumbersome process of tissue acquisition and processing kills the biological factors on the surface and damages the mechanical strength. Immunogenic allograft bone may impede the adhesion and differentiation of cells, and delayed healing, risk of infection, and aseptic looseness occur in clinical applications [4]. Xenograft bones are obtained from animals from a wide range of species, and their natural three-dimensional structure and mechanical properties are very similar to those of human bones. However, they also lose their biological activities, such as osteoinduction, during the process of complete antigen removal and are prone to leaving behind chemical and toxic substances [5,6]. Widely used in clinical settings, medical metals such as stainless steel, titanium alloys, aluminium alloys, degradable magnesium alloys, and various other biomedical metals are notable [7–9], Iron-based degradable materials exhibit significantly enhanced strength [10,11], yet they are hindered by a notably slow corrosion rate within the body. This issue extends to newer stainless steel varieties, which are susceptible to fractures and conspicuous corrosion under complex operational conditions [12,13]. Titanium alloys are distinguished by their superior biocompatibility and robust mechanical characteristics [14]. Nonetheless, their biological inertness and the disparity in elastic modulus compared to bone tissue continue to constrain their advancement. Although a large number of synthetic bone repair materials are used in clinics, their drawbacks have been gradually revealed, such as material brittleness and poor mechanical strength; thus, they can only provide short-term structural support and cannot be used in the treatment of bone defects in load-bearing areas (i.e. hydroxyapatite, calcium phosphate, and biologically active glass) [15]. Other drawbacks include an uncontrolled rate of degradation, degradation products leading to cytotoxicity, and inflammation, resulting in treatment failure (i.e. calcium sulphate bone cement and synthetic polymer materials).

In recent years, biodegradable magnesium alloys with unique humoral degradation properties have become new bone graft materials. Magnesium alloys not only have a similar modulus of elasticity to bone and good biocompatibility and osteogenic activity but can also gradually degrade in the complex human body environment, attracting neighbouring bone tissue to grow into and encircle around them [16–18]. More importantly, the magnesium ions generated by the degradation of the material can stimulate the release of more neurotransmitters (mainly calcitonin gene-related peptide) from sensory nerve terminals at the periosteal site, promote the osteogenic differentiation of stem cells within the periosteum, and form a large amount of new bone at the periosteal site [19]. However, the rapid degradation of magnesium alloys after implantation leads to premature failure of the implant [20–22], while the accumulation of degradation products causes alkalisation of the physiological environment in the vicinity of the implant or even necrosis of the surrounding tissues [23,24], which is still a bottleneck limiting its clinical application. Therefore, there is an urgent need to explore novel corrosion-resistant magnesium alloys systematically.

Amorphous alloys, also known as ‘metallic glass’, have advantages over crystalline magnesium alloys due to their homogeneous structure, absence of crystal defects, and second-phase characteristics, such as high strength, high elastic limit, high fracture toughness, good corrosion resistance, and fatigue resistance [25]. Mg-Zn-Ca amorphous alloys have broad application prospects in the field of biomedical materials owing to their good amorphous-formation ability and biocompatibility. Loffler et al. [26] found that when the Zn composition of amorphous Mg – Zn – Ca alloys reaches a threshold (~28 at. %), corrosion products were deposited on the surface of the alloys to form a protective layer, which was capable of slowing down the rate of corrosion of the material, and no significant hydrogen cavities or inflammatory reactions were observed in implanted animals. Gu et al. [27] concluded that a Mg-Zn-Ca amorphous alloy not only increased the corrosion resistance but also improved the mechanical properties because its compressive strength was three times higher than that of pure magnesium; the surface of the material after immersion in simulated body fluid (SBF) had uniformly distributed micropores, and the in vitro cellular tests showed the adhesion of L-929 and MG63 cells on the surface of the material. Chen et al. [28] showed that the degradation rate of a Mg-Zn-Ca amorphous alloy after immersion was approximately 1/6 of that of pure magnesium, which could avoid the abnormal hydrogen precipitation and osteolysis caused by rapid degradation in vivo. Newborn bone was tightly attached to the surface of the material 2 months after implantation, which confirmed its good osteogenic activity. In our previous study, we found that [29] the glass transition temperature of Mg-Zn-Ca amorphous alloys increases during degradation when subjected to high temperatures owing to the increased interatomic bonding of amorphous group elements and thermal stability. Additionally, with the prolongation of the degradation time, the annihilation and redistribution of the free volume limit the rate of atomic migration, thus improving the corrosion resistance of these alloys.

However, because in vitro corrosion media do not accurately reflect the complex and variable environment of real organisms, uncontrollable factors, such as the physiological flow field and stress corrosion, directly affect the degradation behaviour of the alloy, thus affecting its service life in vivo, resulting in large discrepancies between in vitro immersion and in vivo implantation results. Studies applying Mg-Zn-Ca amorphous alloy implantation in animals are limited to imaging observations and histological evaluations. They also lack quantitative analyses of the bone response around the implant. Furthermore, few studies have used fluorescent markers to track the rate of new bone formation. Therefore, in this study, a Mg-Zn-Ca amorphous alloy was prepared by low-pressure casting in a copper mould, and its degradation properties and biological behaviours as a bone repair material for bone defects were comprehensively evaluated by immersion tests for 28 days and in vivo implantation experiments in animals for 12 weeks to explore the feasibility of its use in the clinical treatment of bone defects.

Materials and methods

Preparation of Mg -Zn-Ca amorphous alloy

A mixture of pure Mg (>99.95%), pure Zn (>99.95%), and Mg-30Ca alloy was selected to be melted in an induction furnace. Argon was used as a protective gas to prepare an alloy ingot with the composition of Mg₇₀Zn₂₅Ca₅. The power of the heating furnace power supply was 2 kw. When the alloy was completely melted and its composition was homogeneous, it was cooled in a furnace to room temperature. The chemical composition of the prepared magnesium alloy elements is presented in Table 1. The above alloy was remelted in a quartz tube in the same induction furnace. Then, the remelted alloy was injected into a copper mould with a hole size of ø 3 × 60 mm, spray-cast into an amorphous rod, and cut by a fast diamond saw to obtain a cylinder with a thickness of 4 mm (Figure 1). The prepared samples were characterised using X-ray diffraction (XRD, D/Max-2500PC, Rigaku,Tokyo, Japan) and differential scanning calorimetry (DSC, 204 F1 Phoenix®, Netzsch, Selb, Germany). Transmission electron microscopy (TEM; Tecnai G2 20, FEI, Hillsboro, Oregon, U.S.A.) was used to observe and analyse the microstructures of the samples. Because Mg-based metals corrode easily and to avoid the crystallization of Mg-based amorphous alloys, an ion thinning apparatus (691 PIPS, Gatan, Pleasanton, U.S.A.) was used in the experiment. The samples were further thinned at-100°C in liquid nitrogen, and the final samples were obtained.

Figure 1. Macrophotograph of the Mg-zn-ca amorphous alloy.

Table 1. The actual composition of Mg-zn-ca amorphous alloy.

Mg (at. %)	Zn (at. %)	Ca (at. %)
69.60	25.36	5.04

In vitro corrosion resistance

Electrochemical test

The circular sample had a cross-sectional area of 0.07 cm², which was encapsulated by epoxy resin. Electrochemical tests were carried out in Hank’s solution (Table 2) at 37°C using a typical three-electrode electrochemical system: the sample was the working electrode, the counter electrode was a platinum electrode, and a standard saturated potassium chloride-glycury electrode was used as the reference electrode. Open circuit potential (OCP) test monitoring was performed for 30 min (dynamic potential polarisation curve voltage measurement range: −0.25 V–1 V, scanning speed: 0.5 mV/s), and each test was repeated three times.

Table 2. Chemical composition of Hank’s solution (g/L). Reprinted from ref [19].

NaCl	KCl	KH2PO4	MgSO4	NaHCO3	CaCL2	Na2HPO4	Glucose
8.0	0.4	0.06	0.20	0.35	0.14	0.12	1.00

Immersion test

The experimental samples were immersed in Hank’s solution (pH 7.4) at 37 ± 0.5°C for 28 days following the guidelines set by the ISO10993–15 standard [30]. Table 2 presents the composition of Hank’s solution. The immersion ratio was maintained at 1.25 cm²/mL, and the solution was refreshed every 24 h. The pH of the solution was measured daily for 28 days. The surface products were examined using scanning electron microscopy (SEM; Merlin Compact, Zeiss, Oberkochen, Germany) after 7, 14, 21, and 28 days of immersion. Additionally, the compositions of the degradation products were analysed using an X-ray photoelectron spectrometer (XPS, Escalab250, Thermo, Waltham, Massachusetts, U.S.A.) after the 28-day immersion period. Subsequently, the samples were ultrasonically cleaned for 15 min in a chromium trioxide solution (200 g/L Cr₂O₃ +10 g/L AgNO₃) to eliminate surface corrosion products. The corrosion rate of the samples after the immersion test, Pw, was calculated according to the following equation [31]: P_w = K·W/(A·T·D), where the unit of the corrosion rate is mm/year, K( = 8.76 × 10⁴) is a constant in mm/a, W is the mass loss in (g), A is the exposed surface area (cm²), T is the immersion time, and D is the density of the implant material (g/cm³). The degraded morphology was observed by SEM and confocal laser scanning microscopy (CLSM; LSM 710, Zeiss, Oberkochen, Germany).

In vivo biological properties

Surgical procedure

All animal experiments were reviewed and approved by the Animal Ethics Committee of the Affiliated Hospital of Shandong University of Traditional Chinese Medicine (approval number: 2021–67; 11 August 2021). A rat lateral femoral condyle cylindrical bone defect repair model was established in this study. Thirty male rats, aged 12 weeks, with an average weight of 326 ± 30.6 g, were included in the experiment. The surgery was performed under sterile conditions. The samples were cylindrical (Ø3 × 4 mm) and sterilised with 75% alcohol and ultraviolet light for 60 min. All rats were intraperitoneally injected with ketamine (10 mg/kg) (Shanghai Ziyuan Pharmaceutical Co., Ltd., Shanghai, China) and 2% xylazine (10 mg/kg) (Shanghai Ziyuan Pharmaceutical Co., Ltd.). In this study, a longitudinal incision of approximately 15 mm was made on the lateral side of the patellar ligament. A cylindrical defect area with a diameter of Ø3 × 4 mm was drilled in the direction of the through-condylar line, as shown in Figure 2a. The fragmented bone was thoroughly rinsed with saline solution before the implantation procedure was carried out using either the Mg-Zn-Ca amorphous alloy or pure titanium material, as depicted in Figure 2b,c. Following implantation, the surgical site was rinsed again with saline solution, and the incision was closed using layered sutures. For postoperative analgesia, buprenorphine (Temgesic; Reckitt & Colman, Hull, UK) was administered subcutaneously at a dose of 0.3 mg/kg. The rats were euthanised 4, 8, and 12 weeks after surgery to collect their femurs. Two groups were included in the study: the Mg-Zn-Ca amorphous alloy experimental group, which received an implant made of Mg-Zn-Ca amorphous alloy, and the pure Ti control group, which received an implant made of pure titanium to repair the bone defect. Each group consisted of 15 rats (Mg-Zn-Ca amorphous alloy and pure Ti), with five rats euthanised at each time point (4, 8, and 12 weeks) for femur collection.

Figure 2. A critical surgical procedure in rats involves creating a cylindrical defect area (a); implanting Mg-Zn-Ca amorphous alloy (b), and using pure titanium (c).

General condition of the animals and the in vivo biosafety

The post-surgical conditions of each rat, including their body temperature, body weight, and wound healing, were observed daily. Twelve weeks after surgery, arterial blood samples were randomly collected from both the experimental and control groups using the cardiac blood sampling method. Serum concentrations of Mg²⁺, Zn²⁺, and Ca²⁺ were measured using inductively coupled plasma mass spectrometry (ICP-MS,NexION 300,PerkinElmer,Waltham,Massachusetts,U.S.A.). Additionally, at 12 weeks postoperatively, tissue samples were obtained from the heart, liver, spleen, and kidneys of the rats in both groups. The collected organ tissues were fixed, embedded, and stained with haematoxylin and eosin (H&E) to identify potential pathological changes. Each group was subjected to routine blood and blood biochemistry tests, and the 15 indexes included white blood cells (WBC), red blood cells (RBC), platelets (PLT), alanine aminotransferase (ALT), aspartate aminotransferase (AST), direct bilirubin (D-BIL), alkaline phosphatase (ALP), albumin (ALB), gamma-glutamic acid basotransferase (γ-GT), blood urea nitrogen (BUN), creatinine (CR), uric acid (UA), creatine kinase (CK), lactate dehydrogenase (LDH-L), and lactate dehydrogenase isoenzyme (LDH-I).

Micro-CT evaluation of the osteogenic properties of Mg-Zn-Ca amorphous alloy in vivo

The right femur of each euthanised rat was completely excised and fixed in 4% paraformaldehyde. The distal femur and femoral condyle were scanned using micro-computed tomography (Micro-CT,viva CT 80, Scanco, Brüttisellen, Switzerland). Medium-resolution mode and an isotropic voxel size of 15 μm (mCT80, Scanco Medical AG, Bassersdorf, Switzerland) were used to evaluate new bone formation around the implants within 1 mm. The horizontal plane of all samples had to be oriented perpendicular to the axis of the X-ray beam. The 2D radiographic images were used to reconstruct and calculate the corresponding bone volume fraction (bone volume/total volume, BV/TV), bone mineral density, trabecular number (TB.n), trabecular spacing (TB.SP), and trabecular thickness (TB.TH) using Scanco software (Scano Medical μCT Evaluation Program 6.6) by thresholding (n = 3). We evaluated the osteogenic activity and degradation behaviour of the Mg-Zn-Ca amorphous alloy in vivo.

Cross-sectional and histological observations

Following the micro-CT scan, a hard tissue slicing procedure was performed on the rat femur specimen. Prior to slicing, the specimens were fixed, washed in water, dehydrated with ethanol, cleaned with xylene, and embedded in methyl methacrylate. Four to five sagittal slices were created by cutting along the long axis of the femur vertically relative to the implant located in the femoral condyle. A single slice from each specimen was polished to 7000 grit, and a gold coating was applied to the surface. Using an FE-SEM (Hitachi S4800, Tokyo, Japan) equipped with an EDS (energy-dispersive spectrometer), an elemental analysis was conducted to investigate the distribution and composition of the remaining material in addition to the tissues surrounding the degradation products, focusing particularly on the presence and distribution of elements such as Ca and P, which are crucial for osteogenesis. After being ground and polished to a thickness of 50 μm, the remaining slices of each specimen were subjected to magenta-methylene blue, Van Gieson, and toluidine blue staining. The slices were observed and imaged (three images per slice) using a high-resolution microscope (Nikon Eclipse E200, Tokyo, Japan) with the following parameters: a full-view image, low-magnification image (20×), and high-magnification image (100×) of the bone defect area.

Fluorescent tracing of new bone formation

The rate of new bone formation was assessed using fluorescent labelling with calcein xanthophyll-dimethylphenol orange-tetracycline hydrochloride. The animals were injected intramuscularly with calcitonin (5 mg/mL, 2 mL/kg), dimethylphenol orange (50 mg/mL, 2 mL/kg), and tetracycline hydrochloride (30 mg/mL, 2 mL/kg) 21, 14, and 7 days prior to the execution of the animals. The specimens were collected and subjected to hard tissue sectioning, and then processed sections were placed under a confocal laser microscope and photographed for documentation.

Data analysis

Data were analysed using the statistical software GraphPad Prism 9.5.1 (GraphPad Software,San Diego,California,U.S.A.). Quantitative data are expressed as mean ± standard deviation (SD), and data were analysed using the independent samples t-test and one-way analysis of variance (ANOVA). Differences were considered to be statistically significant if p < 0.05 or p < 0.01.

Results

Characterization of the materials

Figure 3a shows the XRD curve of the Mg-Zn-Ca amorphous alloy. As seen in the figure, the Mg-Zn-Ca amorphous alloy exhibited typical amorphous diffuse peaks and sharp crystal peaks at 37° with low intensity. The physical phase analysis was Mg_0.97Zn_0.03. In Figure 3b,a a clear exothermic peak can be seen in the DSC thermal analysis, indicating that the material was in an amorphous state.

Figure 3. XRD image (a) and DSC image (b) of the Mg-Zn-Ca amorphous alloy. a.U., arbitrary units; XRD, X-ray diffraction.

To further characterise the microstructure of the prepared Mg-Zn-Ca amorphous alloy, bright-field and high-resolution images of the samples were obtained by TEM. The results of this analysis are shown in Figure 4. The transmission bright-field image of the Mg-Zn-Ca amorphous alloy (Figure 4a) shows a uniform contrast, while a small number of nanocrystals with sizes less than 10 nm can be observed in the high-resolution image (Figure 4b), as shown by the arrows in the figure. At the same time, there was a very small amount of crystal phase in the matrix, as shown in Figure 4c. Energy spectrum analysis was performed, and the results are shown in Table 3; these results are consistent with the phase Mg_0.97Zn_0.03 measured by XRD. There was no difference between the dendrite axis and dendrite composition, and there was no segregation in the crystal. According to the Mg-Zn binary phase diagram, the maximum solubility of Zn in Mg was 2.4 at.%, and the eutectic temperature was 340 ± 1°C. Combined with the DSC results (Figure 3b), the first melting peak in the curve corresponds to the melting peak of the Mg_0.97Zn_0.03 crystal phase, which is a low-melting alloy phase. The results show that the amorphous Mg-Zn-Ca alloy with Φ3 mm prepared by copper mould low-pressure casting contains a small amount of nanocrystalline and a very small amount of Mg_0.97Zn_0.03 crystalline phase.

Figure 4. TEM bright field image (a) and high-resolution TEM image (b) of the Mg-Zn-Ca amorphous alloy, and TEM bright field images with an electron diffraction pattern of the crystal phase (c,d). TEM, transmission electron tomography.

Table 3. Chemical composition of the crystal phase.

Element	Wt. %	at. %
Mg	93.19	97.36
Zn	6.81	2.64
Total	100.00	100

In vitro corrosion resistance

The change in pH over 28 d of immersion is shown in Figure 5a. The pH of Mg-Zn-Ca amorphous alloy rose to 8.48 ± 0.20 on the first day, stabilised over the next 10 days, and then decreased from the 12th day onwards, dropping to 7.05 ± 0.07 on the 28th day. Through the weight loss experiment to calculate the corrosion rate of the samples, as shown in Figure 5b,c, we can see that after 28 days of immersion, the degradation rate of the material was relatively low, which indicates that the prepared Mg-Zn-Ca amorphous alloy samples with a small number of fine nanocrystals and a very small amount of Mg_0.97Zn_0.03 crystal phase possess a uniform microstructure, no segregation, and thus excellent corrosion resistance. The weight loss ratios for 7, 14, 21, and 28 days were 0.70 ± 0.15%, 2.66 ± 0.64%, 7.28 ± 1.66% and 11.64 ± 1.85%, respectively. Furthermore, the corrosion rates for 7, 14, 21, and 28 days were 0.03 ± 0.01, 0.06 ± 0.01, 0.10 ± 0.02, and 0.12 ± 0.02 mm/year, respectively. Electrochemical tests were performed on the amorphous Mg-Zn-Ca alloy, as shown in Figure 5d. The corrosion potential (Ecorr) of the amorphous rods was −1.28 V, and the corrosion current density (Icorr) was 5.7 µA/cm².

Figure 5. pH value (a), weight loss (b), and corrosion rate (c) of the Mg-zn-ca amorphous alloy immersed in Hank’s solution for 28 days and potentiodynamic polarization curves of the Mg-zn-ca amorphous alloy in Hank’s solution (d).

Figure 6 shows the degradation morphology of the amorphous Mg – Zn – Ca alloy immersed for 7, 14, 21, and 28 d. With increasing time, the surface products (degradation products and deposits) gradually increased. Furthermore, the surface of the sample had no obvious corrosion pits, and surface corrosion was uniform. Shallow cracks on the surfaces of the samples were caused by dehydration.

Figure 6. Degradation morphology of the Mg-Zn-Ca amorphous alloy immersed in Hank’s solution for 7 (a), 14 (b), 21 (c), and 28 days (d), respectively.

The corrosion products on the samples after 28 days of immersion were examined by XPS, and the results are shown in Figure 7(a–c). The corrosion products mainly include Mg(OH)₂, CaCO₃, and a small amount of ZnO. The morphology of the substrate after removing its corrosion products is shown in Figure 7d, where the surface of the substrate was relatively flat, no corrosion pits or pitting occurred, and the morphology of the substrate remained intact.

Figure 7. XPS analysis of the degradation products (a–c) and surface morphology after removing the degradation products (d) on the surface of the Mg-Zn-Ca amorphous alloy immersed in Hank’s solution for 28 days. a.U., arbitrary units; XPS, X-ray photoelectron spectrometer.

In vivo biological properties

In vivo biosafety assessment

Histological examination of vital organs is the gold standard for assessing the biosafety of biodegradable materials in vivo. Figure 8 shows the H&E staining results of vital organs (heart, spleen, liver, and kidney) after the implantation of the Mg-Zn-Ca amorphous alloy material in rats. There were no significant changes in the H&E staining results of the Mg-Zn-Ca amorphous alloy group compared to the control group throughout the observation cycle. Additionally, no prominent pathological changes were observed in the tissues of critical organs. This shows that the Mg-Zn-Ca amorphous alloy material did not negatively affect the circulatory, immune, or urinary systems of the animal’s body during implantation.

Figure 8. H&E staining of the heart, spleen, liver, and kidney tissues from rats in the Mg-Zn-Ca amorphous alloy and pure Ti groups. H&E, hematoxylin and eosin.

When magnesium alloys are implanted into the body, their degradation products are inevitably released and enter the bloodstream to circulate throughout the body. Therefore, routine blood tests, blood biochemistry, and serum concentrations of Mg²⁺, Zn²⁺, and Ca²⁺ in rats 12 weeks after surgery were tested, and the results are shown in Figures 9 and 10. The biochemical indices and serum Mg²⁺, Zn²⁺, and Ca²⁺ levels in all experimental groups at 12 weeks postoperatively did not differ significantly from those of the control group and were all within the normal reference ranges. Therefore, the metal ions or degradation products produced by the in vivo degradation of the Mg-Zn-Ca amorphous alloy do not affect the liver and kidney functions of the organism, which indicates good biosafety and further confirms the ability of the experimental animals to metabolise the magnesium ions produced by the degradation of the magnesium alloy.

Figure 9. Levels of the main serum biochemical indicators of liver and kidney function. (a) ALT, (b) AST, (c) DBIL, (d) ALB, (e) ALP, (f) γ-GT, (g) BUN, (h) CREA, (i) UA, (j) CK, (k) LDH, and (l) LDH1 serum magnesium. ALT, alanine aminotransferase; AST, aspartate aminotransferase; DBIL, direct bilirubin; ALB, albumin; ALP, alkaline phosphatase; γ-GT, gamma-glutamic acid basotransferase; BUN, blood urea nitrogen; CREA, creatine; UA, uric acid; CK, creatine kinase; LDH, lactate dehydrogenase; LDH1, lactate dehydrogenase isoenzyme.

Figure 10. Blood cell counts and ion concentrations of Mg²⁺, Zn²⁺, and Ca²⁺ in the Mg-Zn-Ca amorphous alloy group and the pure Ti group; (a) WBC, (b) RBC, (c) PLT (d) Mg²⁺, (e) Zn²⁺, and (f) Ca²⁺. WBC, white blood cell; RBC, red blood cell; PLT, platelets.

Micro-CT results

Images of femoral condylar defect sites in rats implanted with Mg-Zn-Ca amorphous alloy and pure Ti were reconstructed in three dimensions using micro-CT, showing the implantation location of the materials, new bone generation, and in vivo degradation of the Mg-Zn-Ca amorphous alloy. Figure 11 shows a small amount of new bone generation around the Mg-Zn-Ca amorphous alloy and pure Ti at 4 weeks postoperatively and mild degradation of the Mg-Zn-Ca amorphous alloy material, with the overall shape and contour remaining largely intact. A small amount of new bone tissue was generated around the pure Ti at eight weeks postoperatively, and there was still a gap between the pure Ti material and the surrounding bone, indicating poor integration of the material with the surrounding bone tissue. The Mg-Zn-Ca amorphous alloy underwent further degradation after eight weeks of implantation, with blurring of the material boundaries accompanied by a large amount of granular debris. At 12 weeks postoperatively, a large amount of new bone tissue was visible around the pure Ti and Mg-Zn-Ca amorphous alloys; the amount of new bone gradually increased with time, and the amount of new bone around the Mg-Zn-Ca amorphous alloy was significantly higher than the amount of new bone around the pure Ti. Meanwhile, the Mg-Zn-Ca amorphous alloy was completely degraded, whereas the pure Ti group showed no significant change in material volume.

Figure 11. Sagittal, coronal, and three-dimensional reconstruction of bone defect repair in vivo. The amount of new bone (red) increased gradually with time and was significantly higher, and the implants (yellow) gradually degraded.

Figure 12 shows the micro-CT reconstruction of a tomographic slice scan of the bone defect area with two slices per defect per time period. For the Mg-Zn-Ca amorphous alloy at all stages, that is, different degrees of localised degradation, from 4 weeks onwards, the material can be seen in the internal cracks generated; the material’s peripheral profile is basically regular, and with the prolongation of time, the amount of new bone around the material increased, with a good combination of the bone tissue, indicating that the material in vivo has an excellent ability to contribute to the bone.

Figure 12. Computed tomography showing the repair of a bone defect in a two-dimensional section.

The results of the quantitative analysis are shown in Figure 13. The BV/TV indices were higher in the Mg-Zn-Ca amorphous alloy than in the pure Ti group at 4 and 8 weeks. The 12-week measurements of Tb.N and Tb.Th were also higher in the Mg-Zn-Ca alloy group than in the pure Ti group. The remaining volume of the material was approximately 73.50 ± 5.33% at 4 weeks, with most of the degradation occurring at 8 weeks, resulting in a remaining volume of approximately 40.62 ± 13.53%. At 12 weeks, almost complete degradation was observed, with a remaining volume of 14.94 ± 5.05%. These findings further support the outstanding osteogenic and biodegradable properties of the Mg-Zn-Ca amorphous alloy in vivo, which was characterised by a consistent degradation rate. By comparing our in vitro corrosion results, we observed that the corrosion rate of the Mg-Zn-Ca amorphous alloy varied between the in vivo and in vitro environments.

Figure 13. Quantitative analyses of the osteogenesis indices, including BMD, BV/TV, Tb.Sp, Tb.N, Tb.Th, and the Mg-Zn-Ca volume (%) at 4, 8, and 12 weeks. Significance levels are indicated by * p < 0.05 and ** p < 0.01. BMD, bone mineral density; BV/TV, bone volume/total volume; Tb.Sp, trabecular spacing; Tb.N, trabecular number; Tb.Th, trabecular thickness.

Histological evaluation

Figure 14a shows that the original mineralised bone trabeculae were light in colour, and the new bone trabeculae were brightly coloured in purple-red. A layer of bone tissue growing along the material can be seen around the pure titanium rods at 4 weeks postoperatively, but the adherence is not tight; most of the area between the bone tissue and the material is still separated by a layer of interstitial space, and only a small amount of the bone tissue is directly bonded to the surface of the material. In contrast, the Mg-Zn-Ca amorphous alloy underwent significant biodegradation with blurring of the material contour and regeneration of new bone tissue in the voids created by material degradation. At 8 and 12 weeks postoperatively, a further increase in neoplastic bone tissue was observed around both the pure titanium and Mg-Zn-Ca amorphous alloy materials compared to the preoperative period; the neoplastic bone in the Mg-Zn-Ca amorphous alloy group was embedded in the surface of the remaining material, and more bone tissue was observed to grow into the voids created by the degradation of the material under high-magnification microscopy. The outline of the pure Ti material remains clear; however, there is still a large gap between the surface of the implant and the surrounding neoplastic bone tissue, and the two are not tightly integrated. The Van Gieson staining results (Figure 14b) show that the yellow area is an unmineralised bone-like material, and the orange colour indicates the newborn bone; the amount of new bone in the Mg-Zn-Ca amorphous alloy and pure Ti both increases with time, and high magnification shows that the Mg-Zn-Ca implant has good osseointegration, presenting good bone defect repair performance and osteogenic advantages. In Figure 14c, toluidine blue staining shows that the neoplastic bone is dark blue, the osteoid is light blue, and there is sparse neoplastic bone tissue around pure Ti; however, there is still a gap between the surface of the implant and the neoplastic bone. At 8 weeks postoperatively, there was more new bone tissue around the Mg-Zn-Ca amorphous alloy material, and at 12 weeks postoperatively, degradation products were visible around the Mg-Zn-Ca amorphous alloy material. A large amount of new bone tissue could be seen under high-magnification microscopy, growing into the void created by the degradation of the implant.

Figure 14. Microstructure analysis results of (left) the Mg-Zn-Ca amorphous alloy and (right) pure Ti group. In each group of stained images, full-view, low-magnification (20-fold), and high-magnification (100-fold) images of the bone defect area are arranged in order from left to right. (a) Magenta-methylene blue staining, (b) Van Gieson staining, (c) toluidine blue staining.

Overall, the amount of new bone at the bone defect site increased with implantation time in both groups, and the Mg-Zn-Ca amorphous alloy group had a larger amount of new bone, showing good bone growth and osseointegration characteristics. This indicates that the biodegradation of the Mg-Zn-Ca amorphous alloy material promoted the regeneration and growth of new bone into the bone defect site. In addition, the volume of pure Ti at each time point remained the same and did not undergo biodegradation.

Cross-sectional results

Figure 15 depicts the SEM images and energy spectroscopy results of both the metal implant and the surrounding bone tissue. The analysis revealed that, in addition to Mg and Ti, several other predominant elements were detected in the bone tissue, including Zn, Ca, P, O, and C. Notably, the distribution of elemental Ti in the pure Ti group showed distinct boundaries at 4, 8, and 12 weeks, indicating that the material remained intact over time. However, valuable insights were obtained from the EDS spectrum during the biodegradation of the Mg-Zn-Ca amorphous alloy material. Specifically, the EDS spectra enabled the identification of three distinct components, each characterised by unique elemental signals. The first layer was primarily composed of Mg, which is an element commonly found in plants. Strong C and O signals were observed near the implant because these elements are typically present in biological tissues. Another notable component displayed signals for Ca and P, both of which are crucial for hydroxyapatite formation. These minerals play a vital role in animal metabolism and skeletal development and promote bone growth. Notably, Ca and P are interdependent, and their absence or imbalance can hinder osteoblast differentiation and prevent new bone formation.

Figure 15. EDS mapping of the Mg-Zn-Ca amorphous alloy (a,b) and pure ti (c) at postoperative intervals of 4, 8, and 12 weeks. In the mapping, Mg and Ti, Zn, Ca, Sr, O, P, and C are depicted in yellow, orange, purple, yellow, green, blue, and red, respectively.

Rate of new bone formation

Fluorescent labelling tracing was performed by injecting fluorescent stains directly into the animal and by detecting the range of fluorescence and the spacing between different fluorescent bands to understand the rate of formation and growth of bone tissue. As shown in Figure 16, obvious fluorescence staining bands were observed around the Mg-Zn-Ca amorphous alloy and pure Ti at all time points, forming concentric rings or irregular shapes with green, red, and yellow fluorescence sequentially from far to near relative to the implant; the spacing between different fluorescence bands can indirectly indicate the rate of new bone formation, and the fluorescence range can indirectly reflect the amount of osteogenesis. The spacing of the fluorescence bands in the pure Ti group was comparatively narrower than that in the Mg-Zn-Ca amorphous alloy group, with a smaller fluorescence range of 6.16 ± 0.86. This suggests that the rate of osteogenesis and amount of bone formation were lower in the pure Ti group. Conversely, in the Mg-Zn-Ca amorphous alloy group, the spacing of fluorescence bands and the fluorescence range were significantly larger at 8.87 ± 0.60. This indicated that new bone formation was more active in the Mg-Zn-Ca amorphous alloy group than in the pure Ti group. The quantitative analysis presented in Figure 16b also confirmed a significant difference in bone formation between the Mg and Zn – Ca amorphous alloy (12.88 ± 0.49) and pure Ti (5.60 ± 0.59) groups in terms of bone formation.

Figure 16. Osteogenesis rates of the Mg-Zn-Ca amorphous alloy and pure Ti. (a) Fluorescence image showing osteogenesis around the Mg-Zn-Ca amorphous alloy and pure Ti. (b) The resulting data were quantitatively analyzed, showing a significant difference between the Mg-Zn-Ca and the pure Ti groups (* p < 0.05, *** p < 0.001).

Discussion

Corrosion behaviors of Mg-Zn-Ca amorphous alloy

In this study, we found that the Mg-Zn-Ca amorphous alloy is not completely amorphous but rather contains a small number of nanocrystals and an extremely small amount of the crystalline phase. The Mg-Zn-Ca pure amorphous alloy prepared by Gu et al. [17] was exposed to an experimental solution of simulated body fluid (SBF) at 37°C. The pH of the solution increased to approximately 10.5 over a period of 30 days, with a corrosion potential of approximately −1.3 V and a corrosion current density of 11.2 µA/cm². In this study, an Mg-Zn-Ca amorphous alloy containing a small number of nanocrystals was immersed in Hank’s simulated fluid. The pH of the immersion solution increased to approximately 8.5 on day 1, remained stable, and started decreasing on day 12, reaching approximately 7.05 on day 28. Previous studies have indicated that the degradation rate of Mg-alloy implants in vitro is generally 1–5 times higher than that observed in vivo [32]. The weight loss rate of the Mg-Zn-Ca alloy prepared in this study was 11.64 ± 1.85% during the first 4 weeks of immersion in Hank’s simulated body fluid in vitro. This implied that the complete degradation time of the prepared Mg-Zn-Ca amorphous alloy rods in vivo was approximately 40 weeks. However, after 4 weeks of implantation in rats, the residual volume of the Mg-Zn-Ca amorphous alloy was approximately 73.50 ± 5.33%. Most of the material degraded at 8 weeks, with a residual volume of approximately 40.62 ± 13.53%, and the residual volume remained at 14.94 ± 5.05% at 12 weeks. Based on this trend, we predicted that the material would completely degrade within 15 weeks in vivo. Despite the excellent corrosion resistance exhibited by the amorphous Mg – Zn – Ca alloy in the in vitro corrosion test, the degradation rate after implantation was at least three times higher than the in vitro immersion corrosion rate. Witte et al. [23] used an aluminium – zinc – magnesium alloy (AZ91D) and an aluminium- and rare-earth-containing magnesium alloy (LAE442). The corrosion rates of these two magnesium alloys, as observed in the ASTM-D1141-98 solution in vitro, were opposite to the degradation trend observed in guinea pig femurs implanted in vivo for 18 weeks.

Based on the aforementioned data, rods containing fine nanocrystals exhibit superior corrosion resistance compared with pure amorphous rods when tested in vitro. However, it should be noted that no direct experimental comparison was conducted, highlighting the need for further investigation to determine whether the disparity in the degradation performance can be attributed to structural differences. Furthermore, in vivo experiments revealed significant differences in the corrosion behaviour between the Mg-Zn – Ca amorphous alloy observed through in vitro simulations and the degradation observed in animals. Therefore, it is crucial to improve the in vitro test method and identify the factors contributing to the discrepancies between in vivo and in vitro tests. These aspects should be considered carefully in future studies.

Osteogenic properties of Mg-Zn-Ca amorphous alloy in vivo

The repair of bone defects is a complex process involving multiple cytokines, growth factors, and intra- and extracellular signalling pathways [33–35]. Biomaterial implantation triggers a host immune response, and a series of immune cells regulate the repair of peri-implant bone tissue through chemokines and cytokines [36]. In general, M1 macrophages contribute to osteogenesis by initiating the acute inflammatory phase [37], clearing debris from the injury site, and directly secreting chemokines (e.g. C-C chemokine ligand 2 [CCL2], CXC motif ligand 8 [CXCL8], and stromal cell-derived factor 1 [SDF-1]) to recruit bone healing cells, including MSCs, bone progenitors, and vascular progenitors [38,39]. In contrast, M2 macrophages promote the differentiation and function of bone-healing cells by secreting anti-inflammatory cytokines and growth factors such as BMPs, VEGFs, and TGF-β [40,41]. It has also been shown that Mg²⁺ promotes the secretion of higher concentrations of the anti-inflammatory factors IL-4 and IL-10 by M2-type macrophages, the up-regulation of BMP-2 and VEGF expression, as well as the down-regulation of NF-kB signalling to promote osteogenesis, thereby conferring a variety of functions of Mg²⁺ in bone immunomodulation [42–44].

Mg²⁺ also enhances osteogenic differentiation by increasing the expression of Runx-2 and ALP through the TRPM7/PI3K signalling pathway. Knockdown of TRPM7 has been shown to significantly hinder the effects of Mg²⁺ on osteogenesis, mineralisation, migration, and chemotaxis, indicating that TRPM7 mediates the induction of osteogenic differentiation by Mg²⁺ in humans [45,46]. When BMSCs were exposed to conditioned medium with 100 mg/L Mg²⁺, a significant increase in ALP activity and the expression of osteogenic genes (Runx-2, ALP, OPN, and OCN) was observed, indicating the promotion of osteogenic differentiation of BMSCs by Mg²⁺ through the regulation of macrophages [47]. This suggests that the mechanism of bone repair using Mg-Zn-Ca amorphous alloy is not only related to the induction of degradation products to regulate osteogenic differentiation in BMSCs but also potentially involves the modulation of immune cell responses to create a favourable bone immune microenvironment.

In vivo biosafety evaluation of Mg-Zn-Ca amorphous alloy

The Mg²⁺ released from the degradation of Mg-Zn-Ca amorphous alloy materials after implantation is absorbed by the body, most of which is used for the formation of new bone, whereas a small portion is utilised by other tissues and organs; the rest of the Mg²⁺ is excreted through hepatic, intestinal, and renal metabolism [45]. Zn is an essential trace element for the human body to maintain physiological functions. It is a basic component of more than 300 enzymes, has osteogenic and antibacterial characteristics, and is mainly stored in bones and muscles [46,48]. The Zn²⁺ produced during the degradation of the Mg-Zn-Ca alloy is metabolised and excreted from the body without accumulation in the organs. Calcium is an essential dietary element and a major component of bones. Therefore, the combination of Mg²⁺, Zn²⁺, and Ca²⁺ in the Mg-Zn-Ca amorphous alloy composed of all nutrient elements played an important role in cell proliferation, and the in vivo degradation did not bring about significant changes in the serum concentrations of Mg²⁺, Zn²⁺, and Ca²⁺ in the body. Results of the histopathological examination also indicated that it did not produce pathological changes in the heart, spleen, liver, and kidney tissues of the animals, which suggests that Mg-Zn-Ca amorphous alloys have good biological safety.

However, the application of Mg-Zn-Ca amorphous alloys in biomedical implants remains at an early stage. The experimental time of this study was relatively short, and the effect of the Mg-Zn-Ca amorphous alloy on the healing of bone defects was only observed three months after the operation. Therefore, it is necessary to carry out long-term experimental observations to effectively control the degradation rate of Mg-Zn-Ca amorphous alloy in vivo to make it perfectly match the rate of new bone formation.

Conclusions

In this study, a new biodegradable Mg-Zn-Ca amorphous alloy was prepared, and its corrosion resistance and in vivo biological behaviour were investigated. The main conclusions are as follows:

1. The Mg-Zn-Ca amorphous alloy phases degraded slowly and uniformly in Hank^’s solution, and the morphology remained intact after immersion for 1 month, thus showing excellent corrosion resistance.

2. The degradation rate of the amorphous Mg-Zn-Ca alloy remained stable in vivo, with new bone tissue incorporated into the degraded area. The implant showed strong integration with the surrounding bone tissue, indicating positive effects on bone formation and integration. However, the in vivo degradation rate differed significantly from the corrosion rate observed under simulated in vitro conditions.

3. Blood and vital organ tissue tests confirmed that the degradation products did not cause adverse systemic reactions, showing good in vivo biosafety and excellent potential for the future development and application of orthopaedic implants.

Institutional review board statement

The animal study protocol was approved by the Institutional Ethics Committee of the Affiliated Hospital of Shandong University of Traditional Chinese Medicine (protocol code 2021–67, and date of approval is 11 August 2021)

Disclosure statement

No potential conflict of interest was reported by the author(s).

Additional information

Funding

This work was supported by the National Natural Science Foundation of China (No.51971222), the Natural Science Foundation of Shandong Province (No.ZR2022MH021), and the Clinical Medical Science and Technology Innovation Program of the Jinan Science andTechnology Bureau (202019156).