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The in vitro and in vivo evaluation of the biocompatibility of Mg alloys
This content has been downloaded from IOPscience. Please scroll down to see the full text. 2014 Biomed. Mater. 9 015006 (http://iopscience.iop.org/1748-605X/9/1/015006) View the table of contents for this issue, or go to the journal homepage for more
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Biomedical Materials Biomed. Mater. 9 (2014) 015006 (11pp)
doi:10.1088/1748-6041/9/1/015006
The in vitro and in vivo evaluation of the biocompatibility of Mg alloys J Walker 1,4 , S Shadanbaz 1 , T B F Woodfield 2 , M P Staiger 3 and G J Dias 1 1 2 3
Department of Anatomy and Structural Biology, University of Otago, Dunedin, New Zealand Department of Orthopaedic Surgery, University of Otago, Christchurch, New Zealand Department of Mechanical Engineering, University of Canterbury, Christchurch, New Zealand
E-mail: [email protected] Received 8 August 2013, revised 13 October 2013 Accepted for publication 27 November 2013 Published 16 December 2013 Abstract
Magnesium (Mg) and its alloys are being widely investigated for their potential use as resorbable biomaterials for orthopaedic applications. However, the natural corrosion of the metals results in potentially harmful perturbations to the physiological environment, which requires a comprehensive understanding of their biocompatibility. Currently, most investigations proceed directly from in vitro biocompatibility studies to intraosseous implantation. However, this can result in the unnecessary elimination of appropriate materials due to over sensitive in vitro methods or the implantation of potentially harmful materials. This study involved the development of a relevant in vitro cell culture method, and an in vivo soft tissue implantation technique to provide an intermediate step between basic cell culture methods and large animal intraosseous investigations. A Live/Dead fluorescent assay was used to investigate the viability of both L929 and SaOS-2 cells exposed to Mg alloys, with the results compared to those seen with the intramuscular implantation of the same materials in Lewis rats. These methods were able to successfully provide data on the corrosion of Mg alloys, allowing the identification of slowly and safely corroding materials that may be used in future intraosseous investigations. Keywords: in vitro, in vivo, intramuscular, magnesium, corrosion, cell culture (Some figures may appear in colour only in the online journal)
intraosseous environment. This typically requires expensive and complex surgery utilizing larger animal models such as rabbits, dogs, goats, sheep or pigs [1]. It is therefore important both ethically and financially that the materials selected for such investigations demonstrate excellent biocompatibility. The logical first step towards this goal is to investigate the behaviour of the selected materials in an in vitro cell culture environment. Most commonly, this is carried out by either direct or indirect testing methods adapted from International standards. Although these standards seek to regulate testing, the unique corrosion behaviour of Mg that can result in a cytotoxic increase in pH and osmolality, indicates that these methods may not be appropriate for Mg based materials. This does not entirely invalidate the use of in vitro techniques, provided appropriate testing protocols are adopted to take into account potential environmental perturbations due to Mg
1. Introduction Magnesium (Mg) alloys are now widely being researched for their potential application as orthopaedic biomaterials. The appeal lies in the corrosion that occurs when Mg based materials are exposed to a physiological environment. However, it is the corrosion that results in significant difficulties in assessing and predicting the implant behaviour and the subsequent tissue response. For this reason, it is particularly important that a range of appropriate methodologies are designed and utilized to specifically assess the biocompatibility of Mg based materials. The difficulty in the development of orthopaedic biomaterials in particular, is the necessity to investigate the tissue response in an 4
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corrosion. In vitro methods can still allow the comparison of different Mg based materials to one another. However, the results gathered from in vitro testing cannot effectively replicate an in vivo environment, and therefore should not be used alone to assess the potential biocompatibility of a material. In the field of Mg biomaterials, it is common to proceed directly from such in vitro methods to intraosseous investigations. However, this does not generally allow the assessment of a vast number of materials due to ethical and financial barriers. Furthermore, there are significant discrepancies between the behaviour of Mg based materials in in vitro environments when compared to in vivo [2]. It is therefore important that an intermediate step between the in vitro and intraosseous investigation of Mg alloys be developed. The aim of this study was to develop an appropriate in vitro cell culture method, the results of which could be assessed in conjunction with the results from complimentary in vivo investigations of biocompatibility. To this end an in vitro direct contact method using multiple cell lines was developed and utilized alongside an in vivo intramuscular implantation in a rat model to assess the biocompatibility of four Mg alloys.
Rectangular ingots of pure Mg (99.99%), Mg–0.4Ca, Mg– 0.8Ca, Mg–0.5Mn and Mg–1Zn were cast using a purpose built induction furnace. Medical grade Ti6Al4V was supplied by Enztec Limited. For all materials, samples were machined into discs (10 mm diameter, 2 mm height), and polished to 1200 grit with silicon carbide paper. The samples were then cleaned ultrasonically in 100% ethanol, mild detergent and distilled water before drying and sterilization with gammairradiation (standard dosage 25 kGy, Schering-Plough Animal Health).
period for each cell line). The cells were then left to adhere to the discs for 60 min in a 37 ◦ C, 5% CO2 environment, before 5 mL of the appropriate media was added to each well. The cells were maintained for 24, 48 or 96 h with total media replacement every 24 h. At these time points, assessment of the number of both live and dead cells was carried out using the previously mentioned viability/cytotoxicity assay using the following protocol. The media was removed from each well, and the cells washed twice with Dulbecco’s phosphate buffered saline (DPBS). The wash solution was removed before 100 μL of assay solution containing 4 μM calcein AM and 4 μM ethidium homodimer-1 in DPBS was pipetted onto each disc. These were incubated for 30 min at 37 ◦ C prior to visualization with confocal laser-scanning microscopy (Zeis LSM 710 Confocal Microscope; Zen 2009 software, Carl Zeiss MicroImaging GmBH, Germany). Living cells were identified due to the enzymatic conversion of calcein AM to calcein (excitation 494 nm, emission 517 nm). Dead cells were identified due to the binding of ethidium homodimer-1 to the nucleic acids of cell with damaged cell membranes (excitation 528 nm, emission 617 nm). For quantitative analysis, cell counts were performed on a minimum of three randomly identified fields per disc (850 μm2 per field), and for three discs of each material at each time point (n 9). The random identification of fields, and the subsequent image capturing were performed by a blind, independent party. The viability of cells in each image was calculated using the following formula Number of live cells × 100. Cell viability(%) = Total cell number The average viability for each material was then calculated and presented alongside the average total cell number (number of live cells and dead cells) ( ± SE). A one-way ANOVA with Dunnett’s post hoc analysis was used to compare the cell viability and total cell numbers of all Mg based materials to Ti6Al4V.
2.2. Cellular viability/cytotoxicity evaluation
2.3. Animal implantation
The biocompatibility of the Mg alloys was assessed using a modified direct method with two cell lines in conjunction R cell viability/cytotoxicity assay with the LIVE/DEAD (L-3224, Molecular Probes, Invitrogen). The mouse connective tissue cell line L929, was cultured in MEM supplemented with 10% FBS and 2.2 g L−1 NaHCO3 in a humidified, 37 ◦ C, 5% CO2 environment. The human osteosarcoma cell line SaOS-2, was cultured in MEM-α supplemented with 10% FBS and 2.2 g L−1 NaHCO3 in a humidified, 37 ◦ C, 5% CO2 environment. For the assay, discs of pure Mg, Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, and Ti6Al4V were first immersed in the appropriate culture media for 24 h prior to cell exposure. On experimental day 0, the discs were removed and placed in six-well cell culture plates. Cells were then directly seeded onto each disc at a density of either 6000 SaOS-2, or 2000 L929 cells per disc, suspended in 100 μL of appropriate media (cell densities were previously identified as reaching appropriate levels of confluence over the experimental time
For the in vivo analysis of biocompatibility an intramuscular implantation model in the rat was developed and utilized. Ethical approval was gained from the Otago University Animal Ethics committee (73/10). For this study 48 mature male Lewis rats weighing between 310–452 g were randomly allocated to two groups of 24 for the four and eight week time points investigated. Each group of 24 rats was randomly divided into six groups (four animals per group). Each group of four rats received bilateral intramuscular implants of either pure Mg, Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, or Ti6Al4V discs (i.e. each animal received two implants of the same material). There were therefore four animals implanted with two samples each of a single material for each time point. Anaesthesia was induced with a combination of 4% halothane and 2 L min−1 oxygen, and maintained with a reduced level of 2% halothane and 1 L min−1 oxygen. Analgesic relief was provided by subcutaneous administration of a non-steroidal anti-inflammatory analgesic
2. Materials and methods 2.1. Materials
2
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Table 1. Semi-quantitative grading scale applied to assess the tissue reaction to pure Mg, Mg alloys and Ti–6Al–4V. The grade was allocated
per high-powered field of view (1200 × 900 μm).
Scale 0
1
2
3
4
Lymphocytes Macrophages Neo-vascularization
0 0 0
Rare 1–5 cells Rare 1–5 cells 1–3 capillary profiles
Moderate 5–10 cells Moderate 5–10 cells 4–7 capillary profiles
Hydrogen production
None
Minimal distension, no bubbles
Mild distension and bubbles
Heavy infiltrate Heavy infiltrate Broad band of capillaries Moderate distension and bubbles
Packed Packed Extensive band of capillaries Severe distension and bubbles
(carprofen, 5 mg kg−1) 20 min prior to surgery and 24 h post-operatively. Strepsin (0.1 mL, procaine penicillin G and dihydrostreptomycin sulphate) was injected 30 min prior to surgery as a prophylactic antibiotic. The dorsal lower thoracic region of the rat was shaved, disinfected with chlorhexidine gluconate (Hibitane) and covered with a sterile drape exposing only the surgical area. Surgical scissors were then used to make a midline vertical skin incision from the level of the twelfth thoracic vertebra to approximately 10 mm above the level of the sacrum. Ten millimetre vertical incisions were then made bilaterally in the epimysium of the paravertebral muscles. A pocket approximately 12 mm in length was made laterally by bluntly dissecting between muscle fibres within the muscle belly, and the implants were inserted. The external epimysium was then closed with a single interrupted suture (4/0, RB-1 needle, Ethicon Vicryl Rapide). The skin incision was then closed with 5–6 interrupted sutures as required (3/0, FS-1 needle, Ethicon Vicryl, Amtech Medical, New Zealand). The animals were housed individually and monitored for food intake and hydration for 10 days post surgery, before rehoming in group cages. The animals were then monitored primarily for indications of hydrogen evolution and pain behaviour every 48 h until the conclusion of the investigation. At either 4 or 8 weeks post-implantation, the animals were euthanized by carbon dioxide inhalation. The implant and surrounding tissue were then excised and fixed in 10% neutral buffered formalin (10% NBF) for 48 h for histological analysis. After fixation the discs were carefully removed from the surrounding tissue, immersed in a chromic acid solution (200 g L−1 CrO3 and 10 g L−1 AgNO3) for 15 min to remove corrosion products, and weighed to assess corrosion. Tissue samples were then processed using a standard wax embedding protocol, and 4 μM sections were cut 2.5 mm, 5 mm and 7.5 mm from the medial edge of the disc space with a rotary microtome. The sections were then stained with haematoxylin and eosin (H&E) and visualized with light microscopy. For analysis three randomly identified highmagnification fields were examined (40 × objective lens, each field 1200 × 900 μm). A representative field of view is presented in figure 1. In each field the width of the fibrous capsule was assessed by averaging the depth of the fibrous and inflammatory tissue layer adjacent to the implant at three randomly allocated points. A semi-quantitative scale was used to assess the infiltration of lymphoctyes, macrophages and neovascularization based on the scoring system defined by ISO 10993–6 (2007) (table 1). Other features of tissue
Figure 1. A representative field of view used for the assessment of in vivo biocompatibility. The white space towards the top of the image is the area where the implant was present prior to removal. The black arrow indicates the fibrous capsule that has formed adjacent to the implant. This region is lighter in colour as it is primarily composed of collagen as opposed to the darker skeletal muscle.
reaction that were not measured within this study recognized in ISO 10993–6 such as polymorphonuclear cells, plasma cells, and giant cells were identified extremely rarely, and no tissue necrosis was identified in any of the tissue sections examined. The production of hydrogen gas was also assessed at low magnification with a semi-quantitative scale based on the presence of bubble shaped spaces within the tissue, and distension of the implant space for each entire tissue section (table 1). A one-way ANOVA with Dunnett’s post hoc analysis was used to compare the results of all Mg based materials to those of Ti6Al4V. 3. Results 3.1. Assessing the in vitro biocompatibility of magnesium alloys
The viability results for L929 and SaOS-2 cells cultured on the four Mg alloys, pure Mg and Ti–6Al–4V at 24, 48 and 96 h are presented in figures 2 and 3, respectively. The viability results from each time point must be assessed alongside the total cell numbers present to attain a comprehensive view of the cellular response to the materials investigated. This prevents 3
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R Figure 3. Results from the Live/Dead assay 24, 48, and 96 h after the seeding of SaOS-2 cells onto Ti–6Al–4V, pure Mg or Mg alloy discs. ∗ Indicates a statistically significant difference when the Mg based materials were compared to Ti–6Al–4V (p < 0.01) using a 1-way ANOVA with Dunnett’s post hoc analysis.
R Figure 2. Results from the Live/Dead assay 24, 48, and 96 h after
the seeding of L929 cells onto Ti–6Al–4V, pure Mg or Mg alloy discs. ∗ Indicates a statistically significant difference when the Mg based materials were compared to Ti–6Al–4V (p < 0.01) using a 1-way ANOVA with Dunnett’s post hoc analysis.
A similar trend can be identified after 48 h, with statistically significant difference in viability between Ti– 6Al–4V and Mg–0.4Ca, Mg–0.8Ca and Mg–1Zn. Total cell numbers are similar with a significant difference identifiable between Ti6Al4V and Pure Mg, Mg–0.4Ca, Mg–0.8Ca and Mg–0.5Mn. At this stage Mg–1Zn alone exhibited a total cell number that was not significantly different from the titanium alloy. After 96 h, the total cell numbers present on the titanium alloy had reached 1300 with a viability 99.5%. However, Pure Mg is the only material providing a viability significantly lower than Ti–6Al–4V at this time point. In contrast, all the Mg based materials exhibited a statistically significant reduction in total cell number when compared to Ti–6Al–4V. The results of the SaOS-2 assay indicate similar results. At the 24 and 48 h time points, all the magnesium based materials exhibited significantly reduced viability and total cell number when compared to Ti–6Al–4V, with the viability of Pure Mg the only exception. After 96 h, the same trend is followed with the only difference being the lack of a significant reduction
the misrepresentation of a material as cytocompatible, if for example 80% of the cells are viable, but only five cells are present (the low numbers of cells would therefore represent significant toxicity). The results for the L929 cells at 24 h show a statistically significant difference in the viability of cells between Ti– 6Al–4V and Mg–0.4Ca, Mg–0.8Ca and Mg–1Zn (p < 0.01). Whereas at this time point there was no significant difference between Ti–6Al–4V and Pure Mg or Mg–0.5Mn (p > 0.01). However, the total cell numbers indicate a statistically significant difference between Ti–6Al–4V and all of the Mg based materials (p < 0.01). This reduction in total cell number for all Mg based materials at this early stage is indicative of difficulties in the initial adherence of cells to the Mg based substrates. This reduced initial attachment has important ramifications on the total cell numbers present at later time points. Figure 4 displays representative confocal images of live and dead L929 cells after 96 h cultured on Mg based materials or Ti–6Al–4V. 4
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(A)
(B)
(A)
(C)
(D)
(C )
(E)
(F)
(E )
(B)
(D)
(F )
Figure 4. Confocal micrographs of L929 cells cultured on the surface of Mg alloy, pure Mg and Ti–6Al–4V after 96 h. Green profiles indicate living cells (calcein AM). Red profiles indicate dying or dead cells (ethidium homodimer-1). (A) Mg–0.4Ca. (B) Mg–0.8Ca. (C) Mg–0.5Mn. (D) Mg–1Zn. (E). Pure Mg. (F) Ti–6Al–4V.
Figure 5. Confocal micrographs of SaOS-2 cells cultured on the surface of Mg alloy, pure Mg and Ti–6Al–4V after 96 h. Green profiles indicate living cells (calcein AM). Red profiles indicate dying or dead cells (ethidium homodimer-1). (A) Mg–0.4Ca. (B) Mg–0.8Ca. (C) Mg–0.5Mn. (D) Mg–1Zn. (E). Pure Mg. (F) Ti–6Al–4V.
between the total cell numbers on Ti–6Al–4V when compared to Mg–1Zn. Figure 5 displays representative confocal images of live and dead SaOS-2 cells after 96 h cultured on Mg based materials or Ti–6Al–4V.
are presented in figure 7. In concurrence with the results from the fibrous encapsulation, and lymphocyte and macrophage infiltration, there were no statistically significant differences between the average grade allocated for neovascularization when Ti–6Al–4V was compared to the Mg based materials (p > 0.01). In contrast, there was a statistically significant difference between Ti–6Al–4V and all of the Mg based materials for hydrogen production and corrosion (p < 0.01). However, this result was expected as the titanium alloy neither corrodes nor produces hydrogen gas. Figure 8 shows low magnification micrographs indicating tissue distension in response to hydrogen production with the implantation of Ti–6Al–4V and Mg–0.4Ca.
3.2. The biocompatibility of magnesium alloys in an intramuscular environment
The average width of the fibrous capsule, and the average grade allocated for lymphocyte and macrophage infiltration in association with the intramuscular implantation of Ti–6Al–4V, Pure Mg, Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, and Mg–1Zn at 4 and 8 weeks are presented in figure 6. Statistical analysis indicated no significant differences between Ti–6Al–4V and the Mg based materials for any variable at either time point (p > 0.01). The average grade allocated for neovascularization, hydrogen production and corrosion in association with the intramuscular implantation of Ti–6Al–4V, Pure Mg, Mg– 0.4Ca, Mg–0.8Ca, Mg–0.5Mn, and Mg–1Zn at 4 and 8 weeks
4. Discussion 4.1. General comments on the In Vitro methodology
A multitude of in vitro cell culture techniques exist for the assessment of the biocompatibility of magnesium as 5
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Figure 6. The average width of the fibrous capsule, and average grades allocated to lymphocyte infiltration and macrophage infiltration in the tissue adjacent to Ti–6Al–4V, pure Mg or Mg alloy implants. There were no statistically significant differences for any measure when the Mg based materials were compared to Ti–6Al–4V (p > 0.01) using a 1-way ANOVA with Dunnett’s post hoc analysis.
a biomaterial. Primarily, these techniques are based on the methods outlined by international standards with slight variations in experimental set-up and the assessment of cellular
behaviour. In particular, ISO-10993–5 suggests the use of either an indirect method in which cells are exposed to extracts of the materials, or the direct method in which the 6
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Figure 7. The average grade allocated to neovascularization, hydrogen production, and implant corrosion with implantation of Ti–6Al–4V, pure Mg or Mg alloy implants. There were no statistically significant differences between the Mg based materials and Ti–6Al–4V for neovascularization (p > 0.01, 1-way ANOVA with Dunnett’s post hoc analysis). All Mg based materials exhibited statistically significant increases in both hydrogen production and corrosion when compared to Ti–6Al–4V (p < 0.01, 1-way ANOVA with Dunnett’s post hoc analysis).
cells come into contact with the materials [3]. Unfortunately, the standards currently available are designed for the assessment of the in vitro biocompatibility of primarily non-
degrading biomaterials. This leads to difficulties in applying these methods to rapidly corroding materials such as Mg alloys. 7
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factors including both the pH and osmolality of the cell culture environment, reducing the value of the results when used with Mg based materials [20, 24]. For the above reasons, a modified direct method using R the Live/Dead assay was adopted for this investigation. The choice to seed cells directly onto the surfaces of the materials was based upon a desire to assess whether cells could attach and stay viable on an actively corroding material. This was preferable to the protocol outlined in ISO 10993–5 for the direct method in which the test material is placed onto an adherent cell layer, as it was believed that for Mg this was too similar to the indirect method in which the pH change and increase in osmolality would have the greatest influence on cell survival. Additionally, no evidence existed that the corroding Mg materials would interfere with the calcein AM or ethidium homodimer utilized for the assay. Several other aspects of the methodology used within this investigation require comment. Firstly, the materials were pre-soaked to allow the formation of a passivation layer on the surface of the implant. Whilst it could be argued that this step is masking the potentially toxic effects of the rapid initial corrosion of the materials, initial experiments indicated that without this allowance initial cellular adhesion would be greatly diminished. Eliminating this step would drastically reduce any ability to investigate the in vitro biocompatibility of Mg alloys, and it was therefore included in the protocol. Secondly, it is important to note that the media for each material was replaced daily. This step was taken to reduce any toxicity occurring due to an increase in the pH of the static cell culture environment. Again, this could be argued to be masking the toxic effect of the pH change associated with Mg corrosion. However, it is believed that in a dynamic in vivo environment, the pH in the area associated with a Mg alloy implant would not fluctuate as significantly as in an in vitro environment. However, there would still be a marked increase in pH at the surface of the material in vitro that would also occur in an in vivo environment. Therefore, the media replacement regime utilized should not have further increased the innate discrepancies between in vitro and in vivo methodologies. Thirdly, the choice of cells used in the investigation of in vitro biocompatibility is important. In this study two immortalized cell lines were used, L929 (mouse fibroblast) and SaOS-2 (human osteosarcoma). The L929 line was selected as a standardized, and commonly employed cell type for biocompatibility studies [25–28]. The SaOS-2 cells were also included as a cell line derived from the more clinically applicable human osteosarcoma [29]. These cells are also commonly used for the assessment of the biocompatibility of potential orthopaedic materials due to their osteoblastic phenotype, ability to differentiate and produce mineralized extracellular matrix [30–33]. Within this experiment the SaOS2 cells were not exposed to a differentiating media, which in retrospect could have improved adherence, and therefore increased cell number. Although this could potentially have altered the results gathered, the use of both cell lines is still of vital importance as the cellular response to any given material has been observed to differ vastly depending on the cell type [8, 17, 29, 34, 35].
(A)
(B)
Figure 8. Low magnification micrographs of the implant spaces left after the removal of an intramuscular implant (H & E). (A) The white space is the area left after removal of a Ti–6Al–4V implant with clearly identifiable margins within the tissue. This received a grade of 0 on the previously identified scoring system for hydrogen evolution. (B) The space left after removal of a Mg–0.4Ca implant with some distension of the space, and obscured margins due to probable hydrogen bubbles within the tissue. This received a grade 3 on the previously identified scoring system.
The indirect method is used relatively frequently with Mg based materials in one of two ways. Either extracts are produced and diluted to various concentrations [4–16], or a modification to the method is adopted in which metal salts are added to the culture media to replicate the toxicity of the released Mg and alloying component ions [17–19]. The primary issue with the first of these methods is that any toxicity identified is primarily due to the osmolality of the solution [20]. Whilst this can be argued to be of primary importance to the biocompatibility of the material, it is likely that in an in vivo environment constant diffusion would mitigate this effect to a degree. Additionally, when this method is adopted, the extracts are often diluted to a point for which the increased osmolality is no longer toxic [4, 7, 10, 14, 16], somewhat nullifying the use of the method in the first place. The production of an extract solution also requires the immersion of a Mg sample in media for a period of time. The resulting corrosion that occurs would likely cause an increase in the pH of the extract solution [8, 21], which left unadjusted would certainly result in cytotoxicity. However, this point is often left unmentioned in studies using the technique [6, 9, 10, 12–16, 22], or there is reliance on pH adjustment to prevent this occurrence [5]. The aforementioned addition of Mg and alloying component salts dissolved in cell culture media is also problematic due to the effects on the osmolality of the solution. This method can be of use when investigating alloying elements not found within cell culture media such as rare earth elements, but is less useful for assessing the effect of ions already found within physiological solutions such as Mg and Ca. Additionally, the most common technique used to assess the viability of cells with the indirect method is to use a colorimetric assay based on the reduction of tetrazolium salts to formazan dyes within living cells [7–9, 11, 12, 14– 19, 22, 23]. However, this method can be affected by various 8
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the cytocompatibility of pure Mg show variable results, from extremely toxic [21, 38] to highly biocompatible [36, 37]. This highlights the continuing difficulties with comparing the behaviour of Mg based materials between in vitro investigations in which small differences in variables such as surface roughness, cell type, media type or surface area to volume ratio can dramatically affect corrosion behaviour, and therefore cellular behaviour. Even less evidence exists in the literature for the behaviour of cells on the relevant Mg alloys. One study indicated a significantly reduced total protein concentration, for a Mg– 0.5Ca alloy when compared to a negative control (tissue culture slide). The same study also indicated Mg–0.8Ca provided a result statistically indistinct to the control, perhaps suggesting Mg–0.8Ca was more biocompatible than Mg– 0.5Ca [36]. These results are somewhat in contrast to those seen within this investigation in which the lower calcium percentage-containing alloy generally exhibited slightly improved biocompatibility. However this experiment utilized total protein primarily as an indicator of osteogenic capacity rather than to assess cell number, and was also measured after eight days in culture, both of which make comparison to this investigation difficult. The in vitro biocompatibility of a binary Mg–6Zn alloy has also been investigated, with results showing acceptable adhesion and some cellular division [39, 40]. However these studies do not compare the results to an inert control material preventing any comparison to other investigations. No research has been published investigating the in vitro biocompatibility of a binary Mg–Mn alloy. There are many inherent difficulties in both testing the biocompatibility of Mg alloys in vitro, and attempting to compare the results between studies. If a conservative approach was taken in the assessment of the results seen within this investigation, Mg alloys would be assumed highly toxic materials. However, use of similar alloys and pure Mg in an in vivo environment has indicated good biocompatibility [15, 22, 41]. It is therefore important that a more liberal interpretation of the in vitro results be taken, and to ensure that the conclusions from such studies be made alongside other evidence such as the results from in vivo investigations.
The corrosion associated with Mg alloys, and the resulting by-products of this corrosion make the assessment of any aspect of alloy behaviour difficult in an in vitro environment. The assessment of in vitro biocompatibility is particularly difficult, due to the sensitive nature of cells growing in culture. The points mentioned above highlight these issues and the difficulties in developing methods that avoid cell death due to environmental perturbations, without disguising the potential cytotoxic effects of the Mg alloys. For these reasons the assessment of the biocompatibility of the Mg alloys investigated in this study focused on the viability/cytotoxicity. More in depth investigations of cellular behaviour and differentiation are commonly carried out. However, it is believed that the discrepancies and difficulties in replicating an in vivo environment in vitro, is likely to provide results for these studies only barely comparable to those that would be seen with in vivo implantation of the materials. 4.2. Comparison between the in vitro biocompatibility of Mg alloys
The results of the in vitro analysis of the biocompatibility of Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, pure Mg and Ti– 6Al–4V indicate a general trend towards lower viability and total cell number on the Mg based materials when compared to Ti–6Al–4V. This result was not unexpected due to the active corrosion occurring when the Mg based materials are exposed to a physiological solution. The resultant increase in osmolality, increase in pH and the physical disruption to cells by hydrogen gas evolution, are all factors that would reduce cellular adhesion and viability in a closed cell culture environment [21]. However, despite a relatively inhospitable environment, some cellular adhesion and active cell division did occur when cells were seeded directly onto the Mg alloys being investigated. Even at the reduced viability and total cell numbers observed, this can be seen as an exceptionally positive result for the potential biocompatibility of these alloys in the more dynamic and adjustable in vivo environment. Consequently, despite the significant differences between the results from the Ti–6Al–4V and the Mg based materials, the results of these studies can still be used to an extent for the comparison of the in vitro biocompatibility of the Mg alloys. The results from the L929 experiments generally indicated relatively good performance of the Mg–0.5Mn and Mg–1Zn alloys with increasing time. The cells on these two materials appeared to recover after the initial low total cell number seen at 24 h. In contrast, the Ca containing alloys exhibited very low total cell counts for the entirety of the experiment. Similar results were observed with the SaOS-2 cells for which much higher total cell numbers were seen with Mg–0.5Mn, Mg–1Zn and pure Mg when compared to the Ca containing alloys. The assessment of the in vitro biocompatibility of alloys comparable to those investigated in this study is relatively rare, and more infrequent still is the use of a direct method. Most commonly pure Mg is included as a control material in studies investigating a range of alloys [5, 36], surface treatments [21, 37], coatings [38], or fabrication methods [16]. The studies using a direct method for the assessment of
4.3. General comments on the in vivo methodology
Analysis of the biocompatibility of Mg alloys in a soft tissue environment is relatively uncommon, with the majority of studies progressing from in vitro corrosion and biocompatibility investigations directly to their assessment in an intraosseous location. The reason for this avoidance of standard biocompatibility protocols remains unclear. Both ISO and ASTM standards outline protocols for the short-term assessment of the local effects of implantation of biomaterials in either a subcutaneous or intramuscular location [42–44]. For this investigation, the inclusion of a soft tissue study was deemed important for several reasons. Firstly, it was hypothesized that an intramuscular environment would be able to tolerate the production of hydrogen gas better than a rigid intraosseous environment, a factor considered important given the limited knowledge available regarding the effect 9
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of hydrogen production in an in vivo environment. Secondly, soft tissue implantation provided the ability to accurately assess the corrosion of the materials by mass loss. Thirdly, many orthopaedic biomaterials will directly contact muscle and connective tissue as well as bone. The assessment of the reaction of soft tissues to Mg alloys being developed for orthopaedic application is therefore important.
materials, and the inappropriate exclusion of potentially useful materials due to conservative in vitro corrosion analysis. Additionally it was identified that whilst in vitro techniques can be used as an indicator of the biocompatibility of Mg alloys, the results must be analysed with an understanding that the corrosion of the materials can lead to false positive results for toxicity. It is therefore important that in vitro analysis of biocompatibility be assessed alongside the results of a corresponding in vivo method.
4.4. Comparison between the in vivo biocompatibility of Mg alloys
The in vivo biocompatibility of Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, pure Mg and Ti–6Al–4V in an intramuscular environment was analysed through the assessment of the width of the fibrous capsule, the lymphocyte and macrophage infiltration, and neovascularization in the tissue adjacent to the implants. An overwhelming trend could be identified for all these measures indicating no significant differences between any of the Mg based materials and the Ti–6Al–4V. As this Ti alloy is widely used as a medical implant material in both medicine and dentistry [45–47], these results indicate comparable biocompatibility for these Mg alloys. The assessment of in vivo biocompatibility of Mg alloys in a soft tissue environment is rare, with very few articles available for the comparison of results. The majority of those that have been performed rely on brief descriptive results of the histology to indicate the tissue reaction to the materials [13, 48–50]. One study however, has thoroughly assessed the reaction of muscle tissue to a Mg–0.8Ca implant, using a semi-quantitative scale to assess fibrous encapsulation, tissue cavity formation, necrosis and the infiltration of macrophages, giant cells, heterophilic granulocytes, B-lymphocytes and T-lymphoctyes [51]. This extensive analysis indicated a similar result to that seen within this investigation, with Mg–0.8Ca performing similarly to a stainless steel 316L control material.
References [1] Pearce A I, Richards R G, Milz S, Schneider E and Pearce S G 2007 Animal models for implant biomaterial research in bone: a review Eur. Cells Mater. 13 1–10 [2] Witte F et al 2006 In vitro and in vivo corrosion measurements of magnesium alloys Biomaterials 27 1013–8 [3] ISO 1999 Biological Evalutation of Medical Devices Part 5: Tests for In Vitro Cytotoxicity ISO 10993–5 (Geneva: International Organization for Standardization) [4] Fischer J, Pr¨ofrock D, Hort N, Willumeit R and Feyerabend F 2011 Reprint of: Improved cytotoxicity testing of magnesium materials Mater. Sci. Eng. B 176 1773–7 [5] Yang C, Yuan G, Zhang J, Tang Z, Zhang X and Dai K 2010 Effects of magnesium alloys extracts on adult human bone marrow-derived stromal cell viability and osteogenic differentiation Biomed. Mater. 5 045005 [6] Zheng Y F, Gu X N, Xi Y L and Chai D L 2010 In vitro degradation and cytotoxicity of Mg/Ca composites produced by powder metallurgy Acta Biomater. 6 1783–91 [7] Wang D-W, Cao Y, Qiu H and Bi Z-G 2011 Improved blood compatibility of Mg–1.0Zn–1.0Ca alloy by micro-arc oxidation J. Biomed. Mater. Res. A 99 166–72 [8] Gu X, Zheng Y, Cheng Y, Zhong S and Xi T 2009 In vitro corrosion and biocompatibility of binary magnesium alloys Biomaterials 30 484–98 [9] Huan Z G, Leeflang M a, Zhou J, Fratila-Apachitei L E and Duszczyk J 2010 In vitro degradation behavior and cytocompatibility of Mg–Zn–Zr alloys J. Mater. Sci., Mater. Med. 21 2623–35 [10] Li Y et al 2012 Mg–Zr–Sr alloys as biodegradable implant materials Acta Biomater. 8 3177–88 [11] Park R S et al 2012 Corrosion behavior and cytotoxicity of Mg–35Zn–3Ca alloy for surface modified biodegradable implant material J. Biomed. Mater. Res. B 100 911–23 [12] Wang Y et al 2012 In vitro degradation and biocompatibility of Mg–Nd–Zn–Zr alloy Chin. Sci. Bull. 57 2163–70 [13] H¨anzi A C, Gerber I, Schinhammer M, L¨offler J F and Uggowitzer P J 2010 On the in vitro and in vivo degradation performance and biological response of new biodegradable Mg–Y–Zn alloys Acta Biomater. 6 1824–33 [14] Zhang E, Yin D, Xu L, Yang L and Yang K 2009 Microstructure, mechanical and corrosion properties and biocompatibility of Mg–Zn–Mn alloys for biomedical application Mater. Sci. Eng. C 29 987–93 [15] Li Z, Gu X, Lou S and Zheng Y 2008 The development of binary Mg–Ca alloys for use as biodegradable materials within bone Biomaterials 29 1329–44 [16] Gu X N, Zhou W R, Zheng Y F, Liu Y and Li Y X 2010 Degradation and cytotoxicity of lotus-type porous pure magnesium as potential tissue engineering scaffold material Mater. Lett. 64 1871–4 [17] Feyerabend F et al 2010 Evaluation of short-term effects of rare earth and other elements used in magnesium alloys on primary cells and cell lines Acta Biomater. 6 1834–42
4.5. The corrosion of Mg alloys in an intramuscular environment
One of the benefits in performing a soft tissue implantation of Mg alloys was the ability to remove the samples and assess the corrosion by gravimetric analysis. The results indicated Mg–1Zn as corroding slower than the other Mg based materials at the four-week time point. After eight-weeks of implantation, the corrosion of the alloys followed a similar trend, indicating Mg–1Zn to be the slowest corroding of the alloys investigated. 5. Conclusion The primary aim of the in vitro and in vivo investigation of Mg alloy biocompatibility was to develop and refine a series of simple, rapid and reproducible methods to assess Mg alloy biocompatibility. The provision of an intermediate step between in vitro corrosion analysis and intraosseous implantation of Mg alloys allows the investigation of a larger variety of materials in an in vivo location without the expense and complexity of an intraosseous study. This could prevent both the intraosseous implantation of potentially harmful 10
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[34] Wataha JC, Hanks CT and Sun Z 1994 Effect of cell line on in vitro metal ion cytotoxicity Dent. Mater. 10 156–61 [35] Thonemann B, Schmalz G, Hiller K and Schweikl H 2002 Responses of L929 mouse fibroblasts, primary and immortalized bovine dental papilla-derived cell lines to dental resin components Dent. Mater. 18 318–23 [36] Pietak A, Mahoney P, Dias G J and Staiger M P 2008 Bone-like matrix formation on magnesium and magnesium alloys J. Mater. Sci., Mater. Med. 19 407–15 [37] Li L, Gao J and Wang Y 2004 Evaluation of cyto-toxicity and corrosion behaviour of alkali-heat-treated magnesium in simulated body fluid Surf. Coat. Technol. 185 92–8 [38] Jo J-H et al 2011 Hydroxyapatite coating on magnesium with MgF(2) interlayer for enhanced corrosion resistance and biocompatibility J. Mater. Sci., Mater. Med. 22 2437–47 [39] Li J et al 2010 In vitro responses of human bone marrow stromal cells to a fluoridated hydroxyapatite coated biodegradable Mg–Zn alloy Biomaterials 31 5782–8 [40] Zhang S X et al 2009 In vitro degradation, hemolysis and MC3T3-E1 cell adhesion of biodegradable Mg–Zn alloy Mater. Sci. Eng. C 29 1907–12 [41] Xu L P, Yu G N, Zhang E, Pan F and Yang K 2007 In vivo corrosion behavior of Mg–Mn–Zn alloy for bone implant application J. Biomed. Mater. Res. A 83 703–11 [42] ISO 2007 Biological Evaluation of Medical Devices Part 6: Tests for local effects after implantation ISO 10993–6 (Geneva: International Organization for Standardization) [43] ASTM 2004 Standard Practice for Short-Term Screening of Implant Materials ASTM F763–04 (West Conshohocken, PA: American Society for Testing and Materials) [44] ASTM 2008 Standard Practice for Subcutaneous Screening Test for Implant Materials ASTM 1408–97 (West Conshohocken, PA: American Society for Testing and Materials) [45] Deligianni D D, Katsala N, Ladas S, Sotiropoulou D, Amedee J and Missirlis Y F 2001 Effect of surface roughness of the titanium alloy Ti–6Al–4V on human bone marrow cell response and on protein adsorption Biomaterials 22 1241–51 [46] Park J-W, Kim H-K, Kim Y-J, Jang J-H, Song H and Hanawa T 2010 Osteoblast response and osseointegration of a Ti–6Al–4V alloy implant incorporating strontium Acta Biomater. 6 2843–51 [47] Niinomi M 1998 Mechanical properties of biomedical titanium alloys Mater. Sci. Eng. A 243 231–6 [48] Hussl H, Papp C, H¨opfel-Kreiner I, Rumpl E and Wilflingseder P 1981 Resorption time and tissue reactions with magnesium rods in rats and rabbits Eur. J. Plast. Surg. 6 117–26 [49] Xue D, Yun Y, Tan Z, Dong Z and Schulz M J 2012 In vivo and in vitro degradation behavior of magnesium alloys as biomaterials J. Mater. Sci. Technol. 28 261–7 [50] Aghion E, Levy G and Ovadia S 2012 In vivo behavior of biodegradable Mg–Nd–Y–Zr–Ca alloy J. Mater. Sci., Mater. Med. 23 805–12 [51] Erdmann N et al 2010 Evaluation of the soft tissue biocompatibility of MgCa0.8 and surgical steel 316L in vivo: a comparative study in rabbits Biomed. Eng. online 9 63
[18] Feser K, Kietzmann M, B¨aumer W, Krause C and Bach F W 2011 Effects of degradable Mg–Ca alloys on dendritic cell function J. Biomater. Appl. 25 685–97 [19] Zhang E, Yang L, Xu J and Chen H 2010 Microstructure, mechanical properties and bio-corrosion properties of Mg-Si(-Ca, Zn) alloy for biomedical application Acta Biomater. 6 1756–62 [20] Witte F et al 2008 Degradable biomaterials based on magnesium corrosion Curr. Opin. Solid State Mater. Sci. 12 63–72 [21] Lorenz C, Brunner J G, Kollmannsberger P, Jaafar L, Fabry B and Virtanen S 2009 Effect of surface pre-treatments on biocompatibility of magnesium Acta Biomater. 5 2783–9 [22] Zhang S et al 2010 Research on an Mg–Zn alloy as a degradable biomaterial Acta Biomater. 6 626–40 [23] Witte F et al 2007 Biodegradable magnesiumhydroxyapatite metal matrix composites Biomaterials 28 2163–74 [24] Fischer J, Prosenc MH, Wolff M, Hort N, Willumeit R and Feyerabend F 2010 Interference of magnesium corrosion with tetrazolium-based cytotoxicity assays Acta Biomater. 6 1813–23 [25] Turner T D, Spyratou O and Schmidt R J 1989 Biocompatibility of wound management products: standardization of and determination of cell growth rate in L929 fibroblast cultures J. Pharm. Pharmacol. 41 775–80 [26] Acharya C, Ghosh S K and Kundu S C 2008 Silk fibroin protein from mulberry and non-mulberry silkworms: cytotoxicity, biocompatibility and kinetics of L929 murine fibroblast adhesion J. Mater. Sci., Mater. Med. 19 2827–36 [27] Jorge J H, Giampaolo E T, Vergani C E, Machado A L, Pavarina A C and Carlos I Z 2007 Biocompatibility of denture base acrylic resins evaluated in culture of L929 cells. Effect of polymerisation cycle and postpolymerisation treatments Gerodontology 24 52–7 [28] Serrano M C et al 2004 In vitro biocompatibility assessment of poly(epsilon-caprolactone) films using L929 mouse fibroblasts Biomaterials 25 5603–11 [29] Kirkpatrick C J, Krump-Konvalinkova V, Unger R E, Bittinger F, Otto M and Peters K 2002 Tissue response and biomaterial integration: the efficacy of in vitro methods Biomol. Eng. 19 211–7 [30] Anderson H C et al 1995 The mechanism of bone induction and bone healing by human osteosarcoma cell extracts Clin. Orthop. Relat. Res. 313 129–34 [31] Okumura A et al 2001 Substrate affects the initial attachment and subsequent behavior of human osteoblastic cells (Saos-2) Biomaterials 22 2263–71 [32] Gronowicz G and McCarthy M B 1996 Response of human osteoblasts to implant materials: integrin-mediated adhesion J. Orthop. Res. 14 878–87 [33] Yu X, Botchwey E a, Levine E M, Pollack S R and Laurencin C T 2004 Bioreactor-based bone tissue engineering: the influence of dynamic flow on osteoblast phenotypic expression and matrix mineralization Proc. Natl Acad. Sci. USA 101 11203–8
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The in vitro and in vivo evaluation of the biocompatibility of Mg alloys
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Biomedical Materials Biomed. Mater. 9 (2014) 015006 (11pp)
doi:10.1088/1748-6041/9/1/015006
The in vitro and in vivo evaluation of the biocompatibility of Mg alloys J Walker 1,4 , S Shadanbaz 1 , T B F Woodfield 2 , M P Staiger 3 and G J Dias 1 1 2 3
Department of Anatomy and Structural Biology, University of Otago, Dunedin, New Zealand Department of Orthopaedic Surgery, University of Otago, Christchurch, New Zealand Department of Mechanical Engineering, University of Canterbury, Christchurch, New Zealand
E-mail: [email protected] Received 8 August 2013, revised 13 October 2013 Accepted for publication 27 November 2013 Published 16 December 2013 Abstract
Magnesium (Mg) and its alloys are being widely investigated for their potential use as resorbable biomaterials for orthopaedic applications. However, the natural corrosion of the metals results in potentially harmful perturbations to the physiological environment, which requires a comprehensive understanding of their biocompatibility. Currently, most investigations proceed directly from in vitro biocompatibility studies to intraosseous implantation. However, this can result in the unnecessary elimination of appropriate materials due to over sensitive in vitro methods or the implantation of potentially harmful materials. This study involved the development of a relevant in vitro cell culture method, and an in vivo soft tissue implantation technique to provide an intermediate step between basic cell culture methods and large animal intraosseous investigations. A Live/Dead fluorescent assay was used to investigate the viability of both L929 and SaOS-2 cells exposed to Mg alloys, with the results compared to those seen with the intramuscular implantation of the same materials in Lewis rats. These methods were able to successfully provide data on the corrosion of Mg alloys, allowing the identification of slowly and safely corroding materials that may be used in future intraosseous investigations. Keywords: in vitro, in vivo, intramuscular, magnesium, corrosion, cell culture (Some figures may appear in colour only in the online journal)
intraosseous environment. This typically requires expensive and complex surgery utilizing larger animal models such as rabbits, dogs, goats, sheep or pigs [1]. It is therefore important both ethically and financially that the materials selected for such investigations demonstrate excellent biocompatibility. The logical first step towards this goal is to investigate the behaviour of the selected materials in an in vitro cell culture environment. Most commonly, this is carried out by either direct or indirect testing methods adapted from International standards. Although these standards seek to regulate testing, the unique corrosion behaviour of Mg that can result in a cytotoxic increase in pH and osmolality, indicates that these methods may not be appropriate for Mg based materials. This does not entirely invalidate the use of in vitro techniques, provided appropriate testing protocols are adopted to take into account potential environmental perturbations due to Mg
1. Introduction Magnesium (Mg) alloys are now widely being researched for their potential application as orthopaedic biomaterials. The appeal lies in the corrosion that occurs when Mg based materials are exposed to a physiological environment. However, it is the corrosion that results in significant difficulties in assessing and predicting the implant behaviour and the subsequent tissue response. For this reason, it is particularly important that a range of appropriate methodologies are designed and utilized to specifically assess the biocompatibility of Mg based materials. The difficulty in the development of orthopaedic biomaterials in particular, is the necessity to investigate the tissue response in an 4
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Biomed. Mater. 9 (2014) 015006
corrosion. In vitro methods can still allow the comparison of different Mg based materials to one another. However, the results gathered from in vitro testing cannot effectively replicate an in vivo environment, and therefore should not be used alone to assess the potential biocompatibility of a material. In the field of Mg biomaterials, it is common to proceed directly from such in vitro methods to intraosseous investigations. However, this does not generally allow the assessment of a vast number of materials due to ethical and financial barriers. Furthermore, there are significant discrepancies between the behaviour of Mg based materials in in vitro environments when compared to in vivo [2]. It is therefore important that an intermediate step between the in vitro and intraosseous investigation of Mg alloys be developed. The aim of this study was to develop an appropriate in vitro cell culture method, the results of which could be assessed in conjunction with the results from complimentary in vivo investigations of biocompatibility. To this end an in vitro direct contact method using multiple cell lines was developed and utilized alongside an in vivo intramuscular implantation in a rat model to assess the biocompatibility of four Mg alloys.
Rectangular ingots of pure Mg (99.99%), Mg–0.4Ca, Mg– 0.8Ca, Mg–0.5Mn and Mg–1Zn were cast using a purpose built induction furnace. Medical grade Ti6Al4V was supplied by Enztec Limited. For all materials, samples were machined into discs (10 mm diameter, 2 mm height), and polished to 1200 grit with silicon carbide paper. The samples were then cleaned ultrasonically in 100% ethanol, mild detergent and distilled water before drying and sterilization with gammairradiation (standard dosage 25 kGy, Schering-Plough Animal Health).
period for each cell line). The cells were then left to adhere to the discs for 60 min in a 37 ◦ C, 5% CO2 environment, before 5 mL of the appropriate media was added to each well. The cells were maintained for 24, 48 or 96 h with total media replacement every 24 h. At these time points, assessment of the number of both live and dead cells was carried out using the previously mentioned viability/cytotoxicity assay using the following protocol. The media was removed from each well, and the cells washed twice with Dulbecco’s phosphate buffered saline (DPBS). The wash solution was removed before 100 μL of assay solution containing 4 μM calcein AM and 4 μM ethidium homodimer-1 in DPBS was pipetted onto each disc. These were incubated for 30 min at 37 ◦ C prior to visualization with confocal laser-scanning microscopy (Zeis LSM 710 Confocal Microscope; Zen 2009 software, Carl Zeiss MicroImaging GmBH, Germany). Living cells were identified due to the enzymatic conversion of calcein AM to calcein (excitation 494 nm, emission 517 nm). Dead cells were identified due to the binding of ethidium homodimer-1 to the nucleic acids of cell with damaged cell membranes (excitation 528 nm, emission 617 nm). For quantitative analysis, cell counts were performed on a minimum of three randomly identified fields per disc (850 μm2 per field), and for three discs of each material at each time point (n 9). The random identification of fields, and the subsequent image capturing were performed by a blind, independent party. The viability of cells in each image was calculated using the following formula Number of live cells × 100. Cell viability(%) = Total cell number The average viability for each material was then calculated and presented alongside the average total cell number (number of live cells and dead cells) ( ± SE). A one-way ANOVA with Dunnett’s post hoc analysis was used to compare the cell viability and total cell numbers of all Mg based materials to Ti6Al4V.
2.2. Cellular viability/cytotoxicity evaluation
2.3. Animal implantation
The biocompatibility of the Mg alloys was assessed using a modified direct method with two cell lines in conjunction R cell viability/cytotoxicity assay with the LIVE/DEAD (L-3224, Molecular Probes, Invitrogen). The mouse connective tissue cell line L929, was cultured in MEM supplemented with 10% FBS and 2.2 g L−1 NaHCO3 in a humidified, 37 ◦ C, 5% CO2 environment. The human osteosarcoma cell line SaOS-2, was cultured in MEM-α supplemented with 10% FBS and 2.2 g L−1 NaHCO3 in a humidified, 37 ◦ C, 5% CO2 environment. For the assay, discs of pure Mg, Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, and Ti6Al4V were first immersed in the appropriate culture media for 24 h prior to cell exposure. On experimental day 0, the discs were removed and placed in six-well cell culture plates. Cells were then directly seeded onto each disc at a density of either 6000 SaOS-2, or 2000 L929 cells per disc, suspended in 100 μL of appropriate media (cell densities were previously identified as reaching appropriate levels of confluence over the experimental time
For the in vivo analysis of biocompatibility an intramuscular implantation model in the rat was developed and utilized. Ethical approval was gained from the Otago University Animal Ethics committee (73/10). For this study 48 mature male Lewis rats weighing between 310–452 g were randomly allocated to two groups of 24 for the four and eight week time points investigated. Each group of 24 rats was randomly divided into six groups (four animals per group). Each group of four rats received bilateral intramuscular implants of either pure Mg, Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, or Ti6Al4V discs (i.e. each animal received two implants of the same material). There were therefore four animals implanted with two samples each of a single material for each time point. Anaesthesia was induced with a combination of 4% halothane and 2 L min−1 oxygen, and maintained with a reduced level of 2% halothane and 1 L min−1 oxygen. Analgesic relief was provided by subcutaneous administration of a non-steroidal anti-inflammatory analgesic
2. Materials and methods 2.1. Materials
2
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Table 1. Semi-quantitative grading scale applied to assess the tissue reaction to pure Mg, Mg alloys and Ti–6Al–4V. The grade was allocated
per high-powered field of view (1200 × 900 μm).
Scale 0
1
2
3
4
Lymphocytes Macrophages Neo-vascularization
0 0 0
Rare 1–5 cells Rare 1–5 cells 1–3 capillary profiles
Moderate 5–10 cells Moderate 5–10 cells 4–7 capillary profiles
Hydrogen production
None
Minimal distension, no bubbles
Mild distension and bubbles
Heavy infiltrate Heavy infiltrate Broad band of capillaries Moderate distension and bubbles
Packed Packed Extensive band of capillaries Severe distension and bubbles
(carprofen, 5 mg kg−1) 20 min prior to surgery and 24 h post-operatively. Strepsin (0.1 mL, procaine penicillin G and dihydrostreptomycin sulphate) was injected 30 min prior to surgery as a prophylactic antibiotic. The dorsal lower thoracic region of the rat was shaved, disinfected with chlorhexidine gluconate (Hibitane) and covered with a sterile drape exposing only the surgical area. Surgical scissors were then used to make a midline vertical skin incision from the level of the twelfth thoracic vertebra to approximately 10 mm above the level of the sacrum. Ten millimetre vertical incisions were then made bilaterally in the epimysium of the paravertebral muscles. A pocket approximately 12 mm in length was made laterally by bluntly dissecting between muscle fibres within the muscle belly, and the implants were inserted. The external epimysium was then closed with a single interrupted suture (4/0, RB-1 needle, Ethicon Vicryl Rapide). The skin incision was then closed with 5–6 interrupted sutures as required (3/0, FS-1 needle, Ethicon Vicryl, Amtech Medical, New Zealand). The animals were housed individually and monitored for food intake and hydration for 10 days post surgery, before rehoming in group cages. The animals were then monitored primarily for indications of hydrogen evolution and pain behaviour every 48 h until the conclusion of the investigation. At either 4 or 8 weeks post-implantation, the animals were euthanized by carbon dioxide inhalation. The implant and surrounding tissue were then excised and fixed in 10% neutral buffered formalin (10% NBF) for 48 h for histological analysis. After fixation the discs were carefully removed from the surrounding tissue, immersed in a chromic acid solution (200 g L−1 CrO3 and 10 g L−1 AgNO3) for 15 min to remove corrosion products, and weighed to assess corrosion. Tissue samples were then processed using a standard wax embedding protocol, and 4 μM sections were cut 2.5 mm, 5 mm and 7.5 mm from the medial edge of the disc space with a rotary microtome. The sections were then stained with haematoxylin and eosin (H&E) and visualized with light microscopy. For analysis three randomly identified highmagnification fields were examined (40 × objective lens, each field 1200 × 900 μm). A representative field of view is presented in figure 1. In each field the width of the fibrous capsule was assessed by averaging the depth of the fibrous and inflammatory tissue layer adjacent to the implant at three randomly allocated points. A semi-quantitative scale was used to assess the infiltration of lymphoctyes, macrophages and neovascularization based on the scoring system defined by ISO 10993–6 (2007) (table 1). Other features of tissue
Figure 1. A representative field of view used for the assessment of in vivo biocompatibility. The white space towards the top of the image is the area where the implant was present prior to removal. The black arrow indicates the fibrous capsule that has formed adjacent to the implant. This region is lighter in colour as it is primarily composed of collagen as opposed to the darker skeletal muscle.
reaction that were not measured within this study recognized in ISO 10993–6 such as polymorphonuclear cells, plasma cells, and giant cells were identified extremely rarely, and no tissue necrosis was identified in any of the tissue sections examined. The production of hydrogen gas was also assessed at low magnification with a semi-quantitative scale based on the presence of bubble shaped spaces within the tissue, and distension of the implant space for each entire tissue section (table 1). A one-way ANOVA with Dunnett’s post hoc analysis was used to compare the results of all Mg based materials to those of Ti6Al4V. 3. Results 3.1. Assessing the in vitro biocompatibility of magnesium alloys
The viability results for L929 and SaOS-2 cells cultured on the four Mg alloys, pure Mg and Ti–6Al–4V at 24, 48 and 96 h are presented in figures 2 and 3, respectively. The viability results from each time point must be assessed alongside the total cell numbers present to attain a comprehensive view of the cellular response to the materials investigated. This prevents 3
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R Figure 3. Results from the Live/Dead assay 24, 48, and 96 h after the seeding of SaOS-2 cells onto Ti–6Al–4V, pure Mg or Mg alloy discs. ∗ Indicates a statistically significant difference when the Mg based materials were compared to Ti–6Al–4V (p < 0.01) using a 1-way ANOVA with Dunnett’s post hoc analysis.
R Figure 2. Results from the Live/Dead assay 24, 48, and 96 h after
the seeding of L929 cells onto Ti–6Al–4V, pure Mg or Mg alloy discs. ∗ Indicates a statistically significant difference when the Mg based materials were compared to Ti–6Al–4V (p < 0.01) using a 1-way ANOVA with Dunnett’s post hoc analysis.
A similar trend can be identified after 48 h, with statistically significant difference in viability between Ti– 6Al–4V and Mg–0.4Ca, Mg–0.8Ca and Mg–1Zn. Total cell numbers are similar with a significant difference identifiable between Ti6Al4V and Pure Mg, Mg–0.4Ca, Mg–0.8Ca and Mg–0.5Mn. At this stage Mg–1Zn alone exhibited a total cell number that was not significantly different from the titanium alloy. After 96 h, the total cell numbers present on the titanium alloy had reached 1300 with a viability 99.5%. However, Pure Mg is the only material providing a viability significantly lower than Ti–6Al–4V at this time point. In contrast, all the Mg based materials exhibited a statistically significant reduction in total cell number when compared to Ti–6Al–4V. The results of the SaOS-2 assay indicate similar results. At the 24 and 48 h time points, all the magnesium based materials exhibited significantly reduced viability and total cell number when compared to Ti–6Al–4V, with the viability of Pure Mg the only exception. After 96 h, the same trend is followed with the only difference being the lack of a significant reduction
the misrepresentation of a material as cytocompatible, if for example 80% of the cells are viable, but only five cells are present (the low numbers of cells would therefore represent significant toxicity). The results for the L929 cells at 24 h show a statistically significant difference in the viability of cells between Ti– 6Al–4V and Mg–0.4Ca, Mg–0.8Ca and Mg–1Zn (p < 0.01). Whereas at this time point there was no significant difference between Ti–6Al–4V and Pure Mg or Mg–0.5Mn (p > 0.01). However, the total cell numbers indicate a statistically significant difference between Ti–6Al–4V and all of the Mg based materials (p < 0.01). This reduction in total cell number for all Mg based materials at this early stage is indicative of difficulties in the initial adherence of cells to the Mg based substrates. This reduced initial attachment has important ramifications on the total cell numbers present at later time points. Figure 4 displays representative confocal images of live and dead L929 cells after 96 h cultured on Mg based materials or Ti–6Al–4V. 4
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(A)
(B)
(A)
(C)
(D)
(C )
(E)
(F)
(E )
(B)
(D)
(F )
Figure 4. Confocal micrographs of L929 cells cultured on the surface of Mg alloy, pure Mg and Ti–6Al–4V after 96 h. Green profiles indicate living cells (calcein AM). Red profiles indicate dying or dead cells (ethidium homodimer-1). (A) Mg–0.4Ca. (B) Mg–0.8Ca. (C) Mg–0.5Mn. (D) Mg–1Zn. (E). Pure Mg. (F) Ti–6Al–4V.
Figure 5. Confocal micrographs of SaOS-2 cells cultured on the surface of Mg alloy, pure Mg and Ti–6Al–4V after 96 h. Green profiles indicate living cells (calcein AM). Red profiles indicate dying or dead cells (ethidium homodimer-1). (A) Mg–0.4Ca. (B) Mg–0.8Ca. (C) Mg–0.5Mn. (D) Mg–1Zn. (E). Pure Mg. (F) Ti–6Al–4V.
between the total cell numbers on Ti–6Al–4V when compared to Mg–1Zn. Figure 5 displays representative confocal images of live and dead SaOS-2 cells after 96 h cultured on Mg based materials or Ti–6Al–4V.
are presented in figure 7. In concurrence with the results from the fibrous encapsulation, and lymphocyte and macrophage infiltration, there were no statistically significant differences between the average grade allocated for neovascularization when Ti–6Al–4V was compared to the Mg based materials (p > 0.01). In contrast, there was a statistically significant difference between Ti–6Al–4V and all of the Mg based materials for hydrogen production and corrosion (p < 0.01). However, this result was expected as the titanium alloy neither corrodes nor produces hydrogen gas. Figure 8 shows low magnification micrographs indicating tissue distension in response to hydrogen production with the implantation of Ti–6Al–4V and Mg–0.4Ca.
3.2. The biocompatibility of magnesium alloys in an intramuscular environment
The average width of the fibrous capsule, and the average grade allocated for lymphocyte and macrophage infiltration in association with the intramuscular implantation of Ti–6Al–4V, Pure Mg, Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, and Mg–1Zn at 4 and 8 weeks are presented in figure 6. Statistical analysis indicated no significant differences between Ti–6Al–4V and the Mg based materials for any variable at either time point (p > 0.01). The average grade allocated for neovascularization, hydrogen production and corrosion in association with the intramuscular implantation of Ti–6Al–4V, Pure Mg, Mg– 0.4Ca, Mg–0.8Ca, Mg–0.5Mn, and Mg–1Zn at 4 and 8 weeks
4. Discussion 4.1. General comments on the In Vitro methodology
A multitude of in vitro cell culture techniques exist for the assessment of the biocompatibility of magnesium as 5
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Figure 6. The average width of the fibrous capsule, and average grades allocated to lymphocyte infiltration and macrophage infiltration in the tissue adjacent to Ti–6Al–4V, pure Mg or Mg alloy implants. There were no statistically significant differences for any measure when the Mg based materials were compared to Ti–6Al–4V (p > 0.01) using a 1-way ANOVA with Dunnett’s post hoc analysis.
a biomaterial. Primarily, these techniques are based on the methods outlined by international standards with slight variations in experimental set-up and the assessment of cellular
behaviour. In particular, ISO-10993–5 suggests the use of either an indirect method in which cells are exposed to extracts of the materials, or the direct method in which the 6
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Figure 7. The average grade allocated to neovascularization, hydrogen production, and implant corrosion with implantation of Ti–6Al–4V, pure Mg or Mg alloy implants. There were no statistically significant differences between the Mg based materials and Ti–6Al–4V for neovascularization (p > 0.01, 1-way ANOVA with Dunnett’s post hoc analysis). All Mg based materials exhibited statistically significant increases in both hydrogen production and corrosion when compared to Ti–6Al–4V (p < 0.01, 1-way ANOVA with Dunnett’s post hoc analysis).
cells come into contact with the materials [3]. Unfortunately, the standards currently available are designed for the assessment of the in vitro biocompatibility of primarily non-
degrading biomaterials. This leads to difficulties in applying these methods to rapidly corroding materials such as Mg alloys. 7
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factors including both the pH and osmolality of the cell culture environment, reducing the value of the results when used with Mg based materials [20, 24]. For the above reasons, a modified direct method using R the Live/Dead assay was adopted for this investigation. The choice to seed cells directly onto the surfaces of the materials was based upon a desire to assess whether cells could attach and stay viable on an actively corroding material. This was preferable to the protocol outlined in ISO 10993–5 for the direct method in which the test material is placed onto an adherent cell layer, as it was believed that for Mg this was too similar to the indirect method in which the pH change and increase in osmolality would have the greatest influence on cell survival. Additionally, no evidence existed that the corroding Mg materials would interfere with the calcein AM or ethidium homodimer utilized for the assay. Several other aspects of the methodology used within this investigation require comment. Firstly, the materials were pre-soaked to allow the formation of a passivation layer on the surface of the implant. Whilst it could be argued that this step is masking the potentially toxic effects of the rapid initial corrosion of the materials, initial experiments indicated that without this allowance initial cellular adhesion would be greatly diminished. Eliminating this step would drastically reduce any ability to investigate the in vitro biocompatibility of Mg alloys, and it was therefore included in the protocol. Secondly, it is important to note that the media for each material was replaced daily. This step was taken to reduce any toxicity occurring due to an increase in the pH of the static cell culture environment. Again, this could be argued to be masking the toxic effect of the pH change associated with Mg corrosion. However, it is believed that in a dynamic in vivo environment, the pH in the area associated with a Mg alloy implant would not fluctuate as significantly as in an in vitro environment. However, there would still be a marked increase in pH at the surface of the material in vitro that would also occur in an in vivo environment. Therefore, the media replacement regime utilized should not have further increased the innate discrepancies between in vitro and in vivo methodologies. Thirdly, the choice of cells used in the investigation of in vitro biocompatibility is important. In this study two immortalized cell lines were used, L929 (mouse fibroblast) and SaOS-2 (human osteosarcoma). The L929 line was selected as a standardized, and commonly employed cell type for biocompatibility studies [25–28]. The SaOS-2 cells were also included as a cell line derived from the more clinically applicable human osteosarcoma [29]. These cells are also commonly used for the assessment of the biocompatibility of potential orthopaedic materials due to their osteoblastic phenotype, ability to differentiate and produce mineralized extracellular matrix [30–33]. Within this experiment the SaOS2 cells were not exposed to a differentiating media, which in retrospect could have improved adherence, and therefore increased cell number. Although this could potentially have altered the results gathered, the use of both cell lines is still of vital importance as the cellular response to any given material has been observed to differ vastly depending on the cell type [8, 17, 29, 34, 35].
(A)
(B)
Figure 8. Low magnification micrographs of the implant spaces left after the removal of an intramuscular implant (H & E). (A) The white space is the area left after removal of a Ti–6Al–4V implant with clearly identifiable margins within the tissue. This received a grade of 0 on the previously identified scoring system for hydrogen evolution. (B) The space left after removal of a Mg–0.4Ca implant with some distension of the space, and obscured margins due to probable hydrogen bubbles within the tissue. This received a grade 3 on the previously identified scoring system.
The indirect method is used relatively frequently with Mg based materials in one of two ways. Either extracts are produced and diluted to various concentrations [4–16], or a modification to the method is adopted in which metal salts are added to the culture media to replicate the toxicity of the released Mg and alloying component ions [17–19]. The primary issue with the first of these methods is that any toxicity identified is primarily due to the osmolality of the solution [20]. Whilst this can be argued to be of primary importance to the biocompatibility of the material, it is likely that in an in vivo environment constant diffusion would mitigate this effect to a degree. Additionally, when this method is adopted, the extracts are often diluted to a point for which the increased osmolality is no longer toxic [4, 7, 10, 14, 16], somewhat nullifying the use of the method in the first place. The production of an extract solution also requires the immersion of a Mg sample in media for a period of time. The resulting corrosion that occurs would likely cause an increase in the pH of the extract solution [8, 21], which left unadjusted would certainly result in cytotoxicity. However, this point is often left unmentioned in studies using the technique [6, 9, 10, 12–16, 22], or there is reliance on pH adjustment to prevent this occurrence [5]. The aforementioned addition of Mg and alloying component salts dissolved in cell culture media is also problematic due to the effects on the osmolality of the solution. This method can be of use when investigating alloying elements not found within cell culture media such as rare earth elements, but is less useful for assessing the effect of ions already found within physiological solutions such as Mg and Ca. Additionally, the most common technique used to assess the viability of cells with the indirect method is to use a colorimetric assay based on the reduction of tetrazolium salts to formazan dyes within living cells [7–9, 11, 12, 14– 19, 22, 23]. However, this method can be affected by various 8
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the cytocompatibility of pure Mg show variable results, from extremely toxic [21, 38] to highly biocompatible [36, 37]. This highlights the continuing difficulties with comparing the behaviour of Mg based materials between in vitro investigations in which small differences in variables such as surface roughness, cell type, media type or surface area to volume ratio can dramatically affect corrosion behaviour, and therefore cellular behaviour. Even less evidence exists in the literature for the behaviour of cells on the relevant Mg alloys. One study indicated a significantly reduced total protein concentration, for a Mg– 0.5Ca alloy when compared to a negative control (tissue culture slide). The same study also indicated Mg–0.8Ca provided a result statistically indistinct to the control, perhaps suggesting Mg–0.8Ca was more biocompatible than Mg– 0.5Ca [36]. These results are somewhat in contrast to those seen within this investigation in which the lower calcium percentage-containing alloy generally exhibited slightly improved biocompatibility. However this experiment utilized total protein primarily as an indicator of osteogenic capacity rather than to assess cell number, and was also measured after eight days in culture, both of which make comparison to this investigation difficult. The in vitro biocompatibility of a binary Mg–6Zn alloy has also been investigated, with results showing acceptable adhesion and some cellular division [39, 40]. However these studies do not compare the results to an inert control material preventing any comparison to other investigations. No research has been published investigating the in vitro biocompatibility of a binary Mg–Mn alloy. There are many inherent difficulties in both testing the biocompatibility of Mg alloys in vitro, and attempting to compare the results between studies. If a conservative approach was taken in the assessment of the results seen within this investigation, Mg alloys would be assumed highly toxic materials. However, use of similar alloys and pure Mg in an in vivo environment has indicated good biocompatibility [15, 22, 41]. It is therefore important that a more liberal interpretation of the in vitro results be taken, and to ensure that the conclusions from such studies be made alongside other evidence such as the results from in vivo investigations.
The corrosion associated with Mg alloys, and the resulting by-products of this corrosion make the assessment of any aspect of alloy behaviour difficult in an in vitro environment. The assessment of in vitro biocompatibility is particularly difficult, due to the sensitive nature of cells growing in culture. The points mentioned above highlight these issues and the difficulties in developing methods that avoid cell death due to environmental perturbations, without disguising the potential cytotoxic effects of the Mg alloys. For these reasons the assessment of the biocompatibility of the Mg alloys investigated in this study focused on the viability/cytotoxicity. More in depth investigations of cellular behaviour and differentiation are commonly carried out. However, it is believed that the discrepancies and difficulties in replicating an in vivo environment in vitro, is likely to provide results for these studies only barely comparable to those that would be seen with in vivo implantation of the materials. 4.2. Comparison between the in vitro biocompatibility of Mg alloys
The results of the in vitro analysis of the biocompatibility of Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, pure Mg and Ti– 6Al–4V indicate a general trend towards lower viability and total cell number on the Mg based materials when compared to Ti–6Al–4V. This result was not unexpected due to the active corrosion occurring when the Mg based materials are exposed to a physiological solution. The resultant increase in osmolality, increase in pH and the physical disruption to cells by hydrogen gas evolution, are all factors that would reduce cellular adhesion and viability in a closed cell culture environment [21]. However, despite a relatively inhospitable environment, some cellular adhesion and active cell division did occur when cells were seeded directly onto the Mg alloys being investigated. Even at the reduced viability and total cell numbers observed, this can be seen as an exceptionally positive result for the potential biocompatibility of these alloys in the more dynamic and adjustable in vivo environment. Consequently, despite the significant differences between the results from the Ti–6Al–4V and the Mg based materials, the results of these studies can still be used to an extent for the comparison of the in vitro biocompatibility of the Mg alloys. The results from the L929 experiments generally indicated relatively good performance of the Mg–0.5Mn and Mg–1Zn alloys with increasing time. The cells on these two materials appeared to recover after the initial low total cell number seen at 24 h. In contrast, the Ca containing alloys exhibited very low total cell counts for the entirety of the experiment. Similar results were observed with the SaOS-2 cells for which much higher total cell numbers were seen with Mg–0.5Mn, Mg–1Zn and pure Mg when compared to the Ca containing alloys. The assessment of the in vitro biocompatibility of alloys comparable to those investigated in this study is relatively rare, and more infrequent still is the use of a direct method. Most commonly pure Mg is included as a control material in studies investigating a range of alloys [5, 36], surface treatments [21, 37], coatings [38], or fabrication methods [16]. The studies using a direct method for the assessment of
4.3. General comments on the in vivo methodology
Analysis of the biocompatibility of Mg alloys in a soft tissue environment is relatively uncommon, with the majority of studies progressing from in vitro corrosion and biocompatibility investigations directly to their assessment in an intraosseous location. The reason for this avoidance of standard biocompatibility protocols remains unclear. Both ISO and ASTM standards outline protocols for the short-term assessment of the local effects of implantation of biomaterials in either a subcutaneous or intramuscular location [42–44]. For this investigation, the inclusion of a soft tissue study was deemed important for several reasons. Firstly, it was hypothesized that an intramuscular environment would be able to tolerate the production of hydrogen gas better than a rigid intraosseous environment, a factor considered important given the limited knowledge available regarding the effect 9
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of hydrogen production in an in vivo environment. Secondly, soft tissue implantation provided the ability to accurately assess the corrosion of the materials by mass loss. Thirdly, many orthopaedic biomaterials will directly contact muscle and connective tissue as well as bone. The assessment of the reaction of soft tissues to Mg alloys being developed for orthopaedic application is therefore important.
materials, and the inappropriate exclusion of potentially useful materials due to conservative in vitro corrosion analysis. Additionally it was identified that whilst in vitro techniques can be used as an indicator of the biocompatibility of Mg alloys, the results must be analysed with an understanding that the corrosion of the materials can lead to false positive results for toxicity. It is therefore important that in vitro analysis of biocompatibility be assessed alongside the results of a corresponding in vivo method.
4.4. Comparison between the in vivo biocompatibility of Mg alloys
The in vivo biocompatibility of Mg–0.4Ca, Mg–0.8Ca, Mg–0.5Mn, Mg–1Zn, pure Mg and Ti–6Al–4V in an intramuscular environment was analysed through the assessment of the width of the fibrous capsule, the lymphocyte and macrophage infiltration, and neovascularization in the tissue adjacent to the implants. An overwhelming trend could be identified for all these measures indicating no significant differences between any of the Mg based materials and the Ti–6Al–4V. As this Ti alloy is widely used as a medical implant material in both medicine and dentistry [45–47], these results indicate comparable biocompatibility for these Mg alloys. The assessment of in vivo biocompatibility of Mg alloys in a soft tissue environment is rare, with very few articles available for the comparison of results. The majority of those that have been performed rely on brief descriptive results of the histology to indicate the tissue reaction to the materials [13, 48–50]. One study however, has thoroughly assessed the reaction of muscle tissue to a Mg–0.8Ca implant, using a semi-quantitative scale to assess fibrous encapsulation, tissue cavity formation, necrosis and the infiltration of macrophages, giant cells, heterophilic granulocytes, B-lymphocytes and T-lymphoctyes [51]. This extensive analysis indicated a similar result to that seen within this investigation, with Mg–0.8Ca performing similarly to a stainless steel 316L control material.
References [1] Pearce A I, Richards R G, Milz S, Schneider E and Pearce S G 2007 Animal models for implant biomaterial research in bone: a review Eur. Cells Mater. 13 1–10 [2] Witte F et al 2006 In vitro and in vivo corrosion measurements of magnesium alloys Biomaterials 27 1013–8 [3] ISO 1999 Biological Evalutation of Medical Devices Part 5: Tests for In Vitro Cytotoxicity ISO 10993–5 (Geneva: International Organization for Standardization) [4] Fischer J, Pr¨ofrock D, Hort N, Willumeit R and Feyerabend F 2011 Reprint of: Improved cytotoxicity testing of magnesium materials Mater. Sci. Eng. B 176 1773–7 [5] Yang C, Yuan G, Zhang J, Tang Z, Zhang X and Dai K 2010 Effects of magnesium alloys extracts on adult human bone marrow-derived stromal cell viability and osteogenic differentiation Biomed. Mater. 5 045005 [6] Zheng Y F, Gu X N, Xi Y L and Chai D L 2010 In vitro degradation and cytotoxicity of Mg/Ca composites produced by powder metallurgy Acta Biomater. 6 1783–91 [7] Wang D-W, Cao Y, Qiu H and Bi Z-G 2011 Improved blood compatibility of Mg–1.0Zn–1.0Ca alloy by micro-arc oxidation J. Biomed. Mater. Res. A 99 166–72 [8] Gu X, Zheng Y, Cheng Y, Zhong S and Xi T 2009 In vitro corrosion and biocompatibility of binary magnesium alloys Biomaterials 30 484–98 [9] Huan Z G, Leeflang M a, Zhou J, Fratila-Apachitei L E and Duszczyk J 2010 In vitro degradation behavior and cytocompatibility of Mg–Zn–Zr alloys J. Mater. Sci., Mater. Med. 21 2623–35 [10] Li Y et al 2012 Mg–Zr–Sr alloys as biodegradable implant materials Acta Biomater. 8 3177–88 [11] Park R S et al 2012 Corrosion behavior and cytotoxicity of Mg–35Zn–3Ca alloy for surface modified biodegradable implant material J. Biomed. Mater. Res. B 100 911–23 [12] Wang Y et al 2012 In vitro degradation and biocompatibility of Mg–Nd–Zn–Zr alloy Chin. Sci. Bull. 57 2163–70 [13] H¨anzi A C, Gerber I, Schinhammer M, L¨offler J F and Uggowitzer P J 2010 On the in vitro and in vivo degradation performance and biological response of new biodegradable Mg–Y–Zn alloys Acta Biomater. 6 1824–33 [14] Zhang E, Yin D, Xu L, Yang L and Yang K 2009 Microstructure, mechanical and corrosion properties and biocompatibility of Mg–Zn–Mn alloys for biomedical application Mater. Sci. Eng. C 29 987–93 [15] Li Z, Gu X, Lou S and Zheng Y 2008 The development of binary Mg–Ca alloys for use as biodegradable materials within bone Biomaterials 29 1329–44 [16] Gu X N, Zhou W R, Zheng Y F, Liu Y and Li Y X 2010 Degradation and cytotoxicity of lotus-type porous pure magnesium as potential tissue engineering scaffold material Mater. Lett. 64 1871–4 [17] Feyerabend F et al 2010 Evaluation of short-term effects of rare earth and other elements used in magnesium alloys on primary cells and cell lines Acta Biomater. 6 1834–42
4.5. The corrosion of Mg alloys in an intramuscular environment
One of the benefits in performing a soft tissue implantation of Mg alloys was the ability to remove the samples and assess the corrosion by gravimetric analysis. The results indicated Mg–1Zn as corroding slower than the other Mg based materials at the four-week time point. After eight-weeks of implantation, the corrosion of the alloys followed a similar trend, indicating Mg–1Zn to be the slowest corroding of the alloys investigated. 5. Conclusion The primary aim of the in vitro and in vivo investigation of Mg alloy biocompatibility was to develop and refine a series of simple, rapid and reproducible methods to assess Mg alloy biocompatibility. The provision of an intermediate step between in vitro corrosion analysis and intraosseous implantation of Mg alloys allows the investigation of a larger variety of materials in an in vivo location without the expense and complexity of an intraosseous study. This could prevent both the intraosseous implantation of potentially harmful 10
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[18] Feser K, Kietzmann M, B¨aumer W, Krause C and Bach F W 2011 Effects of degradable Mg–Ca alloys on dendritic cell function J. Biomater. Appl. 25 685–97 [19] Zhang E, Yang L, Xu J and Chen H 2010 Microstructure, mechanical properties and bio-corrosion properties of Mg-Si(-Ca, Zn) alloy for biomedical application Acta Biomater. 6 1756–62 [20] Witte F et al 2008 Degradable biomaterials based on magnesium corrosion Curr. Opin. Solid State Mater. Sci. 12 63–72 [21] Lorenz C, Brunner J G, Kollmannsberger P, Jaafar L, Fabry B and Virtanen S 2009 Effect of surface pre-treatments on biocompatibility of magnesium Acta Biomater. 5 2783–9 [22] Zhang S et al 2010 Research on an Mg–Zn alloy as a degradable biomaterial Acta Biomater. 6 626–40 [23] Witte F et al 2007 Biodegradable magnesiumhydroxyapatite metal matrix composites Biomaterials 28 2163–74 [24] Fischer J, Prosenc MH, Wolff M, Hort N, Willumeit R and Feyerabend F 2010 Interference of magnesium corrosion with tetrazolium-based cytotoxicity assays Acta Biomater. 6 1813–23 [25] Turner T D, Spyratou O and Schmidt R J 1989 Biocompatibility of wound management products: standardization of and determination of cell growth rate in L929 fibroblast cultures J. Pharm. Pharmacol. 41 775–80 [26] Acharya C, Ghosh S K and Kundu S C 2008 Silk fibroin protein from mulberry and non-mulberry silkworms: cytotoxicity, biocompatibility and kinetics of L929 murine fibroblast adhesion J. Mater. Sci., Mater. Med. 19 2827–36 [27] Jorge J H, Giampaolo E T, Vergani C E, Machado A L, Pavarina A C and Carlos I Z 2007 Biocompatibility of denture base acrylic resins evaluated in culture of L929 cells. Effect of polymerisation cycle and postpolymerisation treatments Gerodontology 24 52–7 [28] Serrano M C et al 2004 In vitro biocompatibility assessment of poly(epsilon-caprolactone) films using L929 mouse fibroblasts Biomaterials 25 5603–11 [29] Kirkpatrick C J, Krump-Konvalinkova V, Unger R E, Bittinger F, Otto M and Peters K 2002 Tissue response and biomaterial integration: the efficacy of in vitro methods Biomol. Eng. 19 211–7 [30] Anderson H C et al 1995 The mechanism of bone induction and bone healing by human osteosarcoma cell extracts Clin. Orthop. Relat. Res. 313 129–34 [31] Okumura A et al 2001 Substrate affects the initial attachment and subsequent behavior of human osteoblastic cells (Saos-2) Biomaterials 22 2263–71 [32] Gronowicz G and McCarthy M B 1996 Response of human osteoblasts to implant materials: integrin-mediated adhesion J. Orthop. Res. 14 878–87 [33] Yu X, Botchwey E a, Levine E M, Pollack S R and Laurencin C T 2004 Bioreactor-based bone tissue engineering: the influence of dynamic flow on osteoblast phenotypic expression and matrix mineralization Proc. Natl Acad. Sci. USA 101 11203–8
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