Biocompatibility of magnesium implants in primary human reaming debris-derived cells stem cells in vitro
© The Author(s) 2015
Received: 8 December 2014
Accepted: 15 June 2015
Published: 8 July 2015
Use of magnesium for resorbable metal implants is a new concept in orthopaedic and dental medicine. The majority of studies on magnesium’s biocompatibility in vitro have assessed the short-term effect of magnesium extract on cells. The aim of this study was to evaluate the influence of direct exposure to magnesium alloys on the bioactivity of primary human reaming debris-derived (HRD) cells.
Materials and methods
Pure Mg, Mg2Ag, WE43 and Mg10Gd were tested for biocompatibility. The study consisted of assessment of cell viability by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) test, evaluation of alkaline phosphatase (ALP) content, and study of cell morphology under light microscopy, scanning electron microscopy (SEM) and transmission electron microscopy (TEM), along with determination of calcification and pH changes induced by magnesium.
The number of viable cells in the presence of Mg2Ag was high over the entire observation period. Inhibition of ALP content in osteogenic differentiating HRD was caused by pure Mg at day 14 and 28. All other magnesium alloys did not affect the ALP content. Exposure of HRD to magnesium increased the amount of lysosomes and endocytotic vesicles. Cellular attachment was generally the best for those crystals that formed on the surface of all materials. A decrease was observed in Ca2+ in the medium from day 1 to day 14.
In terms of cell morphology, cell viability and differentiation, cell density and the effect on the surrounding pH, Mg2Ag showed the most promising results. All magnesium materials induced calcification, which is beneficial for orthopaedic and dental applications.
KeywordsBiocompatibility Magnesium Human reaming debris-derived cells
Temporary metal implants that would resorb after bone healing has completed would minimise secondary surgery for implant removal and thus decrease postoperative infections [1–3]. This in turn would decrease high costs related to repeated surgery, reduce recovery periods and thus promote higher quality of life for patients.
A perfect resorbable metal implant should provide enough strength to the healing bone; it should resorb after a set time period, preferably no earlier than 12 weeks , and should be non-toxic and cause no damage to the body. Magnesium is a suitable material for biodegradable metal implants because it is biocompatible  and does not cause toxicological tissue response. Magnesium occurs naturally in humans since our organism contains about 25 g of this element, about 50–60 % of which is found in bone . Magnesium has been shown to stimulate bone formation, since its ions enhance cell attachment and proliferation . In previous studies, new bone was seen forming in direct contact with magnesium [8, 9]. Finally, magnesium’s mechanical properties are closer to those of healthy bone than titanium’s .
However, the problem with magnesium is the formation of hydrogen gas on contact with fluids . In water, magnesium hydroxide accumulates on the surface of the magnesium implant to form a corrosion layer, also known as a degradation layer. While this film slows corrosion under aqueous conditions, it reacts with chlorine ions present in blood to produce highly soluble MgCl2 and hydrogen gas . Hydrogen gas is undesirable as it interferes with normal tissue healing and affects primary implant stability in bone.
At the same time, it has been shown that magnesium facilitates calcification and formation of calcium phosphates . Calcium is needed by the body to ensure that bone laid down by osteoblasts is normally mineralised . It has been shown in previous studies that magnesium increases the pH and that high pH promotes Ca2+ binding [13, 14]. This is especially important for implants that are to be used for bone fractures.
The majority of studies on biocompatibility of magnesium in vitro have assessed the short-term effects of magnesium extract on cells [15–17]. The aim of this study was to evaluate the influence of magnesium alloys on the bioactivity of primary human reaming debris-derived (HRD) cells with up to 21 days of direct exposure. This, in our opinion, will mimic in vivo conditions more closely.
Materials and methods
The following materials were used to produce alloys for this study: magnesium (99.99 %, Xinxiang Jiuli Magnesium Co. Ltd., China), yttrium (99.95 % Y, Grirem Advanced Materials Co. Ltd., China), gadolinium (99.95 % Gd, Grirem Advanced Materials Co. Ltd., China), rare-earth mixture (Grirem Advanced Materials Co. Ltd., China) and silver (99.99 % Ag, ESG Edelmetall-Handel GmbH & Co. KG, Germany).
Three magnesium-based materials were produced: Mg2Ag (1.89 % Ag, the rest was Mg), Mg10Gd (8.4 % Gd, the rest was Mg) and WE43 (3.45 % Y, 2.03 % Nd, 0.84 % Ce, the rest was Mg). Pure magnesium (99.99 % Mg) was used as control. The concentrations of magnesium (Mg), Y, neodymium (Nd) and cerium (Ce) were determined by spark emission spectrometer (Spectrolab M, Spektro, Germany), and the concentrations of Ag and Gd were determined by X-ray fluorescence spectrometer (Bruker AXS S4 Explorer, Bruker AXS GmbH, Germany). The materials were cast at Helmholtz-Zentrum Geesthacht Magnesium Innovation Center (HZG-MagIC).
The three magnesium alloys (Mg2Ag, Mg10Gd and WE43) were produced by permanent mould gravity casting. After melting pure Mg, the melt was held at 720 °C and the preheated alloying elements were added with continuous stirring for 15 min. The melt was poured into a preheated (550 °C) permanent steel mould treated with boron nitride. During the casting process, cover gas was used (SF6 and Ar mixture). The alloys were homogenised with a T4 heat treatment prior to extrusion in Ar atmosphere at 550 °C (Mg10Gd and WE43) or 420 °C (Mg2Ag) for 6 h. Afterwards, the alloys were extruded indirectly with extrusion ratio of 4:25. The chamber of the extrusion machine was set to 370 °C, and the 30-mm-diameter billets were preheated for 1 h at 370 °C (Mg2Ag), 390 °C (WE43) or 430 °C (Mg10Gd). The extrusion speed was between 3 and 4.5 mm/s. Pure Mg was cast by permanent mould direct chill casting.27 The cast billet (d = 110 mm) was extruded indirectly with an extrusion ratio of 1:84. The billet temperature was 340 °C, and the speed of the extrusion was 0.7 mm/s. Discs (10 mm diameter, 1.5 mm thickness) were machined from the extruded bars.
Sample sterilisation and pre-incubation
The samples were sonicated for 20 min in dry isopropanol, dried and gamma-sterilised at the BBF Sterilisation Service GmbH facility (Kernen, Germany) with total dose of 29 kGy. Before seeding the cell culture, the samples were pre-incubated to form a protective degradation layer. Per 0.2 g of sample, 3 mL of F12 K (Gibco®, Life Technologies, USA) was used. The samples were kept at 37 °C in 5 % CO2 atmosphere for 12 h.
Isolation of human reaming debris-derived (HRD) cells
HRD cells were cultured from various patients, with the approval of the local Ethics Commission, as described by Wenisch et al. . The adult patients were of different genders and ages and did not display any disease related to bone metabolism. In total, cells from six different patients were taken for this study.
The reaming debris was cultured in Petri dishes with F12 K medium including 20 % foetal calf serum (FCS), 100 U/ml penicillin and 100 μg/g streptomycin. After 4–7 days, HRD cells started to grow out of the debris. When the cells reached confluence after 2–3 weeks, they were trypsinised and transferred to cell culture flasks. All cells were kept at 37 °C in 5 % CO2 atmosphere.
To determine cell viability, an MTT assay was conducted according to Mosmann et al. . Briefly, 10,000 cells/cm2 were seeded into 12-well plates containing pre-incubated magnesium discs and F12 K medium with 20 % FCS and 100 μg/g streptomycin. Controls with no magnesium discs but only HRDs were used as well. The cell medium was changed every second day during the experiment. Duplicates were used for each material and patient, resulting in testing of 12 wells per specimen. After 24 h, 7 and 21 days, MTT solution was added to cell medium. The cells were then incubated in the dark for 4 h at 37 °C. Subsequently, the cell medium was discarded, and the cells were lysed with 0.004 N HCl in isopropanol. The cell lysates were centrifuged, and supernatants were transferred as triplets to a 96-well plate. Adsorption was measured at 570 and 630 nm using a Synergy HT microplate reader (BioTek, Bad Friedrichshall, Germany). The MTT assay was also performed for magnesium discs not seeded with cell culture to exclude the material’s effect on the test and see only how the cells reacted during the assay.
Additionally, cell morphology was studied by inverted light microscopy using a Leica microscope type 090-135.002 (Leica Microsystems GmbH, Wetzlar, Germany) equipped with a Nikon Ds-Fi1 digital camera (Nikon, Düsseldorf, Germany).
Alkaline phosphatase (ALP) content
As an indicator of changes in the differentiation behaviour of the bone-forming cells caused by the test substances, a SensoLyte® pNPP alkaline phosphatase assay (AnaSpec, Fremont, CA) was applied after 24 h and 7, 14, 21 and 28 days of culturing in Dulbecco’s modified Eagle’s medium (DMEM), low glucose with l-glutamine, 10 % FCS, 100 U/ml penicillin, 100 μg/g streptomycin, 0.1 μM dexamethasone, 0.005 μM ascorbic acid and 10 mM β-glycerol phosphate to induce osteogenic differentiation. The cell medium was changed every second day during the experiment. Duplicates were used for each material and patient, for a total of 12 wells per specimen. Controls with no magnesium discs but only HRDs were used as well.
The cells were washed and frozen at −80 °C. After thawing, the cell number was measured using a PicoGreen® dsDNA quantitation assay (Invitrogen, Eugene, OR) according to the manufacturer’s protocol. Cells were lysed with 1 % Triton X-100 in phosphate-buffered saline. The cell lysates were centrifuged, and the supernatants were mixed with PicoGreen® working solution in a 96-well plate. The samples were excited at 485 nm, and the fluorescence emission intensity was measured at 528 nm. The cells that were lysed for the PicoGreen® assay were centrifuged, and the supernatants were diluted in a specific assay buffer. ALP substrate was applied to the diluted samples, and the absorbance was measured at 405 nm. The absolute amounts of ALP were correlated with the cell numbers obtained from the PicoGreen® assay. Both ALP and PicoGreen® assays were additionally performed for magnesium discs not seeded with cell culture to exclude the material’s effect on the test and see only how the cells reacted during the assays.
Transmission electron microscopy
HRD cells seeded in chamber slides (Nalge Nunc International, Rochester, NY) were incubated with and without magnesium discs for 21 days. The cell layer was fixed for 30 min with 2 % paraformaldehyde in 0.1 M sodium phosphate buffer (pH 7.2–7.4) with 2 % glutaraldehyde and 0.02 % picric acid, followed by 20 min fixation with 1 % osmium tetroxide in 0.1 M sodium cacodylate buffer (pH 7.2–7.4). The samples were dehydrated and embedded in Epon before ultrathin sections (80–100 nm) were applied to collodion-coated copper grids. Analysis was done with a LEO 912 transmission electron microscope (Carl Zeiss AG, Oberkochen, Germany) at 80 kV accelerating voltage, equipped with a TRS SharpEye slow-scan dual-speed charge-coupled device (CCD) camera (Albert Troendle Prototypentwicklung, Moorenweis, Germany).
Scanning electron microscopy
HRD cells were cultivated on magnesium discs for 7 and 21 days. Controls with no magnesium discs but only HRDs were used as well. Subsequently, the cells were fixed in 2 % glutaraldehyde in 0.1 M Na-phosphate buffer (pH 7.4) for 1 h at room temperature, followed by dehydration in a graded series of ethanol and critical-point drying. The specimens were mounted together on aluminium pin stubs with the help of adhesive carbon pads. The specimens were then sputter-coated with gold/palladium (SC7640 sputter coater, VG Microtech, Uckfield, East Sussex, UK) and assessed using a LEO 1530 (LEO Elektronenmikroskopie GmbH, Oberkochen, Germany) field-emission scanning electron microscope operated at 7.5 or 15 kV.
Determination of Ca2+ consumption and pH
At established time points, medium was collected and analysed for Ca2+ in solution and pH. The concentration of Ca2+ was measured using a calcium analyser (9180 electrolyte analyzer, Roche, Mannheim, Germany), and pH measurements were carried out using a pH-meter (Titan X, Fisher Scientific GmbH, Schwerte, Germany) for each time point. The control group for this investigation consisted of well plates that contained only HRDs and medium but no magnesium.
Data were analysed using the Statistical Package for the Social Sciences (SPSS, v18, SPSS Inc., Chicago, USA). The significance level was set at 5 %. The treatment groups were compared with the control group without magnesium by Mann–Whitney U-test. The results are expressed as means. The graphs were plotted with SPSS.
MTT results for HRDs after exposure to different magnesium materials over time
Viability (% of control) ± SD (%), day 1
Viability (% of control) ± SD (%), day 7
Viability (% of control) ± SD (%), day 21
93.4 ± 25.3
13.9 ± 5.0
24.0 ± 19.5
57.1 ± 14.4
30.5 ± 9.2
26.3 ± 3.6
37.0 ± 13.1
18.1 ± 7.1
54.7 ± 7.2
113.4 ± 29.8
63.3 ± 11.0
98.5 ± 12.0
Alkaline phosphatase content
Transmission electron microscopy
Scanning electron microscopy
It was seen that all magnesium materials increased the pH of the medium compared with the control group. The following general pattern was observed for all groups: the pH values were stable up to day 7, a sudden pH drop occurred on day 14, then the pH tended to increase slightly up to day 28. The pH values for Mg2Ag were most similar to the control. Pure Mg caused the greatest increase in pH among all groups, and this increase was statistically significant compared with Mg2Ag (p ≤ 0.003) but not compared with Mg10Gd (p ≥ 0.02) or WE43 (p ≥ 0.02). No correlation between pH and Ca2+ consumption was found in this study.
This study looked at the direct long-term effect of magnesium alloys on primary HRDs. Cell viability, differentiation and morphology as well as pH and calcium uptake were analysed to assess the overall biocompatibility of the tested materials. We evaluated the long-term effects of magnesium on human cells to simulate the in vivo situation as closely as possible.
Mg2Ag had high cell viability from day 1 (113.4 ± 29.8 %) to day 21 (98.5 ± 12.0 %). It was also shown in previous works  that Mg-Ag alloys have negligible cytotoxicity and sound cytocompatibility. Pure Mg had high viability at the very first day (93.4 ± 25.3 %), but then the viability decreased to 24.0 ± 19.5 % at day 21. Mg10Gd and WE43 impaired cell viability in this study. Previous studies have shown higher values for cell viability measured by MTT test compared with the present study [15–17]. The difference between this work and previous publications is that the present study applied the longest in vitro incubation times for magnesium alloys tested up to now. The HRDs were kept in direct contact with the magnesium samples and not in magnesium extract, as done in most studies [15–17].
An important drawback of tetrazolium-based tests is that the difference between cytotoxic (cell death) and cytostatic (reduced growth rate) effects cannot be distinguished . We thus looked at cell morphology under light microscopy, TEM and SEM.
After examination under SEM and light microscopy, it was revealed that the number of cells decreased in the presence of pure Mg, Mg10Gd and WE43. These materials seem to have long-term cytotoxic effects on HRD when placed in direct contact with the cells. This explains the low viability values.
The cell number was high and the cells had normal morphology in Mg2Ag groups. However, the cell viability was lowest at day 7 (63.3 ± 11.0 %) for Mg2Ag. TEM analysis revealed an elevated amount of lysosomes which contained degraded magnesium particles. Degradation particles were also found in the cytoplasm. The presence of high amounts of degradation products inside the HRDs could explain the lower cell viability values for Mg2Ag at day 7. It was shown in previous studies that uptake of material particles leads to induction of cell stress which triggers cytotoxicity .
ALP activity in HRDs is an important factor in bone mineral formation and shows a range of changes during differentiation. Inhibition of ALP activity in osteogenic differentiating HRD was caused by pure Mg at day 14 and 28. All other magnesium alloys did not affect the ALP activity. In this respect, our study shows similar results to previous research in this area , despite the fact that we observed osteogenic differentiation over much longer periods and using direct contact of cells with magnesium.
SEM analysis revealed that the cellular attachment was generally best to crystals generated by degradation products on the material surface. Crystals have been seen forming on magnesium alloys such as Mg-Ag in previous studies . Formation of calcium phosphates [Ca x (PO4) x ] was also observed in previous publications . Interestingly, the crystal distribution was not homogeneous throughout the corrosion layer. In this sense, our results are similar to earlier findings [11, 15].
The fact that the cells attached to the crystalline structures more readily than to the overall material surface and developed numerous pseudopodia can be explained by the rough structure of crystals, and by the chemical composition of these crystals. It was previously shown that cells attach better to certain surfaces with preferable average surface roughness of ~0.5 μm up to ~8.5 μm . Values below or above this range diminish the cells’ ability to bind to the surface.
The chemical composition of the crystals and the degradation layer formed on the magnesium’s surface can also explain the better attachment of the cells to these structures. Their chemical composition consists of calcium, phosphorus, magnesium and oxygen . Thus, the cells attach to already reacted material where they are not mechanically disturbed by hydrogen gas produced as a by-product of degradation. The formation of the degradation layer could also explain the increase in cell density around Mg10Gd and WE43 after 21 days of incubation.
Based on previous findings, the following model for the formation of a corrosion layer has been suggested : (1) initial metal corrosion based on contact with water molecules leads to release of Mg ions, and a thin Mg(OH)2 and MgCO3 layer is formed; (2) The corrosion slows down, and a second layer consisting of amino acids and organic matter is formed; (3) Both layers together shield the sensitive environment around the material and enable cells to grow on the material . Such a complex process of corrosion layer formation could explain the cell viability increase at day 21 for Mg2Ag, Pure Mg and Mg10Gd observed in our experiments.
All magnesium-based materials decreased the amount of Ca2+ in this study. As shown in previous studies, Mg2+ promotes formation of calcium phosphates and consequently decreases the amount of free Ca2+ ions in the medium [11, 14]. In this regard, our results are consistent with earlier works. Sufficient supply of calcium is vital to ensure that bone laid down by osteoblasts is normally mineralised . Calcification is thus advantageous for bone implants that are to be used in the orthopaedic and maxillofacial fields.
It was shown in previous studies that magnesium increases the pH and that high pH promotes Ca2+ binding [13, 14]. In this study, it was also revealed that pH shifts to alkaline values in the presence of magnesium, but to somewhat different degrees for the different alloys. However, no statistical correlation was observed between pH and consumed Ca2+.
In conclusion, our study reveals the long-term effects of magnesium materials on human HRDs seeded directly onto magnesium discs. In respect to cell morphology, cell density and the effect on the surrounding pH, Mg2Ag showed the most promising results. However, the mechanism of cell stress induction and cytotoxicity needs to be further studied to enable prediction of possible health risks.
This project receives funding from the People Programme (Marie Curie Actions) of the European Union’s Seventh Framework Programme FP7 (2007–2013) under REA grant agreement no. 289163. Special thanks are due to Iris Schütz, Justus-Liebig University Giessen, Laboratory for Experimental Trauma Surgery for technical assistance.
Compilance with Ethical Standards
Conflict of interest
The authors declare that they have no conflict of interest related to the publication of this manuscript.
The study was authorized by the local ethical committee and was performed in accordance with the ethical standards of the 1964 Declaration of Helsinki as revised in 2000. All patients gave informed consent prior to being included in the study. The patients gave informed consent for publication of the case study.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.
- Quinn A, Hill AD, Humphreys H (2009) Evolving issues in the prevention of surgical site infections. Surgeon 7(3):170–172View ArticlePubMedGoogle Scholar
- Hachenberg T, Sentürk M, Jannasch O, Lippert H (2010) Postoperative Wundinfektionen. Pathophysiologie, Risikofaktoren, und Praventive Konzepte. [Postoperative wound infections. Pathophysiology, risk factors and preventive concepts]. Anaesthesist 9(59):851–866 (in German) View ArticleGoogle Scholar
- Jannasch O, Lippert H (2011) Perioperative Prophylaxe und Therapie von Infektionen-Postoperative Wundinfektionen. [Surgical site infections]. Anasthesiol Intensivmed Notfallmed Schmerzther 46(10):664–673 (in German) View ArticlePubMedGoogle Scholar
- Staiger MP, Pietak AM, Huadmai J, Dias G (2006) Magnesium and its alloys as orthopedic biomaterials: a review. Biomaterials 27:1728–1734View ArticlePubMedGoogle Scholar
- Sandler G, Nguyen L, Lam L, Manglick MP, Soundappan SS, Holland AJ (2011) Trampoline trauma in children: is it preventable? Pediatr Emerg Care 27(11):1052–1056View ArticlePubMedGoogle Scholar
- Goyer RA, Clarkson TW (2001) Toxicity of metals. In: Klaassen CD (ed) Casarett and Doull’s Toxicology: The Basic Science of Poisons. McGraw-Hill, USA, pp 811–868Google Scholar
- Institute of Medicine Standing Committee on the Scientific Evaluation of Dietary Reference Intakes (1997) Dietary Reference Intakes for Calcium, Phosphorus, Magnesium, Vitamin D, and Fluoride. National Academy Press, Washington DCGoogle Scholar
- Li Z, Gu X, Lou S, Zheng Y (2008) The development of binary Mg–Ca alloys for use as biodegradable materials within bone. Biomaterials 29(10):1329–1344View ArticlePubMedGoogle Scholar
- Xu L, Yu G, Zhang E, Pan F, Yang K (2007) In vivo corrosion behavior of Mg–Mn–Zn alloy for bone implant application. J Biomed Mater Res A 83(3):703–711View ArticlePubMedGoogle Scholar
- Witte F, Kaese V, Haferkamp H, Switzer E, Meyer-Lindenberg A, Wirth CJ, Windhagen H (2005) In vivo corrosion of four magnesium alloys and the associated bone response. Biomaterials 26(17):3557–3563View ArticlePubMedGoogle Scholar
- Feyerabend F, Drücker H, Laipple D, Vogt C, Stekker M, Hort N, Willumeit R (2012) Ion release from magnesium materials in physiological solutions under different oxygen tensions. J Mater Sci Mater Med 23(1):9–24View ArticlePubMedGoogle Scholar
- Reid IR (2014) Should we prescribe calcium supplements for osteoporosis prevention? J Bone Metab 21(1):21–28View ArticlePubMedPubMed CentralGoogle Scholar
- Witte F, Hort N, Vogt C, Cohen S, Kainer KU, Willumeit R, Feyerabend F (2008) Degradable biomaterials based on magnesium corrosion. Curr Opin Solid State Mater Sci 12:63–72View ArticleGoogle Scholar
- Willumeit R, Fischer J, Feyerabend F, Hort N, Bismayer U, Heidrich S, Mihailova B (2011) Chemical surface alteration of biodegradable magnesium exposed to corrosion media. Acta Biomater 7(6):2704–2715View ArticlePubMedGoogle Scholar
- Tie D, Feyerabend F, Müller WD, Schade R, Liefeith K, Kainer KU, Willumeit R (2013) Antibacterial biodegradable Mg-Ag alloys. Eur Cell Mater 16(25):284–298Google Scholar
- Feyerabend F, Fischer J, Holtz J, Witte F, Willumeit R, Drücker H, Vogt C, Hort N (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(5):1834–1842View ArticlePubMedGoogle Scholar
- Yang L, Hort N, Laipple D, Höche D, Huang Y, Kainer KU, Willumeit R, Feyerabend F (2013) Element distribution in the corrosion layer and cytotoxicity of alloy Mg-10Dy during in vitro biodegradation. Acta Biomater 9(10):8475–8487View ArticlePubMedGoogle Scholar
- Wenisch S, Trinkaus K, Hild A, Hose D, Herde K, Heiss C, Kilian O, Alt V, Schnettler R (2005) Human reaming debris: a source of multipotent stem cells. Bone 36(1):74–83View ArticlePubMedGoogle Scholar
- Mosmann T (1983) Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J Immunol Methods 65(1–2):55–63View ArticlePubMedGoogle Scholar
- Plumb JA (2004) Cell sensitivity assays: the MTT assay. Methods Mol Med 88:165–169PubMedGoogle Scholar
- Pauksch L, Hartmann S, Rohnke M, Szalay G, Alt V, Schnettler R, Lips KS (2014) Biocompatibility of silver nanoparticles and silver ions in primary human mesenchymal stem cells and osteoblasts. Acta Biomater 10(1):439–449View ArticlePubMedGoogle Scholar
- Li RW, Kirkland NT, Truong J, Wang J, Smith PN, Birbilis N, Nisbet (2014) The influence of biodegradable magnesium alloys on the osteogenic differentiation of human mesenchymal stem cells. J Biomed Mater Res A 14(12):4346–4357. doi:https://doi.org/10.1002/jbm.a.35111 (Epub ahead of print) Google Scholar
- Stout KJ, Sullivan PJ, Dong WP, Mainsah E, Luo N, Mathia T, Zahouani H. Development of Methods for Characterisation of Roughness in Three Dimensions. European Community Contract No 3374/1/0/170/90/2 Birmingham: Univ of Birmingham, 1993Google Scholar