Biomimetic Apatite Mineralization Mechanisms of Mesoporous

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Biomimetic Apatite Mineralization Mechanisms of Mesoporous Bioactive Glasses as Probed by Multinuclear 31P, 29Si, 23Na and 13C Solid-State NMR Philips N. Gunawidjaja,† Andy Y. H. Lo,† Isabel Izquierdo-Barba,‡,§ Ana Garcıá,‡,§ Daniel Arcos,‡,§ Baltzar Stevensson,† Jekabs Grins,| Marıá Vallet-Regı´,‡,§ and Mattias Edeń*,† Physical Chemistry DiVision, Department of Materials and EnVironmental Chemistry, Arrhenius Laboratory, Stockholm UniVersity, SE-106 91, Stockholm, Sweden, Departamento de Quı´mica Inorganica´ y Bioinorgańica, Facultad de Farmacia, UniVersidad Complutense de Madrid, 28040 Madrid, Spain, Networking Research Center on Bioengineering, Biomaterials and Nanomedicine, CIBER-BBN, Madrid, Spain, and Inorganic and Structural Chemistry DiVision, Department of Materials and EnVironmental Chemistry, Arrhenius Laboratory, Stockholm UniVersity, SE-106 91, Stockholm, Sweden ReceiVed: June 12, 2010; ReVised Manuscript ReceiVed: September 9, 2010

An array of magic-angle spinning (MAS) nuclear magnetic resonance (NMR) spectroscopy experiments is applied to explore the surface reactions of a mesoporous bioactive glass (MBG) of composition Ca0.10Si0.85P0.04O1.90 when subjected to a simulated body fluid (SBF) for variable intervals. Powder X-ray diffraction and 31P NMR techniques are employed to quantitatively monitor the formation of an initially amorphous calcium phosphate surface layer and its subsequent crystallization into hydroxycarbonate apatite (HCA). Prior to the onset of HCA formation, 1H f 29Si cross-polarization (CP) NMR evidence dissolution of calcium ions; a slightly increased connectivity of the speciation of silicate ions is observed at the MBG surface over 1 week of SBF exposure. The incorporation of carbonate and sodium ions into the bioactive orthophosphate surface layer is explored by 1H f 13C CPMAS and 23Na NMR, respectively. We discuss similarities and distinctions in composition-bioactivity relationships established for traditional melt-prepared bioglasses compared to MBGs. The high bioactivity of phosphorus-bearing MBGs is rationalized to stem from an acceleration of their surface reactions due to presence of amorphous calcium orthophosphate clusters of the MBG pore wall. 1. Introduction Intense efforts have for decades been devoted to finding biocompatible nontoxic materials that have the ability to bond to soft (e.g., muscles) as well as to hard (e.g., bone and tooth) tissues.1–6 One option is to employ silicate-based bioactiVe glasses (BGs), whose “bioactiVity” stems from the biomimetic formation of an amorphous calcium phosphate (ACP) layer on the glass surface upon its exposure to body fluids.1–6 The layer subsequently crystallizes into calcium-deficient nanocrystalline hydroxycarbonate apatite (HCA), which has a similar composition as the inorganic constituents of bone and tooth. The higher the bioactivity (“tissue-bonding ability”) of the BG, the faster its formation rate of HCA. The bioactivity is often probed in vitro by subjecting the glass to a simulated body fluid (SBF) that mimics acellular human plasma.7 Hench and co-workers introduced the first melt-prepared “45S5 Bioglass” composition, 24Na2O-27CaO-46SiO23P2O5, and demonstrated its apatite-forming ability.1–3 Significant progress has since been made for developing improved BGs and assessing their bioactive properties. In particular, composition-bioactivity correlations have been explored experimentally in the Na2O-CaO-SiO2-P2O5 glass system,8–12 as well as in the limiting Na2O-SiO2-(P2O5)13,14 and CaO* Corresponding author. E-mail: [email protected]. Fax: +46 8 152187. Phone: +46 8 162375. † Physical Chemistry Division, Stockholm University. ‡ Universidad Complutense de Madrid. § Networking Research Center on Bioengineering, Biomaterials and Nanomedicine, CIBER-BBN. | Inorganic and Structural Chemistry Division, Stockholm University.

SiO2-(P2O5)11,15–22 systems. The sol-gel technique is usually exploited for preparing SiO2-richer compositions than those attainable by standard melt-quench procedures.15–21 The bioactivity of melt-prepared BGs (MPBGs) generally increases with their Na and/or Ca contents,3–5 while the presence of a few mol % of P is recognized to markedly enhance the apatite formation rate,3,4,10,17,23 although for not fully understood reasons. Computer modeling constitutes a valuable tool for predicting structures of BGs and their bearing on the bioactivity.24–30 The group of Hench has proposed a five-step mechanism, which accounts for the main surface reactions that lead to HCA formation from MPBGs both in vivo and in vitro.2,3,31 It is outlined in Table 1 and will herein be referred to as the Hench mechanism (HM). Its first step involves a depletion of glass modifier ions through exchange with H+, followed by defragmentation and repolymerization processes of the silicate network. This results in a continuously thickened silica gel surface layer, as reported in several studies of BGs subjected to (simulated) body fluids.8,9,14,15,32 The HM next proceeds by ACP formation (stage 4) and its crystallization into HCA (step 5). A multitude of investigations have explored/discussed various aspects of these surface layers, using primarily X-ray diffraction (XRD), Fourier transform infrared (FTIR) spectroscopy, and electron microscopy,8–11,14–20,23,33–35 and to a much lesser extent solid-state NMR.12,13,21,22,36 Despite being introduced for Na2O-CaO-SiO2-P2O5 MPBGs, the HM is formulated quite generally and is readily adaptable to essentially any silicate-based biomaterial. To our knowledge, the HM is overall corroborated by existing studies. Yet, several details of the bioactive processes are debated, such

10.1021/jp105408c  2010 American Chemical Society Published on Web 10/21/2010

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TABLE 1: HCA Formation Mechanism According to Hencha MBGb stage 1 2 3 4c 5

interpretation +

+

Cation dissolution (M T H exchange) leading to the formation of SiOH groups Silica network defragmentation (“hydrolysis”); formation of SiOH groups but may involve partial loss of Si as Si(OH)4(aq) Network repolymerization; consumption of SiOH groups Formation of ACP

reaction

path A

path B

V

(V)

(-)

Si-O-Si + H2O f 2Si-OH

V

(V)

(-)

2Si-OH f Si-O-Si + H2O

V

(V)

(-)

Attraction of Ca2+ and PO34 either from solution or by migration from glass network

V

V

(V)

V

V

V

-

+

MPBG

b

+

Si-O M + H2O f Si-OH + M (aq) + OH

ACP f HCA crystallization; uptake of CO23 , OH , and other cations from the solution

-

Based on ref 3 and assuming for simplicity an M-Si-(P)-O BG system with monovalent M+ ions. The reactions are formulated assuming a (physiological) pH > 7 for steps 1-3 and also to convey the typically observed increased pH of the solution. These steps are often formulated using H+ or H3O+ ions. b These columns contrast the relevance of each HM step for the case of melt-prepared BGs (MPBGs) or for MBGs. An arrow pointing down, V, signifies that the reactions must proceed through this step, whereas (V) indicates that this stage is necessary but proceeds faster and (-) marks that the step is redundant. Note that phosphorus-bearing MBGs may follow two essentially independent pathways (A and B), as discussed in section 4. c The detailed mechanism of this step remains unsettled; e.g., see refs 13, 27, 28, and 37 for concrete suggestions. a

as the precise nature of the surface sites that most efficiently promote the onset of ACP formation, as well as the detailed elementary reactions involved; here, dissolved SiO44- ions,13,32 silanols, or their negatively charged deprotonated counterparts,13,32,37 as well as two- or three-membered rings of SiO4 tetrahedra,26–29,31 have been proposed as candidates. Also, the precise conversion pathway between ACP and the final HCA product remains unsettled.3 The recently introduced “mesoporous bioactive glasses” (MBGs)5,38–46 are prepared by using surfactants as structuredirecting agents through an evaporation-induced self-assembly (EISA) process.47 Their large surface area, stemming from an ordered arrangement of uniformly sized mesopores, is believed to contribute significantly to their superior bioactivity compared to both gel-prepared and (especially) melt-deriving BGs. Some investigations on composition-bioactivity relations are reported for MBGs.38,39,44,45 With the use of 29Si, 31P, and 1H magicangle spinning (MAS) NMR spectroscopy, we recently provided the first model of the pore-wall constitution of phosphorusbearing MBGs.48 It was found to comprise two primary components: a main silica-rich CaO-SiO2 glass and a minor amorphous calcium orthophosphate component; the latter is henceforth referred to as CaP and constitutes (disordered) Ca3(PO4)2 clusters, incorporating water molecules and OH- ions. As proposed in ref 48, and verified further in ref 41, the CaP clusters provide natural nucleation environments for a further growth into ACP/HCA phases. They should contribute significantly, or even predominantly, to the superior bioactivity observed in vitro from MBGs (and likely also phosphoruscontaining gel-prepared BGs) compared to their MPBGs counterparts. Additional support for the dual-phase pore-wall model, as well as the role of CaP clusters for accelerating the HCA formation of SBF-soaked MBGs, was provided by Jagadeesan et al.,46 who demonstrated that the bioactivity of MBGs may be enhanced further by incorporation of crystalline calcium phosphate into the silica-based pore walls. In this work, we employ complementary 31P, 29Si, 23Na, and 13 C MAS NMR experiments to quantitatively probe both the surface reactions at the silicate portion of the pore wall (by 29Si NMR) and the formation of amorphous and crystalline surfacelayer components (using 31P) for progressively increased SBF-

exposure intervals of an “S85” MBG of composition Ca0.10Si0.85P0.04O1.90. Further, we provide the first reports on the local 23Na and 13C environments of Na+ and CO32- ions incorporated into the phosphate surface layer formed on a bioglass in vitro. We also discuss our results in relation to the HM and identify two alternative mechanistic routes that phosphorus-bearing MBGs may follow when subjected to body fluids to finally form HCA; this feature is identified as the primary reason for their enhanced bioactivity compared to MPBGs. 2. Materials and Methods 2.1. Preparation and Characterization of Samples. An MBG sample of nominal composition 10CaO-85SiO2-5P2O5 was prepared at 40 °C using an EISA process47 with the P123 triblock copolymer as structure-directing agent. Si, P, and Ca were introduced by the precursors tetraethyl orthosilicate (TEOS), triethyl phosphate (TEP), and Ca(NO3)2 · 4H2O, respectively. The resulting homogeneous colorless membranes were calcined at 700 °C for 6 h to remove organic species and nitrate ions; this final product is henceforth denoted “S85” to specify its atom % of Si out of the cations. Reference 44 provides further details on the synthesis. The in vitro bioactivity was assessed on powders of S85 in an SBF solution according to Kokubo et al.,7 prepared by dissolving NaCl, KCl, NaHCO3, K2HPO4 · 3H2O, MgCl2 · 6H2O, CaCl2, and Na2SO4 into distilled water. The solution was buffered at pH ) 7.4 using tris(hydroxymethyl)aminomethane/ HCl and was filtered (0.22 µm Millipore filters) to avoid bacterial contamination. The SBF treatment was carried out at 37 °C, using a sealed polyethylene container under continuous orbital stirring (600 rpm) in an Ecotron HT incubator. This procedure was repeated for each of six samples for increasing time intervals between 30 min and 1 week. Each sample was filtered and gently washed with water to quench the surface reactions and remove potentially precipitated salts. It was subsequently vacuum-dried at 37 °C for several days. Each member of the resulting set of samples is labeled “sbfN”, with the index N denoting the exposure interval in hours (h) or days (d).

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TABLE 2: Analyzed Sample Compositionsa

1

sample

xSi

xCa

xP

stoichiometric formulab

Na (wt %)c

S85 sbf0.5h sbf1h sbf4h sbf1d sbf3d sbf7d

0.853 0.870 0.871 0.878 0.869 0.873 0.868

0.104 0.0855 0.0841 0.0768 0.0775 0.0697 0.0755

0.0431 0.0446 0.0452 0.0454 0.0530 0.0564 0.0563

Ca0.12SiP0.051O2.25 Ca0.098SiP0.051O2.23 Ca0.097SiP0.052O2.23 Ca0.088SiP0.052O2.22 Ca0.089SiP0.061O2.24 Ca0.080SiP0.066O2.24 Ca0.087SiP0.065O2.25

0.00 0.30 0.28 0.28 0.35 0.40 0.47

a XRF-analyzed cation compositions, where xE is the atomic fraction of element E out of the cations. b Charge-balanced stoichiometric formula, normalized to a Si coefficient of unity and ignoring unknown, but relatively small, amounts of surfaceassociated protons. c Analyzed Na content of the sample. All sbfN samples also comprise minute amounts of other cations and anions, such as K+, Mg2+, and Cl-.

Chemical compositions were determined by X-ray fluorescence (XRF) spectroscopy, using a Philips PANalytical AXIOS spectrometer (Philips Electronics NV), with X-rays generated by the Rh KR line at λ ) 0.614 Å. The XRF-analyzed compositions are listed in Table 2. X-ray powder diffraction patterns were recorded with Cu KR radiation using a PANalytical X’Pert PRO MPD diffractometer equipped with an X’Celerator detector. Powdered samples were dispersed on zerobackground Si plates and XRD patterns were recorded using variable slits and 4 cm2 irradiated area in the 2θ range of 10-80° for a total time of 4-12 h. 2.2. Solid-State NMR. All NMR spectra were acquired at a magnetic field of 9.4 T using a Varian/Chemagnetics Infinity400 spectrometer, giving the following Larmor frequencies: -161.98 MHz for 31P, -105.84 MHz for 23Na, -100.61 MHz for 13C, and 79.49 MHz for 29Si. Finely ground powders were filled in 6 mm zirconia rotors and spun at 9.0 kHz for 31P, 8.0 kHz for 29Si, 8.5 kHz for 23Na, and 5.25 kHz for 13C NMR experimentation, respectively, using a double-resonance 6 mm Chemagnetics apex probehead. Single-pulse experiments used rf flip angles, nutation freX /2π (X ) 23Na, 29Si, 31P), and relaxation delays as quencies ωnut 29 follows: for Si (70°; 37 kHz; 720-1080 s); 31P (90°; 48 kHz; 600 s); 23Na (90°; 93 kHz central-transition nutation frequency; 1 s). Relaxation delays were chosen based on separate T1 saturation-recovery measurements on a selected set of samples, so as to provide quantitative NMR spectra for each nucleus. Each 29Si and 31P acquisition was divided in blocks of around 20 transients, using a longer equilibration period of at least twice the relaxation delay prior to starting each block. The total numbers of accumulated signal transients were typically 10 800, 260, and 80-256 for 23Na, 29Si, and 31P experiments, respectively. Ramped49 1H f X (X ) 31P, 29Si, 13C) cross-polarization (CP) was established at the modified Hartmann-Hahn condition H X ωnut - ωnut ) ωr, where ωr is the spinning frequency. Nutation frequencies of ≈20 kHz for 29Si and ≈32 kHz for 13C and 31P were employed. All CPMAS experiments used relaxation delays of 5 s, with 6000-11280 and 5040-8640 coadded signal transients for 29Si and 31P, respectively, whereas those for 13C are listed in the caption of Figure 9. The 1H f 31P CP-based 2D HETCOR data of Figure 7 was recorded by employing a contact interval of 1.5 ms and spectral windows of 27.0 and 9.0 kHz for the direct (31P, horizontal) and indirect (1H, vertical) dimensions, respectively. A purely absorptive 2D NMR spectrum with frequency sign-discrimination along both axes was achieved with time-proportional phase incrementation (TPPI), providing an effective frequency range of 4.5 kHz along the

H dimension. A total of 50 t1 (1H) and 350 t2 (31P) points were acquired, zero-filled to (128 × 1024) points, and apodized with 50 Hz Lorentzian (t1) and 80 Hz Gaussian (t2) broadening prior to 2D Fourier transformation. A total of 1664 signal transients/ t1-value were accumulated, using relaxation delays of 4.0 s. The total experimental time was 92 h. We verified that applying high-power 1H decoupling did not narrow the 29Si or 31P NMR peak widths perceptibly, and all acquisitions were performed without 1H decoupling. No signal apodization was used in the processing of 1D 31P NMR data, whereas 150 Hz Gaussian broadening was employed for that of 29Si and 250 Hz for those of 13C and 23Na. Chemical shifts are quoted relative to neat tetramethylsilane for 13C and 29Si, whereas 85% H3PO4(aq) and 1 M NaCl(aq) were used for 31P and 23Na, respectively. All remaining experimental details are provided in the figure captions. Despite using large sample volumes of 240 µL (i.e., masses ≈ 180 mg), the low effective amounts of NMR-active nuclei in the specimens, coupled with very long spin-lattice relaxation times of 29Si and 31P for the case of directly excited spectra, lead to very long experimental times. For example, the total acquisition times (i.e., excluding time spent for optimizing NMR parameters) for the set of 29Si NMR spectra of the left and right columns of Figure 1 amounted to 25 days and 100 h, respectively, whereas the corresponding 31P NMR data in Figure 4 required total acquisition times of 10 days (left column) and 60 h (right column). The long data-collection intervals required for completing even the standard experimentation employed herein, combined with the absence of triple-resonance capabilities of our current 6 mm NMR probehead,48 unfortunately precluded the possibility of performing some potentially more informative experiments. 3. Results 3.1. 29Si NMR. As the MBG pore-wall composition is homogeneous over ∼10 nm,38,42,44 it is impossible to experimentally determine the individual compositions of each CaP and silica-glass component. However, a key feature of the porewall model proposed in ref 48 is the significant consumption of Ca by the CaP clusters (see Table 2). If a stoichiometric Ca3(PO4)2 composition is assumed for the latter,48 the present S85 stoichiometry may be cast as (Ca0.05SiO2.05)-0.025[Ca3(PO4)2]. Hence, little Ca remains for depolymerizing the silicate glass network and this qualitative conclusion remains for all physically conceivable xCa/xP ratios between 1.0 and 1.7. As discussed further in section 4.1, the silicate network is dominated by Q4 building blocks, accompanied by minor 2 3 n and QCa tetrahedra: the notation QCa implies n amounts of QCa Si-O-Si bonds involving bridging oxygen (BO) atoms,50,51 whereas the subscript indicates that calcium ions charge-balance the (4 - n) nonbridging oxygen (NBO) ions. However, although this structural picture of the CaO-SiO2 pore-wall component accounts for its bulk part, the large surface area of the S85 mesoporous material also implies a diversity of SiOH surface moieties.50–52 The amounts of such QH3 and QH2 silanol groups (where H specifies charge balancing of NBOs by protons) are n 48 . significantly larger than that of QCa 29 Si NMR allows for monitoring the changes in the {Q4, QHn , QnCa} speciation during SBF exposure of S85. The experimental NMR spectra in Figure 1 are shown together with their deconvolutions into component peaks. NMR acquisitions employing direct excitation by single pulses (left column) quantitatively reflect the Qn speciation of the entire sample, whereas those recorded by 1H f 29Si CP with a contact interval τCP )

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Figure 1. Experimental 29Si NMR spectra recorded at 9.4 T from (a and b) pristine and (c-n) SBF-treated samples, using either direct excitation (left panel) or CP from protons with a contact interval τCP ) 2.0 ms (right panel). Deconvoluted signal components are represented by gray lines, with peak assignments displayed at the top of each column. The curve beneath each spectrum represents the deviation between the experimental and best-fit spectrum.

2.0 ms (right column) semiquantitatively reveal the 29Si species at the MBG surface, as they emphasize 29Si in close proximity to protons of OH groups and physisorbed water molecules.50–52 The associated NMR parameters are listed in Table 3. The 29Si peak assignments used herein (see top of Figure 1) and in ref 41 were rationalized and thoroughly explained in ref 48. Note that the spectral region around -92 ppm may carry minor NMR 3 units, besides those of the dominating QH2 signals from QCa 50,51 groups. As expected, the directly excited MAS NMR spectrum from S85 (Figure 1a) reveals a clear dominance of

Gunawidjaja et al. Q4 tetrahedra, accompanied by minor fractions of QH3 and QH2 units. The NMR spectra of the current S85 sample (Figure 1, parts a and b) agree well with those obtained previously from a similar S85 specimen/composition.48 The primary distinction is that the present CPMAS NMR spectrum also reveals small 2 units. This is readily explained by the slightly signals of QCa higher amount of Ca2+ in the present silicate component, Ca0.05SiO2.05, relative to that previously employed, Ca0.03SiO2.03.48 The 29Si NMR spectra following 0.5 h of SBF exposure (Figure 1, parts c and d) display strong alterations relative to those of the pristine S85 sample in Figure 1a and 1b. Whereas only minor differences are discernible between spectra from different sbfN samples, they consistently manifest a higher NMR spectral resolution than that of Figure 1a. This is attributed to a slight narrowing of the Si-O-Si bond angle distribution of the tetrahedral network units on SBF immersion, which results in a minor “ordering” of the MBG pore-wall structure. However, the mean Si-O-Si angles remain almost unchanged and thereby also the respective 29Si NMR peak maximum from each Qn species. By using the set of deconvoluted 29Si NMR signal components displayed in Figure 1, we onward focus on the relative amounts of the QnM (M ) H, Ca) building blocks and their evolution for increasing SBF-soaking intervals (τSBF), as displayed in Table 3 and represented graphically in Figure 2. The most obvious trend during the first 4 h of SBF soaking is a 2 units due to dissolution of Ca2+ ions from the depletion of QCa surface (see Figure 2, parts a and b). This process is primarily responsible for a reported increase of the [Ca2+] in the SBF solution following short exposure-intervals of BGs.3,4,8,11,19,23,39 However, as the (less soluble) CaP clusters consume most of the available amount of Ca of the sample, the resulting low Ca dissolution from the silicate pore-wall part implies a marginally increased [Ca2+] of the SBF medium, as observed experimentally for a similar S85 MBG sample.44 Note the trend of a slightly increased network connectivity revealed both in the bulk and surface regions: this is reflected by the enhanced amount 2 tetrahedra over 1 of QH3 species at the expense of QH2 and QCa week of SBF treatment (Figure 2a-d). A potential problem with CP-acquired NMR spectra is their dependence on the particular choice of the contact interval τCP. In our present comparison of samples subjected to distinct SBFexposure intervals, τCP must be kept relatively short to ensure the selective probing of the material surface, by restricting direct 1 H f 29Si magnetization transfers within a surface thickness of a few angstroms, while simultaneously minimizing spin diffusion among protons and relaxation processes during CP. Note that our conclusions drawn from Figure 2 (Table 3) are based on results about the relatiVe integrated signal-intensities of the various {Q4, QnH, Q2Ca} units for each as-recorded CPMAS NMR spectrum, which is independent of its total integrated intensity and thereby also significantly less affected by the content of physisorbed water. Over time, however, the latter cannot be ensured to remain constant for each sample (or between samples), despite that all specimens were prepared and stored under identical conditions. The question then arises as to whether variable water contents could also lead to distinct CP dynamics among the various SiO4 surface units: this would have implications for our conclusions drawn from Figure 2. Fortunately, we have not observed any indications of such problems, as demonstrated below. Figure 3 reveals the CP-acquired 29Si NMR signal intensities of the various SiO4 surface species for increasing τCP-values, as recorded from the pristine S85 (Figure 3a) and sbf7d (Figure

Apatite Formation Studied by Solid-State NMR TABLE 3:

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29

Si NMR Dataa Q4

QH3

QH2b

sample

-δ (ppm)

population (%)

fwhm (ppm)

-δ (ppm)

population (%)


109.7 110.5 109.6 110.9 110.9 111.1 111.0

75 75 75 72 73 73 76

11.2 10.2 9.7 9.3 9.1 9.0 9.1

100.2 100.8 100.0 101.4 101.3 101.5 101.3

18 18 21 24 24 24 21


109.4 110.3 110.5 110.4 110.8 110.9 111.1

25 27 30 27 28 25 28

11.7 9.8 9.7 9.8 9.2 8.9 9.1

100.2 100.6 100.7 100.7 100.9 101.0 101.0

55 56 56 57 61 63 63

fwhm (ppm)

-δ (ppm)

2 c QCa

population (%)

fwhm (ppm)

92.2 93.9 92.0 93.0 91.9 93.0 93.0

7 7 4 4 3 3 3

12.0 12.0 8.0 7.9 7.3 8.8 8.8

Cross Polarization 7.9 91.1 7.2 92.2 7.1 92.4 6.9 92.6 6.8 92.3 6.7 92.4 6.8 92.3

15 14 12 15 11 12 9

6.8 8.1 8.1 8.9 8.0 8.1 7.5

Single Pulse 8.5 7.1 7.4 7.8 7.8 7.3 7.3

-δ (ppm)

population (%)

fwhm (ppm)

83.0 82.0 83.0 83.0 82.0

5 3 2 1 0.4

11.6 12.0 10.3 5.0 5.1

a

Results of deconvoluting the experimental NMR spectra recorded either by single pulses or cross-polarization (Figure 1). Each peak component is characterized by a chemical shift (δ; accurate within (0.25 ppm), relative population (uncertainty (2 percentage units), and full 3 tetrahedra. c Only a few width at half-maximum (fwhm; accuracy (0.3 ppm). b This NMR signal may comprise minor contributions from QCa CP-acquired NMR spectra reveal significant signals from this unit.

2 Figure 2. Plots of the speciation of (a and b) QH2 and QCa , (c and d) Q3H, and (e and f) Q4 SiO4 units against the SBF-soaking interval (τSBF), with relative populations (listed in Table 3) obtained from spectra as acquired by single pulses (left panel) and by CP (right panel). Note the use of a logarithmic horizontal scale and that the vertical scales of all plots span the same range (despite that the starting and ending values vary). The dotted vertical lines mark the data of the pristine S85 sample.

Figure 3. Integrated signal intensities from the various silicate surface units for a series of 1H f 29Si CPMAS experiments conducted for increasing values of τCP and recorded from (a) S85 and (b) sbf7d. The experimental conditions were identical to those of Figure 1, except for the number of accumulated signal transients employed, which varied between samples and τCP-values so as to achieve comparable S/N in all NMR spectra. To improve clarity, error bars are not included for data associated with τCP < 1 ms.

3b) specimens, i.e., the two extreme members of our series of samples. Essentially identical NMR signal buildup is (within experimental error) observed in both cases. These results further corroborate our peak assignments: 29Si in Q4 tetrahedra display the slowest CP dynamics due to their largest distances to surrounding protons. The silicate units involving directly attached OH groups, QH3 and QH2, provide the most rapid NMR signal growth and reach their maximum intensities for τCP ≈ 2 2.5 ms. The minor amounts of QH2 and QCa tetrahedra lead to weak NMR signals and thereby relatively large uncertainties

in their best-fit fractions; yet, the comparatively fast signal 2 tetrahedra from S85 in Figure 3a clearly buildup of QCa corresponds to the expected behavior from a 29SiO4 surface unit devoid of Si-OH bonds but still in close proximity to neighboring protons of silanol groups and physisorbed water molecules. Over the time scale up to τCP ) 12 ms probed in Figure 3, signatures of relaxation during CP is only observed for τCP > 5 ms; this is most clearly revealed by the NMR signal decay of the QH3 units. The contact interval of 2.0 ms used for recording the NMR spectra in Figure 1 optimizes the two (weakest) signals

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Figure 4. 31P MAS NMR spectra of (a and b) pristine and (c-l) SBFexposed S85 samples, obtained either by direct excitation (left panel) or cross-polarization with τCP ) 4.0 ms (right panel). Numbers at the outermost spectral portions specify the respective maximum (in ppm) of the main orthophosphate 31P NMR peak. The asterisks in spectra a and b mark a minor resonance assigned to P-O-Si moieties (ref 48).

2 stemming from QH2 and QCa moieties, while being sufficiently short to avoid influences from relaxation (see Figure 3) and thereby providing spectra that semiquantitatively reflect the relative SiO4 surface-speciation of each sample. We stress that the CP-derived results of Figure 2 only allows for establishing trends of the evolution of the Qn speciation for increasing SBF exposure. Yet, it is gratifying that the results of Figure 2 were reproduced (within experimental error) when instead using the shorter value τCP ) 0.5 ms (see Figure S2 of the Supporting Information). 3.2. 31P NMR. Figures 4a and 4b depict 31P MAS NMR spectra of the pristine S85 sample, as recorded using either single pulses or 1H f 31P CP. They display excellent agreement with previous results.48 The CaP clusters feature a broad NMR signal around 3 ppm, which is typical of an amorphous orthophosphate phase.53–56 Both spectra in Figures 4a and 4b also manifest a minor resonance around -6 ppm, assigned to P-O-Si moieties.48 Their amount diminishes rapidly during an hour of SBF immersion and is ignored in the subsequent analyses. All other results of Figure 4 were obtained from the corresponding set of SBF-exposed samples. Very minor alterations of the NMR spectra occur for increased SBF-soaking intervals up to 24 h. This evidence the close similarity between the local 31P environments of the CaP clusters present in the MBG structure and those of the ACP surface layer initially forming during SBF soaking, thereby corroborating the proposed role of the CaP clusters to promote ACP formation.48 The ACP/HCA creation is revealed in Figure 5a by a clear increase in the phosphorus content of the sample after 24 h of SBF soaking and by a pronounced 31P NMR peak narrowing in the spectra of Figure 4. Figure 5b plots the full width at half-

Figure 5. Plots of various parameters associated with the bioactive layer for increasing values of τSBF: (a) total phosphorus content of the sample in atom % out of the cations. (b) The fwhm of the 31P NMR peaks shown in the left panel of Figure 4, measured directly without reference to a specific peak-shape function. (c) The 31P NMR-derived relative amounts (see Table 4) of crystalline HCA and ACPtot, the latter representing the net contributions from CaP and ACP components. Dotted lines mark the data of the pristine S85 sample. Whenever not specified, the error bar is within the size of the symbol.

maximum (fwhm) height of the NMR signals versus τSBF: the 31 P peak narrows from 6.46 ppm (1045 Hz) in the spectrum of S85 down to 2.44 ppm (395 Hz) from that of sbf7d. This reflects a progressive ordering of the ACP layer into crystalline HCA. These results may be compared with previously reported reductions of 31P NMR peak-widths, from ≈6.1 ppm (of the formed ACP) to ≈4.1 ppm, as observed from SBF soaking of CaO-SiO2 glasses.21 Further, the onset of HCA formation around τSBF ≈ 24 h suggested by the 31P NMR peak-width alterations in Figure 5b is confirmed by the powder XRD results shown in Figure 6: the sample sbf1d reveals weak, yet clearly discernible, HA-type reflections (Figure 6h) that enhance significantly during the subsequent week of SBF immersion. The nearly identical NMR spectra recorded by single pulses (that quantitatively reflect the entire 31P content of the sample) and those obtained by cross-polarization from nearby protons verify that all components (CaP, ACP, HCA) of the MBG specimen and its SBF-induced surface layer comprise 1H-bearing species, such as water molecules and OH groups. The presence of the latter in the CaP clusters was previously demonstrated,48 whereas ACP is known to comprise structural water.56 The 1H NMR spectrum of HA (or HCA), on the other hand, is known to reveal a characteristic 1H resonance around 0 ppm from OH groups, together with NMR signals from physisorbed water.57,58


Figure 6. Powder X-ray diffractograms for the as-indicated S85deriving samples, displayed together with that of crystalline HA.82 All identifiable “sharp” peaks in (a-g) stem from HCA, except those (encircled) from a minor quartz impurity in (a).

Figure 7a shows a 1H-31P HETCOR NMR spectrum recorded from the sbf7d sample. Such a 2D NMR spectrum reveals which 1 H structural sites mediate magnetization to which 31P sites. The spectrum manifests two main 2D NMR peaks from protons associated with chemical shifts around 0.1 and 4.6 ppm, respectively, and may be contrasted with that obtained from

J. Phys. Chem. C, Vol. 114, No. 45, 2010 19351 pristine S85 in ref 48. It is noteworthy that although the primary source of the 31P magnetization derives from OH groups of HCA, this 1H NMR resonance (≈0.1 ppm) is not easily discernible in the directly acquired 1D NMR spectrum shown in Figure 7b; on a vertically expanded scale, this peak is identifiable as a weak and broad component. The spectrum in Figure 7b displays a main peak stemming from water molecules, accompanied by minor resonances from organics molecules, as discussed in the Supporting Information. The strikingly different signal intensities observed in the 1H projection of the 2D spectrum and the directly excited 1D MAS counterpart may be rationalized from the relatively low amount of P compared to Si, coupled to that the water reservoir of the sample is primarily associated with its silica portion. Focusing on the two distinct 31 P NMR signals, we observe a narrow peak around 3.0 ppm (Figure 7c; top trace) assigned to nanocrystalline HCA, and a significantly broader resonance (bottom trace) that stems from the amorphous ACP and CaP components of the sample. To summarize, the 2D HETCOR spectrum establishes, as expected, 31 P NMR signals from a (nano)crystalline HCA phase and amorphous phosphorus-bearing components. To gain more quantitative insight into the ACP/HCA formation, we deconvoluted the 31P MAS spectra (Figure 4; left column) into two signals, accounting for the amorphous CaP and ACP components on one hand (“ACPtot”) and the more ordered HCA phase on the other. The best-fit 31P NMR spectra are presented in the Supporting Information, whereas the resulting NMR parameters are listed in Table 4. Owing to the very similar NMR characteristics of the CaP clusters present at the S85 pore wall and the ACP layer forming on SBF exposure, it is not possible to separately distinguish their 31P NMR contributions by spectral deconvolution. Indeed, attempts to include a third peak in the iterative fitting, consistently reduced to the results of the two-peak fits of Table 4. The relative fractions of HCA and amorphous orthophosphate components are plotted in Figure 5c. As was concluded qualitatively by monitoring trends of 31P fwhm peak-widths (Figure 5b) and results from XRD (Figure 6), the sbf1d sample only comprises a minor amount (22%) of HCA out of all

Figure 7. (a) 2D 1H-31P HETCOR NMR spectrum recorded at 9.0 kHz MAS frequency from the sbf7d specimen and shown together with sum projections along each spectral dimension. Contour levels are drawn from 2% of the maximum 2D peak amplitude. (b) 1H NMR spectrum acquired with a Hahn spin-echo at 9.0 kHz MAS. It reveals a primary resonance around 4.6 ppm from water molecules and a set of narrow peaks around 3.6 ppm (OCH2) and 1.1 ppm (CH3), stemming from minor amounts of organic molecules; see the Supporting Information. (c) Slices taken through the as-indicated 1H chemical shifts at 0.1 ppm (top trace) and 4.6 ppm (bottom) of the 2D spectrum. Each slice reveals a narrow (top) and broad (bottom) 31P NMR peak component, stemming from nanocrystalline HCA and amorphous ACP/CaP sample components, respectively. The bottom trace is displayed with a twofold vertical expansion relative to that at the top.

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TABLE 4:

Gunawidjaja et al.

31

P NMR Dataa ACPtotb

HCA

sample

δ (ppm)

population (%)

fwhm (ppm)

δ (ppm)

population (%)

fwhm (ppm)

S85c sbf1hc sbf4h sbf1d sbf3d sbf7d

3.1 3.1 3.3 3.4 3.5 3.8

100 100 100 78 55 43

6.4 5.9 5.7 5.7 5.4 4.8

3.2 3.1 3.0

22 45 57

2.8 2.3 2.0

a Obtained by deconvoluting the MAS spectra in the left column of Figure 4; the best-fit spectra are presented in the Supporting Information. The uncertainty of each parameter is as follows: δ ((0.2 ppm); fwhm ((0.25 ppm); relative population ((1.5%), except for sbf3d ((2.5%) and sbf7d ((4.0%). b This component represents the net contribution from the SBF-induced ACP phase and the CaP clusters of S85. c These NMR spectra also comprise a minor peak stemming from P-O-Si structural fragments (ref 48) characterized by δ ≈ -6 ppm, fwhm ≈ 8 ppm, and largest relative fraction 7% (in S85). This peak intensity decreases rapidly on SBF exposure and was ignored by renormalizing the fraction of ACPtot to unity.

phosphorus-bearing phases. However, it grows significantly as the SBF treatment proceeds and becomes the major component after 1 week. The 31P NMR peak maximum (3.0 ppm) and width (2.0 ppm) from the HCA phase of the sbf7d surface layer accord well with previously reported parameters from (nano)crystalline (carbonate-bearing) calcium orthophosphates53–55,58 and may also be contrasted with our NMR spectrum of crystalline HA that revealed a 31P peak with δ ) 2.80 ppm and a fwhm of 0.93 ppm (displayed in the Supporting Information). On the basis of both our 31P NMR and XRD observations, we conclude that the nanocrystalline HCA particles of the sbf7d specimen have reached a relatively high degree of maturity; compare, for instance, with published powder XRD results from HCA.59,60 3.3. 23Na NMR. Table 2 reveals small but significant amounts (0.3-0.5 wt %) of Na in all sbfN samples, stemming from the incorporation of sodium ions into the ACP and HCA phases. All 23Na (spin 3/2; 100% natural abundance) MAS NMR spectra in Figure 8 display a broad peak (fwhm ≈ 14 ppm), centered around a mean 23Na shift of -7.5 ppm and tailing asymmetrically toward lower shift values. The latter feature reflects distributions of 23Na quadrupolar coupling constants, as typically observed from quadrupolar spins in disordered phases.50 The very similar NMR characteristics of all samples verify an essentially unaltered 23Na environment within the phosphate surface layer for progressively increasing values of τSBF. This is, to our knowledge, the first 23Na NMR report on sodium environments of an SBF-treated bioglass. The NMR spectra also display minor background signals from the probehead and impurities in the zirconia rotors (that were found to vary slightly between different rotors). These signals are overall broad (see Figure 8, spectra h and i), except for a sharp component around +7 ppm, which incidentally is close to the 23Na shift of NaCl. Hence, the consistently low intensity of this peak in all NMR spectra precludes the possibility of NaCl precipitation that could stem from an incomplete washing of the specimens after the SBF treatment; this observation is important to bear in mind when considering the 13C NMR results presented below. As described in ref 61, we generated numerically calculated 23 Na MAS NMR spectra employing all relevant experimental parameters and assuming uncorrelated Gaussian distributions

Figure 8. (a-f) Experimental 23Na MAS NMR spectra obtained from the as-indicated samples at 9.4 T and 8.50 kHz MAS. The signal at ∼7 ppm marked by an asterisk in spectrum a stems from impurities of the zirconia rotors; their extents are most clearly seen from the NMR spectra recorded from two nominally sodium-free specimens, i.e., the pristine S85 sample (h) and tetrakis(trimethylsilyl) silane (i; “blank”). (g) Best-fit numerically simulated 23Na spectrum, employing the j iso ) -4.2 following average NMR parameters and distribution widths: δ j Q ) 1.7 MHz, σiso ) 4.0 ppm, and σQ ) 1.2 MHz. All NMR ppm, C spectra resulted from 10 800 accumulated signal transients, except for using 3600 transients in (h) and (i); these spectra are therefore displayed with a threefold vertical expansion.

of 23Na quadrupolar coupling constants and isotropic chemical shifts. Fitting of the experimental NMR spectra resulted in an j iso ) -4.2 ( 1.5 ppm, average 23Na isotropic chemical shift of δ j Q ) 1.7 ( 0.6 MHz, a mean quadrupolar coupling constant C and associated widths σiso ) 4.0 ppm and σQ ) 1.2 MHz for the distributions in chemical shifts and quadrupolar coupling constants, respectively. The best-fit spectrum is displayed in Figure 8g. These 23Na parameters may be compared with those j iso ≈ -4 ppm; recently reported from sodium-bearing HCA (δ j iso ≈ -2 j Q ≈ 1.2 MHz),62 as well as from tooth and bone (δ C j Q ≈ 1.5 MHz).63 The close agreement, which is within ppm; C experimental errors between the different studies, suggest very similar local 23Na environments in the SBF-induced MBG surface layer to those found in HCA and/or natural bone. However, extraction of more detailed structural information is complicated; for instance, the 23Na chemical shift may not directly convey the Na coordination number, since it additionally carries an explicit dependence on the Na-O distances,64 as also pointed out by Laurencin et al.63 3.4. 13C NMR. It is well-known that carbonate ions may substitute into hydroxyapatite structures (such as bone and tooth)


J. Phys. Chem. C, Vol. 114, No. 45, 2010 19353 HCA:62 169.8 ppm; fwhm ) 2.8 ppm. However, it is at this stage difficult to make firm structural analogies as the chemical shift range of 13CO32- ions in inorganic phases is relatively small.50 4. Discussion

Figure 9. 13C CPMAS NMR spectra recorded from the as-indicated samples. They reveal weak signals from carbonate ions of the ACP/ HCA components of the SBF-induced bioactive layer. By accounting for the different numbers of accumulated transients of (a) 12 288, (b) 17 408, and (c) 33 792, the spectra are scaled vertically so as to quantitatively reveal the relative carbonate content of each specimen.

in amounts up to several mol %.59,60,62,65–68 The replacements are classified as “A” or “B” types, depending on whether CO32replaces OH- or PO43- ions, respectively, or combinations 2thereof, “AB”. They are often accompanied by PO34 f HPO4 substitutions and generally lead to a calcium deficiency in the HCA structure.62,65–68 The consequences of these substitutions on the local phosphorus environments have been explored by 31 P NMR on HCA and natural bone.53,55,62,66,69–71 While recent 13 C NMR studies have explored the organic/inorganic interface of bones and teeth,72,73 the dominating carbon contributions from organic constituents generally mask the smaller signals from carbonate ions. Consequently, the current insight from NMR into the local 13CO32- environments in naturally biomineralized tissues is very limited.66 More surprising is the present void of investigations targeting carbonate ions of phosphate phases generated in vitro, such as the surface layer of BGs subjected to SBF. To our knowledge, the 1H f 13C CPMAS NMR results in Figure 9 represent the first report from a biomimetic apatite-like layer. In the Supporting Information, we motivate further that the 13C NMR signals of Figure 9 indeed stem from carbonate ions within the ACP/HCA phases. Due to the low carbonate content of the samples and the low natural abundance of 13C (1%), the attainable spectral S/N is very poor, despite spending between 1 and 2 days per sample for data collection. Nevertheless, it may be concluded that the 13C resonance of the ACP/HCA components is characterized by a peak around 169.5 ( 0.5 ppm and a fwhm of 4 ppm (≈400 Hz): this is representative for carbonate ions in a disordered structure.62,66,74 Within the experimental S/N, no significant changes are revealed in the 13 CO32- environments over 1 week of SBF immersion (Figure 9). Our mean 13C chemical shift value may be contrasted with those reported by Beshah et al. of 166.5 and 170.2 ppm for A and B type HCA, respectively, whereas carbonate-containing ACP revealed a peak around 168.2 ppm with a fwhm of 3 ppm (≈300 Hz).66 Hence, the presently observed 13C peak position ≈169.5 ppm is very similar to that of B-type HCA66 but also to one of the two 13CO32- environments reported from AB-type

Here, we start by reviewing the main structural/topological distinctions between “traditional” MPBGs and MBGs of the ternary CaO-SiO2-P2O5 system (section 4.1). Focusing on steps 1-3 of the HM, section 4.2 discusses the surface reactions of the silicate part of the MBG pore wall, as probed by 29Si NMR of the sbfN samples. We conclude by highlighting the two subsequent processes of ACP (HM, stage 4) and HCA (HM, stage 5) formation. 4.1. Composition-Bioactivity Correlations for BGs and MBGs: Similarities and Distinctions. Here we contrast the structural and topological distinctions between MPBGs and MBGs and their bearing on the observed bioactivity. The bioactivity of MPBGs is dictated solely by their composition, whereas the ACP/HCA formation on MBG surfaces may in principle be controlled by several textural effects, such as the specific surface area and pore volume, the (distribution of) pore sizes, and the mesoporous arrangement.4,5,18,42 However, the last feature has recently been demonstrated to be inconsequential for the bioactivity.41 The impact from other textural properties on the bioactivity is not well-understood; however, for MBG compositions devoid of P, the total number of silanol groups (in turn dictated by the surface area) is most likely the most influential factor. The composition-texture-bioactivity correlations of sol-gel prepared BGs are intermediate between those of MPBGs and MBGs.19,20 For MPBGs, the amount of glass modifier(s) relative to that of Si is predominantly dictating the bioactivity through their effect on the aVerage number of BO atoms (nj) per SiO4 tetrahedron, which is related to the ratio between the oxygen and silicon molar fractions associated with the silicate network:50,51

nj ) 2(4 - xO /xSi)

(1)

nj is sometimes referred to as the “network connectivity”51 or Stevel’s parameter Y.75 It has been demonstrated that MPBGs only display bioactivity for a compositional range associated with nj e 3.76,77 Such silicate networks correspond to twodimensional ring structures, built primarily by Q2 and Q3 units.50,51 However, except for ref 36, an equal partitioning of BOs between SiO4 and PO4 tetrahedra has generally been assumed,76–78 which leads to an underestimated value of nj for the silicate network. Only very recent work appears to have accounted fully for the consumption of NBO and Ca2+ by the orthophosphate ions when assessing the average silicate network connectivity from the sample composition.26,34,48,79,80 The underlying reasons for the restriction of nj e 3 may be rationalized by the first three steps of the HM (Table 1); “bioactive” MPBGs display “open” structures that provide high access of Ca and Si to aqueous attack in order to form a modifier-ion-depleted silica gel layer.2,3,8,9,14,23 The gel provides both (i) an increased surface area of the material and (ii) a significant number of SiOH groups (that are not initially present in the MPBG structure). Both features i and ii are intrinsic to the MBG surface on its contact with water. Hence, the requirement of high amounts of glass-modifier ions of an MBG material for ensuring its bioactive property should be significantly relaxed (or even eliminated).18,20 Consequently, concepts such as “network con-

19354


nectivity” becomes immaterial. Indeed, by using nj ) ∑nnxn, where xn is the fraction of Qn units from Table 3, we conclude that the silicate network of S85 is characterized by a high average number of BOs (nj ≈ 3.68). The surface Qn speciation derived from 29Si CPMAS NMR, on the other hand, is characterized by a significantly lower value nj ≈ 3.04. These results may be contrasted with the average number of BOs ≈3.90, as obtained by directly applying eq 1 to the pore-wall composition Ca0.05SiO2.05 (see section 3.1). Obviously, 29Si CPMAS should be the most useful NMR tool to experimentally assess structure-bioactivity correlations, provided that care is taken to employ a relatively short contact interval τCP and keeping it constant for all experiments applied to different samples in a series, as discussed in section 3.1. These examples underline the (unsurprising) fact that the network connectivity, as estimated either from the sample composition or determined experimentally by the single-pulseexcited 29Si NMR spectrum, cannot explain the high bioactivity of the present MBG specimen. Yet, the value nj ≈ 3.04, reflecting the surface connectivity as obtained from 29Si CPMAS NMR, is only around the threshold value expected for a bioactive material.76,77 The reason for this seeming discrepancy is that the role of P for the bioactivity has been ignored. Indeed, a phosphorus-free CaO-SiO2 MBG sampleshaving similar textural properties as well as composition with respect to Ca, Si, and O as the present S85 specimenshas been demonstrated to give negligible ACP/HCA formation.41 Hence, the explanation for the high bioactivity of S85 cannot solely be found from its large number of SiOH groups or favorable textural properties, but from the presence of CaP clusters that open a route to significantly accelerate the apatite formation,41,46,48 as discussed further below. 4.2. Silicate Surface Reactions. 29Si solid-state NMR studies of MPBGs generally evidence stage 3 of the HM by a markedly increased silicate network polymerization following the gradual conversion of the initially present Q2 (and Q3) tetrahedra into higher-connectivity Q3 and Q4 units.12,13,22 For the present S85 MBG specimen, having an oVerall low Ca2+ content (xCa ) 0.10; Table 2) and a negligible amount participating in the silicate network, steps 1 and 2 of the HM are not expected to be ratelimiting reactions, as the MBG surface is already rich in silanol groups. Indeed, the 29Si NMR results (Table 3; Figure 2) merely 2 reveal a condensation of QH2 and QCa into QH3 (and to a much lesser extent Q4) surface units, according to the following schematic reactions:

Table 1 identifies reaction 2 as a part of stage 1 of the HM. 2 The net reaction of 2 and 3 result in QCa f QH3 conversions, 2 3 whereas process 4 reflects QH f QH condensations. Note that reactions 2-4 are formulated to convey the obserVed net

Gunawidjaja et al. transformations; the processes are dynamic and involve all Qn species. As follows from Table 3 and Figure 2, 29Si NMR 2 monitors reactions 2 and 3 through the depletion of QCa tetrahedra that, however, initially only amounts to ≈5% of all surface-associated SiO4 units. Processes 2 and 3 are completed over the first 4 h of SBF immersion, whereas the QH2 f QH3 polymerization reaction 4 proceeds throughout the entire week of SBF soaking (Figure 2). Altogether, reactions 2-4 are responsible for the small net increase in the relative fraction of QH3 species from ≈55% at the pristine S85 surface to ≈63% at that of sbf7d. The rather limited alterations of the CaO-SiO2 portion of the MBG surface on SBF soaking are partially a manifestation of its intrinsically high abundance of SiOH groups. Nevertheless, while accelerating (or even partially avoiding) the initial three stages of the HM, the surface reactions of MBGs devoid of P are overall expected to follow the same mechanistic pathway to crystalline HCA as do the BGs. As illustrated schematically in Table 1, however, for the case of phosphorus-bearing MBG samples subjected to SBF, the HCA formation may follow two (essentially) independent pathways: one proceeds through all five stages of the HM (path A), of which the first three steps are significantly facilitated, whereas path B completely circumvents stages 1-3 by exploiting already present CaP clusters41,48 for a further growth and HCA crystallization. We argue that route B constitutes the key to the significantly enhanced bioactivity of the MBGs. For phosphorus-free MBGs, only path A is relevant. 4.3. Formation and Composition of the ACP/HCA Components. The phosphorus content increases only marginally over the first 4 h of SBF exposure relative to the initial value of S85 (Table 2; Figure 5a). The two formation processes of ACP and HCA partially occur simultaneously, with the former being most active over the initial 24 h of SBF immersion and the latter dominating between 3 and 7 days (Figures 5 and 6). Between 1 and 3 days of SBF immersion, both amounts of ACP and HCA increase, as verified independently from Figures 5a, 5b and 6, as well as semiquantitatively from the pronounced 31P peak narrowing in Figure 5b. Beyond 3 days of SBF exposure, the total amount of P in the solution is consumed. Both Na+ and CO32- ions are present in the sbfN samples already after 30 min of immersion of S85 in SBF. Their incorporation at the earliest ACP formation stages is intriguing and at variance with the common view that these ions enter the HCA phase during its crystallization.3–5 The sodium content increases from 0.3 to 0.5 wt % over 1 week of SBF treatment (see Table 2). However, the integrated 13C signal intensities of Figure 9 indicate that the uptake of carbonate is maximized already after 30 min of SBF immersion, whereupon it reduces, such that the CO32- content of the sbf7d specimen is about half of that in sbf0.5h. The nearly constant 13C peak position and fwhm observed, coupled with the decreased amount of carbonates for prolonged SBF exposure, may reflect that CO32- ions are readily incorporated into the ACP phase but that they are partially expelled during the subsequent crystallization process. This implies that the carbonate content is much lower in HCA relative to its parent ACP phase. More work is required to verify (or possibility disprove) this suggestion. Despite the close resemblance of 31P NMR parameters between the CaP clusters and the ACP surface layer formed during short intervals τSBF e 4 h, Table 4 manifests a gradual increase of the 31P chemical shift from ACP during prolonged SBF treatment. We tentatively attribute this trend as stemming from an increasing uptake of Na+ from the solution (Table 2)

Apatite Formation Studied by Solid-State NMR that induces a slight 31P deshielding. Analogously, it has been reported36,78 that the simultaneous association of Na+ and Ca2+ around the orthophosphate ions in Na-Ca-Si-P-O MPBGs gives an intermediate 31P chemical shift between 3 and 14 ppm observed from the compounds Ca3(PO4)2 and Na3PO4, respectively.50,54 Naturally, the 31P deshielding is much less pronounced for the present case, due to the minute extent of Na+ for Ca2+ substitutions. 5. Conclusions We have investigated the gross reaction sequence occurring at the surface of an SBF-immersed S85 MBG sample using 29Si and 31P solid-state NMR. 29Si CPMAS results verify Ca2+ 2 dissolution by the depletion of QCa tetrahedra during the first 24 h of SBF treatment. However, only minor alterations occur in the SiO4 speciation of the bulk material and at its surface during 1 week of soaking: the net result is a minor condensation of Q2Ca and Q2H units into Q3H. 31P NMR, combined with analyzed sample compositions and powder XRD results, reveal that the formation of ACP and its subsequent crystallization into HCA starts within the first 4 and 24 h of SBF exposure, respectively. Whereas both ACP and HCA grow simultaneously between 1 and 3 days of SBF soaking, the latter process dominates beyond 3 days. After 1 week, nanocrystalline HCA constitutes the main fraction (≈60%) of the total amount of phosphorus-bearing phases. We also probed the incorporation of Na+ and CO32- ions into the ACP/HCA phases by 23Na and 13C NMR. The local environments of each respective ion display similar NMR characteristics to those reported from synthetic HCA and natural biomineralized tissues.62,63,66 Our results evidence significant sodium and carbonate uptake already after 30 min in SBF. However, while the sodium content of the phosphate layer increased slightly over 1 week, we observed a clear decrease of CO32- ions. Our study of an SBF-soaked S85 MBG sample overall corroborates the relevance of the five-step mechanism proposed for melt-derived BGs by Hench and co-workers.2,3,31 However, two primary distinctions between MBGs and MPBGs need to be considered in relation to the Hench mechanism depicted in Table 1: (i) Fairly well-established relationships between the MPBG composition and its bioactivity10,76,77 are not important for porous bioglasses associated with a large surface area: their surface is already inherently “prepared” to accelerate the first three stages of the HM. For MPBGs, on the other hand, these stages are of crucial importance to first arrange such a surface (i.e., a “silica gel”).2,3,8,9,14,23 This feature explains the loss of bioactivity for MPBGs associated with “too condensed” (i.e., nj g 3) silicate networks.76,77 (ii) Phosphorus-bearing MBGs possess an intrinsic bioactivity-boosting property in the guise of amorphous CaP clusters.41,48 They provide natural environments for the ACP/HCA growth, thereby allowing a circumvention of steps 1-3 according to pathway B of Table 1. We note that (nano)crystalline calcium phosphates have previously been reported to constitute structural components of some gel-prepared BGs20,81 and very recently also in magnesiumbearing MPBGs.34 Nevertheless, their relationship to the presently discussed amorphous CaP clusters of MBGs41,48 is unclear. Molecular dynamics simulations consistently point toward a rather uniform dispersion of PO34 ions over the silicate network of (Na)-Ca-Si-P-O MPBGs, whereas an observed phosphorus clustering merely coincides with the transition to bioinactiVe compositions.24–26 However, whereas the relatively low connectivity of an MPBG network should readily allow

J. Phys. Chem. C, Vol. 114, No. 45, 2010 19355 for its accommodation of orthophosphate ions, the dense silicadominated pore-wall component of calcium-poor MBGs (such as that of the present S85 sample) does not leave much space for such PO43- “defects”: this may be the driving force toward a local nanoscale phase separation of the MBG pore wall. Future work will hopefully shed further light onto the possibly similar or distinct structural role(s) of phosphorus in the MPBG and MBG groups of biomaterials. Acknowledgment. This work was supported by the Swedish Research Council (VR), the Magn. Bergvall Foundation, the Faculty of Natural Sciences at Stockholm University, CICYT Spain (project MAT 2008-00736), and the Comunidad Autońoma de Madrid (project S2009/MAT-1472). P.N.G. and A.Y.H.L. were supported by postdoctoral Grants from the Carl Trygger Foundation and the Wenner-Gren Foundations, respectively. We thank the anonymous reviewers for helpful comments. Supporting Information Available: Additional figures and discussion of 13C and 29Si CPMAS experiments and deconvoluted 31P NMR spectra. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Hench, L. L.; Splinter, R. J.; Allen, W. C.; Greenlee, T. K. J. Biomed. Mater. Res. 1971, 2, 117. (2) Clark, A. E.; Pantano, C. G.; Hench, L. L. J. Am. Ceram. Soc. 1976, 59, 37. (3) Hench, L. L. J. Am. Ceram. Soc. 1991, 74, 1487. (4) Vallet-Regı´, M.; Ragel, C. V.; Salinas, A. J. Eur. J. Inorg. Chem. 2003, 1029. (5) Vallet-Regı´, M.; Colilla, M.; Izquierdo-Barba, I. J. Biomed. Nanotechnol. 2008, 4, 1. (6) Vallet-Regı´, M. J. Chem. Soc., Dalton Trans. 2001, 97. (7) Kokubo, T.; Kushitani, H.; Sakka, S.; Kitsugi, T.; Yamamuro, T. J. Biomed. Mater. Res. 1990, 24, 721. (8) Kim, C. Y.; Clark, A. E.; Hench, L. L. J. Non-Cryst. Solids 1989, 113, 195. ¨ . H.; Karlsson, K. H. J. Non-Cryst. Solids 1991, 129, (9) Andersson, O 145. (10) Lebecq, I.; De´sanglois, F.; Leriche, A.; Follet-Houttemane, C. J. Biomed. Mater. Res., Part A 2007, 83, 156. (11) Saravanapavan, P.; Jones, J. R.; Pryce, R. S.; Hench, L. L. J. Biomed. Mater. Res., Part A 2003, 66, 110. (12) Dietrich, E.; Oudadesse, H.; Le Floch, M.; Bureau, B.; Gloriant, T. AdV. Eng. Mater. 2009, 11, B98. (13) Hayakawa, S.; Tsuru, K.; Ohtsuki, C.; Osaka, A. J. Am. Ceram. Soc. 1999, 82, 2155. (14) Ogino, M.; Hench, L. L. J. Non-Cryst. Solids 1980, 38-39, 673. (15) Ebisawa, Y.; Kokubo, T.; Ohura, K.; Yamamuro, T. J. Mater. Sci. Mater. Med. 1990, 1, 239. (16) Skipper, L. J.; Sowrey, F. E.; Rashid, R.; Newport, R. J.; Lin, Z.; Smith, M. E. Phys. Chem. Glasses 2005, 46, 372. (17) Salinas, A. J.; Martin, A. I.; Vallet-Regı´, M. J. Biomed. Mater. Res., Part A 2002, 61, 524. (18) Arcos, D.; Greenspan, D. C.; Vallet-Regi, M. J. Biomed. Mater. Res., Part A 2003, 65, 344. (19) Peltola, T.; Jokinen, M.; Rahiala, H.; Leva¨nen, E.; Rosenholm, J. B.; Kangasniemi, I.; Yli-Urpo, A. J. Biomed. Mater. Res., Part A 1999, 44, 12. (20) Jokinen, M.; Rahiala, H.; Rosenholm, J. B.; Peltola, T.; Kangasniemi, I. J. Sol-Gel Sci. Technol. 1998, 12, 159. (21) Lin, K. S. K.; Tseng, Y. H.; Mou, Y.; Hsu, Y. C.; Yang, C. M.; Chan, J. C. C. Chem. Mater. 2005, 17, 4493. (22) Hayakawa, S.; Tsuru, S.; Iida, H.; Ohtsuki, C.; Osaka, A. Phys. Chem. Glasses 1996, 37, 188. (23) Ohtsuki, C.; Kokubo, T.; Yamamuro, T. J. Non-Cryst. Solids 1992, 143, 84. (24) Pedone, A. J. Phys. Chem. C 2009, 113, 20773. (25) Tilocca, A.; Cormack, A. N. J. Phys. Chem. B 2007, 111, 14256. (26) Tilocca, A. Proc. R. Soc. London, Ser. A 2009, 465, 1003. (27) Sahai, N.; Tossell, J. A. J. Phys. Chem. B 2000, 104, 4322. (28) Sahai, N.; Anseau, M. Biomaterials 2005, 26, 5763. (29) Tilocca, A.; Cormack, A. N. Langmuir 2010, 26, 545. (30) Tilocca, A.; de Leeuw, N. H. J. Phys. Chem. B 2006, 110, 25810.

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