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J Dent Res. 2010 April; 89(4): 405–410.
PMCID: PMC2840178

In vitro Remineralization of Severely Compromised Bonded Dentin


Biomimetic remineralization is potentially useful for the remineralization of incompletely resin-infiltrated collagen matrices created by etch-and-rinse adhesives. In this study, we tested the hypothesis that structurally altered dentin collagen cannot be remineralized to the same hierarchical order and dimension seen in structurally intact dentin collagen. The remineralization medium consisted of a set Portland cement/simulated body fluid system containing polycarboxylic acid and polyvinylphosphonic acid as biomimetic analogs. Remineralization of air-dried, collapsed hybrid layers was apparent after one month, with hybrid layers remineralized to 80-90% of their thickness after 2-4 months. A hypermineralized layer was seen on the hybrid layer surface, and tubular orifices were occluded with apatite deposits that resembled those present in non-carious cervical dentin. Structurally altered collagen is unlikely to be remineralized to the same hierarchical order and dimension as seen in intact dentin. The aggressively air-dried acid-etched dentin remineralization model also sheds light on the mechanism of sclerotic dentin formation.

Keywords: remineralization, biomimetic analogs, intrafibrillar remineralization, structurally altered collagen, hypermineralized dentin


Natural mineralized dentin exhibits excellent strength and toughness because the collagen fibrils are reinforced by intrafibrillar and interfibrillar minerals (Kinney et al., 2003). These carbonated apatite nanoplatelets have to be sufficiently small and arranged in a specific order to be incorporated into the gap zones of collagen fibrils (Xu et al., 2007). Recent biomineralization studies have shown that fluidic amorphous nanoprecursors and mesocrystalline intermediates play critical roles in hard tissue formation (Cai and Tang, 2008; Cölfen, 2008).

A Portland-cement-based (Tay et al., 2007) biomimetic remineralization protocol has recently been developed (Tay and Pashley, 2008). This protocol involves the binding of two biomimetic analogs to dentin collagen so that the doped collagen can guide the scale and distribution of apatite remineralization. In this protocol, a low-molecular-weight polyacrylic acid (PAA) was used to mimic the stabilization function of dentin matrix protein 1 (DMP1) on amorphous calcium phosphate precursors (He et al., 2005). The PAA reduces these precursors released by interaction of set Portland cement with the simulated body fluid to a nanoscale (Olszta et al., 2003). This sequestration step prevents the nanoprecursors from aggregation and precipitation (Cai and Tang, 2008). The other biomimetic analog, polyvinylphosphosphonic acid (PVPA), is a polyanion that mimics the negative charges of phosphoproteins such as DMP1, phosphophoryn, or bone sialoprotein (Gajjeraman et al., 2007; Baht et al., 2008) and their roles in organizing aspects of mineralization.

Resin-dentin bonds created by contemporary etch-and-rinse adhesives are not durable (Pashley et al., 2007), since degeneration of denuded collagen fibrils within hybrid layers can occur in vivo (Carrilho et al., 2007). Biomimetic remineralization provides a potential means for extending the longevity of these bonds by remineralizing incompletely infiltrated hybrid layers (Tay and Pashley, 2009). Demineralized collagen fibrils may collapse via inter-peptide hydrogen bonding when they are inadvertently air-dried during bonding, which restricts the infiltration of adhesive resin monomers (Pashley et al., 2007). It is unknown whether a similar process prevents the penetration of the PAA-stabilized amorphous calcium phosphate nanoprecursors into the resin-dentin interfaces produced after extensive air-drying.

There has been a recent paradigm shift on the clinical treatment of deep caries lesions. It is now acceptable to leave behind caries-affected or even caries-infected dentin during restorative procedures (Thompson et al., 2008). Carious dentin may contain structurally altered collagen of various thicknesses, depending on the clinician’s perception of the end-point of caries excavation. Since the order between gap zones and collagen molecules is irreversibly altered in denatured collagen (i.e., gelatin) (Zeugolis et al., 2008), the hierarchy of apatite remineralization within the latter may be different from that observed in intact mineralized collagen. In the present study, the limits of biomimetic remineralization of resin-dentin interfaces were examined by means of a demineralized dentin model consisting of a thin surface layer of structurally altered collagen and a subsurface network of collapsed, but structurally intact, collagen fibrils. Extensive air-drying of acid-etched dentin is far removed from clinical practice. However, an understanding of the mechanisms of biomimetic remineralization in collapsed and structurally altered collagen matrices provides critical information that facilitates future attempts to remineralize thick layers of carious dentin. Thus, the hypothesis tested was that structurally altered dentin collagen cannot be remineralized to the same hierarchical order and dimension as seen in structurally intact dentin collagen.

Materials & Methods

Remineralization Medium

Type I white Portland cement (Lehigh Cement Company, Allentown, PA, USA) was sieved and mixed with de-ionized water in a water-to-powder ratio of 0.35:1, placed in flexible silicone molds, and allowed to set and age at 100% relative humidity for 1 wk before use (Tay et al., 2007). We prepared a simulated body fluid (SBF) by dissolving 136.8 mM NaCl, 4.2 mM NaHCO3, 3.0 mM KCl, 1.0 mM K2HPO4·3H2O, 1.5 mM MgCl2·6H2O, 2.5 mM CaCl2, and 0.5 mM Na2SO4 in de-ionized water (Kokubo et al., 1990) and adding 3.08 mM sodium azide to prevent bacterial growth. The SBF was buffered to pH 7.4 with 0.1 M Tris Base and 0.1 M HCl. For biomimetic remineralization, 500 µg/mL of polyacrylic acid (Mw 1800; Sigma-Aldrich, St. Louis, MO, USA) and 200 µg/mL of polyvinylphosphonic acid (Mw 24,000; Sigma-Aldrich) were added to the SBF as biomimetic analogs, with the pH of the latter adjusted to 7.4.

Dentin Bonding

Twenty recently extracted human third molars were collected after patients’ informed consents were obtained under a protocol reviewed and approved by the Human Assurance Committee of the Medical College of Georgia. A flat dentin surface was prepared perpendicular to the longitudinal axis of each tooth by means of a slow-speed Isomet saw (Buehler Ltd., Lake Bluff, IL, USA) under water-cooling. The exposed tooth surface was further wet-polished with a 320-grit silicon carbide paper (Oliveira et al., 2003) to create an enamel-free bonding surface in mid-coronal dentin. Two etch-and-rinse adhesives were used: One-Step (Bisco, Schaumburg, IL, USA) and Single Bond Plus (3M ESPE, St. Paul, MN, USA), with 10 teeth randomly assigned to each adhesive. Each dentin surface was etched with 32% phosphoric acid (Uni-Etch, Bisco) for 15 sec, rinsed with water, and air-dried with 0.207 MPa compressed air for 60 sec applied at 5 mm away from the etched dentin. Each adhesive was then applied to the air-dried dentin with 2 adhesive coats and light-cured for 20 sec by means of a light-curing unit with an output intensity of 600 mW/cm2. This was followed by the incremental placement of two 2-mm-thick layers of a resin composite. Each layer was light-cured separately for 40 sec. Each tooth was then sectioned occluso-gingivally to produce four 1-mm-thick slabs, each containing the resin-dentin interface (i.e., 4 x 10 = 40 specimen slabs per adhesive). These 40 specimens were randomly assigned to experimental (20 specimens/adhesive) and control groups (20 specimens/adhesive).

Biomimetic Remineralization

Each experimental specimen slab was placed over a set Portland cement block (ca. 1 g) inside a glass scintillation vial. The latter was filled with 15 mL of SBF containing the 2 biomimetic analogs and incubated at 37°C. The remineralization medium was changed every month, with its pH (after inclusion of the Portland cement blocks) monitored weekly so that it was above 9.25. This ensured that apatite was formed instead of octacalcium phosphate (Meyer, 1983; Eanes, 2001). The set-up for the control specimens was the same, except that the liquid medium was replaced with SBF only. For each adhesive, 4 experimental specimens were retrieved after 1-4 mos for ultrastructural examination of the extent of remineralization (i.e., 4 x 4 specimens per mo = 16 non-demineralized experimental specimens). For each adhesive, 2 control specimen slabs were examined at the baseline period (i.e., before immersion) for the extent of incomplete resin infiltration via the ammoniacal silver nitrate tracer protocol reported previously (Tay et al., 2002). In addition, 4 different control specimen slabs were examined after 1-4 mos of immersion in the Portland cement/SBF for evaluation of the extent of remineralization (i.e., 2 baseline control specimens + 4 x 4 control specimens/mo = 18 non-demineralized control specimens).

Transmission Electron Microscopy (TEM)

Following reduction of the diamine silver ions ([Ag(NH3)2]+), the silver-impregnated control specimen slabs were fixed in Karnovsky’s fixative, post-fixed in 1% OsO4, dehydrated in ascending ethanol (50-100%), immersed in propylene oxide, and embedded in epoxy resin. For examination of the extent of remineralization, experimental specimen slabs were processed for epoxy resin embedding in the manner described previously, but without silver impregnation. Non-demineralized, 90-nm-thick sections were examined without staining in a JEM-1230 TEM (JEOL, Tokyo, Japan) operated at 110 kV.

To examine the conditions of the collagen matrix in the heavily remineralized experimental specimens, the last 4 experimental slabs in each adhesive that had undergone 4 mos of biomimetic remineralization were completely demineralized in a formic acid/sodium formate buffer (i.e., 16 non-demineralized experimental specimens + 4 demineralized experimental specimens = 20 experimental specimens/adhesive). The last 2 control slabs in each adhesive that had been immersed in SBF were completely demineralized in a similar manner (i.e., 18 non-demineralized control specimens + 2 demineralized control specimens = 20 control specimens/adhesive). After TEM processing, 90- nm-thick sections were stained with phosphotungstic acid/uranyl acetate before examination.


Control Specimens

Intense air-drying resulted in extensive areas of incomplete resin infiltration in hybrid layers created by both adhesives (Figs.1A, ,1B),1B), with extensive silver uptake within the denuded collagen matrices. Control specimens examined after 4 mos of immersion in the remineralization medium exhibited no remineralization within the hybrid layers (Figs. 1C, ,1D1D).

Figure 1.
Transmission electron micrographs (TEMs) of non-demineralized, silver-impregnated resin-dentin interfaces created by the application of (A) One-Step and (B) Single Bond Plus to intensively air-dried acid-etched dentin. Extensive regions of incomplete ...

Experimental Specimens

Resin-dentin interfaces exhibited evidence of remineralization as early as 1 mo (Figs. 2A, ,2B),2B), which became very distinct after 2 mos (Figs. 2C, ,2D),2D), with both intrafibrillar and interfibrillar remineralization within the denuded collagen matrix. Initially, intrafibrillar minerals appeared as nanocrystals (Fig. 2E; Tay and Pashley, 2008) that eventually coalesced to produce larger mineral platelets within the collagen fibrils (Fig. 2F). These platelets were identified by selected-area electron diffraction to be apatite (Fig. 2F-inset).

Figure 2.
TEMs taken from unstained, non-demineralized sections of representative experimental resin-dentin slabs that had undergone biomimetic remineralization (BR) for 1-2 mos. (A) One-Step after 1 mo of BR; (B) Single Bond Plus after 1 mo of BR; (C) One-Step ...

By 3-4 months, most of the hybrid layers had remineralized to 80-90% of their entire thickness, except for a thin basal portion that appeared well-infiltrated with resin and did not exhibit any signs of remineralization. Additional features of hypermineralization (Kwong et al., 2000) could be seen. One-Step-bonded specimens in which a hypermineralized layer was present along the surface of the remineralized hybrid layer are shown in Figs. 3A and and3B.3B. Cohesive failure of the hypermineralized layers occurred during ultramicrotomy for all One-Step-bonded specimens. These hypermineralized layers lacked the hierarchical crystallite arrangement found in a mineralized, intact collagen matrix. An atypical ordered arrangement could be identified (Fig. 3C), with the larger mineral platelets (ca. 25-30 nm) aligning linearly and parallel to one another (Tay et al., 2000). Along the crystallization front (Fig. 3C- inset), each platelet was formed by the fusion of multiple nanocrystals, providing evidence for involvement of mesocrystalline phases in the crystallization process. Hypermineralization features in the Single Bond Plus specimens appeared more subtle (Fig. 3D). The dentinal tubule orifices in these specimens exhibited a phenomenon similar to that in the One-Step specimens, in that they were often occluded by electron-dense mineral casts (Figs. 3D, ,3E).3E). The larger platelets that formed these mineral casts were identified via selected-area electron diffraction to be apatites (Fig. 4F).

Figure 3.
Representative unstained, non-demineralized TEMs of resin-dentin interfaces prepared from intensely air-dried acid-etched dentin that had remineralized after more than 2 mos of biomimetic remineralization (BR). (A) A One-Step-bonded specimen that had ...
Figure 4.
TEMs of phosphotungstic acid/uranyl acetate-stained, demineralized sections taken from experimental specimens that had undergone biomimetic remineralization (BR) for 4 mos revealed the presence of a denatured surface collagen layer created by intense ...

Structurally Altered Surface

Stained demineralized sections of four-month remineralized specimens revealed a 500- to 750-nm-thick structurally altered surface layer created by intense air-drying of acid-etched dentin (Figs. 4A, ,4B).4B). Fibrillar architecture and banding characteristics were absent within this layer, and a bed of unraveled microfibrillar strands (gelatin) was observed (Figs. 4B). A similar altered layer was seen around dentinal tubules that were located directly beneath the etched dentin surface (Fig. 4C). Conversely, collagen fibrils were intact in dentinal tubules that were located more than 10 µm beneath the etched dentin (Fig. 4D).


When collapsed air-dried specimens were immersed in the aqueous remineralization medium, re-expansion of the collagen matrix created a sponge- like effect (Inaba et al., 1995) that probably absorbed the biomimetic analogs from the SBF into the denuded matrix. The intrafibrillar remineralization noted in this investigation provides indirect evidence to support that rehydration of a collapsed, but intact, collagen matrix is a reversible process (Pashley et al., 2007). These principles do not apply to the remineralization of the surface layer of structurally altered, demineralized collagen. This layer is only 500-750 nm thick, despite the presence of a 5- to 8-µm-thick layer of acid-etched dentin. Dehydration of the collagen fibrils could have introduced additional reversible interpeptide hydrogen bonding that increases the denaturing resistance of the bulk of the demineralized collagen matrix (Miles et al., 2005; Armstrong et al., 2006).

The remineralized structurally altered surface layer bore a morphologic resemblance to the surface of naturally occurring non-carious cervical dentin lesions (Kwong et al., 2000; Tay and Pashley, 2004). For both entities, dentinal tubules were occluded with apatite aggregates. Tubular occlusion by apatite crystallites is a feature also shared by transparent dentin beneath caries lesions (Daculsi et al., 1987) and age-induced transparent dentin (Kinney et al., 2005). The etiology of tubular occlusion has been ascribed to a “dissolution and reprecipitation” mechanism (Porter et al., 2005; Zavgorodniy et al., 2008). The results of the present study further suggest that structurally altered collagen plugs must be present in the tubular orifices before tubular occlusion can occur via a “dissolution and reprecipitation” mechanism.

Apatite crystallites within natural hypermineralized intertubular dentin exhibit an atypical head-to-tail alignment (Tay et al., 2000; Tay and Pashley, 2004). Similar features were seen in the remineralized surface layer of intensively air-dried bonded dentin. The association of the remineralized surface layer with a layer of structurally altered collagen provides an alternative mechanism to account for the etiology of hypermineralized dentin. Binding of phosphoproteins to collagen results in conformation changes in the collagen triple helix (Dahl et al., 1998), possibly increasing the flexibility of the hole zones (Landis et al., 2006) to receive calcium phosphate nanoprecursors. Interaction of biomimetic phosphoprotein analogs such as PVPA with intact dentin collagen may achieve similar results. Apatite deposition was considerably more difficult to achieve in type I collagen gels than gelatin gels due to the steric effects of intact collagen on apatite nucleation and growth (Blumenthal et al., 1991). Presumably, alignment of the collagen molecules in a quarter-staggered configuration imposes steric constraints on the apposition of apatites in the hole zone regions. When collagen is denatured, most of these triple helical arrangements are lost as the latter progressively dissociates into the 3 randomly coiled peptide α-chains (gelatin) (Johns and Courts, 1977). When these steric constraints are removed, phosphoprotein analogs such as PVPA may bind to alternative sites along the dissociated gelatin peptide chains. Since there may be as many as 3 times the number of α-chains compared with the original number of triple helices, this may result in hypermineralization when apatite nanocrystals are deposited around the PVPA-bound gelatin.

The results of the present study indicated that although structurally altered collagen matrices are amenable to remineralization in the presence of biomimetic analogs, they are unlikely to be remineralized to the same hierarchical order and dimension as seen in structurally intact dentin collagen. It is important to recognize, however, that these morphologic results were based on two-dimensional TEM images. We are aware that native tissues that have similar morphologic features may have different properties and compositions (Katz et al., 2007). Thus, the nanomechanical properties of remineralized hybrid layers in structurally intact and structurally altered collagen matrices should be further investigated by nanoscopic Dynamic Mechanical Analysis.

Biomimetic remineralization represents a revolutionary approach to improving the durability of resin-dentin bonds by a particle-mediated, non-classic crystallization strategy. We are still at a proof-of-concept stage, since planar remineralization was achieved via placement of a sectioned specimen slab on top of a Portland cement block. Understanding how compromised resin-bonded dentin responds to biomimetic remineralization will help establish the foundation for more sophisticated translational strategies to be designed for delivery of the different components of biomimetic remineralization in a three-dimensional manner to resin-dentin interfaces.


We thank Michelle Barnes for secretarial support.


This study was supported by Grant R21 D019213-01 from the National Institute of Dental and Craniofacial Research (PI. Franklin R. Tay).


  • Armstrong SR, Jessop JL, Winn E, Tay FR, Pashley DH. (2006). Denaturation temperatures of dentin matrices. I. Effect of demineralization and dehydration. J Endod 32:638-641 [PubMed]
  • Baht GS, Hunter GK, Goldberg HA. (2008). Bone sialoprotein-collagen interaction promotes hydroxyapatite nucleation. Matrix Biol 27:600-608 [PubMed]
  • Blumenthal NC, Cosma V, Gomes E. (1991). Regulation of hydroxyapatite formation by gelatin and type I collagen gels. Calcif Tissue Int 48:440-442 [PubMed]
  • Cai Y, Tang R. (2008). Calcium phosphate nanoparticles in biomineralization and biomaterials. J Mater Chem 18: 3775-3787
  • Carrilho MR, Geraldeli S, Tay F, de Goes MF, Carvalho RM, Tjäderhane L, et al. (2007). In vivo preservation of the hybrid layer by chlorhexidine. J Dent Res 86:529-533 [PubMed]
  • Cölfen H. (2008). Single crystals with complex form via amorphous precursors. Angew Chem Int Ed Engl 47:2351-2353 [PubMed]
  • Daculsi G, LeGeros RZ, Jean A, Kerebel B. (1987). Possible physicochemical processes in human dentin caries. J Dent Res 66: 1356-1359 [PubMed]
  • Dahl T, Sabsay B, Veis A. (1998). Type I collagen-phosphophoryn interactions: specificity of the monomer-monomer binding. J Struct Biol 123:162-168 [PubMed]
  • Eanes ED. (2001). Amorphous calcium phosphate. In: Octacalcium phosphate. Monographs in oral science. Vol. 18 Chow LC, Eanes ED, editors. , Editors. Basel: Karger, pp.130-147 [PubMed]
  • Gajjeraman S, Narayanan K, Hao J, Qin C, George A. (2007). Matrix macromolecules in hard tissues control the nucleation and hierarchical assembly of hydroxyapatite. J Biol Chem 282:1193-1204 [PubMed]
  • He G, Gajjeraman S, Schultz D, Cookson D, Qin C, Butler WT, et al. (2005). Spatially and temporally controlled biomineralization is facilitated by interaction between self-assembled dentin matrix protein 1 and calcium phosphate nuclei in solution. Biochemistry 44:16140-16148 [PMC free article] [PubMed]
  • Inaba D, Iijima Y, Takagi O, Ruben J, Arends J. (1995). The influence of air-drying on hyperremineralization of demineralized dentine: a study on bulk as well as on thin wet section of bovine dentine. Caries Res 29:231-236 [PubMed]
  • Johns P, Courts A. (1977). Relationship between collagen and gelatin. In: The science and technology of gelatin. Ward AG, Courts A, editors. , Editors. New York: Academic Press, pp. 164-165
  • Katz JL, Misra A, Spencer P, Wang Y, Bumrerraj S, Nomura T, et al. (2007). Multiscale mechanics of hierarchical structure/property relationships in calcified tissues and tissue/material interfaces. Mater Sci Eng A Struct Mater 27:450-468 [PMC free article] [PubMed]
  • Kinney JH, Habelitz S, Marshall SJ, Marshall GW. (2003). The importance of intrafibrillar mineralization of collagen on the mechanical properties of dentin. J Dent Res 82:957-961 [PubMed]
  • Kinney JH, Nalla RK, Pople JA, Breuning TM, Ritchie RO. (2005). Age-related transparent root dentine: mineral concentration, crystallite size, and mechanical properties. Biomaterials 26:3363-3376 [PubMed]
  • Kokubo T, Kushitani H, Sakka S, Kitsugi T, Yamamuro T. (1990). Solutions able to reproduce in vivo surface-structure changes in bioactive glass-ceramic A-W. J Biomed Mater Res 24:721-734 [PubMed]
  • Kwong SM, Tay FR, Yip HK, Kei LH, Pashley DH. (2000). An ultrastructural study of the application of dentine adhesives to acid-conditioned sclerotic dentine. J Dent 28:515-528 [PubMed]
  • Landis WJ, Silver FH, Freeman JW. (2006). Collagen as a scaffold for biomimetic mineralization of vertebrate tissues. J Mater Chem 16: 1495-1503
  • Meyer JL. (1983). Phase transformation in the spontaneous precipitation of calcium phosphate. Croat Chem Acta 56:753-767
  • Miles CA, Avery NC, Rodin VV, Bailey AJ. (2005). The increase in denaturation temperature following cross-linking of collagen is caused by dehydration of the fibres. J Mol Biol 346:551-556 [PubMed]
  • Oliveira SS, Pugach MK, Hilton JF, Watanabe LG, Marshall SJ, Marshall GW., Jr (2003). The influence of the dentin smear layer on adhesion: a self-etching primer vs. a total-etch system. Dent Mater 19:758-767 [PubMed]
  • Olszta MJ, Odom DJ, Douglas EP, Gower LB. (2003). A new paradigm for biomineral formation: mineralization via an amorphous liquid-phase precursor. Connect Tissue Res 44(Suppl l):326-334 [PubMed]
  • Pashley DH, Tay FR, Carvalho RM, Rueggeberg FA, Agee KA, Carrilho M, et al. (2007). From dry bonding to water-wet bonding to ethanol- wet bonding. A review of the interactions between dentin matrix andsolvated resins using a macromodel of the hybrid layer. Am J Dent 20:7-20 [PubMed]
  • Porter AE, Nalla RK, Minora A, Jinschek JR, Kisielowskia C, Radmilovic V, et al. (2005). A transmission electron microscopy study of mineralization in age-induced transparent dentine. Biomaterials 26:7650-7660 [PubMed]
  • Tay FR, Pashley DH. (2004). Resin bonding to cervical sclerotic dentin: a review. J Dent 32:173-196 [PubMed]
  • Tay FR, Pashley DH. (2008). Guided tissue remineralisation of partially demineralised human dentine. Biomaterials 29:1127-1137 [PubMed]
  • Tay FR, Pashley DH. (2009). Biomimetic remineralization of resin-bonded acid-etched dentin. J Dent Res 88:719-724 [PMC free article] [PubMed]
  • Tay FR, Kwong SM, Itthagarun A, King NM, Yip HK, Moulding KM, et al. (2000). Bonding of a self-etching primer to non-carious cervical sclerotic dentin: interfacial ultrastructure and microtensile bond strength evaluation. J Adhes Dent 2:9-28 [PubMed]
  • Tay FR, Pashley DH, Yoshiyama M. (2002). Two modes of nanoleakage expression in single-step adhesives. J Dent Res 81:472-476 [PubMed]
  • Tay FR, Pashley DH, Rueggeberg FA, Loushine RJ, Weller RN. (2007). Calcium phosphate phase transformation produced by interaction of the Portland cement component of white MTA with a phosphate-containing fluid. J Endod 33:1347-1351 [PubMed]
  • Thompson V, Craig RG, Curro FA, Green WS, Ship JA. (2008). Treatment of deep carious lesions by complete excavation or partial removal: a critical review. J Am Dent Assoc 139:705-712 [PMC free article] [PubMed]
  • Xu A-W, Ma Y, Cölfen H. (2007). Biomimetic mineralization. J Mater Chem 17:415-449
  • Zavgorodniy AV, Rohanizadeh R, Swain MV. (2008). Ultrastructure of dentine carious lesions. Arch Oral Biol 53:124-132 [PubMed]
  • Zeugolis DI, Khew ST, Yew ES, Ekaputra AK, Tong YW, Yung LY, et al. (2008). Electrospinning of pure collagen nano-fibres—just an expensive way to make gelatin? Biomaterials 29:2293-2305 [PubMed]

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