Acta Biomaterialia 7 (2011) 1173–1183

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Acta Biomaterialia

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Bone nodules on chitosan–polygalacturonic acid–hydroxyapatite nanocompositefilms mimic hierarchy of natural bone

Rohit Khanna, Kalpana S. Katti ⇑, Dinesh R. KattiDepartment of Civil Engineering, North Dakota State University, Fargo, ND 58105, USA

a r t i c l e i n f o a b s t r a c t

Article history:Received 20 May 2010Received in revised form 15 October 2010Accepted 22 October 2010Available online 26 October 2010

Keywords:Bone noduleHierarchyCollagenMineralModulus mapping

1742-7061/$ - see front matter � 2010 Acta Materialdoi:10.1016/j.actbio.2010.10.028

⇑ Corresponding author. Tel.: +1 701 231 9504; faxE-mail address: kalpana.katti@ndsu.edu (K.S. Katti

The ultimate goal of bone tissue engineering is to develop bony tissues on tissue engineered constructsthat mimic the native bone. Nanoscale characterization of in vitro generated bony tissues on engineeredscaffolds is essential to understanding both the physical and mechanical characteristics of the engineeredbone. Bone nodule formation, a typical early indicator of bone formation was observed on chitosan–polygalacturonic acid–hydroxyapatite (Chi–PgA–HAP) nanocomposite films without the use of differen-tiating media. Thus, the Chi–PgA–HAP substrates designed are osteoinductive and provide an appropriatemicroenvironment for cell organization and tissue regeneration. Imaging using atomic force microscopyrevealed several levels of hierarchical structures of bone in the bone nodules, consisting of mineralizedcollagen fibers, fibrils and mineral deposits in extrafibrillar spaces. The nanoscale elastic properties ofthe collagen and mineral crystals were found to be in close agreement with the experimental and simu-lations results on natural bone reported in the literature. Fourier transform infrared spectroscopy exper-iments indicate the presence of collagen and biological apatite in bone nodules exhibiting thecharacteristics of newly precipitated, immature bone. Collectively, our structural, chemical, and mechan-ical analyses support the conclusion that synthetic bone nodules mimic the hierarchy of natural bone.

� 2010 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

1. Introduction

In vitro regeneration of biological tissues such as liver, heart,cartilage, skin, and bone is an active area of research in tissue engi-neering. Tissue engineering offers a promising alternative to tradi-tional transplants by employing synthetic or naturally derivedengineered biomaterials to replace damaged tissues [1–3]. Materi-als used for hard tissue regeneration such as bone are made mostlyfrom biodegradable and biocompatible natural and synthetic poly-mers, which are used in combination with hydroxyapatite (HAP) todevelop three-dimensional (3D) porous matrices [4–7]. Porousmatrices serve as a template to guide cell organization and growthand allow diffusion of nutrients to transplanted cells, which ulti-mately leads to generation of complex architectures which mimicnative tissues [8]. Appropriate mechanical properties of the scaf-fold are also required to maintain cell–scaffold load transfer mech-anisms during tissue regeneration, which is one of the key factorsin the selection and design of biomaterials, especially in generatinghard tissues such as bone. Various nanoscale reinforcements,including silicates, have also been investigated to enhance themechanical response of the scaffolds [9,10].

ia Inc. Published by Elsevier Ltd. A

: +1 701 231 6185.).

Biological tissues are complex and exhibit structural hierarchy,and tissue regeneration on synthetic 3D scaffolds requires appro-priate design and control of the physical, mechanical, and chemicalbehavior over the macro-, micro-, and nanoscale of scaffolds, to mi-mic the 3D hierarchical architecture of tissues [3,11]. For bonemineralization studies, scaffolds are often treated with variousgrowth factors, such as ascorbic acid, dexamethasone, b-glycero-phosphate, and bone morphogenetic protein-2 (BMP-2), to stimu-late osteogenesis as they would otherwise not form bone[12–14]. Thus, the engineered scaffolds are required to possessosteogenic and osteoinductive properties [15].

Extensive studies of the tissue structure, function, and physiol-ogy of bone exist in the literature. Human bone exhibits seven or-ders of hierarchical structure that spans from sub-nanostructuresto macrostructures [16–19]. At lower levels of hierarchy, bone isa nanocomposite of collagen and mineral crystals of carbonatedHAP, also termed ‘‘biological apatite’’ [20]. The organic matrix ofbone consists of mainly type I collagen and non-collagenous pro-teins. These are the smallest structural building blocks of bonestructure, which are repeated several times leading to functionalbone. Hierarchical structures serve many biological functions, suchas providing mechanical support to cells and promoting and regu-lating cellular functions such as cell adhesion, proliferation, differ-entiation, and vascularization in a physiological environment [21].The unique mechanical properties of bone are derived primarily

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from its hierarchical structure and organization at all levels of hier-archy [17,22]. The size, shape, orientation, and organization ofmineral crystals in collagen fibrils regulate the mechanical func-tion of bone. Often, small misorientations of the mineral crystalsin bone can lead to bone diseases like osteogenesis imperfecta, abrittle bone disease.

Hierarchical structures of various calcifying tissues (turkey andchicken bone) have been studied by various researchers usingmodern experimental techniques such as high voltage electronmicroscopy (HVEM), computed tomography [19,23,24], and smallangle X-ray scattering (SAXS) [25–30]. SAXS and HVEM experi-ments have revealed the size, geometry, orientation, and spatialdistribution of mineral crystals both within and on the surface ofcollagen fibrils. These studies examined thick sections of calcifyingtissues of chicken, turkey, etc., and performed 3D imaging usingHVEM or SAXS to evaluate their structure, and also conducted mi-cro-mechanical analyses.

In our prior work we designed and developed chitosan–polyga-lactouronic acid–hydroxyapatite (Chi–PgA–HAP) biopolymernanocomposites for bone tissue engineering. Biomimetically syn-thesized Chi–PgA–HAP nanocomposite samples exhibited im-proved mechanical strength due to stronger interfacialinteractions [31]. These nanocomposites also exhibited swellingcharacteristics over time when soaked in cell culture medium, asrevealed by in situ physico-chemical and mechanical analyses onsoaked substrates [32]. Further, 3D porous Chi–PgA–HAP scaffoldswere found to be osteogenic and osteoinductive and exhibited for-mation of bone nodules in the absence of differentiating medium[33]. The current work reports nanoscale characterization of bonenodules generated by human osteoblasts on Chi–PgA–HAP nano-composite films. The specific objectives of the present work are:(a) nanoscale structural analysis of bone nodules using AFM tocharacterize the size, morphology, and spatial distribution ofphases present on the surface of bone nodules; (b) to determinethe nanoscale elastic properties of different constituents of bonenodules by making shallow indentations (�2–3 nm) using a mod-ulus mapping technique; (c) to characterize the chemical struc-tures of bone nodules using Fourier transform infrared (FTIR)spectroscopy.

2. Materials and methods

2.1. Synthesis of nanocomposite films

Disodium hydrogen phosphate (J.T. Baker) and calcium chloride(EM Sciences) were used to prepare nanohydroxyapatite particlesby the wet chemistry route. The detailed processing route is dis-cussed elsewhere [34]. Chitosan (molecular weight 190,000, >85%deacetylation) and polygalacturonic acid (molecular weight25,000) were obtained from Sigma–Aldrich chemicals. For filmpreparation, chitosan and PgA powders were dissolved separatelyin 100 ml of deionized (DI) water. The solution pH values were ad-justed to 4.5. The pH of the chitosan and PgA solutions were ad-justed by adding acetic acid and NaOH, respectively. Chitosan–PgA solution was prepared by drop-wise addition of chitosan solu-tion to PgA solution followed by sonication for 3 min. To synthesisethe Chi–PgA–HAP nanocomposite, HAP nanoparticles (1 g per100 ml of DI water) were sonicated for 45 min and then mixedwith sonicated Chi–PgA solution. The resulting solution was soni-cated again for 90 s to obtain a uniform dispersion of HAP nanopar-ticles in the nanocomposite. Further, films of two compositionswere made (Chi–PgA–10% HAP and Chi–PgA–20%HAP) on tissueculture polystyrene (TCPS) petri dishes by the solvent evaporationroute in a clear air laboratory environment at room temperature.For film preparation Chi–PgA–HAP solution was diluted in the ratio

1:10 with DI water and 3 ml of the resulting solution was air driedon TCPS petri dishes. Films made of both compositions were foundto be biocompatible. Chi–PgA–20%HAP was selected for biocom-patibility studies in the present work. 3D porous scaffolds wereprepared using a freeze drying technique described elsewhere [35].

2.2. Cell culture and Alizarin Red S staining

Human osteoblasts (CRL 11,732, ATCC) were cultured inDulbecco’s modified Eagle’s medium (DMEM/F12, Hyclone), sup-plemented with 2.5 mM l-glutamine (without phenol red), 10% fe-tal bovine serum (FBS, ATCC) and 1% antibiotic (G418, JR ScientificInc.), referred as complete medium. Film samples were UV steril-ized for 1 h. Subsequently, 1 � 105 cells were seeded onto filmsamples and placed under standard cell culture conditions (37 �C,5% CO2) in a humidified atmosphere. The complete medium wasreplaced every 3 days. Cell cultures were monitored for 2 and12 h and 10 and 22 days. At the end of predetermined time, phasecontrast/fluorescence micrographs were recorded using an in-verted microscope (Axiovert, Zeiss, Germany). For mineralizationstudies, cell seeded samples were removed from the incubatorafter 22 days of culture, washed three times with phosphate-buf-fered saline (PBS), and fixed with 2.5% glutraldehyde for 6 h. Fixedcultures were washed twice with PBS and then stained with Aliza-rin Red S dye (ARS) (5 mg ml–1) for 5 min. Cultures were rinsedthoroughly with water to reduce non-specific ARS staining. Thepresence of red color is an indicator of calcium deposits in the cul-ture. For nanoscale characterization of bone nodules four nodulesamples were carefully obtained from the same culture before fix-ing the samples for ARS staining. Unstained and unfixed bone nod-ule samples were immediately taken for structural, mechanicaland chemical analyses.

2.3. Cell proliferation determined by MTT assay

Cell proliferation studies were carried out using MTT reagent(3-(4, 5-dimethylthiazol-2-yl)-2,5-diphenyl-terazolium bromide)obtained from EMD Biosciences. The MTT assay reduces the yellowtetrazolium salt to purple formazan crystals by mitochondrialdehydrogenase enzymes of metabolically active cells. Chi–PgA–HAP nanocomposite films were prepared directly on 24-well platesby solvent evaporation. Completely dried samples were then sub-jected to UV sterilization for 1 h. Subsequently, 1.5 ml of cell cul-ture medium containing 2 � 104 cells were seeded into each ofthe 24 wells. Cell proliferation experiments on Chi–PgA–HAP filmswere carried out for 1, 4, 8, and 22 days. For cell proliferation stud-ies on Chi–PgA–HAP scaffolds, 1.5 ml of cell culture medium con-taining 1 � 105 cells were seeded onto scaffolds and incubatedfor 2, 4, 8, 12, and 16 days. At predetermined times 150 ll ofMTT solution (5 mg ml–1 in PBS) was added to each of the wellsand incubated for 6 h. The solution was aspirated and 500 ll ofdimethylsulfoxide (DMSO) (Mediatech Inc., USA) was added to dis-solve formazan crystals. Mean absorbance values were read at570 nm using a microplate reader (Bio-Rad Laboratories, USA).The resulting absorbance values give a quantitative measure of via-ble cells in terms of the cell population. All the experiments wererun in triplicate.

2.4. Nanostructural analysis

The size, morphology, and spatial distribution of the constitu-ents of the bone nodule were characterized by multimode AFM(Veeco Metrology Group, Santa Barbara, CA), equipped with aNanoscope IIIa controller and J-type piezo scanner. The equipmenthad a z-axis resolution of around 0.5 Å. Silicon nitride cantileverprobes (model NP-S20) with pyramidal tips of �20 nm radius of

Fig. 1. AFM height image indicating the topography of micro-sized bundles of Chi–PgA fibers randomly distributed on the surface of the Chi–PgA–20HAPnanocomposite.

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curvature and a nominal stiffness of 0.06 N m–1, were used forimaging under contact mode in a clear air laboratory environment(room temperature, 40–50% relative humidity). Height and deflec-tion images were obtained from several regions of bone nodulesamples to evaluate the topography and sub-surface structures.Sectional analysis was used to measure the feature sizes of constit-uents on nodule surfaces using nanoscope software (AFM TechnicalManual, Veeco Metrology, Santa Barbara, CA).

2.5. Nanoscale modulus mapping

A Hysitron triboscope nanomechanical instrument (Minneapo-lis, MN) equipped with a Nanoscope IIIa controller (VeecoMetrology, Santa Barbara, CA) was used to obtain modulus mapsof the surface of bone nodule samples with a Berkovich (three-sided pyramid, 100–200 nm tip radius) diamond indenter tip.Modulus mapping is a surface nanomechanical characterizationtechnique that enables measurement of the purely elasticproperties of a material surface by applying extremely shallow dis-placements (�2–3 nm). Detailed descriptions of the principle ofmodulus mapping, its mechanistic formulation and applicationcan be found in the literature [36,37]. In a typical experiment flatregions of sample surface were identified by imaging in contactmode and height images were obtained. Further, modulus mapswere acquired from the same area by applying a quasi-static forceof 3 lN with a superimposed 1 lN sinusoidal force at a frequencyof 200 Hz. High resolution modulus maps were obtained on scanareas of 5 � 5 lm and 2.5 � 2.5 lm. Tip calibration was performedusing a standard quartz sample of known elastic modulus.

2.6. Chemical analysis using micro-FTIR

Micro-FTIR was used to evaluate the chemical structure on thesurface of bone nodule samples. In many mineralization studies,bone specimens are fixed to prevent bacterial growth and degrada-tion of the sample before the data is acquired. However, this leadsto concerns that bone sample treatment with a fixative solutioncan potentially alter the chemistry of the mineral phase [38]. Inour study, bone nodules were not treated with fixative solutions.Spectra were acquired on various regions of the bone nodule sam-ples using a Nicolet micro-FTIR spectrometer equipped with a Con-tinulm™ spectroscopy accessory with an internal liquid nitrogencooled MCT/B detector. Spectra were collected at a frequency res-olution of 4 cm–1 using Omnic™ software and processed usingGrams/AI7™ software from Galactic Industries.

3. Results

3.1. Chi–PgA–HAP microstructure and cellular response to the Chi–PgA–HAP nanocomposite

Fig. 1 shows AFM height image (100 � 100 lm) of a Chi–PgA–HAP nanocomposite film. Bundles of Chi–PgA fibers were observedto be uniformly distributed in the Chi–PgA–HAP microstructure.The Chi–PgA fibers were found to be several microns long and 1–2 lm in diameter. Nano-HAP particles with a mean particle sizeof 50 nm were observed as before [39]. A phase contrast image ofosteoblasts seeded on Chi–PgA–HAP nanocomposite films after2 h culture is shown in Fig. 2a. As can be seen, the majority of cellsare observed to have a rounded morphology, indicating poor cellattachment to the substrate, although most of the seeded cells re-mained attached. Fluorescence images were obtained to ascertaincell density and cell viability after 12 h culture. Fig. 2b shows thefluorescence image of a cell seeded Chi–PgA–HAP nanocompositeafter 12 h culture obtained after live/dead cell assay. As can be

seen, the cells appear to be viable on the synthetic Chi–PgA–HAPsubstrate. A rounded cell mass, as observed in Fig. 2b, is indicativeof cell assemblage and organization. The same cell density was ob-tained after 2 h and 12 h culture. This observation suggests thatcells migrate, assemble and organize themselves during 12 h cul-ture to form rounded cell masses (Fig. 2b). With an increase in cellculture time to 8–10 days the cells appear to-collect themselves to-wards a rounded cell mass from all directions as revealed in thephase contrast image taken on the cell seeded Chi–PgA–HAP nano-composite after 10 days of culture (Fig. 2c). Cellular growth on theChi–PgA–HAP nanocomposite was continuously monitoredthroughout the cell culture duration. Osteoblasts generated densestructures (described as bone nodules) on the Chi–PgA–HAP nano-composites without the use of a differentiating medium, whichwere observed and analyzed for mineralization after 22 days ofculture. Fig. 2d shows a phase contrast image of an unstained bonenodule on the Chi–PgA–HAP substrate after 22 days of culture. Alarge number of cells with rounded morphology appear to sur-round bone nodule. A phase contrast image of a stained bone nod-ule is shown in Fig. 2e. Here the appearance of red color on thenodule indicates the presence of calcium deposits. Bone nodulesappear to grow in size from less than 200 lm to a maximum sizein the range 500–800 lm. Cell proliferation studies were per-formed for cell culture durations of 1, 4, 8, and 22 days, using theMTT assay (Fig. 2f). Mean absorbance values increase graduallyfrom day 1 to day 8, followed by higher absorbance values ob-served after 22 days of culture.

3.2. Hierarchical micro/nanostructures of bone nodule

AFM analyses were performed on unstained bone nodule sam-ples. Fig. 3a shows the 3D AFM height image of a 10 � 10 lm scanarea on the surface of a bone nodule sample. The observed heightimage reveals a topography of hierarchical structures in the extra-cellular matrix of a bone nodule synthesized by osteoblasts on theChi–PgA–HAP nanocomposites after 22 days of culture. As seen inFig. 3a, lower levels of hierarchical bone structure are observed,consisting of an array of repeating collagen fibers (�1.7 lm width)and collagen fibrils (�300 nm width) aligned along the long axes ofthe collagen fibers. Mineral deposits are observed to lie on the sur-faces of collagen fibers and fibrils along the long axes of the colla-gen fibers. Collagen fibrils and the spatial distribution of mineral

Fig. 2. Phase contrast image of osteoblasts after 2 h incubation. (a) A fluorescence image indicates that cells are viable and assemble into a cell mass after 12 h cell culture. (b)A phase contrast image indicating cell assemblage and alignment of cells as a rounded cell mass, observed after 10 days culture. (c) A phase contrast image of an unstainedbone nodule after 22 days cell culture. The periphery of the bone nodule appears to be surrounded by a large cell mass with a rounded morphology. (d). A phase contrastimage of a stained bone nodule on the substrate after 22 days culture. (e) Red color on the nodule indicates the presence of calcium deposits. (f) MTT assay results indicatingcell proliferation after 1, 4, 8, and 22 days cell culture.

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deposits in these regions were closely examined by taking highresolution height and deflection images (5 � 5 lm) of the regionsindicated by a box in Fig. 3a. The height image indicated a topog-raphy of mineralized collagen fibrils with a banded appearance,aligned parallel to each other and to the long axes of the collagenfibers, as revealed in Fig. 3b. Irregularly shaped micro/nanosizedmineral patches (�50–1200 nm) were observed on the surface ofcollagen fibrils. A schematic representing regions (1)–(4) is shown

in Fig. 3b. The illustration demonstrates the hierarchical organiza-tion of mineralized collagen fibrils (300 nm diameter) in a singlecollagen fiber (1.7 lm diameter), aligned parallel to each otherand to the long axes of the collagen fibrils. The illustration demon-strates the spatial arrangement of mineral crystals/patches, theirshape, size, location, and orientation on the collagen fibrils. Theschematic representation is scaled relative to the AFM images ob-tained on the regions indicated in Fig. 3a. An AFM deflection image

Fig. 3. (a) 3D AFM height image of the surface of a bone nodule sample indicating the topography of hierarchical structures in the extracellular matrix of the bone nodule. (b)Hierarchical structures of a bone nodule consisting of an array of repeating collagen fibers (�1.7 lm width) and collagen fibrils (�300 nm width) aligned along the long axesof collagen fibers. Schematic diagram of the hierarchical organization of collagen fibrils in a single collagen fiber. The illustration demonstrates the spatial arrangement ofmineral patches, their shape, size, location and orientation on the collagen fibrils. (c) Mineral patches were observed to lie on the surfaces of collagen fibrils parallel andperpendicular to the long axes of collagen fibrils.

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(Fig. 3c) resolved the sharp surface features of the irregular mineralpatches on the surface of the collagen fibrils. Some micro-sizedmineral patches were observed to extend over adjoining group offibrils, as indicated by an arrow in the deflection image of Fig. 3c.Mineral patches on the surface of collagen fibrils were observedto be very thin (13–25 nm) and overlay the collagen fibrils, as re-vealed by the banded signatures of collagen fibrils underneaththe mineral patch, as shown in Fig. 3c.

A hierarchical organization of collagen fibrils and mineral crys-tals oriented along the collagen fibrils was also observed on thebone nodule surface, as shown in the deflection image in Fig. 4.Individual collagen fibrils were oriented parallel to each otherand to the long axes of the collagen fibrils. The mineral crystals ap-pear to be nearly flat (length 167 ± 19 nm, width 129 ± 6.0 nm) andwere oriented parallel to each other in each fibril and aligned alongthe long axes of the collagen fibrils. A schematic representationillustrates the spatial arrangement, shape, size, location, and orien-tation of mineral crystals along the long axes of the collagen fibrils,as observed in Fig. 4. Mineral crystals are demonstrated to be ori-ented parallel to each other in each fibril and aligned along the col-lagen fibril. In the schematic, mineral crystals are shown to beparallel to spatially spaced fibrils with an interfibrillar distance of�130 nm. The illustration has been drawn for the regions shownby a dotted box in the deflection image.

Hierarchical bone nodule structures were observed on the Chi–PgA–HAP fibrous scaffold after 16 days of culture, as shown inFig. 5a. Oriented collagen fibers are observed in the height image.Nanostructured mineral crystals appear to lie along the long axesof the collagen fibrils. Fig. 5b shows the cell proliferation resultsfor the Chi–PgA–HAP fibrous scaffold for cell culture durations of2, 4, 8, 12, and 16 days, respectively. The MTT assay results indicatethat the cell population was almost constant until day 8 of cell cul-ture. Rapid cell proliferation was observed after 12 and 16 days cellculture.

3.3. Nanoscale elastic properties of bone nodule

Fig. 6 shows the height image and corresponding modulus map(5 � 5 lm) of the surface of a bone nodule sample. The diagonal ar-rows in both images indicate the direction of the long axes of thecollagen fibrils. The height image (Fig. 6a) indicates an apparent ta-pered topography of an array of cylindrical collagen fibers alignedparallel to the long axes of the collagen fibers. The modulus mapimage (Fig. 6b) represents the elastic responses of the bone noduleconstituents, spatially distributed over an area of 5 � 5 lm.Modulus values were in the range 4–60 GPa. In Fig. 6 a lighter coloris related to the elastic responses of the soft collagenous phasewith a lower modulus while a darker color corresponds to stiffer

Fig. 4. (a) AFM deflection image indicating the individual collagen fibrils oriented parallel to each other. Flat mineral crystals are parallel to each other in each fibril andaligned along the long axis of the collagen fibrils. Mean repeat distance between the mineral crystals oriented along the individual fibrils is observed to be 74 nm. (b)Schematic representation illustrating the spatial arrangement, shape, size, location and orientation of mineral crystals along the sides of collagen fibrils enclosed in a singlecollagen fiber.

Fig. 5. (a) AFM height and corresponding deflection image of the surface of a bone nodule on a Chi–PgA–HAP fibrous scaffold after 16 days cell culture. Oriented collagenfibers are revealed in the height image. (b) Nanostructured mineral crystals appear to lie along the long axes of collagen fibrils. MTT assays indicate cell proliferation after 2, 4,8, and 16 days cell culture.

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mineral components with a higher elastic modulus. In betweengrayscale shades on the modulus map are indicative of collectiveelastic responses of the constituents of the bone nodule, mainlycollagen and mineral structures. Further, selected modulus vs. po-sition plots were used to separate the individual responses of the

constituents distributed spatially on the surface of a bone nodule.Plot 1 (4000–5000 nm regions, dotted arrows on modulus map)and plot 4 (3400–5000 nm regions, dotted arrows on modulusmap) indicate moduli in the ranges 6.4–12.7 and 5.3–10.4 GPa,respectively as also indicated by arrows in plots 1 and 4. These

Fig. 6. Height image (a), and corresponding modulus map image (b) of a 5 � 5 lm scan area on the surface of a bone nodule sample. Arrows shown in both the imagesindicate the direction of the long axes of the collagen fibrils. (c) Modulus map image showing the elastic responses of the bone nodule constituents, spatially distributed overan area of 5 � 5 lm. The lighter color is related to the elastic response of a soft collagenous phase with a lower modulus while the darker color corresponds to a stiffer mineralcomponent with a higher modulus. Mixed colors on the modulus map are related to the collective elastic responses of the constituents of the bone nodule, mainly collagenand mineral structures. Selected modulus vs. position plots (1–4) with dotted and solid arrows indicate the elastic responses of the major components of the bone nodule, i.e.collagen fibrils, collagen–mineral complexes and mineral lying parallel (Mk) and perpendicular (M\) to the long axes of the collagen fibrils.

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regions are represented by lighter colors on the modulus map andindicate the nanoscale elastic responses of collagen with a minimalinfluence of mineral. On these plots the lowest values of �4 GPaindicate the elastic property of pure collagen fibrils/molecules.Close assessments of both the height and modulus map images re-vealed the signatures of parallel arrays of collagen fibers and fibrils,oriented parallel to the long axes of the collagen fibers. Although,mineral deposits were not resolved in the height image, their elas-tic responses parallel and perpendicular to the long axes of the col-lagen fibers were distinctly marked by darker colored regions(higher modulus) along the diagonal bands in the modulus map

image. Plot 2 indicates the elastic properties of constituents spa-tially distributed along the sectional profile 2. Sectional profile 2in the modulus map represents the elastic responses of mineralcrystals in the parallel and perpendicular directions, as indicatedby the dotted arrows. Along the collagen fibril axes mineral depos-its (Mk) exhibited elastic moduli of 30–60 GPa, with most of thehigh modulus values centered around 30–40 GPa (plot 2). In addi-tion, evaluation of the elastic properties of mineral deposits on thesurface of the collagen fibrils oriented in the perpendicular direc-tion was also done. The surfaces of mineralized collagen fibrilswere resolved in the regions enclosed by stars in the modulus

Fig. 7. Micro-FTIR spectrum of a bone nodule sample in the region 2350–800 cm–1. The major bands for collagen are amide I, amide II and amide II and those of biologicalapatite m1, m3 PO4

3–, m3 HPO43–, and m2 CO3

2–, respectively.

Table 1FTIR band assignment for a nodule sample.

Band position(cm–1)

Band assignment

2315 NH vibrations of aminea

2027 NH3+ symmetric stretchinga

1805 Amides containing –CO–NH–CO–a

1678 Amide I1560 Amide II1470 Type A carbonate1415 m COO– and m3 CO3

2–, type AB carbonate1273 Amide III1188 m3 HPO4

3–

1127 m3 PO43–, non-stoichiometric HAP, newly precipitated

apatites1079 Weak m3 PO4

3–

976 m1 PO43–

873 m2 CO32–

a Bands from cell culture medium.

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map image, which represents the elastic responses of constituentson the surface of the collagen fibrils (10–60 GPa). Although individ-ual collagen fibrils were not resolved in the height image, thewidths of these regions were observed to be same as that of a sin-gle collagen fiber (Fig. 3b). Also, more than 50% of the area enclosedby stars in the modulus map are represented by darker colors inthe modulus map, indicating the presence of mineral crystals inseveral regions on the surfaces of the collagen fibrils. Elastic mod-uli on the surface of the collagen fibrils (2800–4000 nm regions onsectional profile 2) varied in the range 12.5–36 GPa, which isindicative of responses from primarily stiff mineral deposits(M\ = 20–40 GPa) and a composite response of collagen fibril–min-eral with a mean modulus of 15 GPa. Combined elastic responses(�15 GPa) from collagen and mineral constituents were observedin several regions of the modulus map, as represented by mixedcolors, mostly lying in the regions between the collagen fibers.These regions are selectively shown in sectional profiles 1, 3, and4 (solid arrows) in the modulus map image and have also beenmarked on plots 1, 3, and 4, respectively.

3.4. Chemical analysis of bone nodule

A FTIR spectrum of a bone nodule sample in the region 800–2350 cm–1 is shown in Fig. 7. Band assignments are shown inTable 1. The bands at 2315, 2027, and 1805 cm–1 are atrributedto N–H vibrations, NH3

+ symmetric stretching, and amides contain-ing –CO–NH–CO–, respectively. All the three bands originate fromthe cell culture medium. To confirm this fact, separate experimentswere carried out by soaking a clean gold substrate in cell culturemedium. All the bands in the bone nodule were compared with mi-cro-FTIR spectroscopy results reported in the literature for pediat-ric human bone [40]. The bands at 1678, 1560, and 1273 cm–1 wereassigned to amide I, amide II and amide III, respectively. In biolog-ical apatite carbonate groups substitute for PO4

3– (type A substitu-tion) or OH– (type B substitution) groups in HAP (Ca10(PO4)6(OH)2).The band at 1470 cm–1 can be assigned to type A carbonate and theband at 1415 cm–1 can be assigned to molecular vibrations of mCOO– and m3 CO3

2– type AB carbonate species. Phosphate bandsare observed in the region 900–1200 cm–1. Both inorganic ortho-phosphate ions, i.e. PO4

3– and HPO42– ions, have been reported to

occur in biological apatites of mineralized tissue [40]. The presenceof a band at 1188 cm–1 indicates the existence of HPO4

2– ions inapatite. The band at 1127 cm–1 has been previously observed inthe spectra of newly precipitated apatites, and is attributed to m3

PO43–, which is a non-stoichiometric HAP. A weak band at

1079 cm–1 can be attributed to molecular vibrations of m3 PO43–

ions. The band at 976 cm–1 can be assigned to the m1 vibrationmode of PO4

3– ions. The bands from 850 to 900 cm–1 are assignedto m2 CO3

2–; in the spectrum shown this band was observed at873 cm–1. Our microspectroscopic analysis indicates that all themajor bands of the bone nodule correspond to bands observed inhuman bone [40]. This is suggestive of the fact that bone nodulesgenerated on Chi–PgA–HAP nanocomposite films are chemicallysimilar to human bone.

4. Discussion

Previously we synthesized Chi–PgA–HAP nanocomposite filmsand scaffolds at solution pH values of 7.0 and 8.5 that resulted ina nano-colloidal morphology of the Chi–PgA material system[41]. The Chi–PgA–HAP nanocomposite films and scaffolds

Table 2Elastic properties of collagen.

Tissue type/experimental conditions Technique Elastic modulus (GPa) Reference

Bone nodule Modulus mapping �4–8 (Collagen) Present work�15 (Collagen-mineral) Present work20–60 (Mineral) Present work

Collagen Steered molecular dynamics 4.5–6.2 [43]Collagen of rat tail tendon (varying dehydration states) AFM indentation 3.75–11.5 [40]Collagen of bovine Achilles tendon (0.15 M NaCl solution) X-ray diffraction 2.9 ± 0.1 [42]Collagen-like peptide Simulation 4.8 ± 1.0 [44]Hydrated cross-link-free collagen fibril Simulation: atomistic and steered molecular dynamics 5–7 [45]

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synthesized at a solution pH of 4.5 resulted in a fibrous morphologyof the Chi–PgA material system (Fig. 1). Bone nodules were formedon both kinds of nanocomposites, i.e. on nano-colloidal [33] as wellas fibrous Chi–PgA–HAP nanocomposites, in the absence of anyosteogenic supplements. AFM imaging of bone nodules formed onthe fibrous Chi–PgA–HAP nanocomposite revealed a hierarchicalstructural organization of mineralized collagen fibers and fibrils(Figs. 3 and 4). The structural hierarchy, molecular structures. andnanomechanical properties of the bone nodules can be furtherdiscussed in the light of the existing literature.

4.1. Synthetic bone nodule mimics natural bone’s hierarchy

Nature has designed the complex and mechanically robust hier-archical structures of bone by repeating the smallest structuralbuilding blocks (i.e. collagen molecules, fibrils, and mineral crys-tals) several times to generate functional bone. In this work, we ob-served lower levels of hierarchical structures (collagen fibers,fibrils, and mineral crystals) of bone nodules formed on bothtwo-dimensional film substrates and 3D scaffolds. Mineral depos-its on the surface of collagen fibrils were first reported by Landisand co-workers in the case of calcifying leg tendons of turkeyand chicken bone using 3D electron microscopic tomography[24,42]. For the first time, we observed mineral crystals on the sur-face of collagen fibrils of in vitro generated bone nodules on an arti-ficial substrate, as revealed by high resolution AFM images (Figs. 3and 4). Crystals are deposited in a highly organized manner in thehole and overlap zones of collagen fibrils with characteristic repeatdistances of 67 nm [18,43]. As can be seen in Fig. 4, the mineralcrystals occur at almost equi-spaced distances along individual col-lagen fibrils with a repeat distance of 74 ± 7.1 nm, which very clo-sely matches the repeat distance of �67 nm between mineralcrystals, reported in the case of trabecular bone [44]. This repeatdistance between the mineral crystals corresponds to the distanceby which adjacent collagen molecules are staggered. In humandentin, the collagen fibrils are randomly oriented and the diame-ters of hydrated and dehydrated collagen fibrils have been foundto be in the range 75–105 nm [45]. In contrast, collagen fibrils inbone nodules possess a high degree of structural organizationand are oriented perfectly parallel to the long axes of collagen fi-bers with a diameter of � 300 nm. Mineral crystals in the bonenodule are observed to be larger in size, which suggests that min-eral crystals are not only confined to 40 nm hole zones. These min-eral crystals are believed to nucleate initially in gap zones andgrow in length along their crystallographic c-axis parallel to thecollagen fibril axes and extend to the surface of the collagen fibrilsand extrafibrillar spaces by fusion of smaller crystals into largerones, as suggested by Landis et al. [23,46]. This suggests that theformation of mineral crystals is site dependent and occurs as a re-sult of independent nucleation events in local zones of collagen, asalso observed by Landis et al. [23,42,47]. Recently, mineralizedbone nodules generated by osteoblasts were characterized byTEM, but no collagen–mineral hierarchical organization was ob-

served [48]. The major conclusion from the AFM structural analysisis that synthetic bone nodules mimic the hierarchy of natural bone.

4.2. Nanoscale elastic modulus of collagen and mineral of bone nodules

In human bone, the basic building blocks are designed at thenanoscale, with hard nanosized mineral crystals embedded in asoft collagenous matrix. Evaluation of the nanoscale mechanicalproperties of these structural elements generated by human oste-oblasts on synthetic biomaterials can serve as guidelines for scaf-fold design and development. The elastic moduli of collagen andmineral of the bone nodules obtained in the present work are inclose agreement with experimental and simulations results re-ported in the literature for various collagenous and calcifying tis-sues of different animal species and human bone [49–51](Table 2). In our results, the maximum modulus value of �60GPa, with a mean value of 35 GPa, was measured, which appearsto be a nanoscale mechanical property of mineral crystals at extre-mely shallow penetration depths of 2–3 nm. The minimum elasticmodulus of 4–8 GPa for collagen corresponds well with that re-ported in the case of rat tail tendon (E = 3.75–11.5 GPa, for a pen-etration depth of 2–5 nm) [50] and human dentin containinginterfibrilar mineral (E = 2GPa) [52] using AFM and nanoindenta-tion. Our results are also in close agreement with X-ray diffractionresults for bovine Achilles tendon (E = 2.9 ± 0.1 GPa) [53]. Steeredmolecular dynamics (SMD) simulations have been conducted oncollagen by our group [54–56]. The simulations results also indi-cate that the elastic modulus of collagen is in the range 4.5–6.2GPa. The elastic modulus of collagen is dependent on the rate ofpulling, stiffness of the spring used in pulling and the length ofthe molecule. The elastic moduli of collagen (5–7 GPa) reportedin the literature using SMD simulations [57,58] are in close agree-ment with the elastic moduli obtained from our experimental andSMD simulation results.

4.3. FTIR spectra of bone nodules

Tissue engineered bone should be chemically similar to humanbone to regulate bone physiology and mechanical functions duringthe healing or repair of tissue. For instance, it has been reported thatin carbonated or biological apatites carbonate ion substitution forphosphate ions alter the crystal shape and arrangement in thematuring bone, leading to altered mechanical properties [59]. OurFTIR analysis indicates that the developing bone nodule is chemi-cally similar to human bone [40], as revealed by the presence of col-lagen (amide I, amide II and amide III bands) and carbonated HAP(Fig. 7). Evidence of three major bands i.e. m3 HPO4

3– (1188 cm–1),m3 PO4

3– (1127 cm–1) and m2 CO32– (873 cm–1), confirmed the pres-

ence of immature biological apatite in bone nodules. Biological apa-tites are both chemically and structurally different from thesynthetic nanohydroxyapatite used in the preparation of the Chi–PgA–HAP nanocomposites [39,60], as revealed by the distinctivesizes, shapes, and chemistries of the as-synthesized HAP and

1182 R. Khanna et al. / Acta Biomaterialia 7 (2011) 1173–1183

mineral deposits of bone nodules. The differences in physico-chemical characteristics can arise due to the different physiologicalenvironments in which they are generated. It suggests that theenvironment essentially mediates the structural and biologicalfunctions of functional tissues.

5. Conclusions

1. The prepared Chi–PgA–HAP fibrous nanocomposites are osteo-conductive and osteoinductive and thereby provide an appro-priate microenvironment for cell organization and bonenodule formation.

2. The bone nodules formed on the biomaterial systems describedin this work mimic natural bone’s hierarchy in the light of fol-lowing results.a) High resolution AFM imaging revealed hierarchical struc-

tures of a bone nodule, consisting of an array of repeatingcollagen fibers (�1.7 lm width) and collagen fibrils(�300 nm width) aligned along the long axes of collagenfibers. Mineral deposits are observed to lie on the surfacesof collagen fibers and fibrils along the long axes of the col-lagen fibers, and are reported for the first time in anin vitro generated bone nodule. Structural organization ofthe bone nodule constituents was found to be similar to thatof calcifying tissues, which supports the conclusion thatsynthetic bone nodules mimic the natural bone hierarchy.

b) The elastic properties of mineral deposits parallel and per-pendicular to the long axes of collagen fibrils is reportedfor the first time using a modulus mapping technique. Thenanoscale elastic properties of collagen fibrils and mineralcrystals closely match the experimental and simulationresults for bone reported in the literature, which supportsthe conclusion that bone nodules mimic the nanoscalemechanical properties of natural bone.

c) FTIR analysis indicates the presence of collagen and biolog-ical apatite in the bone nodules, which supports the conclu-sion that bone nodules are chemically similar to humanbone and possess the characteristics of newly precipitatedor immature bone.

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