International Journal of Polymer Science

International Journal of Polymer Science / 2010 / Article
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Polymeric Biomaterials for Tissue Engineering Applications

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Research Article | Open Access

Volume 2010 |Article ID 175264 |

Rohit Khanna, Kalpana S. Katti, Dinesh R. Katti, "In Situ Swelling Behavior of Chitosan-Polygalacturonic Acid/Hydroxyapatite Nanocomposites in Cell Culture Media", International Journal of Polymer Science, vol. 2010, Article ID 175264, 12 pages, 2010.

In Situ Swelling Behavior of Chitosan-Polygalacturonic Acid/Hydroxyapatite Nanocomposites in Cell Culture Media

Academic Editor: Shanfeng Wang
Received12 Feb 2010
Revised22 Mar 2010
Accepted22 Mar 2010
Published03 Jun 2010


The molecular and mechanical characteristics of in situ degradation behavior of chitosan-polygalacturonic acid/hydroxyapatite (Chi-PgA-HAP) nanocomposite films is investigated using Fourier Transform Infrared spectroscopy (FTIR), Atomic Force Microscopy (AFM), and modulus mapping techniques for up to 48 days of soaking in cell culture media. The surface molecular structure of media-soaked samples changes over the course of 48 days of soaking, as indicated by significant changes in phosphate vibrations (1200–900  ) indicating apatite formation. Chitosan-Polygalacturonic acid polyelectrolyte complexes (PECs) govern structural integrity of Chi-PgA-HAP nanocomposites and FTIR spectra indicate that PECs remain intact until 48 days of soaking. In situ AFM experiments on media-soaked samples indicate that soaking results in a change in topography and swelling proceeds differently at the initial soaking periods of about 8 days than for longer soaking. In situ modulus mapping experiments are done on soaked samples by probing 1–3 nm of surface indicating elastic moduli of 4 GPa resulting from proteins adsorbed on Chi-PgA-HAP nanocomposites. The elastic modulus decreases by 2 GPa over a long exposure to cell culture media (48 days). Thus, as water enters the Chi-PgA-HAP sample, surface molecular interactions in Chi-PgA-HAP structure occur that result in swelling, causing small changes in nanoscale mechanical properties.

1. Introduction

Recent developments in design of novel polymeric biomaterials have allowed researchers to confront many of the challenges dealing with the design of novel biomaterials. The tremendous potential of biodegradable polymers for tissue engineering and medical devices results from their biocompatibility, ease in processing, and capability of controlled degradation in response to biological environment. Candidate materials for bone tissueengineering include natural polymers (chitosan, collagen, hyaluronan, fibrin), synthetic polymers (polycaprolactone, polylactic acid, polyglycolic acid, and their copolymers) and inorganic materials (tricalcium phosphate, hydroxyapatite). Also, the natural biodegradable and biofunctional biopolymer, chitosan, has been considered as a potential candidate material for numerous biomedical applications including controlled drug release [1, 2], wound dressing [3], and more recently, for tissue engineering application [413]. Chitosan [ (1,4)-linked 2-amino-2-deoxy-D glucan] is a cationic polysaccharide obtained by N-acetylation of chitin. Chitosan provides improved cell attachment, since the polysaccharide backbone of chitosan is structurally similar to glycoaminoglycans, the major component of ECM of cartilage and bone [14]. Due to cationic character of chitosan, it interacts with a number of polyanions and has been used in preparation of various polyelectrolyte complexes with natural polyanions such as alginic acid, dextran sulphate, heparin, pectin, and xanthan [15, 16]. Another natural biodegradable biopolymer, polygalacturonic acid (PgA), is a water-soluble polyanionic polysaccharide, consisting of a linear chain of D-galacturonic acid units with (1-4) glycosidic linkages. This is a principal component of pectin that exists in cellular walls of plants. Pectin is extensively used in food industry as a gelling agent and widely used in the production of jams and jellies, confectionary products and for stabilization of yogurts [17] and drug production such as for antidiarrheic, detoxicants and as protectors of gastrointestinal tract [18, 19].

Appropriate mechanical property of bone tissue engineering scaffold is one of the key factors in selection and design of biomaterials, which may limit the independent use of natural chitosan/pectin-based materials. Chitosan, has been reported to possess improved biocompatibility, mechanical strength, hard tissue regeneration and tailored degradation kinetics by incorporation of hydroxyapatite (HAP) [20, 21] and gelatin [20, 22, 23], alginate [24], and so forth. By employing electrostatically complementary nature of chitosan and PgA and by adopting a biomimetic synthesis route, we designed and developed novel chitosan-PgA-hydroxyapatite (Chi-PgA-HAP) nanocomposites [2527] which exhibited a substantial improvement in mechanical properties. Our FTIR results demonstrated that strong interfacial interactions in Chi-PgA-HAP structures are believed to be the key factors in mechanical property improvement [2527]. Biocompatibility results demonstrated that human osteoblasts (CRL 11732) generated mineralized bone nodules on Chi-PgA-HAP fibrous nanocomposites without using any differentiating media suggesting that Chi-PgA-HAP substrates are osteoinductive and provide an appropriate microenvironment for cell organization and tissue regeneration [27].

For suitability of Chi-PgA-HAP nanocomposites for bone tissue engineering, their degradation behavior needs to be evaluated. Ideally, material degradation rate should match the rate of tissue regeneration, while generating nontoxic degradation products. Both chitosan and pectin are well known to be enzymatically degradable and water resistant [18, 19, 28, 29]. Several investigations on enzymatic degradation of chitosan (with varying degree of deacetylation) for human use have been carried out in lysozyme [3034], since it is found in human body fluid, including serum (concentration 4–13 mg/L) and tears ( 450–1230 mg/L) [35]. Chitosan-pectin membranes have been reported to swell considerably in water and their swelling characteristics depend on pH, temperature, ionic strength, and degree of polyelectrolyte complex formation [18, 29, 36]. Swelling behavior of chitosan-pectin membranes is an attractive property for drug delivery systems [37, 38]. In situ swelling behavior of Chitosan-PgA-HAP nanocomposites in terms of change in surface properties has not been investigated. Here, we present the in situ swelling behavior of Chi-PgA-HAP nanocomposite films over the soaking period of 48-day in cell culture media. In situ swelling experiments include the evaluation of topography, chemistry and nanomechanical properties of soaked 2D substrates in a fully hydrated state. 2D substrates have been utilized due to their relative ease as compared to 3D scaffolds. In situ AFM experiments were performed in cell culture media at a temperature of 37°C, to obtain the topography of soaked samples. Modulus mapping technique has been recently used to obtain the surface elastic properties of dentin-enamel junction [39] and biomimetic nanohydroxyapatite particles [40]. However, this technique has not been used till now to evaluate the purely elastic properties of constituents of soaked samples under fully immersed conditions. In the present work, in situ modulus mapping experiments were designed and elastic properties of soaked samples were obtained in fully submerged conditions (cell culture media; 37°C) by applying extremely shallow displacements (~2-3 nm). Molecular structure of both dry and soaked specimen was investigated by Fourier Transform Infrared spectroscopy.

2. Materials and Methods

2.1. Film Preparation

HAP nanoparticles were prepared by wet precipitation route from sodium phosphate (Na2HPO4; J. T. Baker), and calcium chloride (CaCl2; EM Sciences) by maintaining pH at 7.44. The detailed processing procedures have been discussed elsewhere [41]. Chitosan (MW 190,000, >85% deacetylation) and polygalacturonic acid (MW 25000) were obtained from Sigma-Aldrich chemicals. Chitosan and PgA solutions were prepared in deionized water at a concentration of 1 g/100 mL. The pH of chitosan solution was adjusted by adding acetic acid and that of PgA solution was adjusted by adding NaOH. The pH of resulting solutions was adjusted to 4.5. Chitosan-PgA solution was prepared by adding chitosan solution dropwise to PgA solution followed by sonication for 3 minutes. For synthesis of Chi-PgA-HAP nanocomposite, HAP nanoparticles (1 g) were suspended in 100 mL of deionized water and sonicated for 1 hour to obtain a uniform dispersion of nanoparticles. Subsequently, sonicated HAP nanoparticles were mixed with sonicated Chitosan-PgA solution and further sonicated for 90 seconds to obtain a homogeneous solution. Films of two compositions were prepared (Chi-PgA-10% HAP and Chi-PgA-20% HAP) on TCPS (Tissue Culture Polystyrene) petridishes by solvent evaporation at room temperature under clear-air laboratory environment. For making films, Chi-PgA-HAP solution was diluted to 1 : 10 ratio with deionized water and 3 mL of resulting solution was air-dried on tissue culture polystyrene petridishes. Both the nanocomposite substrates were found to be biocompatible and Chi-PgA-20% HAP composition was selected for soaking study.

2.2. Soaking Experiments

All the samples were UV sterilized and subsequently soaked in 5 mL of cell culture media for 1, 2, 8, 24, and 48-day, respectively, under standard incubator conditions (37°C, 5% CO2). Cell culture media was prepared by adding DMEM (Dulbecco’s modified eagle medium; Hyclone), supplemented with 2.5 mM l-glutamine (without phenol red), 10% FBS (fetal bovine serum; ATCC) and 1% antibiotic (G418; J R Scientific Inc.). At the end of predetermined time, films samples were taken out of incubator, and washed twice with PBS and immediately taken for the AFM/FTIR/nanoindentation analyses. Culture media were replaced after every 3 days.

2.3. FTIR Experiments

FTIR experiments were carried out using a Nicolet 850 FT-IR spectrometer with a KBr beam splitter using reflectance accessory (45° angle of incidence). Spectra were collected in the range of 4000–500 cm-1 at a spectral resolution of 4 cm-1. Film samples of chitosan, PgA and Chi-PgA-HAP nanocomposites were prepared on the clean gold substrates and allowed to dry overnight. FTIR spectra were obtained on soaked samples which were air-dried before data acquisition. A gold-coated metal plate was used to collect the background spectrum.

2.4. In Situ AFM Experiments Using Multimode Heater

Topography of dry and soaked samples was characterized by Atomic Force Microscope (multimode AFM: Veeco Metrology Group, Santa Barbara, CA), equipped with a Nanoscope IIIa controller and J-type piezo scanner. The equipment has -axis resolution of around 0.5 A°. Silicon nitride cantilever probes (Model: NP-S20) with pyramidal tips of ~20 nm radius of curvature and a nominal stiffness of 0.06 N/m were used for imaging under contact mode in clean-air laboratory environment with 40%–50% relative humidity. For performing in situ AFM experiments in fully immersed conditions, Multimode low temperate heater from Veeco Metrology (Santa Barbara, CA) was used. It enabled in situ AFM analyses of soaked samples in a controlled liquid environment. Major components of the system include a fluid cell, a heating element, Thermal Applications Controller (TAC), and spacer block. For a detailed description of components available for Multimode Scanning Probe Microscopes, AFM technical support note 392 can be referred. The sample temperature was set and maintained at 37°C by regulating the heating element and tip heater voltage with the help of Thermal Applications Controller (TAC). The whole sample-fluid-tip was thermally equilibrated for at least 20 minutes to enable the tests to be performed in a stable and controlled environment (cell culture media; 37°C).

2.5. In Situ Modulus Mapping Technique

A Hysitron triboscope nanomechanical instrument (Minneapolis, MN) equipped with nanoscope IIIa controller (Veeco Metrology, Santa Barbara, CA) was used to test the dry and soaked samples with a Berkovich (three-sided pyramid; a 100–200 nm tip radius) diamond indenter fluid tip. The fluid tip with a longer steel shaft allows a liquid level up to 3-4 mm to be maintained above the sample surface and also ensures the machine calibration in the liquid. In order to avoid any meniscus forces, the tip was fully submerged in the liquid prior to testing. In situ modulus mapping experiments were performed on soaked samples in cell culture fluid and at a temperature of 37°C to simulate incubator conditions. Modulus mapping is a surface nanomechanical characterization technique that enables the measurement of purely elastic properties of material surface by applying extremely shallow displacements (~2-3 nm). Detailed description of principles of modulus mapping technique, its formulation, and application can be found elsewhere [39, 42]. In force modulation mode, a quasistatic force of 3  N was superimposed by a 2  N sinusoidal force at a frequency of 200 Hz. A total of 256 256 tests were done to obtain a modulus map in only 10–15 minutes.

For testing in liquid environment, a special fluid cell was designed by using O-rings (9/ O.D. 3/ 3/ ) glued onto 15 mm steel discs. Samples (0.5 cm 0.5 cm) were glued and kept inside the fluid cell and then, flushed with liquid level up to 3-4 mm for testing in fully submerged conditions. The whole tip-sample-fluid assembly was then heated and maintained to 37°C using Multimode heater as described in Section 2.4. Before data acquisition, the entire setup was allowed to equilibrate for at least 20 minutes. Before starting modulus mapping experiment, the tip was wiped with acetone soaked tissue wipes to remove any ions/salts which may have adsorbed from the previous measurements. Tip calibration was done using a standard quartz sample with known elastic modulus. Extreme care was taken to keep the samples under the wet conditions before the in situ testing to avoid the effect of drying on nanomechanical measurements.

All the AFM, modulus mapping and FTIR experiments were performed using on least 3 samples for each set of test conditions. All the tests were repeated at least twice to account for repeatability and reproducibility of test measurements.

3. Results and Discussion

3.1. Molecular Nature of Surfaces of Soaked Chi-PgA-HAP Nanocomposite Samples

FTIR spectroscopy was used in a reflection mode to characterize the surface molecular structure of soaked samples. Figure 1 shows the FTIR spectra of dry and soaked Chi-PgA-HAP samples within the range 4000–500 cm-1. The band positions and their assignments have been summarized in Tables 1, 2, and 3. The band at 1666 cm-1, as seen in Figure 1, is a characteristic vibration of N-acetylated chitin and is assigned to Amide I band. It can also overlap with Amide I band from carbonyl stretching in protein structure, as proteins from cell culture media can get adsorbed onto surface of Chi-PgA-HAP samples. The band at 1622 cm-1 is attributed to asymmetric stretching vibration of dissociated carboxylate (COO-) [26, 43]. This band shifts slightly to lower wavenumber, to 1605 cm-1 in soaked samples, which indicates that the change in environment of Chi-PgA-HAP structure through its interaction with water, possibly through hydrogen bonds with Chi-PgA-HAP structure. As water enters the Chi-PgA-HAP sample which contains polyelectrolyte complexes with –NH3+, –COO- ions, surface structure of Chi-PgA-HAP structure can get modified. In such a case, swelling can occur due to mutual repulsion between positively/negatively charged species along with interactions of these charges species with water. Swelling characteristics of chitosan-pectin have been reported to be modified by preparing the samples at varying pH [44].

Band assignmentBand position as per the testing conditions
Dry1 day2 days8 days24-day48-day

Asymmetric N–H stretching338633773365336533683360
symmetric N–H stretching3290328432923288not observednot observed
Asymmetric C–H stretching293129362933293229342933
Symmetric C–H stretching287828782874287828662869

Band assignmentBand position as per the soaking conditions
Dry1 day2 days8 days24-day48-day

Amide I16661665166616651665
Asymmetric COO- stretching162216051624160416051603
NH3+ deformation153415321534153315341539
symmetric COO- stretching141414171418142014161421
C–H bending vibration of ring133013311329132813321336

Band assignmentBand position as per the soaking conditions (cm-1)
Dry1 day2 days8 days24-day48-day

C–O stretching in (C–O–C)115511501154115111491152
asymmetric PO stretching1113, 10701107, 10611110, 106711021107, 1072, 10431110, 1077, 1049
symmetric PO stretching1009, 954996, 9531001, 9521021, 9561009, 9561010, 960
P–O(H) stretching894895894895896896
O–H bending in HAP structure645645645644643645
O–P–O bending569569568566563564

PgA-HAP interactions were also affected as a result of soaking, as revealed in 1100–1000 cm-1 region of FTIR spectra of soaked samples (Figure 2). PgA interacts with HAP through dissociated carboxylate groups. As carboxylate groups carry negative charge, calcium ions in HAP can act as binding sites for PgA-HAP complexes [26]. To investigate PgA-HAP interactions, phosphate regions were carefully observed for both dry and soaked samples, as shown in Figure 2(a–h). Figure 2(a, b) shows the FTIR spectrum obtained from dry chitosan and dry PgA film, respectively. Bands observed at 1086 cm-1, 1055 cm-1, and 1032 cm-1 in PgA (Figure 2(b)), originate from skeletal vibrations involving C–O stretching, which are characteristic of saccharide structure of PgA [18]. Comparing FTIR spectrum of PgA (Figure 2(b)) with that of dry Chi-PgA-HAP (Figure 2(c)), bands originating from –C–O–C– glycosidic linkages overlap with bands from phosphate regions and occur as a broad band (1100–1000 cm-1) in Figure 2(c). Some of the characteristic bands of HAP structure, that is, asymmetric PO stretching vibrations in HAP structure occur at 1113 cm-1, 1070 cm-1, and 1009 cm-1 regions can be clearly observed in dry Chi-PgA-HAP samples. Significant changes in band positions in phosphate regions (1200–900 cm-1) were not observed throughout the soaking duration. However, significant changes in shape of bands in phosphate regions (1200–900 cm-1) in soaked samples, over a soaking duration of 48-day indicate that interactions in PgA-HAP structure changes as a result of soaking. Comparing spectrum of PgA (Figure 2(b)), dry Chi-PgA-HAP (Figure 2(c)), and pure HAP [45] with that of 48-day soaked sample (Figure 2(h)), characteristic shape of phosphate regions observed on 48 soaked sample matched with that observed in HAP structure. It appears, that as soaking proceeds, more HAP nanoparticles may get exposed onto the surface of soaked samples. Similar observations of exposed HAP particles were made in AFM topography image of 48-day soaked samples, as discussed in the next section. There can be another possibility of apatitic growth on soaked film samples, which accounted for the visibility of characteristic bands of HAP structure on 48-day soaked Chi-PgA-HAP sample.

Overall, FTIR spectroscopy results on soaked samples indicate that the degradation of Chi-PgA-HAP nanocomposite films was not severe in cell culture medium until 48-day of soaking. There were apparent changes in surface chemistry of soaked Chi-PgA-HAP substrates as compared to dry substrates. Severe degradation can be expected on dissociation of Chi-PgA polyelectrolyte complexes (PECs) and can also be due to breaking of –C–O–C– glycosidic linkages in Chitosan/PgA structure. Spectral changes in both Chi-PgA-HAP complexes and –C–O–C– glycosidic linkages were not observed in soaked samples. It can be due to the fact that both chitosan and PgA have been reported to be enzymatically degradable [19, 30, 31] and degrade primarily through lysozyme-mediated hydrolysis. Moreover, fully deacetylated chitosan has low degradation rates due to inability of hydrolytic enzymes to penetrate the crystalline microstructures. Slow degradation characteristics in our study can be due to high deacetylation degree of chitosan (>85%) and maintenance of structural integrity of PECs in cell culture media.

3.2. AFM Topographic Characterization

Topographical signatures of swelling/degradation were investigated by carrying out systematic in situ AFM experiments in hydrated environment using the Multimode heater, as described in Section 2.4. Representative AFM topographic images of dry and soaked film samples are shown in Figure 3. Topography of soaked samples was investigated for 1, 2, 8, 24 and 48-day, respectively. For investigating the major changes in substrate topography at the microscale, large scan sizes (100  m 100  m) were obtained on both dry and soaked regions. Figure 3(a) shows a smooth 3D topography of dry Chi-PgA-HAP nanocomposite. Further, significant changes in topography were observed after soaking in cell culture medium, as indicated in Figures 3(b)3(f). After 1 day of soaking, a rough corrugated appearance as indicated by the uneven distribution of sharp ridges and valleys in the microstructure (Figure 3(b)) is observed. Figure 3(c) shows the 3D topography after 2 days of soaking. Ridges appear to grow in height and were observed to have nonuniform distribution over the entire scan area. After 8 days of soaking, some of sharp ridges appear to decrease in height or flatten out and more valleys or grooves were observed (Figure 3(d)). The ridges appear to flatten out and take the shape of circular features (bright spots), were observed to be unevenly distributed over the microstructure.

Soaking experiments were carried out for longer periods to observe the topographical changes of Chi-PgA-HAP samples over a long term exposure. Figure 3(e) indicates the 3D topography of 24-day soaked substrate. As seen in AFM topographic image, deep grooves and pits appear to be present over the entire area. In the literature, swelling characteristics of chitosan and pectin based materials have been commonly reported in terms of the water absorption and water retention [18, 28, 30, 32, 46]. Our AFM results demonstrate that topography changes dramatically upon soaking in cell culture media (1 day) which can be due to swelling of Chi-PgA-HAP biodegradable polymer nanocomposite. Irregularities in topographic features indicate that swelling profiles are site-specific, depending on the access of water into the polymer structures. Due to hydrophilic nature of Chi-PgA-HAP nanocomposite, diffusion of water into the matrix of nanocomposite is faster than degradation, which can result in swelling of the Chi-PgA-HAP nanocomposite prior to degradation. Swelling appears to proceed in a nonuniform fashion during the initial soaking period up to 8 days as indicated by dramatic changes in ridges and valleys in AFM topography images. Sizes and shapes of circular features also appear to be distorted and the irregular shaped larger pits appear to be formed by the coalescence of smaller pits, as revealed by more diffused patterns of light and dark regions in the AFM topographic image of 48-day soaked sample (Figure 3(f)).

The topographic changes can occur due to change in molecular interactions as demonstrated in FTIR spectra of soaked samples (Figure 1). Selected 2D AFM topographic images (100  m 100  m) were taken for dry (Figure 4(a)), 8 days (Figure 4(b)) and 48-day (Figure 4(c)) soaked nanocomposite samples to correlate the topographic observations with the chemistry of soaked samples. As seen in Figure 4(a), bundles of Chi-PgA fibers were observed and HAP particles were barely visible on the surface of dry sample. After 8 days of soaking in cell culture media, HAP nanoparticles were observed on the surface of soaked sample, as seen in AFM topographic image (Figure 4(b)). Figure 4(c) indicates the AFM topography image of 48-day soaked Chi-PgA-HAP sample. It appears that ~80% of the scan area is covered with HAP nanoparticles and FTIR spectrum taken on this sample exhibited the characteristic shape of phosphate bands which match closely with that observed in case of pure HAP samples [45]. This suggests that as soaking proceeds, molecular interactions in Chi-PgA-HAP structures change (phosphate regions; Figure 2), thereby, exposing more HAP nanoparticles to surface of soaked samples and these observations were not made in dry Chi-PgA-HAP samples (both AFM images & FTIR results).

3.3. Nanoscale Elastic Properties by In Situ Modulus Mapping Technique

In situ elastic properties of soaked samples were obtained in presence of cell culture media and at a temperature of 37°C (using Multimode heater) using modulus mapping technique. The description of test procedures is given in Section 2.4. Figure 5(a) shows the modulus map of a scan area of 10  m 10  m on surface of dry Chi-PgA-HAP nanocomposite film. In modulus map image, lighter color corresponds to lower elastic moduli (~4 GPa) due to relatively softer polymer phase, that is, Chi-PgA phase. Darker color corresponds to higher elastic moduli (~80 GPa), which indicate the elastic responses from the stiffer hydroxyapatite phase or from the regions having a very thin polymer layer. A combined elastic response (~10–30 GPa) from both the polymer and HAP phase is indicated by mixed color patches observed in the modulus map image. The composite elastic modulus can be obtained from the surfaces of biomimetic nanohydroxyapatite, as reported in our prior work [40]. The variation in modulus values along the scan length of 10 m indicates the local distribution of constituents in the microstructure (Figure 5(b)).

Figures 6(a)6(d) show the modulus data of samples soaked in cell culture media for 1, 2, 8, and 48-day, respectively. Representative line scans were selected from modulus maps obtained as per each soaking experiment conditions, and modulus values were plotted as function of position. In situ elastic moduli of 3.5–8 GPa have been obtained from the surfaces of soaked substrates for upto 2 days of soaking. These are the nanoscale elastic responses of the swollen phases or constituents of Chi-PgA-HAP substrate. An observed scatter in modulus values within 4–8 GPa is consistently observed during the initial soaking duration (Figures 6(a) and 6(b)). Over the longer soaking duration (Figures 6(c) and 6(d)), less scatter in moduli was observed and moreover, nanoscale modulus is observed to decrease by ~2 GPa.

Elastic modulus in the range of 2–4 GPa [47] has been measured from the gold nanoparticles functionalized with bovine serum albumin and streptavidin using nanoindentation technique (load = 5  N). The elastic modulus of ~3-4 GPa measured using modulus mapping technique (load = 5  N) can result from the nanoscale elastic response from the proteins adsorbed on the surface of soaked samples, which also match closely with that reported in literature. Protein adsorption on Chi-PgA-HAP nanocomposites is more likely due to its hydrophilic nature. Specifically, chitosan’s cationic nature allows various electrostatic interactions with negatively charged species such as proteins, anionic glycoaminoglycans [4]. Chi-PgA-HAP structure with mainly –NH2, –COOH and –(COO-NH3+) complexes can interact with proteins and modify the surfaces. Performing in situ elastic property measurement in cell culture fluid and at 37°C can give the nanoscale properties of proteins adsorbed on Chi-PgA-HAP substrates in their natural environment. However, in a modulus mapping technique, it is not easy to identify the elastic responses of individual sites, due to a potential overlap between the modulus values of proteins and sites of soaked substrate.

Further, additional experiments were done on soaked but air dried samples in clear air laboratory environment to see the effect of dehydration from the soaked polymer nanocomposite samples. Selected results are presented herein for modulus mapping experiments on 24-day soaked samples and tested in a fully hydrated state and unhydrated (dried) environment. Figure 7(a) indicates the modulus map obtained after 24-day of soaking and tested in a hydrated state (cell culture media; 37°C). Modulus values of 4–6 GPa were observed with a mean value of ~5 GPa (Figure 7(b)). Figure 8(a) indicates the modulus map obtained after 24-day of soaking, and completely dried in air, and then, tested in clean-air laboratory environment. Modulus data were plotted as a function of position over a length of 10  m (Figure 8(b)). Modulus values in the range of ~4–80 GPa were observed. Significantly large difference in modulus values observed by testing under the hydrated and unhydrated environment can result from dehydration effects from the surface of the swollen polymer specimen which makes its surface more stiff. Effect of sample dehydration on nanomechanical properties is less important for hard materials like metals, ceramics and some hard tissues. However, soft and spongy polymers (e.g., Chi-PgA-HAP biodegradable polymer nanocomposites) which exhibit swelling characteristics, have entrapped water in their structures. Hence, testing such samples in a dry environment can severely affect the mechanical properties, as seen in our modulus mapping results. It has been shown that even in case of hard tissue like bone, upon dehydration, hardness can be increased by 17%–30% and modulus by 15%–50% [48, 49]. Therefore, it is important that in situ nanomechanical mesurements are carried out in a liquid environment to get in situ elastic properties of water-swollen polymeric structures. Our results suggest that modulus mapping is a promising technique to evaluate the purely elastic properties of proteins and that of soaked substrates.

4. Conclusions

The in situ degradation/swelling behavior of novel Chi-PgA-HAP nanocomposites is characterized by FTIR, AFM and modulus mapping technique. FTIR results demonstrated that molecular structure of soaked samples changes over the course of 48-day of soaking in cell culture media, as indicated by significant variations in phosphate vibrations. As water enters the Chi-PgA-HAP sample, interactions in Chi-PgA-HAP structure are modified that can result in swelling. PECs govern the structural integrity in Chi-PgA-HAP nanocomposites and FTIR spectra indicated that PECs were still in-tact until 48-day of soaking. In situ AFM experiments on soaked samples indicate that soaking results in a change in topography due to swelling of Chi-PgA-HAP substrates. AFM topographic images indicate that swelling proceeds in an inhomogeneous fashion during the initial soaking period up to 8 days and then, occurs in a more gradual fashion during the longer soaking periods. In situ modulus mapping experiments on soaked samples represent the elastic properties of surface of soaked samples. Elastic moduli of ~4 GPa are observed, which may result from nanoscale elastic responses of proteins adsorbed on Chi-PgA-HAP nanocomposites. Modulus mapping results of soaked samples indicate that nanoscale elastic modulus decreases by ~2 GPa over a long exposure to cell culture media (48-day) and there were no major changes in surface mechanical properties over 48-day of soaking in cell culture media.

This work has important implications for tissue engineering. For instance, mostly biodegradable materials are used in the fabrication of tissue engineering scaffolds. Biodegradable materials can undergo physic chemical-mechanical changes under the wet conditions, as also indicated by our study. It is now understood that cells do respond to such physico-chemical changes which can also affect cell-biomaterial interactions. In order to correlate the changes in physical properties of degrading substrate to cellular responses (biochemical/biomechanical), a systematic evaluation of in situ material properties of biodegradable materials in nearly physiological environment is required, which merits scientific interest.


  1. S. Miyazaki, K. Ishii, and T. Nadai, “The use of chitin and chitosan as drug carriers,” Chemical & Pharmaceutical Bulletin, vol. 29, no. 10, pp. 3067–3069, 1981. View at: Google Scholar
  2. K. Aiedeh, E. Gianasi, I. Orienti, and V. Zecchi, “Chitosan microcapsules as controlled release systems for insulin,” Journal of Microencapsulation, vol. 14, no. 5, pp. 567–576, 1997. View at: Google Scholar
  3. G. Kratz, C. Arnander, J. Swedenborg et al., “Heparin-chitosan complexes stimulate wound healing in human skin,” Scandinavian Journal of Plastic and Reconstructive Surgery and Hand Surgery, vol. 31, no. 2, pp. 119–123, 1997. View at: Google Scholar
  4. S. V. Madihally and H. W. T. Matthew, “Porous chitosan scaffolds for tissue engineering,” Biomaterials, vol. 20, no. 12, pp. 1133–1142, 1999. View at: Publisher Site | Google Scholar
  5. W.-C. Hsieh, C.-P. Chang, and S.-M. Lin, “Morphology and characterization of 3D micro-porous structured chitosan scaffolds for tissue engineering,” Colloids and Surfaces B, vol. 57, no. 2, pp. 250–255, 2007. View at: Publisher Site | Google Scholar
  6. Y. Wan, H. Wu, and D. Wen, “Porous-conductive chitosan scaffolds for tissue engineering, 1: preparation and characterization,” Macromolecular Bioscience, vol. 4, no. 9, pp. 882–890, 2004. View at: Publisher Site | Google Scholar
  7. J. Nakamatsu, F. G. Torres, O. P. Troncoso, M. L. Yuan, and A. R. Boccaccini, “Processing and characterization of porous structures from chitosan and starch for tissue engineering scaffolds,” Biomacromolecules, vol. 7, no. 12, pp. 3345–3355, 2006. View at: Publisher Site | Google Scholar
  8. A. Wang, Q. Ao, W. Cao et al., “Porous chitosan tubular scaffolds with knitted outer wall and controllable inner structure for nerve tissue engineering,” Journal of Biomedical Materials Research Part A, vol. 79, no. 1, pp. 36–46, 2006. View at: Publisher Site | Google Scholar
  9. L. Peng, X. R. Cheng, J. W. Wang, D. X. Xu, and G. Wang, “Preparation and evaluation of porous chitosan/collagen scaffolds for periodontal tissue engineering,” Journal of Bioactive and Compatible Polymers, vol. 21, no. 3, pp. 207–220, 2006. View at: Publisher Site | Google Scholar
  10. Y. Wan, A. X. Yu, H. Wu, Z. X. Wang, and D. J. Wen, “Porous-conductive chitosan scaffolds for tissue engineering II. In vitro and in vivo degradation,” Journal of Materials Science: Materials in Medicine, vol. 16, no. 11, pp. 1017–1028, 2005. View at: Publisher Site | Google Scholar
  11. H. F. Liu, F. L. Yao, Y. Zhou et al., “Porous poly (DL-lactic acid) modified chitosan-gelatin scaffolds for tissue engineering,” Journal of Biomaterials Applications, vol. 19, no. 4, pp. 303–322, 2005. View at: Publisher Site | Google Scholar
  12. L. Ma, C. Y. Gao, Z. W. Mao et al., “Collagen/chitosan porous scaffolds with improved biostability for skin tissue engineering,” Biomaterials, vol. 24, no. 26, pp. 4833–4841, 2003. View at: Publisher Site | Google Scholar
  13. J. Yang, T. W. Chung, M. Nagaoka, M. Goto, C.-S. Cho, and T. Akaike, “Hepatocyte-specific porous polymer-scaffolds of alginate/galactosylated chitosan sponge for liver-tissue engineering,” Biotechnology Letters, vol. 23, no. 17, pp. 1385–1389, 2001. View at: Publisher Site | Google Scholar
  14. E. Khor and L. Y. Lim, “Implantable applications of chitin and chitosan,” Biomaterials, vol. 24, no. 13, pp. 2339–2349, 2003. View at: Publisher Site | Google Scholar
  15. S. Dumitriu and E. Chornet, “Inclusion and release of proteins from polysaccharide-based polyion complexes,” Advanced Drug Delivery Reviews, vol. 31, no. 3, pp. 223–246, 1998. View at: Publisher Site | Google Scholar
  16. C. Peniche and W. Arguelles-Monal, “Chitosan based polyelectrolyte complexes,” Macromolecular Symposia, vol. 168, pp. 103–116, 2001. View at: Google Scholar
  17. W. G. T. Willats, P. Knox, and J. D. Mikkelsen, “Pectin: new insights into an old polymer are starting to gel,” Trends in Food Science & Technology, vol. 17, no. 3, pp. 97–104, 2006. View at: Publisher Site | Google Scholar
  18. P. Bernabé, C. Peniche, and W. Argüelles-Monal, “Swelling behavior of chitosan/pectin polyelectrolyte complex membranes. Effect of thermal cross-linking,” Polymer Bulletin, vol. 55, no. 5, pp. 367–375, 2005. View at: Publisher Site | Google Scholar
  19. L. S. Liu, M. L. Fishman, J. Kost, and K. B. Hicks, “Pectin-based systems for colon-specific drug delivery via oral route,” Biomaterials, vol. 24, no. 19, pp. 3333–3343, 2003. View at: Publisher Site | Google Scholar
  20. F. Zhao, W. L. Grayson, T. Ma, B. Bunnell, and W. W. Lu, “Effects of hydroxyapatite in 3-D chitosan-gelatin polymer network on human mesenchymal stem cell construct development,” Biomaterials, vol. 27, no. 9, pp. 1859–1867, 2006. View at: Publisher Site | Google Scholar
  21. B. Q. Li, Q. L. Hu, X. Z. Qian, Z. P. Fang, and J. C. Shen, “Bioabsorbable chitosan/hydroxyapatite composite rod prepared by in-situ precipitation for internal fixation of bone fracture,” Acta Polymerica Sinica, no. 6, pp. 828–833, 2002. View at: Google Scholar
  22. W. Y. Xia, W. Liu, L. Cui et al., “Tissue engineering of cartilage with the use of chitosan-gelatin complex scaffolds,” Journal of Biomedical Materials Research Part B, vol. 71, no. 2, pp. 373–380, 2004. View at: Publisher Site | Google Scholar
  23. V. Chiono, E. Pulieri, G. Vozzi, G. Ciardelli, A. Ahluwalia, and P. Giusti, “Genipin-crosslinked chitosan/gelatin blends for biomedical applications,” Journal of Materials Science: Materials in Medicine, vol. 19, no. 2, pp. 889–898, 2008. View at: Publisher Site | Google Scholar
  24. Z. S. Li, H. R. Ramay, K. D. Hauch, D. M. Xiao, and M. Q. Zhang, “Chitosan-alginate hybrid scaffolds for bone tissue engineering,” Biomaterials, vol. 26, no. 18, pp. 3919–3928, 2005. View at: Publisher Site | Google Scholar
  25. D. Verma, K. S. Katti, D. R. Katti, and B. Mohanty, “Mechanical response and multilevel structure of biomimetic hydroxyapatite/polygalacturonic/chitosan nanocomposites,” Materials Science and Engineering C, vol. 28, no. 3, pp. 399–405, 2008. View at: Publisher Site | Google Scholar
  26. D. Verma, K. S. Katti, and D. R. Katti, “Effect of biopolymers on structure of hydroxyapatite and interfacial interactions in biomimetically synthesized hydroxyapatite/biopolymer nanocomposites,” Annals of Biomedical Engineering, vol. 36, no. 6, pp. 1024–1032, 2008. View at: Publisher Site | Google Scholar
  27. D. Verma, K. S. Katti, and D. R. Katti, “Osteoblast adhesion, proliferation and growth on polyelectrolyte-complex-hydroxyapatite nanocomposites,” Philosophical Transactions. Series A, vol. 368, no. 1917, pp. 2083–2097, 2010. View at: Google Scholar
  28. M.-W. Lee, C.-L. Hung, J.-C. Cheng, and Y.-J. Wang, “A new anti-adhesion film synthesized from polygalacturonic acid with 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide crosslinker,” Biomaterials, vol. 26, no. 18, pp. 3793–3799, 2005. View at: Publisher Site | Google Scholar
  29. W. Argüelles-Monal, O. L. Hechavarría, L. Rodríguez, and C. Peniche, “Swelling of membranes from the polyelectrolyte complex between chitosan and carboxymethyl cellulose,” Polymer Bulletin, vol. 31, no. 4, pp. 471–478, 1993. View at: Publisher Site | Google Scholar
  30. S. M. Lim, D. K. Song, S. H. Oh, D. S. Lee-Yoon, E. H. Bae, and J. H. Lee, “In vitro and in vivo degradation behavior of acetylated chitosan porous beads,” Journal of Biomaterials Science, Polymer Edition, vol. 19, no. 4, pp. 453–466, 2008. View at: Publisher Site | Google Scholar
  31. W. Tachaboonyakiat, T. Serizawa, and M. Akashi, “Inorganic-organic polymer hybrid scaffold for tissue engineering—II: partial enzymatic degradation of hydroxyapatite-chitosan hybrid,” Journal of Biomaterials Science, Polymer Edition, vol. 13, no. 9, pp. 1021–1032, 2002. View at: Publisher Site | Google Scholar
  32. K. Tomihata and Y. Ikada, “In vitro and in vivo degradation of films of chitin and its deacetylated derivatives,” Biomaterials, vol. 18, no. 7, pp. 567–575, 1997. View at: Publisher Site | Google Scholar
  33. S. Hirano, H. Tsuchida, and N. Nagao, “N-acetylation in chitosan and the rate of its enzymic hydrolysis,” Biomaterials, vol. 10, no. 8, pp. 574–576, 1989. View at: Google Scholar
  34. W. L. Cao, M. Y. Cheng, Q. Ao, Y. D. Gong, N. M. Zhao, and X. F. Zhang, “Physical, mechanical and degradation properties, and schwann cell affinity of cross-linked chitosan films,” Journal of Biomaterials Science, Polymer Edition, vol. 16, no. 6, pp. 791–807, 2005. View at: Publisher Site | Google Scholar
  35. D. W. Ren, H. F. Yi, W. Wang, and X. J. Ma, “The enzymatic degradation and swelling properties of chitosan matrices with different degrees of N-acetylation,” Carbohydrate Research, vol. 340, no. 15, pp. 2403–2410, 2005. View at: Publisher Site | Google Scholar
  36. A. Cárdenas, W. Argüelles-Monal, F. M. Goycoolea, I. Higuera-Ciapara, and C. Peniche, “Diffusion through membranes of the polyelectrolyte complex of chitosan and alginate,” Macromolecular Bioscience, vol. 3, no. 10, pp. 535–539, 2003. View at: Publisher Site | Google Scholar
  37. J. R. R. de Souza, J. I. X. de Carvalho, M. T. S. Trevisan, R. C. M. de Paula, N. Ricardo, and J. P. A. Feitosa, “Chitosan-coated pectin beads: characterization and in vitro release of mangiferin,” Food Hydrocolloids, vol. 23, no. 8, pp. 2278–2286, 2009. View at: Publisher Site | Google Scholar
  38. K. D. Yao, H. L. Tu, F. Cheng, J. W. Zhang, and J. Liu, “pH-sensitivity of the swelling of a chitosan-pectin polyelectrolyte complex,” Angewandte Makromolekulare Chemie, vol. 245, pp. 63–72, 1997. View at: Google Scholar
  39. G. Balooch, G. W. Marshall, S. J. Marshall, O. L. Warren, S. A. S. Asif, and M. Balooch, “Evaluation of a new modulus mapping technique to investigate microstructural features of human teeth,” Journal of Biomechanics, vol. 37, no. 8, pp. 1223–1232, 2004. View at: Publisher Site | Google Scholar
  40. R. Khanna, K. S. Katti, and D. R. Katti, “Nanomechanics of surface modified nanohydroxyapatite particulates used in biomaterials,” Journal of Engineering Mechanics, vol. 135, no. 5, pp. 468–478, 2009. View at: Publisher Site | Google Scholar
  41. K. S. Katti, P. Turlapati, D. Verma, R. Bhowmik, P. K. Gujjula, and D. R. Katti, “Static and dynamic mechanical behavior of hydroxyapatite-polyacrylic acid composites under simulated body fluid,” American Journal of Biochemistry and Biotechnology, vol. 2, no. 2, pp. 73–79, 2006. View at: Google Scholar
  42. S. A. S. Asif, K. J. Wahl, R. J. Colton, and O. L. Warren, “Quantitative imaging of nanoscale mechanical properties using hybrid nanoindentation and force modulation,” Journal of Applied Physics, vol. 90, no. 3, pp. 1192–1200, 2001. View at: Publisher Site | Google Scholar
  43. K. Kato, Y. Eika, and Y. Ikada, “In situ hydroxyapatite crystallization for the formation of hydroxyapatite/polymer composites,” Journal of Materials Science, vol. 32, no. 20, pp. 5533–5543, 1997. View at: Google Scholar
  44. F. Bigucci, B. Luppi, T. Cerchiara et al., “Chitosan/pectin polyelectrolyte complexes: selection of suitable preparative conditions for colon-specific delivery of vancomycin,” European Journal of Pharmaceutical Sciences, vol. 35, no. 5, pp. 435–441, 2008. View at: Publisher Site | Google Scholar
  45. D. Verma, K. Katti, and D. Katti, “Experimental investigation of interfaces in hydroxyapatite/polyacrylic acid/polycaprolactone composites using photoacoustic FTIR spectroscopy,” Journal of Biomedical Materials Research Part A, vol. 77, no. 1, pp. 59–66, 2006. View at: Publisher Site | Google Scholar
  46. W. W. Thein-Han and R. D. K. Misra, “Biomimetic chitosan-nanohydroxyapatite composite scaffolds for bone tissue engineering,” Acta Biomaterialia, vol. 5, no. 4, pp. 1182–1197, 2009. View at: Publisher Site | Google Scholar
  47. H. P. Wampler and A. Ivanisevic, “Nanoindentation of gold nanoparticles functionalized with proteins,” Micron, vol. 40, no. 4, pp. 444–448, 2009. View at: Publisher Site | Google Scholar
  48. J.-Y. Rho and G. M. Pharr, “Effects of drying on the mechanical properties of bovine femur measured by nanoindentation,” Journal of Materials Science: Materials in Medicine, vol. 10, no. 8, pp. 485–488, 1999. View at: Publisher Site | Google Scholar
  49. S. Hengsberger, A. Kulik, and Ph. Zysset, “Nanoindentation discriminates the elastic properties of individual human bone lamellae under dry and physiological conditions,” Bone, vol. 30, no. 1, pp. 178–184, 2002. View at: Publisher Site | Google Scholar

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