imaging of bone ultrastructure using atomic force microscopy · completely and neither are the...

Imaging of Bone Ultrastructure using Atomic Force Microscopy

P. J. Thurner*,1,2, E. Oroudjev2, R. Jungmann2, C. Kreutz2, J. H. Kindt2, G. Schitter2, T. O. Okouneva3, M. E. Lauer2, G. E. Fantner2, H. G. Hansma2, and P. K. Hansma2 1Bioengineering Science Research Group, School of Engineering Science, University of Southampton,

Highfield, Southampton SO17 1BJ, UK 2Department of Physics, University of California, Santa Barbara, CA 93106, USA 3Neuroscience Research Institute, Santa Barbara, CA 93106, USA Atomic force microscopy (AFM) is a superb tool for imaging of bone ultrastructure in a close to physiological state. AFM provides similar spatial resolution as transmission electron microscopy without the need of excessive sample dehydration and preparation. Due to this fact AFM imaging of bone ultrastructure can be performed in a functional manner, i.e. imaging can be combined with in situ experiments such as chemical treatments or mechanical deformation, or it can be used as a quality control instrument to follow up on tissue engineered bone matrix. In this communication we present an overview of bone ultrastructure assessment using AFM. We present protocols for sample preparation, provide images of the different surfaces encountered in bone, and show examples of functional imaging experiments. Overall we conclude that AFM is a very valuable tool for bone structure assessment and that we are far from exploiting all the possibilities this technique offers for the investigation of bone ultrastructure and mechanical competence.

Keywords bone; ultrastructure; atomic force microscopy; functional imaging; tissue engineering; deformation; chemical treatment; osteoblast

1. Introduction

Bone is a remarkable natural nano-composite material, being lightweight and both stiff and tough. At the macroscopic scale bone is present in two different forms: cortical (or compact) bone forms the solid shell of all bones providing shape and structure, whereas trabecular (or cancellous) bone, a spongy structure, fills the ends of the long bones and all space within vertebrae [1]. Trabecular bone provides for a transfer of loads from joint faces to the midshaft of long bones and 90% of the load transfer in vertebrae avoiding high stresses and strains that could lead to fracture. Both cortical and trabecular bone are covered with an outer surface called osteoid, which differs from the bulk material. Below this osteoid layer bone is usually arranged in layers or packets, which can be concentric in compact bone (osteons) and are irregular in trabecular bone. Detailed knowledge about bone ultrastructure below this level and its relation to bone material properties is essential for the development of new biomimetic materials as well as complementary diagnostic tools and therapies for bone diseases. Such investigations on bone are mainly motivated by the immense social impact in health care due to osteoporosis in post-menopausal women and the aged [2, 3] as well as the enormous medical costs (13.8 billion US$ in the USA alone in 1995). Osteoporosis results in bone loss and change of trabecular architecture and tissue composition, causing a decrease in bone strength. It is generally accepted that besides bone density also bone microarchitecture and bone matrix material properties, a combination referred to as bone quality, are also important factors influencing fracture resistance. Whereas there exist numerous studies on the microarchitecture of trabecular bone and how these could be exploited for fracture risk diagnosis [4-6], there are almost no studies addressing the bone ultrastructure and the related bone matrix material properties of trabecular bone. This is despite the facts that, at least for cortical bone, bone composition does change with age and disease [7, 8], and that fracture toughness decreases due to age and disease [9]. Despite these empirical findings the exact nano- or macromolecular structure of bone is not resolved

* Corresponding author: e-mail: [email protected], Phone: +1-415-502-0809

Modern Research and Educational Topics in Microscopy. A. Méndez-Vilas and J. Díaz (Eds.) ©FORMATEX 2007 _______________________________________________________________________________________________

37

completely and neither are the mechanisms that are responsible making bone a stiff, tough and durable nanocomposite material. From the view of the materials scientist this knowledge is essential. It seems that only a small fraction of noncollagenous proteins (NCPs) in bone tissue guides and controls the assembly of the major components, collagen type I fibrils, about 100 nm in diameter and up to 10 µm and more in length, as well as carbonated hydroxyapatite (Ca10(PO4)6(OH)2) crystals with typical sizes of 50 nm x 25 nm x 3 nm [10-12]. However, how the self-assembly is achieved and what the properties and functions of the NCPs both for development and later for the material properties are, remains unclear. Only recently it was proposed that these NCPs could form a matrix layer in between mineralized collagen fibrils that can repeatedly dissipate energy without failure through a self-healing capability [13-16]. In the recent past also atomic force microscopy (AFM) has been used for the assessment of ultrastructure and nanomechanical properties of bone [12, 13, 17-27]. AFM [28] has become one of the most important tools to image, probe, and manipulate on the nanoscale. It has been used extensively for investigations of biological materials [29] such as DNA [30], globular proteins and protein networks [31], cells [32] and more. The most important reason why AFM is so attractive for investigations of biological materials is the fact that samples of interest can be probed and imaged in conditions very close to their natural state, e.g. in liquid or saline. This can be done with a spatial resolution similar as achieved in transmission electron microscopy, but without the need to dry samples and put them in a vacuum. AFM is thus a unique and complementary tool for the assessment of bone and other tissues. In this chapter we show the different ultrastructural arrangements that can be uncovered in bone using AFM and how functional imaging approaches can help to uncover more details about bone ultrastructure and development. AFM is not, of course, the only tool useful for characterizing bone ultrastructure. But to explore all recent breakthroughs with techniques other than AFM is beyond the scope of this chapter.

2. Sample preparation and imaging methods

2.1 Trabecular and cortical bone

In general all bone samples used for the work presented here were of bovine origin. Parts of femora and vertebrae where freshly obtained from a local grocery store (Gelson’s Market, Santa Barbara, USA). The bone was cleaned of soft tissue and sawed into smaller pieces using a butcher’s bandsaw and/or a smaller bandsaw (Marmed Inc. Cleveland, OH, USA). Subsequently bone marrow was cleaned out with a jet of pressurized water. Prior to imaging all samples were briefly rinsed with HPLC grade water (EMD Chemicals Inc., Gibbstown, NJ), blotted dry on a Kimwipe, and desiccated in vacuum for 30 min. In general there are two types of surfaces that best resemble the natural state of bone. These are native, i.e. osteoid and fracture surfaces. Native surfaces are generally smooth so that sample preparation is relatively simple. Trabecular bone from vertebra was cut to cuboids measuring 4 mm x 4 mm x 5 mm. For fracture surface investigations, the surfaces of the bone cuboids were stained with Coomassie Blue R-250 (Sigma-Aldrich, 0.2 g/ml) in Na-buffer (150 mM NaCl, 10 mM Hepes, pH 7.0) for 5 min, and then rinsed in Ca-buffer (110 mM NaCl, 40 mM CaCl2, 10 mM Hepes, pH 7.0) to remove any excess stain. Subsequently, a stained bone cuboid was clamped in a vice, with the bone load axis parallel to the vice grip edge. The half of the cuboid extending from this vice was clamped into a second vice. The cuboid was then pulled apart, i.e. fractured in tension and dried as described above. To locate an area suitable for AFM imaging, the dried cube was then placed under a dissection microscope. The Coomassie stain allowed to clearly distinguish between stained (blue) external surfaces and un-stained (white) fracture surfaces. Fracture areas appearing translucent underneath a dissection microscope were most smooth and suitable for AFM investigation. Pieces with apparent smooth fracture surfaces were then removed with a scalpel, and glued onto a custom sample disc using epoxy resin (2-Ton Clear Epoxy, Devcon, Danvers, MA, USA) with the fracture surface facing upward, being oriented roughly parallel to the sample disc.

©FORMATEX 2007Modern Research and Educational Topics in Microscopy. A. Méndez-Vilas and J. Díaz (Eds.) _______________________________________________________________________________________________

38

Fig. 1 AFM height images obtained in tapping mode in air. (A) Native cortical bone and (B) native trabecular bone surfaces both exhibit pristine collagen fibrils with their typical corrugation of ~67 nm. The inset in (B) is an AFM height image of a similar native surface imaged in Na-buffer. In liquid the 67 nm banding pattern is not observed. In contrast to the native surfaces fractured cortical bone (C), and fractured trabecular bone (D) surfaces both exhibit fibrilar features coated with small particulates – platelet-shaped hydroxyapatite crystals. All images were obtained in tapping mode at 1Hz line frequency. The inset was obtained in contact mode and a liquid environment at 1Hz Pieces of bovine femora were cut along and perpendicular to the long axis of the bone. After marrow extraction single trabeculae with rod-like shape were directly cut out using a pair of scissors. Fracture surfaces in single trabeculae were produced by “peeling” off a top layer of their outer surface using a scalpel in physiological solution. A small angle notch was made on the sample surface and the piece sticking out of the surface layer was then clamped with a pair of tweezers and peeled off. All samples were then washed and dried as described above, and subsequently either glued to sample discs using 2-ton epoxy or directly inserted into the in situ three-point bending device (cp. Section 4) Cortical bone samples from the femur were cut to small pieces containing native surfaces and washed and dried as described above and glued to sample discs using 2-ton epoxy. In order to obtain a smooth fracture surface of cortical bone, pieces of the femur were first cut into rectangular sheets, 1mm in thickness, 30 mm in length and, 10 mm in width, with the long axis of the sheet being oriented parallel to


39

the long axis of the bone, using a diamond wafering blade (TechCut 4, Allied High Tech Products Inc., Rancho Dominguez, CA, USA) under constant irrigation with tap water. Subsequently two notches, 5 mm in length, in the middle of both shorter sides (thus the notches were also oriented parallel to the long axis of the bone) were created with the same wafering blade. Subsequently the samples were snap frozen in liquid nitrogen and a crack was initiated by spreading apart one of the notches. The produced samples had smooth fracture surfaces oriented roughly parallel to the long axis of the sheet. They were rinsed and dried as described before and were glued to sample discs using two-ton epoxy.

2.2 Tissue engineered bone

Bone-like tissue can be produced in vitro by cultured osteoblasts in a similar manner to osteoblasts that produce bone de novo in a native bone matrix. The MC3T3-E1 cell line was derived from murine calvaria bone tissue and subsequently subcloned into a few cell lines that demonstrate relatively high or low osteoblast differentiation potential and corresponding ability to produce highly mineralized extracellular matrix (ECM) [33]. In our experiments we used the MC3T3-E1 (subclone 14) cell line, which was shown to differentiate into mature osteoblasts upon induction with ascorbic acid and then to deposit well-mineralized bone-like ECM. MC3T3-E1-14 cells were maintained in culture in incomplete Minimal Essential Medium Alpha Medium (AMEM) without ascorbic acid (custom-made product, Invitrogen) supplemented with 10% fetal bovine serum (FBS) in an incubator kept at 370C and in 5% CO2. Atomically flat surfaces for cell culturing were made through cleaving 12 mm mica discs, and sterilization was achieved by 15 minutes of dry cycle in an autoclave. Care was taken to keep the cleaved side of mica discs free from contact with any other surface or from deposits from other materials/contaminants in the autoclave. Sterilized mica discs were placed into wells of a sterile 12-well plate with one mica disc per well (cleaved side up) and MC3T3-E1-14 cells were seeded into wells at a desired density (30000 cells/ml or more depending on targeted tissue thickness) in 2 ml of culturing medium. After 24 hours of recovery time, medium was exchanged to a 10% FBS in complete medium (AMEM) supplemented with ascorbic acid (50 µg/ml) and glycerol 2-phosphate (10 mM) to induce osteoblast differentiation and supply osteoblasts with a source of phosphate for production of the inorganic phase of ECM. Differentiated osteoblasts were supported in culture by regular medium exchange every 3-4 days. Osteoblasts can be fixed on mica disc surface by formaldehyde to study cell-to-cell or cell-to-surface contacts or to produce snaps of early stages of ECM deposition. For this purpose mica discs were first transferred into a petri dish and gently washed with three changes of phosphate-buffered saline (PBS). Cells were fixed by substitution of PBS with freshly prepared 3.7% formaldehyde solution in PBS and subsequent incubation at room temperature for 1 hour. Fixation solution was then removed by gentle wash with four changes of sterile ultra pure water. Samples can be stored in sterile ultra pure water in refrigerated conditions for at least one week. Samples intended for imaging in air were taken out of the petri dish and excess of water was removed by blotting the sample substrate on Whatman paper. Then these samples were dried in a dust free environment and kept in low humidity conditions in refrigerator. Significant parts of the grown tissue can become deposited at the mica disc edges and connect to the plastic surfaces of the well plate. In such cases the samples were removed carefully by cutting the deposited ECM at the disc edges using a scalpel. Samples intended to study the deposited ECM where washed three times in PBS and four times in sterile ultra pure water (each washing lasts approximately 10 minutes). They can be stored in water or dried in air in a manner similar to samples with osteoblasts fixed on mica surface. For investigation of the “smooth” side of the bone-like tissue, the deposited ECM was peeled from mica surface using forceps, turned upside down, and then dried in air.


40

Fig. 2 AFM image of single collagen fibrils on a native trabecular bone surface obtained in contact mode. (A) Deflection image, revealing collagen ultra�structure on top of the banding pattern (white arrows). (B) 3D view of the highlighted area in (A) reveals the collagen fine structure (image size 350 nm). Imaged in air using contact mode at 1Hz.

2.3 AFM imaging

Atomic force microscopy was performed on two different AFM systems, a Dimension 3100 (Veeco Inc., Santa Barbara, CA, USA) with a Nanoscope IIIa controller and a Multimode with Nanoscope IV controller (Veeco Inc.). Images were captured in both tapping and in contact mode using MLCT-AUHW (Veeco Inc.) and RTESPW (Veeco Inc.) cantilevers respectively. The individual imaging parameters are given in the corresponding figure captions. Generally all images were processed using the Nanoscope software (Version 7.00R2.sr2, Veeco Inc.). Height images were processed using a 1st order planefit and a 0th order flattening command, whereas amplitude images were processed using only a 1st order planefit.

3. Native and fractured bone surfaces

Native and fractured bone surfaces of cortical as well as trabecular bone were prepared as described in Section 2.1. Imaging experiments on both surfaces are summarized in Fig. 1. Bare collagen fibrils were found on both native surfaces, whereas fractured surfaces exhibited fibrilar morphologies covered with small particles – hydroxyapatite crystals. Such morphologies have previously been found on trabecular bone surfaces investigated with AFM [12, 13, 23] and other techniques [10, 34-37], but so far no investigation on cortical bone was reported. The hydroxyapatite crystals seen on the fracture surfaces in Fig. 1D and C range in their largest dimension from about ~50 nm to ~250 nm. This is in agreement of particle sizes previously reported [10-12, 35], although it seems that the crystals seen in the images presented here are mostly at the upper end of this spectrum. The bare collagen fibrils (cp. Fig. 1A and B) exhibit the typical corrugation or D-banding with 67 nm periodicity. This corrugation is in some cases well aligned in neighboring fibrils as can be seen in Fig. 1 B, yet it is not clear whether the corrugation itself as well as the alignment is an intrinsic effect of the tissue or a drying artifact. Fig. 1 B and the included inset show a comparison between a native trabecular bone surface imaged in air (cp. Fig. 1B) with tapping mode AFM and in Na-buffer (inset) with contact mode AFM. The fact that the 67 nm collagen banding pattern is observed in air but not in liquid could be due to the fact that the pattern is a drying artifact. Hence, it could be that the structure of collagen in a hydrated environment is changed like proposed in [38] due to protein hydration. Although Fig. 1A was obtained using tapping mode AFM and the inset using contact mode, it is not plausible that this is the reason for not observing the banding


41

pattern. The structure of water surrounding collagen has been studied by a wide variety of techniques and is considered a paradigm for protein hydration [38, 39]. It is commonly accepted that at least some water in collagen fibrils differs from bulk, ambient water. Within the most simple scheme, this interstitial water is separated into two distinct classes tightly bound and “free” or bulk-like. More elaborate models imagine three or even more fractions. Tightly bound waters are believed to stabilize the triple helix by participating in the H�bond backbone. In a larger context, however, the specific role of water in collagen assembly, structure and stability is still an open question [40]. Fig. 2 shows a native trabecular bone surface imaged in contact mode at high magnification uncovering some of the ultrastructure of the collagen fibrils (cp. white arrows in Fig. 3A). A section analysis in the vicinity of Fig. 2A (not shown) resulted in an average height of the banding of molecules of (5.12 ± 0.84) nm and an average banding periodicity of (66.79 ±1.24) nm, which correlates well with reported findings [41].

4. Functional imaging of bone

Fig. 3A shows a fractured surface of vertebral trabecular bone. The fracture surface exhibits fibrilar features similar as the ones presented in Fig. 1C and D, coated with hydroxyapatite particles. Chemical treatment of this fracture surface with a saturated aqueous solution of NaF (pH 7.0) in situ, for about 20 s, and rinsing in a steady flow of HPLC grade water for 2 min, completely changed the morphology. Following treatment, bare collagen fibril networks, characterized by their typical D-banding (~67 nm periodicity), were clearly resolved. Similar morphological changes can be observed, when the freshly prepared bovine samples were exposed to solutions containing EDTA [12]. In contrast to the EDTA-treated samples, images obtained from samples treated with NaF generally reveal the presence of a few remaining mineral platelets. These platelets exhibited similar sizes and shapes as those seen during initial imaging of the untreated fracture surface (cp. Fig. 4 A). We have previously reported an additional analysis of larger, mm-sized samples treated with both NaF and EDTA solutions as well as bone powder treated with NaF protein extraction [12]. We found that larger samples treated with saturated solutions of NaF for 18 h only loose a few percent of their initial weight and that more protein is liberated from bone powder when following NaF treatment. Hence, NaF detaches hydroxyapatite crystals whereas they dissolve in the presence of EDTA.

Fig. 3 : Time-resolved sequence of AFM images of a bovine trabecular bone fracture surface, imaged in N2 gas in tapping mode at 1Hz, exhibiting collagen fibrils coated with mineral platelets (A), and the same sample location after treatment with a saturated (1M) solution of NaF exhibiting bare collagen fibrils (B).


42

Fig. 4 (A) AFM mixed height and amplitude image of a dry bovine single trabecula loaded in three-point bending. Overlaid is a displacement vector field evaluated using digital image correlation on a set of two images (in unloaded and in bent state). Image obtained in air using tapping mode (0.5 Hz). (B) Same surface with a contour plot overlay of regions with same relative displacement in x-direction (fast scan axis). There seems to be a border region (arrows) haphazardly dividing the surface in two domains sliding along each other. Using a newly developed prototype for in situ three-point bending of small bone specimens [42] we investigated a single trabecula bent in dry condition. A 20 µm region on the surface of the sample was imaged in air, before and after bending to a displacement of 99 µm and a force of 1.9 N. Subsequently a surface displacement measurement was performed using the amplitude the images obtained before and after bending using a regional interrogation-window-based particle-image-velocimetry algorithm [43]. Dividing the image in 16x16 px large quadratic interrogation windows and allowing 50% overlap from one image to the next was adequate. For the actual correlation calculation, a freeware module (MatPIV, [44]) for MATLAB (The MathWorks, Natick, MA, USA) was used. From the obtained displacement information, we calculated the local strain in the direction parallel to the long axis of the trabecula (fast scan axis), by computing the gradient in this direction: εx = du/dx. For validation purposes we also scanned a calibration grating with 5 µm pitch size (Veeco Inc.) and subjected the retrieved data to the surface displacement measurement, which resulted in negligible strains [42]. Fig. 4A and B show the results of the bending experiment, evaluated with the surface displacement algorithm. The white arrows in Fig. 4A show a collagen fiber bundle spanning from the top left corner almost diagonal over the image. Fig. 4B represents the surface displacement contour map in the bending direction (x-axis and fast scanning axis). It seems that the image is divided into two domains separated by this fiber bundle, sliding along each other. We calculated the average strain du/dx in the bending direction to be 0.024 ± 0.006.

Fig. 5 Series of AFM images of a native trabecular surface recorded at 2000 lines or 8 images per second. Every 8th image of the series is shown, showing a zoom onto bare collagen fibrils on this surface.


43

Fig. 5 shows a series of AFM images of a native surface of a single trabecula, taken a line rate of 2 KHz or 8 frames/s using a custom-made high-speed AFM [45]. Next generation AFMs [46, 47] allow to image significantly faster than current commercial AFM-systems, and will enable to study dynamic processes on the nanoscale with a much higher time resolution. This will open the door for real-time functional imaging of bone, e.g. during chemical treatment or mechanical testing, and may allow new insights into the function of this highly complex nano-composite material.

5. Tissue engineered bone

Samples of differentiated murine osteoblasts (Fig. 6A) and bone-like ECM deposited by these cells on cleaved mica surfaces were prepared as described above and imaged using AFM. After first three days of stimulating osteoblast differentiation, deposition of biopolymers with fibrilar appearance can be clearly seen in Fig.6B in the vicinity of most cells. These initial deposits can serve as an anchor points for cell adhesion and as a foundation for the first layer of the ECM deposited by osteoblast in the next few days. In order to investigate these very first layers of ECM, deposited by osteoblasts attached directly to mica surface, before other osteoblasts had a chance to modify their appearance/morphology, we took relatively “young” samples of deposited ECM (10-14 days of cultivation) and imaged the parts of mica that were not yet covered by ECM. At the border of those areas where open mica surface would meet areas covered with ECM we were able to find a very thin deposits of ECM (Fig. 6A) that had significant morphological differences (cheese-like appearance) from more mature (old) deposits (Fig. 7). Interestingly, the thickness of these initial deposits remains more or less constant (150-200 nm on average, cp. Fig 6 B) on distances up to 40 µm (data not shown) from the edge of the ECM. This could be a result of uniform deposition of ECM by one or few “priming” osteoblasts at their entire side facing the mica substrate.

Fig. 6 (A) AFM amplitude (tapping mode) image of fixed, differentiated murine osteoblasts MC3T3-E1-14 after 3 day cultivation on cleaved mica surface in air. Deposits of fibrilar matter (ECM molecules) in vicinity of cells can be clearly seen. (B) AFM height (tapping mode) image of naive ECM matrix deposited by murine osteoblasts after 14 days of cultivation on cleaved mica surface in liquid. Cells and water-soluble components of ECM were removed by multiple washes in ultra-pure water. Mica surface in a left bottom corner is still not covered with ECM. On the rest of the mica surface ECM was deposited as a network of amorphous matter. A cross section analysis of this network (inset) shows that it has relatively uniform thickness (around 200 nm) at least on 10-20 µm distances from the edge of network. Openings in ECM network generally reach the mica surface. Numerous openings of a different size that are clearly seen in Fig. 6B do penetrate all the way down to the mica surface and most likely are results of contacts of focal adhesion points on the cell membrane of


44

osteoblasts facing the mica substrate [48]. The majority of these contacts seem to persist through the entire period of initial ECM deposition. ECM material is presumably deposited in a pockets formed between these focal adhesion points and the mica surface. More mature (old) ECM deposits (15-35 days) have no signs of this “cheese-like” appearance on the either side of tissue as shown in Fig. 7.

Fig. 7 Mica-oriented (bottom) side of ECM matrix deposited by murine osteoblasts MC3T3-E1-14 after 21 days of cultivation on cleaved mica surface. AFM amplitude (tapping mode) images in air. Cells and water-soluble components of ECM were removed by multiple washes in ultra-pure water. ECM surface at mica-ECM interface (A and B) is rich in fibrils of different size (width) that span ECM in irregular manner. We speculate that these fibers are collagen fibrils covered with non-collagenous ECM proteins and hydroxyapatite crystals. At higher magnification (B) a more or less regular pattern (~25 nm) can be observed on these fibrils, possibly reflecting the internal structural/functional pattern in these mineralized collagen fibrils. (C and D) Hydroxyapatite crystals have been partially removed by brief wash (5 seconds) with EDTA followed by extensive wash with ultra-pure water. Collagen fibrils (exposed by EDTA treatment) with characteristic 67 nm banding pattern can clearly be seen in both images. In the higher magnification image (B) the ultrastructure of the banding in collagen fibrils can be observed. Significant increase in the thickness (up to ~200 µm, data not shown) is usually observed for such mature ECM deposits. These findings suggest that as more ECM is deposited by osteoblasts, initial contacts between mica surface and cell is lost, possibly due to osteoblast relocation. The relatively


45

smooth surface of mica-oriented side of ECM bears signs of numerous fibrils spanning the whole ECM body in all directions. At higher magnifications (cp. Fig. 7B) characteristic signs of mineralization of these fibrils with hydroxyapatite crystals become evident. A relatively uniform band pattering of these crystals along the fibrils (~25 nm average band width, data not shown) is most likely a result of regular placement of crystallization initiation centers along fibrils long axis. Short exposure (~5 seconds) of these surfaces to concentrated EDTA solution (100 mM) removes the majority of mineral (inorganic/hydroxyapatite) deposits from ECM surface (and probably some organic deposits that were associated with it), as demonstrated in Fig 7C and D. This treatment leaves most of fibrils fully or partially striped from their mineral coating and reveals on their surface a characteristic (~67 nm) band pattering that is a sign of collagen type I fibrils [17, 23, 41]. Generally the morphology of these “mature” deposits very closely resemble woven bone tissues that are initially formed at the places of bone repair or growth [1]. In general no significant increase in ECM mass was observed after 4- 5 weeks of osteoblast culturing despite a significant number of viable cells (data not shown). Overall, we successfully used AFM to study morphological, structural and temporal aspects of bone-like ECM deposition by osteoblasts cultured in vitro. The further application of AFM in combination with cell biology methods and other imaging techniques should significantly enhance our knowledge of ultra-structural organization of bone.

7. Summary

The presented results demonstrate how the versatility of AFM technique and its ability to combine it with in situ and ex situ time-lapsed experiments aide in the assessment of bone ultrastructure in a complementary fashion. Hence, AFM can be used for the inspection of bone surfaces and offers the possibility of quantification of local geometries, such as hydroxyapatite crystal sizes, collagen fibril diameters, or collagen ultrastructure. In additional functional imaging approaches based on chemical treatments or mechanical testing allow to assess bone ultrastructure in a 3D fashion and to investigate fundamental mechanisms involved in bone plasticity and failure. Such approaches become more and more important with the advent of high-speed or video-rate AFM imaging. Then AFM investigations might even uncover the dynamics of damage and plasticity processes, or chemical reactions at the nanometer scale. In the context of functional imaging, we also demonstrate that AFM can be used as a tool to study tissue engineered constructs and the in vitro development of bone. In the future AFM will surely become one of the most important tools in the realm of medical research for imaging, mechanical probing and manipulation of bone and other tissue constructs at the macromolecular scale in a real-time fashion.

Acknowledgements This work was supported by National Institutes of Health under Award R01 GM065354, by the NASA University Research, Engineering and Technology Institute on Bio-inspired Materials under Award No. NCC-1-02037 and by a research agreement with Veeco #SB030071. PJT acknowledges SNF fellowship PA002—111445, and RJ acknowledges DAAD scholarship No. D/05/42569. GS acknowledges TU Delft, faculty 3mE grant PAL 614.

References

[1] Baron, R., General Principles of Bone Biology, in Primer on the Metabolic Bone Diseases and Disorders of Mineral Metabolism, M.J. Favus, Editor. 2005, American Society for Bone and Mineral Research.

[2] Cooper, C., E.J. Atkinson, S.J. Jacobsen, W.M. O'Fallon, and L.J. Melton, 3rd. Am J Epidemiol, 1993. 137(9): p. 1001-5.

[3] Ray, N.F., J.K. Chan, M. Thamer, and L.J. Melton, 3rd. J Bone Miner Res, 1997. 12(1): p. 24-35. [4] Carballido-Gamio, J. and S. Majumdar. Curr Osteoporos Rep, 2006. 4(2): p. 64-70. [5] Ladinsky, G.A. and F.W. Wehrli. Curr Osteoporos Rep, 2006. 4(4): p. 140-7. [6] Ulrich, D., T. Hildebrand, B. Van Rietbergen, R. Muller, and P. Ruegsegger. Stud Health Technol Inform,

1997. 40: p. 97-112.


46

[7] Grynpas, M.D., J.H. Tupy, and J. Sodek. Bone, 1994. 15(5): p. 505-513. [8] McCreadie, B.R., M.D. Morris, T.C. Chen, D. Sudhaker Rao, W.F. Finney, E. Widjaja, and S.A. Goldstein.

Bone, 2006. 39(6): p. 1190-5. [9] Nalla, R.K., J.J. Kruzic, J.H. Kinney, and R.O. Ritchie. Bone, 2004. 35(6): p. 1240-1246. [10] Weiner, S. and W. Traub. Faseb J, 1992. 6(3): p. 879-85. [11] Eppell, S.J., W. Tong, J.L. Katz, L. Kuhn, and M.J. Glimcher. J Orthop Res, 2001. 19(6): p. 1027-34. [12] Kindt, J.H., P.J. Thurner, M.E. Lauer, B.L. Bosma, G. Schitter, G.E. Fantner, M. Izumi, J.C. Weaver, D.E.

Morse, and P.K. Hansma. Nanotechnology, 2007. 18(13): p. 135102 (8pp). [13] Fantner, G.E., T. Hassenkam, J.H. Kindt, J.C. Weaver, H. Birkedal, L. Pechenik, J.A. Cutroni, G.A.G.

Cidade, G.D. Stucky, D.E. Morse, and P.K. Hansma. Nature Materials, 2005. 4(8): p. 612-616. [14] Gupta, H.S., P. Fratzl, M. Kerschnitzki, G. Benecke, W. Wagermaier, and H.O. Kirchner. J R Soc Interface,

2007. 4(13): p. 277-82. [15] Gupta, H.S., J. Seto, W. Wagermaier, P. Zaslansky, P. Boesecke, and P. Fratzl. Proc Natl Acad Sci U S A,

2006. 103(47): p. 17741-6. [16] Gupta, H.S., W. Wagermaier, G.A. Zickler, D. Raz-Ben Aroush, S.S. Funari, P. Roschger, H.D. Wagner, and

P. Fratzl. Nano Lett, 2005. 5(10): p. 2108-11. [17] Sasaki, T., K. Debari, and M. Hasemi. Journal of Electron Microscopy, 1993. 42(5): p. 356-359. [18] Reilly, G.C., H.F. Knapp, A. Stemmer, P. Niederer, and M.L. Knothe Tate. Ann Biomed Eng, 2001. 29(12):

p. 1074-81. [19] Baranauskas, V., I. Garavello-Freitas, Z. Jingguo, and M.A. Cruz-Hofling. Journal of Vacuum Science &

Technology A, 2001. 19(4): p. 1042-1045. [20] Sasaki, N., A. Tagami, T. Goto, M. Taniguchi, M. Nakata, and K. Hikichi. Journal of Materials Science-

Materials in Medicine, 2002. 13(3): p. 333-337. [21] Xu, J., J.Y. Rho, S.R. Mishra, and Z. Fan. J Biomed Mater Res A, 2003. 67(3): p. 719-26. [22] Tong, W., M.J. Glimcher, J.L. Katz, L. Kuhn, and S.J. Eppell. Calcif Tissue Int, 2003. 72(5): p. 592-8. [23] Hassenkam, T., G.E. Fantner, J.A. Cutroni, J.C. Weaver, D.E. Morse, and P.K. Hansma. Bone, 2004. 35(1):

p. 4-10. [24] Hassenkam, T., H.L. Jorgensen, M.B. Pedersen, A.H. Kourakis, L. Simonsen, and J.B. Lauritzen. Micron,

2005. 36(7-8): p. 681-7. [25] Bozec, L., J. de Groot, M. Odlyha, B. Nicholls, and M.A. Horton. IEE Proc Nanobiotechnol, 2005. 152(5): p.

183-6. [26] Fantner, G.E., O. Rabinovych, G. Schitter, P. Thurner, J.H. Kindt, M.M. Finch, J.C. Weaver, L.S. Golde,

D.E. Morse, E.A. Lipman, I.W. Rangelow, and P.K. Hansma. Composites Science and Technology, 2006. 66(9): p. 1205-1211.

[27] Hassenkam, T., H.L. Jorgensen, and J.B. Lauritzen. Anat Rec A Discov Mol Cell Evol Biol, 2006. 288(10): p. 1087-94.

[28] Binnig, G., C.F. Quate, and C. Gerber. Physical Review Letters, 1986. 56(9): p. 930-933. [29] Jena, B. and J.K. Horber, eds. Atomic Force Microscopy in Cell Biology. Methods in Cell Biology, ed. L.

Wilson and P. Matsudaira. Vol. 68. 2002, Academic Press: San Diego. [30] Hansma, H.G., K. Kasuya, and E. Oroudjev. Curr Opin Struct Biol, 2004. 14(3): p. 380-5. [31] Poggi, M.A., E.D. Gadsby, L.A. Bottomley, W.P. King, E. Oroudjev, and H. Hansma. Anal Chem, 2004.

76(12): p. 3429-43. [32] Allison, D.P. and M.J. Doktycz. J Cell Mol Med, 2006. 10(4): p. 847-56. [33] Wang, D., K. Christensen, K. Chawla, G. Xiao, P.H. Krebsbach, and R.T. Franceschi. J Bone Miner Res,

1999. 14(6): p. 893-903. [34] Landis, W.J., K.J. Hodgens, J. Arena, M.J. Song, and B.F. McEwen. Microsc Res Tech, 1996. 33(2): p. 192-

202. [35] Rho, J.Y., L. Kuhn-Spearing, and P. Zioupos. Medical Engineering & Physics, 1998. 20(2): p. 92-102. [36] Weiner, S. and H.D. Wagner. Annual Review of Materials Science, 1998. 28: p. 271-298. [37] Thurner, P.J., B. Erickson, R. Jungmann, Z. Schriock, J.C. Weaver, G.E. Fantner, G. Schitter, D.E. Morse,

and P.K. Hansma. Engineering Fracture Mechanics, 2007. 74: p. 1928-1941. [38] Leikin, S., V.A. Parsegian, W.H. Yang, and G.E. Walrafen. Proceedings of the National Academy of

Sciences of the United States of America, 1997. 94(21): p. 11312-11317. [39] Chapman, G.E., S.S. Danyluk, and Mclauchl.Ka. Proceedings of the Royal Society of London Series B-

Biological Sciences, 1971. 178(1053): p. 465-&. [40] Fraser, P.E., D.R. Mclachlan, W.K. Surewicz, C.A. Mizzen, A.D. Snow, J.T. Nguyen, and D.A. Kirschner.

Journal of Molecular Biology, 1994. 244(1): p. 64-73.


47

[41] Venturoni, M., T. Gutsmann, G.E. Fantner, J.H. Kindt, and P.K. Hansma. Biochem Biophys Res Commun, 2003. 303(2): p. 508-13.

[42] Jungmann, R., G. Schitter, G.E. Fantner, M.E. Lauer, P.K. Hansma, and P.J. Thurner. Real-Time Microdamage and Strain Detection during Micromechanical Testing of Single Trabeculae. in Annual Meeting of the Society for Experimental Mechanics. 2007. Springfield, MA: Society for Experimental Mechanics Inc.

[43] Raffel, M., C. Willert, S. Wereley, and J. Kompenhans, Particle Image Velocimetry: A Practical Guide. 1998, Berlin: Springer.

[44] Sveen, J.K., An introduction to MatPIV v. 1.6.1, in Mechanics and Applied Mathematics. 2004, Dept. of Math. University of Oslo,.

[45] Schitter, G., K.J. Aström, B. DeMartini, P.J. Thurner, K.L. Turner, and P.K. Hansma. IEEE Transactions on Control Systems Technology Special Issue: Dynamics and Control of Micro- and Nano-scale Systems, 2007. in press.

[46] Ando, T., N. Kodera, E. Takai, D. Maruyama, K. Saito, and A. Toda. Proceedings of the National Academy of Sciences of the United States of America, 2001. 98(22): p. 12468-12472.

[47] Hansma, P.K., G. Schitter, G.E. Fantner, and C. Prater. Science, 2006. 314(5799): p. 601-602. [48] Khatiwala, C.B., S.R. Peyton, and A.J. Putnam. Am J Physiol Cell Physiol, 2006. 290(6): p. C1640-50.


48

imaging of bone ultrastructure using atomic force microscopy · completely and neither are the...

Documents