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Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)

Microscopic methods to study the structure of scaffolds in bone tissue engineering: a brief review Mazeyar Parvinzadeh Gashti1*, Farbod Alimohammadi2, Jürg Hulliger1, Matthias Burgener1, Hanane Oulevey-Aboulfadl and Gary L. Bowlin3 1

Department of Chemistry and Biochemistry, University of Berne, Freiestrasse 3, CH-3012 Berne, Switzerland Young Researchers Club, Textile Department, Islamic Azad University- South Tehran Branch, Tehran, Iran 3 Department of Biomedical Engineering, School of Engineering, Virginia Commonwealth University, USA

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Bone composition in the body is well known to be a multiphasic, heterogeneous and anisotropic in structure at all scales. There has been an increasing focus in bone tissue engineering because it presents a new approach for bone repair and regeneration. Scaffolds have been applied as structural supports in combination with cells in an attempt to engineer bone tissue. A wide range of biodegradable polymers and fabrication technologies have been applied to bone tissue regeneration, also in effective tissue engineering scaffolds. Microscopy techniques are fundamental tools for the characterization of scaffolds used in solid biomaterials. In this regard, the use of microscopy techniques are critical for defining fiber size distributions, polar alignment, pore size distributions, overall scaffold homogeneity, surface properties and thickness of layers in scaffolds. Different microscopic techniques have been utilized to characterize scaffolds for bone tissue engineering, including scanning electron microscope (SEM), transmission electron microscope (TEM), atomic force microscope (AFM), confocal microscope (CM), fluorescent microscope (FM), microcomputed tomography (microCT), second and third harmonic optical microscopy, scanning pyro- and piezoelectric microscopy. Keywords: Scaffolds; tissue engineering; bone; regeneration; microscopic techniques

1. Introduction Bones are rigid organs acting as mineral reserves for hydroxyapatite, Ca10(PO4)6(OH)2 (HAp). HAp forms composite materials with a wide variety of organic matter (mostly type I collagen and other proteins). Organic parts of bone play an important role in templating the structure of deposited minerals. The study of mineralization in bone is not only important to gain an understanding of mechanisms for creation of mineral-rich tissues in vivo but also for designing of advanced materials for bone tissue repair. Recently, novel nanofibrous collagen scaffolds produced by two main fabrication techniques of electrospinning and thermally-induced phase separation resemble the non-mineral portion of bone extracellular matrices. Incorporation of such structures with synthetic HAp can improve cell adhesion, stem cell differentiation, and bone tissue formation [1]. To understand interactions which may occur between collagen scaffolds and minerals, it is important to gain information on the structure and composition of bone at atomic and macromolecular levels [1, 2]. Surface and bulk techniques prove to be valuable aids to study such interactions. Optical microscopic techniques are the traditional in vitro methods employed for visualization of multi-dimensional and multiparameter data of bone regeneration. Valuable growth of bone tissue has been observed for measuring physical parameters such as concentration, tissue properties and surface area in order to gain temporal insight on biological functions of bone [2]. Many of these methods can be employed for timely monitoring the steps for bone tissue repairing within a defect site. Microscopic methods can be conducted not only to visualize the gross anatomical structure of bone, but also to visualize substructures of minerals depositions and interactions at the molecular level. Thus, the imaging modalities of light and electron microscopes play an important role. Various light as well as fluorescence microscopy techniques allow the visualization of bone tissues [1, 2]. Such traditional optical microscopes are limited to visualize the bone tissue surface for high resolution images because at greater depths light scattering blurs the images. Confocal and non-linear microscopes are less sensitive to scattering so they are more suitable for high resolution records. Other highresolution techniques are also required for a detailed insight into the ultrastructure of the collagen tissue and the surrounding extracellular matrix. Electron microscopy has become one of the most important tools to image, probe, and manipulate on the micro- and nanoscale levels of biological tissues. The most important reason why electron microscopy is so attractive is the fact that samples of interest can be probed and imaged in conditions very close to their natural state, but with the need to dry or freeze them. However, some alternative procedures are required which do not destroy the structure of the bone tissue or cause artifacts by drying and freezing. Recently, combination of microscopy methods such as fluorescent and electron microscopic information have gained a significant impact in the fields of bone tissue engineering for gathering structural information. Apart from these transmission, confocal and electron scanning techniques, new surface active scanning methods relying on the pyroelectric or piezoelectric effect were developed. These instruments allow for the first time to reveal information on the spatial distribution of polarity in tissues [3]. In this review, various microscopic techniques will be presented which are used to reveal characteristics of bone tissues.

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2. Microscopic methods 2.1 Scanning electron microscope (SEM): Different microscopy techniques have been applied to characterize the structures of bone tissue engineering scaffolds [4]. Electron microscopy is a precise tool that can image bio-nanostructures and it can track bio-materials in nanomaterials, and finally measure physical properties and determine composition [5]. SEM and TEM are two main types of electron microscopes, and SEM is one of the most used microscopes in tissue engineering [6]. SEM images surface with a beam of electrons which produce three-dimensional image of a sample, so SEM micrograph is useful for understanding structure of surface [7,8]. SEM is considered in characterization of different biomaterials. In this regard, carbon nanotubes (CNTs)/biopolymer composites are recently evaluated [9, 10]. CNT incorporated biopolymers were applied for preparation of scaffolds and it has been proved that CNTs are useful for reinforcing such polymeric bioscaffolds [11]. Moreover, the CNTs coated collagen sponge had high biocompatibility with bone [12]. As one example of the use of SEM in bone scaffold analysis, nano composite containing CNTs and poly(propylene fumarate) (PPF) were characterized. SEM images in Fig. 1 showed highly porous scaffolds surrounded by thin walls of nanocomposites and well interconnected. It was suggested that CNTs/polymer nanocomposites are proper for in-situ injection and cross-linking [13], SEM images were applied to observe porosity contain pore networks connecting scaffold interiors to surface openings. However, it is difficult to perform quantitative measurements from SEM images and samples cannot be reused due to sectioning and coating during sample preparation.

Fig.1 SEM images of scaffolds incorporate of (A1–4) PPF, (B1–4) ultra-short single-walled carbon nanotube (US-tube) nanocomposite, and (C1–4) dodecylated US-tube (F-US-tube) nanocomposite with different porogen fractions of 75, 80, 85, and 90 vol%.

Electrospinning is an attractive method for fabrication of different polymeric materials in order to prepare advanced tissue engineering scaffolds [14]. Silk fibroin-based fibers were prepared from aqueous silk worm silk (Bombyx mori) solution with poly(ethylene oxide) PEO by electrospinning process. It was shown by SEM that adhesion, spreading, and proliferation of human bone marrow stormal cells have been improved on silk fibroin-based fibers [15]. In a research project done by the Bowlin group, HAp/polydioxanone (PDO) and HAp/poly(glycolide: lactide) composite scaffolds were introduced to increase overall mineral content [16]. It was shown by SEM that a PLGA scaffold has surface or layered deposition. However PDO scaffolds showed mineralization of individual fibers (Fig. 2). This study presented that the SEM method has an important role for designing porous three-dimensional temporary scaffolds for bone repair. Such structures can manipulate cell function and guide of new organ formation. SEM method can be used to study several design criteria for scaffolds such as biocompatibility, biodegradability, porosity, homogeneity, reproducibly and processablity [16].

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Fig. 2 Scanning electron micrographs of electrospun PLGA (top) and PDO (bottom) scaffolds containing 0% and 50% of nanoHAp incubated in ionic simulated body fluid (i-SBF) and revised simulated body fluid (r-SBF) for 2 weeks. These conditions represent respectively the combination of factors that resulted in minimal (A) and maximal (H) scaffold mineralization

In another example of bone scaffold design and analysis, polycaprolactone (PCL) nanofiber coated with biomimetic calcium phosphate achieved a highly bioactive scaffold for bone tissue engineering. Nanofibrous scaffolds based on (PCL) with a bone-like calcium phosphate (CaP) coaing were prepared that had uniform fibrous structure. In this procedure before Cap coating for improving calcium and phosphate ion grafting, a plasma treatment was done to clean and active the PCL surface and then scaffolds were coated by simulated body fluid. The porous and fibrous structure can be observed and the diameter of the fibers increased from 1.5 to 2.2 µm (Fig. 3). After 3h, the fiber diameter continuously increased to 3.2 µm. The surface of the scaffold was fully covered, after 6h of incubation, and the fibrous structure was no longer visible. It seems a 2h incubation creates the most appropriate material for tissue engineering. It was shown that the scaffolds provide pore for cell migration and nutrient exchange and the coating process improved hydrophilic properties as compared to the uncoated scaffold. The SEM images were applied to gauge the surface structure and appropriate mineralization period [17].

Fig. 3 SEM of the electrospun scaffolds after different mineralization period; a)1h; b) 2h; c) 3h; d) 6h; and e) 7day.

A research was carried out by Ciapetti et al. to study osteoblast-like Saos-2 cells growth in porous PCL matrices for bone repair. In this regard, SEM imaging showed timely steps of the spreading and growth of HAp and cells onto matrices. Intercellular connections were maintained through cytoplasmic elongations. During 3 weeks of cell culture, polygonal cells were covering the surface of the micro/macroporous PCL, as well as the HA-added microporous polymer (Fig. 4a). After 4 weeks of culture, the surfaces of the PCL polymers were fully covered by osteoblast-like cells and Hap (Fig. 4b). It was suggested that the addition of HAp particles has proven useful to promote bone formation by cells covering HAp-added PCL samples [18].

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Fig. 4 SEM images form saos-2 cells spreading onto HA-added microporous PCL: (a) Cells are polygonal and cover most of the polymer surface ( bar=100 μm) at 3 weeks, (b) Cytoplasmic extensions going deeper in the pores of the polymeric matrix (asterisks) after 4 weeks: (bar=10 μm).

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2.2 Transmission electron microscope (TEM): TEM images have been widely applied for specifying both size and morphology of synthesized nanoparticles for tissue engineering applications [19, 20]. HAp is the major inorganic compound that is found for repairing hard tissues in human body and has been widely characterized by TEM [21, 22]. It was reported that polymer incorporation nanoHAp can improve the activity and viability of cells cultured on them so nanoHAp has been widely used in bone tissue engineering [23]. Many organic and inorganic composites, including collagen– HAp [24], gelatine–HAp [25], chitosan–HAp, [25] and Polylactide–HAp [26] composites have been reported that have significant advantages over pure HA. In one study, porous bone regeneration composite scaffolds with surface-immobilized nanoHAp were promoted. In this procedure, surface colloidal nanoHAp with carbonyl groups was fabricated through in-situ polyvinyl pyrrolidone (PVP)-grafting synthesis and then scaffolds were prepared by special freezing and lyoplizing of PVP-grafted nanoHAp and chitosan mixture. Therefore, modified nanoHAp had effective colloidal stability and surface reactivity with chitosan [27]. Moreover, nano fibers incorporating nanoHAp were prepared for bone tissue engineering [28-30]. NanoHAp crystals resembling the shape and chemical composition of natural bone minerals were prepared in the presence of poly(ethylene glycol) (PEG) with different molecular weight by hydrothermal method. Transmission electron microscopy (TEM) showed that the presence of PEG increased the size of nanoHAp crystals and PEG improved the growth of nanoHAp crystals and also increased the crystallinity as shown in Fig. 5 [31]. Moreover with the increase of PEG molecular weight, the crystals grow was enhanced. Thus, it illustrates the usefulness of TEM to investigate both size and morphology of synthesized nano particles for tissue engineering applications. On the other hand, highresolution transmission electron microscopy (HRTEM) makes possible the imaging of the crystallographic structure of a sample at an atomic scale.

Fig. 5 TEM images of the nanoHAp crystals; a) control; b) PEG400; c) PEG 6000; d) PEG 20000, the length distribution of the samples is shown in the top right corner. HRTEM results; Ⅰ) control and Ⅱ) PEG 20000, FFT patterns of the HRTEM fringes using the Digital Micrograph software is shown in the top right corner

In terms of the scaffold incorporating HAp, TEM images can be used for specifying the diameter of the nanofibers and dispersion of the nanoparticles within [32, 33]. Fig. 6 shows TEM images of the poly(L-lactide) (PLLA) nano fibers containing nanoHAp particle produced by direct deposition of electrospun fibers onto a copper grid coated with carbon film. It was shown that HAp particles were well dispersed with almost perfect axial orientation through the fibers. Thus, the experimental confirmed stress results and importance of this technique [33].

Fig. 6 The TEM images of electrospun fibers contain; a) nanoHAp and b) HAp particles.

In another study, TEM images were applied to investigate biomineralization of nanofibrous PLLA scaffolds. In this procedure, nano-fibrous PLLA scaffolds were seed with osteoblastic cells, cultured for 14 days, and rinsed in

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phosphate-buffered saline (PBS) and embedded in polymer matrix. Although mineralization deposition was not seen on osteoblast-seeded solid-walled scaffolds within this culture period time, large amount of small globular mineral deposits added to nano fibers (Fig. 7). It was observed that the mineralization in nano-fibrous scaffolds could be physiologic, not dystrophic [34]. Thus, the importance of the use of TEM to characterize bone scaffolds is represented.

Fig. 7 TEM images of solid-walled, a) nano-fibrous PLLA scaffolds (×5000), b-d) murine calvarial osteoblasts cells cultured after 14 days (×12,000), E) NIH3T3 cells cultured for 14 days, F) scaffolds immersed in culture medium in the absence of cells after 14 days.

2.3 Atomic force microscope (AFM): AFM allows measuring surface topography with a sharp probing tip and can provide three-dimensional high-resolution images of samples under a variety of environmental conditions [33-35]. AFM has made significant contribution in the biomaterials surface characterization and provides information about the surfaces and interfaces at nano scale and is usually used to corroborate observations made by SEM and TEM. AFM has found various applications to surface texture without previous coating of the specimens and also is an effective tool to gain an improved cognition of the structure and property of biological tissues [36, 37]. As an example, the structural and mechanical heterogeneity of dentin at different scale were analyzed by AFM images. It was reported that all demineralization protocols exposed the gap and overlap zones of dentin collagen fibrils [38]. Compared to SEM, several advantages of AFM compared to SEM are: the ability to image in an air or solution environment with little or no sample preparation or fixation, the capacity for direct quantification of image features, such as structural heights and depths, surface hardness or elasticity and measuring the interactions in tissue by the tip used. In terms of bone tissue engineering scaffolds, AFM has been utilized to scan the surface of PLGA and PLGA/collagen electrospun scaffolds and also has been investigated the surface of mineralized nanofibers using an alternate calcium-phosphate dipping method. AFM images in Fig. 8 indicated nanotexturing fibers after forming nanoHAp during mineralization. The nanoHAp formed during mineralization was estimated approximately 50-70nm diameter by AFM. Thus, AFM provide a very high resolution and three dimensional images of surface which allows providing topographical scan of surface [39].

Fig. 8 AFM images of mineralized fibers; (a) 3D surface topography of PLGA, (b) 3D surface topography of PLGA/Col, (c) 3D surface topography PLGA+nanoHAp and (d) 3D surface topography PLGA/Col+nqnoHAp.

Recently, direct investigation of the 3D nucleation and growth of the HAp crystal faces in pseudo-physiological solutions becomes possible by AFM. The growth mechanism of crystals and step velocities can be measured by determining the inclusion of a growth unit at the step front. AFM showed the growth rate of HAp to be 1 or 2 orders of magnitude larger than that of the a-face (in agreement with the elongation of the crystal in the [0 0 0 1] direction as observed in bone matrix and tooth enamel). AFM technique is also able to show reduction of lattice-mismatch strain as growth progressed and electrical charge build up at the surface (Fig.9) [40].

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Fig. 9 AFM images showing nucleation and growth of hydroxyapatite from pseudo-physiological solution: (a) HA seed surface prior to growth; (b) after 10 min in solution; (c) after 2 h; (d) after 24 h.

2.4 Laser scanning confocal microscope (LSCM): LSCM is as useful tool in bone tissue engineering scaffold characterization because it allows one to obtain high resolution optical images with depth discrimination in three-dimensions (scaffolds and regenerating tissue) and has become widely established as a research instruments [41, 42]. Furthermore, the use of LSCM allows for the detail examination of cell colonization of scaffolds, tissue regeneration, and capillary formation within the structures. LSCM is also able to collect simultaneously multiple images in digital form from serial sections of thick tissue specimens. All these are critical aspects to be monitored during tissue regeneration/development. As one example, LSCM was use for investigating the dispersion of nanoHAp particles in nanofibers, in this study PLGA/nanoHAp composite were stained with calcein solution and imaged. Fig. 10 shows the calcein stained nano composite scaffold with a fluorescent green color. It indicated that nanoHAp has a good dispersion in fibers without any visible agglomeration within the resolution of the microscope [43]. Fig. 10 LSCM images of nanofibers showing dispersion of nanoHAp.

In one study, extracellular matrix were simulated by combining nano and micro fibers scaffolds that allowed for a nano-network to promote cell adhesion, and a micro-fiber mesh that provides the mechanical integrity. In their study, they suggested that the nano/micro fiber scaffold might be a solution for the formation of vascularized bone. In their procedure, the micro-fibers were fabricated by a fiber bonding methodology using PCL and then the scaffolds were impregnated with nano fibers by electrospinning. Also PCL fiber scaffold without the nano fibers were prepared as control. Additionally, the scaffolds were coated with a fibronectin and seeded with endothelial cells (ECs) with the cellular interaction investigated by CLSM after 3 and 7 days. Fig. 11 shows the cell growth of human dermal microvascular EC (HDMEC) and human umbilical vein EC (HUVEC) on the nano/micro fibers and micro fibers (control). The growth of cells were seen on both micro and nano fibers. After 7 days, nearly full growth on the surface area of the scaffolds was seen. HDMECs were observed on both micro and nano fibers and remained viable as confirmation by the ability to convert Calcein into a green fluorescent compound. On the scaffolds without the nanonetwork, the cells were seen on the micro fibers but no cells were observed to span among the fibers [44].

Fig. 11 CLSM images of scaffolds; a,b,d,e) viable HDMECs; c,f) viable HUVECs. Left column contains nano/micro fiber composites and right column contains micro fiber scaffolds.

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2.5 Fluorescent microscope (FM): FM is one of the common utilized techniques to enhance imaging capabilities in bone tissue research that is not possible with other traditional optical microscopes. This method enables one to visualize the fluorescent stained tissues against non-stained background/scaffold to observe inhomogeneity of tissue that is not identifiable by standard imaging methods. This method is utilized in conjunction with any material that absorbs excitation light at a certain wavelength and emits light at a longer wavelength. The emitted light is then recorded as a photographic image, video, fluorescence decay trace, or as “photo multiplier tube” signals from bone tissue that is displayed and analyzed. The aim of this method is to bind fluorescent stains to biomolecules which the distribution can be accurately identified [45]. As one example of this technique, Kruyt et al. investigated the potential of tissue engineered bone to heal a critical sized defect in the goat. Tissue engineered constructs were prepared from goat bone marrow stromal cells and porous biphasic calcium phosphate ceramic scaffolds. Fluorescent microscopy indicated that bone was growing away from the scaffold surface and confirmed that no bone had formed in the middle of the defect in the first 9 weeks (Fig. 12) [46]. One of the major limitations of this method is the inability of visible light to pass well through the bone tissue. In this regard the distances traveled in tissue by the probe should be small (micrometers in length) and the bone tissues grown in laboratory cultures need to have the thickness from 1 to 20 microns, to be viewed with a fluorescence light microscope.

Fig. 12 High magnification fluorescence microscopy of a peripheral area of the implant (a). Bone formation originated before 3 weeks on the outer side and inner side of the ceramic, as indicated by the Calcein green (b, 3 week) label. Fusion between surrounding bone and the scaffold was accomplished around 5 weeks as indicated by the yellow OTC label (bar = 50 μm). (c) High magnification fluorescence microscopy of a more central part. Only the Xylenol orange (X, 7 weeks) was present close to the BCP scaffold (bar = 50 μm).

2.6 Microcomputed tomography (microCT): The microCT is an X-ray technique showing high-resolution assessments of density, geometry and 3D microarchitecture of mineralized tissues, such as bones and teeth, calcification as result of pathology, or soft tissues and biomaterials stained with radiographic contrast media. This method may improve our ability to estimate the quality of newly formed bone. Other applications include fracture healing, skeletal phenotyping, and segmental bone defect repair, age-related effects on bone, joint degeneration, vascular remodeling, developmental biology, and microstructure of materials. By this method, users are able to extract quantitative morphometric and density data along with threedimensional models of their specimens [47]. In this regard, Kanczler et al. used microCT to examine the temporal modulation of embryonic skeletal development in a three dimensional framework. Isolated embryonic chick femurs were organotypic (air/liquid interface) cultured for 10 days in either basal, chondrogenic, or osteogenic supplemented culture conditions. The growth development and modulating as well as the structural morphology effects in embryonic bone were investigated using microCT (Fig. 13). Significant differences in the structural architecture of the embryonic bones were observed between basal, chondrogenic, and osteogenic organotypic cultures over a 10 day culture period. MicroCT showed that embryonic chick femurs cultured in chondrogenic conditions are shorter in stature with a thicker diaphysis and an enlarged epiphyseal region compared with basal cultured femurs. In the osteogenic culture conditions embryonic chick femurs were comparable to the basal cultured femurs; however, the levels of bone mineralization were increased compared with the embryonic bones cultured in basal and chondrogenic conditions. It was presented that this multidimensional technique has the potential to yield integral information on bone development for therapeutic strategies for skeletal regenerative technology and the possible understanding for bone diseases [48].

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Fig. 13 microCT analysis of organotypic cultured embryonic femurs in basal, chondrogenic, and osteogenic conditions.

Harry van Lenthe et al. reviewed the application of time-lapsed microCT technique for monitoring the spatial and temporal mineralization in individual scaffolds cultured in bioreactors. They monitored the formation of bone-like tissue from human mesenchymal stem cells (hMSC) cultured in osteogenic medium on SF scaffolds over a 44-day period (Fig. 14). MicroCT imaging allowed to quantify bone volume density, bone surface density, trabecular thickness and spacing. It was shown that bone volume is steadily increasing over which indicated the importance of longitudinal studies, rather than cross-sectional ones [49].

Fig. 14 Time-lapsed μCT images demonstrating bone-like tissue formation in two silk fibroin scaffolds seeded with human mesenchymal stem cells. Full top view and side view on the center part of the disk-shaped constructs are displayed.

In an article, Bouxsein et al. reviewed recent applications of microCT for characterization of bone microstructure. It was mentioned that high-resolution microCT imaging can assess trabecular bone morphology (Fig. 15). They provided the guidelines recommendations regarding standardized terminology and units in microCT technique, information to be included in describing the methods for a given experiment, and a minimal set of outcome variables that should be reported by this method. With this regard, they mentioned the minimal set of variables that should be used to describe trabecular bone morphometry include bone volume fraction and trabecular number, thickness, and separation. The minimal set of variables that should be used to describe cortical bone morphometry was recommended as total crosssectional area, cortical bone area, cortical bone area fraction, and cortical thickness. This review demonstrated several advantages of microCT including the excellent reproducibility and accuracy of bone morphology measurement, direct 3D measurement of trabecular thickness and separation, significantly faster and larger volume measurement compared to typical histologic analyses, nondestructive assessment of bone morphology and, finally estimation of bone tissue mineralization by comparing with that of hydroxyapatite standards. However, the major limits on application of microCT, are direct measurement of bone micro-architecture without relying on stereologic models, inadequate resolution of mCT images relative to the trabecular size, the use of a plate model to estimate trabecular thickness in 2D histomorphometry versus direct 3D methods in microCT and poor threshold selection of bone tissue [50].

Fig. 15 MicroCT image used for calculating trabecular thickness (A) and separation (B). 3D distances are computed by fitting spheres inside the structure (ie, to assess averagetrabecular thickness) or inside the background (marrow space, ie, to assess average trabecular separation). The average diameter of the spheres represents the object thickness, and the standard deviation of the diameter represents the variability in the object thickness

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2.7 Pyroelectric scanning microscope (SPEM): The existence of polar properties in different biological materials is known for nearly 50 years [51]. These properties stems from structural ordering of biopolymers. Materials containing a permanent polarization have the ability for pyroelectrical-, piezoelectrical- and SHG-effects. HAp crystallizes in centrosymmetric space group P63/m and therefore shows no pyroelectric, piezoelectric or SHG signal. In contrast, partially polar alignment of collagen fibers occurring in calcified structures such as bones or teeth belong to the texture group ∞ [52] being pyro-, piezo- and SHG-active. A first theoretical model [53] explains polar alignment of fibrils by a Markov chain, due to selective prolongation reactions of fibrils. In the following section three microscopic techniques are presented using these physical effects for mapping polarity in biological tissues. SPEM has proved to detect the permanent polarization in organic materials [54]. In SPEM a sample is placed into a capacitor. A modulated laser beam heats up a small area and causes a local electrical displacement D. The change in spontaneous polarization Ps is compensated within the capacitor. The resulting discharge current is described as [55]

where Q is the charge flowing between the electrodes and the heated area A. Considering a constant pyroelectric coefficient for a small temperature change dT, the discharge current is proportional to the net polarization of the heated area A. This current is typically in the order of femto to pico ampere depending on the magnitude of the pyroelectric coefficient. For a human femur the pyroelectric coefficient is 3.6·10-13 C cm-2 K-1 [52]. By scanning the surface, a 2D map of polarization distribution is obtained. The direction of the discharge current contains information about the absolute orientation of polarization. Recently, a new approach using an interdigitated comb-electrode (IDE) has improved the sensitivity of SPEM allowing to measure low displacement currents of biological tissues [56]. Fig. 16 shows the polarization distribution of the cross section of a tooth recorded by SPEM. The difference between the enamel and dentine can be attributed to a higher content of parallel collagen in the dentil section. b)

a)

Fig 16. (a) Polarization amplitude map of the radial cross section of a human tooth. The polarization distribution map shows several peaks which lie more or less on a circle. These peaks cover regions of dentine surrounded by non-polar enamel [56]. (b) Schematic view of an IDE electrode. The dashed red line indicates the radial cross section of a sample. The pyroelectric current is measured between electrodes (black lines).

SPEM provides typically a resolution of ~1 μm [55]. Only a thin layer of the surface region is measured. By changing the modulation frequency the penetration depth of heat energy can be adjusted according to [55]:

where ω is the modulation frequency and α is the thermal diffusion coefficient. By measuring at different modulation frequencies information in the direction normal to surfaces can be obtained. 2.8 Piezoresponse force microscope (PFM): In the last decade the piezoresponse force technique has been developed to a powerful tool to reveal the structure of biological tissues at submicron level. Piezoelectricity describes the connection between an external electric field and an induced material strain. In PFM a scanning probe tip is in contact with the substrate. An applied electrical bias Vtip = Vdc + Vac cos(ωt) between tip and bottom electrode induces periodic displacement of the surface in form of d = d1ω cos(ωt + φ) [57]. The deformation is recorded by the scanning probe tip, whereby the amplitude and phase of the

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deflection allows determining the strength of a piezoresponse and the absolute orientation of polarization. PFM has an estimated spatial resolution of about 5 nm to distinguish two different piezoelectric domains [58]. The potential of PFM has been demonstrated with success for the investigation of teeth samples [58]. Fig 17 b shows the piezoelectric strength and Fig. 17 c the phase of a chemically treated dentin sample. Phase contrast from bright to dark reveals random orientation of collagen fibers.

Fig. 17. Visualization of a pattern of collagen fibrils in a chemically treated dentin sample: (a) topography, (b) PFM amplitude, and (c) PFM phase. Collagen fibrils visible in the topographic image appear as elongated bright regions in the PFM amplitude image and as bright and dark regions in the PFM phase image [58].

In PFM measurements also lateral deformations are detectable because in-plane polarization components causes shear forces leading to torsion oscillations of a cantilever [57-59]. Combination of vertical (V) and lateral (L) PFM (VPFM and LPFM, respectively) can provide insight to the 3D piezoelectricity environment in thin layers. Conditions and limitations of the method are discussed elsewhere [60]. As collagen fibrils show a rotation axis, measurement of the vertical and lateral deformation suffices to reveal their spatial orientation [57]. PFM constitute an interesting tool for the exploration of collagen alignment in biological tissues. This also opens up a suitable way to study arrangements of fibrils in HAp. Fig 18 shows VPFM and LPFM measurements performed at the dentil-enamel junction (DEJ) [61]. The piezoelectric response can be attributed to protein inclusion in the DEJ region. The in-plane and out-plane electromechanical response vector can be identified in different piezoelectric domains showing a complex collagen alignment in enamel.

Fig. 18. a) vertical PFM and b) lateral PFM images of protein inclusions in tooth enamel. Blue and red markers indicates the orientation of the electromechanical response vector, while the intensity provides its magnitude (maximum is 7.5 pm/V) [61].

2.9 Second harmonic generation microscope (SHGM): Second harmonic generation microscopy (SHGM) is a nonlinear technique. SHG involves the conversion of an intense laser pulse of frequency ω in its harmonic 2ω. At the molecular level, this response is related to the presence of a hyperpolarizable electronic system. Coherent construction of the signal requires the absence of centrosymmetry at the macroscopic level [62]. In practice, SHG is usually studied for molecules possessing a permanent dipole moment which align in an acentric fashion. The wave is generated though the second order nonlinear polarization PNL:

where is the second order nonlinear susceptibility tensor setting the strength of the SHG effect and is the electric field of the fundamental wave. is the induced SHG polarization generating radiation at twice the angular frequency . SHG microscopy was first demonstrated for the study of nonlinear crystals [63], surfaces/interfaces [64], field distribution in semiconductors [65,66], and was then applied to biological systems. SHG responses have already been observed as early as 1980 for fibrillar collagen [65,67]. Since then, SHG microscopy has increasingly been used as an emerging microscopic technique for a wide range of biological and clinical imaging tool. SHG can be of further help to

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investigate fundamental biological processes. Strong SH signals are reported for muscles [68], skin [69], teeth [70], bones [71,72], blood vessels [73] and tendon [74]. Collagen molecules play an important role to determine the morphology and functional properties of tissues and organs. The collagen fibrils in connective tissues or the actomyosin complexes in muscles are sufficiently structured and ordered to generate SHG responses. That permits SHG microscopy to image a variety of tissue structures. It was suggested by Fukada and Yasuda [75] that tendon has an overall macroscopic polarity, which can only arise from acentric molecular arrangements maintained over macroscopic distances. Due to the fact that SHG signals arise from an induced polarization rather than from absorption, SHG provides direct information about the collagen structure in tissues without photobleaching and photoxicity due to additional staining [76]. This technique employs pulsed lasers in the infrared wavelength range, permitting deep optical penetration of several hundred microns with minimal tissue damage and little signal loss [77]. These are major advantages of the SHG method. SHG microscopy is suited as well for determination of the overall collagen fiber orientation in biological tissues [78]. The SHG light induced by collagen molecules is strong when the polarization direction of the incident light is parallel to the longitudinal direction of collagen fibers. However, the SHG light almost disappears if the laser polarization is perpendicular to the collagen orientation. By taking advantage of these characteristics, SHG microscopy has been applied for example to visualize detailed structures of dermal collagen fibers in skin (Fig. 19) [79]. SHG imaging of porcine ear cartilage [80] and veal muscle [81] has been reported showing in both cases oriented collagen fibers (Fig. 20). Many diseases are characterized by an abnormally organized or defect fibrillar assembly of collagen relative to normal tissue [82]. For example connective tissue disorders, musculoskeletal disorder, cancers that are characterized by changes in the collagen structure (increase of collagen amount or altered fibril orientation in malignant tissues), bone fractures, progression of diabetic complications, abnormalities in wound healing, organ fibrosis and many other pathologies related to collagen-comprised tissues can be investigated by this optical technique. Furthermore, it may serve useful as a diagnostic tool for providing real time spatial guidance for diseases and biopsies or monitoring changes associated with disease progression.

Fig. 19: SHG images of collagen dermis of human keloids in a) the superficial dermis: Collagen fibrils are organized in a parallel manner. b) the medial dermis: Collagen fibrils are randomly oriented. c) the deep dermis: Collagen fibrils are highly oriented in one direction [79].

Fig. 20: Image of the SHG response of a) porcine tissue from ear cartilage [80], b) and c) muscle tissue from a veal escalope [81].

Application of phase sensitive second harmonic generation microscopy [83,84] to tissues may open up the possibilities to determine the absolute polarity of tissue in order to understand the fundamental mechanism leading to a polar matter in nature in relation to its biological function (Fig. 21). In particular the effect of a disease on the polarity distribution may be relevant for healing.

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Fig. 21: Phase sensitive second harmonic generation microscopy image of human cementum. a) Optical image illuminated by day light. b) SHG image with polarization parallel (arrow) to the fibril axis. c) and d) phase sensitive responses demonstrate that aligned fibrils can show a bi-polar structure in tissues. (H. Aboulfadl, J. Hulliger to be published).

4. Future Trends: Using microscopy techniques to study the structure of scaffold in bone tissue engineering opens up several perspectives: 1. Upgrading of microscopic instruments for better evaluation of scaffolds. 2. Application of new materials for bone tissue generation in order to make higher qualified images. 3. Developing new types of microscopes with combination of two or more techniques. Therefore, the progress in microscopic tools can overshadow bone tissue scaffolds which may provide the door to observation of different structural and morphological phenomena in bone tissue repairing.

References [1] Puppi D, Chiellini F, Piras AM, Chiellini E. Polymeric materials for bone and cartilage repair. Progress in Polymer Science. 2010;35:403–440. [2] Holzwartha JM, Maa PX. Biomimetic nanofibrous scaffolds for bone tissue engineering. Biomaterials. 2011;32:9622-9629. [3] Armentano I, Dottori M, Fortunati E, Mattioli S, Kenny JM. Biodegradable polymer matrix nanocomposites for tissue engineering: A review. Polymer Degradation and Stability. 2010;95:2126-2146. [4] Marx KA. Introduction: Enabling Nanomanufacturing Using Microscopy Techniques. Microscopy Research and Technique. 2007;70:493–496. [5] Mao C. Nanomaterials Characterization: Structures, Compositions, and Properties. Microscopy Research and Technique. 2006;69:519–521. [6] Cruthers N, Carr D, Niven B, Girvan E, Laing R. Methods For Characterizing Plant Fibers. Microscopy Research and Technique. 2005;67:260–264. [7] Mao C. Introduction: Nanomaterials Characterization Using Microscopy. Microscopy Research and Technique. 2004;64:345– 346. [8] Mao C, Introduction: Bio and Nano Imaging and Analysis. Microscopy Research and Technique. 2011;74:559–562. [9] Alimohammadi F, Parvinzadeh M, shamei A. Carbon Nanotube Embedded Textiles. US-Patent 2011, 0171413. [10] Alimohammadi F, Parvinzadeh Gashti M, Shamei A. A novel method for coating of carbon nanotube on cellulose fiber using 1,2,3,4-butanetetracarboxylic acid as a cross-linking agent. Progress in Organic Coatings. doi:10.1016/j.porgcoat.2012.01.012. [11] Sitharaman B, Shi X, Walboomers XF, Liao H, Cuijpers V, Wilson LJ, Mikos AG, Jansen JA. In vivo biocompatibility of ultrashort single-walled carbon nanotube/biodegradable polymer nanocomposites for bone tissue engineering. Bone. 2008;43:362– 370. [12] Hirata E, Uo M, Takita H, Akasaka T, Watari F, Yokoyama A. Multiwalled carbon nanotube-coating of 3D collagen scaffolds for bone tissue engineering. Carbon. 2011;49:3284–3291. [13] Shi X, Sitharaman B, Pham QP, Liang F, Wu K, Billups WE, Wilson LJ, Mikos AG. Fabrication of porous ultra-short singlewalled carbon nanotube nanocomposite scaffolds for bone tissue engineering. Biomaterials. 2007;28:4078–4090. [14] Jang JH, Castano O, Kim HW. Electrospun materials as potential platforms for bone tissue engineering. Advanced Drug Delivery Reviews. 2009;61:1065–1083. [15] Jin HJ, Chen J, Karageorgiou V, Altman GH, Kaplan DL. Human bone marrow stromal cell responses on electrospun silk fibroin mats. Biomaterials. 2004;25:1039–1047. [16] Madurantakam PA, Rodriguez IA, Cost CP, Viswanathan R, Simpson DG, Beckman MJ, Moon PC, Bowlin GL. Multiple factor interactions in biomimetic mineralization of electrospun scaffolds. Biomaterials. 2009;30:5456–5464. [17] Yang F, Wolke JGC, Jansen JA. Biomimetic calcium phosphate coating on electrospun poly (e-caprolactone) scaffolds for bone tissue engineering. Chemical Engineering Journal. 2008;137:154–161. [18] Ciapetti G, Ambrosio L, Savarino L, Granchi D, Cenni E, Baldini N, Pagani S, Guizzardi S, Causa F, Giunti A. Osteoblast growth and function in porous poly ε-caprolactone matrices for bone repair: a preliminary study. Biomaterials. 2003 ;24:3815– 382.

© 2012 FORMATEX

636

Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)

[19] Miyauchi T, Yamada M, Yamamoto A, Iwasa F, Suzawa T, Kamijo R, Baba K, Ogawa T. The enhanced characteristics of osteoblast adhesion to photofunctionalized nanoscale TiO2 layers on biomaterials surfaces. Biomaterials. 2010;31:3827–3839. [20] Jurczyk MU, Jurczyk K, Miklaszewski A, Jurczyk M. Nanostructured titanium-45S5 Bioglass scaffold composites for medical applications. Materials and Design. 2011;32:4882–4889. [21] Considine GD. Encyclopedia of Chemistry. 5th ed. John Wiley and Sons press: 2005. [22] Holzwarth JM, Ma PX. Biomimetic nanofibrous scaffolds for bone tissue engineering. Biomaterials. 2011;32:9622-9629. [23] Li J, Dou Y, Yang J, Yin Y, Zhang H, Yao F, Wang H, Yao K. Surface characterization and biocompatibility of micro- and nano-hydroxyapatite/chitosan-gelatin network films. Materials Science and Engineering C. 2009;29:1207–1215. [24] Li X, Feng Q, Cui F. In vitro degradation of porous nano-hydroxyapatite/collagen/PLLA scaffold reinforced by chitin fibres. Materials Science and Engineering C. 2006;26:716 – 720. [25] Li J, Chen YP, Yin Y, Yao F, Yao K. Modulation of nano-hydroxyapatite size via formation on chitosan–gelatin network film in situ. Biomaterials. 2007;28:781–790. [26] Kim SS, Parkc MS, Jeonc O, Choi CY, Kima BS. Poly(lactide-co-glycolide)/hydroxyapatite composite scaffolds for bone tissue engineering. Biomaterials. 2006;27:1399–1409. [27] Song Z, Yin Z, Li C, Yang Z, Ning C, Zhou D, Wang R, Xu Y, Qiu J. Efficient and toxicity-free surface immobilization of nano-hydroxyapatite for bone-regenerative composite scaffolds by grafting polyvinyl pyrrolidone. Materials Science and Engineering C. 2012; doi:10.1016/j.msec.2012.02.025. [28] Peng F, Yu X, Wei M. In vitro cell performance on hydroxyapatite particles/poly(L-lactic acid) nanofibrous scaffolds with an excellent particle along nanofiber orientation. Acta Biomaterialia. 2011;7:2585–2592. [29] Wan YZ, Hong L, Jia SR, Huang Y, Zhu Y, Wang YL, Jiang HJ. Synthesis and characterization of hydroxyapatite–bacterial cellulose nanocomposites. Composites Science and Technology. 2006;66:1825–1832. [30] Mavis B, Demirtas TT, sderelioglu MG, Gunduz G, Colak U. Synthesis, characterization and osteoblastic activity of polycaprolactone nanofibers coated with biomimetic calcium phosphate. Acta Biomaterialia. 2009;5:3098–3111. [31] Zhang H, Liu M, Fan H, Zhang X. Carbonated Nano Hydroxyapatite Crystal Growth Modulated by Poly(ethylene glycol) with Different Molecular Weights. Crystal Growth & Design. dx.doi.org/10.1021/cg200917y. [32] Son WK, Youk JH, Park WH. Antimicrobial cellulose acetate nanofibers containing silver nanoparticles. Carbohydrate Polymers. 2006;65:430–434. [33] Peng F, Yu X, Wei M. In vitro cell performance on hydroxyapatite particles/poly(L-lactic acid) nanofibrous scaffolds with an excellent particle along nanofiber orientation. Acta Biomaterialia. 2011;7:2585–2592. [34] Woo KM, Jun JH, Chen VJ, Seo J, Baek JH, Ryoo HM, Kim GS, Somerman MJ, Ma PX. Nano-fibrous scaffolding promotes osteoblast differentiation and biomineralization. Biomaterials. 2007;28:335–343. [35] Thrrien J, Dindar A, Smith D. AFM Studies of Nanoparticle Deposition via Electrospray Ionization. Microscopy Research and Technique. 2007;70:530–533. [36] Greene ME, Kinser C, Kramer DE, Pingree LSC, Hersam MC. Application of Scanning Probe Microscopy to the Characterization and Fabrication of Hybrid Nanomaterials. Microscopy Research and Technique. 2004;64:415–434. [37] Wagner M, Kaehler D, Anhenn O, Betz T, Yawad S, Shamaa A, Theegarten D, Linder R. Nanostructural Analysis by Atomic Force Microscopy Followed by Light Microscopy on the Same Archival Slide. Microscopy Research and Technique. 2009;72:471–481. [38] Bertassoni LE, Marshall GW, Swain MV. Mechanical heterogeneity of dentin at different length scales as determined by AFM phase contrast. Micron. dx.doi.org/10.1016/j.micron.2012.03.021. [39] Ngiam M, Liao S, Patil AJ, Cheng Z, Chan CK, Ramakrishna S. The fabrication of nano-hydroxyapatite on PLGA and PLGA/collagen nanofibrous composite scaffolds and their effects in osteoblastic behavior for bone tissue engineering. Bone. 2009;45:4–16. [40] Kanzaki N, Onuma K, Ito A, Teraoka K, Tateishi T, Tsutsumi S. Direct growth rate measurement of hydroxyapatite single crystal by Moire phase shift interferometry. Journal of Physical Chemistry B. 1998;102:6471–6476. [41] Wilson T. Fluorescence imaging modes in fibre-optic based confocal scanning microscopes. Optics Communications. 1993;96:133-141. [42] Heinrich L, Freyria AM, Melin M, Tourneur Y, Maksoud R, Bernengo JC, Hartmann DJ. Confocal Laser Scanning Microscopy Using Dialkylcarbocyanine, Dyes for Cell Tracing in Hard and Soft Biomaterials. Journal of Biomedical Materials Research Part B: Applied Biomaterials. DOI 10.1002/jbmb. [43] Jose MV, Thomas V, Johnson KT, Dean DR, Nyairo E. Aligned PLGA/HA nanofibrous nanocomposite scaffolds for bone tissue engineering. Acta Biomaterialia. 2009;5:305–315. [44] Santos MI, Tuzlakoglu K, Fuchs S, Gomes ME, Peters K, Unger RE, Piskin E, Reis RL, Kirkpatrick CJ. Endothelial cell colonization and angiogenic potential of combined nano- and micro-fibrous scaffolds for bone tissue engineering. Biomaterials. 2008;29: 4306–4313. [45] Martin I, Mastrogiacomo M, Leo GD, Muraglia A, Beltrame F, Cancedda R, Quarto R. Fluorescence Microscopy Imaging of Bone for Automated Histomorphometry. Tissue Engineering. 2002;8:847-852. [46] Kruyt MC, Dhert WJA, Yuan H, Wilson CE, van Blitterswijk CA, Verbout AJ, de Bruijn JD, Bone tissue engineering in a critical size defect compared to ectopic implantations in the goat. Journal of Orthopaedic Research. 2004;22:544–551. [47] Davis GR, Wong FS. X-ray microtomography of bones and teeth. Physiological Measurement. 1996;17:121-146. [48] JM Kanczler, Smith EL, Roberts CA, Oreffo ROC. A Novel Approach for Studying the Temporal Modulation of Embryonic Skeletal Development Using Organotypic Bone Cultures and Microcomputed Tomography. Tissue Engineering: C. 2012;18:114. [49] van Lenthe GH, Hagenmüller H, Bohner M, SJ Hollister, L Meinel, Müller R. Nondestructive micro-computed tomography for biological imaging and quantification of scaffold–bone interaction in vivo. Biomaterials. 2007;28:2479–2490. [50] Bouxsein ML, Boyd SK, Christiansen BA, Guldberg RE, Jepsen KJ, Muller R. Guidelines for Assessment of Bone Microstructure in Rodents Using Micro–Computed Tomography. Journal of Bone and Mineral Research. 2010;25:1468–1486.

© 2012 FORMATEX

637

Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)

[51] Athenstaedt H. Spontaneous polarization and pyroelectric behavior of organisms. Ferroelectrics. 1987;73:455-466. [52] Lang S B. Pyroelectric effect in bone and tendon. Nature. 1966;212:704-705. [53] Hulliger J. Connective tissue polarity unraveled by a Markov-chain mechanism of collagen fibril segment self-assembly. Biophysical Journal. 2003;84:3501-3507. [54] Quintel A, Hulliger J, Wübbenhorst M. Analysis of the polarization distribution in a polar perhydrotriphenylene inclusion compound by scanning pyroelectric microscopy. Journal of physical chemistry B. 1998;102:4277-4283. [55] Batagiannis A, Wübbenhorst M, Hulliger J. Piezo- and pyroelectric microscopy. Current Opinion in Solid State and Materials Science. 2010;14:107-115. [56] Wübbenhorst M, Putzeys T, Burgener M, Hulliger J. Observation of permanent polarization distributions in bioorganic materials using a highly sensitive scanning pyroelectric microscope. Proceedings 14th International Symposium on Electrets (ISE). 2011;75-76. [57] Kalinin S V, Rodriguez B J, Shin J, Jesse S, Grichko V, Thundat T, Baddorf A P, Gruverman A. Bioelectromechanical imaging by scanning probe microscopy: Galvani`s experiment at the nanoscale. Ultramicroscopy. 2006;106:334-340. [58] Gruverman A, Wu D, Rodriguez B J, Kalinin S V, Habelitz S. High-resolution imaging of proteins in human teeth by scanning probe microscopy. Biochemical and Biophysical Research Communications. 2007;352:142-146. [59] Rodriguez B J, Gruverman A, Kingon A I, Nemanich R J, Cross J S. Three-dimensional high-resolution reconstruction of polarization in ferroelectric capacitors by piezoresponse force microscopy. Journal of Applied Physics. 2004;95:1958-1962. [60] Kalinin S V, Rodriguez B J, Jesse S, Shin J, Baddorf A P, Gupta P, Jain H, Williams D B, Gruverman A. Vector piezoresponse force microscopy. Microscopy and Microanalysis. 2006;12:206-220. [61] Rodriguez B J, Kalinin S V, Shin J, Jesse S, Grichko V, Thundat T, Baddorf A P, Gruverman A. Electromechanical imaging of biomaterials by scanning probe microscopy. Journal of Structural Biology. 2006;153:151-159. [62] Boyd RW. Nonlinear optics. 2nd ed. San Diego, CA: Academic Press; 2003. [63] Gannaway JN, Sheppard CJR. Second-harmonic imaging in the scanning optical microscope. Optical and Quantum Electronics. 1978;10:435–439. [64] Shen YR. Surface properties probed by second harmonic and sum frequency generation. Nature. 1989;337:519–525. [65] Sun C-K, Chu S-W, Tai S-P, Keller S, Mishra UK, DenBaars SP. Scanning second-harmonic-generation and third-harmonicgeneration microscopy of GaN. Applied Physics Letters. 2000;77:2331–2333. [66] Sun C-K, Chu S-W, Tai S-P, Keller S, Abare A, Mishra UK, et al. Mapping piezoelectric-field distribution in gallium nitride with scanning second-harmonic generation microscopy. Scanning. 2001;23:182–192. [67] Freund I, Deutsch M, Sprecher A. Connective tissue polarity: optical second-harmonic microscopy, crossed-beam summation, and small-angle scattering in rat-tail tendon. Biophysical Journal. 1986;50:693–712. [68] Plotnikov SV, Millard AC, Campagnola PJ, Mohler WA. Characterization of the myosin-based source for second-harmonic generation from muscle sarcomeres. Biophysical Journal. 2006;90:693–703. [69] Zhuo SM, Chen JX, Luo TS, Chen HL, Zhao JJ. High-contrast multimodel nonlinear optical imaging of collagen and elastin. Journal of Physics: Conference Series. 2006, Vol. 48. pp. 1476–1481. [70] Chen M-H, Chen W-L, Sun Y, Fwu PT, Dong C-Y. Multiphoton autofluorescence and second-harmonic generation imaging of the tooth. Journal of Biomedical Optics. 2007;12:064018–1. [71] Ambekar R, Chittenden M, Jasiuk I, Toussaint Jr. KC. Quantitative second-harmonic generation microscopy for imaging porcine cortical bone: Comparison to SEM and its potential to investigate age-related changes. Bone. 2012;50:643–650. [72] Scaglione S, Giannoni P, Bianchini P, Sandri M, Marotta R, Firpo G, et al. Order versus disorder: in vivo bone formation within osteoconductive scaffolds. Scientific Reports. 2012. [73] Zoumi A, Lu X, Kassab GS, Tromberg BJ. Imaging coronary artery microstructure using second-harmonic and two-photon fluorescence microscopy. Biophysical Journal. 2004;87:2778–2786. [74] Roth S, Freund I. Optical second-harmonic scattering in rat-tail tendon. Biopolymers. 1981;20:1271–1290. [75] Fukada E, Yasuda I. Piezoelectric effects in collagen. Japanese Journal of Applied Physics. 1964;3:117–121. [76] Campagnola PJ, Millard AC, Terasaki M, Hoppe PE, Malone CJ, Mohler WA. Three-dimensional high-resolution secondharmonic generation imaging of endogenous structural proteins in biological tissues. Biophysical Journal. 2002;81:493–508. [77] Mohler W, Millard AC, Campagnola PJ. Second harmonic generation imaging of endogenous structural proteins. Methods. 2003;29:97–109. [78] Yasui T, Tohno Y, Araki T. Determination of collagen fiber orientation in human tissue by use of polarization measurement of molecular second-harmonic-generation light. Applied Optics. 2004;43:2861–2867. [79] Yu HB, Chen S, Zhu XQ, Yang HQ, Chen JX. Second harmonic generation imaging of dermal collagen component in human keloid tissue. Journal of Physics: Conference Series. 2011, Vol. 277. p. 012046. [80] Rao RA, Mehta MR, Toussaint, Jr. KC. Fourier transform-second-harmonic generation imaging of biological tissues. Optics Express. 2009;17:14534–14542. [81] Odin C, Guilbert T, Alkilani A, Boryskina OP, Fleury V, Le Grand Y. Collagen and myosin characterization by orientation field second harmonic microscopy. Optics Express. 2008;16:16151–16165. [82] Nadiarnykh O, Plotnikov S, Mohler WA, Kalajzic I, Redford-Badwal D, Campagnola PJ. Second harmonic generation imaging microscopy studies of osteogenesis imperfecta. Journal of Biomedical Optics. 2007;12:051805. [83] Rechsteiner P, Hulliger J, Flörsheimer M. Phase-sensitive second harmonic microscopy reveals bipolar twinning of markov-type molecular crystals. Chemistry of Materials. 2000;12:3296–3300. [84] Aboulfadl H, Hulliger J. One-hundred eighty degree domain detection by surface phase sensitive second harmonic generation microscopy of polar materials. Crystal Growth & Design. 2011;11:3045–3048.

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