Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)
L. Tayebi 1,2 , A. Nozari 1 , D. Vashaee 3 and M. Mozafari 1
1 Helmerich Advanced Technology Research Center, School of Materials Science and Engineering, Oklahoma state
University, Tulsa, OK, USA
2 School of Chemical Engineering, Oklahoma State University, Stillwater, OK, USA
3 Helmerich Advanced Technology Research Center, School of Electrical and Computer Engineering, Oklahoma state
University, Tulsa, OK, USA
Tissue engineering involves three-dimensional (3D) culturing of various cells on 3D micro-structured scaffolds. Imaging the features of cells, scaffolds and the cell-scaffold interfaces require more advanced methods compare to the ordinary two-dimensional cell imaging. Different techniques have been used for this purpose include: phase contrast microscopy, fluorescent microscopy, multiphoton microscopy, confocal laser scanning microscopy, confocal Raman microscopy, magnetic resonance microscopy, optical coherence tomography, scanning electron microscopy stereoimaging and other kinds of electron microscopy. Different microscopy methods illuminate different features of tissue-engineered constructs.
In this review we describe and discuss the application of these complementary microscopy techniques for explanatory imaging of engineered tissues.
Keywords Microscopy; Tissue engineering; Tissue-engineered construct imaging; Three-imentional imaging
Optical imaging has a long and eminent history in biology and medicine. Since the primary observations of the interior structure of living tissues and subsequent discovery of micro-orgasm in 17 th century [1], the microscope has been an indispensable tool in biomedical research. Regenerative medicine or tissue engineering is an emerging multidisciplinary approach involving engineering, biomaterials and medicine. In this approach a biocompatible scaffold provides an adhesive substrate for efficient and healthy regeneration of cells [2].
Tissue engineering materials are a combination of more than one cell type in a three-dimensional (3D) scaffold.
Compare to the ordinary two-dimensional (2D) cell imaging, analyzing the features of cells, scaffolds and the cellscaffold interfaces require more advanced methods due to the these three characteristics of the tissue engineered constructs: 1) opacity of the scaffolds 2) 3D structure of the cell/scaffold organization 3) possible existence of more than one cell type in the unit structure. Also it is partly necessary to analyze the tissue engineered organizations by realtime imaging. The non-destructive and minimally-invasive imaging methodologies are also of importance in tissue engineering and regenerative medicine.
In this chapter we discuss about the different techniques of tissue engineering microscopy include: phase contrast microscopy, fluorescent microscopy, multiphoton microscopy, confocal laser scanning microscopy, confocal Raman microscopy, magnetic resonance microscopy, optical coherence tomography, and electron microscopy. The capabailties, advantages and disadvantage of each technique in imaging the in vivo and in vitro 3D constructs of cells/scaffold in tissue engineering will be discussed.
Phase contrast microscopy is often being employed to view cells in two dimensions or in highly transparent thin constructs. It has been used extensively to infer morphometric features of cells without the need for external contrast agents [3]. However, this method is qualitative in terms of optical path-length measurement. In the other words, the relationship between the irradiance and phase of the image field is generally nonlinear [4]. Normal bright field microscopy renders most of the detail of living cells invisible as there is little or no contrast between the structures inside the cell. However, when light passes through a cell, it will pass through many lipid membranes and subsequently will be bent as the light passes through materials with varying refractive indices. The contrast in these images can be increased by staining the cells with a compound such as MTT (3-(4,5-Dimethyl-2-thiazolyl)-2,5-diphenyltetrazolium bromide) which is often used to measure cell viability. The MTT is reduced to an insoluble purple colored formazan product inside viable cells. It remains inside the cell until it is eluted when the cell membrane is dissolved. In broad terms, the more dye produces greater metabolic activity of the cells [5].
Although this metabolic indicator provides good contrast, it is toxic to the cells. That’s why it is only used at the end of the experiments. Similarly if cells are grown in a 3D construct which light can penetrate through (e.g., a collagen gel or a hydrogel), phase contrast microscopy can also be used to image cells although it is very difficult to identify unstained cells in 3D. Again addition of an agent to provide visible contrast can help, as with 2D culture the metabolic indicator MTT provides good contrast but kills the cells. An alternative indicator Alamar blue is converted by living
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Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.) cells from an initial blue to a pink end-point, which is nontoxic to cells. This means cells can continue to be cultured without loss of viability [5].
In brief, phase contrast microscopy is usually available in cell culture laboratories and is easy to use but getting highresolution photographs are difficult in this method. Although this method can be used in very limited applications in
3D, it works best in 2D. The samples cannot be very thick and they must be virtually transparent [5,6].
In fluorescence or phosphorescence phenomenon, specimens absorb and consequently reradiate light. In fluorescence phenomenon, the time delay between photon absorption and emission is usually less than a microsecond. Thus, we can consider that fluorescent emission and absorption of the excitation light are simultaneous. However, in phosphorescence phenomenon the emission stays longer after the excitation light is extinguished [7].
Fluorescence microscopy is a strong method of imaging in cell and molecular biology at sub-cellular levels of resolution. Direct live cell imaging to investigate the physiological processes of cell or tissue is among the capabilities of this method of microscopy [8]. The main benefit of fluorescence microscopy is its superior sensitivity and range compare with other microscopy tools that work on the basis of alterations in optical density or chemiluminescent emission. In chemiluminescent emission one photon emits per molecule while a few hundreds of photons can be emitted by one fluorochrome [9].
An ordinary fluorescence microscope transports excitation light of the suitable wavelengths to the sample and distincts the excitation light and the emitted fluorescence. After collecting the emitted fluorescence, the imagining of the sample is possible with high degree of details as a single fluorescing molecule is possible to be pictured. [10].
Using multiple stains gives the fluorescent microscope the capability of simultaneously identifying a number of targets. Imaging with spatial resolution below the diffraction limit is not possible with fluorescence microscopy however it can detect the fluorescing molecules below these limits [7]. Although fluorescence microscopes are relatively expensive, imaging with high contrast and without manipulation of samples are worth the expenses [5].
Fluorescence microscopy enables the study of diverse processes including many features in biological systems such as protein location and associations, motility, and other phenomena such as ion transport and metabolism. Resolution and light collection of wide-field fluorescence microscopy (WFFM) involves with excitation of fluorophore(s) in the sample of interest. It is also possible to reverse this process to a certain extent by computer-based methods commonly known as deconvolution microscopy. Deconvolution fluorescence microscopy (DFM) is a kind of WFFM, which needs prior science of the point spread function (PSF). There are various algorithms available for 2D or 3D deconvolution. Its measurement technology is based on using a fluorescent bead with the size of less than 0.2 μ m. Stimulated emission depletion (STED) microscopy has been recently developed as a new super resolution method that modified fluorescence microscopy resolution to levels of electron microscopy techniques by approximately an order of magnitude over traditional techniques. . This superior advanced microscopy can be used for detailed observations of biological processes [8].
Fluorescence microscopy can be employed to image 3D objects such as tissue engineered constructs by adopting systems such as AXON ImageXpress (Axon Instruments/Molecular Devices, Union City, CA). Although such systems are suitable to detect the general location and number of cells, the quality of images weakens with depth. That’s why only thin and transparent objects can be practically used in this method. Among the scaffold used in tissue engineering the fibrin gels or thin electrospun scaffolds are ideal for this technique. The images obtained are usually en face slices through the sample. This means that 3D experiments with a fluorescence microscopy end point have to be carefully thought out to increase the chances of getting good images [5].
Fig. 1 shows 3D fluorescence images of two different types of poly(D,L-lactide-coglycolide) (PLGA) scaffolds with uniform and non-uniform pores. Fibroblast cells were seeded into the scaffolds to investigate the distribution of cells them. After 7 days of cell culture, 4'-6-diamidino-2-phenylindole (DAPI) was used to stain the nuclei of the cells in the scaffolds and then sectioned the constructs, around 500 μ m in depth, using a microtome. The images showed that the cells were uniformly distributed throughout the inverse opal scaffold, which can assist the seeding efficiency, migration of cells, and transportation of nutrients [11].
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Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)
Fig. 1 Fluorescence images of fibroblast cells after 7 days of culture from the center plane of (a) an inverse opal scaffold and (b) a non-uniform scaffold
[11].
Since the biological tissues scatter light strongly, deep imaging with high-resolution is not easily possible by traditional fluorescence microscopy. The microscopy techniques that are sued one-photon or linear absorption for contrast generation, can be only employed to image tissues that are in 100 μ m distance from the surface. At higher depths, multiple light scattering hazes the images especially for high-resolution imaging. Confocal microscopy is particularly influenced by scattering. This limitation is resolved by nonlinear optical microscopy, in particular two photon-excited fluorescence microscopes that provide large depth penetration and imaging of various organs of living animals within several hundred microns depth. In this kind of microscopy due to the localized nonlinear signal generation, the multiply scattered signal photons can be assigned to their origin and allows the sharp 3D imaging of tissues [12, 13].
Multiphoton microscopy is a growing technique for bio-imaging. It has been one of the most accurate microscopy methods for many biological applications specifically in biomedicine [14-16]. Using near infrared laser excitation source has several benefits to image the structures and record the dynamic interactions of biological tissues. The key advantages of multiphoton imaging include enhanced axial depth discrimination, decreased photodamage, and increased imaging depth.
The multiphoton fluorescence and second harmonic generation (SHG) microscopes make a very useful combination in getting complementary information on tissue engineered constructs. The SHG signals initiate from structures lacking a centrosymmetry and the autofluorescence signals results from the molecular excitation of intrinsic fluorophores [17-
23]. Fig. 2 shows a collagen scaffold in which the autofluorescence and SHG images shows that corresponding signals originate from different quantities of the sample [12].
One of the most growing methods in imaging the intact tissue is employing the two photon-excited fluorescence laser-scanning microscopy (2PLSM) when the fluorescence markers are used for the in vivo studies [24, 25]. In many aspects, a 2PLSM and confocal microscope are similar in many aspects except for the excitation laser and the detection pathway. Confocal microscope might be able to convert to multiphonton microscope by major modification and rebuilding of some parts [14].
Fig. 2 3D multiphoton images of collagen scaffolds. The autofluorescence (left panel), SHG (middle panel), and combined
(right panel) images [12].
The restriction of wide field microscopy of low resolution of fluorescence microscopy can be compensated by Confocal laser scanning microscopy (CLSM) which has the ability of image generation in 3 dimentional. [26-30]. Laser scanning confocal microscope provides a non-destructive imaging method which simplifies the in situ characterization of living cells and tissues [26, 31]. Using CLSM one can reveal the presence of single molecule. Typically imaging of thin films of tissues with thickness of up to 100 μ m is possible with CLSM.
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Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)
It is suggested that CLSM imaging of live cell can cause the light activation of intracellular calcium transients and oscillations. It is harmful for cells and may cause the cell death reduce their viability. This effect can be more pronounced when some fluorescent materials such as fluo-4 AM calcium indicator have to be used in the experiments.
However, different tricks may be used for different materials to increase the cell viability (e.g. treatment with the antioxidant ascorbate if not harmful for the xeprimnet) [32].
The confocal laser scanning microscope is one of the most popular equipment for producing the 3D image of live cells and thin tissues. Some modern CLSM systems are equipped with software to perform complex 3D ( z -stack, or xy images taking sequentially from top to bottom of the sample), 4D ( z -stack over time), or even 5D ( z -stack over time including spectral imaging) experiments. Such advanced systems offer a variety of advantages. They enable rapid data acquisition for complex methodologies such as fluorescence resonance energy transfer (FRET), spectral deconvolution and fluorescence recovery after photobleaching [8]. Fig. 3 represents how fluorescently labeled polymeric vesicles penetrate into tissue engineered skin [33].
Although CLSM provides higher resolution than conventional fluorescence microscopy, confocal microscope can be
10-20 times more expensive than fluorescence microscope. It is the big disadvantage of CLSM. In addition, the resolution of CLSM highly depends on the camera and the lens numerical aperture. Another limitation which is specifically important in tissue engineering is the fact that imaging by CLSM involves penetration of light through the specimens. If the tissue engineering scaffolds are very dense or not thin enough light penetration will be blocked and there is a finite length of depth penetration that is generally of the order of 200–300 μ m. [5].
Fig. 3 (a) CLSM image of a tissue engineered oral mucosa model exposed to rhodamine-labeled PMPC25-PDPA70 for 48 h. Images on the left show x-y and x-z sections. The image on the right is a 3D projection of this model. (b) Hematoxylin and eosin stained section of tissue engineered mucosa cultured for 10 days [33].
Confocal microscope is a strong instrument that can produce sharp images of samples that would otherwise look fuzzy and hazy when observed under a conventional phase contrast microscope. In confocal microscopy most of the light is excluded from part of the sample that is not from the microscope’s focal plane. Therefore, the image obtained has less blurred with better contrast than that of a conventional microscope. Confocal microscope can represent a thin cross-section of the sample
[34-39].
It has lots of applications in biology especially in dynamic processes. Capturing short-timescale dynamics of events is possible by employing video-rate confocal microscopy [40].
The advantage of confocal microscope over conventional ones is its ability to create sharper, more detailed 2D images, and gathering data for 3D imaging. The capability of a confocal microscope to generate sharp optical cross sectional image of samples and superposition of them makes it possible to create
3D reconstruction of the samples. Confocal microscope employs special software to combine the 2D images for producing a 3D rendition [36].
An effective way to investigate the influence of the decomposed scaffolds on cells and the decomposition kinetics of them is using Raman spectroscopy. It is a powerful technique for characterization of cellular events such as cell cycle, differentiation, proliferation, extracellular matrix (ECM) production, toxicity, cell death and etc. Raman spectroscopy can be applied to monitor the dynamic in vitro and in vivo changing of decomposable biomaterials and biodegradable polymers in histological sections. Light penetration is one of the challenges of Raman spectroscopy in many scaffolds. Hence, finding the best part of the sample, laser focusing and the interpretation of scaffold features become difficult. In this regard, SEM combined with Raman spectroscope offers good solutions for this problem. [41].
Confocal microscopy can be joined with other methods such as Raman microscopy to accomplish further analysis of the light signal with respect to the spectral composition or time dependence. Confocal Raman microscopy can image the chemical composition of a sample by recording the intensity of characteristic Raman lines of the materials in the sample.
Thus the homogeneity of the materials can be investigated on the length scale of micrometers and above. In Raman confocal microscope the composition of biological products can be mapped to visualize the macro-phase separation of polymer mixtures [42]. Confocal Raman microscopy is a non-destructive technique to investigate live cells. In order to
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Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.) develop a phenotypic identification method, confocal Raman microscopy was successfully employed to distinguish different types of bone cell lines commonly used in bone tissue engineering [43].
This technique has been used to obtain chemical data about the degradation rate of polymer and hydrogel microsphere inside the macrophages [41].
The confocal Raman image displayed in Fig. 4a, constructed from the 1656 cm –1 Raman band of unsaturated lipids, shows the presence of many lipid droplets in a macrophage that has been incubated with a 2-hydroxyethylmethacrylatederivatized dextran (dex-HEMA) microsphere for 7 days, and Fig. 4b shows a hierarchical cluster image. According to these images, clusters 3, 5, and 6 are assigned to lipid droplets, the dextran microsphere, and the cell nucleus, respectively.
As shown in Fig. 4c, the strong Raman bands in the spectrum of cluster 3 suggest that the lipid droplets are rich in unsaturated lipids [44].
Fig. 4 (a) Confocal Raman image of a RAW 264.7 macrophage that has been incubated with a DS8 dex-HEMA microsphere for 7 days. (b) Corresponding hierarchical cluster analysis image. Clusters 3, 5, and 6 are assigned to lipid droplets, the dextran microsphere, and the cell nucleus, respectively. (c) Average Raman spectrum of cluster 3 in image (b) [44] .
An important class of tissue engineering scaffolds is made of absorbable materials. Due to their sensitivity to histological processing, tissue engineered constructs are very tricky before implantation [45]. As we discussed in section 5, light penetration through the specimens is the requirement for many microscopic techniques, which limits the view to the surface of the sample or a couple of hundred microns into the tissue engineered constructs. For example confocal microscope, which is the popular instrument for assessment of porous constructs to observe cellular interaction in tissue engineered porous scaffolds, is depth and opacity dependent as it works based on laser penetration into the material. Thus, almost no information about the central region of thick samples can be provided by such methods [46,
47].
Such information can be provided by Magnetic resonance microscopy (MRM). It has the ability to detect and quantify the approximate motion of molecules inside the tissues and also the interaction of molecules with restraining boundaries. Different endogenous image contrasts such as relaxation time, molecular diffusion and magnetic susceptibility can be achieved using MRM. Also, sit is possible to develop exogenous contrast agents that target specific functions or structures inside the tissue [48-50].
MRM and magnetic resonance imaging (MRI) follow same principles common in clinical practice. Unpaired nuclei possess a magnetic moment arising from the spin angular moment of the unpaired nucleon. The unpaired proton in 1 H is the most common source of signal biological imaging because of the abundance of protons in most tissues. It uses the thermodynamic properties for imaging. Thus it doesn’t depend on the thickness or opacity of samples and is the contingents of the applied magnetic field strength [45]. Using the parameters extracted by MRI or MRM, we can distinguish the types of tissues such as differentiating fat from smooth muscle [51]. Low sensitivity of conventional
MRI is the important drawback of this imaging method in tissue engineering. The porous structure of tissue engineered constructs can weaken the signals required for imaging [52].
For tissue engineering applications, MRM can be used to optimize the microstructure of polymers or the proliferation techniques in porous and absorbable tissue-engineered builds. In a specific investigation, the uneven dynamic cell spreading in the scaffolds were recognized and analyzed by employing the MRM methods in a cellular porous polylactide (PLLA) system. Different cellular numbers in dynamic loading were identified and distinguished using
MRM. MRM can take images of the microstructure of tissue engineering scaffolds at any depth and produce the 3D pictures without the limitation of other microscopy method about the thickness of the samples [45]. MRM is also very useful for in vivo studies of tissue engineered constructs of fixed specimens [53, 54]. Anatomical phenotyping of genetically manipulated organisms such as transgenic mice can be studies very well using MRM [55-58].
In brief, the aspects that differentiate the MRM over the existing optical histology are the nondestructive, 3D and digital nature of the images. Imaging with MRM does not require the dehydration of tissues and it uses the unique
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Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)
“proton” stains [53]. The disadvantages of MRM include: 1) limit of spatial resolution 2) lack of staining flexibility and
3) costly equipment [53, 59].
Advances in fiber optics enabled direct examination of the organs in the body with the use of endoscopes. The new advances in optical imaging allow chemical and quantitative examination of tissues and blood cells. Optical imaging may be done by revealing optical contrasts such as reflection, absorption, scattering, or birefringence. It is a preferred method for its ease of use, inexpensive compared with most other microscopy techniques, and safety reasons due to its nonionizing characteristics.
Today there are only few optical microscopes that use the coherent properties of light for imaging in biomedical applications [60]. Due to the high scattering characteristics of most biological tissues [61, 62], it is often difficult to obtain high quality images with non-coherent lights. Optical coherence tomography (OCT) is a more recent technique developed to overcome this difficulty [63-66]. OCT which is based on advanced photonics technology is a novel imaging technique that allows imaging of optical scattering materials such as biological tissues. It uses near infrared light for which most tissues are transparent to produce high resolution 3D images. With the aid of OCT, the crosssectional images of living tissues can now be observed with micrometer resolution [60, 67, 68].
In OCT the 3D internal structure is reconstructed by optically slicing the sample. For this purpose, a Michelson interferometer is used where the interferometer is illuminated with a broad spectrum light. In this interferometric technique, rather than the intensity, the amplitude of the backscattered light from the sample is measured, which is the key to its high detection sensitivity. The coherent length of the illumination source also determines the axial resolution of the image. A highly improved axial resolution down to 1 μ m has been recently shown using femtosecond lasers with extremely broad spectral bandwidth [69, 70]. In conventional OCT the illuminated spot is scanned laterally across the sample in one or two directions to acquire cross-sectional or en face images, respectively.
OCT was used in medicine and tissues initially about two decades ago [71-73]. Owing to its potential to be as one of the first diagnostic imaging tools based on coherent optics features, OCT has attracted much attention in the photonic area [68]. Compared with acoustic microscopy, OCT functions similarly with using light instead of sound to distinguish the intrinsic differences in tissue structures. However, rather than the time-of-flight measurement, it uses coherent gating to find the location of the reflected optical signal. In contrast, OCT has a relatively limited depth of penetration due to the turbid nature of the biological tissues [67]. With today’s technology, it is possible to image small blood vessels and other structures as deep as 1-3 mm under the surface [67, 73-77]. High frequency ultrasonic imaging can achieve greater penetration depth but with lower resolution [78], which makes it a competing technology to OCT.
However, in addition to higher resolution, OCT has the advantages due to its relative simplicity and lower cost of hardware.
OCT has been used in surgical endoscopy where the probe is inserted through the laryngoscope with its tip placed either near or in gentle contact with the area of interest. It can be accompanied with a rigid endoscope to view the video simultaneously with the OCT image on a separate monitor [79].
OCT has been used to image a large variety of plants, animals, and human tissues. As an example, En Face (XY) image of a fixed human esophagus sample is shown in Fig. 5. This image is recorded at an average depth of 100 μ m below the sample surface. Despite the highly scattering nature of the medium, one can visibly see the cell membrane and nuclei in the captured image [62].
Fig. 5.
En face (XY) OCT image of a fixed human esophagus epithelium.
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For tissue engineering applications, various 3D techniques are currently used to study the interaction of cells and biomaterials. There have been several methods for visualization of these events on the surface of biomaterials. Recently stereoimaging scanning electron microscopy is frequently used to characterize the surface topography. Stereo imaging by scanning electron microscopy (SEM) can provide a fast and robust procedure for observation of 3D surface reconstructions. Stereoimaging SEM has been usully used to fill the obvious gap between the micro-CT and routine
SEM techniques. This technique is used for different purposes [80]. It seems that there is no other instrument with this wide range of abilities to monitor porous materials in comparison with SEM. This technique is critical in all fields that require characterization of porous materials for tissue engineering applications. As an advantage for many applications the samples require minimal sample preparation. SEM techniques can be used to analyze radiationally transparent materials such as human tissues and biomaterials. It can create highly magnified gray-scale images.
Recently it was found that the morphology and microstructure of fibrous scaffolds for neural tissue engineering can be estimated by using SEM. According to Naghavi et al. [81], an image analyzer program such as ImageJ, which uses grayscale level processing based on image structure, can be used to characterize the SEM micrographs (Fig. 6). They calculated the average diameters of fibers and the porosity of various layers by this software package. Using different thresholds, the SEM micrographs were converted to binary images, and porosity of scaffolds was calculated in various layers.
Fig. 6 (a) SEM micrograph of electrospun PVA/chitosan nanofibers; (b) fiber diameter distribution of PVA/chitosan nanofibers [81].
When the grayscale images were converted to binary form by ImageJ software, various layers of nanofibers could be seen by applying different thresholds. As can be seen in Fig. 7, various layers of nanofibers were applied by application of thresholds. Threshold 1 eliminated the upper layer, and so, the surface layers were captured, a representative sum of surface and middle layers was captured using threshold 2, and threshold 3 was used for capture of all the visible layers.
After converting the original image to various binary images, the porosity of each binary image was calculated.
Interestingly, they reported that the porosity of the same layers in the scaffolds fabricated with PVA and
PVA/chitosan blend did not differ significantly. However, the pore morphologies were different. They proposed and used an effective method for measurement of porosity at different layers of a scaffold using SEM micrographs [81].
Fig. 7 Various binary images with different thresholds for PVA and PVA/chitosan samples. (a) and (b) original images, (c) and (d) binary images with threshold 1, (e) and (f) binary images with threshold 2, (g) and (h) binary images with threshold 3 [81].
Literature data shows that the recent progress in adaptation of SEM techniques for observation of partially hydrated samples relies on some improvements. However, the goal of imaging in wet environments has not been addressed yet
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[82]. The images of biological samples produced by the electron microscope are inherently of very low contrast due to the organic parts of the samples. The organic elements are made of relatively light atoms of carbon, oxygen and hydrogen, which have low electron scattering power. To overcome this problem, some staining methods can be applied by the introduction of heavy atoms into the sample in order to reveal ultrastructural detail [83, 84].
Transmission electron microscopy (TEM) can also be used for imaging of tissues and biomaterials in tissue engineering. This powerful technique is not only used to visualize the cells but also to look inside the cells. For this purpose the samples need to be very thin about 100 nm at the region of interest to get good TEM images. The advantage of using thin layers is minimizing the number of atoms in the path of the electron beam, which even allows taking high quality images of epithelium, lamina propria, and basement membrane [5, 85]. Desantis et al. [86] have displayed regional differences in the mare oviductal epithelium by using SEM and TEM investigations (Fig. 8).
Fig. 8 Electron images of ampullar epithelial cells of a mare oviduct at the oestrus phase. (a), non-ciliated cells showed apical protrusions with secretory granules; (b), apical protrusions contained mostly granules with dark homogeneous matrix, some granules with moderate electron-dense matrix, few electronlucent vesicles and showed short and sparse microvilli; (c), supra-nuclear region of three non-ciliated cells showing several dyctiosomes in the central non-ciliated cell, round mitochondria and thin profiles of RER were scattered throughout the cytoplasm. CC, ciliated cells; ci, cilia; G, Golgi apparatus; m, mitochondria; mv, microvilli; n, nucleus; NC, non-ciliated cells; nu, nucleolus; sg, secretory granules; v, vesicle; arrow, apical protrusion; asterisk, rough endoplasmic reticulum [86].
Generally, SEM has a resolution of between 1 and 5 nm but TEM provides a resolution of down to 0.2 nm. However, the fixing process required for biological samples gives an effective resolution of 2 nm [87]. Nevertheless, taking TEM images of biological samples are difficult due to their low contrast. Scanning transmission microscopy (STEM) imaging is a direct approach that has been developed to solve this problem, and it is aimed to be used for biological applications
[88, 89]. The primary method currently used for obtaining insight into the 3D scaffolds and cellular structures is tiltseries TEM [90,91].The 3D STEM exhibits several advantages over tilt-series TEM due to the absence of mechanical tilt [92]:
(1) Tilt-series TEM suffers from missing information due to the missing wedge, and a range of spatial frequencies is absent in the data [93]. The 3D STEM, on the contrary, contains all the information in horizontal slices. The 3D STEM misses information in the vertical (z) direction due to shadowing and the missing cone is much larger than for tomography [94, 95].
(2) Tilt-series TEM has a limited resolution with thick samples.
(3) The total area that can be imaged with tilt-series TEM is limited by the amount of pixels and the maximal focus changes [96]. However, 3D STEM collects images pixel-by-pixel and without the need of tilt.
(4) The order of magnitude shorter time required to record the data, making it suitable for high-throughput imaging.
The 3D STEM imaging and data processing are already implemented in a fully automated procedure.
(5) The contrast mechanism of STEM can be used to detect nano-sized particles of a high atomic number inside an embedding medium of a low atomic number [97, 98].
Acknowledgements This chapter book is partially based upon work supported by Air Force Office of Scientific Research
(AFOSR) High Temperature Materials program under grant no. FA9550-10-1-0010 and the National Science Foundation (NSF) under grant no. 0933763.
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