Impulse acoustic microscopy: a new approach for investigation of polymer and natural scaffolds.
A rapid development of tissue engineering and regenerative medicine over the past decades has led to the need for the development of a new method, which could be suitable for providing control at all stages of formation of tissue engineered materials and should be non-invasive. On the other hand, this method should enable one to be used for opaque specimens with a thickness from 1 mm to several cm. In this case, the methods based on ultrasound look most promising.
The high frequency ultrasound microscopy has routinely been used for small multicomponent objects in material science. Now this class of instruments alters the course towards biological and medical needs. It can be successfully applied for living objects because of using low-intensive ultra-short pulses of ultrasound beams. The advantage of the ultrasound microscopy investigation consists in getting the unique data about elastic properties of the samples. Also this kind of microscope can be easily used as a flaw detector. Quality control of specimens at the stage of developing new and prospective materials will allow one to provide their fastest introduction into experimental and clinical practice.
Only a few studies were focused on ultrasound investigation in regenerative medicine [1--3]. It has been shown that scanning acoustic microscopy is a useful method with a big potential in medical research. It may be applied for measuring attenuation, acoustic impedance, and other physical and elastic properties of biological tissues at the microscopic level. So far, investigations have been confined to the analysis of cartilage implants, tissue-engineered cartilage [1,2] and inner-surface morphology of the artificial heart .
Scaffold is the major component of an artificial organ that provides support for cell components, reproduces morphology, and structure of the organ. Thus, the important stage of tissue engineering product development is selection of the scaffold material and the way of its fabrication. A well-known bioidentical scaffold is based on the natural extracellular matrix (ECM) fabricated by decellularization of native tissues.
ECM is a derivative from donor organs and has some problems (such as infection and immune response to cell debris) frequently encountered during its application. Research interests are shifting to the development of synthetic matrixes. They have a comparatively higher quality, more available and cheaper. The most popular scaffold materials are natural and synthetic polymers. Widely used natural polymers for matrixes are polysaccharides (such as cellulose, chitin, etc.) and proteins (collagen, fibrin, etc.). Usually, polylactic acid, polyglycolic acid, and their derivates are used as biodegradable polymers for scaffolds. Artificial non-biodegradable scaffolds are made from polyamide, polyethylene, and so forth. The product should be non-toxic, be characterized by durability, adhesiveness, histocompatibility and especially biodegradability [4, 5], It is very important to control the structure of the materials during all procedures: fabrication, sterilization, wetting, post processing, and cell seeding, both in vitro and in vivo testing.
However, their biological properties and structure are quite different from native tissues, as well as ECM. Existing control methods for studies of the structure of matrixes, like scanning electron microscopy, optical microscopy, histological staining and others, are very laborious. Each of these methods has its own constraints. For scanning electronic microscopy--high cost, need of sputtering of a metal film, need of water elimination, small field of view, only surface observation; for histological researches--need of alcohol and organic solvent treatment, need of micro-slices, need of selective staining.
In this regard, one of the promising methods is acoustic microscopy, which has several advantages: being noninvasive, intravital, non-toxic, and allows observation in dynamics. Accordingly, the aim of this work is to show the possibilities of acoustic microscopy for the study of a variety of scaffolds for tissue engineering and regenerative medicine. In this article, we give results of investigating the decellularized rat diaphragm and nonwoven fibrous materials based on natural and synthetic polymers, which is similar in morphological and mechanical characteristics to natural tissues.
MATERIALS AND METHODS
Diaphragm harvesting was performed according to the previously described protocol [6, 7]. Briefly, the diaphragm was dissected from the surrounding adipose tissue, then cannulated and fixed in a customized bioreactor, allowing a combined decellularization, both perfusion and agitation. The retrograde perfusion of the diaphragm was performed (6 ml per min) through the vena cava into the right and left caudalis phrenic veins. The total perfusion time was 24 h: in 4% sodium deoxycholate for 3 h (Sigma Aldrich, USA), in PBS for 10 min (Gibco, Life Technologies, USA), in DNase I 2,000 kU (Sigma Aldrich, USA) diluted in PBS with calcium and magnesium (Invitrogen, Life Technologies, USA) for 1 h, and two washes of each of 2 mM EDTA (Sigma Aldrich, USA) in MilliQ water for 30 min. The final step of decellularization was washing in PBS (Gibco, Life Technologies, USA). All steps were performed at room temperature, with the reagent final volume of 200 ml.
Artificial scaffolds were represented by nonwovens from cellulose diacetate (CDA), chlorinated polyvinyl chloride (CPVC) and polysulfone (PSU). CDA is a synthetic polymer made by treating cellulose with acetic acid; it consists of two acetate radicals per each unit of D-anhydroglucopyranose of the cellulose molecule. CDA fibers are popular not only because of their good mechanical properties but also for their electrospinning processability. Cellulose-based materials have poor biodegradability and solubility because of their highly ordered structure; however, acetylation decreases the degree of crystallinity, which helps to overcome such problems. CDA is the most important cellulose derived biopolymer, natural and biodegradable [4, 5], Chlorinated polyvinyl chloride and polysulfone are widely used bio inert polymers with a high chemical resistance to alkalis, acids and solvents. Also they are well-known electrospun materials. The architecture of such materials is a highly porous microfibrillar network with intercalated pores. The fiber diameter was 4.3-6.2 pm. The fibers were stacked in layers with a thickness of a few fibers in diameter.
Table 1 shows the main parameters of the nonwoven materials used. All of these materials are selected from the following considerations; hydrophobicity/hydrophilicity, the number of defects and inhomogeneities in the structure, different packing density of the material. All samples were wetted by bidistilled water and stored in it.
The essence of the electrospinning technique is formation of a polymer fiber from a liquid solution. The polymer composition is forced through a metering capillary under high voltage in electrostatic field. The fiber repeatedly elongates and splits into branched jets as it moves to the collector. At the last stage-- fiber-gel solidifies due to the final solvent evaporation from its surface.
A significant part of stretching a polymer stream takes place at the base of a metering capillary. Selection of the collector type largely determines the architecture of the material. The collector can be implemented in the form of a grounded plane, a movable disk, a rod, or a drum committing rotational and reciprocating motions or other types of receivers. A distinctive feature of electrospinning is that the process is carried out in one step and the finished material is immediately formed on the receiving electrode. There is also a possibility of obtaining a multilayer and multi-component material in one technological operation. The diameter of the fibers produced by electrospinning ranges from 100 nm to 15 pm. These nanofibers can form even three-dimensional structured scaffolds with large surface areas and high porosity.
Impulse acoustic microscopy is based on probing specimens by short pulses (1.5-2 oscillations in the pulse) of high-frequency (50-200 MHz) ultrasound. The pulse is reflected from the front surface, elements of the internal structure and the bottom of the specimen. Echoes reflected from the specimen radiate from a narrow focal spot of the probe beam whose diameter gives lateral resolution. During mechanical scanning the echo signals are recorded and stored together with the coordinates of each point. The database is displayed in a quasi tomography regime and allows visualization of the internal microstructure of an object in the form of layer by layer acoustic images (C-scans) at different depths in the specimen volume and its cross-sections (B-scans) with time delay on the specimen depth , The value of an echo signal is displayed as a corresponding gray-scale pixel in acoustic images.
For the thin specimens with heterogeneous areas in the material bulk and the sound speed closer to water, it is profitable to use the B/D scanning procedure . B/D ("brightness/depth") is a modified scanning procedure with a dynamic shift of the probe beam focus position inside the specimen volume. As a result, a cross section of the object with the real elevational size is formed. Dynamic focusing allows one to increase the sensitivity of a microscope to variations of the acoustic impedance and to get a high elevational resolution.
Impulse acoustic microscopes SIAM-2 with operation frequencies of 100 MHz and angular aperture of 11[degrees], produced by the Institute of Biochemical Physics, RAS, was used to visualize the bulk microstructure inside specimens of a few millimeters in thickness with lateral resolution of 15-45 pm. Ultrashort (20-40 ns) probing ultrasonic pulses provide 15-30 pm elevational resolution. The scanning mechanical system of a microscope with the speed of 20-25 mm/sec allows one to get an acoustic image of the specimen volume in 10-15 min. Bidistilled water was used as an immersion liquid.
Investigation of nonwoven scaffold surfaces has been done by optical microscopy Leica DMCM and by scanning electron microscopy (SEM) under a Helios Nanolab (FEI) 600 microscope at a voltage of 2 Kv.
Decellularized diaphragm specimens were fixed in 2.5% glutaraldehyde (Merck, USA) with 0.1M cacodylate buffer (Prolabo, Sweden) at room temperature for 2 h. After that specimens were washed in cacodylate buffer and dehydrated through an ethanol gradient, processed through critical point drying, and sputter coated with gold. Evaluation of the fiber configuration was observed with SEM.
Decellularized diaphragm specimens were fixed in 10% neutral buffered formalin (Histolab, Sweden) at room temperature overnight, processed and embedded in paraffin. Specimens were sectioned at the thickness of 5 mm. To evaluate the quality and effectiveness of the decellularization process and matrix preservation routine histological methods were used. Hematoxylin and Eosin (Histolab, Sweden) staining (H&E) was performed both in native and decellularized tissues according to manufacturers' instructions.
RESULTS AND DISCUSSION
Histological staining and SEM are basic and conventional types of investigation of decellularized tissues. The main feature of acoustic microscopy is a possibility of a nondestructive visualization of the internal structure. A focused ultrasound beam penetrates inside scaffolds and reflects different microstructural elements--single fibers or their conglomerates, air inclusions, and so forth.
The correlation between the structure of fibers, their direction, the architecture of the diaphragm muscle tissue and also fascia, covering the diaphragm on both sides (Fig. 1d), can be clearly seen in the acoustical images (Fig. 1c and d). The prevalent fiber orientation and its packing are shown in the C-scan at the depth of 300 pm (Fig. 1c). These parameters correspond to the morphological data obtained from optical and scanning electron microscopy (Fig. 1a and b). Accordingly, comparative studies of the decellularized diaphragm specimens by routine histological methods, ultrastructure analyses (SEM), and acoustic microscopy suggest correlation between them. Acoustic microscopy can be used as an additional method for quality evaluation of the obtained decellularized matrixes.
Another class of objects is nonwoven scaffolds based on various types of polymers. SEM and optical microscopy without staining are also basic methods of investigating the microstructure of fibrous materials. Images in Figs. 2 and 3 show the surface and internal microstructure of CPVC specimens with different fiber diameters and packing density obtained via different methods. A chaotic distribution of fibers and partial fiber interweaving can be observed in all specimens. Air bubble inclusions are also clearly visible. Figure 2 shows a nonwoven hydrophobic low density material CPVC 5,5. The loose surface and bulk structure are formed because of low density of the specimen (Fig. 2e). Individual fibers and their orientation, fiber packing density and air bubbles were visualized in the specimen volume (Fig. 2c-e). Air bubbles are elongated along the fibers in the subsurface layer and have an oval shape. A low density of this specimen allows one to reach a low concentration of air bubble artifacts in the bulk (Fig. 2c and d). The images of the specimen surface obtained by different methods are similar (Fig. 2a-c). The fiber diameter (4--6 pm) is less than the ultrasound wave length (~15 pm at 100 MHz); nevertheless, acoustic microscopy allows one to observe not only the surface of opaque specimens as optical and electron microscopy do, but also gives layer-by-layer information about their internal microstructure. The acoustic images performed in an ultramicroscopical mode  give information on the presence of small-sized scatterers and their distribution over the specimen bulk, rather than on their real size and shape. Imaging of these elements is possible because of Rayleigh scattering of ultrasound from them. The ultrasonic objective receives its backscattered radiation. Particle sizes and shape have an impact on the efficiency of scattering and the amplitude of echo; respectively, the brightness of corresponding spots in the image also changes . The acoustic ultramicroscopy mode is similar to the dark field in optical microscopy.
Figure 3 shows a higher density hydrophobic material CPVC5,9. Optical and SEM (Fig. 3a and b) images give information only about the dense fiber packing. The acoustic image (B-scan) shows the distribution of the areas of different density over the specimen volume (Fig. 3e). Furthermore, air bubbles were visualized inside the sample (Fig. 3c and e). It is clear that the acoustic beam fails to separate the individual fibers both at the surface and at the depth inside the dense nonwoven material (Fig. 3c and d).
Figure 4 represents another nonwoven material made from polysulfone (PSU). PSU is also hydrophobic. Air bubbles at the subsurface layer are the same as in the CPVC specimen: oval shaped and elongated along the individual fibers. Two largest air bubbles, whose presence is caused by polymer specification, are clearly seen in the volume (Fig. 4d and e). All air bubbles, especially big ones, are critical defects for the matrix decontamination and future cells migration. Accordingly, the detection of any air bubbles inside a specimen is one of the basic questions. Such air defects, whose size a is larger than the focal spot dF of the probe ultrasound beam (a^lOdf), generate an acoustic shadow in the volume under the inclusion. The only way to visualize the microstructure under a large inclusion is specimen scanning from the bottom side.
Figure 5 shows a hydrophilic nonwoven material made from CDA. Due to the low density of the specimen individual fibers can be easily seen at the surface (Fig. 5e) and at the depth of the specimen (Fig. 5d). Swelling of the fibers is not detected because its diameter is still smaller than the ultrasound wave length. Air bubbles are more spherical and located under the interweaving of the fibers all over the specimen thickness.
Wetting of all artificial matrixes with the high surface area is an important but difficult task in regenerative medicine; it can be applied to both hydrophobic and hydrophilic polymers. Wetting (both local and macroscopic) is a critical parameter for cells. Also, contamination and formation of cavities are possible in the deficient areas.
Detection of non-wet scaffold parts can be performed the most objectively and precisely because of the meaningful difference between the acoustic impedance of the polymer media and air bubbles. The acoustic microscopy technique makes it possible to find the smallest quantity of artifacts in comparison with the other aforesaid techniques.
The acoustic microscopy allows one not only to detect the defects in wetted matrixes, but also to identify the characteristics of these defects. So, in the hydrophobic patterns (CPVC, PSU) air bubbles with an elongated shape were detected along the length of the fibers; in the hydrophilic specimens (CDA) air defects with a spherical shape were delocalized too. At the same time, air bubbles are completely absent on the acoustic images of the decellularized diaphragm, which adds support to the fact.
It has been shown that impulse acoustic microscopy is a noninvasive tool suitable for investigating the surface and internal structure of opaque scaffolds. Also it allows one to diagnose and examine the heterogeneous regions and defects inside the specimen bulk. All defects can impede the humidification of a matrix, which will be critical for cell colonization.
It should be noted that none of the used methods, except acoustic microscopy, allow one to analyze defects in the specimen bulk. Also the acoustic microscopy method permits examination of specimens with the area more than 1 [cm.sup.2] in one session and without their long previous preparation.
The method provides monitoring and evaluation of tissue engineering products both in vitro and in vivo, and even investigation of the engraftment process of the tissue implant in situ and in some cases in real time. These features define the essential novelty of the approach and the significant advantages of this method in comparison with other techniques currently used in tissue engineering. Future investigation will facilitate understanding the mechanisms of migration and proliferation of cell cultures within the matrix.
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Elena Khramtsova, (1) Egor Morokov, (1) Kseniya Lukanina, (2) Timofei Grigoriev, (2,3) Yulia Petronyuk (iD), (1,4) Alexey Shepelev, (2) Elena Gubareva, (5) Elena Kuevda, (5) Vadim Levin, (1) Sergey Chvalun (2)
(1) N.M. Emanuel Institute of Biochemical Physics Russian Academy of Sciences, Moscow, Russia
(2) National Research Centre "Kurchatov Institute, " Moscow, Russia
(3) A. N. Nesmeyanov Institute of Organoelement Compounds Russian Academy of Sciences, Moscow, Russia
(4) Scientific and Technological Center of Unique Instrumentation, Russian Academy of Sciences, Moscow, Russia
(5) Kuban State Medical University, Krasnodar, Russia
Correspondence to: Y.S. Petronyuk; e-mail: firstname.lastname@example.org
The material was partially presented at the TOP 2016 Conference.
The reported study was partially funded by RFBR according to the research projects 15-33-20986, 16-02-00855a, 17-03-01361a.
Published online in Wiley Online Library (wileyonlinelibrary.com).
Caption: FIG. 1. Decellularized rat diaphragm: (a) Histological evaluation. H&E staining shows the absence of cell nuclei and preservation of extracellular matrix, magnification X10; (b) SEM evaluation of the surface, magnification X7500; (c) acoustic image (C-scan) at the depth of 300 [micro]m, bar--1 mm: (d) acoustic image (B/D-scan) corresponds to the histological section across the diaphragm tissue and along the fibers, bar--500 mkm.
Caption: FIG. 2. Images of the CPVC5,5 specimen: (a) optical image of the surface microstructure, bar--100 pm; (b) SEM evaluation of the surface, bar--200 [micro]m; (c) acoustic image (C-scan) of the subsurface layer, bar--1 mm; (d) acoustic image (Cscan) of the internal microstructure at the depth of 300 pm from the surface, bar--1 mm; (e) acoustic image (B-scan), cross-section in the central part of the specimen; 1--air bubble. [Color figure can be viewed at wileyonlinelibrary.com]
Caption: FIG. 3. Images of the CPVC5,9 specimen: (a) Optical image of the surface microstructure, bar--100 pm; (b) SEM evaluation of the surface, bar--200 [micro]m; (c) acoustic image (C-scan) of the subsurface layer, bar--1 mm; (d) acoustic image (C-scan) of the internal microstructure at the depth of 300 pm from the surface, bar--1 mm; (e) acoustic image (B-scan), cross-section in the central part of the specimen; 1--air bubble. [Color figure can be viewed at wileyonlinelibrary.com]
Caption: FIG. 4. Images of the PSU specimen: (a) Optical image of the surface microstructure, bar--100 [micro]m; (b) SEM evaluation of the surface, bar--200 [micro]m; (c) acoustic image (C-scan) of the subsurface layer, bar--1 mm; (d) acoustic image (C-scan) of the internal microstructure at the depth of 300 pm from the surface, bar--1 mm; (e) acoustic image (B-scan), cross-section in the central part of the specimen; 1--air bubble; 2--large internal air bubbles.
Caption FIG. 5. Images of the CDA specimen: (a) Optical image of the surface microstructure, bar--100 [micro]m; (b)-- SEM evaluation of the surface, bar--200 [micro]m; (c) acoustic image (C-scan) of the subsurface layer, bar--1 mm; (d) acoustic image (C-scan) of the internal microstructure at the depth of 300 [micro]m from the surface, bar--1 mm; (e) acoustic image (B-scan), cross-section in the central part of the specimen; 1--air bubble.
TABLE 1. Parameters of used nonwoven materials. Average fibers Sample Compound diameter ([micro]m) CPVC5.5 Chlorinated polyvinyl chloride 5.5 CPVC5.9 Chlorinated polyvinyl chloride 5.9 PSU Polysulfone 4.3 CDA Cellulose diacetate 6.2 Packing Thickness Sample density (%) (mm) CPVC5.5 7 0.5 CPVC5.9 11 0.5 PSU 3 1.5 CDA 4 0.8
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|Author:||Khramtsova, Elena; Morokov, Egor; Lukanina, Kseniya; Grigoriev, Timofei; Petronyuk, Yulia; Shepelev,|
|Publication:||Polymer Engineering and Science|
|Date:||Jul 1, 2017|
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