Characterization and in vivo study of decellularized aortic scaffolds using closed sonication system

ABSTRACT

Extracellular matrix (ECM) based bioscaffolds prepared by decellularization has increasingly emerged in tissue engineering application because it has structural, biochemical, and biomechanical cues that have dramatic effects upon cell behaviors. Therefore, we developed a closed sonication decellularization system to prepare ideal bioscaffolds with minimal adverse effects on the ECM. The decellularization was achieved at 170 kHz of ultrasound frequency in 0.1% and 2% Sodium Dodecyl Sulphate (SDS) solution for 10 hours. The immersion treatment as control was performed to compare the decellularization efficiency with our system. Cell removal and ECM structure were determined by histological staining and biochemical assay. Biomechanical properties were investigated by the indentation testing to test the stiffness, a residual force and compression of bioscaffolds. Additionally, in vivo implantation was performed in rat to investigate host tissue response. Compared to native tissues, histological staining and biochemical assay confirm the absence of cellularity with preservation of ECM structure. Moreover, sonication treatment has not affected the stiffness [N/mm] and a residual force [N] of the aortic scaffolds except for compression [%] which 2% SDS significantly decreased compared to native tissues showing higher SDS has a detrimental effect on ECM structure. Finally, minimal inflammatory response was observed after 1 and 5 weeks of implantation. This study reported that the novelty of our developed closed sonication system to prepare ideal bioscaffolds for tissue engineering applications.

INTRODUCTION

The end-stage of cardiovascular diseases such as myocardial infarction and coronary artery diseases often lead to demands for bypass surgery. Limitation on the availability of appropriate autologous vessels because of the concomitant disease or previous use causes the artificial graft as a standard clinical alternative to autologous are preferred and commonly used such as polyethylene terephthalate (Dacron) and polytetrafluoroethylene (Teflon).¹ However, these materials have proven inadequate and unsuitable for generating grafts with increased thrombosis and infection, low compliance and poor elasticity.^2,3 Moreover, these synthetic materials have a lower rate of cell proliferation and ECM secretion when implanted long term.⁴

Vascular graft’s function in the normal physiological response to prevent the thrombosis and inflammation has guided attempts to construct the graft that mimic the native vascular tissues. The recent study has shown the use of xenogeneic tissues to construct the ECM based bioscaffolds with the preservation of the ECM.^5,6 Using xenogeneic tissues is advantageous because providing the possibility of off-the-shelf bioscaffolds. Various xenogeneic tissues have been isolated to construct the bioscaffolds for a variety of tissue engineering application including small intestine reconstruction, skin reconstruction, and orthopaedic application to mimic the entire native tissues.⁷ These features include the significant removal of the cells with the preservation of the ECM properties which provides the structural integrity that further could determine the biomechanical properties of bioscaffolds.^8,9 Previously, Various techniques have been used for decellularization that involves any combination of physical, chemical and enzymatic methods with various solvents, detergent, and enzyme to lyse the cells while creating the free volume spaces upon which native host cells can proliferate.¹⁰ SDS is the most used of the chemical agent in the decellularization process because of their capabilities in cells removal by dissolving the cell membranes. However, it also disrupts the integrity structure of ECM that resulted in weak biomechanical properties of tissues.¹¹

To facilitate the penetration of SDS into the tissues, we have developed the sonication decellularization system to increase the efficiency of the decellularization process. Our sonication decellularization system incorporates the use of ultrasound and SDS solution to promote a more efficient and homogenous cellular removal during decellularization. Previous work using agitation method on vascular tissue need to use over one reagent, high concentration and longer treatment time to prepare significant removal of cells from bioscaffolds with different thickness of tissues.¹² To remove cells significantly from the deep structure of tissues, the tissues should excessively expose to the single reagent which resulting in the ECM structure to disrupt, altering the collagen and elastin fibril network.¹³ Thus, it is important to improve the permeability of the SDS which can decrease the treatment time and potential to destruct the ECM’s structure.

Considering the decellularization treatment would introduce changes in biomechanical and structural properties of bioscaffolds, it is crucial to study the effects of sonication treatment on biomechanical and functionality of the bioscaffolds. The ball indention test was used to identify the viscoelasticity parameters including stiffness, compressibility and a residual force which can be derived from indentation load-displacement curve.¹⁴ The ball indention test can measure the deformation of the materials based on the indentation depth of the ball penetrate the sample’s surface with an applied load.¹⁵ Cyclic indentation testing was performed as an extension of automated ball indentation to provide the physiological loading condition to obtain viscoelastic properties of bioscaffolds.^16,17 The value of these viscoelasticity properties provides information regarding the biomechanical compliance of native aortic tissue and decellularized aortic scaffolds. Changes in biomechanical properties of an aorta are associated with modification of load-bearing structural components. This modification involves a complex interaction between the structural properties of ECM such as elastin, collagen, proteoglycan, and glycoprotein with cell signalling pathways.¹⁸

Moreover, the intense investigation is in progress to confirm that implanted bioscaffolds are biocompatible and elicit only minimal immune responses. The host reaction outcome to ECM based bioscaffolds can be attributed to factors such as decellularization methods, the use of crosslinking agents and age of source animal tissues.¹⁹ Not effective decellularization might cause inflammatory remodelling which promotes dense scar tissue which likely related to cellular debris remains in bioscaffolds, not in contact with surrounding tissues, less blood supply and not appropriate microenvironmentally cues following implantation.²⁰ The effective decellularization procedure is important to enhance the reduction of tissue antigenicity in order to succeed in xenotransplantation. Thus, the main goal of this study was to prepare the aortic scaffolds by sonication decellularization system with significant removal of the cellular components, preserved ECM structure and biomechanical properties of bioscaffolds with the minimal inflammatory response. The novelty of this study is residing on the preparation the bioscaffolds using sonication decellularization system which holds all the main aorta properties and characteristics.

MATERIALS AND METHODS

Aortic tissue preparation

Fresh aortas from 7 months old porcine were obtained from a local slaughterhouse and the excess blood and fat surrounding the aorta were removed. The aorta was dissected using a scalpel by cutting into the size of 15 × 15 x 3 mm³. An equal number of aortae were divided into the native (non-decellularized) and decellularized aorta groups. Native aortas were stored at −20°C while the remaining of the aorta underwent decellularization.

Sonication decellularization treatment

The developed closed sonication decellularization system was innovated to improve the previous open sonication decellularization system aiming to reduce the risk of contamination during decellularization. The customized sonication decellularization system comprises two main parts which are the sonication bioreactor and chiller (SS-600, Sonitech) that integrated by the main controller as shown in Fig. 1. The sonication bioreactor comprises an ultrasonic transducer, an ultrasonic generator, water bath, sample holder, temperature monitor, and an incubator. In addition, the chiller is connected to sonication bioreactor for controlling the temperature inside the sonication bioreactor during decellularization. The samples were sonicated 10mm from the ultrasonic transducer in 0.1% and 2% SDS. The frequency and power output were set at 170 kHz and 15W respectively, with a temperature of 36°C for 10 hours.²¹

FIGURE 1.

Developed closed sonication decellularization system comprises the incubator and chiller main part.

Immersion decellularization treatment

An immersion treatment was conducted to compare the decellularization efficiency with our developed closed sonication system. The samples were incubated in 0.1% and 2% SDS with 70 rpm agitations to ensure a constant and homogenous condition throughout the treatment. The treatment time was set for 10 hours at 36°C.

Washing process

All decellularized samples were rinsed in 1x PBS for 5 days under constant shaking to remove the residual SDS. The PBS solution was changed every 24 hours.

Histological analysis for observation of residual cells and structural preservation of collagen and elastin

To test the efficiency of cells removal and the general histological characterization of aortic scaffolds, the H&E staining was performed. Native and decellularized aortic scaffolds were fixed in Neutral buffered saline (NBF), dehydrated with the increased alcohol concentration (70–100%), and washed with Xylene overnight. Then, the samples were embedded in a paraffin block (EG1160, Leica, USA) and sectioned 3-µm thick using microtome (Leica RM2255, USA) for H&E, PSR, and VVG staining.

DNA quantification for observation of residual DNA

Further tested to evaluate the quality of decellularization was performed by DNA quantification. Native and decellularized samples were incubated in proteinase K with lysis buffer solution (Bioneer, Korea) at 60°C for overnight. The DNA was isolated using the AccuPrep Genomic DNA Extraction Kit from Bioneer (Korea) following a standard protocol recommended by the manufacturer. The total amount of DNA was quantified by a spectrophotometer at 260 nm and normalized to the tissue weight of 25 mg.

Scanning electron microscopy for observation of ultrastructure’s integrity

To characterize the ultrastructure of the ECM surface and fibre networks of aortic scaffolds, SEM images were used. The tissue samples were fixed with McDowell-Trump fixative at 4ᵒC for 24 hours. Samples were then dehydrated with a series of increasing concentrations of ethanol (50% to 100%). Dehydrated samples were immersed in Hexamethyldisilazane (HMDS) solution for 10 minutes and leave in a desiccator to air-dry at room temperature. Next, dried samples were mounted on SEM stub of metal and coated with Gold/Palladium using Sputter Coater (Leica Biosystem, US) for 60 seconds and visualized by using Scanning electron microscope (Carl Zeiss, Germany) with 5000x magnification.

Biochemical assay to evaluate collagen and elastin content

Biochemical assays were performed to quantify the protein of native tissues and aortic scaffolds such as collagen and elastin. The samples were first freeze-dried to obtain the Nano size of samples.

Collagen content

Collagen content was quantified using Sircol Soluble collagen assays kit following the manufacturer recommendations (Biocolor, Belfast, Northern Ireland). The collagen was extracted by acid pepsin at 4^◦C overnight. Then, the extracted collagen was incubated in the colorimetric reagent (the dye Sirius red in picric acid) for 30 minutes. After that, the Sirius red dye will release from the collagen-dye complex with alkali reagent (0.5M NaOH). The absorbance was measured at a wavelength of 555nm.

Elastin content

Elastin content was quantified using the Fastin Elastin assay kit (Biocolor, Belfast, Northern Ireland). The samples were freeze-dried and weighed, and then the alpha elastin was isolated following the manufacturer’s standard protocol. Aorta tissues were incubated in 0.25 M oxalic acid at 100°C for 1 hour. Then the extracted alpha elastin was incubated with a colorimetric reagent (TPPS in a citrate-phosphate buffer) for 90 minutes. Then, the elastin bound dye will release with Dye Dissociation Reagent. The absorbance was measured at a wavelength of 513 nm.

Indentation test to evaluate biomechanical preservation

Biomechanical testing was assessed and quantified for native and aortic scaffolds using indentation testing device as depicted previously.²² Prior to the testing, the sample was cut into the size of 6 mm X 6 mm with a thickness of 2 mm. It was placed into a customized metallic plate with a surface parallel to the base. The universal testing machine (MX-500N, IMADA) was used to perform the constraint compression-relaxation test. The test was analyzed using the digital force analyzer (FA PLUS, IMADA) which was recorded by the Recorder software (F-S Recorder, IMADA).

Four cycles were performed using 3 mm steel ball indenter at with the interval of the 30 seconds. At each cycle, the ball indenter was calibrated to positioning at the zero level base. The indenter test was started with pre-loading on the samples and proceeds with three phases include: (1) Dynamic compression with a constant load velocity of 10 mm/min until 3 N, (2) Static compression of the sample for 60 s with a load of 3 N and (3) Relaxation of the sample with a constant unload velocity of 10 mm/min until the load of 0 N. The stiffness value was obtained from the linear elastic slope of the loading curve in the load-displacement graph between 0.5 and 1.5 N as shown in Fig. 2. As for the relative sample compression value was measured by the indenter position in relation to the absolute sample height. While a residual force value was determined at the end of static compression in the load-time graph as shown in Fig. 3.

FIGURE 2.

Schematic representation of the indentation load-displacement graph illustrating the four repetitive cycles: A) Preloading, B) 1^st cycle, C) 2^nd cycle and D) 3^rd cycle. Red dotted line represents the stiffness.

FIGURE 3.

Schematic representation of the indentation load-time graph showing the four repetitive cycles: a) Preloading, b) 1^st cycle, c) 2^nd cycle and d) 3^rd cycle. The blue triangle and red square represent the compression and the residual force, respectively.

Subcutaneous implantation

Eighteen Wistar rat were group according to native tissues and sonication treatment for 1-week and 5 weeks implantation. Each rat received a piece of bioscaffold subcutaneously. Before subcutaneous implantation of bioscaffolds, all animals were anesthetized by injecting 75mg/kg Ketamine and 25mg/kg Xylazine. After 15 minutes, square pieces of bioscaffolds were implanted into a subcutaneous back pocket of the rat. All surgical interventions were performed under sterile conditions. All procedures were based on International Islamic University Malaysia on Animal Guideline in accordance with the law of experimental animal protection.

Histological evaluation of explanted bioscaffolds

After post-implantation at 1 and 5 weeks, the bioscaffolds were explanted and immersed in 10% NBF for further histological analysis by H&E staining.

Statistical analysis

All values were expressed as mean ± standard deviation (SD). One way ANOVA was used to determine the significant differences between the native tissues and aortic scaffolds using the SPSS17.0 (SPSS GmbH Software, Munchen, Germany). Statistical difference between the groups was determined using Turkey post-hoc test. P < .05 were considered as statistically significant.

RESULTS

Evaluation of cell removal

The cell removal and matrix structure preservation were verified through the observation with histological staining as portrayed in Fig. 4. The nuclei in the native sample show consistently spread and distributed with the typical histological structure of aortic tissues by the basophilic dark purple staining. In the immersion treatment tissues, the nuclei were not removed as shown in Fig. 4B and 4C with the cell remnants presented by the basophilic dark purple staining. In contrast, it confirmed the effectiveness of sonication decellularization system by the removal of all cells and preservation the matrix of the aortic scaffold with displaying no basophilic purple staining.

FIGURE 4.

Hematoxylin-Eosin staining of A) native tissue, B) 0.1% immersed aortic scaffolds, C) 2% immersed aortic scaffolds, D) and F) 0.1% sonicated aortic scaffolds, and E) 2% sonicated aortic scaffolds. The arrows indicate the nuclei stained.

Evaluation of DNA residual

Further assessment to verify and strengthen the effectiveness of sonication decellularization system, residual DNA content were quantified (Fig. 5). The average DNA content of the native aortic tissues was 3.9 ± 0.22 µg/mg. These values were significantly reduced to 0.6 ± 0.15 µg/mg (p < .01) and 0.3 ± 0.07 µg/mg (p < .01) resulting in an 87% and 92% decreased in 0.1% and 2% sonicated aortic scaffolds DNA content, respectively. In contrast, the average content of the 0.1% and 2% immersed aortic scaffolds showed only reduce about 74% and 64% compared to native tissues which showed residual DNA of 1.2 ± 0.20 µg/mg and 0.9 ± 0.08 µg/mg, respectively.

FIGURE 5.

DNA quantification of native aortic tissues and decellularized aortic scaffolds. **p < .01 indicate native versus other treatments, *p < .05 indicates sonicated with 0.1% SDS versus sonicated with 2% SDS.

Evaluation of ECM’s structure

For a more complete verification of ECM structure, collagen and elastin structure in native tissues and aortic scaffolds were identified by PSR and VVG staining, respectively. For elastic fibre detection, native, immersed and sonicated aortic samples were stained with VVG as shown in Fig. 6. Tunica media were characterized by a high amount of elastic fibres, whereas tunica adventitia and intima had few elastic fibres. The staining revealed that sonication and immersion treatment could maintain elastic fibres of aortic tissues although some differences existed among the treatments. In native aortic tissues, a small wave-like shape of lamellar elastic fibrils was crimped and stained as dark purple while collagen fibres that lie between the two fenestrated sheets of elastin were stained as pink. In contrast, the lamellar elastin fibrils of the decellularized aortic scaffolds were straight and had a large wavelike shape. The inter-lamellar spaces of the decellularized aortic scaffolds were enlarged because of the loss of inter-lamellar fine elastic fibrils and collagen fibres. The lamellar elastin fibrils at the tunica intima surface of the immersed aortic scaffolds with 2% SDS appeared to be fragmented.

FIGURE 6.

Verhoeff-van Gieson staining, Picro Sirius Red staining and polarized images of A) native aortic tissues, B) 0.1% immersed aortic scaffolds, C) 2% immersed aortic scaffolds, D) 0.1% sonicated aortic scaffolds, and E) 2% sonicated aortic scaffolds.

For collagen fibre detection, native aortic tissues and decellularized aortic scaffolds were stained with PSR. The major constituent of collagen fibrillar in tunica intima, media, and adventitia is collagen type III and I. In native tissues, the staining under bright field light showed the collagen fibres of type I (red) and type III (pink) were crimped with a wavy shape. In contrast, the collagen fibres of the scaffolds treated with higher SDS concentration were straight with the enlargement of interfibrillar space because of the loss of fine collagen fibres type III.

Collagen fibres were evaluated under polarized light to observe the color and intensity of birefringence in each layer of aortic tissues. From the polarized images, the collagen fibres in the tunica intima and media were portrayed as the red to yellow (type I) and green (type III) color, while in tunica adventitia, the fibres were demonstrated as the yellow color. In native tissues, the staining under polarized light showed the low-intensity color for all type of collagen fibres. In contrast, the collagen fibres in immersed aortic scaffolds showed high-intensity color in yellow to red with less green color because of partial loss of fine collagen fibre type III. While in sonicated aortic scaffolds, the intense color of collagen fibres was like in native aortic tissue.

Evaluation of ultrastructure integrity

SEM was performed to examine the morphology and structure of decellularized aortic scaffolds. The dense structure of ECM network with wavy collagen fibres in native tissues was demonstrated in Fig. 7A. After decellularization with 0.1% SDS of sonication treatment, the network of fibres was revealed with the detailed porous structure which appeared to be distinct holes that were previously occupied by cells. In contrast, the immersed and 2% sonicated aortic scaffolds showed the smooth surface suggesting the uncramping of fibres. In addition, the intact fibres of decellularized aortic scaffolds give the impression that sonication treatment did not damage the ECM.

FIGURE 7.

SEM images of A) native aortic tissues, B) 0.1% immersed aortic scaffolds, C) 2% immersed aortic scaffolds, D) and F) 0.1% sonicated aortic scaffolds, and E) 2% sonicated aortic scaffolds.

Evaluation of collagen and elastin content

The effect of decellularization treatment on the extracellular matrix of collagen and elastin content of the aorta was quantified using samples from the native and decellularized aorta as shown in Fig. 8.

FIGURE 8.

The collagen (A) and elastin (B) content of the native aortic tissues and decellularized aortic scaffolds. *p < .05 shows the native versus immersed and sonicated scaffolds.

Collagen preservation after decellularization

Average of soluble collagen content of native aortic tissues was 0.004 ± 0.0005 µg/mg. Soluble collagen content for decellularized aortic tissues showed significantly increased compared to native aortic tissue. Immersed aortic scaffolds with 0.1% (0.0057 ± 0.0009 µg/mg) and 2% (0.0055 ± 0.0008 µg/mg) SDS had higher elastin content than sonicated aortic scaffolds with 0.1% (0.0056 ± 0.0006 µg/mg) and 2% (0.0053 ± 0.0006 µg/mg) SDS.

Elastin preservation after decellularization

Average of elastin content of native tissues was 0.011 ± 0.0025 µg/mg. Elastin content in decellularized aortic scaffolds showed insignificantly decreased compared to native aortic tissues. Immersed aortic scaffolds with 0.1% (0.0097 ± 0.0013 µg/mg) and 2% (0.0092 ± 0.0005 µg/mg) SDS had lower elastin content than sonicated aortic scaffolds with 0.1% (0.01 ± 0.0002 µg/mg) and 2% (0.0099 ± 0.0003µg/mg) SDS. This shows the sonication treatment caused minor damage on the major elastin content.

Evaluation of biomechanical properties

This is important to evaluate the comparison of biomechanical properties between native tissues and aortic scaffolds to determine the long-term success of a vascular graft. Stiffness, compression and a residual force of native aortic tissues and decellularized aortic scaffolds were demonstrated in Fig. 9. Based on the graph, there was increment throughout the testing from cycle 1 to cycle 3 on stiffness, compression and a residual force for all the samples. For compression and a residual force, the value of immersed and sonicated scaffolds was higher than native tissues. However, there was a significant difference for 2% of immersed and sonicated scaffolds in compression data. In contrast, native tissues had a higher value than immersed and sonicated scaffolds with no significant difference in residual force data.

FIGURE 9.

Stiffness (A), a residual force (B) and compression (C) of native aortic tissues and decellularized aortic scaffolds. *p < .05 indicates the significant difference between native tissues and aortic scaffolds.

Histological analysis of explanted bioscaffolds

We evaluated at 1 and 5 weeks response of sonicated bioscaffolds after implantation into subcutaneous of the rat as shown in Fig. 10. The clear boundaries between the sonicated bioscaffolds and host tissues with different thickness can be observed at 1-week post-implantation. The thickness of boundaries separating the bioscaffolds with surrounding tissues in both sonicated bioscaffolds was smaller than in native tissues. A large number of cells can be observed in the boundaries surrounds the bioscaffolds with developed small neovasculature. Most of a large number of cells surround the sonicated bioscaffolds have a round-shape cell with polymorphonuclear and lymphocyte morphology indicating a mild inflammatory response surrounding the implantation sites.

FIGURE 10.

H&E staining at 1- week (top panel) and 5 weeks (bottom panel) following (A), (D) native tissues, (B), (E) 0.1% sonicated bioscaffold and (C), (F) 2% sonicated bioscaffold implantation into Wistar rat.

By weeks 5, the cellular response to sonicated bioscaffolds has dramatically changed. The boundaries in sonicated bioscaffolds become vague and had almost amalgamated into the surrounding tissues. Meanwhile, a clear boundary could be observed separating the native tissues with surrounding tissues. There were fewer cells in the boundaries surrounding the sonicated bioscaffolds with a larger developed vasculature. In addition, there is a remarkable difference in cells shape populated between native tissues and sonicated bioscaffolds. Most of the cells surround and infiltrate the native tissues has a round-shape cell with mononuclear morphology. Meanwhile, the majority of predominant spindle-shaped morphology and elongated nuclei aligned with collagen orientation were observed infiltrating to the center 0.1% and 2% sonicated bioscaffolds.

DISCUSSION

The main finding of the present study is biomechanically characterized the sonicated aortic scaffolds with the minimal inflammatory response. It is important to evaluate the bioscaffolds after decellularization process to anticipate the possibility of the clinical utility of sonicated bioscaffolds for tissue engineering application. Our result showed the preservation of structural integrity and biomechanical properties of sonicated aortic scaffolds with a minimal inflammatory response as determined by histological, biochemical, biomechanical and in vivo studies.

The histological analysis through H&E staining showed that the primary structure of sonicated aortic scaffolds was maintained on the microscopic scale with the intact of ECM fibres without the presence of any nuclei. VVG staining showed that elastin, a major ECM component of aortic tissues, was preserved. This was further confirmed through elastin quantification that resulted in an insignificant reduction of elastin content within the sonicated aortic scaffolds. The reduction of elastin content was expected as the outcome of the decellularization process. In previous studies of the varying type of tissue, elastin has been shown significantly reduced after decellularization.^23–26 However, the level of elastin and other cellular components can be restored in vivo after scaffolds reincorporate into the body as reported by Reimer et al.²³ From picrosirius staining, collagen fibres were preserved in sonicated aortic tissues. Biochemical quantification of collagen hydroxyproline has shown the increased level of collagen content in aortic scaffolds. The large loss of total proteins with the retention of the collagen can explain the increment of collagen content in aortic scaffolds following decellularization that made up a higher percentage of collagen content. This result is supported by the previous finding that made the same observation.^27–29

The sonication treatment did not affect the biomechanical properties of the aorta tissues which insignificantly increased the stiffness and decreased the compression and the residual force. Stiffness is an important parameter of biomechanical properties of the aorta to determine the pulse pressure that depends on the distensibility of the aortic walls to eject the blood intermittently for efficient tissue perfusion.¹⁸ According to Avolio et al., the stiffness of aorta has a relation to the “passive” mechanism that is related to material properties such as elastin, collagen, proteoglycan and glycoprotein and “active” mechanism that is related to cellular and molecular signalling.¹⁸ Hence, the stiffening of aortic scaffolds might correlate to the increase of collagen concentration in aortic scaffolds because of the loss of other components in aortic scaffolds. According to Berrilis et al. that studied the age-related changes in the amount and concentration of collagen, the increment of collagen concentration and decrement of the collagen amount during aging increased the stiffness of aorta.⁸ Other study hypothesized that change in the interface between collagen, elastic and smooth muscle cells during aging lead to the change in stiffness of the vessel wall. Moreover, the previous study on the biomechanics of decellularized carotid arteries, which obtained the same results suggested that the stiffer of the decellularized scaffolds attribute to the loosening and uncrimping of collagen network.³⁰ Furthermore, the previous study on biomechanical properties of the pulmonary artery in pulmonary hypertension demonstrated the stiffening of the conduit was strongly associated with the change in collagen content without found any change in SMC content and contraction.³¹ In addition, the less compacted of aortic scaffolds lead to increasing of fibres motility that allows fibres to recruit and reorient easily towards the direction of the applied load, resulting in increased stiffness of aortic scaffolds. Meanwhile, native tissues had compacted of the structure with the fibre-fibre interaction and cell-fibre interaction that hinder the recruitment and reorientation towards the direction of the applied load which would lead to less stiffness.³²

Residual stress is an important parameter of mechanics walls because they counteract stresses from blood pressure to increase resistance to failure which is attributed to elastin and vascular smooth muscle cell (VSMC).³³ As the histological analysis showed the preservation of elastic fibres in sonicated aortic scaffolds that similar to native aortic tissues, decreasing of a residual force is most likely because of the removal of cells which lead to the increased porosity and disruption of cells fibre interaction, resulting in residual force to be released. The insignificantly decreased of a residual force shows more viscous properties in aortic scaffolds as suggested by the statement of Sandmann et al.³⁴

Compressibility is an important parameter of mechanics walls over the pressure change during systolic and diastolic that is attributed to the elastic fibres.³⁵ The occurrence of disruption and alteration of elastic fibres after decellularization might be responsible for low compression of aortic scaffolds. The insignificantly decreased of compression of aortic scaffolds showed that the elastic fibres were not disrupted by decellularization treatment with a lower percentage of SDS. Based on this result, decellularization treatment was not affecting the compression of aortic scaffolds but SDS concentration. Previous studies reported that the addition of the small molecular weight of various solutions including SDS could dramatically affect the biomechanical properties of elastin.^36,37 For example, Kagan et al. studied the proteolysis of elastin-SDS complexes that reported it to stimulate the rate of digestion of elastin. The stimulation rate depends upon the SDS concentration and incubation time. The maximal stimulation to proteolyze the elastin is obtained when incubating with 1% SDS. In contrast, incubation with 0.03% SDS will inactivate the elastase to proteolyze the elastin.³⁸ These suggest that 2% SDS used in both treatment study was dramatically stimulating the elastase by binding to the elastin yielding the substrate complex that more susceptible to proteolysis which affects the compression of the aortic scaffold after decellularization treatment.

For in vivo studies of the host tissue response after implantation, native tissues and sonicated aortic scaffolds were implanted subcutaneously into Wistar rat for 1 and 5 weeks. At 1 week of implantation, staining with H&E revealed the dense accumulation of the polymorphonuclear cells with the appearance of primarily mononuclear cells within all of the bioscaffolds. The production of boundaries as a provisional matrix surround the bioscaffolds were seen as a result as inflamed appearance. These provisional matrices revealed that sonicated aortic scaffolds elicit minimal inflammatory response compared to native tissues. In the five weeks of implantation, the population of the mononuclear cells was observed within all bioscaffolds. Most of the fibroblast-like cells were seen in sonicated bioscaffolds. Meanwhile, most of the lymphocytes/monocytes like cells with multinucleated giant cells were observed in untreated bioscaffolds. The boundaries of host-sonicated bioscaffolds are nearly similar to host tissues in sonicated bioscaffolds and return to its original physiology. Hence, these results imply the acceptance of sonicated aortic scaffolds by host tissues.

To the best of our knowledge, this is the first report of biomechanical properties of aortic scaffolds using sonication treatment. Previous works have studied the factors that might influence the effectiveness of sonication in decellularization process, including solution parameter [pH, dissolved oxygen (DO), conductivity], sonication power and frequency, SDS concentration, and a distance of irradiation and treatment time.^39–43 The most effective cavitation intensity occurred during the 10 hours of 170 kHz sonication when DO and pH in the lowest value while conductivity in the highest value because of the production of the highly reactive free radical that responsible to destruct the cells^22,42 Surfactant concentration also contributes to the production of highly reactive free radical.⁴⁴ Both 0.1% and 2% of the SDS concentration showed complete decellularization by a previous study.⁴⁵ The acoustic intensity distribution of sonication is non-uniform with a higher at the center of irradiation. Increase the distance of the sample to the center of irradiation resulting in the non-uniform decellularization which affects the cell removal effectiveness.⁴⁶ Based on these previous studies, the sonication decellularization system enhances the SDS capacity to infiltrate into the tissue structure, which will accordingly reduce the treatment time and improve the decellularization process. Overall, the aortic scaffolds decellularized by sonication decellularization system in this study can be considered as the initial platform for further studies on other small-diameter blood vessels for tissue engineering blood vessels since it maintains the structure and biomechanical properties of native tissues with a minimal inflammatory response.

CONCLUSION

In conclusion, we developed a novel sonication decellularization system with minimal deteriorative effects on the structure and biomechanical properties of bioscaffolds for use in tissue engineering. The ideal bioscaffolds with preserve the composition and functional characteristic were obtained when treated with 0.1% SDS. In addition, 0.1% sonicated aortic scaffolds elicit minimal inflammatory response while providing a suitable environment for repopulation of host cells. This work showed the capability of our developed closed sonication decellularization system to prepare the ideal bioscaffolds for tissue engineering application. More investigation needs to be done on the biocompatibility to affirm the safety of bioscaffolds decellularized by sonication decellularization system.

Acknowledgments

The authors are grateful to the Ministry of Higher Education for financial support through the Fundamental Research Grant Scheme (FRGS/1/2017/STG05/UIAM/02/6), Prototype Research Grant Scheme (PRGS16-002-0033) and Trans disciplinary Research Grant Scheme (TRGS16-02-001-0001).