Annals of Biomedical Engineering, Vol. 42, No. 6, June 2014 ( 2014) pp. 1305–1318 DOI: 10.1007/s10439-014-1000-1

On the Decellularization of Fresh or Frozen Human Umbilical Arteries: Implications for Small-Diameter Tissue Engineered Vascular Grafts HO-YI TUAN-MU,1 CHEN-HSIANG YU,2 and JIN-JIA HU1,3 2

1 Department of Biomedical Engineering, National Cheng Kung University, #1 University Rd., Tainan 701, Taiwan; Department of Gynecology and Obstetrics, National Cheng Kung University Medical Center, Tainan, Taiwan; and 3Medical Device Innovation Center, National Cheng Kung University, Tainan, Taiwan

(Received 16 October 2013; accepted 22 March 2014; published online 29 March 2014) Associate Editor Dan Elson oversaw the review of this article.

INTRODUCTION

Abstract—Most tissues, including those to be decellularized for tissue engineering applications, are frozen for long term preservation. Such conventional cryopreservation has been shown to alter the structure and mechanical properties of tissues. Little is known, however, how freezing affects decellularization of tissues. The purpose of this study was two-fold: to examine the effects of freezing on decellularization of human umbilical arteries (HUAs), which represent a potential scaffolding material for small-diameter tissue-engineered vascular grafts, and to examine how decellularization affects the mechanical properties of frozen HUAs. Among many decellularization methods, hypotonic sodium dodecyl sulfate solution was selected as the decellularizing agent and tested on fresh HUAs to optimize decellularization conditions. The efficiency of decellularization was evaluated by DNA assay and histology every 12 up to 48 h. The optimized decellularization protocol was then performed on frozen HUAs. The stiffness, burst pressure, and suture retention strength of fresh HUAs and frozen HUAs before and after decellularization were also examined. It appeared that freezing decreased the efficiency of decellularization, which may be attributed to the condensed extracellular matrix caused by freezing. While the stiffness of fresh HUAs did not change significantly after decellularization, decellularization reduced the compliance of frozen HUAs. Interestingly, the stiffness of decellularized frozen HUAs was similar to that of decellularized fresh HUAs. Although little difference in stiffness was observed, we suggest avoiding freezing if more efficient and complete decellularization is desired.

Atherosclerotic vascular diseases are the leading cause of morbidity and mortality in modern society. In some patients, arterial bypass surgery is required to restore blood flow and preserve functions of downstream tissues. Synthetic grafts (e.g., expanded polytetrafluoroethylene and Dacron), which work well in replacing high-flow, large-diameter arteries, are not suitable for small vessel reconstructions due to acute thrombogenicity and anastomotic intimal hyperplasia.24,42 Autologous native vessels remain the gold standard for small-diameter arterial replacement. Unfortunately, many patients do not have suitable autologous vessels for surgery because of previous harvest or other vascular disease.43 Moreover, additional surgery for vessel harvest is required for using autologous vessels. A variety of tissue-engineering strategies have thus been developed for functional small-diameter vascular grafts.5,38 The mechanical properties of a tissue-engineered vascular graft (TEVG) are critical for its success. Specifically, compliance mismatch between the graft and the host artery was thought to affect graft patency.4,16 TEVGs fabricated based on natural polymers such as collagen and fibrin usually suffer from insufficient mechanical strength35 while those made from biodegradable synthetic polymers such as poly(lactic acid)41 and polycaprolactone28 are generally too stiff. This issue is less of a concern if decellularized arteries are used as a scaffolding material as decellularization preserves the majority of extracellular matrix (ECM) and hence the mechanical properties of native arteries.17,23,38 Decellularized human umbilical vessels, particularly, represent a potential scaffolding

Keywords—Human umbilical arteries, Decellularization, Small-diameter tissue-engineered vascular grafts, Mechanical properties, Conventional cryopreservation, Freezing.

Address correspondence to Jin-Jia Hu, Department of Biomedical Engineering, National Cheng Kung University, #1 University Rd., Tainan 701, Taiwan. Electronic mail: [email protected]

1305 0090-6964/14/0600-1305/0

 2014 Biomedical Engineering Society

1306

TUAN-MU et al.

material for small-diameter TEVGs because their dimensions are comparable to target arteries and sufficiently long grafts without branches are achievable. Cryopreservation is an important tool for long term storage of blood vessels that are to be used as vascular grafts with an aim to preserving their functional characteristics. In a typical conventional cryopreservation protocol, blood vessels were equilibrated with culture medium or buffer solution containing cryoprotectants [e.g., dimethyl sulfoxide (DMSO), sucrose, etc.], frozen at a defined cooling rate, and then stored in deep subzero temperatures.3,27 Although it has been shown that the conventional cryopreservation influences the structure and mechanical properties of arterial tissues.27,30,44 To the best of our knowledge, the effects of freezing on decellularization of human umbilical arteries (HUAs) have not been studied. Note, particularly, that some of previous studies used frozen human umbilical vessels for decellularization.1,11 The purpose of this study was two-fold: to examine the effects of freezing on decellularization of HUAs and to examine how decellularization affects the mechanical properties of frozen HUAs. It is known that the efficiency of decellularization depends on the origin of target tissues and specific methods that are used.15 A few decellularization methods specific for human umbilical blood vessels have been reported.1,11,17 For example, sequential treatments of 3-[(3-Cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS) buffer, sodium dodecyl sulfate (SDS) buffer, and endothelial growth media-2 were used to decellularize HUAs.17 Detergents are commonly used in decellularization as they solubilize cell and nuclear membranes and dissociate DNA from proteins. Specifically, SDS is one of the most commonly used ionic detergents and appears to be more effective than other detergents for removing cell residuals from dense tissues.10 SDS solution has been used alone1,11,34 or combined with enzymatic treatments17,25 for decellularizing blood vessels. The concentrations of SDS used in these studies and their efficiency on decellularization are not consistent, however.1,11,17,34 In this study, hypotonic SDS solution was selected as the decellularization agent and tested on fresh HUAs to optimize decellularization conditions. The efficiency of decellularization was evaluated by DNA assay and histology every 12 up to 48 h. The optimized decellularization protocol was then performed on frozen HUAs. The stiffness, burst pressure, and suture retention strength of fresh HUAs and frozen HUAs before and after decellularization were also examined.

METHODS Preparation of HUAs Human umbilical cords were obtained from Department of Obstetrics and Gynecology, National Cheng Kung University Hospital with patients’ consent. Immediately after delivery, the cords were cut into 10-cm segments and placed in cold normal saline. The cords were then transferred to the laboratory for processing. Briefly, intact HUAs were isolated from the Wharton’s jelly using blunt dissection. The HUAs were cut into half; one part (fresh HUA) was either subjected to decellularization or mechanical testing, and the other part (frozen HUA) was frozen in excess volume of normal saline (volume ratio of saline to tissue is greater than 20) at a cooling rate of 1 C/min. The frozen HUAs were stored at 220 C for 1 week prior to further processing or testing. Decellularization of HUAs We found in our preliminary experiments that hypertonic or isotonic SDS solution is less efficient in cell removal so hypotonic SDS solution was selected to be optimized. SDS (Mallinckrodt Baker, Phillipsburg, NJ) was dissolved in deionized water to prepare SDS solutions with three concentrations (0.1, 0.5, and 1% (w/v)). The effect of duration of treatment (12, 24, 36, and 48 h) for each concentration was examined. Fresh HUAs from five donors were used (N = 5); one HUA from each donor was cut into twelve 2-mm segments (one for each concentration and each time point). The segment was then incubated in excess volume of SDS solution with agitation at room temperature. After the specified duration, the segment was washed in agitated deionized water for at least five times until no bubbles were found in the solution to remove residual SDS and then processed for either histology or residual DNA quantification. Upon the determination of optimal decellularization protocol, frozen HUAs from another three donors were subjected to the optimal protocol and the results were compared with their fresh counterparts (N = 3).

Histology and Image Quantification Segments of HUAs were fixed in 10% neutral-buffered formalin in an unloaded configuration overnight at room temperature, dehydrated through a series of graded alcohol overnight, and then embedded in paraffin to enable examination of cross sections. Five

On the Decellularization of Fresh or Frozen HUAs

micron sections were cut using a microtome (Leitz 1512, Leica, Germany) and collected on positively charged slides. After removal of paraffin and re-hydration, sections were stained with H&E, Alcian blue, and picrosirius red (PSR) for illustration of nuclei, glycosaminoglycans (GAGs), and collagen, respectively. Histological images were acquired by an optical microscope (DM2500P, Leica, Germany) with a CCD camera (DFC295 digital camera, Leica, Germany). In particular, PSR-stained sections were imaged under polarized light. Specifically for quantitative analysis, ten representative 24-bit color images of 2048 9 1563 pixel resolution were acquired per specimen with a 409 objective and saved in tagged-image file format (TIFF). Relative collagen content and collagen staining intensity were analyzed by a LabVIEW routine previously developed for quantifying collagen in PSR-stained sections.19 Relative GAG content was analyzed by another similar LabVIEW routine. In addition to histology, the rehydrated sections were also imaged by multiphoton microscopy (see below). Multiphoton Microscopy Second harmonic generation (SHG) images (1024 9 1024 pixels) were acquired by a multiphoton microscope (FluoView FV 1000, Olympus, Japan) with a 259 or 409 water immersion objective (XLUMPLFLN 1.12 NA, Olympus, Japan). The microscope system was fitted to a mode-locked Ti-sapphire laser (Maitai-HP DeepSee-OL, Newport Corporation, Irvine, CA). Excitation wavelength was set at 900 nm and laser power was set at 35 mW throughout image acquisition. The SHG signal was collected by a non-descanted detector through a 420–460 nm bandpass filter and a 485 dichroic mirror. The pixel dwell time was 40 ls. For collagen quantification, ImageJ (NIH) was used to measure the signal intensity per area of the media. Data from three images for each group were averaged for statistical analysis. DNA Quantification Quant-iT PicoGreen dsDNA assay kit (Invitrogen, USA) was used to quantify residual DNA in the processed HUAs. Briefly, the specimen was lyophilized at 240 C for 24 h and its dry weight was measured. The dried specimen was incubated in a papain solution, which contains 20 U/mL papain (Worthinton, Lakewood, NJ), 1.1 mM EDTA (Panreac, Spain), 5.5 mM cysteine–HCl (Panreac, Spain) and 0.067 mM 2mercaptoethanol (Alfa Aesar, England) overnight at 60 C until the specimen was completely digested. The solution was then diluted with 0.2 M Tris–EDTA buffer and then incubated with the working solution of the kit in a 96-well plate. The fluorescence of the sample was

1307

measured using a fluorometer (excitation: 485 nm, emission: 538 nm; Fluoroskan Ascent, Thermo Fisher Scientific, Waltham, MA) and values were compared with a k dsDNA standard (0–10 ng/mL) to determine the weight of the residual DNA. Finally, the weight of the residual DNA was normalized by the dry weight.

Mechanical Properties of the HUAs The stiffness of fresh HUAs, decellularized fresh HUAs, frozen HUAs, and decellularized HUAs was examined by pressure–diameter tests using a custombuilt mechanical tester, which consists a stepper motor with a motion control system (MID-7604 and PXI7330, National Instruments, Austin, TX), a syringe pump (KDS-210, KD Scientific, Holliston, MA), a pressure transducer (Model 209, 0-5psig, Setra, Boxborough, MA), a load cell (LTS-200GA, Kyowa, Japan), a 1394 CCD camera (656 9 492, Stingray F033B, Germany) with a TV lens (HF25HA-1B, Fujinon, Japan), and a custom-made loading frame.20,21 Pressure–diameter tests have been regarded as the best method to assess the mechanical properties of blood vessels as the tubular shape of the vessel is preserved.22 HUAs from seven donors were used. One sufficiently long HUA from each donor was cut into four segments of ~25 mm long; each of which underwent one of the four treatments prior to mechanical testing (N = 7 for each treatment). Briefly, the HUA was cannulated with luer adaptors using 3-O suture and coupled to the loading frame. The HUA was submerged in a chamber filled with normal saline at room temperature and then air in the tubing was expelled. The HUA was pressurized cyclically between 0 and 150 mmHg for ten times using a syringe pump at a flow rate of 0.2 mL/ min for preconditioning. After preconditioning, the HUA was decoupled from the loading frame and recoupled at its unloaded configuration (luminal pressure = ~10 mmHg and axial load = ~0 mN). The dimensions of the HUA (i.e., the outer radius and the length between the two suture ties) were recorded at the unloaded configuration; circumferential and axial stretches were calculated based on these dimensions. The axially constrained HUA was then subjected to cyclic pressurization at its unloaded length. Note that preconditioned HUAs did not buckle when pressurized at their unloaded length, which is unusual for native arteries. Data from the loading phase of the cycle were analyzed for the stiffness of the HUA. The compliance was calculated using Compliance ð% per 100 mmHgÞ ¼

ðDsys  Ddias Þ  104 ; Ddias ðPsys  Pdias Þ

1308

TUAN-MU et al.

where Psys and Dsys are the systemic pressure and the corresponding outer diameter of the HUA respectively, and Pdias and Ddias are the diastolic pressure and the corresponding outer diameter of the HUA respectively. Herein, the compliance was calculated between 70 and 120 mmHg. As the compliance can be influenced by the wall thickness of the HUA, it is more appropriate to examine mechanical behavior of the HUA in terms of stress–stretch relationship. The determination of stress requires dimensions of the HUA at its deformed state. The unloaded thickness of the HUA, H, was measured from the histological section of the HUA using ImageJ (NIH) to determine the wall volume of the HUA. That is, V ¼ pðB2  A2 ÞL; where B is the unloaded outer radius, A(= B 2 H) is the unloaded inner radius, and L is the unloaded length. Although not measurable, the deformed inner radius, a, at any deformed state can be computed given the on-line measurement of deformed outer radius, b, with the assumption of incompressibility. That is, sffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi  2  B  A2 a ¼ b2  ; kz where kz is the axial stretch ratio (l/L) and l is the deformed length of the HUA. Once a and b are known, the pressure–diameter data can be used to calculate the mean circumferential stress, rh, as rh ¼

Pa ; h

where P is the luminal pressure, and h(= b 2 a) is the deformed wall thickness. The associated mean circumferential stretch ratio, kh, can be computed as kh ¼

ða þ bÞ=2 ðA þ BÞ=2

HUAs from another five donors were used to measure the burst pressure and the suture retention strength of fresh HUAs, decellularized fresh HUAs, frozen HUAs, and decellularized frozen HUAs (N = 5 for each treatment). The burst pressure and suture retention strength were examined using the same custom-built mechanical tester except that a 100 psi pressure transducer (Model 80A, 0–100 psi, Sensermate, Taiwan) and a 50 N load cell (MOB-10, Transducer Technique, Temecula, CA) were used as larger maximum pressure and force were expected. The burst pressure of HUAs, the maximum load that the specimen could bear before failure, was measured after

preconditioning. Suture retention tests were performed on ~20-mm long HUA segments. A 6-0 polypropylene suture connected to the load cell was placed 2 mm from one end of the segment. The other end was fixed by cannulation. The segment was stretched at an extension rate of 150 mm/min until the suture pulled through the HUA. Statistical Analysis Data are presented as mean ± standard deviation. Differences in mechanical properties of fresh HUAs, decellularized fresh HUAs, frozen HUAs, and decellularized frozen HUAs were examined by two-way ANOVA with repeated measures in conjunction with Holm-Sidak post hoc procedure. Significance level was set at p < 0.025.

RESULTS Figure 1 reveals that nuclei still remained on the sections of fresh HUAs treated by 0.1% and 0.5% SDS solutions for 48 h and that apparently no nuclei left on the section of fresh HUAs treated by 1% SDS solution for 48 h. It is noteworthy that nuclei still remained on the section of frozen HUAs treated by 1% SDS solution for 48 h. Less intense eosin staining (i.e., less pink) was observed on the sections of decellularized HUAs, indicating a loss of ECM components. The intensity of Alcian blue staining appeared to decrease on the sections of decellularized fresh HUAs, frozen HUAs, and decellularized frozen HUAs, indicating that GAGs were removed by freezing or decellularization (Figs. 2a–2d). Results of image quantification also showed that the GAG content in decellularized fresh HUAs, frozen HUAs, and decellularized frozen HUAs was significantly lower than that in fresh HUAs (Fig. 2e). Figure 3 shows that residual DNA in fresh HUAs treated by 1% SDS solution was significantly lower than that in fresh HUAs treated by 0.1% SDS solution for all of the time points (N = 5). It appeared that 1% SDS solution was the most effective in HUA decellularization compared to 0.5% SDS and 0.1% SDS solutions; no residual DNA was detected in fresh HUAs treated by 1% SDS solution for 48 h. The DNA quantification supported the histological findings that cells in fresh HUAs were completely removed after being treated in 1% SDS solution for 48 h. Figure 4 shows that when fresh HUAs and frozen HUAs underwent the optimal decellularization protocol (i.e., 1% SDS solution for 48 h) the residual DNA in decellularized fresh HUAs was significantly lower than that in decellularized frozen HUAs (N = 3). Note that

On the Decellularization of Fresh or Frozen HUAs

(a)

1309

(b)

L

(c)

L

L

(d)

L

(e)

L

FIGURE 1. Representative H&E stained sections of fresh HUAs (a) and those treated by 0.1% (b), 0.5% (c), and 1% (d) SDS for 48 h. A representative H&E stained section of frozen HUAs treated by 1% SDS solution for 48 h (e). Insets show areas with higher magnitude. L denotes the lumen of the artery. Solid arrows indicate cells. Scale bar represents 100 lm.

0.13 ± 0.03 ng/mg residual DNA still remained in frozen HUAs treated by 1% SDS solution for 48 h. PSR images revealed that collagen intensity on the section of decellularized fresh HUAs was weaker than that on the section of fresh HUAs (Figs. 5a and 5b). Also, collagen intensity on the section of decellularized frozen HUAs was weaker than that on the section of frozen HUAs (Fig. 5c and 5d). Image quantification of the PSR images showed that collagen content of HUAs reduced significantly after decellularization for both fresh and frozen HUAs while no significant differences in collagen content were found between fresh and

frozen HUAs (Fig. 5e); results of collagen staining intensity appeared to be similar (Fig. 5f). It is noteworthy that SHG images illustrated that decellularization but not freezing decreased undulation of collagen fibers; this phenomenon appeared to be more apparent when frozen HUAs were decellularized (Figs. 6a–6d). Image quantification of the SHG images showed that decellularization but not freezing reduced collagen intensity (Fig. 6e), consistent with the finding of Fig. 5f. Figures 5 and 6 also show that freezing altered the morphology of HUAs. Specifically, PSR staining

1310

TUAN-MU et al.

(b)

(a)

L L (d)

(c)

L

L

(e) Relative GAG Content

0.08

0.06

*

*

*

0.04

0.02

0.00

0 UAs UAs UAs UAs nH nH sh H sh H roze c. froze Fre ec. fre F D De

FIGURE 2. Representative Alcian blue stained sections of fresh HUAs (a), decellularized fresh HUAs (b), frozen HUAs (c) and decellularized frozen HUAs (d). Comparisons of relative GAG content among the four groups (e; N 5 3). L denotes the lumen of the artery. Data are presented as mean 6 SD. Scale bar represents 100 lm. *p < 0.025.

illustrated that collagen was condensed at the vicinity of the pores on the section of frozen HUAs, indicating that pore formation may have compressed the surrounding collagen in frozen HUAs (Fig. 5c). Also, the intensity of SHG was found to be more intense around the pores on the section of frozen HUAs (Fig. 6c). Figure 7a shows pressure–diameter curves of fresh HUAs before and after decellularization (fresh HUAs vs. decellularized fresh HUAs; N = 7). The pressure– diameter curve of HUAs only slightly shifted to the left after decellularization; the two curves are not

significantly different. The pressure–diameter curve of a blood vessel represents its structural stiffness; it is influenced by the wall thickness of the vessel. We thus prepared stress–stretch plots for the comparison of material stiffness. Figure 7b shows the stress–stretch curves of fresh HUAs before and after decellularization. No significant differences were found in the stress– stretch curves, indicating that the material stiffness did not change significantly after decellularization. While decellularization had little effect on the stiffness of fresh HUAs, the stiffness of HUAs changed

On the Decellularization of Fresh or Frozen HUAs

Residual DNA (ng/mg)

2.5 2.0

0.1% SDS 0.5% SDS 1% SDS

*†

1.5

*

1.0

*

0.5

*

*

0.0 12

24

36

48

Duration of decellularization (hours) FIGURE 3. Time course of residual DNA in fresh HUAs treated by 0.1, 0.5 and 1% SDS solutions. Data are presented as mean 6 SD with N 5 5 per concentration per time point. *p < 0.025 vs. 1% SDS;  p < 0.025 vs. 0.5% SDS.

Residual DNA (ng/mg)

2.5 Fresh HUAs Frozen HUAs

2.0 1.5

1311

observed (Figs. 7e and 7f; N = 7). Surprisingly, the pressure–diameter and the stress–stretch relationships of decellularized frozen HUAs appeared to be similar to those of decellularized fresh HUAs. Representative pressure–diameter and stress–stretch curves of fresh HUAs, decellularized fresh HUAs, frozen HUAs, and decellularized frozen HUAs are shown in Supplemental Fig. 1. Serving as a basis for comparison with other studies, the compliance of the HUAs was calculated and analyzed. The compliance of frozen HUAs was found to be significantly greater than that of both fresh HUAs and decellularized fresh HUAs (Fig. 7g). Figure 8 shows the burst pressure and the suture retention strength of fresh HUAs, decellularized fresh HUAs, frozen HUAs, and decellularized frozen HUAs. The burst pressure decreased significantly after decellularization for either fresh HUAs or frozen HUAs; no significant differences between fresh HUAs and frozen HUAs were found, however (Fig. 8a). There were no significant differences in the suture retention strength among the four groups (Fig. 8b). Table 1 shows the unloaded wall thickness and the inner diameter of HUAs. There were no significant differences in the wall thickness and the inner radius among the four groups.

1.0

DISCUSSION

0.5

* 0.0 12

24

36

48

Duration of decellularization (hours) FIGURE 4. Time course of residual DNA in fresh HUAs and frozen HUAs treated by 1% SDS solution. Data are presented as mean 6 SD with N 5 3 per group per time point. *p < 0.025.

significantly after freezing. Figures 7c and 7d show the structural stiffness and the material stiffness of fresh HUAs before and after freezing (fresh HUAs vs. frozen HUAs; N = 7). Both pressure–diameter and stress–stretch curves shifted to the right after freezing; that is, frozen HUAs manifested a softer mechanical behavior than fresh HUAs. Statistical analysis showed significant differences between fresh HUAs and frozen HUAs in pressure–diameter curves (70–150 mmHg) and in stress–stretch curves (50–100 kPa). Unlike the non-significant effect of decellularization on the stiffness of fresh HUAs, both pressure–diameter and stress–stretch curves shifted to the left significantly when frozen HUAs were decellularized. Significant differences between frozen HUAs and decellularized frozen HUAs in pressure–diameter curves (70–150 mmHg) and in stress–stretch curves (30–100 kPa) were

We optimized the decellularization protocol involving only SDS solution as our first step. SDS solution only was shown not to be effective in arterial cell removal compared to decellularization methods involving both enzymes and SDS solution.14,17 The SDS solutions used in these studies, however, were essentially hypertonic17 or isotonic.14 Consistent with our finding, Rosario et al.31 found that 0.1% SDS in hypotonic solution is more effective in decellularization of porcine urinary bladder than 1 M NaCl hypertonic solution. Note that osmotic shock has been used as a common method for tissue decellularization.15 For example, tissues were treated sequentially using hypotonic and hypertonic solutions for decellularization.6 Hypotonic solutions effectively enhance cell lysis47 while hypertonic solutions dissociate DNA from proteins.9 Additional enzymatic or chemical agents are required to further remove cellular residues, however. The use of fresh HUAs, compared to frozen ones, was found to be better with respect to the efficiency of decellularization. The decreased efficiency of decellularization in frozen HUAs may be attributed to the condensed ECM caused by ice crystal formation during freezing. Venkatasubramanian et al.44 found

1312

TUAN-MU et al.

(b)

(a)

L

L (c)

(d)

L

L (f) 1.0 0.8

* *

*

0.6 0.4 0.2 0.0

0 UAs UAs UAs UAs nH nH sh H sh H roze c. froze Fre ec. fre F D De

Collagen Staining Intensity

Relative Collagen Content

(e)

140 120 100

*

* *

*

80 60 40 20 0

0 UAs UAs UAs UAs n H zen H sh H fresh H e e z r o o F ec. Fr ec. fr D D

FIGURE 5. Representative PSR stained sections of fresh HUAs (a), decellularized fresh HUAs (b), frozen HUAs (c), and decellularized frozen HUAs (d). Significant pore formation (arrowheads) and condensed collagen surrounding the pores (arrows) were found on the section of frozen HUAs. Comparisons of relative collagen content (e) and collagen staining intensity (f) among the four groups (N 5 3). L denote lumen of the artery. Data are presented as mean 6 SD. Scale bar represents 100 lm. *p < 0.025.

nuclear condensation in a freezing process. Although not examined in this study, nuclear condensation, similar to the ECM condensation, may also decrease mass transfer and compromise the efficiency of decellularization. It is interesting to note that Ketchedjian et al.25 tested cryopreserved, cryopreserved/ decellularized, and fresh/decellularized pulmonary trunks as patches for reconstruction of great vessel defects. While decellularization was found to be essential for reducing antigenicity and calcification, the

influence of the use of fresh or frozen tissues on the efficiency of decellularization was not investigated. To the best of our knowledge, this is the first study observing that the efficiency of decellularization was influenced by freezing. Pressure–diameter data are essentially structure related. That is, a thicker HUA expands less than a thinner one when pressurized. The stress–stretch curves, on the other hand, represent the material stiffness of the HUAs.13 The greater variance in the

On the Decellularization of Fresh or Frozen HUAs

(a)

1313

(b)

L

L

L

(c)

Collagen Intensity

(e) 80 60

*

(d)

*

L

*

40

20

0

0 UAs HUAs UAs UAs n nH sh H sh H roze c. froze Fre ec. fre F D De

FIGURE 6. Representative SHG images of fresh HUAs (a), decellularized fresh HUAs (b), frozen HUAs (c), and decellularized frozen HUAs (d). Significant pore formation (arrowheads) and condensed collagen surrounding the pores (arrows) were found on the section of frozen HUAs. Comparisons of the collagen intensity among the four groups (e; N 5 3). L denotes the lumen of the artery. Data are presented as mean 6 SD. Scale bar represents 100 lm. *p < 0.025. The insets in each image (403) illustrate the undulation of the collagen fibers (arrows with dot line).

pressure–diameter curves (compared to the stress– stretch curves) might be due to the variance of wall thickness among specimens. Because the wall thickness

varied among HUAs, the highest stress, which was converted from the maximum pressure in the test (i.e., 150 mmHg) based on the Laplace law, varied among

1314

TUAN-MU et al.

(b)

160

Pressure (mmHg)

140 120 100 80 60 40 Fresh HUAs Dec. fresh HUAs

20 0

Circumferential Stress (kPa)

(a)

120 100 80 60 40 20 0

1.00 1.02 1.04 1.06 1.08 1.10 1.12 1.14 1.16

(d) 120

140

Pressure (mmHg)

1.00 1.02 1.04 1.06 1.08 1.10 1.12 1.14 1.16

120

*

100 80 60 40

Fresh HUAs Frozen HUAs

20 0

Circumferential Stress (kPa)

(c) 160

100 80

40 20 0

120

*

80 60 40 Frozen HUAs Dec. frozen HUAs

20 0

1.00 1.02 1.04 1.06 1.08 1.10 1.12 1.14 1.16

Compliance (% per 100 mmHg)

1.00 1.02 1.04 1.06 1.08 1.10 1.12 1.14 1.16

OD/OD0

Circumferential Stress (kPa)

Pressure (mmHg)

140

(g)

Fresh HUAs Frozen HUAs

(f) 120

160

100

*

60

1.00 1.02 1.04 1.06 1.08 1.10 1.12 1.14 1.16

(e)

Fresh HUAs Dec. fresh HUAs

100 80

*

60 40 20

Frozen HUAs Dec. frozen HUAs

0 1.00 1.02 1.04 1.06 1.08 1.10 1.12 1.14 1.16

Circumferential Stretch

8

* 6

*

4

2

0

0 UAs UAs UAs UAs n H zen H sh H fresh H e z e r o o F ec. Fr ec. fr D D

FIGURE 7. Comparisons of pressure–diameter curves (a, c, and e), circumferential stress–stretch curves (b, d, and f) and compliance (g) among the four groups (N 5 7). Data are presented as mean 6 SD. *p < 0.025.

specimens (100–180 kPa). Simply for statistical comparison purposes, we presented our stress–stretch data up to 100 kPa, which is much smaller than maximum

stress shown in previous studies (up to 20 MPa).12,14,33 Note that the ratio of wall thickness to inner diameter is relatively high in HUAs compared to other arteries,

On the Decellularization of Fresh or Frozen HUAs

leading to lower circumferential stress given the same pressure. Also note that the wall thickness measured from histology may not be as precise as that obtained using other methods. The process of fixation may lead to shrinkage of the vessel and hence overestimate the stress. As all the HUAs (fresh or processed) were subjected to the same procedure for histology, the shrinkage should not affect the comparison of stress– stretch behavior of the HUAs. The freezing process appeared to have significant influence on the stiffness of HUAs. As both collagen content and fiber undulation did not change after freezing, the reduced stiffness may be attributed to pore formation in frozen HUAs. That the frozen

(a) Burst Pressure (mmHg)

1400 1200

*

*

*

1000 800 600 400 200 0

Suture Retention Strength (g)

UAs UAs sh H fresh H e r F . c De

(b)

*

0 As UAs n H zen HU e z o o r f Fr . Dec

100 80 60 40 20 0

UAs HUAs h sh H Fre c. fres De

s s 0 HUA n HUA e zen Fro c. froz De

FIGURE 8. Comparisons of burst pressure (a) and suture retention strength (b) among the four groups (N 5 5). Data are presented as mean 6 SD. *p < 0.025.

1315

HUAs were significantly more compliant than their fresh counterparts was consistent with the finding of Blondel et al.7 Some studies, however, reported that cryopreservation26,30 or freezing2,39 have no significant effects on the mechanical properties of blood vessels while others reported that blood vessels become stiffer after cryopreservation.14,40 The controversy may be due to the different arterial structure, which depends on the specific location of vessels within the vasculature. Indeed, Rosset et al.32 reported that common carotid arteries are significantly stiffer after cryopreservation, but not superficial femoral arteries. Interestingly, the burst pressure and the suture retention strength of HUAs did not change after freezing. The results were consistent with many other studies.18,29,39 Consistent with our finding that decellularization did not change the stiffness of fresh HUAs, Gui et al.17 evaluated the mechanical properties of HUAs before and after decellularization by pressure–diameter tests and found slight shift in the stress–strain curve while maximum stress, elastic modulus and compliance did not change significantly after decellularization. Also, Zou and Zhang48 determined the mechanical properties of porcine thoracic aorta before and after decellularization by equibiaxial planar tensile tests and reported no significant changes in stress–strain curves after decellularization. Although many previous studies reported that the compliance of the arteries decreases significantly after decellularization,12,33,36,45 the arteries they used are all elastic arteries which are essentially different from muscular HUAs. Frozen HUAs, however, seemed to be more sensitive to decellularization as the shift in stress–stretch curves was greater than that of fresh HUAs. The change of the stiffness might be attributed to the removal of smooth muscle cells as well as some relatively loose ECM components during decellularization.17 Note that the reduced undulation of collagen fibers in decellularized frozen HUAs may also explain the reduced compliance. Decellularization has been shown to reduce collagen fiber undulation in various blood vessels and hence to increase their stiffness.45,48 Recently, Merina et al.45 further established a strong correlation between waviness of collagen fibers and mechanical properties in cardiac tissues. Surprisingly, no significant differences in compliance were observed between decellularized fresh HUAs and decellularized

TABLE 1. The unloaded wall thickness and inner radius of the HUAs.

Wall thickness (lm) Inner radius (mm)

Fresh HUAs

Decellularized HUAs

Frozen HUAs

Decellularized frozen HUAs

234.42 ± 48.89 1.44 ± 0.26

205.46 ± 24.55 1.47 ± 0.32

216.86 ± 37.42 1.45 ± 0.28

210.62 ± 40.70 1.49 ± 0.28

Data were presented as mean ± SD.

1316

TUAN-MU et al.

frozen HUAs although decellularized frozen HUAs appeared to be slightly more compliant than decellularized fresh HUAs. The reduction in burst pressure due to decellularization may be attributed to the reduced collagen content. The finding was consistent with that of Xiong et al.,46 who showed that the burst pressure of porcine saphenous arteries significantly reduces after decellularization. Some studies, however, reported that the burst pressure does not change significantly after decellularization17,29; the controversy may be due to different decellularization methods used in these studies, which possibly have different effects on ECM components.15 Despite its decrease after decellularization, the burst pressure was still three times greater than the physiological blood pressure. The maximum pressure we used for the preconditioning and in our pressure–diameter tests was 150 mmHg which is significantly higher than the physiological pressure of HUAs (~50 mmHg)8 and might potentially damage the HUAs. Note, however, that the mechanical behavior of HUAs after preconditioning was found to be repeatable. Moreover, since decellularized HUAs are to be used as a scaffolding material for small-diameter TEVGs, the mechanical properties examined in the range of physiological blood pressure of systemic circulation are desired. As to the decellularization method, we tested only SDS solution in this study. Although SDS is an effective detergent for decellularization, it tends to remove GAGs and disrupt ultrastructure of tissues.10 Many other agents were reported to be effective in arterial decellularization and their efficiency of decellularization on frozen HUA may be worth investigation. Nevertheless, it is unlikely that the difficulty of removing residual DNA from the condensed matrix (caused by freezing) can be resolved by other decellularization methods. Finally, we did not use any cryoprotectants when freezing the HUAs as they may interfere the subsequent decellularization and complicate data interpretation. Note that cryoprotectants are always used in conventional cryopreservation. The freezing in conventional cryopreservation, however, inevitably damages the structure of ECM even with cryoprotectants.27,37 In a preliminary study, we found that, as expected, freezing with 15% DMSO solution did not alleviate pore formation (Supplemental Fig. 2). It is worth noting that vitrification has been used to cryopreserve complex, multicellular tissues. Unlike the conventional cryopreservation that involves freezing, vitrification essentially prevents nucleation and growth of ice crystals and thus better preserves the ECM in tissues. Nevertheless, significant amount of work on preserving tissues still involves freezing; for example,

Abousleiman et al.1 and Crouzier et al.11 used frozen human umbilical vessels to prepare their decellularized matrix. CONCLUSION In this study, we found that freezing decreased the efficiency of decellularization and decellularization reduced the compliance of both fresh HUAs and, more significantly, frozen HUAs. Surprisingly, the stiffness of decellularized frozen HUAs was similar to that of decellularized fresh HUAs. Although little difference in the stiffness was observed, we suggest avoiding freezing if more efficient and complete decellularization is desired. ELECTRONIC SUPPLEMENTARY MATERIAL The online version of this article (doi: 10.1007/s10439-014-1000-1) contains supplementary material, which is available to authorized users.

ACKNOWLEDGMENT This research was supported by grants from the National Science Council (NSC-100-2221-E-006-097) and the National Health Research Institute (NHRIEX102-10217EC) in Taiwan.

REFERENCES 1

Abousleiman, R. I., Y. Reyes, P. McFetridge, et al. The human umbilical vein: a novel scaffold for musculoskeletal soft tissue regeneration. Artif. Organs 32:735–742, 2008. 2 Adham, M., J. P. Gournier, J. P. Favre, et al. Mechanical characteristics of fresh and frozen human descending thoracic aorta. J. Surg. Res. 64:32–34, 1996. 3 Bakhach, J. The cryopreservation of composite tissues: principles and recent advancement on cryopreservation of different type of tissues. Organogenesis 5:119–126, 2009. 4 Ballyk, P. D., C. Walsh, J. Butany, et al. Compliance mismatch may promote graft-artery intimal hyperplasia by altering suture-line stresses. J. Biomech. 31:229–237, 1998. 5 Bergmeister, H., M. Strobl, C. Grasl, et al. Tissue engineering of vascular grafts European Surgery. Acta Chirurgica Austriaca 45:187–193, 2013. 6 Bishopric, N. H., L. Dousman, and Y. M. M. Yao. Matrix Substrate for a Viable Body Tissue-Derived Prosthesis and Method for Making the Same. Saint Paul: St. Jude Medical Inc, 1999. 7 Blondel, W. C. P. M., B. Lehalle, G. Maurice, et al. Rheological properties of fresh and cryopreserved human arteries tested in vitro. Rheol. Acta 39:461–468, 2000. 8 Couet, F., S. Meghezi, and D. Mantovani. Fetal development, mechanobiology and optimal control processes can

On the Decellularization of Fresh or Frozen HUAs improve vascular tissue regeneration in bioreactors: an integrative review. Med. Eng. Phys. 34:269–278, 2012. 9 Cox, B., and A. Emili. Tissue subcellular fractionation and protein extraction for use in mass-spectrometry-based proteomics. Nat. Protoc. 1:1872–1878, 2006. 10 Crapo, P. M., T. W. Gilbert, and S. F. Badylak. An overview of tissue and whole organ decellularization processes. Biomaterials 32:3233–3243, 2011. 11 Crouzier, T., T. McClendon, Z. Tosun, et al. Inverted human umbilical arteries with tunable wall thicknesses for nerve regeneration. J. Biomed. Mater. Res. A 89:818–828, 2009. 12 Dahl, S. L. M., J. Koh, V. Prabhakar, et al. Decellularized native and engineered arterial scaffolds for transplantation. Cell Transplant. 12:659–666, 2003. 13 Ferruzzi, J., M. R. Bersi, and J. D. Humphrey. Biomechanical phenotyping of central arteries in health and disease: advantages of and methods for murine models. Ann. Biomed. Eng. 41:1311–1330, 2013. 14 Fitzpatrick, J. C., P. M. Clark, and F. M. Capaldi. Effect of decellularization protocol on the mechanical behavior of porcine descending aorta. Int. J. Biomater., 2010. doi: 10.1155/2010/620503. 15 Gilbert, T. W., T. L. Sellaro, and S. F. Badylak. Decellularization of tissues and organs. Biomaterials 27:3675–3683, 2006. 16 Greenwald, S. E., and C. L. Berry. Improving vascular grafts: the importance of mechanical and haemodynamic properties. J. Pathol. 190:292–299, 2000. 17 Gui, L. Q., A. Muto, S. A. Chan, et al. Development of decellularized human umbilical arteries as small-diameter vascular grafts. Tissue Eng. Part A 15:2665–2676, 2009. 18 Hoenicka, M., K. Lehle, V. R. Jacobs, et al. Properties of the human umbilical vein as a living scaffold for a tissueengineered vessel graft. Tissue Eng. 13:219–229, 2007. 19 Hu, J. J., A. Ambrus, T. W. Fossum, et al. Time courses of growth and remodeling of porcine aortic media during hypertension: a quantitative immunohistochemical examination. J. Histochem. Cytochem. 56:359–370, 2008. 20 Hu, J. J., W. C. Chao, P. Y. Lee, et al. Construction and characterization of an electrospun tubular scaffold for small-diameter tissue-engineered vascular grafts: a scaffold membrane approach. J. Mech. Behav. Biomed. Mater. 13:140–155, 2012. 21 Hu, J. J., T. W. Fossum, M. W. Miller, et al. Biomechanics of the porcine basilar artery in hypertension. Ann. Biomed. Eng. 35:19–29, 2007. 22 Humphrey, J. D. Cardiovascular Solid Mechanics: Cells, Tissues, and Organs. New York: Springer, 2002. 23 Isenberg, B. C., C. Williams, and R. T. Tranquillo. Smalldiameter artificial arteries engineered in vitro. Circ. Res. 98:25–35, 2006. 24 Kannan, R. Y., H. J. Salacinski, P. E. Butler, et al. Current status of prosthetic bypass grafts: a review. J. Biomed. Mater. Res. Part B 74B:570–581, 2005. 25 Ketchedjian, A., A. L. Jones, P. Krueger, et al. Recellularization of decellularized allograft scaffolds in ovine great vessel reconstructions. Ann. Thorac. Surg. 79:888–896, 2005. 26 Masson, I., A. Fialaire-Legendre, C. Godin, et al. Mechanical properties of arteries cryopreserved at 280 degrees C and 2150 degrees C. Med. Eng. Phys. 31:825–832, 2009. 27 Muller-Schweinitzer, E. Cryopreservation of vascular tissues. Organogenesis 5:97–104, 2009. 28 Nseir, N., O. Regev, T. Kaully, et al. Biodegradable scaffold fabricated of electrospun albumin fibers: mechanical

1317

and biological characterization. Tissue Eng. Part C 19:257– 264, 2013. 29 Pellegata, A. F., M. A. Asnaghi, I. Stefani, et al. Detergentenzymatic decellularization of swine blood vessels: insight on mechanical properties for vascular tissue engineering. Biomed. Res. Int. 20:918753, 2013. 30 Pukacki, F., T. Jankowski, M. Gabriel, et al. The mechanical properties of fresh and cryopreserved arterial homografts. Eur. J. Vasc. Endovasc. Surg. 20:21–24, 2000. 31 Rosario, D. J., G. C. Reilly, E. A. Salah, et al. Decellularization and sterilization of porcine urinary bladder matrix for tissue engineering in the lower urinary tract. Regen. Med. 3:145–156, 2008. 32 Rosset, E., A. Friggi, G. Novakovitch, et al. Effects of cryopreservation on the viscoelastic properties of human arteries. Ann. Vasc. Surg. 10:262–272, 1996. 33 Roy, S., P. Silacci, and N. Stergiopulos. Biomechanical properties of decellularized porcine common carotid arteries. Am. J. Physiol. Heart Circ. Physiol. 289:H1567– H1576, 2005. 34 Schaner, P. J., N. D. Martin, T. N. Tulenko, et al. Decellularized vein as a potential scaffold for vascular tissue engineering. J. Vasc. Surg. 40:146–153, 2004. 35 Schutte, S. C., Z. Z. Chen, K. G. M. Brockbank, et al. Cyclic strain improves strength and function of a collagenbased tissue-engineered vascular media. Tissue Eng. Part A 16:3149–3157, 2010. 36 Sheridan, W. S., G. P. Duffy, and B. P. Murphy. Mechanical characterization of a customized decellularized scaffold for vascular tissue engineering. J. Mech. Behav. Biomed. Mater. 8:58–70, 2012. 37 Song, Y. C., B. S. Khirabadi, F. Lightfoot, et al. Vitreous cryopreservation maintains the function of vascular grafts. Nat. Biotechnol. 18:296–299, 2000. 38 Stegemann, J. P., S. N. Kaszuba, and S. L. Rowe. Review: advances in vascular tissue engineering using protein-based Biomaterials. Tissue Eng. 13:2601–2613, 2007. 39 Stemper, B. D., N. Yoganandan, M. R. Stineman, et al. Mechanics of fresh, refrigerated, and frozen arterial tissue. J. Surg. Res. 139:236–242, 2007. 40 Thakrar, R. R., V. P. Patel, G. Hamilton, et al. Vitreous cryopreservation maintains the viscoelastic property of human vascular grafts. FASEB J. 20:874–881, 2006. 41 Vaz, C. M., S. van Tuijl, C. V. C. Bouten, et al. Design of scaffolds for blood vessel tissue engineering using a multilayering electrospinning technique. Acta Biomater. 1:575– 582, 2005. 42 Veith, F. J., S. K. Gupta, E. Ascer, et al. 6-Year prospective multicenter randomized comparison of autologous saphenous-vein and expanded polytetrafluoroethylene grafts in infrainguinal arterial reconstructions. J. Vasc. Surg. 3:104– 114, 1986. 43 Veith, F. J., C. M. Moss, S. Sprayregen, et al. Preoperative saphenous venography in arterial reconstructive surgery of the lower-extremity. Surgery 85:253–256, 1979. 44 Venkatasubramanian, R. T., E. D. Grassl, V. H. Barocas, et al. Effects of freezing and cryopreservation on the mechanical properties of arteries. Ann. Biomed. Eng. 34: 823–832, 2006. 45 Williams, C., J. Liao, E. M. Joyce, et al. Altered structural and mechanical properties in decellularized rabbit carotid arteries. Acta Biomater. 5:993–1005, 2009. 46 Xiong, Y., W. Y. Chan, A. W. C. Chua, et al. Decellularized porcine saphenous artery for small-diameter tissueengineered conduit graft. Artif. Organs 37:E74–E87, 2013.

1318 47

TUAN-MU et al.

Xu, C. C., R. W. Chan, and N. Tirunagari. A biodegradable, acellular xenogeneic scaffold for regeneration of the vocal fold lamina propria. Tissue Eng. 13:551–566, 2007.

48

Zou, Y., and Y. H. Zhang. Mechanical evaluation of decellularized porcine thoracic aorta. J. Surg. Res. 175:359– 368, 2012.

On the decellularization of fresh or frozen human umbilical arteries: implications for small-diameter tissue engineered vascular grafts.

Most tissues, including those to be decellularized for tissue engineering applications, are frozen for long term preservation. Such conventional cryop...
6MB Sizes 0 Downloads 3 Views