Skip to main content
NCBI home page
Organogenesis logoLink to Organogenesis
. 2015 Mar 2;11(1):30–45. doi: 10.1080/15476278.2015.1022009

Freezing/Thawing without Cryoprotectant Damages Native but not Decellularized Porcine Renal Tissue

Nafiseh Poornejad 1, Timothy S Frost 1, Daniel R Scott 1, Brinden B Elton 1, Paul R Reynolds 2, Beverly L Roeder 3, Alonzo D Cook 1,*
PMCID: PMC4594613  PMID: 25730294

abstract

Whole organ decellularization of porcine renal tissue and recellularization with a patient's own cells would potentially overcome immunorejection, which is one of the most significant problems with allogeneic kidney transplantation. However, there are obstacles to achieving this goal, including preservation of the decellularized extracellular matrix (ECM), identifying the proper cell types, and repopulating the ECM before transplantation. Freezing biological tissue is the best option to avoid spoilage; however, it may damage the structure of the tissue or disrupt cellular membranes through ice crystal formation. Cryoprotectants have been used to repress ice formation during freezing, although cell toxicity can still occur. The effect of freezing/thawing on native (n = 10) and decellularized (n = 10) whole porcine kidneys was studied without using cryoprotectants. Results showed that the elastic modulus of native kidneys was reduced by a factor of 22 (P < 0.0001) by freezing/thawing or decellularization, while the elastic modulus for decellularized ECM was essentially unchanged by the freezing/thawing process (p = 0.0636). Arterial pressure, representative of structural integrity, was also reduced by a factor of 52 (P < 0.0001) after freezing/thawing for native kidneys, compared to a factor of 43 (P < 0.0001) for decellularization and a factor of 4 (P < 0.0001) for freezing/thawing decellularized structures. Both freezing/thawing and decellularization reduced stiffness, but the reductions were not additive. Investigation of the microstructure of frozen/thawed native and decellularized renal tissues showed increased porosity due to cell removal and ice crystal formation. Orcein and Sirius staining showed partial damage to elastic and collagen fibers after freezing/thawing. It was concluded that cellular damage and removal was more responsible for reducing stiffness than fibril destruction. Cell viability and growth were demonstrated on decellularized frozen/thawed and non-frozen samples using human renal cortical tubular epithelial (RCTE) cells over 12 d. No adverse effect on the ability to recellularize after freezing/thawing was observed. It is recommended that porcine kidneys be frozen prior to decellularization to prevent contamination, and after decellularization to prevent protein denaturation. Cryoprotectants may still be necessary, however, during storage and transportation after recellularization.

Keywords: biomechanical properties, cryopreservation, decellularized ECM, freezing/thawing, kidney, renal tissue

Introduction

Approximately one in 15 Americans, nearly 20 million people, are afflicted with chronic kidney disease and many of those diagnosed progress to kidney failure.1 The two treatments available for kidney failure are dialysis and kidney transplantation. An organ transplant is the preferred method among patients with kidney failure, and is fairly successful as 83% of patients who receive a kidney transplant are still alive 5 y after the transplant.2 However, relatively few kidneys are available for patients to receive.

Tissue engineered organs are a promising alternative for whole organ transplantation.3–7 Creating a transplantable, bioengineered organ to be used for those with chronic kidney failure hinges on the ability to develop a viable extracellular matrix (ECM) to give basic structure to the organ.8–9 The ECM serves as scaffolding that provides both structure and signals to the cells that attach to it upon recellularization.4,5 The ECM for an organ, especially a kidney, is extremely difficult to create synthetically due to its complex composition of amino acids, collagens, glycoproteins, glycosaminoglycans (GAGs), and sensitive microstructures needed for filtration.8–10 The most efficient method to produce this ECM is by decellularizing a native mammalian organ. This decellularization process strips the ECM of all living cells leaving just the collagenous structure of the ECM with greatly reduced immunogenic potential.10–12 The ECM can then be repopulated with cells harvested from the recipient to create a hybrid organ transplant that will have minimal probability of being rejected as the organ is recellularized using the recipient's own cells.13,14

One of the problems in creating these hybrid organs is the transportation and maintenance of the organ before transplantation. One possible process is to prepare the decellularized ECM scaffold, freeze the ECM until needed, and then proceed to thaw and recellularize the ECM with human cells when needed. Porcine kidneys can be used since they are comparable to human kidneys in size and structure. Understanding the effects of the freezing and thawing cycles is critical to allow this process to be plausible. Freezing the kidneys and thawing them when needed prevents bacterial growth, preserves protein structure, and allows the ECM to last for longer durations of time, making it a more viable option to be used for organ transplant. However, one concern is that freezing/thawing may cause damage to the microstructure of the ECM due to ice crystal formation that would disrupt the ability to perfuse the kidneys under pressure.15

There are several previous studies that have dealt with different methods to cryopreserve whole organs before being transplanted.16–19 One method is vitrification, which is a process that uses high concentrations of cryoprotectants to convert living biological materials into a glassy state without ice crystal formation. Fahy et al. used vitrification to cryopreserve a whole rabbit kidney prior to transplantation.19 They reported several obstacles for using high concentrations of cryoprotectants since these materials might be toxic. However, they were able to preserve the structure and function of the whole kidney. This research group also studied cryopreservation of slices of rat kidneys and livers through rapid freezing and warming using 18% dimethyl sulfoxide (DMSO), and commercially available cryoprotectant mixtures.20 Vitrification of both liver and kidney slices using these cryoprotectants was not successful and histological evaluation demonstrated irreversible damage to the structure even without ice crystal formation.

The effects of cryoprotection on mechanical properties has also been studied in many other tissues. For example, Woo et al. determined the effect of freezing on ligament tensile behavior.21 They performed freezing/thawing cycles and observed no significant change in stress-strain curves, tensile strength and ultimate strain of ligaments following freezing compared to non-frozen ligaments. In another study, Chan et al. measured the effects of cryogenic grinding on soft-tissue optical properties of calf aorta, rat jejunum, and rabbit sciatic nerve.22 They reported that within the 400-nm to 850-nm spectrum, optical properties of the tissues were approximately intact. Nonaka et al. also determined freezing/thawing cycle effects on decellularized rat lungs by mechanical properties measurements.23 They observed no changes in resistance and elasticity on lung tissues after freezing and thawing.

To our knowledge, there is no previous study for evaluation of freezing/thawing cycle effects on decellularized kidneys compared to native renal tissue. Therefore, the purpose of this paper was to determine the extent of damage due to cycles of freezing and thawing compared to the decellularization process. In this study, freezing/thawing cycles were used without any cryoprotectant, freezing at −20°C for at least 24 h, cooling to −80°C for 12 h, slow warming at −20°C for another 24 h, and finally thawing slowly at 4°C for 12 h. Compression and arterial pressure measurements were used to evaluate structural integrity of the ECM. Microstructures of renal tissue were examined through histological staining and scanning electron microscopy (SEM). Frozen/thawed and non-frozen decellularized ECM samples were also recellularized with human renal cortical tubular epithelial (RCTE) cells to compare the capacity of ECM to support cell attachment and proliferation.

Results

In this study, the microstructure and mechanical properties of native porcine kidneys (n = 10) and decellularized porcine ECM (n = 10) were examined to investigate the effect of freezing/thawing on the tissue. The 20 porcine kidneys were analyzed as 4 groups: native non-frozen, native frozen/thawed, decellularized non-frozen, and decellularized frozen/thawed (n = 5 per group). Some of the kidneys in the decellularized groups were stored at −20°C before decellularization.

Ten kidneys were decellularized by perfusion of sodium dodecyl sulfate (SDS) through the inherent vasculature of the kidney for 7 h. Cell removal was confirmed by hematoxylin and eosin (H&E) staining (Fig. 1) and no nuclear materials were detected in the decellularized ECM. Residual DNA quantification within the decellularized ECM also showed more than 98% removal of nuclear materials. Native (n = 5) and decellularized kidneys (n = 5) were subjected to a freezing/thawing cycle and tested for preservation of mechanical and microstructural properties, including compression (Figs. 2 and 3), arterial pressure (Fig. 4), SEM (Fig. 5), histology (Figs. 6–9) and cell viability (Fig. 10).

Figure 1.

Figure 1.

Open in a new tab

Native (A-1) and decellularized (B-1) porcine kidneys. Cells are visible in the native renal ECM (A-2). H&E staining confirmed complete cell removal from the decellularized ECM (B-2).

Figure 2.

Figure 2.

Open in a new tab

Stress/strain curves representing the upper and lower bounds for frozen/thawed and non-frozen samples (n = 20). Freezing/thawing significantly reduced the modulus of elasticity of native kidneys (P value < 0.0001); however, there was not an additive effect of freezing/thawing for decellularized renal ECM (P value = 0.0636).

Figure 3.

Figure 3.

Open in a new tab

Elastic moduli for frozen/thawed and non-frozen samples (n = 15-22 per group) were calculated as the slope of the linear part of the stress/strain curves at low strains. Freezing/thawing caused a significant reduction in elastic modulus of native kidneys (NK), while no significant reduction was observed for decellularized kidneys (DK). Data are reported in log-transformed format for a more clear comparison and reliable statistical analysis.

Figure 4.

Figure 4.

Open in a new tab

Arterial pressure measurements in frozen/thawed and non-frozen native kidneys (NK) and decellularized kidneys (DK) (n = 12) were indicative of renal vasculature integrity. Arterial pressure was highly reduced for NK after freezing/thawing (P < 0.0001), while it was essentially unchanged for DK (p = 0.18).

Figure 5.

Figure 5.

Open in a new tab

Scanning electron microscopy images of frozen/thawed and non-frozen kidney samples at 250X (A1-D1) and 500X (A2-D2) magnifications. No microstructural damage was detected after freezing/thawing. Renal corpuscles, distal and proximal tubules all were preserved during freezing/thawing cycles.

Figure 6.

Figure 6.

Open in a new tab

Orcein stain imaging for frozen/thawed and non-frozen native kidney samples at magnifications of 10X (A-1 & B-1) and 20X (A-2 & B-2). Generally, the elastin fibers were damaged as a result of freezing/thawing (affinity for Orcein stain color was lower for frozen/thawed samples which resulted in lighter color in images). Because of fibril damage the structure was more porous and had less integrity as indicated by the white spaces that were more frequent in frozen/thawed samples (arrows show renal corpuscles).

Figure 9.

Figure 9.

Open in a new tab

Sirius red stain imaging representative of general collagen for frozen/thawed and non-frozen decellularized kidney samples at 10X (A-1 & B-1) and 20X (A-2 & B-2) magnifications. General collagen seemed to be partially damaged through freezing/thawing since stain color was lighter for frozen/thawed samples; however, the difference was not as stark as for native kidneys after freezing/thawing. Also, the porosity was greater in frozen/thawed decellularized kidney samples, which is again indicative of fibril damage and diminished integrity (arrows show renal corpuscles).

Figure 10.

Figure 10.

Open in a new tab

H&E staining of recellularized non-frozen (A) and frozen/thawed (B) decellularized ECM with human RCTE cells after 12 d. Both non-frozen and frozen/thawed decellularized ECM showed the potential to support proliferation of human cells.

Compression test

To evaluate the mechanical properties of native and decellularized tissues, samples (n = 15-22 per group) of renal cortex were excised and subjected to compression. The stress/strain curves representing the upper and lower bounds of the measurements are shown in Figure 2. The elastic modulus representing the elasticity of renal tissue was calculated as the slope of the linear section of the stress/strain curves at low strain. The results of calculating the elastic moduli are shown in Figure 3. The average elastic modulus of decellularized kidneys was 6.4 ± 2.7 kPa before and 4.6 ± 3.0 kPa after freezing/thawing. The average elastic modulus of non-frozen native kidneys was 41.6 ± 22.4 kPa, which is in the range of previously reported averages.24 The elastic modulus of native kidneys was reduced 22 times (with 95% confidence interval of 12.4 - 32.9), to 5.6 ± 2.0 kPa, after being subjected to one freeze/thaw cycle (p-value < 0.0001). The data obtained after freezing/thawing are in complete agreement with Ternifi et al.'s observations of a decrease in renal cortex elastic modulus after storage at −18°C, −34°C, and −80°C.25

The average elastic modulus for decellularized ECM was approximately the same before and after being frozen/thawed (p = 0.0636); however, it was slightly smaller for frozen/thawed ECMs, which can be attributed to elastin and collagen fiber damage as demonstrated by elastin and collagen staining (Figs. 6-9). Similarly, Nonaka et al. reported no significant changes in mechanical properties of decellularized lungs after several freeze/thaw cycles.23 Both decellularization and freezing/thawing resulted in a decrease in stiffness, but the 2 processes were not additive in their damage to the mechanical properties of the porcine kidneys.

Arterial pressure measurement

All frozen/thawed and non-frozen kidneys (n = 3 per group) were perfused with 1X phosphate buffered saline to measure arterial pressure. PBS perfusion was performed over a range of flow rates to determine the preservation and integrity of renal capillaries in native kidneys (NK) and ECM from decellularized kidneys (DK) after being subjected to freezing/thawing cycles (see Fig. 4). The approximately linear trend of arterial pressure versus flow rate was preserved during freezing/thawing for both native and decellularized kidneys. The average vasculature pressure for non-frozen native kidneys (range: 40 – 70 mmHg) was significantly larger than for frozen/thawed native kidneys (range: 1 – 15 mmHg, p-value < 0.0001); however, this difference was not significant for decellularized ECMs (non-frozen range 3–30; frozen/thawed range: 2–29; p-value = 0.18). The average reduction in pressure after decellularization was 42.6 ± 2.2 mmHg (P < 0.0001), compared to the average reduction in pressure after freezing/thawing of 52.4 ± 3.5 mmHg (P < 0.0001) for native kidneys. In contrast, the effect of freezing/thawing on decellularized kidneys was only 4.2 ± 1.5 mmHg (P < 0.0001). These results are in agreement with Orlando et al.'s observations for non-frozen native and decellularized kidneys.26,27

Microstructure of frozen/thawed and non-frozen samples

Scanning electron microscopy of frozen/thawed and non-frozen kidney samples (n=4 per group) was performed and the results are shown in Figure 5. No detectable damage, in terms of dilation or increased porosity, was observed at the scale of these images. Renal glomeruli, distal and proximal convoluted tubules, and Bowman's capsules all were preserved during the freezing/thawing cycle.

Histological evaluation

Renal cortex biopsies of all frozen/thawed and non-frozen samples (n = 8 per group) were fixed in 4% paraformaldehyde solution (PFA), sectioned and stained with Orcein elastin and Sirius Red specific to elastic and general collagen tissue, respectively. Some damage to elastin occurred from the freezing/thawing process in native kidneys, as shown in Figure 6 at 10X and 20X magnification. Figure 7 shows elastin staining for frozen/thawed and non-frozen decellularized ECM with all imaging performed under the same conditions in terms of focus and light intensity. As a result of freezing/thawing, previously frozen samples stained a lighter color compared to non-frozen samples, which was attributed to a reduction in total elastin. However, the lighter color might also be due to ice formation and elastin fiber damage that occurred during freezing. As was also shown by the elastic modulus data (Fig. 3), the damage caused by freezing/thawing was more pronounced for native kidneys, and produced a greater color difference than was found with frozen/thawed and non-frozen decellularized samples.

Figure 7.

Figure 7.

Open in a new tab

Orcein stain imaging for frozen/thawed and non-frozen decellularized kidney samples at magnifications of 10X (A-1 & B-1) and 20X (A-2 & B-2). Generally, the elastin fibers were damaged as a result of freezing/thawing (affinity for Orcein stain color was lower for frozen/thawed samples which resulted in lighter color in images). Also it appeared that because of fibril damage the structure was more porous and had less integrity as white spaces were more frequent in frozen/thawed samples (arrows show renal corpuscles).

General collagen distribution of frozen/thawed and non-frozen native kidneys is shown in Figure 8 and the collagen distribution for decellularized kidneys can be seen in Figure 9. The red color represents general collagen, and was less detectable for previously frozen native or decellularized ECM compared to non-frozen samples. It is suspected that collagen fibers were damaged during the freezing/thawing cycle because of ice formation, leading to reduced stiffness of the ECM and reduced arterial pressure. In all the histology images, the arrows depict preserved renal corpuscles.

Figure 8.

Figure 8.

Open in a new tab

Sirius red stain imaging representative of general collagen for frozen/thawed and non-frozen native kidney samples at 10X (A-1 & B-1) and 20X (A-2 & B-2) magnifications. General collagen appeared to be partially damaged through freezing/thawing as evidenced by decreased stain color in frozen/thawed samples, which was detected as lighter colors. Void spaces were more detectable in frozen/thawed samples, which was again indicative of fibril damage and less integrity (arrows show renal corpuscles).

Human cell viability on the ECM

Repopulation of both frozen/thawed and non-frozen decellularized ECM (n = 3 per group) was performed using RCTEs. After 12 days, the repopulated ECM samples were stained with H&E for cell detection and the results are presented in Figure 10. As shown in this figure, there was no detectable difference between frozen/thawed and non-frozen ECM in terms of supporting cell attachment and growth. Cells proliferated and covered the whole surface in both cases.

Discussion

Freezing/thawing effects were studied for both native and decellularized porcine kidneys. Whole native and decellularized kidneys were sequentially frozen at −20°C for at least 24 h, then deep-frozen at -80°C for 12 h, followed by slow warming (without phase transition) at −20°C for 24 h, and finally thawed at 4°C for 12 h. The mechanical properties of all frozen/thawed and non-frozen, native and decellularized kidney samples were examined to assess the effect of ice crystal formation in the extracellular matrix. Previous research had demonstrated the detrimental effects of freezing/thawing on the biomechanical properties of native kidneys.25 Some loss of mechanical strength was observed for decellularized kidneys in preliminary experiments in our laboratory. One of the key motivations for this work was to determine if decellularized ECM would be further damaged by freezing to −80°C followed by thawing.

Mechanical properties of renal tissue were the most important characteristic for this evaluation, since the mechanical properties of biological tissue are essential for normal function and stability of an organ during recellularization. Previous researchers have focused on measurements of the biomechanical properties of abdominal organs24,28 but not frozen/thawed kidneys. Another reason for these experiments, which is critical for decellularized tissues, is that the stiffness of the ECM can dictate cell differentiation during repopulation with stem cells.29,30 Engler et al. showed that mesenchymal stem cells could be differentiated to brain cells on soft matrices, to muscle cells on stiffer matrices, and to bone cells on comparatively rigid matrices.29 Therefore it is desired to preserve the decellularized ECM stiffness and elasticity as close as possible to natural tissue.

Cell survival is essential for both whole organ transplantation and regenerative medicine. There are several previous papers that have described methods for safe preservation of organs before transplantation.31–33 In most of these methods high concentrations of cryoprotectants were used to avoid phase transition and crystal formation at low temperature (referred to as vitrification instead of freezing) while preserving cells. However, it has been shown that hearts and kidneys do not function properly after thawing if they are kept at temperatures lower than −20°C.34,35 In this study, the damage that occurs to native tissue from freezing/thawing without any cryoprotectants was assessed. It was found that frozen/thawed kidneys demonstrated significantly lower stiffness than non-frozen native kidneys, indicating that intact cells contribute significantly to mechanical properties. Elastic modulus reduction has also been observed for several other tissues after treatment with freezing/thawing such as porcine aortic tissue,36 decellularized lung,23 porcine kidney,25 rabbit tendons,37 and bovine liver.38

In addition to the cellular disruption caused by freezing and thawing, ice formation can damage both elastin and collagen fibers. Giannini et al. used TEM imaging to characterize collagen fibers of human posterior tibial tendons after being frozen at −80°C.39 They reported an increase in the mean of collagen fibrils' diameter, while the mean number of fibrils was also shown to be decreased. In another study, Chen et al. used H&E staining to assess the freezing/thawing cycle effect on Achilles tendons of rabbits.37 They observed more disordered collagen fibrils, and after several freezing/thawing cycles, gaps were apparent between tendon bundles because of ice crystal formation. However, they did not report any change in elastic modulus of their samples. All of these results are in complete agreement with what was observed in the present studies by histological staining representative of collagen and elastin.

In addition to collagen and elastin damage, increased porosity was detected after freezing and thawing. Several previous researchers have also noted this kind of dilation after freezing/thawing.36,38–42 For example, O'Leary et al. investigated the effect of long term freezing on the mechanical properties of porcine aortic tissue.36 They reported increased porosity and reduced density of the aortic tissue after being frozen and thawed. Chow et al. also reported such dilation for aortic tissue.41 They quantified the amount of total collagen and soluble collagen in their frozen and non-frozen samples, and reported a significant reduction or denaturation of collagen fibers in their frozen samples. Similarly, increased porosity has also been observed for frozen/thawed heart tissue.43

The arterial pressure experiments demonstrated a loss of integrity of the ECM structure, leading to decreased arterial pressures during perfusion. This was most likely due to the denaturation of collagen fibers, however, ice formation can also damage microcapillaries, and that might account for the highest pressure drop during perfusion. Another reason for reduced pressure drop is cellular plasma membrane disruption. This effect is intense enough that some researchers have used freezing/thawing for decellularization, or at least partial decellularization, of tissues and organs.7,9,44

Overall, cell and protein damage leading to increased porosity can account for the observed alterations in elastic modulus and arterial pressure that occurred after freezing/thawing. Porosity contributes more to loss of mechanical properties in native kidneys since the tissue is more densely packed prior to freezing/thawing. These effects were less important for decellularized ECM since porosity was intentionally created as a consequence of decellularization.

In summary, for decellularized organs, results from the current experiments predict that freezing and thawing without cryoprotectants will be acceptable both prior to and after decellularization. When organs are preserved for transplant, cyroprotectants are required to preserve the living cells and prevent proteins from denaturing. In the case of decellularized whole organs, it is not required to maintain cell viability, and there is minimal impact of freezing/thawing on the ECM proteins. Freezing may even prevent denaturation of the proteins during long-term storage prior to recellularization, and the reduced arterial pressure achieved by freezing/thawing kidneys before decellularization may improve the process by reducing SDS exposure time. The freeze/thaw process described herein is therefore expected to be satisfactory for whole porcine kidney decellularization. In support of these conclusions, recellularization studies with human RCTE cells on slices of frozen/thawed decellularized ECM demonstrated substantial cell growth. Future work includes investigating the recellularization of whole porcine kidneys after freezing and thawing.

Materials and Methods

Kidney retrieval

Porcine kidneys were obtained at a local abattoir using special care to preserve sufficient lengths of the renal vasculature. A catheter was inserted into the renal artery and the kidneys were perfused with heparinized 1X phosphate buffered saline (PBS) solution. The kidneys were also gently massaged during perfusion to ensure complete exposure to the heparin solution. The kidneys were then placed in a 5 gallon container of heparinized 1x PBS for transportation back to the laboratory. Upon arrival, some kidneys were frozen at −20°C, then later thawed and subjected to decellularization or deep frozen to −80°C as samples for the full freezing/thawing cycle; while other kidneys were used as controls for the characterization tests (native kidneys) without being frozen.

Decellularization and freezing/thawing cycle

To prepare kidneys for decellularization, fat and excess tissues were trimmed and the kidneys were cannulated via the renal artery with white nylon tubing (Value Plastics MTLS210–1, male luer slip to 200 series barbed coupler, 1/16" tube ID). Then the kidneys were perfused with a 0.5% solution of sodium dodecyl sulfate (SDS) for 7 hours for complete decellularization. The process began with low flow rates and the flow was gradually increased during the perfusion while keeping the arterial pressure below the natural diastolic pressure of the body (80 mmHg). After decellularization, the kidneys were perfused with deionized water for approximately 2 d to remove SDS detergent from the ECM.

To evaluate the freezing/thawing effect on native and decellularized ECM, several native and decellularized whole kidneys (n = 10) were subjected to freezing/thawing cycles as follows: the kidneys were frozen at −20°C for 24 h, then at −80°C for 12 h, then for 24 h at −20°C and finally, they were thawed at +4°C. These parameters were selected since they had previously been used to preserve lung tissue.23

Compression test

To compare the effects of freezing on the structural integrity of the extracellular matrix, an Instron 3342 Single Column Universal Testing System and Instron Model 1321 were used to perform compression tests. Samples (n = 15-22) of the renal cortex (10 mm × 10 mm with a height of 7 mm) were taken from both the native and decellularized kidneys for compression measurements. The control samples were compressed at a rate of 0.07 mm/sec until a compression force of 45 N was reached. The samples subjected to freezing cycles were frozen and thawed in the process described above and then subjected to the same compression tests. Using the compression data, the elastic moduli were then calculated for each sample at low stress values. All statistical analysis was performed using JMP Statistical Discovery Software from SAS.

Arterial pressure measurement

A simple apparatus was configured to measure the arterial pressure of the kidney. The kidney was cannulated through the renal artery and then connected to a peristaltic pump, which drew 1X PBS from a 1000 mL graduated cylinder. The pump was then set to the lower limit of 10 rpm. The pressure observed was recorded manually, and the flow rate was calculated by determining the time required to drain 10 mL from the graduated cylinder. The peristaltic pump speed was then incrementally increased by 5 rpm and the process was repeated. The pump speed was increased until it reached 70 rpm while maintaining less than the maximum pressure (< 80 mmHg). A total of 12 frozen/thawed and non-frozen native and decellularized kidneys were tested (n=3 per group).

Scanning Electron Microscopy

Five mm cube samples were taken from the cortex tissue of frozen/thawed and non-frozen, native and decellularized kidneys (n = 4 per group). Samples were subsequently soaked in 2% glutaraldehyde in a sodium cacodylate buffer (pH 7.3) for 24 h at room temperature. Samples were then rinsed in a buffer wash process involving 6 15 min washes in Millonig's phosphate buffer (pH 7.3) on a lab shaker. Samples were placed in a solution consisting of 25% sucrose and 10% glycerol in 0.05 M PBS for 2 h, washed in fresh solution, immersed in liquid nitrogen and fractured. Samples were then exposed to the Millonig's buffer wash process, followed by 1.5 h in a 1% OsO4, sodium cacodylate buffer (pH 7.3) solution. Samples were dehydrated using graded (10, 30, 50, 70, 95, 100%) ethanol baths inside critical dryer baskets. Each sample was submerged for 15 min up to 70% concentration ethanol solution and stored for several days at 4°C. The dehydration was completed with 70, 95 and 3 × 100% ethanol solution baths. Samples were then placed under 100% ethanol in a CO2 critical dryer and subsequently mounted on aluminum stands. Fifteen nm of a gold-palladium (Au-Pd) alloy were sputter-coated onto samples and images were taken using a scanning electron microscope (Phillips/FEI XL30ESEMSEG).

Histology

Excised renal cortex samples were taken from frozen/thawed and non-frozen, native and decellularized kidneys (n = 8 per group). The samples were fixed in 4% paraformaldehyde in 1X PBS for 12 h and partially dehydrated in 30%, 50%, and 70% ethanol prior to complete dehydration in an automated processor. The samples were then soaked in xylene and embedded in paraffin. Paraffin blocked samples were sectioned into 5 μm slices and placed on microscope slides. The sectioned samples were heated at 60°C for 12 h, dehydrated, deparaffinized, then stained either with standard hematoxylin and eosin (H&E, Thermo Scientific), Orcein (O7380; Sigma) or Picro-sirius red stain (Direct Red 80, Sigma-Aldrich) according to the manufacturer's instructions. H&E staining was performed to confirm there were no cellular components in the decellularized ECM. Orcein stain and Sirius red stain were used to examine elastin and collagen fibers, respectively. A standard light microscope equipped with a digital camera (Olympus America Inc..) at magnifications of 10X and 20X was used to image all stained samples.

Human cell viability and growth

Slices of frozen/thawed and non-frozen decellularized ECM (n = 3 per group) 5 cm in diameter were sterilized with 70% ethanol for 2 h. The samples were rehydrated in DMEM cell culture media supplemented with 10% FBS and 1% Pen-Strep (Gibco) at 37°C for 2 d before cell seeding. Human RCTEs were grown in cell culture media and were detached on the day of recellularization using TrypLE Express (Life Technology, Grand Island, NY). Approximately 8 ± 1.5 × 106 cells were seeded on each piece of tissue. The media was changed every other day and after 12 d the tissues were removed, sectioned, fixed, and prepared for H&E staining.

Conclusion

In this study, the effects of freezing/thawing on native and decellularized whole porcine kidneys in the absence of additional cryoprotectants were investigated. The adverse effects of freezing/thawing for native kidneys can lead to destructive effects that need to be mitigated or avoided using cryoprotectants in order to preserve the functionality of a whole organ. However, it was shown that the damaging effects of ice crystal formation are much less severe for decellularized ECM scaffolds, and freezing/thawing without any cryopreservation can be considered a promising option for long-term preservation of decellularized porcine kidneys. Our recommendation is that the porcine kidneys be frozen prior to decellularization to prevent spoilage by bacterial growth and after decellularization to prevent proteins from denaturing. The decellularized organs can then be preserved for months without cryoprotectants and thawed just prior to recellularization. Ideally, the repopulated kidneys would then be implanted without freezing again, but transportation and scheduling may require preservation of cell function by freezing. If needed, the use of current protocols using cryoprotectants to preserve cell function is recommended for freezing recellularized kidneys.

DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST

No potential conflicts of interest were disclosed.

Acknowledgments

We acknowledge the laboratory assistance of Ryan J. Morris, Jason R. Gassman, Tyson R. Jergensen, Robert M. Fuller, Benjamin Buttars, Spencer Baker, Jonathan Thibaudeau, and Angela Nakalembe.

Funding

Funding for this project was provided by Brigham Young University.

DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST

No potential conflicts of interest were disclosed.

REFERENCES

  • 1. Prevention CFDCA 2014 National chronic kidney disease fact sheet. 2014; Available from: http://www.cdc.gov/diabetes/pubs/factsheets/kidney.htm. [Google Scholar]
  • 2. H.R.S.A The need is real: Data. Donate the Gift of Life 2012; Available from: http://www.organdonor.gov/about/data.html. [Google Scholar]
  • 3. Orlando G, Cocco PDi, D'Angelo M, Clemente K, Famulari A, Pisani F. Regenerative medicine applied to solid organ transplantation: where do we stand?. Transplant Proc 2010; 42: 1011-1013; http://dx.doi.org/ 10.1016/j.transproceed.2010.03.066. [DOI] [PubMed] [Google Scholar]
  • 4. Soto-Gutierrez A, Wertheim JA, Ott HC, Gilbert TW. Perspectives on whole-organ assembly: Moving toward transplantation on demand. J Clini Invest 2012; 122: 3817-3823; PMID:23114604; http://dx.doi.org/ 10.1172/JCI61974. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5. Badylak SF, Taylor D, Uygun K. Whole-organ tissue engineering: Decellularization and recellularization of three-dimensional matrix scaffolds. Annu Rev Biomed Eng 2011; 13: 27-53; PMID:21417722; http://dx.doi.org/ 10.1146/annurev-bioeng-071910-124743. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Orlando G, Soker S, Stratta RJ. Organ bioengineering and regeneration as the new Holy Grail for organ transplantation. Ann Surg 2013; 285: 221-32; PMID:23782908; http:// dx.doi.org/ 10.1097/SLA.0b013e31829c79cf. [DOI] [PubMed] [Google Scholar]
  • 7. Crapo PM, Gilbert TW, Badylak SF. An overview of tissue and whole organ decellularization processes. Biomaterials 2011; 32: 3233-3243; PMID:21296410; http://dx.doi.org/ 10.1016/j.biomaterials.2011.01.057. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8. Badylak SF, Freytes DO, Gilbert TW. Extracellular matrix as a biological scaffold material: Structure and function. Acta Biomater 2009; 5: 1-13; PMID:18938117; http://dx.doi.org/ 10.1016/j.actbio.2008.09.013. [DOI] [PubMed] [Google Scholar]
  • 9. Gilbert TW, Sellaro TL, Badylak SF. Decellularization of tissues and organs. Biomaterials 2006; 27: 3675-3683; PMID:16519932; http://dx.doi.org/ 10.1016/j.biomaterials.2006.02.014. [DOI] [PubMed] [Google Scholar]
  • 10. Badylak SF. Xenogeneic extracellular matrix as a scaffold for tissue reconstruction. Transpl Immunol 2004; 12: 367-377; PMID:15157928; http://dx.doi.org/ 10.1016/S0966-3274(03)00117-5. [DOI] [PubMed] [Google Scholar]
  • 11. Mirmalek-Sani S-H, Sullivan DC, Zimmerman C, Shupe TD, Petersen BE. Immunogenicity of decellularized porcine liver for bioengineered hepatic tissue. Am J Pathol 2013; 183: 558-565; PMID:23747949; http://dx.doi.org/ 10.1016/j.ajpath.2013.05.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12. Caralt M, Uzarski JS, Iacob S, Obergfell KP, Berg N, Bijonowski BM, Kiefer KM, Ward HH, Wandinger-Ness A, Miller WM. Optimization and critical evaluation of decellularization strategies to develop renal extracellular matrix scaffolds as biological templates for organ engineering and transplantation. Am J Transplant 2015; 15(1):64-75; PMID:25403742; http://dx.doi.org/ 10.1111/ajt.12999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13. Badylak SF, Gilbert TW. Immune response to biologic scaffold materials. Semin Immunol 2008; 20: 109-116; PMID:18083531; http://dx.doi.org/ 10.1016/j.smim.2007.11.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14. Salvatori M, Peloso A, Katan R, Soker Sh, Lerut JP, Stratta RJ, Orlando G. Semi-xenotransplantation: the regenerative medicine-based approach to immunosuppression-free transplantation and to meet the organ demand. Xenotransplantation 2014; 17: 243-256; http://dx.doi.org/ 10.1016/10.1111/xen.12122. [DOI] [PubMed] [Google Scholar]
  • 15. Karlsson JOM, Toner M. Long-term storage of tissues by cryopreservation: Critical issues. Biomaterials 1996; 17: 243-256; PMID:8745321; http://dx.doi.org/ 10.1016/0142-9612(96)85562-1. [DOI] [PubMed] [Google Scholar]
  • 16. Adem CG, Harness JB. Computer control of a modified langendorff perfusion apparatus for organ preservation using cryoprotective agents. J Biomed Eng 1981; 3: 134-139; PMID:7015013; http://dx.doi.org/ 10.1016/0141-5425(81)90006-6. [DOI] [PubMed] [Google Scholar]
  • 17. Rabin Y, Taylor MJ, Wolmark N. Thermal expansion measurements of frozen biological tissues at cryogenic temperatures. J Biomech Eng 1998; 120: 259-266; PMID:10412388; http://dx.doi.org/ 10.1115/1.2798310. [DOI] [PubMed] [Google Scholar]
  • 18. Fahy GM. Biological effects of vitrification and devitrification, in The biophysics of organ cryopreservation. Springer 1987; 147: 265-297; http://dx.doi.org/ 10.1007/978-1-4684-5469-7_13. [DOI] [Google Scholar]
  • 19. Fahy GM, Wowk B, Pagotan R, Chang A, Phan J, Thomson B, Phan L. Physical and biological aspects of renal vitrification. Organogenesis 2009; 5: 167; PMID:20046680; http://dx.doi.org/ 10.4161/org.5.3.9974. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20. De Graaf IaM, Draaisma AL, Schoeman O, Fahy GM, Groothuis GMM, Koster HJ. Cryopreservation of rat precision-cut liver and kidney slices by rapid freezing and vitrification. Cryobiology 2007; 54: 1-12; PMID:17166492; http://dx.doi.org/ 10.1016/j.cryobiol.2006.09.002. [DOI] [PubMed] [Google Scholar]
  • 21. Woo SLY, Orlando CA, Camp JF, Akeson WH. Effects of postmortem storage by freezing on ligament tensile behavior. J Biomech 1986; 19: 399-404; PMID:3733765; http://dx.doi.org/ 10.1016/0021-9290(86)90016-3. [DOI] [PubMed] [Google Scholar]
  • 22. Chan E, Menovsky T, Welch AJ. Effects of cryogenic grinding on soft-tissue optical properties. Appl Opt 1996; 35: 4526-4532; PMID:21102871; http://dx.doi.org/ 10.1364/AO.35.004526. [DOI] [PubMed] [Google Scholar]
  • 23. Nonaka PN, Campillo N, Uriarte JJ, Garreta E, Melo E, De Oliveira LV, Navajas D, Farré R. Effects of freezing/thawing on the mechanical properties of decellularized lungs. J Biomed Mater Res A 2014; 102: 413-419; PMID:23533110; http://dx.doi.org/ 10.1002/jbm.a.34708. [DOI] [PubMed] [Google Scholar]
  • 24. Umale S, Deck C, Bourdet N, Dhumane P, Soler L, Marescaux J, Willinger R. Experimental mechanical characterization of abdominal organs: Liver, kidney & spleen. J Mech Behav Biomed Mater 2013; 17: 22-33; PMID:23127642; http://dx.doi.org/ 10.1016/j.jmbbm.2012.07.010. [DOI] [PubMed] [Google Scholar]
  • 25. Ternifi R, Gennisson JL, Tanter M, Beillas P. Effects of storage temperature on the mechanical properties of porcine kidney estimated using shear wave elastography. J Mech Behav Biomed Mater 2013; 28: 86-93; PMID:23973616; http://dx.doi.org/ 10.1016/j.jmbbm.2013.07.012. [DOI] [PubMed] [Google Scholar]
  • 26. Orlando G, Farney AC, Iskandar SS, Mirmalek-Sani SH, Sullivan DC, Moran E, Aboushwareb T, Paolo D, Wood KJ, Stratta RJ, et al. Production and implantation of renal extracellular matrix scaffolds from porcine kidneys as a platform for renal bioengineering investigations. Ann Surg 2012; 256: 363-370; PMID:22691371; http://dx.doi.org/ 10.1097/SLA.0b013e31825a02ab. [DOI] [PubMed] [Google Scholar]
  • 27. Orlando G, Booth C, Wang Z, Totonelli G, Ross CL, Moran E, Salvatori M, Maghsoudlou P, Turmaine M, Delario G, et al. Discarded human kidneys as a source of ECM scaffold for kidney regeneration technologies. Biomaterials 2013; 34: 5915-5925; PMID:23680364; http://dx.doi.org/ 10.1016/j.biomaterials.2013.04.033. [DOI] [PubMed] [Google Scholar]
  • 28. Nasseri S, Bilston LE, Phan-Thien N. Viscoelastic properties of pig kidney in shear, experimental results and modelling. Rheologica acta 2002; 41: 180-192; http://dx.doi.org/ 10.1007/s003970200017. [DOI] [Google Scholar]
  • 29. Pek YS, Wan AC, Ying JY. The effect of matrix stiffness on mesenchymal stem cell differentiation in a 3d thixotropic gel. Biomaterials 2010; 31: 385-391; PMID:19811817; http://dx.doi.org/ 10.1016/j.biomaterials.2009.09.057. [DOI] [PubMed] [Google Scholar]
  • 30. Engler A, Sweeney H, Discher D, Schwarzbauer J. Extracellular matrix elasticity directs stem cell differentiation. J Musculoskelet Neuronal Interact 2007; 7: 335; PMID:18094500. [PubMed] [Google Scholar]
  • 31. Fahy GM, Wowk B, Wu J. Cryopreservation of complex systems: The missing link in the regenerative medicine supply chain. Rejuvenation Res 2006; 9: 279-291; PMID:16706656; http://dx.doi.org/ 10.1089/rej.2006.9.279. [DOI] [PubMed] [Google Scholar]
  • 32. Kheirabadi BS, Fahy GM. Permanent life support by kidneys perfused with a vitrifiable (7.5 molar) cryoprotectant solution. Transplantation 2000; 70: 51-57; PMID:10919575. [PubMed] [Google Scholar]
  • 33. Fahy GM, Wowk B, Wu J, Phan J, Rasch C, Chang A, Zendejas E. Cryopreservation of organs by vitrification: Perspectives and recent advances. Cryobiology 2004; 48: 157-178; PMID:15094092; http://dx.doi.org/ 10.1016/j.cryobiol.2004.02.002. [DOI] [PubMed] [Google Scholar]
  • 34. Elami A, Gavish Z, Korach A, Houminer E, Schneider A, Schwalb H, Arav A. Successful restoration of function of frozen and thawed isolated rat hearts. J Thorac Cardiovasc Surg 2008; 135: 666-672; PMID:18329491; http://dx.doi.org/ 10.1016/j.cryobiol.2004.02.002. [DOI] [PubMed] [Google Scholar]
  • 35. Pegg DE, Green CJ, Walter CA. Attempted canine renal cryopreservation using dimethyl sulphoxide helium perfusion and microwave thawing. Cryobiology 1978; 15: 618-626; PMID:369772; http://dx.doi.org/ 10.1016/0011-2240(78)90086-X. [DOI] [PubMed] [Google Scholar]
  • 36. O'Leary SA, Doyle BJ, Mcgloughlin TM. The impact of long-term freezing on the mechanical properties of porcine aortic tissue. J Mech Behav Biomed Mater 2014; 37: 165-173; PMID:24922621; http://dx.doi.org/ 10.1016/j.jmbbm.2014.04.015. [DOI] [PubMed] [Google Scholar]
  • 37. Chen L, Wu Y, Yu J, Jiao Z, Ao Y, Yu C, Wang J, Cui G. Effect of repeated freezing–thawing on the achilles tendon of rabbits. Knee Surg Sports Traumatol Arthrosc 2011; 19: 1028-1034; PMID:20938643; http://dx.doi.org/ 10.1007/s00167-010-1278-y. [DOI] [PubMed] [Google Scholar]
  • 38. Santago AC, Kemper AR, Mcnally C, Sparks JL, Duma SM. Freezing affects the mechanical properties of bovine liver- Biomed 2009. Biomed Sci Instrum 2008; 45: 24-29; PMID:19369734. [PubMed] [Google Scholar]
  • 39. Giannini S, Buda R, Di Caprio F, Agati P, Bigi A, De Pasquale V, Ruggeri A. Effects of freezing on the biomechanical and structural properties of human posterior tibial tendons. Int Orthop 2008; 32: 145-151; PMID:17216243; http://dx.doi.org/ 10.1007/s00264-006-0297-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40. Gabriel M, Wachal K, Dzieciuchowicz L, Pawlaczyk K, Krasiński Z, Oszkinis G. The influence of cryopreservation on changes in diameter and compliance of allografts in an animal experimental model. Eur J Vasc Endovasc Surg 2006; 32: 169-175; PMID:16564709; http://dx.doi.org/ 10.1016/j.ejvs.2006.01.025. [DOI] [PubMed] [Google Scholar]
  • 41. Chow M-J, Zhang Y. Changes in the mechanical and biochemical properties of aortic tissue due to cold storage. J Surg Res 2011; 171: 434-442; PMID:207019; http://dx.doi.org/ 10.1016/j.jss.2010.04.007. [DOI] [PubMed] [Google Scholar]
  • 42. Han B, Miller JD, Jung JK. Freezing-induced fluid-matrix interaction in poroelastic material. J Biomech Eng 2009; 131: 021002-1; PMID:19102561; http://dx.doi.org/ 10.1115/1.3005170. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43. Schenke-Layland K, Xie J, Heydarkhan-Hagvall S, Hamm-Alvarez SF, Stock UA, Brockbank KGM, Maclellan WR. Optimized preservation of extracellular matrix in cardiac tissues: Implications for long-term graft durability. Ann Thorac Surg 2007; 83: 1641-1650; PMID:17462373; http://dx.doi.org/ 10.1016/j.athoracsur.2006.12.005. [DOI] [PubMed] [Google Scholar]
  • 44. Soto-Gutierrez A, Zhang L, Medberry C, Fukumitsu K, Faulk D, Jiang HB, Reing J, Gramignoli R, Komori J, Ross M, et al. A whole-organ regenerative medicine approach for liver replacement. Tissue Eng Part C Methods 2011; 17: 677-686; PMID:21375407; http://dx.doi.org/ 10.1089/ten.tec.2010.0698. [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from Organogenesis are provided here courtesy of Taylor & Francis

RESOURCES