Preparation of Decellularized Kidney Scaffolds in Rats

Summary

This protocol introduces a method to develop a scaffold using decellularized rat kidneys. The protocol includes decellularization and recellularization processes to confirm bioavailability. Decellularization is performed using Triton X-100 and sodium dodecyl sulfate.

Abstract

Tissue engineering is a cutting-edge discipline in biomedicine. Cell culture techniques can be applied for regeneration of functional tissues and organs to replace diseased or damaged organs. Scaffolds are needed to facilitate the generation of three-dimensional organs or tissues using differentiated stem cells in vivo. In this report, we describe a novel method for developing vascularized scaffolds using decellularized rat kidneys. Eight-week-old Sprague-Dawley rats were used in this study, and heparin was injected into the heart to facilitate flow into the renal vessels, allowing heparin to perfuse into the renal vessels. The abdominal cavity was opened, and the left kidney was collected. The collected kidneys were perfused for 9 h using detergents, such as Triton X-100 and sodium dodecyl sulfate, to decellularize the tissue. Decellularized kidney scaffolds were then gently washed with 1% penicillin/streptomycin and heparin to remove cellular debris and chemical residues. Transplantation of stem cells with the decellularized vascular scaffolds is expected to facilitate the generation of new organs. Thus, the vascularized scaffolds may provide a foundation for tissue engineering of organ grafts in the future.

Introduction

Cell culture techniques are applied for regeneration of functional tissues and organs to replace diseased or damaged organs. Allogenic organ transplantation is currently the most common treatment for irreversible organ damage; however, this approach requires the use of immunosuppression to prevent rejection of the transplanted organ. Moreover, despite advances in transplant immunology, 20% of transplant recipients may experience acute rejection within 5 years, and within 10 years after transplantation, 40% of recipients may lose their transplanted graft or die¹.

Advances in tissue engineering technologies have yielded in a new paradigm for transplantation of new organs without immune rejection using differentiated stem cells. After stem cell differentiation, a scaffold, called a synthetic extracellular matrix, is needed to facilitate the generation of three-dimensional organs and enable the new tissue to thrive within the recipient. Scaffolds from decellularized native organs have advantages, including a more effective environment for establishment of cells and enhancement of stem cell proliferation, although these mechanisms have not been fully elucidated². In particular, the kidney is a suitable organ for scaffold generation because it has abundant circulation and a niche for stem cell establishment. Additionally, because of the complex structure of the kidney, it is difficult to artificially regenerate kidneys for organ transplantation.

In this report, we introduce a method of developing vascularized scaffolds using decellularized organs in a rat model to facilitate future animal studies for tissue engineering purposes.

Protocol

This study was approved by the administration of Pusan National University of Medicine and was conducted in accordance with ethical guidelines for the use and care of animals. (certificate no. 2017-119). Prior to any animal studies, institutional approval should be obtained.
NOTE: All surgical and anesthetic instruments/equipment and reagents recommended for successful surgical presentation and imaging of abdominal organs are detailed in Table 1.

1. Preparation procedures for harvesting of rat kidneys

1. In preparation for surgery, place 8-week-old Sprague-Dawley rats (weighing 200–250 g) on a warming pad. Place a rectal thermometer probe in the rectum to monitor core temperature.
2. Anesthetize the rat with a 5% mixture of isoflurane gas (induction: 5%, maintenance: 3%).
3. To start the operation, place the rat in a supine position after administration of anesthesia. Mount the four limbs of the rat on the operation table with tape.
4. Shave and clean the abdomen of the donor rat with germicidal soap. Apply 2% betadine for at least 1–2 min, and wipe with a 70% ethanol solution. Repeat this sequence three times.
5. Cover the operative field with a sterile fenestrated drape.
6. Make a vertical abdominal incision and expose the left kidney, ureter, abdominal aorta, and inferior vena cava.
7. Visualize and dissect the left kidney, ureter, abdominal aorta, and inferior vena cava just before cutting the pedicle.

2. Transcardial perfusion

1. Before surgery, prepare the perfusion solution.
   1. Make 50 mL of perfusion solution per rat.
   2. Mix 1x PBS with approximately 10 U/mL heparin (1 25 kU vial will make 2.5 L of PBS+Hep).
   3. Mix equal volumes of 8% paraformaldehyde with 1x PBS to make the 4% PFA/1xPBS solution.
      NOTE: 8% PFA made in water can be stored at 4 °C for up to 2 months. However, 4% PFA diluted in PBS is only stable for 1 week at 4 °C. Make the dilution fresh.
2. Extend the vertical abdominal incision cranially. Be sure to draw the scissors away from the organs when cutting to avoid damaging the internal organs.
3. Continue the incision through the rib cage, and then cut through the diaphragm by lifting the sternum.
4. Pin the loose flap of skin out of the way, and free the heart by tearing any connective tissue with the forceps.
   CAUTION: Do not use the scissors to free the heart and this could result in unwanted bleeding.
5. Open the phosphate-buffered saline (PBS) line and ensure that the line is flowing before placing the needle into the left ventricle. Hold the heart gently with blunt forceps, and use a hemostat to control the needle. The needle should be inserted no more than 1/4 inch.
   NOTE: Insertion greater than 1/4 inch may result in perforation to the other side of the tissue.
6. While supporting the heart with the needle and hemostat, locate the right atrium and snip through it with iridectomy scissors. Rest the hemostat on the rat’s body, and make sure the needle is still positioned inside the heart.
   NOTE: If the cut is sufficient, there should be blood in the body cavity as the pressure from the PBS flowing into the rat is relieved.
7. Carefully unpin the front feet and skin flap.
8. Continue perfusing PBS for 4 min or longer if there is still blood visible in the kidney and liver.

3. Kidney harvesting and decellularization

1. Harvest the left kidney with the abdominal aorta and inferior vena cava.
2. Ligate the ureter, thoracic aorta, superior vena cava, and branches of the abdominal aorta.
3. Keep the organ hydrated in Dulbecco’s PBS (DPBS) in a 10 cm Petri dish.
4. Cannulate the abdominal aorta and inferior vena cava with a 23 Gauge catheter. To remove residual blood, connect the cannula with a peristaltic pump, and wash with DPBS (500 mL) and 16 U/mL heparin for 90 min at a rate of 5 rpm at 37 °C.
5. To decellularize the kidney, perfuse the kidney with 1% Triton X-100 (1 L) for 3 h and then with 0.75% sodium dodecyl sulfate (SDS) solution (2 L) for 6 h at a constant pressure of 40 mmHg.
   NOTE: The kidney will become transparent after 8 h.
6. To remove residual SDS, perfuse the sample with 1% penicillin in distilled water (6 L) for 18 h (overnight) and then with sterile DPBS (500 mL) and 16 U/mL heparin for 90 min.

Representative Results

The gross morphology of rat kidneys was dark red (Figure 1A). After decellularization, the kidney became pale and translucent (Figure 1D). Residual genomic DNA was assessed with a commercial kit according to the manufacturer’s instructions, in decellularized kidney scaffolds and compared with that in native kidneys (control). Quantitative analysis confirmed that tissue genomic DNA was almost eliminated after decellularization. From 14 cases, the average DNA contents were 115.05 ng/µL for the control and 1.96 ng/µL for the decellularized scaffold. In total, 98.3% of DNA was removed (Figure 2), although the three-dimensional structure was maintained, and acellular gromeruli were preserved in the cortical parenchyma (Figure 3).

Figure 1: Rat kidneys subjected to renal arterial perfusion decellularization. (A) Immediately after the start of decellularization. (B) After Triton X-100 treatment. (C) After SDS buffer treatment. (D) After overnight scaffold washing. Please click here to view a larger version of this figure.

Figure 2: DNA concentrations in control and decellularized rat kidneys, showing reduced DNA contents after decellularization. Please click here to view a larger version of this figure.

Figure 3: Hematoxylin and eosin staining of control and decellularized kidney samples. (A) control cortex (A`) decellularized cortex (B) control medulla (B`) decellularized medulla (C) control vein (C`) decellularized vein. Scale bar, 100 µm. Please click here to view a larger version of this figure.

Discussion

Various protocols have been used for decellularization of organs and other tissues. The optimal decellularization protocol should preserve the three-dimensional architecture of the extracellular matrix (ECM). In general, such protocols consist of lysing the cell membrane by physical processing or ionic solutions, dissociating the cytoplasm and nucleus from the ECM by enzymatic processing or detergents, and then removing cellular debris from the tissue³. Physical processes include scraping, solution agitation, pressure gradients, snap-freezing, nonthermal permanent electroporation, and supercritical fluids². Cells on the external surface of a tissue or organ, such as the skin or small intestine, can be efficiently removed by mechanical processes combined with enzymes⁴. Ionic or nonionic detergents dissolve DNA/protein interactions, lipids, and lipoproteins, but can damage the ECM structure⁵. Enzymes remove the dissociated cytoplasm and nuclear material, but leave these materials in the ECM, which can cause an immune response⁶. The optimal agents for decellularization are determined by tissue thickness and density or the clinical use of the decellularized tissue.

For decellularization, we used a combination of nonionic and ionic detergents: Triton X-100 and SDS. Triton X-100, as a nonionic detergent, effectively disrupts lipid/lipid and lipid/protein interactions. However, Triton X-100 may also destroy the ECM ultrastructure owing to loss of glycosaminoglycan (GAG), laminin, and fibronectin contents. SDS, as an ionic detergent, effectively removes nuclear remnants and cytoplasmic proteins, but also disrupts the ECM ultrastructure by loss of GAG and collagen³. Although these agents destroy the microstructure of the ECM, SDS and Triton X-100 successfully remove all DNA contents^7,8. This is essential because remaining DNA content within a scaffold can cause immune rejection. In tissue that has been properly decellularized, the DNA content should be less than 50 ng/mg^9,10.

In the method, the pressure of the decellularization perfusion was 40 mmHg. Pressure control is required for decellularization perfusion. The optimal perfusion pressure varies from organ to organ, and 60 mmHg is the optimal pressure for human and porcine kidney or heart decellularization¹¹. In rats, 40 mmHg is considered sufficient for decellularization perfusion¹².

One promising treatment for replacing allograft transplantation is transplantation of stem cells using a vascularized scaffold. We hope that this protocol for organ decellularization may provide a foundation for future tissue engineering studies.

Materials

Name	Company	Catalog Number	Comments
1 cc syringe (inject probes and vehicle solutions	Becton Dickinson	305217
10-0 ethilon for vessel anastomosis	Ethicon	9032G
25 gauge inch guide needle(for vascular catheters)	Becton Dickinson	305145
3-0 PDS incision closure rat	Ethicon	Z316H
3-0 Prolene incision closure rat	Ethicon	8832H
3-0 silk spool vascular access/ligation in rat	Braintree Scientific	SUT-S 110
4-0 PDS incision closure mouse	Ethicon	Z773D
4-0 Prolene incision closure mouse	Ethicon	8831H
5-0 silk spool vascular access/ligation in mouse	Braintree Scientific	SUT-S 106
Fine Scissors to cut fascia/connective tissue	Fine Science Tools	14058-09
Halsey needle holder	Fine Science Tools	12001-13
Kelly Hemostat for rats: muscle clamp to minimize bleeding when cut	Fine Science Tools	13018-14
Polyethelyne 50 tubing, catheter tubing 100 ft	Braintree Scientific	.023" × .038”
Schwartz microserrefine vascular clamps	Fine Science Tools	18052-01 (straight)
		18052-03 (curved)
Surgical Scissors to cut skin	Fine Science Tools	14002-12
Vannas-Tubingen Spring scissors for arteriotomy	Fine Science Tools	15003-08