Microdissection of Primary Renal Tissue Segments and Incorporation with Novel Scaffold-free Construct Technology

Summary

Tissue-engineered renal constructs provide a solution for the organ shortage and deleterious effects of dialysis. Here, we describe a protocol to micro dissect murine kidneys for isolation of cortico-medullary segments. These segments are implanted into scaffold-free cellular constructs, forming renal organoids.

Abstract

Kidney transplantation is now a mainstream therapy for end-stage renal disease. However, with approximately 96,000 people on the waiting list and only one-fourth of these patients achieving transplantation, there is a dire need for alternatives for those with failing organs. In order to decrease the harmful consequences of dialysis along with the overall healthcare costs it incurs, active investigation is ongoing in search of alternative solutions to organ transplantation. Implantable tissue-engineered renal cellular constructs are one such feasible approach to replacing lost renal functionality. Here, described for the first time, is the microdissection of murine kidneys for isolation of living corticomedullary renal segments. These segments are capable of rapid incorporation within scaffold-free endothelial-fibroblast constructs which may enable rapid connection with host vasculature once implanted. Adult mouse kidneys were procured from living donors, followed by stereoscope microdissection to obtain renal segments 200 - 300 µm in diameter. Multiple renal constructs were fabricated using primary renal segments harvested from only one kidney. This method demonstrates a procedure which could salvage functional renal tissue from organs that would otherwise be discarded.

Introduction

Chronic kidney disease (CKD) is one of the current major public health challenges worldwide¹. The prevalence of CKD in the United States is over 14% of the total population, with over 600,000 Americans suffering from the most severe form, end-stage renal disease (ESRD)². The current treatment options available for those with ESRD include dialysis and kidney transplantation. Although approximately 25,000 patients undergo renal transplantation each year, a significant number of patients are added annually leading to a large disparity between those awaiting a life-saving organ and those receiving transplantation³. In addition to its serious negative effects on longevity and quality of life, dialysis is associated with an astonishing financial burden. In 2014, Medicare paid claims totaled over $30 billion for ESRD patients². With a limited organ supply and no apparent downtrend in patients requiring dialysis, research efforts aimed at identifying alternative solutions to dialysis and transplantation are ever important. Even a relatively short delay in the need for dialysis increases a patient's number of quality-adjusted life years and productivity substantially while postponing dialysis-related costs^4,5,6.

Solutions for functional tissue loss, like that in ESRD, are currently being studied in tissue engineering and regenerative medicine laboratories, with widely varied approaches ranging from scaffold-based organoid fabrication to whole organ engineering using decellularized organ structures for cellular implantation^7,8,9,10,11. Recapitulating complex renal structures from marginal or discarded kidneys has only partially been investigated. In fact, nearly 20% of kidneys procured for transplantation are discarded for various reasons^12,13. The functional renal tissue from these putative grafts could be utilized and incorporated into one or many tissue-engineered constructs. Prior studies have demonstrated the feasibility of working with these discarded organs, utilizing kidneys for the extra-cellular matrix for tissue engineering purposes^14,15. However, few have used primary nephronal tissue from healthy kidneys for tissue-engineering purposes^16,17,18.

One method previously described by Kim et al. involves isolation of renal "segments" from healthy rat kidneys, which were then seeded on polyglycolic acid (PGA) scaffolds for construct fabrication¹⁶. However, little information is given regarding exact dissection methodology, and segments were obtained from a combination of fine mincing and filtration. We describe a modification of this protocol, which similarly produces discrete renal segments with intact nephronal architecture, but instead relies on microdissection techniques. Nephrectomies are performed on living adult mice, after which the kidneys are transferred to the dissection microscope where the renal capsule is removed, and the tissue is further dissected. Small-bore 30½ G needles are used as cutting instruments and also as guides aiding in dissection, as the needle diameter is equal to the target diameter of the renal segments. The isolated, in this case murine, renal segments maintain viability in culture and incorporate with scaffold-free endothelial-fibroblast cellular constructs¹⁹. These constructs have previously been used to engineer other organs, including a bio-artificial pancreas²⁰.

Protocol

All animal surgical procedures described below were approved by the Institutional Animal Care and Use Committee (IACUC) at the Medical University of South Carolina prior to any animal surgeries or use of any animal tissues.

1. Murine Nephrectomy

1. Don a surgical mask and bouffant cap to minimize the risk of contamination. Maintain sterility during set-up of the surgical area.
   1. Place non-fenestrated surgical drapes on the operating table.
   2. Open the pack of autoclaved instruments onto the sterile drapes. The instruments required for nephrectomy include 3 small hemostats, fine forceps with teeth, fine forceps without teeth, and straight iris scissors.
   3. Place another non-fenestrated surgical drape in the center of the operating table.
   4. Prepare 5 mL of Dulbecco's Phosphate Buffered Saline (DPBS) with calcium and magnesium, supplemented with 1% penicillin/streptomycin in a 15-mL sterile conical tube. Place this solution on ice next to the operating table.
2. Obtain an 8 - 12 week old C57BL/6 mouse (male or female) from the animal housing complex.
3. Place the mouse inside an anesthesia induction chamber and induce with 3.5% inhaled isoflurane. Place a nose cone on the mouse, using 2% isoflurane for maintenance anesthesia.
   1. Monitor for adequate depth of anesthesia periodically throughout the surgical procedure by assessing for pedal reflex (firm toe pinch).
4. Remove all of the hair from the ventral torso using hair removal cream. Rinse this area with distilled water to ensure all hair has been removed from the abdomen.
5. Transfer the mouse to the sterile operating table in supine position under the top non-fenestrated drape. Cut out a 2 x 2 cm² fenestration in the drape just overlying the mouse abdomen.
6. Apply povidone-iodine followed by 70% ethanol to the abdomen using sterile gauze or cotton balls.
7. Using fine forceps with teeth and iris scissors, make a 3 - 4 cm vertical abdominal skin incision parallel to the midline, 0.5 cm to the left of midline
   Note: If attempting to remove the right kidney, this will be an incision 0.5 cm to the right of midline.
   1. Grasp the peritoneum with fine forceps without teeth and incise with iris scissors to enter the abdominal cavity. Apply hemostats to the superior and inferior lateral edges of skin and peritoneum to place tension laterally and enhance exposure.
   2. Shift the intestinal contents delicately to the right side of the abdomen to expose the left kidney. Gently place traction on the kidney to elevate it out of the abdomen.
   3. Place a hemostat across the hilum of the left kidney. Use iris scissors to transect the ureter and renal vessels.
   4. Place the left kidney in the 1% Penicillin/Streptomycin DPBS Solution on ice.
   5. Transfer the tube containing the kidney to the sterile tissue culture hood. Transfer the kidney from the 15-mL conical tube to a 60-mm petri dish (Figure 1). Rinse twice using 5 mL DPBS with calcium and magnesium. Keep the kidney suspended in 5 mL DPBS in the 60-mm dish.
      1. Prepare an additional 60 mm petri dish with 5 mL DPBS to be used during the microdissection protocol.
   6. Euthanize the mouse by thoracotomy and exsanguination following the procurement of the kidney, according to an approved IACUC animal protocol.

2. Murine Kidney Microdissection for Renal Segment Isolation

1. Continue using sterile technique for this portion of the protocol, including sterile gloves, surgical mask, and bouffant to minimize contamination risk.
   1. Place sterile drapes on the stereo microscope platform as well as to the areas just left and right of the microscope to have a sterile field for instruments. Place plastic adhesive sheets on microscope focus knobs and spray with 70% ethanol. Turn on the dual gooseneck illuminator to light the stage.
   2. Open sterilized instruments (fine forceps without teeth, scalpel handle, #15 scalpel blade, two straight hemostats, two 30½ gauge hollow-bore needles) onto sterile drapes. Place the #15 blade onto the scalpel handle now and attach one hemostat to each needle adaptor/hub to create needle dissection instruments for later use, as shown in Figure 1.
2. Move the 60-mm petri dishes, one containing the kidney in DPBS and the other only DPBS, from the tissue culture hood to the microscope area.
   1. Place the 60-mm petri dish under the objective and remove the lid to expose the kidney. Leave the DPBS-containing dish to the side on the sterile field for later use.
   2. Move the dish so that only the anterior or posterior surface of the kidney is in view (i.e., the large flat face of the kidney, with the other face touching the bottom of the dish). Zoom to 3.2 - 4X for this portion of the procedure. Using the non-dominant hand, use fine forceps without teeth to pierce and pin the kidney against the dish near the inferior and superior poles of the kidney.
   3. Keep the non-dominant hand in place to pin the kidney. With the dominant hand, use the #15 blade to remove the translucent fibrous capsular layer from the exposed surface. Shave the most superficial 0.5 mm from the exposed surface of the kidney to remove the remainder of the capsule.
   4. Use the #15 blade to dissect an inverted pyramid of tissue from the decapsulated area, approximately 2 mm³ in size. Retrieve the 60-mm dish containing only DPBS and place the pyramid of tissue into the clean dish. Discard the remainder of the kidney.
   5. Decrease magnification to 1.5 - 2.0X for the rest of the dissection. Using two hemostat-needle instruments as cutting instruments, slice the tissue fragment into progressively smaller pieces until the tissue segments are less than or equal to the diameter of the needle tips. A renal tissue 2 mm³ in size will produce more than 50 segments once completely dissected.
   6. Move the 60-mm dish with renal segments to the tissue culture hood. Remove DPBS with 1000 µL pipette. Add 5 mL DMEM, supplemented with 10% Fetal Bovine Serum (FBS) and 1% penicillin/streptomycin.
   7. Culture for 3 days at 37 °C in an incubator or implant into cellular constructs immediately.

3. Renal Segment Cellular Construct Fabrication

1. Culture normal human dermal fibroblasts (NHDF) and human adipose microvascular endothelial cells (HAMEC) for several weeks to obtain 7.2 x 10⁵ fibroblasts and 1.8 x 10⁵ endothelial cells per construct desired. In the tissue culture hood, seed the appropriate ratio of fibroblasts to endothelial cells (4:1) into rod-shaped wells in agarose molds to form scaffold-free pre-vascular endothelial-fibroblast constructs (SPEC) as previously described by Czajka et al¹⁹.
2. Obtain a sterilized hemostat, 30½ G needle, and a sterile spatula with flat end.
   1. Remove the majority of the media from the 60-mm plate containing renal segments, being careful not to aspirate and remove the renal segments.
   2. Equip surgical Loupes to visualize implantation of the segments.
   3. Scrape the end of the spatula gently along the bottom of the 60-mm dish to obtain 10 - 15 segments, spread out evenly along the very tip of the spatula. Place the tip of the spatula as close as possible to the lip of the well where the SPEC was seeded.
   4. Use a hemostat-30½ G needle instrument in the other hand to move each individual segment from the spatula tip to the edge of the well. Gently move each segment down into the well using the needle tip until slightly submerged in the previously seeded cellular suspension.
   5. Culture renal segment-SPEC in 2:1:1 ratio of FGM-2/EGM-2/DMEM medium (2.5 mL volume total). Change culture media every 24 h for 3 days. Remove the constructs from the molds at 72 h and fix or flash-freeze for processing.

Results

The protocol described produces approximately 50 renal segments per pyramidal 2 mm³ section of renal tissue. The renal segments that have been processed and imaged have tubular and glomerular components in differing proportions (see Figure 2). The intact segments were subjected to an assay in order to determine the viability of different segments once every 24 h for three days. Green-fluorescent calcein-AM is present with intracellular esterase act...

Discussion

Methods used to engineer living renal tissue constructs vary widely with regard to both the type of cells and biomaterials utilized, and in many cases, are outdated or not well-characterized in the literature⁷. While many are using stem cell approaches or recapitulating individual components of the renal architecture in isolation, the prospect of artificially recreating an entire organ with over 26 different differentiated cell types from cellular suspensions is overwhelming to consider

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Non-fenestrated Sterile Field	Busse Hospital Disposables	696
Fenestrated Sterile Field	Busse Hospital Disposables	697
Halsted Mosquito Forceps 5 Curved	Miltex	Mil-7-4	"Hemostat" in manuscript
Extra Fine Graefe Forceps, Curved with teeth	Fine Science Tools	11155-10	Fine forceps with teeth
Extra Fine Graefe Forceps, Serrated (without teeth)	Fine Science Tools	11152-10	Fine forceps without teeth
Fine Scissors - Tungsten Carbide	Fine Science Tools	14568-09	Iris Scissors
Betadine Surgical Scrub with Pump, Povidone-iodine 7.5%	Purdue Products L.P.	67618-151-17
Sterile Cotton Gauze Pad (4" x 4")	Fisher Healthcare	22-415-469
Dulbecco's Phosphate Buffered Solution	Corning	21-030-CV
Penicillin/Streptomycin Solution, 100X	Corning	30-002-Cl
Isoflurane, USP	Manufacturer: Piramal, Distributor: McKesson	2254845
Nair Hair Remover	Nair	22600-23307	Hair Removal Cream in text
200 Proof Ethanol	Decon Laboratories	2705	Diluted to 70% Ethanol Solution
BioLite 60mm Tissue Culture Dish	Themo-Scientific	130181
Press'n Seal	Glad	12587-70441	Applied to Stereoscope
SZX16 Stereo Microscope	Olympus	SZX16
Fiber Optic Illuminator	Cole Parmer	41720-20
Self-Supporting Dual-Light Pipe, 23" L Gooseneck	Cole Parmer	EW-41720-60
Scalpel Handle #3	Miltex	Mil-4-7
Sterile Rib-Back Carbon Steel Blade, Blade Size 15	Bard-Parker	371115
31 1/2 Gauge Needle	ThermoFisher Scientific	14-826F	Becton Dickinson 305106
Dulbecco's Modified Eagle's Medium	Corning	10-017-CV
Fetal Select 100% Bovine Serum	Atlas Biologicals	FS-0500-AD
Normal Human Dermal Fibroblasts	Lonza	CC-2511
Human Adipose Microvascular Endothelial Cells	Sciencell Research Laboratories	7200
Surgical Loupes (2.5x)	Orascoptic	(N/A) Custom Order
FGM-2 (Fibroblast Basal Medium with FGM-2 SingleQuots Added)	Lonza	CC-3131, CC-4126
EGM-2 (Endothelial Basal Medium with EGM-2 SingleQuots Added)	Lonza	CC-3156, CC-4176
Live/Dead Viability/Cytotoxicity Kit for Mammalian Cells	ThermoFisher Scientific	L3224
Anti-Cytokeratin-18 Antibody	Abcam	ab668
Goat anti-Mouse IgG, Alexa Fluor 633	ThermoFisher Scientific	A-21052
Goat anti-Rabbit IgG, Alexa Fluor 546	ThermoFisher Scientific	A-11010
Anti-Von Willebrand Factor Antibody	Abcam	ab6994
Albumin, Fluorescein isothiocyanate Conjugate	Sigma Aldrich	A9771-50MG
Hoescht 33342	BD Pharmingen	561908
Background Buster	Innovex Biosciences	NB306