NIH Institutional Postdoctoral Training Grant, NIH-HL-007260

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
<ol>
	<li>Don a surgical mask and bouffant cap to minimize the risk of contamination. Maintain sterility during set-up of the&#160;surgical area.
	<ol>
		<li>Place non-fenestrated surgical drapes on the operating table.</li>
		<li>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.</li>
		<li>Place another non-fenestrated surgical drape in the center of the operating table.</li>
		<li>Prepare 5 mL of Dulbecco&#39;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.</li>
	</ol>
	</li>
	<li>Obtain an 8 - 12 week old C57BL/6 mouse (male or female) from the animal housing complex.</li>
	<li>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.
	<ol>
		<li>Monitor for adequate depth of anesthesia periodically throughout the surgical procedure by assessing for pedal reflex (firm toe pinch).</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>Apply povidone-iodine followed by 70% ethanol to the abdomen using sterile gauze or cotton balls.</li>
	<li>Using fine forceps with teeth and iris scissors, make a 3 - 4&#160;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.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>Place a hemostat across the hilum of the left kidney. Use iris scissors to transect the ureter and renal vessels.</li>
		<li>Place the left kidney in the 1% Penicillin/Streptomycin DPBS Solution on ice.</li>
		<li>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.
		<ol>
			<li>Prepare an additional 60 mm petri dish with 5 mL DPBS to be used during the microdissection protocol.</li>
		</ol>
		</li>
		<li>Euthanize the mouse by thoracotomy and exsanguination following the procurement of the kidney, according to an approved IACUC animal protocol.</li>
	</ol>
	</li>
</ol>
2. Murine Kidney Microdissection for Renal Segment Isolation
<ol>
	<li>Continue using sterile technique for this portion of the protocol, including sterile gloves, surgical mask, and bouffant to minimize contamination risk.
	<ol>
		<li>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.</li>
		<li>Open sterilized instruments (fine forceps without teeth, scalpel handle, #15 scalpel blade, two straight hemostats, two 30&#189; 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.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Move the 60-mm dish with renal segments to the tissue culture hood. Remove DPBS with 1000 &#181;L pipette. Add 5 mL DMEM, supplemented with 10% Fetal Bovine Serum (FBS) and 1% penicillin/streptomycin.</li>
		<li>Culture for 3 days at 37 &#176;C in an incubator or implant into cellular constructs immediately.</li>
	</ol>
	</li>
</ol>
3. Renal Segment Cellular Construct Fabrication
<ol>
	<li>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.</li>
	<li>Obtain a sterilized hemostat, 30&#189; G needle, and a sterile spatula with flat end.
	<ol>
		<li>Remove the majority of the media from the 60-mm plate containing renal segments, being careful not to aspirate and remove the renal segments.</li>
		<li>Equip surgical Loupes to visualize implantation of the segments.</li>
		<li>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.</li>
		<li>Use a hemostat-30&#189; 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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>

<ol>
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U.S.R.D Available from: <a target="_blank" href="https://www.usrds.org/2016/download/v2_ESRD_16.pdf">https://www.usrds.org/2016/download/v2_ESRD_16.pdf</a> (2016)</li><li>Hart, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=OPTN/SRTR+2015+Annual+Data+Report:+Kidney.">OPTN/SRTR 2015 Annual Data Report: Kidney.</a> American journal of transplantation : official journal of the American Society of Transplantation and the American Society of Transplant Surgeons. 17, 21-116 (2017).</li><li>de Vries, E. F., Rabelink, T. J., van den Hout, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modelling+the+Cost-Effectiveness+of+Delaying+End-Stage+Renal+Disease.">Modelling the Cost-Effectiveness of Delaying End-Stage Renal Disease.</a> Nephron. 133 (2), 89-97 (2016).</li><li>Lefebvre, P., Duh, M. 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S., Russo, S., Marcellusi, A., Quintaliani, G., Fouque, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Economic+effects+of+treatment+of+chronic+kidney+disease+with+low-protein+diet.">Economic effects of treatment of chronic kidney disease with low-protein diet.</a> Journal of renal nutrition : the official journal of the Council on Renal Nutrition of the National Kidney Foundation. 24 (5), 313-321 (2014).</li><li>Moon, K. H., Ko, I. K., Yoo, J. J., Atala, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kidney+diseases+and+tissue+engineering.">Kidney diseases and tissue engineering.</a> Methods. 99, 112-119 (2016).</li><li>Wobma, H., Vunjak-Novakovic, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+Engineering+and+Regenerative+Medicine+2015:+A+Year+in+Review.">Tissue Engineering and Regenerative Medicine 2015: A Year in Review.</a> Tissue engineering. 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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diagnosing+the+Decades-Long+Rise+in+the+Deceased+Donor+Kidney+Discard+Rate+in+the+United+States.">Diagnosing the Decades-Long Rise in the Deceased Donor Kidney Discard Rate in the United States.</a> Transplantation. 101 (3), 575-587 (2017).</li><li>Mohan, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+weekend+effect+alters+the+procurement+and+discard+rates+of+deceased+donor+kidneys+in+the+United+States.">The weekend effect alters the procurement and discard rates of deceased donor kidneys in the United States.</a> Kidney international. 90 (1), 157-163 (2016).</li><li>Orlando, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Discarded+human+kidneys+as+a+source+of+ECM+scaffold+for+kidney+regeneration+technologies.">Discarded human kidneys as a source of ECM scaffold for kidney regeneration technologies.</a> Biomaterials. 34 (24), 5915-5925 (2013).</li><li>Katari, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+bioengineering+with+scaffolds+generated+from+human+kidneys.">Renal bioengineering with scaffolds generated from human kidneys.</a> Nephron. Experimental nephrology. 126 (2), 119 (2014).</li><li>Kim, S. S., Park, H. J., Han, J., Choi, C. Y., Kim, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Renal+tissue+reconstitution+by+the+implantation+of+renal+segments+on+biodegradable+polymer+scaffolds.">Renal tissue reconstitution by the implantation of renal segments on biodegradable polymer scaffolds.</a> Biotechnology letters. 25 (18), 1505-1508 (2003).</li><li>Guimaraes-Souza, N. K., Yamaleyeva, L. M., AbouShwareb, T., Atala, A., Yoo, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+reconstitution+of+human+kidney+structures+for+renal+cell+therapy.">In vitro reconstitution of human kidney structures for renal cell therapy.</a> Nephrology, dialysis, transplantation : official publication of the European Dialysis and Transplant Association - European Renal Association. 27 (8), 3082-3090 (2012).</li><li>Kelley, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tubular+cell-enriched+subpopulation+of+primary+renal+cells+improves+survival+and+augments+kidney+function+in+rodent+model+of+chronic+kidney+disease.">Tubular cell-enriched subpopulation of primary renal cells improves survival and augments kidney function in rodent model of chronic kidney disease.</a> American journal of physiology. Renal physiology. 299 (5), 1026-1039 (2010).</li><li>Czajka, C. A., Drake, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Self-assembly+of+prevascular+tissues+from+endothelial+and+fibroblast+cells+under+scaffold-free,+nonadherent+conditions.">Self-assembly of prevascular tissues from endothelial and fibroblast cells under scaffold-free, nonadherent conditions.</a> Tissue engineering. Part A. 21 (1-2), 277-287 (2015).</li><li>Rhett, J. M., Wang, H., Bainbridge, H., Song, L., Yost, M. 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The authors have nothing to disclose.

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<sup class=...

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&#39;s number of quality-adjusted life years and productivity substantially while postponing dialysis-related costs4,5,6.
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 implantation7,8,9,10,11. 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 reasons12,13. 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 purposes14,15. However, few have used primary nephronal tissue from healthy kidneys for tissue-engineering purposes16,17,18.
One method previously described by Kim et al. involves isolation of renal &#34;segments&#34; 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 30&#189; 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.

<table><tbody><tr><td>Non-fenestrated Sterile Field</td><td>Busse Hospital Disposables</td><td>696</td><td></td></tr><tr><td>Fenestrated Sterile Field</td><td>Busse Hospital Disposables</td><td>697</td><td></td></tr><tr><td>Halsted Mosquito Forceps 5 Curved</td><td>Miltex</td><td>Mil-7-4</td><td>&quot;Hemostat&quot; in manuscript</td></tr><tr><td>Extra Fine Graefe Forceps, Curved with teeth</td><td>Fine Science Tools</td><td>11155-10</td><td>Fine forceps with teeth</td></tr><tr><td>Extra Fine Graefe Forceps, Serrated (without teeth)</td><td>Fine Science Tools</td><td>11152-10</td><td>Fine forceps without teeth</td></tr><tr><td>Fine Scissors - Tungsten Carbide</td><td>Fine Science Tools</td><td>14568-09</td><td>Iris Scissors</td></tr><tr><td>Betadine Surgical Scrub with Pump, Povidone-iodine 7.5%</td><td>Purdue Products L.P.</td><td>67618-151-17</td><td></td></tr><tr><td>Sterile Cotton Gauze Pad (4&quot; x 4&quot;)</td><td>Fisher Healthcare</td><td>22-415-469</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Solution</td><td>Corning</td><td>21-030-CV</td><td></td></tr><tr><td>Penicillin/Streptomycin Solution, 100X</td><td>Corning</td><td>30-002-Cl</td><td></td></tr><tr><td>Isoflurane, USP</td><td>Manufacturer: Piramal, Distributor: McKesson</td><td>2254845</td><td></td></tr><tr><td>Nair Hair Remover</td><td>Nair</td><td>22600-23307</td><td>Hair Removal Cream in text</td></tr><tr><td>200 Proof Ethanol</td><td>Decon Laboratories</td><td>2705</td><td>Diluted to 70% Ethanol Solution</td></tr><tr><td>BioLite 60mm Tissue Culture Dish</td><td>Themo-Scientific</td><td>130181</td><td></td></tr><tr><td>Press&#039;n Seal</td><td>Glad</td><td>12587-70441</td><td>Applied to Stereoscope</td></tr><tr><td>SZX16 Stereo Microscope</td><td>Olympus</td><td>SZX16</td><td></td></tr><tr><td>Fiber Optic Illuminator</td><td>Cole Parmer</td><td>41720-20</td><td></td></tr><tr><td>Self-Supporting Dual-Light Pipe, 23&quot; L Gooseneck</td><td>Cole Parmer</td><td>EW-41720-60</td><td></td></tr><tr><td>Scalpel Handle #3</td><td>Miltex</td><td>Mil-4-7</td><td></td></tr><tr><td>Sterile Rib-Back Carbon Steel Blade, Blade Size 15</td><td>Bard-Parker</td><td>371115</td><td></td></tr><tr><td>31 1/2 Gauge Needle</td><td>ThermoFisher Scientific</td><td>14-826F</td><td>Becton Dickinson 305106</td></tr><tr><td>Dulbecco&#039;s Modified Eagle&#039;s Medium</td><td>Corning</td><td>10-017-CV</td><td></td></tr><tr><td>Fetal Select 100% Bovine Serum</td><td>Atlas Biologicals</td><td>FS-0500-AD</td><td></td></tr><tr><td>Normal Human Dermal Fibroblasts</td><td>Lonza</td><td>CC-2511</td><td></td></tr><tr><td>Human Adipose Microvascular Endothelial Cells</td><td>Sciencell Research Laboratories</td><td>7200</td><td></td></tr><tr><td>Surgical Loupes (2.5x)</td><td>Orascoptic</td><td>(N/A) Custom Order</td><td></td></tr><tr><td>FGM-2 (Fibroblast Basal Medium with FGM-2 SingleQuots Added)</td><td>Lonza</td><td>CC-3131, CC-4126</td><td></td></tr><tr><td>EGM-2 (Endothelial Basal Medium with EGM-2 SingleQuots Added)</td><td>Lonza</td><td>CC-3156, CC-4176</td><td></td></tr><tr><td>Live/Dead Viability/Cytotoxicity Kit for Mammalian Cells</td><td>ThermoFisher Scientific</td><td>L3224</td><td></td></tr><tr><td>Anti-Cytokeratin-18 Antibody</td><td>Abcam</td><td>ab668</td><td></td></tr><tr><td>Goat anti-Mouse IgG, Alexa Fluor 633</td><td>ThermoFisher Scientific</td><td>A-21052</td><td></td></tr><tr><td>Goat anti-Rabbit IgG, Alexa Fluor 546</td><td>ThermoFisher Scientific</td><td>A-11010</td><td></td></tr><tr><td>Anti-Von Willebrand Factor Antibody</td><td>Abcam</td><td>ab6994</td><td></td></tr><tr><td>Albumin, Fluorescein isothiocyanate Conjugate</td><td>Sigma Aldrich</td><td>A9771-50MG</td><td></td></tr><tr><td>Hoescht 33342</td><td>BD Pharmingen</td><td>561908</td><td></td></tr><tr><td>Background Buster</td><td>Innovex Biosciences</td><td>NB306</td><td></td></tr></tbody></table>

microdissection of primary renal tissue segments and incorporation with novel scaffold-free construct technology

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 &#181;m 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.

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...

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

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.

Kidney transplantation is now a mainstream therapy for end-stage renal disease. However, with approximately 96,000 people on the waiting list and only ...

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Universidad de Cartagena UDEC

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JoVE Journal

Bioengineering

7.3K Views.  Medical University of South Carolina. 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.

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

Minced Tissue in Compressed Collagen: A Cell-containing Biotransplant for Single-staged Reconstructive Repair

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped human cell culture facilities. These methods include isolation of cells in single cell suspension and several laborious and time-consuming events before transplantation back to the patient. Previous studies suggest that the body itself could be used as a bioreactor for cell expansion and regeneration of tissue in order to minimize ex vivo manipulations of tissues and cells before transplanting to the patient. The aim of this study was to demonstrate a method for tissue harvesting, isolation of continuous epithelium, mincing of the epithelium into small pieces and incorporating them into a three-layered biomaterial. The three-layered biomaterial then served as a delivery vehicle, to allow surgical handling, exchange of nutrition across the transplant, and a controlled degradation. The biomaterial consisted of two outer layers of collagen and a core of a mechanically stable and slowly degradable polymer. The minced epithelium was incorporated into one of the collagen layers before transplantation. By mincing the epithelial tissue into small pieces, the pieces could be spread and thereby the propagation of cells was stimulated. After the initial take of the transplants, cell expansion and reorganization would take place and extracellular matrix mature to allow ingrowth of capillaries and nerves and further maturation of the extracellular matrix. The technique minimizes ex vivo manipulations and allow cell harvesting, preparation of autograft, and transplantation to the patient as a simple one-stage intervention. In the future, tissue expansion could be initiated around a 3D mold inside the body itself, according to the specific needs of the patient. Additionally, the technique could be performed in an ordinary surgical setting without the need for sophisticated cell culturing facilities.

Tissue engineering often includes in vitro expansion in order to create autografts for tissue regeneration. In this study a method for tissue expansion, regeneration, and reconstruction in vivo was developed in order to minimize the processing of cells and biological materials outside the body.

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped ...

Epithelial Cell Repopulation and Preparation of Rodent Extracellular Matrix Scaffolds for Renal Tissue Development

This protocol details the generation of acellular, yet biofunctional, renal extracellular matrix (ECM) scaffolds that are useful as small-scale model substrates for organ-scale tissue development. Sprague Dawley rat kidneys are cannulated by inserting a catheter into the renal artery and perfused with a series of low-concentration detergents (Triton X-100 and sodium dodecyl sulfate (SDS)) over 26 hr to derive intact, whole-kidney scaffolds with intact perfusable vasculature, glomeruli, and renal tubules. Following decellularization, the renal scaffold is placed inside a custom-designed perfusion bioreactor vessel, and the catheterized renal artery is connected to a perfusion circuit consisting of: a peristaltic pump; tubing; and optional probes for pH, dissolved oxygen, and pressure. After sterilizing the scaffold with peracetic acid and ethanol, and balancing the pH (7.4), the kidney scaffold is prepared for seeding via perfusion of culture medium within a large-capacity incubator maintained at 37 &#176;C and 5% CO2. Forty million renal cortical tubular epithelial (RCTE) cells are injected through the renal artery, and rapidly perfused through the scaffold under high flow (25 ml/min) and pressure (~230 mmHg) for 15 min before reducing the flow to a physiological rate (4 ml/min). RCTE cells primarily populate the tubular ECM niche within the renal cortex, proliferate, and form tubular epithelial structures over seven days of perfusion culture. A 44 &#181;M resazurin solution in culture medium is perfused through the kidney for 1 hr during medium exchanges to provide a fluorometric, redox-based metabolic assessment of cell viability and proliferation during tubulogenesis. The kidney perfusion bioreactor permits non-invasive sampling of medium for biochemical assessment, and multiple inlet ports allow alternative retrograde seeding through the renal vein or ureter. These protocols can be used to recellularize kidney scaffolds with a variety of cell types, including vascular endothelial, tubular epithelial, and stromal fibroblasts, for rapid evaluation within this system.

This protocol describes decellularization of Sprague Dawley rat kidneys by antegrade perfusion of detergents through the vasculature, producing acellular renal extracellular matrices that serve as templates for repopulation with human renal epithelial cells. Recellularization and use of the resazurin perfusion assay to monitor growth is performed within specially-designed perfusion bioreactors.

This protocol details the generation of acellular, yet biofunctional, renal extracellular matrix (ECM) scaffolds that are useful as small-scale model ...

Three-dimensional Biomimetic Technology: Novel Biorubber Creates Defined Micro- and Macro-scale Architectures in Collagen Hydrogels

Tissue scaffolds play a crucial role in the tissue regeneration process. The ideal scaffold must fulfill several requirements such as having proper composition, targeted modulus, and well-defined architectural features. Biomaterials that recapitulate the intrinsic architecture of in vivo tissue are vital for studying diseases as well as to facilitate the regeneration of lost and malformed soft tissue. A novel biofabrication technique was developed which combines state of the art imaging, three-dimensional (3D) printing, and selective enzymatic activity to create a new generation of biomaterials for research and clinical application. The developed material, Bovine Serum Albumin rubber, is reaction injected into a mold that upholds specific geometrical features. This sacrificial material allows the adequate transfer of architectural features to a natural scaffold material. The prototype consists of a 3D collagen scaffold with 4 and 3 mm channels that represent a branched architecture. This paper emphasizes the use of this biofabrication technique for the generation of natural constructs. This protocol utilizes a computer-aided software (CAD) to manufacture a solid mold which will be reaction injected with BSA rubber followed by the enzymatic digestion of the rubber, leaving its architectural features within the scaffold material.

An innovative biofabrication technique was developed to engineer three-dimensional constructs that resemble the architectural features, components, and mechanical properties of in vivo tissue. This technique features a newly developed sacrificial material, BSA rubber, which transfers detailed spatial features, reproducing the in vivo architectures of a wide variety of tissues.

Tissue scaffolds play a crucial role in the tissue regeneration process. The ideal scaffold must fulfill several requirements such as having proper ...

Optimization of Renal Organoid and Organotypic Culture for Vascularization, Extended Development, and Improved Microscopy Imaging

Embryonic kidney organotypic cultures, and especially pluripotent stem cell-derived kidney organoids, are excellent tools for following developmental processes and modelling kidney disease. However, the models are limited by a lack of vascularization and functionality. To address this, an improved protocol for the method of xenografting cells and tissues to the chorioallantoic membrane (CAM) of an avian embryo to gain vascularization and restoration of blood flow was developed. The grafts are overlaid with custom-made minireservoirs that fix the samples to the CAM and supply them with culture medium that protects the grafts from drying. The improved culture method allows xenografts to grow for up to 9 days. The manuscript also describes how to provide optimal conditions for long-term confocal imaging of renal organoids and organotypic cultures using the previously published Fixed Z-Direction (FiZD) method. This method gently compresses an embryonic organ or organoid between a glass coverslip and membrane in a large amount of medium and provides excellent conditions for imaging for up to 12 days. Together, these methods allow vascularization and blood flow to renal organoids and organotypic kidney cultures with improved confocal imaging. The methods described here are highly beneficial for studying fundamental and applied functions of kidneys ex vivo. Both methods are applicable to various types of tissues and organoids.

This work describes two methods for studying organ development, an improved xenotransplantation setup on chorioallantoic membrane (CAM) from avian embryos that allows for vascularization of cultured embryonic organs and organoids and a novel fixed z-direction organ culture method with modified experimental conditions that allows for high-resolution time-lapse confocal imaging.

Embryonic kidney organotypic cultures, and especially pluripotent stem cell-derived kidney organoids, are excellent tools for following developmental ...

Developmental Biology

Renal Cancer and Healthy Tissue Co-Culture Using Microphysiological Systems

Microfluidic Co-culture of Renal Healthy and Tumor Epithelium to Model Kidney Cancer Progression

Microphysiological systems (MPS) have enabled the introduction of more complex and relevant physiological elements into in vitro models, recreating intricate morphological features in three-dimensional environments with dynamic interactions lacking in conventional models. We implemented a renal cell carcinoma (RCC) co-culture model to recreate the cross-talk between healthy and malignant renal tissue. 
This model is based on the referenced multi-organ platform and consists of co-culturing a reconstructed renal proximal tubule with RCC spheroids. Custom-designed 3D-printed chambers were used to culture human renal epithelial proximal tubule cells (RPTEC) and facilitate their self-assembly into a renal tubule contained in a collagen type I matrix. Caki-1 RCC cells were embedded in an agar collagen matrix, subsequently forming cancer spheroids. Both collagen and agar/collagen gels were optimized to maintain their integrity during cyclic perfusion and withstand shear stress during a minimum culture period of 7 days. 
The gels also enable adequate nutrient supply and cell secretions. Moreover, the agar/collagen gels limit the overproliferation of RCC cells, ensuring relatively homogeneous spheroid size. The MPS chip microfluidic circuits comprise two independent culture chambers with the size of a standard 96-microplate well. The renal tubule and RCC gels populate separate chambers and share the same culture media, which is recirculated approximately twice per minute. Under these conditions, we observed upregulation of immune factor expression and secretion in the renal tubules (interleukin-8 and tumor necrosis factor-alpha). The renal tubules also shift their metabolic activity towards glycolysis under the influence of RCC. This novel approach demonstrates that a co-culture-based MPS can amplify the complexity of RCC in vitro and be employed to study the impact of cancer on non-tumor cells.

Here, we present protocols that detail instructions for implementing 3D cell cultures using collagen and collagen-agarose matrixes in a microphysiological system. These protocols support renal proximal tubule and renal cell carcinoma spheroids co-culture, simulating in vivo conditions and enabling advanced investigation of kidney cancer cell interactions.

Microphysiological systems (MPS) have enabled the introduction of more complex and relevant physiological elements into in vitro models, recreating ...

Cancer Research

Isogenic Kidney Glomerulus Chip Engineered from Human Induced Pluripotent Stem Cells

Chronic kidney disease (CKD) affects 15% of the U.S. adult population, but the establishment of targeted therapies has been limited by the lack of functional models that can accurately predict human biological responses and nephrotoxicity. Advancements in kidney precision medicine could help overcome these limitations. However, previously established in vitro models of the human kidney glomerulus-the primary site for blood filtration and a key target of many diseases and drug toxicities-typically employ heterogeneous cell populations with limited functional characteristics and unmatched genetic backgrounds. These characteristics significantly limit their application for patient-specific disease modeling and therapeutic discovery. 
This paper presents a protocol that integrates human induced pluripotent stem (iPS) cell-derived glomerular epithelium (podocytes) and vascular endothelium from a single patient to engineer an isogenic and vascularized microfluidic kidney glomerulus chip. The resulting glomerulus chip is comprised of stem cell-derived endothelial and epithelial cell layers that express lineage-specific markers, produce basement membrane proteins, and form a tissue-tissue interface resembling the kidney's glomerular filtration barrier. The engineered glomerulus chip selectively filters molecules and recapitulates drug-induced kidney injury. The ability to reconstitute the structure and function of the kidney glomerulus using isogenic cell types creates the opportunity to model kidney disease with patient specificity and advance the utility of organs-on-chips for kidney precision medicine and related applications.

Presented here is a protocol to engineer a personalized organ-on-a-chip system that recapitulates the structure and function of the kidney glomerular filtration barrier by integrating genetically matched epithelial and vascular endothelial cells differentiated from human induced pluripotent stem cells. This bioengineered system can advance kidney precision medicine and related applications.

Chronic kidney disease (CKD) affects 15% of the U.S. adult population, but the establishment of targeted therapies has been limited by the lack of ...

An Efficient Sieving Method to Isolate Intact Glomeruli from Adult Rat Kidney

Preservation of glomerular structure and function is pivotal in the prevention of glomerulonephritis, a category of kidney disease characterized by proteinuria which can eventually lead to chronic and end-stage renal disease. The glomerulus is a complex apparatus responsible for the filtration of plasma from the body. In disease, structural integrity is lost and allows for the abnormal leakage of plasma contents into the urine. A method to isolate and examine glomeruli in culture is critical for the study of these diseases. In this protocol, an efficient method of retrieving intact glomeruli from adult rat kidneys while conserving structural and morphological characteristics is described. This process is capable of generating high yields of glomeruli per kidney with minimal contamination from other nephron segments. With these glomeruli, injury conditions can be mimicked by incubating them with a variety of chemical toxins, including protamine sulfate, which causes foot process effacement and proteinuria in animal models. Degree of injury can be assessed using transmission electron microscopy, immunofluorescence staining, and western blotting. Nephrin and Wilms Tumor 1 (WT1) levels can also be assessed from these cultures. Due to the ease and flexibility of this protocol, the isolated glomeruli can be utilized as described or in a way that best suits the needs of the researcher to help better study glomerular health and structure in diseased states.

The main focus of this protocol is to efficiently isolate viable primary glomeruli cultures with minimal contaminants for use in a variety of downstream applications. The isolated glomeruli retain structural relationships between component cell types and can be cultured ex vivo for a short time.

Preservation of glomerular structure and function is pivotal in the prevention of glomerulonephritis, a category of kidney disease characterized by ...

Medicine

Accessing Collagen Scaffold for Single-Stage Urinary Diversion Surgery in Pigs

Surgical Model for Single-Staged Tissue-Engineered Urothelial Tubes in Minipigs

Reconstructive surgeries are often challenged by a lack of grafting tissue. In the treatment of urogenital malformations, the conventional solution has been harvesting gastrointestinal tissue for non-orthotopic reconstruction due to its abundance to reestablish normal function in the patient. The clinical outcomes after rearranging native tissues within the body are often associated with significant morbidity; thus, tissue engineering holds specific potential within this field of surgery. Despite substantial advances, tissue-engineered scaffolds have not yet been established as a valid surgical treatment alternative, mainly due to the costly and complex requirements of materials, production, and implantation. In this protocol, we present a simple and accessible collagen-based tubular scaffold embedded with autologous organ-specific tissue particles, designed as a conduit for urinary diversion. The scaffold is constructed during the primary surgical procedure, comprises commonly available surgical materials, and requires conventional surgical skills. Secondly, the protocol describes an animal model designed to evaluate the short-term in vivo outcomes post-implantation, with the possibility of additional variations to the procedure. This publication aims to demonstrate the procedure step-by-step, with special attention to the use of autologous tissue and a tubular form.

Tissue-engineered implants for reconstructive surgery rarely progress beyond preclinical trials due to laborious ex vivo culturing, which includes complex and expensive scaffold components. Here, we present a single-staged procedure designed for urinary diversion with an accessible collagen-based tubular scaffold containing autologous micrografts.

Reconstructive surgeries are often challenged by a lack of grafting tissue. In the treatment of urogenital malformations, the conventional solution has ...

Preparation of Decellularized Kidney Scaffolds in Rats

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.

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.

Tissue engineering is a cutting-edge discipline in biomedicine. Cell culture techniques can be applied for regeneration of functional tissues and organs ...

Application of Laser Microdissection to Uncover Regional Transcriptomics in Human Kidney Tissue

Gene expression analysis of human kidney tissue is an important tool to understand homeostasis and disease pathophysiology. Increasing the resolution and depth of this technology and extending it to the level of cells within the tissue is needed. Although the use of single nuclear and single cell RNA sequencing has become widespread, the expression signatures of cells obtained from tissue dissociation do not maintain spatial context. Laser microdissection (LMD) based on specific fluorescent markers would allow the isolation of specific structures and cell groups of interest with known localization, thereby enabling the acquisition of spatially-anchored transcriptomic signatures in kidney tissue. We have optimized an LMD methodology, guided by a rapid fluorescence-based stain, to isolate five distinct compartments within the human kidney and conduct subsequent RNA sequencing from valuable human kidney tissue specimens. We also present quality control parameters to enable the assessment of adequacy of the collected specimens. The workflow outlined in this manuscript shows the feasibility of this approach to isolate sub-segmental transcriptomic signatures with high confidence. The methodological approach presented here may also be applied to other tissue types with substitution of relevant antibody markers.

We describe a protocol for laser microdissection of sub-segments of the human kidney, including the glomerulus, proximal tubule, thick ascending limb, collecting duct and interstitium. The RNA is then isolated from the obtained compartments and RNA sequencing is carried out to determine changes in the transcriptomic signature within each sub-segment.

Gene expression analysis of human kidney tissue is an important tool to understand homeostasis and disease pathophysiology. Increasing the resolution and ...