The authors would like to acknowledge the Lynn and Mike Garvey Imaging Laboratory at the Institute for Stem Cell and Regenerative Medicine and LifeCenter NorthWest. They would also like to acknowledge the financial support of National Institutes of Health grants, UH2/UH3 TR000504 (to J.H.) and DP2DK102258 (to Y.Z.), NIH T32 training grant DK0007467 (to R.J.N.), and an unrestricted gift from the Northwest Kidney Centers to the Kidney Research Institute.

Human kidneys were isolated by LifeCenter Northwest following ethical guidelines set by the Association of Organ Procurement Organizations. This protocol follows animal care and cell culture guidelines outlined by the University of Washington.
1. Preparation of Human Kidney Tissue
<ol>
	<li>Preparation of decellularization solution
	<ol>
		<li>Sterilize a 5000 mL beaker and a 70 x 10 mm stir bar.</li>
		<li>Mix 1:1000 (weight:volume) sodium dodecyl sulfate (SDS) in autoclaved deionized water in the beaker. Leave the solution on a stir plate at approximately 200 rpm for 24 h or until the SDS is completely dissolved. 
		Note: Typically, 2500 mL of 1% SDS solution is sufficient to decellularize a single human kidney.</li>
		<li>Transfer the solution to a 500 mL sterile vacuum filter and filter it into sterilized sealable containers.</li>
	</ol>
	</li>
	<li>Processing of kidney tissue
	<ol>
		<li>Wash and autoclave a pair of forceps, two hemostat clamps, a pair of general service grade scissors, two scalpel blade handles, a 1000 mL beaker covered with aluminum foil, and a 36 x 9 mm stir bar.</li>
		<li>Line a tissue culture hood with underpad. Place the beaker, a sterile tissue culture dish (150 x 25 mm), and the whole kidney organ into the hood. Fill the beaker with 500 mL of 1% SDS solution. 
		Note: Human kidneys were received on ice from LifeCenter NorthWest.</li>
		<li>Place the kidney in the sterile tissue culture dish (Figure 1A). Remove all perirenal fat by lightly shaving around the renal capsule with a scalpel (Figure 1B).</li>
		<li>Make a shallow 8-10 cm incision with the scalpel, just deep enough to break open the renal capsule without damaging the underlying cortex tissue, across the superior end of the kidney. Remove the renal capsule by peeling it away from the cortex tissue with two hemostat clamps (Figure 1C).</li>
		<li>Bisect the kidney along the coronal plane by using the scalpel along the lateral side of the kidney (Figure 1D). Isolate cortex tissue from both halves by carving out the medullar region with the scalpel (Figure 1E) and dice the cortex tissue into 0.5 cm3 pieces (Figure 1F). Remove any large visible vessels.</li>
	</ol>
	</li>
	<li>Isolation of extracellular matrix
	<ol>
		<li>In a tissue culture hood, fill a 1000 mL beaker with 500 mL of 1% SDS solution. Place the diced cortex tissue and stir bar into the beaker containing SDS solution. Cover the beaker with autoclaved aluminum foil and place it on a stir plate at approximately 400 rpm outside of the tissue culture hood.</li>
		<li>After the cortex tissue has been on the stir plate for 24 h, bring the beaker into a tissue culture hood and add a 40 &#181;m sterile cell strainer made with nylon mesh. Fill a separate 1000 mL beaker with 200 mL of bleach and place it in the tissue culture hood.</li>
		<li>Pipette out the SDS solution through the cell strainer into the beaker containing bleach. Pipette out all SDS solution until only decellularized tissue and the cell strainer remain in the beaker. 
		Note: The cell strainer should prevent any tissue from being removed during solution aspiration.</li>
		<li>Leave the cell strainer in the beaker and fill with 500 mL of fresh SDS solution. Cover the beaker with the same aluminum foil and place onto a stir plate at the same speed as before.</li>
		<li>Repeat steps 1.3.1-1.3.3 every 24 hours with fresh SDS solution for a total of five days.</li>
		<li>Rinse decellularized tissue with autoclaved DI water every 24 h for 3 days total, following the technique outlined in steps 1.3.1-1.3.3.</li>
		<li>Rinse decellularized tissue with cell culture grade water every 24 h for 2 days total, following the technique outlined in steps 1.3.1-1.3.3.</li>
		<li>Repeat steps 1.3.1-1.3.2. Transfer the decellularized tissue (referred to as kECM from this point on) into a 30 mL self-standing conical tube and fill it with cell culture grade water until all the tissue is submerged.</li>
	</ol>
	</li>
</ol>
2. Fabrication of Hydrogel Stock Solution
<ol>
	<li>Mechanical processing of decellularized tissue
	<ol>
		<li>In a tissue culture hood, mechanically homogenize the kECM within the conical tube with a tissue homogenizer for 2 min. 
		Note: Homogenized kECM should resemble an opaque solution with no visible pieces of ECM.</li>
		<li>Submerge the conical tube containing the kECM in liquid nitrogen until boiling surrounding the tube no longer persists. Store the kECM at -4 &#730;C overnight.</li>
	</ol>
	</li>
	<li>Lyophilization of frozen decellularized tissue
	<ol>
		<li>Slightly loosen the conical tube cap to allow for gas exchange and place the tube into a lyophilization machine. Lyophilize the kECM for three days or until it resembles a fine white powder. Store at -4 &#730;C.</li>
	</ol>
	</li>
	<li>Chemical digestion and solubilization of gel
	<ol>
		<li>Autoclave a 20 mL scintillation vial and cap, a 15.9 x 7.9 mm stir bar, and one pair of fine-tip forceps.</li>
		<li>Weigh the lyophilized kECM and calculate the volume of HCl and mass of pepsin needed to solubilize the kECM to a 3% (30 mg/mL) solution using the following equations, where mpepsin is the mass of pepsin, mtissue is the mass of lyophilized tissue, and VHCl is the volume of 0.01 N HCl: 
		<img alt="Equation 1" src="/files/ftp_upload/58314/58314eq1.jpg" /> 
		<img alt="Equation 2" src="/files/ftp_upload/58314/58314eq2.jpg" /></li>
		<li>In a tissue culture hood, add porcine gastric pepsin, 0.01 N HCl, and the stir bar to the scintillation vial, and leave it on a stir plate at approximately 500 rpm until all the pepsin has dissolved. Transfer the lyophilized kECM to the scintillation vial and leave the solution on a stir plate at approximately 500 rpm for three days.</li>
	</ol>
	</li>
</ol>
3. Hydrogel Gelation
<ol>
	<li>Kidney ECM hydrogel preparation
	<ol>
		<li>Gel the hydrogel by mixing the kECM hydrogel stock solution with 1 N NaOH, 10x Media Supplement (M199), and cell culture media. Keep all the solutions on ice. 
		Note: Final gel concentrations of 7.5 mg/mL were used for cell culture. 1 mL of kECM gel was sufficient for cell culture experiments presented.</li>
		<li>Determine the volume of workable kECM gel produced and volume of stock kECM hydrogel needed by using the following equation, where Vfinal is the volume of gel created, Vstock kECM is the volume of stock kECM hydrogel needed, Cstock kECM is the concentration of the stock kECM hydrogel, and Cfinal is the concentration of the final gel: 
		<img alt="Equation 3" src="/files/ftp_upload/58314/58314eq3v2.jpg" /></li>
		<li>Determine the volume of neutralizing reagents needed by using the following equations, where VNaOH is the volume of 1 N NaOH, V10X is the volume of M199 10X media supplement, and V1X is the volume of cell culture media: 
		<img alt="Equation 4" src="/files/ftp_upload/58314/58314eq4.jpg" /> 
		<img alt="Equation 5" src="/files/ftp_upload/58314/58314eq5.jpg" /> 
		<img alt="Equation 6" src="/files/ftp_upload/58314/58314eq6.jpg" /></li>
		<li>In a tissue culture hood, pipette the neutralizing reagents (NaOH, M199, and cell culture media) into a sterile 30 mL self-standing conical tube. Mix the neutralizing reagent solution with a microspatula.</li>
		<li>Use a sterile 1 mL syringe to transfer the appropriate volume of stock kECM hydrogel to the neutralizing reagent solution. Use a microspatula to gently mix the solution until a homogeneous in color hydrogel solution is obtained. 
		Note: Avoid introducing air bubbles by stirring slowly and gently.</li>
		<li>To incorporate cells into the kECM hydrogel, subtract 10 &#181;L of cell culture media (V1X) from the neutralizing solution volume calculations in step 3.1.1.3.
		<ol>
			<li>Suspend cells into 10 &#181;L of cell culture media. Determine the number of cells to be suspended by using the following equation, where #cells implies the number of cells to suspend and Vfinal is the volume of gel created: 
			<img alt="Equation 7" src="/files/ftp_upload/58314/58314eq7.jpg" /> 
			Note: 300,000 cells/mL is the concentration of cells used in the kECM gel.</li>
			<li>Pipette the 10 &#181;L of cell suspended solution into the final kECM gel after the kECM stock solution has been mixed with neutralizing reagent solution. Stir the solution with a microspatula until the cells are evenly distributed.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Use a 1 mL syringe to fill a desired cell culture device with the kECM hydrogel.</li>
	<li>Allow the gel to set at 37 &#730;C for 1 h before transferring or plating cells.</li>
</ol>

The authors have nothing to disclose.

Matrices provide important mechanical and chemical cues that govern cell behavior. Synthetic hydrogels are able to support complex 3-dimensional patterning but fail to provide the diverse extracellular cues found in physiological matrix microenvironments. Hydrogels derived from native ECM are ideal materials for both in vivo and in vitro studies. Previous studies have used decellularized ECM hydrogels to coat synthetic biomaterials to prevent host immunological responses33,34, to differentiate stem cells35,36,37,38, as a substrate for 2D and soft lithography cell culture39,40,41,42, and in preparation of bioinks for 3-dimensional printing43,44,45,46. ECM hydrogels provide cells with proper signaling to initiate adhesion and control further proliferation and differentiation2,47.
The ratios and composition of major components of the ECM, such as collagens, laminins, elastin, fibronectin, and glycosaminoglycans, are highly dependent on the anatomical location and functionality of the tissue48,49,50. It is well established that using nonspecific or nonnative ECM-derived hydrogels will elicit improper responses from cells21,51,52,53,54. In the kidney, ECM composition varies widely between anatomical locations1,2. It is, therefore, important to differentiate between regions such as the cortex, medulla, or papilla before fabricating hydrogels for experimental use.
The kECM matrix fabricated here preserves native kidney cortex ECM protein ratios following decellularization while also enabling gelation to a physiologically relevant mechanical stiffness3,25,28,29,30. Serum albumin, a blood plasma protein, was detected in trace amounts within the decellularized cortex tissue, possibly due to binding to heparin that escaped decellularization and rinsing55,56. Although pepsin, a non-native chemical to kidney cortex ECM, is present in the stock kECM solution, it only accounts for 2.5% (w/v) of the final gelled product. Furthermore, pepsin becomes deactivated with the addition of the neutralizing reagent solution57.
The kECM hydrogel serves as a substrate upon which kidney cortex-specific cells can be maintained under physiological conditions. We demonstrated that this matrix can support HKMEC growth on a planar surface. These HKMECs maintained a quiescent physiological state as determined through functional assays, phenotypic expression, and genetic expression58. In contrast, these cells became activated when collagen-I, a matrix component correlated with kidney fibrosis, was added to the kECM hydrogel at a 1:1 ratio59. By comparison, when human umbilical vein endothelial cells were cultured on collagen-I, they were quiescent, and when mixed with kECM they became activated12. Accordingly, collagen-I is known to be an integral component of the basement membrane composition within the umbilical cord and is decreased under pathological states such as preeclampsia38,60. These results highlight the importance of providing cells with tissue-specific cues to maintain quiescence and also how manipulation of the ECM milieu can affect homeostasis.
Whereas all steps in the outlined procedure are important, several steps are critical in ensuring the viability of the fabricated hydrogel. The degree of homogenization of the decellularized cortex tissue will vary based on the technique or equipment used. It is important to find a technique that will best homogenize the tissue. During solubilization, the pH should be kept at 3-4 to ensure the pepsin is active. When mixing the hydrogel with neutralizing reagents, the gel must be mixed thoroughly and without the introduction of air bubbles. If a uniform mixture is difficult to obtain, check the pH of the gel solution to ensure it is neutral.
The representative results presented in this method demonstrate how a physiologically relevant matrix for the in vitro study of kidney cells can be achieved. Decellularized kidney cortex ECM provides an ideal base material to support kidney-derived cell growth as ECM protein ratios are conserved in the final hydrogel product3,25. The gelled product can also achieve physiologically relevant mechanical properties28,29,30. The kECM hydrogel allows for proper cell-matrix interactions and has been shown to elicit greater physiological behavior from kidney cells than collagen-I when cultured on a planar surface. Furthermore, the hydrogel can be seeded with kidney-specific cells prior to gelation to model a more physiologically relevant 3-dimensional environment. The composition of the kECM hydrogel can also be easily manipulated. For example, mixing the hydrogel with varying amounts of collagen-I prior to gelation could produce matrices that mimic progressive states of renal fibrosis31,32. The tunability of this ECM-derived hydrogel to mimic known diseased ECM compositions warrants further investigation and will enable the future study of kidney diseases.

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. The complex molecular composition regulates both structural and functional properties of tissue. Structural proteins provide cells with spatial awareness and allow for adhesion and migration1. Bound ligands interact with cell surface receptors to control cell behavior2. Kidney ECM contains a plethora of molecules whose composition and structure varies depending on anatomical location, developmental stage, and disease state3,4. Recapitulating the complexity of ECM is a key aspect in studying kidney-derived cells in vitro.
Previous attempts at replicating ECM microenvironments have focused on decellularizing whole tissue to create scaffolds capable of recellularization. Decellularization has been performed with chemical detergents such as sodium dodecyl sulfate (SDS) or non-ionic detergents, and it utilizes either whole organ perfusion or immersion and agitation methods5,6,7,8,9,10,11,12,13. The scaffolds presented here preserve the structural and biochemical cues found in native tissue ECM; furthermore, recellularization with donor-specific cells has clinical relevance in reconstructive surgery14,15,16,17,18,19. However, these scaffolds lack structural flexibility and are therefore incompatible with many current devices used for in vitro studies. To overcome this limitation, many groups have further processed decellularized ECM into hydrogels20,21,22,23,24. These hydrogels are compatible with injection molding and bioink and circumvent micrometer scale spatial constraints that decellularized scaffolds place on cells. Furthermore, molecular composition and ratios found in native ECM are preserved3,25. Here we demonstrate a method to fabricate a hydrogel derived from kidney cortex ECM (kECM).
The purpose of this protocol is to produce a hydrogel that replicates the microenvironment of the kidney cortical region. Kidney cortex tissue is decellularized in a 1% SDS solution under constant agitation to remove cellular matter. SDS is commonly used to decellularize tissue because of its ability to quickly remove immunological cellular material6,7,9,26. The kECM is then subject to mechanical homogenization and lyophilization5,6,9,11,26. Solubilization in a strong acid with pepsin results in a final hydrogel stock solution20,27. Native kECM proteins that are important for structural support and signal transduction are preserved3,25. The hydrogel can also be gelled to within one order of magnitude of native human kidney cortex28,29,30. This matrix provides a physiological environment that has been used to maintain the quiescence of kidney-specific cells compared to hydrogels from other matrix proteins. Furthermore, matrix composition can be manipulated, for example, through the addition of collagen-I, to model disease environments for the study of renal fibrosis and other kidney diseases31,32.

<table><tbody><tr><td>&lt;strong&gt;Preparation of Kidney Tissue&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>5000 mL Beaker</td><td>Sigma-Aldrich</td><td>Z740589</td><td></td></tr><tr><td>Sodium Dodecyl Sulfate (SDS)</td><td>Sigma-Aldrich</td><td>436143</td><td></td></tr><tr><td>Sterile H&lt;sub&gt;2&lt;/sub&gt;O</td><td></td><td></td><td>Autoclaved DI H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Stir Bar (70 x 10 mm)</td><td>Fisher Science</td><td>14-512-128</td><td></td></tr><tr><td>500 mL Vacuum Filter</td><td>VWR</td><td>97066-202</td><td></td></tr><tr><td>Stir Plate</td><td>Sigma-Aldrich</td><td>CLS6795420D</td><td></td></tr><tr><td>1000 mL Beaker</td><td>Sigma-Aldrich</td><td>CLS10031L</td><td></td></tr><tr><td>Forceps</td><td>Sigma-Aldrich</td><td>F4642</td><td>Any similar forceps may be used</td></tr><tr><td>Scissor-Handle Hemostat Clamp</td><td>Sigma-Aldrich</td><td>Z168866</td><td></td></tr><tr><td>Dissecting Scissors</td><td>Sigma-Aldrich</td><td>Z265977</td><td></td></tr><tr><td>Scalpel Handle, No. 4</td><td>VWR</td><td>25859-000</td><td>Any similar scalpel handle may be used</td></tr><tr><td>Scalpel Blade, No. 20</td><td>VWR</td><td>25860-020</td><td>Any similar scalpel blade may be used</td></tr><tr><td>Stir Bar (38.1 x 9.5 mm)</td><td>Fisher Science</td><td>14-513-52</td><td></td></tr><tr><td>Absorbent Underpad</td><td>VWR</td><td>82020-845</td><td></td></tr><tr><td>Petri Dish (150 x 25 mm)</td><td>Corning</td><td>430597</td><td></td></tr><tr><td>Autoclavable Biohazard Bag</td><td>VWR</td><td>14220-026</td><td></td></tr><tr><td>Sterile Cell Strainer (40 um)</td><td>Fisher Science</td><td>22-363-547</td><td></td></tr><tr><td>Cell Culture Grade Water</td><td>HyClone</td><td>SH30529.03</td><td></td></tr><tr><td>30 mL Freestanding Tube</td><td>VWR</td><td>89012-778</td><td></td></tr><tr><td>&lt;strong&gt;Fabrication of ECM Gel&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Tissue Homogenizer Machine</td><td>Polytron</td><td>PCU-20110</td><td></td></tr><tr><td>Freeze Dryer</td><td>Labconco</td><td>7670520</td><td></td></tr><tr><td>20 mL Glass Scintillation Vials and Cap</td><td>Sigma-Aldrich</td><td>V7130</td><td></td></tr><tr><td>Stir Bar (15.9 x 8 mm)</td><td>Fisher Science</td><td>14-513-62</td><td></td></tr><tr><td>Pepsin from Porcine Gastric Mucosa</td><td>Sigma-Aldrich</td><td>P7012</td><td></td></tr><tr><td>0.01 N HCl</td><td>Sigma-Aldrich</td><td>320331</td><td>Dilute to 0.01 N HCl with cell culuture water</td></tr><tr><td>&lt;strong&gt;Kidney ECM Gelation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1 N NaOH (Sterile)</td><td>Sigma-Aldrich</td><td>415413</td><td>Dilute to 1 N in cell culture grade water</td></tr><tr><td>Medium 199</td><td>Sigma-Aldrich</td><td>M4530</td><td></td></tr><tr><td>15 mL Conical Tube</td><td>ThermoFisher</td><td>339651</td><td></td></tr><tr><td>Cell Culture Media</td><td>ThermoFisher</td><td>11330.032</td><td>Dulbecco&#039;s Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F12)</td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>Gibco</td><td>10082147</td><td></td></tr><tr><td>Antibiotic-Antimycotic 100X</td><td>Life Technologies</td><td>15240-062</td><td></td></tr><tr><td>Insulin, Transferrin, Selenium, Sodium Pyruvate Solution (ITS-A) 100X</td><td>Life Technologies</td><td>51300-044</td><td></td></tr><tr><td>1 mL Syringe</td><td>Sigma-Aldrich</td><td>Z192325</td><td></td></tr><tr><td>Microspatula</td><td>Sigma-Aldrich</td><td>Z193208</td><td></td></tr></tbody></table>

fabricating a kidney cortex extracellular matrix-derived hydrogel

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust mechanical support for in vitro cell culture but lack the necessary protein and ligand composition to elicit physiological behavior from cells. This manuscript describes a fabrication method for a kidney cortex ECM-derived hydrogel with proper mechanical robustness and supportive biochemical composition. The hydrogel is fabricated by mechanically homogenizing and solubilizing decellularized human kidney cortex ECM. The matrix preserves native kidney cortex ECM protein ratios while also enabling gelation to physiological mechanical stiffnesses. The hydrogel serves as a substrate upon which kidney cortex-derived cells can be maintained under physiological conditions. Furthermore, the hydrogel composition can be manipulated to model a diseased environment which enables the future study of kidney diseases.

The kECM hydrogel provides a matrix for kidney cell culture with similar chemical composition as the native kidney microenvironment. To fabricate the hydrogel, kidney cortex tissue is mechanically isolated from a whole kidney organ and diced (Figure 1). Decellularization with a chemical detergent (Figure 2A.1-A.3) followed by rinses with water to remove detergent particles (Figure 2A.4-A.6) yields isolated kidney cortex ECM. Histological evaluation confirms typical basal laminar proteins such as collagen-IV and laminin and structural proteins such as collagen-I are preserved, with vitronectin as the only noted exception (Figure 2B). Furthermore, protein composition, including preservation of isoforms, in the ECM remains consistent with observed values in native kidney ECM (Figure 3). The ECM (Figure 4A) is mechanically homogenized and lyophilized (Figure 4B) then solubilized to produce the final kECM hydrogel (Figure 4C). The kECM hydrogel appeared opaque with small amounts of visible tissue and was not as viscous as traditional collagen-I hydrogel. Rheological measurement of gelled kECM at a concentration of 15 mg/mL revealed a complex modulus (dynamic elastic modulus) of around 800 Pa over the linear range of strain values, significantly greater than that of 7.5 mg/mL collagen-I (p = 1.602E-14). Human kidney peritubular microvascular endothelial cells (HKMECs) cultured on collagen-I, kECM, and a 1:1 mixture gel showed differences in phenotype, specifically in CD31 expression around cell surfaces and junctions (Figure 5). HKMECs cultured on collagen-I displayed uniform CD31 expression while HKMECs cultured on the two gels containing kECM displayed reduced CD31 expression in uneven distributions. Matrix type did not appear to affect the high PV1 and low VWF expression in the HKMECs.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58314/58314fig1.jpg" /> 
Figure 1: Isolation of kidney cortex tissue. Mechanical processing of a whole kidney organ to isolate the cortex tissue as a base material for the kECM hydrogel. Mechanical isolation begins with (A) the removal of the perirenal adipose tissue. Large pieces of adipose tissue can be torn away from the kidney with hemostat clamps. Remaining pieces of adipose tissue can be removed by running a scalpel against the renal capsule at an angle. (B) The renal capsule is best removed by making a shallow incision along the superior end of the kidney and (C) peeling the renal capsule away from the underlying tissue with hemostat clamps. (D) Bisecting the kidney along the coronal axis allows for the visualization of the cortex and medulla regions. (E) Isolation of the cortex tissue is best done by carving out pieces of the medulla tissue with a scalpel. The color of the cortical region is noticeably darker than that of the medullar region and can be used differentiate the two anatomically distinct tissues. Final processing of the cortex tissue involves (F) dicing the tissue into 0.5 cm3 pieces to aid in subsequent decellularization. <a href="https://www.jove.com/files/ftp_upload/58314/58314fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58314/58314fig2.jpg" /> 
Figure 2: Decellularization of cortex tissue. Visual and histological characterization of decellularized tissue. (A.1) Submerging diced cortex tissue in 1% SDS solution causes lysing of cells and removal of cellular material. After (A.2) 1 h and (A.3) 24 h, the tissue begins to lose color, indicating cellular matter is being removed. By (A.4) 120 h the tissue is blanched and only the ECM remains. Rinses with water at (A.5) 24 h and (A.6) 120 h show no visible changes to the tissue. (B) Immunofluorescence staining of untreated and decellularized cortex tissue reveals near complete removal of cellular matter and preservation of major structural proteins (collagen-IV = Col-IV; laminin = LAM; fibronectin = FN; heparin sulfate proteoglycans = HSPG; and vitronectin = VN). Scale = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/58314/58314fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58314/58314fig3.jpg" /> 
Figure 3: Mass spectroscopy of decellularized tissue. Analysis of decellularized cortex tissue by mass spectrometry to determine ECM protein composition. All ratios are measured as mass percent. (A) Structural and other proteins associated with the basal lamina were present with collagen-IV and -I being the most highly represented. (B.1) Collagen-IV A1 and A2 chains, ubiquitous in all basement membranes, were conserved. Collagen-IV A3 and A5 chains, present only in basement membranes of the glomerulus, were also detected. (B.2) Common isoforms of laminins and (B.3) collagen-I were also detected. This figure was reproduced with permission12. <a href="https://www.jove.com/files/ftp_upload/58314/58314fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58314/58314fig4.jpg" /> 
Figure 4: Fabrication of kECM hydrogel. Mechanical and chemical processing of decellularized cortex tissue yields a workable kECM hydrogel with sufficient mechanical properties following gelation with neutralizing reagents. (A) Decellularized cortex tissue is mechanically homogenized with a tissue homogenizer until no visible pieces of ECM remain. (B) A coarse powder was yielded after 3 days under lyophilization. (C) Solubilization in HCl and chemical digestion with pepsin in a scintillation vial resulted in a workable kECM hydrogel. The hydrogel was opaque and of a low viscosity. (D) Physical characterization of the kECM hydrogel following gelation with neutralizing reagents. Rheological experiments were performed with a 30 mm diameter parallel plate system. The sample edges were protected with mineral oil and the loading platform was set at 37 &#730;C. The kECM hydrogel was allowed to gel for 1 h prior to testing. Viscoelastic properties of the gel were measured with a strain sweep between 0.01 to 20%. Three samples of both kECM and collagen-I were tested (n = 3). <a href="https://www.jove.com/files/ftp_upload/58314/58314fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58314/58314fig5.jpg" /> 
Figure 5: Cell growth characterization. Characterization of morphological differences in HKMECs grown on different matrices. Hydrogels were mixed with neutralizing reagents and set at 37 &#730;C for 45 min in open-faced polydimethylsiloxane molds. HKMECs were seeded on the surface of the gels and kept in culture for 72 h before being fixed and stained. Immunofluorescent images of HKMECs cultured in (A and D) 7.5 mg/mL collagen-I gel; (B and E) 7.5 mg/mL kECM gel; and (C and F) 7.5 mg/mL 1:1 mixture gel. (A-C): red = CD31; green = VWF; blue = nuclei; scale bar = 50 &#181;m. (D-F): red = F-actin; green = PV1; blue = nuclei; scale bar = 50 &#181;m. This figure was reproduced with permission12. <a href="https://www.jove.com/files/ftp_upload/58314/58314fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel at JoVE.com

Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel

Here we present a protocol to fabricate a kidney cortex extracellular matrix-derived hydrogel to retain the native kidney extracellular matrix (ECM) structural and biochemical composition. The fabrication process and its applications are described. Finally, a perspective on using this hydrogel to support kidney-specific cellular and tissue regeneration and bioengineering is discussed.

Extracellular matrix (ECM) provides important biophysical and biochemical cues to maintain tissue homeostasis. Current synthetic hydrogels offer robust ...

fabricating-a-kidney-cortex-extracellular-matrix-derived-hydrogel

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JoVE Journal

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10.4K Views.  University of Washington. Here we present a protocol to fabricate a kidney cortex extracellular matrix-derived hydrogel to retain the native kidney extracellular matrix (ECM) structural and biochemical composition. The fabrication process and its applications are described. Finally, a perspective on using this hydrogel to support kidney-specific cellular and tissue regeneration and bioengineering is discussed.

Video: Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel

Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field of 3D templates is rising due to the potential resemblance of those materials to the natural extracellular matrix. Protein-based hydrogels are particularly promising because they can easily be functionalized and can achieve defined structures with adjustable physicochemical properties. However, the production of macroporous 3D templates for cell culture applications using natural materials is often limited by their weaker mechanical properties compared to those of synthetic materials. Here, different methods were evaluated to produce macroporous bovine serum albumin (BSA)-based hydrogel systems, with adjustable pore sizes in the range of 10 to 70 &#181;m in radius. Furthermore, a method to generate channels in this protein-based material that are several hundred microns long was established. The different methods to produce pores, as well as the influence of pore size on material properties such as swelling ratio, pH, temperature stability, and enzymatic degradation behavior, were analyzed. Pore sizes were investigated in the native, swollen state of the hydrogels using confocal laser scanning microscopy. The feasibility for cell culture applications was evaluated using a cell-adhesive RGD peptide modification of the protein system and two model cell lines: human breast cancer cells (A549) and adenocarcinomic human alveolar basal epithelial cells (MCF7).

Different methods to manipulate three-dimensional architecture in protein-based hydrogels are evaluated here with respect to material properties. The macroporous networks are functionalized with a cell-adhesive peptide, and their feasibility in cell culture is evaluated using two different model cell lines.

Hydrogels are recognized as promising materials for cell culture applications due to their ability to provide highly hydrated cell environments. The field ...

Biochemistry

Processing of Human Cardiac Tissue Toward Extracellular Matrix Self-assembling Hydrogel for In Vitro and In Vivo Applications

Acellular extracellular matrix preparations are useful for studying cell-matrix interactions and facilitate regenerative cell therapy applications. Several commercial extracellular matrix products are available as hydrogels or membranes, but these do not possess tissue-specific biological activity. Because perfusion decellularization is usually not possible with human heart tissue, we developed a 3-step immersion decellularization process. Human myocardial slices procured during surgery are first treated with detergent-free hyperosmolar lysis buffer, followed by incubation with the ionic detergent, sodium dodecyl sulfate, and the process is completed by exploiting the intrinsic DNase activity of fetal bovine serum. This technique results in cell-free sheets of cardiac extracellular matrix with largely preserved fibrous tissue architecture and biopolymer composition, which were shown to provide specific environmental cues to cardiac cell populations and pluripotent stem cells. Cardiac extracellular matrix sheets can then be further processed into a microparticle powder without further chemical modification, or, via short-term pepsin digestion, into a self-assembling cardiac extracellular matrix hydrogel with preserved bioactivity.

Processing of Human Cardiac Tissue Toward Extracellular Matrix Self-assembling Hydrogel for In Vitro and In Vivo Applications

This protocol describes the complete decellularization of human myocardium while preserving its extracellular matrix components. Further processing of the extracellular matrix results in the production of microparticles and a cytoprotective self-assembling hydrogel.

Acellular extracellular matrix preparations are useful for studying cell-matrix interactions and facilitate regenerative cell therapy applications. ...

Developmental Biology

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 ...

Construction of Modular Hydrogel Sheets for Micropatterned Macro-scaled 3D Cellular Architecture

Hydrogels can be patterned at the micro-scale using microfluidic or micropatterning technologies to provide an in vivo-like three-dimensional (3D) tissue geometry. The resulting 3D hydrogel-based cellular constructs have been introduced as an alternative to animal experiments for advanced biological studies, pharmacological assays and organ transplant applications. Although hydrogel-based particles and fibers can be easily fabricated, it is difficult to manipulate them for tissue reconstruction. In this video, we describe a fabrication method for micropatterned alginate hydrogel sheets, together with their assembly to form a macro-scale 3D cell culture system with a controlled cellular microenvironment. Using a mist form of the calcium gelling agent, thin hydrogel sheets are easily generated with a thickness in the range of 100 - 200 &#181;m, and with precise micropatterns. Cells can then be cultured with the geometric guidance of the hydrogel sheets in freestanding conditions. Furthermore, the hydrogel sheets can be readily manipulated using a micropipette with an end-cut tip, and can be assembled into multi-layered structures by stacking them using a patterned polydimethylsiloxane (PDMS) frame. These modular hydrogel sheets, which can be fabricated using a facile process, have potential applications of in vitro drug assays and biological studies, including functional studies of micro- and macrostructure and tissue reconstruction.

We describe the fabrication of micropatterned hydrogel sheets using a simple process, which can be assembled and manipulated in a freestanding form. Using these modular hydrogel sheets, a simple macro-scaled 3D cell culture system can be generated with a controlled cellular microenvironment.

Hydrogels can be patterned at the micro-scale using microfluidic or micropatterning technologies to provide an in vivo-like three-dimensional (3D) tissue ...

Bioprinting Cellularized Constructs Using a Tissue-specific Hydrogel Bioink

Bioprinting has emerged as a versatile biofabrication approach for creating tissue engineered organ constructs. These constructs have potential use as organ replacements for implantation in patients, and also, when created on a smaller size scale as model "organoids" that can be used in in vitro systems for drug and toxicology screening.
Despite development of a wide variety of bioprinting devices, application of bioprinting technology can be limited by the availability of materials that both expedite bioprinting procedures and support cell viability and function by providing tissue-specific cues. Here we describe a versatile hyaluronic acid (HA) and gelatin-based hydrogel system comprised of a multi-crosslinker, 2-stage crosslinking protocol, which can provide tissue specific biochemical signals and mimic the mechanical properties of in vivo tissues. 
Biochemical factors are provided by incorporating tissue-derived extracellular matrix materials, which include potent growth factors. Tissue mechanical properties are controlled combinations of PEG-based crosslinkers with varying molecular weights, geometries (linear or multi-arm), and functional groups to yield extrudable bioinks and final construct shear stiffness values over a wide range (100 Pa to 20 kPa). Using these parameters, hydrogel bioinks were used to bioprint primary liver spheroids in a liver-specific bioink to create in vitro liver constructs with high cell viability and measurable functional albumin and urea output. This methodology provides a general framework that can be adapted for future customization of hydrogels for biofabrication of a wide range of tissue construct types.

We describe a set of protocols that together provide a tissue-mimicking hydrogel bioink with which functional and viable 3-D tissue constructs can be bioprinted for use in in vitro screening applications.

Bioprinting has emerged as a versatile biofabrication approach for creating tissue engineered organ constructs. These constructs have potential use as ...

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

Ultrathin Porated Elastic Hydrogels As a Biomimetic Basement Membrane for Dual Cell Culture

The basement membrane is a critical component of cellular bilayers that can vary in stiffness, composition, architecture, and porosity. In vitro studies of endothelial-epithelial bilayers have traditionally relied on permeable support models that enable bilayer culture, but permeable supports are limited in their ability to replicate the diversity of human basement membranes. In contrast, hydrogel models that require chemical synthesis are highly tunable and allow for modifications of both the material stiffness and the biochemical composition via incorporation of biomimetic peptides or proteins. However, traditional hydrogel models are limited in functionality because they lack pores for cell-cell contacts and functional in vitro migration studies. Additionally, due to the thickness of traditional hydrogels, incorporation of pores that span the entire thickness of hydrogels has been challenging. In the present study, we use poly-(ethylene-glycol) (PEG) hydrogels and a novel zinc oxide templating method to address the previous shortcomings of biomimetic hydrogels. As a result, we present an ultrathin, basement membrane-like hydrogel that permits the culture of confluent cellular bilayers on a customizable scaffold with variable pore architectures, mechanical properties, and biochemical composition.

Current bilayer culture models do not allow for functional in vitro studies that mimic in vivo microenvironments. Using polyethylene glycol and a zinc oxide templating method, this protocol describes the development of an ultrathin biomimetic basement membrane with tunable stiffness, porosity, and biochemical composition that closely mimics in vivo extracellular matrices.

The basement membrane is a critical component of cellular bilayers that can vary in stiffness, composition, architecture, and porosity. In vitro studies ...

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

Synthesis of Decellularized Cartilage Extracellular Matrix Hydrogels

Decellularized cartilage extracellular matrix (DC-ECM) hydrogels are promising biomaterials for tissue engineering and regenerative medicine due to their biocompatibility and ability to mimic natural tissue properties. This protocol aims to produce DC-ECM hydrogels that closely mimic the native ECM of cartilage tissue. The protocol involves a combination of physical and chemical disruption and enzymatic digestion to remove the cellular material while preserving the structure and composition of the ECM. The DC-ECM is cross-linked using a chemical agent to form a stable and biologically active hydrogel. The DC-ECM hydrogel has excellent biological activity, spatial structure, and biological induction function, as well as low immunogenicity. These characteristics are beneficial in promoting cell adhesion, proliferation, differentiation, and migration and for creating a superior microenvironment for cell growth. This protocol provides a valuable resource for researchers and clinicians in the field of tissue engineering. Biomimetic hydrogels can potentially enhance the development of effective tissue engineering strategies for cartilage repair and regeneration.

This paper introduces a new method for the synthesis of decellularized cartilage extracellular matrix (DC-ECM) hydrogels. DC-ECM hydrogels have excellent biocompatibility and provide a superior microenvironment for cell growth. Therefore, they can be ideal cell scaffolds and biological delivery systems.

Decellularized cartilage extracellular matrix (DC-ECM) hydrogels are promising biomaterials for tissue engineering and regenerative medicine due to their ...

Comparative Study of Decellularization Techniques for Bovine Lung ECM Hydrogels

Development and Characterization of Decellularized Lung Extracellular Matrix Hydrogels

The use of extracellular matrix (ECM)-derived hydrogels in tissue engineering has become increasingly popular, as they can mimic cells&#39; natural environment in vitro. However, maintaining the native biochemical content of the ECM, achieving mechanical stability, and comprehending the impact of the decellularization process on the mechanical properties of the ECM hydrogels are challenging. Here, a pipeline for decellularization of bovine lung tissue using two different protocols, downstream characterization of the effectiveness of decellularization, fabrication of reconstituted decellularized lung ECM hydrogels and assessment of their mechanical and cytocompatibility properties were described. Decellularization of the bovine lung was pursued using a physical (freeze-thaw cycles) or chemical (detergent-based) method. Hematoxylin and Eosin staining was performed to validate the decellularization and retention of major ECM components. For the evaluation of residual collagen and sulfated glycosaminoglycan (sGAG) content within the decellularized samples, Sirius red and Alcian blue staining techniques were employed, respectively. Mechanical properties of the decellularized lung ECM hydrogels were characterized by oscillatory rheology. The results suggest that decellularized bovine lung hydrogels can provide a reliable organotypic<span style="font-size: 11pt; font-family: Calibri, sans-serif; color: black; background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">&#160;alternative to commercial ECM products by retaining most native ECM components. Furthermore, these findings reveal that the decellularization method of choice significantly affects gelation kinetics as well as the stiffness and viscoelastic properties of resulting hydrogels.

Author Spotlight: Optimizing Bovine Lung Decellularization for Organotypic Hydrogels

The protocol elucidates two distinct decellularization methodologies applied to native bovine pulmonary tissues, providing a comprehensive account of their respective characterizations.

The use of extracellular matrix (ECM)-derived hydrogels in tissue engineering has become increasingly popular, as they can mimic cells&#39; natural ...

Fabricating a Kidney Cortex Extracellular Matrix-Derived Hydrogel

Summary

Explore More Videos

Easy Manipulation of Architectures in Protein-based Hydrogels for Cell Culture Applications

Processing of Human Cardiac Tissue Toward Extracellular Matrix Self-assembling Hydrogel for In Vitro and In Vivo Applications

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

Construction of Modular Hydrogel Sheets for Micropatterned Macro-scaled 3D Cellular Architecture

Bioprinting Cellularized Constructs Using a Tissue-specific Hydrogel Bioink

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Ultrathin Porated Elastic Hydrogels As a Biomimetic Basement Membrane for Dual Cell Culture

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

Synthesis of Decellularized Cartilage Extracellular Matrix Hydrogels

Development and Characterization of Decellularized Lung Extracellular Matrix Hydrogels