Isolation of Primary Human Proximal Tubule Epithelial Cells and Their Use in Creating a Microphysiological Model of the Renal Proximal Tubule

We present an optimized protocol for processing whole human kidneys to isolate and culture primary renal proximal tubule epithelial cells and the application of these cells in a three-dimensional, microfluidic, microphysiological platform to recapitulate the renal proximal tubule.

Kidney disease affects over 850 million people globally, including 37 million Americans. Risk factors for chronic kidney disease include environmental influences, genetic predispositions, co-existing medical conditions, and a history of acute kidney injury. These factors often take months or years to develop, complicating longitudinal studies of disease etiology and pathophysiology. Advanced kidney models are needed to improve our understanding of disease mechanisms and enhance nephrotoxicity prediction in drug development. Proximal tubule epithelial cells (PTECs) in the kidney play a critical role in xenobiotic and toxin clearance as well as the reabsorption of essential nutrients. We have previously demonstrated that three-dimensional (3D) microphysiological system (MPS) platforms, populated with isolated primary PTECs, can be used to investigate renal drug interactions, assess nephrotoxicity of compounds, and predict drug clearance. Here, we present protocols for isolating and culturing primary PTECs from whole human kidneys and for seeding them into a 3D MPS platform that mimics in vivo renal physiology. This protocol enables long-term studies supporting PTEC viability, physiological morphology, and functional polarization of key transporter proteins in MPS devices for up to 6 months.

The kidney plays a critical role in the clearance and elimination of a wide array of xenobiotics, toxins, and endogenous compounds from the body. This is achieved by filtering blood to remove waste products and by regulating electrolyte balance, fluid levels, and pH. Each human kidney contains around one million nephrons, the structural and functional units of the kidney¹. Within these nephrons, specialized epithelial cells in the proximal tubules, known as proximal tubule epithelial cells (PTECs), are responsible for reabsorbing essential molecules such as glucose, amino acids, and ions, as well as for secreting drug substrates and potentially toxic substances into the urine^2,3,4. In some cases, PTECs may also reabsorb compounds from the urine back into the bloodstream⁴. Due to their critical role in drug and toxin interactions, primary PTECs isolated from human kidneys provide a valuable tool for studying renal drug-drug interactions (DDIs) and assessing the nephrotoxicity of compounds.

Drug-induced nephrotoxicity poses a significant clinical challenge, as it can lead to acute kidney injury and chronic kidney disease⁵. Therefore, a deeper understanding of the renal proximal tubule physiology is essential for accurately predicting and characterizing the nephrotoxic potential of drugs and toxins. Traditional in vitro models, including immortalized renal cell lines (e.g., RPTEC-TERT1, HK-2), have limitations in mimicking the complex structure and function of the human proximal tubules⁶, which operate under dynamic, laminar flow (with a low Reynolds number) and a uniform extracellular matrix (ECM) in vivo^7,8. Additionally, traditional two-dimensional (2D) models often fail to functionally express major renal transporters (e.g., organic anion transporter 1 and 3 (OAT1 and OAT3), organic cation transporter 2 (OCT2)) due to rapid degradation and internalization of these proteins^6,9,10,11. Animal models, while informative, may not fully replicate human renal physiology and often lack translatability due to species differences in transporter expression and activity¹². For example, mOct1 is basolaterally expressed in mouse PTECs, while in humans, the membrane protein expression of OCT1 in the kidney is undetectable^13,14.

Advancements in microphysiological systems (MPS) and organ-on-a-chip technologies have enabled researchers to develop in vitro models that closely mimic the three-dimensional (3D) architecture and dynamic fluid flow conditions of human organs¹⁵. Our group has previously characterized two MPS models with PTECs^16,17, and has utilized these models to conduct toxicity studies^18,19,20 and predict drug disposition accurately²¹. The use of primary PTECs in these models provides significant advantages due to their ability to retain functional characteristics observed in vivo.

Here, we present the protocols for the isolation of human PTECs from an intact human kidney obtained from a deceased donor via a United Network for Organ Sharing (UNOS)-approved organ procurement organization and applying these cultured human PTECs within an MPS platform.

All work was conducted in compliance with the University of Washington's human tissue handling guidelines. Suitable donors meet the following requirements: less than 36 h of cold ischemic time (CIT), no known history of kidney diseases, dialysis, or any other medical conditions (e.g., diabetes mellitus type 1 or 2, hepatitis B, hepatitis C, human immunodeficiency viruses (HIV), viral/bacterial meningitis, methicillin-resistant Staphylococcus aureus infection, syphilis, sepsis, or Covid-19). The whole human kidneys used in this study were sourced through a UNOS-approved OPO.

1. Biosafety cabinet preparation

1. Spray the hood down with 70% ethanol. Decontaminate all surfaces and items inside the cabinet to create a sterile environment.
2. Lay down blue absorbent pads. Perform all tissue work over these pads.
3. Ensure that necessary equipment is available in the biosafety cabinet and properly sterilized (razor blades, serological pipette tips, serological pipette gun, 100 µm cell strainers, p1000 tips, 1000 µL micropipette, 15 cm circular culture dishes, and conical tubes).

2. Kidney pre-processing preparation

1. Prepare kidney digestion solution by adding 0.75 mg/mL collagenase IV + 0.75 mg/mL dispase II into Dulbecco's Phosphate Buffered Saline (DPBS) with calcium and magnesium. Filter the solution through a 0.2 µm sterile membrane filter.
   NOTE: For a whole human adult kidney, approximately 14 50-mL conical tubes, each containing ~35 mL of digestion solution (totaling 500 mL), are required.
2. Prepare PTEC media by adding 4.425 g of Dulbecco's Modified Eagle Medium (DMEM)/F12 powder without glucose, 0.5 g of D-glucose powder, 0.6 g of sodium bicarbonate powder, 5 mL of 100x insulin-transferrin-selenium supplement (containing 1000 mg/L insulin, 550 mg/L transferrin, and 0.67 mg/L sodium selenite), 0.5 mL of 50 µM hydrocortisone, and 5 mL of 100x antibiotic-antimycotic solution (containing 10000 units/mL of penicillin G sodium, 10000 µg/mL streptomycin sulfate, and 25 µg/mL amphotericin B in 0.85% saline) into 500 mL of sterile autoclaved ultrapure (Type 1) water. Filter the media through a 0.2 µm sterile membrane filter. Thaw fetal bovine serum (FBS) and make 10 mL aliquots in 14 50 mL conical tubes.
3. Gather and label an additional set of 50-mL conical tubes. At this step, ensure there are 3 sets of 14 50 mL tubes.
4. Pre-warm the orbital shaker to 37 °C.

3. Isolation of PTECs from the whole kidney

1. Using a sterile technique, place the shipping container with the tissue sample into the biosafety hood. Remove the bag from the box and spray the outside with 70% ethanol before putting it in the hood.
2. Place the kidney on a round 15-cm culture dish. Aspirate/Discard the remaining media.
3. Remove the surrounding fat and the renal capsule using sterile razor blades. Gently score the renal capsule to create a slit down the center.
4. Using tweezers, pull off the capsule and use razor blades to slice off any fat attached to the kidney. Discard both the capsule and fat into another 15 cm culture dish.
5. Separate the cortex (~1 cm thick) from the medulla. Discard the medulla.
   NOTE: The cortex has a lighter brownish-yellow color compared to the medulla.
6. Using a razor blade, mince the tissue into <1 cm³ pieces until a slurry-like appearance.
7. Add a small amount of kidney digestion buffer into the dish and transfer the slurry into the first set of 50 mL tubes containing 35 mL of the kidney digestion solution. Distribute evenly and do not exceed a total volume of 45 mL for each tube.
8. Transfer all tubes onto the 37 °C orbital shaker and incubate for 30 min at the highest speed that does not cause the tubes to fall out. Transfer the tubes back into the biosafety cabinet after spraying them with 70% ethanol.
9. Invert the tubes to mix and allow the larger pieces of tissue to settle to the bottom. Transfer as much of the solution as possible to the 50 mL conical tubes containing 10 mL of fetal bovine serum (FBS) without transferring intact tissue. Distribute evenly and do not exceed a total volume of 45 mL for each tube.
10. Centrifuge the tubes at 200 g for 7 min. Move the tubes back into the biosafety cabinet after spraying them with 70% ethanol.
11. Carefully aspirate out the supernatant. Add 10 mL of PTEC media into each tube and resuspend the pellets.
12. Strain the resulting cell suspensions through 100 µm cell strainers into new 50-mL conical tubes.
13. Centrifuge the resulting cell filtrates at 400 g for 5 min. Move the tubes back into the biosafety cabinet after spraying them with 70% ethanol.
14. Wash the pellets with 5 mL of DPBS. Resuspend pellets using a p1000 tip.
15. Centrifuge the tubes at 400 g for 5 min. Move the tubes back into the biosafety cabinet after spraying them with 70% ethanol, then aspirate off the supernatants.
16. Repeat step 3.15 two more times for a total of three washes. Then, resuspend cell pellets with 15 mL of PTEC media and plate the cells on sterile cell culture T-75 flasks.
17. Label the flasks appropriately and allow the cells to grow in a sterile incubator at 37 °C and 5% CO₂. Allow PTECs to grow undisturbed in the incubator for 48 h before the first media change.

4. Media changes

1. Aspirate media from the flasks.
2. Wash cells with 5 mL of pre-warmed DPBS.
3. Add 5 mL of pre-warmed PTEC media to the T-25 flasks.
4. Return the flask to the incubator.
5. Change the media every 48 h until the flask reaches at least 70% confluency for passaging or use in experiments.

5. Passaging PTECs

1. Aspirate media from the flask. Add 5 mL of pre-warmed 0.05% Trypsin-EDTA into each T-25 flask and incubate at 37 °C for 1 to 2 min to allow trypsin digestion.
2. Once cells are detached, neutralize trypsin with 5 mL of pre-warmed defined trypsin inhibitor solution.
3. Resuspend 5 times to dislodge any cells still attached to the flask. Then, transfer cell suspension to 15 mL conical tubes and centrifuge at 400 g for 5 min.
4. Aspirate out the supernatant.
   NOTE: Stop here to cryopreserve PTECs and move to that protocol.
5. Resuspend cell pellets with 14 mL of PTEC media per tube. Transfer the cell suspensions to T-75 flasks (1 tube per flask).
6. Perform a T-shake to distribute cells across the flask evenly. Monitor cells and change the media every 48 h.

6. Cryopreserving PTECs

1. After step 5.6., resuspend the cell pellets in 2 mL of 10% DMSO and 90% low glucose PTEC media for each 15 mL conical tube, and transfer 1 mL into one cryovial.
2. Transfer the cryovials to a cell freezing container that allows for temperature-controlled freezing and place the container in a -80 °C freezer for 24 h. After 24 h at -80 °C, transfer cryovials to a liquid nitrogen tank for long-term storage.

7. Thawing PTECs

1. Thaw the vial from -80 °C in the 37 °C water bath. Do not thaw at room temperature (RT). Once the vial has been partially thawed (there should be a small piece of ice visible), add 1 mL of cell suspension to a 15 mL conical tube with 10 mL of pre-warmed PTEC media.
2. Spin down the cell suspension at 400 g for 5 min. Aspirate the supernatant and resuspend cells in 5 mL of pre-warmed low-glucose PTEC media.
3. Transfer the 5 mL cell suspension into a T-25 flask and perform a T-shake to distribute cells across the flask evenly. From here, follow section 4 for media changes.

8. Coating the MPS device with Type I Collagen matrix

1. Aspirate the PBS from the MPS device and remove the device from the package. Wipe the device dry with medical wipes and label it as desired.
2. Place the MPS device in a 15-cm circular culture dish and place it in 4 °C until ready for use. Keep medical wipes on the bottom of the dish to prevent moisture on the MPS device.
3. Keep 1 mL syringes with 22 G Luer-Lok needle blunts attached at -20 °C until use for collagen I injection.
4. For every four MPS devices that need to be filled, keep one 15-mL conical tube on ice. Use one 1-mL syringe with the needle blunt attached for every 15-mL conical tube.
5. Using a sterile technique in a biosafety cabinet, open all the ports on the MPS device. Make sure the slits at the top of each screw valve align vertically with the port.
6. Using a 1 mL syringe/needle blunt assembly, flush the ECM chamber with 1 mL of ethanol into Ports #1, #3, and #6. Aspirate out the ethanol and allow it to evaporate completely (at least 30 s) so it does not penetrate the MPS device from Ports #2, #4, and #5.
7. Place the MPS device back at 4 °C, and keep PTEC media, M199, and 1 N NaOH on ice.
8. Calculate the matrix composition for 1 mL of the Type I collagen mixture for approximately four MPS devices according to Table 1 (scale as needed). In the biosafety cabinet, mix PTEC media, M199, and 1 N NaOH in this order, according to the above calculations, and keep all the tubes on ice.
9. Carefully add the Type I collagen stock with a 1 mL syringe to the premade mix. Mix the collagen mixture by pipetting up and down until the mixture has a stable salmon orange-pink color.
10. Briefly centrifuge the tube to collect the solution at the bottom. Ensure that there are no bubbles in the solution.
11. Place the pre-chilled 1 mL syringes with 22 G Luer-Lok needle blunts and the MPS device into the biosafety cabinet and on ice. Fill the pre-chilled 1 mL syringe with the collagen mixture and avoid bubbles completely.
12. Very slowly and gently inject the collagen solution into Ports #1, #3, and #6 of the chilled device chambers. Hold the device vertically with ports facing down to allow air bubbles to flow up and out of Ports #2, #4, and #5.
    NOTE: There should be a droplet of the collagen mixture at Ports #2, #4, and #5, which indicates that the chambers are fully coated.
13. Remove the syringe and place it back in the 15 mL tube on ice. Turn the screw valve for Ports #2, #4, and #5 to seal the filled chip and place the chip on the culture dish at 4 °C for 30 min.
14. After 30 min, transfer the chip with the dish into the biosafety cabinet in a sterile manner and spread the chips out evenly and individually so they warm up uniformly.
15. Put a moistened medical wipe into each dish (to prevent the collagen I matrix from drying out and peeling away from the MPS device walls) and seal the dishes with a semi-transparent flexible sealing film. Allow the collagen to polymerize overnight at RT.

9. Creating a tubular lumen with Type IV Collagen as the ECM in an MPS device

1. Sterilize C-flex tubing by flushing the tubes with ultrapure (Type 1) water and pure ethanol, then autoclave the whole line of tubing.
2. In the biosafety cabinet, open all the ports of the Type I collagen-filled MPS device (the screw valve will be vertical with the ports) and remove the wire protruding from Ports #1, #3, and #6 to form the tubular lumens.
3. Insert the metal couplers into Ports #1, #3, and #6.
4. Prepare a 10 mL solution of 5 µg/mL Type IV collagen in PTEC media.
5. Insert 24 inches of sterile C-flex tubing onto the metal blunts protruding from Ports #1, #3, and #6.
6. Fill 1 mL syringes fitted with a 22-G Luer-Lok needle blunt with 0.5 mL of Type IV collagen solution.
7. Place the filled syringe on a syringe infusion pump and connect the tubing from Port #1 to the syringe. Repeat this step for Ports #3 and #6.
8. Using the syringe pump, deliver the Type IV collagen solution into the MPS ports at a rate of 10 µL/min for 30 min. Once the ports have been infused, allow the collagen to solidify in a 37 °C incubator overnight before using them the next day.
   NOTE: Type IV collagen-coated MPS devices can be stored for up to 1 week at 4 °C.

10. Seeding primary PTECs in an MPS device

1. Inspect the MPS device under the microscope to ensure the lumen has formed correctly.
2. Place a pre-sterilized 5 mL syringe with a 22 G needle into a 15 mL tube containing 5 mL of ethanol.
3. Obtain a PTEC cell pellet in a 15-mL conical tube from a T-25 flask by following steps 5.1. to 5.3. Add 80 µL of PTEC media to the cell pellet and gently resuspend.
4. Transfer the cell suspension to a 1.5 mL microtube.
5. Remove the dish containing the MPS device from the incubator and place it in the biosafety cabinet. Close Ports #2, #4, and #5, and turn the screw valve horizontally to the ports.
6. Gently flick the PTEC-containing tube to resuspend the cells. Fill the syringe with cells and insert the needle into the injection port that is farthest from the ports of the screw valves.
7. Carefully push down on the plunger of the syringe to inject the cells. Visualize the cells after injection under a microscope.
   NOTE: Cells should be moving through the lumen. If no cells are observed in the lumen, check if the needle has been securely inserted into the injection port and reinsert the needle if needed. A full syringe of cells can fill all three lumens on the device.
8. Continue cell injections with the other two ports. After the injections, close Ports #2, #4, and #5 and place the MPS device back into the 37 °C incubator. Keep the plate level so the cells do not move out of the lumen by gravity.
9. For additional MPS devices, wash the syringe with ethanol after it is emptied and wash with DPBS before drawing up cells.
10. Allow the MPS devices to sit overnight undisturbed.
11. On the next day, connect the MPS devices to the syringe pumps.
12. In the biosafety cabinet, fill 5 mL syringes with PTEC media. Then, fit 22 G Luer-Lok tip blunts with the 24-inch section of C-flex tubing attached to the syringes.
13. Place the PTEC media-loaded syringes into the syringe pump. Prime the lines to remove any air bubbles.
14. Attach the primed tubing lines to Ports #1, #3, and #6 and turn the screw valves to a vertical position to introduce media flow.
15. Set the syringe pump flow rate at 0.5 µL/min. Place the dish with the MPS device back into the 37 °C incubator and allow the cells to form tubules (within 5 to 7 days) under continuous media perfusion before starting an experiment.

Morphology and confluency of isolated primary PTECs over time in 2D culture
Following isolation from the kidney cortex, PTECs were allowed to grow undisturbed for at least 48 h before the first media change. Approximately a week after culturing the cells, small batches of PTECs should appear throughout the culture flask with a uniform epithelial, cobblestone-like morphology (Figure 1A,B). Depending on the growth rates of PTECs from different donors, cell confluency should reach approximately 50%-60% after 10 days in culture (on average) (Figure 1C). After approximately 14 days in culture, cells should reach full confluency (Figure 1D) and are ready to either be passaged, cryopreserved, or seeded in an MPS device. On average, a whole human kidney is able to provide 14 full T-75 flasks full of PTECs. To prevent contamination, it is essential to employ sterile techniques and include antibiotics in the media during the culture period.

Injection of PTECs into an MPS device
After MPS devices are coated with Type I and Type IV collagen, they are ready for culturing PTECs in a 3D format. As shown in Video 1 and Figure 2, PTECs could be seen flowing in the lumen from the injection port (Figure 3). If there are no cells observed, this could indicate leakage or misalignment of the injection and the syringe needle blunt. As shown in previously published studies^17-28, PTEC tubules will form approximately a week after injection into an MPS device.

Figure 1: Morphology and confluency of isolated primary proximal tubule epithelial cells (PTECs) over time. PTECs isolated from a single adult kidney are typically plated in 14 T-75 cell culture flasks. (A) For this representative kidney, after 6 days in culture, small patches of PTECs (boxed in red) with epithelial and cobblestone-like appearance could be observed throughout the cell culture flask, with blood cells still visually present. (B) On Day 8, defined PTEC patches appeared with reduced blood cells after media change.(C) Confluency reached 50 to 60% by Day 10 on average and (D) reached approximately 100% around 2 weeks after isolation and are ready to either be passaged, cryopreserved, or used in an MPS device. Images were taken at 4x magnification using a microscope. Scale bars represent 100 µm. Please click here to view a larger version of this figure.

Figure 2: Workflow overview for creating a proximal tubule model using PTECs in an MPS platform. This flowchart describes the general workflow for using PTECs in an MPS device. Generally, it takes approximately 3 weeks to isolate PTECs from human kidneys to create a mature and stable in vitro 3D proximal tubule. Please click here to view a larger version of this figure.

Figure 3: PTECs after being injected into an MPS device. After the formation of the lumen in the MPS device with collagen I and IV, PTECs can be injected. As shown in this figure and the video, after correctly injecting the PTECs, the cells could be seen flowing through the lumen from the injection port. Please click here to view a larger version of this figure.

Table 1: Sample calculations for preparing 1 mL of the Type I collagen MPS coating mixture. The volume of collagen stock needed should be adjusted according to the stock concentration. The volume of PTEC media can be adjusted to bring up or down the total volume to approximately 1 mL.

Video 1: PTECs flowing in the lumen from the injection port. Please click here to download this Video.

MPS, or organ-on-a-chip technologies, offer a highly relevant in vitro platform to recapitulate key aspects of human physiology, thereby reducing the reliance on animal models in drug development and toxicological assessments. Recently, the FDA Modernization Act 2.0 (2022) has allowed the inclusion of MPS, among other alternatives, in Investigational New Drug applications²⁹. This protocol details a standard operating procedure for the dissociation of PTECs from human donor cortical tissue and their subsequent use in MPS platforms, aiming to model human proximal tubule function and evaluate renal toxicological responses to xenobiotics (Figure 2).

An important factor for maintaining high PTEC viability is minimizing warm ischemic time (WIT). We recommend using kidneys with under 1 h of WIT. While CIT under 24 h is ideal, up to 36 h can be tolerated when kidney availability is limited. Another critical step is proper tissue preparation, which includes removing outer membranes, visceral fat, and thoroughly separating the cortex from the medulla to maximize PTEC purity. We have found that digesting the tissue with collagenase IV and dispase rather than collagenase I alone^10,11,30,31 offers a gentler dissociation that preserves the cell membrane integrity. Additionally, other blends or the inclusion of DNase may be beneficial when there is significant cellular debris or cell clumping in the suspension. It is also essential to mince tissues into pieces of less than 1 cm³ and keep them cold on ice or at 4 °C immediately after digestion to prevent excessive cell damage.

For short-term applications of PTECs in static culture platforms like transwells, the removal of most red blood cells during tissue processing is key to preventing oxidative stress caused by free hemoglobin³². Different cell types in the kidney cortex, including blood cells, can be separated using a Percoll gradient³³. Alternatively, blood cells can be selectively lysed using an ammonium chloride-based buffer³⁴. On the other hand, when using PTECs in MPS devices, moderate levels of contaminating blood cells can be tolerated as these PTECs need to be grown in a standard culture format until confluency. Additionally, prolonged digestion beyond one hour should be avoided, as it can drastically increase contamination risk that may not be resolved by simply adding antimicrobial agents. A 30-min digestion generally provides adequate single-cell suspensions, and if the yield is lower than expected, washing the digested tissue one or two additional times with isotonic buffers such as PBS can often free PTECs that adhere to partially digested tissue. If cell viability is low or adherence is delayed, it is advisable to re-examine the digestion, ensure minimal WIT, and confirm that the culture medium conditions (hormonally-defined, serum-free, 1 g/L glucose) are correct. Verifying that steps post-digestion are performed on ice or at 4 °C (until plating) is also essential for reducing cellular stress. For subsequent cell seeding in the MPS platforms, it is important not to introduce any air bubbles when coating with collagen solutions to maintain a uniform ECM for the PTECs to form structured tubules. Sterile techniques are especially important during MPS setup since the multicompartmental nature of MPS devices increases the risk of contamination.

One major limitation of this protocol is the reliance on donor tissues, which may vary in quality and availability. Prolonged WIT or high CIT can reduce viable cell yield and limit experimental reproducibility. Another limitation is the finite lifespan of primary PTECs, which can typically be maintained for 3 to 4 passages before cell senescence occurs. Lastly, this method yields a mixed population of cells initially, which contains both PTECs and distal tubule epithelial cells (DTECs), following digestion of the renal cortex. While this may not be ideal for short term applications, the purpose is to propagate the PTECs for subsequent use in MPS platforms. Since we employ a defined culture medium without serum, other cell types of much lower percentages, namely mesangial cells and fibroblasts, will not grow at a significant rate. Additionally, using flow cytometry and immunohistochemical (IHC) staining, we previously demonstrated that the majority of the cultured primary cells using this method are PTECs, with a minority of cells expressing DTEC markers based on the staining data of renal epithelial cell markers. Specifically, the 3D cultured PTECs form polarized tubules with PTEC flow cytometric phenotype for neprilysin (CD10) and alanine aminopeptidase (CD13)²³. The IHC staining data also demonstrated that these cells, in both 2D and 3D formats, express PTEC markers lotus tetragonolobus lectin (LTL), sodium-glucose co-transporter 2 (SGLT2), and aquaporin-1 (AQP1), with minimal expression of distal nephron markers, including prominin-2 (PROM2), uromodulin (UMOD), and aquaporin 2 (AQP2)¹⁶. This is expected since DTECs have been shown to dedifferentiate and transdifferentiate towards PTECs in 2D culture³⁰.

While static renal cell cultures and immortalized cell lines are widely used, they often fail to replicate physiological levels of shear stress and the three-dimensional architecture found in the kidney. In contrast, 3D MPS platforms better approximate the in vivo proximal tubule environment by incorporating flow and ECM scaffolds, thereby enhancing the predictive accuracy for nephrotoxicity assessments, as we have shown before^{17,18,19,20,21,22,23,24,25,26,27,28}. The use of primary PTECs rather than immortalized lines confers enhanced physiological relevance, as these cells retain key transporters and enzymes essential for xenobiotic metabolism and clearance. The combination of careful tissue handling, optimized enzymatic digestion, and proper culture conditions ensures that this MPS method aligns with the overarching goals of reducing animal testing while enhancing the translational value of preclinical kidney research²⁹.

This protocol enables the establishment of robust human proximal tubule models in microphysiological devices, thereby facilitating drug screening, nephrotoxicity predictions, and detailed investigations of kidney pathophysiology, particularly for renal clearance mechanisms, drug-transporter interactions, and renal injury pathways. By reducing or replacing the use of animal models, these primary human-based MPS approaches further the 3Rs principle (reduce, refine, replace) and can accelerate drug development pipelines. At the same time, patient-specific MPS holds promise for precision medicine applications, such as personalized drug screening and toxicity profiling. Overall, this reproducible method for isolating and cultivating PTECs from human donor tissues provides critical steps to preserve cell viability and functionality and, when combined with 3D MPS platforms, significantly improves the predictive power of in vitro renal models while advancing toxicological research and drug discovery.

The authors declare that they have no conflicts of interest or financial disclosures relevant to this study.

Portions of this work were supported by the NASA contract 80ARC023CA001, the National Center for Advancing Translational Sciences (NCATS) (U2CTR004867, UH3TR000504, UG3TR002158), jointly by the NCATS and the Center for the Advancement of Science in Space (CASIS) (UG3TR002178), the National Institute of Environmental Health Sciences (NIEHS) (P30ES00703), The National Institute of General Medical Sciences (T32GM007750), an unrestricted gift from Northwest Kidney Centers to the Kidney Research Institute, and the School of Pharmacy at the University of Washington (Ji-Ping Wang Award and Bradley Fellowship).