Isolation, Characterization, And High Throughput Extracellular Flux Analysis of Mouse Primary Renal Tubular Epithelial Cells

Mitochondrial dysfunction in the renal tubular epithelial cells (TECs) can lead to renal fibrosis, a major cause of chronic kidney disease (CKD). Therefore, assessing mitochondrial function in primary TECs may provide valuable insight into the bioenergetic status of the cells, providing insight into the pathophysiology of CKD. While there are a number of complex protocols available for the isolation and purification of proximal tubules in different species, the field lacks a cost-effective method optimized for tubular cell isolation without the need for purification. Here, we provide an isolation protocol that allows for studies focusing on both primary mouse proximal and distal renal TECs. In addition to cost-effective reagents and minimal animal procedures required in this protocol, the isolated cells maintain high energy levels after isolation and can be sub-cultured up to four passages, allowing for continuous studies. Furthermore, using a high throughput extracellular flux analyzer, we assess the mitochondrial respiration directly in the isolated TECs in a 96-well plate for which we provide recommendations for the optimization of cell density and compound concentration. These observations suggest that this protocol can be used for renal tubular ex vivo studies with a consistent, well-standardized production of renal TECs. This protocol may have broader future applications to study mitochondrial dysfunction associated with renal disorders for drug discovery or drug characterization purposes.


Introduction
Renal tubular epithelial cell (TEC) function is strongly associated with the overall health condition of the kidney. Pathological signaling in the kidney causes the dedifferentiation of TECs, which plays a major role in kidney fibrosis and chronic kidney disease (CKD) 1,2 . As a highly energetic organ, the kidney is second only to the heart in oxygen consumption, primarily through mitochondrial oxidative phosphorylation 3 . Electron microscopy studies have revealed a positive correlation of mitochondrial morphological changes to pathological events in the renal tubules 4 . Mitochondrial dysfunction in TECs causes renal fibrosis through epithelial to mesenchymal transition 5 and defective fatty acid oxidation 6 . Fibrosis is a progressive renal pathology that results in CKD. Therefore, understanding the energetic status of renal TECs is a necessity to uncover the pathophysiology of CKD.
There are > 20 cell types in the adult kidney 7 . To study the function of TECs, a primary culture of the renal epithelial cells is needed as a platform for molecular biology applications such as chemical treatments and genetic manipulations. Importantly, genetic manipulations can be done in vivo in mice via transgenesis or by using AAV gene delivery techniques 8 so that the isolated primary cells would already be genetically manipulated. The isolation of primary renal tubular cells from mice 9,10 , rats 11,12,13 , canines 14 , rabbits 15,16 , and humans 17,18 has been reported with purification steps to yield pure proximal tubular cells. In these previously published protocols that focus on the isolation of proximal tubular cells, gradient centrifugation and sorting experiments were performed for purification purposes 19 . While these protocols are valuable for studying proximal tubules, they are not sufficient when both proximal and distal tubules are needed to be studied. For example, our study on the Alport syndrome has revealed that both proximal and distal renal tubules play important roles in the disease progression 1. Remove the renal capsules and medulla, mince both kidneys into tiny pieces, and incubate them in 10 mL of a digestion buffer in a 37 °C oven with gentle rotation for 5 min. 2. Remove any undigested kidney tissues by passing the buffer through a 70-μm filter. Add 10 mL of culture media to stop the digestion. 3. To collect tubular cells, centrifuge the filtered cell suspension at 50 x g for 5 min to collect the first pellet. Transfer the supernatant to a new tube and add 5 mL of culture media, centrifuge it at 50 x g for 5 min to ensure all tubular cells are collected into the second pellet.
The centrifugation is at a lower speed to primarily pellet heavy tubules. Later, after the cells recover from the isolation, the pure tubular culture is centrifuged at a higher speed during the sub-cultures. 4. Resuspend the first pellet in 20 mL of culture media and centrifuge it at 50 x g for 5 min to collect the third pellet. 5. Resuspend the second and third pellets in 1 mL of culture media. Mix 10 µL of the cell suspension with 10 µL of Trypan blue, load the mixture into chamber A of a counting slide, and record the cell viability from the automatic cell counter (see Table of Materials). 6. Seed up to 10 7 cells (a heterogeneous population) onto a single 60-mm dish pre-coated with collagen and let the tubular cells attach overnight.

Primary Tubular Cells Sub-culture and Characterization
1. On day 1 after the isolation, collect culture media and centrifuge it at 50 x g for 5 min to pellet any floating tubules. Remove the supernatant and resuspend the cell pellet in 4 mL of fresh culture media and plate it back to the same culture dish. 2. On day 4 after the isolation, remove the old culture media and add fresh media. 3. On day 7 after the isolation, detach the cells by incubating them at 37 °C in 2 mL of 0.25% trypsin-EDTA for 5 min. Add 3 mL of culture media to stop the reaction and collect the cells by centrifugation at 250 x g for 5 min. 4. To sub-culture and characterize the cells from P0 to P1, seed 5,000 cells per well onto a 24-well plate coated with collagen I as described above. 5. 24 h after step 4.4, fix the cells at P1 with 4% PFA for 10 min, permeabilize them with 0.2% Triton X-100 for 3 min, and block them with 10% donkey serum (DS) for 1 h at room temperature. 6. Dilute, to 1:100 in 10% DS, each of the following proteins: the proximal tubular markers angiotensinogen (AGT) and aquaporin 1 (AQP1); the distal tubular marker E-cadherin; the mesangial marker CD90/Thy1; and the macrophage markers EGF-like module-containing mucin-like hormone receptor-like 1 (F4/80) and cluster of differentiation 68 (CD68), and incubate them with cells overnight at 4 °C. 7. The next day, detect any staining using 1:200 anti-Rabbit, anti-Mouse, or anti-Rat fluorescent secondary antibodies for 45 min. Take images under confocal microscopy to confirm the expression of the markers, as shown in Figure 2A. 8. On day 3 after the sub-culture of P1, detach the cells for a sub-culture and characterization at P2 by a staining of the tubular, mesangial, and macrophage markers described in step 4.5. Image the staining under confocal microscopy to confirm the expression of the markers, as shown in Figure 2A. 9. On day 3 after the sub-culture of P2, detach the cells for a sub-culture and characterization at P3 by a staining of the tubular, mesangial, and macrophage markers described in step 4.5. Image the staining under confocal microscopy to confirm the expression of the markers, as shown in Figure 2A. 10. Prepare the tissue staining: 1. Air-dry frozen wild-type and Col4a3 -/kidney slides for 1 h at room temperature and fix them with 4% PFA for 10 min. Permeabilize them with 0.2% Triton X-100 for 10 min and block them with 10% donkey serum (DS) for 1 h at room temperature. Add antibodies against the marker proteins described in step 4.5 at 1:200 and incubate them at 4 °C overnight.

Mitochondria Bioenergetics Assay
1. Seed P1 tubular cells at 20,000, 30,000, or 40,000 cells per well in 100 μL of culture media onto a 96-well microplate pre-coated with 5 μg/ cm 2 collagen I the day before the extracellular flux assays.
2. For the hydration of the sensor cartridge, lift the sensor cartridge and fill each well of the plate with 200 μL of a calibration solution. Carefully load the cartridge back to submerge the sensors in the calibration solution. Place the cartridge in a 37 °C oven without CO 2 for at least 7 h prior to use. For the best results, overnight cartridge hydration is recommended. 3. Prepare the compounds: prepare 8 µM oligomycin, 9 µM FCCP, and 20 µM rotenone/antimycin A mixture in both glucose (described in step 1.10) and fatty acid (described in step 1.11) extracellular flux assay media. 4. Change the media: aspirate the cell culture media, add 175 µL of glucose or fatty acid assay media (dependent on the compound that is being worked in, see step 5.3), and incubate them for 1 h in a 37 °C CO 2 -free incubator. 5. Load the cartridge ports with 25 µL of the following compounds: 8 μM oligomycin for port A to achieve a final concentration of 1 µM (note: as each well will contain 175 µL of media, the compound will get diluted 8x), 9 μM FCCP in port B to achieve a final concentration of 1 µM (note: as each well will contain 175 µL of media plus 25 µL of the solution injected from port A, the compound will get diluted 9x), and 20 µM rotenone/antimycin A in port C to achieve a final concentration of 2 µM for each compound (note: as each well will contain 175 µL of media plus 50 µL of the solution injected from ports A and B, the compound will get diluted 10x). 6. Add water to all wells in port D and all other ports of the background wells (no cells). Incubate the cartridge in a 37 °C CO 2 -free incubator for 10 min.
1. Choose Standard Assay. Press Assay Wizard. Using the Compounds tab, assign the compound layout and use the Groups and Labels tab to label the experimental groups. Remember to assign the empty wells (without cells) as background. 2. Under the Protocol tab, set the following mix and measure cycles using the available commands as indicated in Table 1: calibrate, mix for 2 min, wait for 2 min, and measure for 3 min (repeat this cycle 2 -3x); inject port A, mix for 2 min, wait for 2 min, measure for 3 min (repeat this cycle 2 -3x); inject port B, mix for 2 min, wait for 2 min, measure for 3 min (repeat this cycle 2 -3x); inject port C, mix for 2min, wait for 2 min, measure for 3 min (repeat this cycle 2 -3x). Press End Wizard. It is also possible to save the current template for future use. 3. Press Start to begin the calibration. The analyzer then automatically ejects the plate holder and asks for the cartridge plate to be inserted.
9. When the calibration step is done (usually in 20 -25 min), press the prompt command to change the cartridge plate to the cell plate and continue the run. 10. When the run is completed, transfer the data, and remove the 96-well plate. Add Hoechst (1:1,000) to each of the assay wells and incubate them for 5 min at 37 °C. Normalize the OCR data to cell count by a measurement of the Hoechst fluorescence reading at a 355-nm excitation and a 460-nm emission.

Kidney Perfusion and Digestion Yield Highly Viable Tubular Epithelial Cells:
Mouse renal tubular epithelial cells were isolated following the steps outlined in sections 1 -3 of the protocol described above.
After the digestions, a heterogeneous population of kidney cells, incompletely digested tubules, and other tissue debris that is smaller than 70 µm were plated onto the culture dish on isolation day. Changes from day 0 to day 1 after the isolation are usually expected to be seen only in the cell attachment rather than in the cell growth. Looking through the floating heterogenous population, only a few tubular cells were attached at day 1 ( Figure 1A). A re-collection of the cells at day 1, a centrifugation, and a re-plating helped to remove light debris and settle the small tubule pieces for the tubular cell release. From day 1 to day 3, changes were expected not only in a better cell attachment but also in a remarkably improved cell growth rate that was observed with a tripled cell density as compared to day 1 ( Figure 1A). In the initial growth phase, the cells formed several colonies and populated around the colonies. From this point forward, the isolated cells were fully recovered and displayed a healthy proliferation. By day 5, the cells were at an 80 -90% confluency in a 60-mm Petri dish with some spaces between cell to cell and colony to colony (Figure 1A).

Sub-culture and Characterization of the Isolated Tubular Cells:
The isolated renal tubular epithelial cells were sub-cultured for a characterization following the steps outlined in section 4 of the protocol described above.
From day 5, the cells fully recovered from the isolation and started to proliferate vigorously. One week after the isolation, the cells grew to confluency in a 60-mm Petri dish. After 1 week in a culture at passage 0, the cells were ready to be sub-cultured to passage 1 and, subsequently, for 2 more passages. Similar growth patterns were observed in passage 1 and passage 2. Usually, it takes less than a week for cells at passage 1 and 2 to grow to confluency for further sub-cultures ( Figure 1B). The continuous culture of confluent cells from passage 0 to passage 2 showed an extensive dome formation 23,24 (Figure 1B), suggesting that the isolated cells maintained a healthy status where they excreted liquids similar to an in vivo status. This caused the monolayer of the cells to lift off the plate but stay connected via tight junctions.
To characterize the cells in the culture, we performed immunofluorescent staining in the cultured cells from passage 1 through passage 4, as well as in a control cell line-human proximal renal epithelial HK-2 cells. Proximal tubular markers, aquaporin 1 (AQP1) 25 and angiotensinogen (AGT) 9 , the distal tubular marker E-cadherin 25 , the epithelial marker smooth muscle actin (SMA) 26,27,28 , macrophage markers F4/80 29 and CD68 30 , and the mesangial marker thymocyte differentiation antigen 1 (Thy1/CD90) 9 were used for the characterization studies. Both proximal tubular proteins AQP1 and AGT were consistently highly expressed in the isolated tubular cells from passage 1 to passage 4 as well as in the positive control HK-2 proximal epithelial cells (Figure 2A). The distal tubular protein E-cadherin was expressed in the isolated tubular cells through passage 4 and was also observed in the HK-2 cells (Figure 2A). SMA was expressed abundantly in the isolated tubular cells and in the control HK-2 cells, consistent with published reports 26,27 . On the other hand, the mesangial protein Thy1 and macrophage protein F4/80 were absent in both the isolated tubular cells and the control HK-2 cells (Figure 2A). CD68 showed a minimal expression in the HK-2 cells and in the isolated tubular cells at passage 1 and passage 2, and then its expression became undetectable from passage 3 through passage 4 (Figure 2A). The results suggest that the cells isolated following this protocol are a mix of proximal and distal tubular cells. To compare the expressions of these marker proteins in vivo, we performed staining in frozen kidney tissues. Tubular markers, including AQP1, AGT, and E-cadherin, and mesangial protein Thy1 were found highly expressed in kidneys harvested from a wild-type healthy mouse (Figure 2B). Low expressions of F4/80 and CD68 were observed in wild-type kidneys but extensively expressed in kidneys harvested from a Col4a3 -/mouse that developed renal failure with a macrophage infiltration 20,31 ( Figure 2C).

Mitochondrial Bioenergetics Assay on Isolated Primary Tubular Cells:
The mitochondrial respiration assay steps are outlined in section 5 of the protocol described above.
The mitochondrial respiration of isolated primary tubular cells is measured by an extracellular flux analysis of the oxygen consumption rate (OCR) at different plating densities. To titrate the plating density, 20,000, 30,000, and 40,000 primary TECs per well were seeded onto a 96-well XF96 microplate the day before (approximately 20 h before) the extracellular flux assay (Figure 3A). Following the extracellular flux analysis, the OCR measurements were then normalized to the cell counts by a quantification of Hoechst staining. Plating the TECs at 20,000, 30,000, or 40,000 cells/well resulted in an average basal OCR of 25, 45, or 50 pmol/min, respectively ( Figure 3A). Moreover, microscopic images of the plated cells revealed that 40,000 cells/well covered the entire surface of the bottom of the microplate wells better than the other plating densities ( Figure 3B). Even though the maximal OCR did not increase using 40,000 cells compared to the density of 30,000 cells, we recommend a cell density of around 40,000 cells/well for optimal interactions between cells and compounds.
An extracellular flux assay yields a number of important parameters for assessing the bioenergetics of renal TECs. For instance, as fatty acid oxidation is shown to be specifically defective in TECs, the use of media containing fatty acid substrate (palmitate) along with the inhibitor of glycolysis (2DG) can serve as a useful tool to directly evaluate fatty acid oxidation in TECs at passages 1 and 2 ( Figure 3C). In the case of renal fibrosis, the overall respiration capacity of the cells is expected to be lower than that of a healthy kidney even though the glucose-based media may not reveal any differences.
Taken together, the extracellular flux assay, especially to assess the fatty acid oxidation capacity, can be utilized as an informative measure to evaluate the energetic status of renal TECs in which pathological changes affecting the bioenergetics profile play a major role in renal fibrosis and the progression to kidney failure.
Steps suspension and centrifuge it to pellet the unattached tubules and cells the second day after the cell plating. This low-speed centrifugation step further removes other cell types that are lighter than tubular cells and allows for unattached tubules and tubular cells to settle.
The identification of proper cell density is the first and key step for a successful extracellular flux assay. The results showed that 40,000 cells per well on an XF96 microplate is ideal for primary tubular cells in both a fatty acid and a glucose-based respiration assay ( Figure 3C). In this protocol, isolated tubular cells were used for the extracellular flux assay at passages 1 and 2. The cells sub-cultured to passage 3, although they maintained an expression of the tubular markers ( Figure 2) and a decent performance in the bioenergetics assays (Figure 3A), and showed decreased basal respiration levels compared to passage 2 (shown by comparing OCR in the rightmost panels of Figure 3A to Figure 3C). This decrease may not affect substantially healthy tubular cells (for example, ones isolated from young wild-type mice). However, for studies on cells isolated from CKD mouse models that already have a diminished mitochondrial respiration, higher passages of the cells may cause a further decrease in the basal respiration which would affect the results of the extracellular flux assay. In the studies conducted here, the cells from both passage 1 and passage 2 showed high basal respiration levels. Therefore, following this protocol, we recommend using these two early passages for mitochondrial respiration studies with cells isolated from both healthy and diseased animals. Cells from passage 2 should still be taken into consideration if the passage 1 sub-culture does not yield sufficient cells for the flux assay. In addition to bioenergetics studies, our previous research shows that primary TECs at passage 3 can be extremely useful for treatments with compounds followed by protein and RNA studies (data not shown). That being said, we suggest that investigators using this protocol to isolate tubular cells should carefully choose the optimal passage for different research applications.
The working principle of the extracellular flux analysis is based on the interactions between the injected compounds and the respiration chain complexes and the effect of the uncoupler. Oligomycin is an inhibitor of complex V (ATP synthase) and is used to distinguish ATP-linked oxygen consumption and the oxygen consumption that is required to overcome the regular proton leak across a mitochondrial inner membrane 32 . FCCP uncouples oxygen consumption from the ATP production by disrupting the mitochondrial membrane potential. Thus, it provides a measurement of the maximal respiration capacity as it circumvents the limited capacity of a proton ion efflux by ATP synthase by allowing a proton transport through the membrane. Antimycin-A, a complex III inhibitor, and rotenone, a complex I blocker, are used in combination to shut down the entire mitochondrial respiration allowing a differentiation between the mitochondrial vs. the non-mitochondrial oxygen consumption in the cells. These compounds should always be titrated for a specific cell type before the extracellular flux assay to determine the optimal concentrations that yield the optimal OCR curves. Here, we recommend 1 µM of oligomycin, 1 µM of FCCP, and 2 µM of rotenone/antimycin A for the extracellular flux assay on primary TECs.
In conclusion, this protocol provides a simple and cost-effective way to isolate renal primary proximal and distal tubular epithelial cells that can be used for assessing mitochondrial bioenergetics ex vivo. While this protocol can be useful in a wide range of molecular biology studies exploring the biological function of renal tubular epithelial cells, we acknowledge its limitations when applying it to studies needing pure proximal or distal tubules. For example, studies on the Lowe Syndrome, a selective proximal tubular dysfunction 33 , or studies on distal renal tubular acidosis, a distal tubular dysfunction 34 , would require a more sophisticated protocol for cell isolation and purification. However, for the majority of the studies that compare tubules vs. glomeruli, and for studies to screen potential mitochondrial respiration regulators in tubular cells in general, the protocol provides a feasible high throughput approach. Therefore, this protocol may have broad applications to study mitochondrial dysfunction associated with renal disorders for drug discovery or target validation purposes.

Disclosures
The authors have nothing to declare.