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Method Article

Rodent Kidney Explant Culture and Establishment of Primary Ureteric Bud Epithelial Cell Culture

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DOI:

10.3791/69462

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December 30th, 2025

 ,  ,  , 
1Stem Cells and Metabolism Research Program Unit, Faculty of Medicine, University of Helsinki, 2GM Unit, Helsinki Institute of Life Science, University of Helsinki, 3Department of Pediatric Surgery, Section of Pediatric Urology, New Children's Hospital, Helsinki University Hospital, 4Laboratory Animal Centre, University of Helsinki

In This Article

Summary

This protocol describes the establishment of the most commonly used mammalian kidney culture, which allows renal growth and differentiation in a Trowell-type air-liquid interphase system. We also detail how to set up primary ureteric bud epithelial cell cultures from the early kidney rudiments.

Abstract

The developing kidney is a widely used model system to study mammalian organogenesis. Renal differentiation is orchestrated by reciprocal inductive tissue interactions among mesenchyme, stroma, and epithelium. Branching morphogenesis of epithelial ureteric bud promotes not only embryonic kidney growth and 3D patterning but also induces mesenchyme-to-epithelium transitions within the surrounding nephron progenitor pool, thereby ensuring nephron formation required for functional filtration.

Critical cell fate decisions within all three tissue layers, epithelium, mesenchyme, and stroma, underpin a dynamic balance between progenitor self-renewal and differentiation equilibrium. These complex, multilayered regulatory interactions are inherently dependent on the structural integrity of the organ, making intact kidney models essential for studying morphogenetic processes and cellular behaviors in their native context.

Kidney organ culture methods provide insightful platforms for visualizing developmental processes and dissecting morphological, cellular, and molecular mechanisms that govern renal differentiation. A variety of kidney culture methods exist, each offering distinct advantages. In this article, we detail the establishment of a traditional Trowell-type air-liquid interphase culture system for in vitro kidney development. To enhance the resolution of cellular and molecular analyses, we also describe the setup of a primary ureteric bud epithelial cell culture, which enables targeted investigation of epithelial-specific mechanisms.

Finally, we demonstrate that the traditional kidney organ culture technique is applicable to the embryonic kidney of a novel model organism, the spiny mouse (Acomys dimidiatus). This opens interesting innovative avenues for stem cell and regeneration research, as Acomys has a remarkable functional repair capacity across several organ systems studied to date.

Introduction

The practice of culturing organs dates back to the beginning of the 20th century1. Kidneys were among the first organs to be cultured in vitro when Rienhoff introduced the hanging drop method for embryonic chicken kidneys in 19222. In 1950's, embryonic mouse (Mus musculus) kidney cultures established the importance of mesenchyme-epithelium interactions in guiding developmental processes3,4. Since then, tissue and organ cultures have served as a method to study branching morphogenesis5,6,7,8,9,10,11,12, inductive signaling3,4,13,14, differentiation15,16,17,18,19, and their mechanisms. Still, highly relevant in vitro organ culture allows the addition of different signaling molecules, agonists, or antagonists/inhibitors to the cultures to monitor their effects on morphological events20,21,22.

Culturing genetically modified (GM) kidneys has provided valuable information on the renal differentiation regulation6,7,9,20,23,24,25,26,27. Genetic labeling methods and live imaging techniques enable visualization of specific cell populations or even individual cells, facilitating the analysis of their contribution to the morphological processes9,10,12,18,24,25,26,28,29,30.

At least four different types of kidney ex vivo culture systems exist: 1) the hanging drop method, where tissue is cultured in a small medium drop hanging from a coverslip or a petri dish lid2,31; 2) Trowell-type air liquid interphase culture, where explants are placed on a membrane on top of a metal grid or hanging from a plastic frame (commercial inserts) with media underneath the membrane32,33; 3) low-volume culture, where a very small amount (~85 µL) of culture medium and the tissue are placed inside a silicon ring on a glass slide34; and 4) Fixed Z-Direction (FiZD) culture optimized for confocal live imaging and where the kidney is compressed between a glass slide and a membrane separated by spacers35. The Trowell-type kidney culture is presented here for its simplicity, high cost-efficiency, ease of visualization of morphological processes, and since it is not linked to special microscopy6.

Kidney culture methods provide a means to understand organogenesis in a holistic manner. Primary cell cultures present a supplementary approach for studying biochemical and molecular mechanisms, especially those involved in cellular adhesion and its changes, delamination, and motility. For these, in vitro culture of epithelial cells is an excellent model36. The monolayer culture of primary ureteric bud (UB) is established in approximately 48 h, after which the cells do not divide but survive alive for approximately two weeks7. Such a non-proliferative phenotype can be effectively utilized to analyze, e.g., cell migration with scratch assay or inter-cellular and cell-to-matrix adhesion forces7,37,38. Additionally, cells derived from global GM mice may provide essential cell-type-specific information of renal differentiation6,7.

The permanent kidney (metanephros) development starts with the process known as kidney induction, where reciprocal inductive tissue interactions between the epithelial Wolffian duct (WD, also called nephric duct) and the metanephric mesenchyme (MM) induce the formation of the initial ureteric bud (UB)13,39,40,41. The MM secretes signals such as fibroblast growth factors (FGFs) and glial cell line-derived neurotrophic factor (GDNF), causing cells in the WD first to cluster and then to outgrow to form the initial primary UB, which invades into the MM5,40,42,43. In the mouse, the most common mammalian model used to study organogenesis, this happens at embryonic day 10.5 (E10.5) and in humans around gestational week 4-540,44,45,46.

The process of kidney induction is followed by the start of UB branching morphogenesis. Here, the bud tip first elongates, then balloons into an ampulla to finally bifurcate into a T-bud structure with two brand new tips (Figure 1)12,37,39,47,48. The newly formed UB tips not only receive regulatory cues from the MM but also actively influence the fate and behavior of the MM6,42,49. Accordingly, the differentiation of nephron epithelium is induced by each UB tip (Figure 1). In this way, the extent of UB branching determines the final nephron number in the kidney. The branching morphogenesis continues with iterative cycles until birth, but the majority of the branching happens during 10 to 13 rounds, taking place by E15.5 in mice, after which the rate of new tip formation decreases significantly30,49. Interestingly, the nephron formation continues longer, and the morphological and molecular basis for this is only beginning to be elucidated50,51,52,53,54.

The UB is molecularly and functionally divided into two subpopulations, the tip and the stalk, which differ in their gene expression profiles and proliferation capacity12,55,56,57,58,59. Most of the bifurcations that generate new branches occur in the tips, while differentiation takes place in the trunk region where the two types of collecting ductal cells, principal and intercalated cells, are born11,12,60,61,62. Studies heavily dependent on the in vitro kidney cultures have identified that the UB tips host a progenitor population capable of giving rise to new tips, the trunk, the ureter, and the entire collecting duct system5,25. Despite the known tip-trunk regionalization, the mechanisms controlling the differentiation versus self-renewal decisions within collecting duct progenitors are poorly known. Importantly, the in-situ identity of collecting duct progenitors and the means to identify them are only emerging37.

The UB tip forms a fundamental niche for the establishment of the cap mesenchyme, which constitutes the nephron progenitor (NP) pool13,63. On one side, the cap mesenchyme is surrounded by the UB, and on the other side by the stromal progenitors giving rise to interstitial cells, such as pericytes and mesangial cells (Figure 1)27,64,65,66,67,68. During each UB branch cycle, a subset of NP cells self-renew to keep up the NP population, and another portion is induced to differentiate into functional nephrons66. Much less is known about the behavior of stromal progenitors, which though persist around the NP cells in the periphery of the growing embryonic kidney65,69.

The cap mesenchyme hosting the NP cells is highly organized and dynamic in nature28,66. The cells closest to the tips are the true progenitors, while those localizing a bit further away, in the armpits of T-bud, are pre-committed to differentiation18,28,67. Thus, the position of the NP cells within the cap clearly affects their fate, while their dynamic behavior, as evidenced by their capability to migrate in and out of the cap compartment, suggests flexibility in fate decisions at the individual cell level. Recent comparative studies have established that the overarching regulatory themes of renal progenitors are well conserved between mouse and us humans18,46,70,71,72 supporting the use of organ culture in the future as well.

Nephron differentiation occurs in the close vicinity of UB epithelium and starts when signals from the UB, and possibly the surrounding stroma, induce the committed nephron progenitors to form a cell condensation known as the pretubular aggregate (Figure 1)13,47. This cellular cluster then undergoes a mesenchymal-to-epithelial transition to form the renal vesicle, an axially polarized structure with fates of proximal and distal nephron segments established. This means that the cell's position in the vesicle dictates whether it will become part of the renal corpuscle containing podocytes, glomerular tuft, and mesangial cells, or incorporate into the more distal segments of proximal convoluted tubule, descending loop of Henle, ascending loop of Henle, macula densa, and distal convoluted tubule. The eventual identity is determined by the intricate signals emanating from the UB, stroma, and the vesicle itself, and includes, e.g., NOTCH, RHO, and WNT pathways19,21,73,74. The fate determination in the renal vesicle represents a classic example of inductive tissue interactions taking place in an intact organ, which cannot be studied without 3D structural organization.

From the renal vesicle, the nephrogenesis progresses through well-described morphological steps to form comma- and S-shaped bodies, where the regional identities are further strengthened and additional cell types, like endothelia, invade into developing glomeruli48,75. Finally, at the cessation of organogenesis, the undifferentiated progenitor pool is used up to provide the renal filtration capacity for the life18,67. The anatomical and morphological steps of nephrogenesis, including innervation, are well represented in the in vitro nephrogenesis76,77. However, the vascularization of the kidney does not occur in the classic kidney culture systems. This aspect can be studied with certain limitations by kidney grafting onto avian chorioallantontoic membrane78,79.

In addition to whole organ culture, there are methods to generate kidney-resembling organoids from pluripotent cells. Different protocols with specific emphasis on creating nephrons80,81,82,83, ureteric bud84,85,86, and organoids with higher-ordered structures87, as well as tubuloids, the 3D cultures of nephron segment and collecting duct epithelium88,89, exist. These are valuable tools for studying renal morphogenesis in, e.g., humans, where a limited amount of fetal material is available. However, organoid methods cannot recapitulate the intact organogenesis similarly as in vitro organ culture. In this article, we provide detailed instructions for establishing Trowell-type kidney cultures and primary epithelial cell colonies from the UB. These protocols have primarily been used for mouse embryonic kidneys, but also work for developing rat kidneys. We demonstrate that Trowell-type culture is also effective for the embryonic kidneys of spiny mouse (Acomys dimidiatus) - a novel rodent model with exceptional regeneration capacity90,91,92,93,94. The advantage of embryonic kidney culture is its ease of setup and cost-efficient use.

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Protocol

All experiments were performed in accordance with relevant national and European Union guidelines and regulations. The Committee for animal experiments of the University of Helsinki approved the use of rodents for these experiments under the licenses KEK22-016 (valid: 1.7.2022-30.6.2025) and KEK25-014 (valid: 1.7.2025-30.6.2028).

NOTE: Make sure to dispose of all sharps and chemicals according to institutional guidelines and good laboratory practices. Kidney cultures can be established with early kidney rudiments originating from rodent embryos. The typical stages for setting up the kidney culture are E11.5-E13.5. The earlier the organ is, the harder it is to recognize it. Also, very early kidney rudiments (E10-11) grow more poorly in vitro, as they often fail to initiate the UB branching morphogenesis. On the other hand, the complexity of the UB epithelial tree increases upon kidney growth, and later it becomes somewhat more challenging to establish primary UB cell cultures, since this requires both visual recognition and careful separation of the tips from the surrounding MM. Ultimately, the scientific question being addressed will determine the appropriate developmental stage of the kidney to be used in the experiment.

1. Embryonic kidney culture

  1. Collection of embryonic kidneys
    NOTE: Mouse and rat females exhibit a specific reproductive biology feature where they form vaginal clogging, known as a plug, after conception. The day of vaginal plug is designated as 0.5 days post conception, also known as embryonic day 0.5 (E0.5)95. Following this embryonic staging, the kidney cultures are typically established at E11-12.5 by euthanizing the pregnant mouse with gradual filling of the chamber with CO2 followed by cervical dislocation.
    1. Spray the skin of the abdomen with 70% ethanol (Figure 2A). Open abdominal skin and muscle layers, localize the uterus containing embryos (Mus E11.5-E12,5, Acomys E18-E19) (Figure 2B), and cut it out. Place the uterus with embryos in a 100 mm petri dish containing Dulbecco's PBS.
      NOTE: As safety notes, the CO2-chamber should be placed either in the hood or used in a room with proper ventilation to avoid any harm to users. Also, ethanol is easily flammable and should not be used near a potential source of spark or fire. As a general note, Dulbecco's PBS is interchangeable with CO2-independent medium, where embryos and tissues are well-protected for short- and long-term storage96. Also, remember to always use clean and sterile dissection tools to avoid any source of contamination.
    2. Hold the uterus with forceps and simultaneously open its muscle layers by incising with scissors.
    3. Take out the embryos and rip them free from the extraembryonic tissues and placenta by gently pulling with thin forceps (Figure 2C). To best preserve the tissues, keep all other embryos except the one working with on ice during the dissection activity.
    4. By using needles as surgical knives, cut the embryo into two halves diagonally just below the liver (Figure 2D). Spare more tissue from the dorsal than the ventral side. Remove hind limbs, any possible residual pieces of liver, and intestine to get a closer look at the developing kidneys. To help orientation, at E11.5-E12.5 developmental stages, kidneys are located bilaterally on both sides of the dorsal aorta and spinal cord, ventral to the neural tube and somite, at the level of the hind limbs.
    5. Next, dissect the kidneys out of the body cavity. First, remove the dorsal skin and the spinal cord. Then, turn the embryo to lie on its back and gently remove lateral body walls by sliding the needles to the lateral body cavities, simultaneously carefully cutting off these extra tissues. The kidneys are now located as one block in the thin strip of tissue, where the Wolffian duct is on one side and the MM on the opposite side (Figure 2E).
    6. Trim off extra tissues from both the posterior and caudal ends of the stripe, and finally separate the two kidneys from each other by slicing in between them from the Wolffian duct side. Now the two kidneys are freed from the embryo and each other (Figure 2F) and can wait in Dulbecco's PBS for placement on the culture dish (1.2.2).
      NOTE: When studying genetically modified kidneys, process kidneys always pairwise to keep track of their origin and enable their genotyping. This can be done, for example, by placing the two kidneys in one of the 24-well plate wells.
  2. Setting up Trowell-type kidney culture
    1. Prepare culture dish(es). Place the Transwell insert filter of 0.1 µm pore size in the well and add 1.5 mL of culture medium (DMEM/F12 supplemented with 10% FBS, 1x penicillin-streptomycin (100 U/mL penicillin, 100 ug/mL streptomycin), and 1x L-glutamine) to the bottom of the well (Figure 3A). Air bubbles may disturb nutrient and reagent adsorption from the medium. Make sure there are no air bubbles that would be in touch with the filter. To facilitate this, add the culture medium to the Transwell plates by pipetting it through the small lateral holes in between the insert and dish wall.
      NOTE: Commercial Transwell inserts can be replaced with metal grids as a support for pieces of porous filter membrane (pore size 0.1 µm). With grid-supported filters, the culture system can be established in any type of cell culture dish. Note that the porous filter membranes must be pre-rinsed with distilled H2O and stored in 70% EtOH before use. For establishing the culture system, rinse the filters well with distilled H2O or Dulbecco's PBS and place either the entire filter or smaller pieces cut from the filter on top of the grid. Use media just enough so that it is in contact with but not covering the filter. This will ensure proper attachment of the tissue to avoid its floating off.
    2. Use the capillary or a very small pipette (0.5 -10 µL) to place a single kidney at a time on the filter (Figure 3B). Avoid extra liquid when placing the kidney, and remove any visible liquid surrounding the kidney by gently sucking with a capillary. Avoid punching the tissue.
      NOTE: It is extremely important to place the kidneys on a filter in a specific order when establishing kidney cultures originating from the GM mouse lines, since the information on the genotype at the time of culture set-up is lacking. The numbered order of kidneys on the filter allows their recognition during and after the culture. With freely rotating Transwell inserts, use markings on top of the lid but also on the sides of the dish to avoid losing the orientation if the Transwell insert rotates.
    3. Place the culture dish(es) containing the kidney explants into the incubator with +37 °C temperature and 5% CO2 for culture of desired time (Figure 3B).

2. Establishment of primary ureteric bud epithelial cultures

  1. Isolation of ureteric buds and setting up the cell culture
    1. Collect kidneys as described in step 1.1.
    2. Incubate kidneys in 4 mg/mL collagenase in D-MEM/F12 supplemented with 10% stripped fetal calf serum for 15 min at + 37 °C. To spare the enzyme, pipette a small amount (1-2 mL) of collagenase on a dry, clean 30 mm dish so that it forms a nice drop and preheat at 37 ˚C. Add and remove the kidneys with the help of a glass capillary for treatment. When using the collagenase treatment on 24-well plates, it is safer to add and remove the solution rather than move around the kidneys, which become sticky upon this treatment.
    3. Stop collagenase function by adding 25 U/mL of DNase in DMEM/F12 + 10% FBS. Let the kidneys rest for 10-15 min on ice.
    4. Uncouple the UB mechanically free from the MM. For this, use needles (e.g., 26 G) to gently scrape the surrounding mesenchyme away. For the successful isolation of UBs, it is extremely important to understand the precise morphology of the early kidney, and especially where the UB is located within the MM. UB is more clearly visible in the earlier stage kidneys, where MM is not thick and covers it from multiple angles. The UB epithelium appears as a bud- or T-bud shaped hollow tube at E11-11.5, and the shape is repeated at later stages but in a more complicated form.
      1. Start isolation of UB by penetrating the mesenchyme with needles at the stalk region of the bud. Remove the MM by pushing it away, starting from the stalk and moving towards the tips. The stalk would be near the Wolffian duct at E11.5 kidneys and on the medullar side at E12.5 kidneys. Since the E12.5 UB tree is more complex in 3D, it may be more accessible and thus easier to reach from the tips rather than the stalk. It is possible to establish UB primary cell culture from the tips only. Notably, this will generate fewer cells and is therefore recommended only for E11.5 or older kidneys.
        NOTE: To allow comparisons, the number of UBs or the relative amount of UB epithelium used to set up the primary cell culture must be as close to the same as possible between different samples and/or treatments. This establishes comparable cultures for any type of downstream analysis.
    5. Add 0.5-0.75 mL of culture medium (DMEM/F12 supplemented with 10% FBS, 1x penicillin-streptomycin, and 1x L-glutamine) supplemented with GDNF (5 ng/mL), FGF2 (25 ng/mL), and HGF (50 ng/mL) to the fibronectin-coated culture plate in advance. Then, place isolated UBs in the centre of the fibronectin-coated well (Figure 4A). All buds should be close to each other to form a single monolayer culture.
      NOTE: Ready-made fibronectin-coated plates are commercially available. Such plates should be pre-heated at least 30 min prior to use. However, in-house coating with fibronectin is a cheaper option. The plates (or glass cover slips, which are ideal for imaging) are incubated with 10 mg/mL fibronectin for 1 h at room temperature / 30 min at 37 °C. To remove excess fibronectin, wash with 0.2% BSA 3x 15 min at 37 °C followed by culture medium additions for the plates to be ready for use.
    6. Place the plate in the cell culture incubator extra carefully so as not to move UBs that easily float around the well. Be extremely careful when moving the plates after the UBs have been placed in the wells. Adhering tissues/cells to the fibronectin takes a few hours, and thus any sudden movement may push UBs to float all around the plate. This may cause the UBs to spread to the edges of the plate, which is not optimal for the documentation of their growth and behavior during the experiment.
    7. Let the UBs settle and adhere. During 42-48 h culture, they will form a monolayer (Figure 4B). To get an idea of where the UBs are settling in the well, the cultures can be monitored once a day during the first 48 h. If the UBs are from different genetic backgrounds, it might be useful to document their delamination process by time-lapse imaging to estimate possible differences in adhesive properties or monolayer formation capacity.

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Results

The ex vivo kidney culture method can be used to address a wide array of research questions related to kidney development, and the outcome of each experiment depends on the experimental setup. However, with the use of appropriate control samples, it is important to ensure that the culture conditions themself do not affect the development. Ideally, the control samples ex vivo recapitulate the in vivo kidney development. The house mouse has been the standard model of kidney culture experiments. W...

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Discussion

The kidney is a vital organ with significant blood filtration and body fluid balancing functions. The basis for renal functional capacity is built during fetal life, when all nephrons for life are formed44,51,97. Understanding of both the kidney growth through branching morphogenesis and the nephron differentiation per se have extensively benefited from kidney culture methods, which have provided the fundamental insight...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

We thank Mrs. Agnés Viherä for her expert technical help in all our kidney and UB epithelial cell cultures. Animals were housed at the Laboratory Animal Centre of University of Helsinki. The work received funding from, Aamu Pediatric Cancer Foundation; HiLIFE and iCANDOC at University of Helsinki; Väre Children's Cancer Foundation, Liv och hälsä and Perklén Foundations.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
10X PBSLonza Biosciences512261X solution for the use
1ml syringesThermo Fisher Scientific10564431
23-25G disposable injection needlesThermo Fisher Scientific10482792
24-well platesGreiner662160
6-well CellStar plates MerckM8562
70% ethanol (EtOH), diluted in MilliQ-H2OAny commericial providerUsed for sterilizing surfaces and skin
Bovine serum albumin (BSA)Thermo FisherB14Final concentration of 0.2%
Collagenase Invitrogen17018029200mg/ml stock in PBS
DMEM/F12 Thermo Fisher31331028
DnaseFisher10649890
Drummond Wiretrol II Micro Dispenser, 10-20µlTranscat pipettesDRMC064
Dulbecco’s PBS (with Ca2+ and Mg2+)Thermo Fisher Scientific14040117Dulbecco’s PBS is interchangeable with CO2 independent medium (Gibco/ThermoFisher), where embryos and tissues are well-protected for short- and long-term storage. 
E11.5 - E12.5 mouse embryosAny commericial breeder / own stockhttps://janvier-labs.com/en/Mouse embryos are typically produced by in-house plugging. The day of vaginal plug appearance is E0.5. Commercial  female mice with timed pregrancies are also available.
E18-19 spiny mouse embryosN/AN/ACorresponding alternative for mouse embryos. 
Fetal bovine serum (FBS)GibcoA5256801
FibronectinNovus Biologicals1030-FN-05M1mg/ml stock
Glass or plastic dishes (100mm, 60mm and 30mm).Gibco150476, 174888, 153066
GlutaMAXGibco35050061
human recombinant Fibroblast Growth Factor (rFGF2)PeproTech100-18B
Human recombinant Glial Cell Line Derived Neurotrophic Factor (rGDNF)R&D212-GD-010/CF
Human recombinant Hepatocyte Growth Factor (rHGF)PeproTech100-39
penicillin-streptomycinGibco15140122
Small scissors and fine forceps Fine Science Tools14040-10, 11254-20Only use clean and sterile dissection tools to avoid any source of contamination.
StereomicroscopeLeica MZ7.5 or Olympus SZ x9https://www.leica-microsystems.com/
Transwell Permeable Supports, 24mm inserts Fisher10619141

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