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JoVE Journal Biology

Biology

Generation and Maintenance of Primate Induced Pluripotent Stem Cells Derived from Urine

Published: July 28, 2023 doi: 10.3791/64922
Automatic Translation
1, 1, 1, 1, 1,2,3
1Faculty of Biology, Ludwig Maximilian University of Munich, 2Institute for the Advanced Study of Human Biology, Kyoto University, 3Hakubi Center, Kyoto University

Summary

The present protocol describes a method to isolate, expand, and reprogram human and non-human primate urine-derived cells to induced pluripotent stem cells (iPSCs), as well as instructions for feeder-free maintenance of the newly generated iPSCs.

Abstract

Cross-species approaches studying primate pluripotent stem cells and their derivatives are crucial to better understand the molecular and cellular mechanisms of disease, development, and evolution. To make primate induced pluripotent stem cells (iPSCs) more accessible, this paper presents a non-invasive method to generate human and non-human primate iPSCs from urine-derived cells, and their maintenance using a feeder-free culturing method.

The urine can be sampled from a non-sterile environment (e.g., the cage of the animal) and treated with a broad-spectrum antibiotic cocktail during primary cell culture to reduce contamination efficiently. After propagation of the urine-derived cells, iPSCs are generated by a modified transduction method of a commercially available Sendai virus vector system. First iPSC colonies may already be visible after 5 days, and can be picked after 10 days at the earliest. Routine clump passaging with enzyme-free dissociation buffer supports pluripotency of the generated iPSCs for more than 50 passages.

Introduction

Genomic comparisons of human and non-human primates (NHPs) are crucial to understand our evolutionary history and the evolution of human-specific traits1. Additionally, these comparisons allow for the inference of function by identifying conserved DNA sequences2, e.g., to prioritize disease-associated variants3. Comparisons of molecular phenotypes such as gene expression levels are crucial to better interpret genomic comparisons and to discover, for example, cellular phenotypic differences. Furthermore, they have - similar to comparisons at the DNA level - the potential to infer functional relevance, and hence to better interpret medically relevant variation within humans4. The incorporation of comprehensive molecular phenotypic data into these comparative studies requires appropriate biological resources (i.e., orthologous cells across species). However, ethical and practical reasons make it difficult or impossible to access such comparable cells, especially during development. Induced pluripotent stem cells (iPSCs) allow for the generation of such inaccessible cell types in vitro5,6, are experimentally accessible, and have been used for primate comparisons6,7,8,9,10,11,12,13,14.

To generate iPSCs, one needs to acquire the primary cells to be reprogrammed. Cells isolated from urine have the advantage that they can be sampled non-invasively from primates, and that they can be readily reprogrammed, probably due to their stem cell-like molecular profiles15. The culture conditions to maintain primate iPSCs are as important as reprogramming; classically, the culture of human pluripotent stem cells required a non-defined, serum-based medium and co-culture of mouse embryonic fibroblasts - so-called feeder cells - that provide essential nutrients and a scaffold for embryonic stem cells (ESCs)16. Since the development of chemically defined and feeder-free culture systems17,18, there are now various options of commercially available iPSC culture media and matrices. However, most of these culture conditions have been optimized for human ESCs and iPSCs, and hence might work less well in NHP iPSC culture. In this video protocol, we provide instructions to generate and maintain human and NHP iPSCs derived from urinary cell culture.

Since the first report of iPSC generation by the forced expression of defined factors in fibroblasts in 2006, this method has been applied to many different cell types of various origins19,20,21,22,23,24,25,26,27,28,29,30,31,32. Among them, only urine-derived cells can be obtained in a completely non-invasive manner. Based on the previously described protocol by Zhou et al.33, one can isolate and expand cells from primate urine even from non-sterile samples, by supplementing broad-spectrum antibiotics15. Notably, urine-derived cells sampled by this protocol exhibit a high potential to produce iPSCs, within a shorter period of time (colonies become visible in 5-15 days) than the conventional reprogramming of fibroblasts (20-30 days, in our experience), and with a sufficiently high success rate. These urine-derived cells were classified as the mixed population of mesenchymal stem cell-like cells and bladder epithelial cells, causing the high reprogramming efficiency15.

In addition to the variation in primary cells, the reprogramming methods to generate iPSCs also vary according to the purpose of usage. Conventional reprogramming procedures for human somatic cells were carried out by the overexpression of reprogramming factors with retrovirus or lentivirus vectors, which allowed the integration of exogenous DNA in the genome5,34,35. To keep the generated iPSCs genomically intact, researchers have developed a wide variety of non-integration systems - excisable PiggyBac vector36,37, episomal vector38,39, non-integrating virus vectors such as Sendai virus40 and adenovirus41, mRNA transfection42, protein transfection43,44, and chemical compound treatment45. Due to the efficiency and ease in handling, the Sendai virus-based reprogramming vectors are used in this protocol. Infection of primary cells is performed in a 1 h suspension culture of cells and viruses at a multiplicity of infection (MOI) of 5 prior to plating. This modified step could increase the likelihood of contact between cell surfaces and viruses, compared to the conventional method in which the viruses are added directly to the adherent cell culture, and thus yield more iPSC colonies15.

Passaging of human and NHP pluripotent stem cells can be done by clump passaging and single-cell passaging. Ethylenediaminetetraacetic acid (EDTA) is a cost-efficient chelating agent that binds calcium and magnesium ions, and thus prevents the adherent activity of cadherin and integrin. EDTA is also used as a mild, selective dissociation reagent, as undifferentiated cells detach before differentiated cells due to their different adhesion molecules. Complete dissociation induces massive cell death of primate iPSCs via the Rho/Rho-associated coiled-coil containing protein kinase (Rho/Rock)-mediated myosin hyperactivation. Therefore, supplementing the culture medium with a Rho/Rock inhibitor is essential for experiments that require single cells in suspension46,47. In this protocol, we recommend clump passaging as the routine passaging method and recommend single-cell passaging only when it is necessary, e.g., when seeding of defined cell numbers is required, or during sub-cloning.

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Protocol

This experimental procedure was approved by the responsible ethic committee on human experimentation (20-122, Ethikkommission LMU München). All experiments were performed in accordance with relevant guidelines and regulations.
NOTE: Approval must be obtained from the appropriate ethical committee before starting experiments dealing with human and NHP samples. All experimental procedures must be performed in accordance with relevant guidelines and regulations. Each of the following steps should be performed using sterile technique in a biological safety cabinet. All buffer and media compositions can be found in Supplementary Table S1. Ensure that all media are warmed to room temperature (22 °C) before being added to the cells. Each centrifugation step should be performed at room temperature, unless mentioned otherwise.

1. Isolation of cells from urine samples

CAUTION: Ensure that human donors are free from human immunodeficiency virus (HIV), hepatitis B virus (HBV), and hepatitis C virus (HCV). For NHPs, make sure the possible donors/cells are free from specific pathogens-B Virus (BV), Simian Immunodeficiency Virus (SIV), Simian Betaretrovirus (SRV), and Simian T Cell Lymphotropic Virus (STLV).

  1. Prepare a gelatin-coated 12-well plate by adding 500 µL of 0.2% gelatin per well, and distribute the liquid by moving the plate. Place at 37 °C for at least 30 min before needed.
  2. Collect human urine samples in 50 mL conical tubes. For primates, collect urine from the floor of the animal facility with a syringe.
    NOTE: A volume of 5 mL of urine was proven to be sufficient for isolating at least one colony in 42% of the attempts. However, using a higher volume of ~50 mL of urine is recommended to increase the chance of isolating colonies. NHP urine should be sampled as fresh as possible, preferably immediately after urination. The storage of urine samples at 4 °C for 4 h had no negative effect on the success rate of the protocol, but longer storage times were not tested.
  3. Centrifuge the urine-containing tube at 400 × g for 10 min, and carefully aspirate the supernatant, leaving approximately 1 mL in the tube.
  4. Resuspend the pellet in the residual 1 mL of liquid. Pool the suspensions in one tube if multiple tubes of urine were collected.
  5. Wash the cells by adding 10 mL of urine wash buffer (see Supplementary Table S1) containing 2.5 µg/mL amphotericin to the tube, and carefully mix the suspension using a serological pipette.
  6. Centrifuge the tube at 200 × g for 10 min, and carefully aspirate the supernatant, leaving approximately <0.2 mL in the tube.
  7. Resuspend the cell pellet in 1 mL of primary urine medium (see Supplementary Table S1) containing 0.5 µg/mL amphotericin per 50 mL of initially processed urine (resuspend in 1 mL, even if less than 50 mL of urine was processed).
  8. Aspirate gelatin from the wells (prepared in step 1.1), and plate 1 mL of the suspension from step 1.7 into one well of a 12-well plate. Repeat for as many wells as desired, or for as many mililiters of suspension available.
    Optional: To avoid contamination originating from unsanitary sample collection, add 100 µg/mL antimicrobial reagent to the cells from here on, until the first passage.
  9. Place the plate in a 37 °C, 5% CO2 incubator.
  10. Add 1 mL of primary urine medium per well daily until day 5, without removing the existing medium.
  11. On day 5, aspirate 4 mL of medium from the plate, leaving approximately 1 mL of medium. Add 1 mL of REMC medium (see Supplementary Table S1) per well to get a 1:1 mixture with the new culture medium.
  12. Replace half of the medium with REMC medium every day until the first colonies appear (Figure 1A, B). Therefore, remove 1 mL of old medium, and add 1 mL of fresh REMC medium per well.

2. Expansion of urinary cells

NOTE: Urinary cell passaging should be conducted before the culture reaches 90% confluency.

  1. Prepare the desired amount of gelatin-coated 12-well plates, as stated in step 1.1.
  2. Aspirate the old medium, and wash the cells by adding 1 mL of Dulbecco's phosphate buffered saline (DPBS).
  3. Aspirate the DPBS, and add 300 µL of 0.5x dissociation enzyme diluted with DPBS. Incubate the plate at 37 °C for 5 min.
  4. Add 700 µL of REMC medium to stop the enzymatic reaction. Gently pipette the suspension using a P1000 pipette until the cells are dissociated into single cells.
  5. Transfer the cell suspension to a 15 mL tube, and centrifuge the tube at 200 × g for 5 min.
  6. Carefully aspirate the supernatant and resuspend the pellet in 1 mL of REMC medium.
  7. Count the cells using a cell counter (a hemocytometer or an automated cell counter).
  8. For expansion of the urinary cells, plate 1.5 × 104 to 3 × 104 cells in 1 mL of REMC medium into one 12-well plate coated with 0.2% gelatin.
  9. Perform subsequent medium changes every other day until the culture reaches 80%-90% confluency. Therefore, aspirate the old medium and add 1 mL of fresh REMC medium.

3. Generation of iPSCs by Sendai virus vector infection

NOTE: For the workflow of the reprogramming procedure, see Figure 2A. Urinary cells used for reprogramming should be as young as possible, but a remarkable loss of reprogramming efficiency is not observed before passage 4. The Sendai Virus Reprogramming Kit must be used in a BL-2 facility. Handle viruses under a biological safety cabinet with laminar flow, and always use appropriate safety equipment to prevent mucosal exposure.

  1. Prepare a basement membrane matrix-coated 12-well plate by adding 500 µL of basement membrane matrix per well, and distribute the liquid by moving the plate. Incubate the plate at 37 °C for at least 1 h, and replace the basement membrane matrix with 900 µL of REMC medium. Store the plate at 37 °C until use.
  2. Quickly thaw the components of the Sendai Reprogramming Kit in a 37 °C water bath. Mix the Sendai viruses (polycistronic KLF4-OCT3/4-SOX2, cMYC, and KLF4) with a MOI of 5, and add REMC medium up to 100 µL. Use equation (1):
    Equation 1
    NOTE: As virus titers differ between lots, always check the titer in the certificate of analysis that is provided by the manufacturer.
    Optional: Use green fluorescent protein (GFP) Sendai virus in addition as a positive control for the transduction efficiency. For this, prepare an additional 3.5 × 104 cells in a separate tube during step 3.3.
  3. For dissociation of the urinary cells, follow steps 2.2-2.4. Count the cells using the cell counter, and transfer 7 × 104 urinary cells to a 1.5 mL tube.
  4. Centrifuge the tube at 200 × g for 5 min, and carefully remove the supernatant without disrupting the cell pellet. Resuspend the pellet in 100 µL of the SeV mixture prepared in step 3.2. Incubate the tube for 1 h at 37 °C for suspension infection.
  5. Plate the suspension on the basement membrane matrix-coated 12-well plates that were prepared in step 3.1. Routinely, plate 1 × 104 and 2.5 × 104 cells per well in duplicates.
  6. Incubate the cells at 37 °C and 5% CO2. Replace the medium with 1 mL of fresh REMC medium 24 h post-transduction and on day 3.
  7. On day 5 after transduction, change the medium to PSC generation medium (see Supplementary Table S1), with subsequent medium changes every other day. Therefore, remove the old medium and add 1 mL of PSC generation medium per well.
    NOTE: It can take up to 15 days until the first colonies appear.
  8. Pick individual iPSC colonies when the size of the colony exceeds 1 mm. To do this, scrape and carefully collect a single colony with a p10 pipette under a microscope. Transfer the colony into a new well of a 12-well plate coated with a basement membrane matrix containing 750 µL of PSC culture medium.
    Optional: Rinsing the plate with DPBS and treating for 1 min with 0.5 mM EDTA prior to picking could support the robust culture of further steps. If the cells are to be cultured for longer to wait for later emerging colonies, do not perform this EDTA treatment step.
  9. Grow the cells at 37 °C and 5% CO2 with subsequent medium changes every other day, as stated in section 4 of the protocol. When the picked colony reaches a diameter of 2 mm, continue with the routine iPSC passaging, as explained in section 5 of the protocol.

4. Medium change

NOTE: The culture medium should be changed every other day until the colonies grow large enough for passaging.

  1. Aspirate the old medium and add 750 µL of the fresh medium per 12-well plate. To switch to a different type of medium, replace the medium at least 1 day after passaging.

5. Passaging

NOTE: The cells should be passaged when the iPSC colonies grow large enough (diameter > 2 mm), or the colonies are about to touch each other. Routinely, iPSCs can be split approximately every 5 days. Use clump-passaging (step 5.1) for routine maintenance, and single-cell passaging (step 5.2) for experiments where a defined number of cells is needed. In case the iPSCs differentiate a lot, colony picking (step 5.3) can help improve the purity of the cultures.

  1. Clump-passaging
    1. Prepare a basement membrane matrix-coated 12-well plate by adding 500 µL of basement membrane matrix per well, and distribute the liquid by moving the plate. Incubate the plate at 37 °C for at least 1 h. Replace the basement membrane matrix with 500 µL of PSC culture medium and store the plate at 37 °C until use.
    2. Aspirate the medium from the cultured cells, and wash the cells by carefully adding 500 µL of DPBS. Remove the DPBS and add 500 µL of 0.5 mM EDTA to the well.
    3. Incubate the plate at RT for 2-5 min, until the colonies start detaching. Carefully observe the cells under the microscope.
    4. When the edges of the colonies start to peel off and gaps between the cells become visible (Figure 3A), remove the EDTA and carefully add 500 µL of DPBS.
      NOTE: Always pipette against the side wall of the well and never directly onto the cells, so as not to detach the cells from the plate.
    5. Aspirate the DPBS and flush the well with 500 µL of PSC culture medium using a p1000 pipette. Pipette up and down 1x-5x to disperse the colonies into clumps of appropriate size (Figure 3A). Do not pipette too much.
      ​NOTE: If the iPSCs are accidentally pipetted too much, add 10 µM of Rock inhibitor Y-27632 to the medium. This can enhance survival, as iPSCs are not able to survive as single cells.
    6. Transfer 1/10-1/50 of the cell clump suspension to the new wells. The ratio depends on the confluency of the well before splitting, the desired density of the seeded cells, and iPSC clonal preference.
    7. Distribute the clumps evenly in the well by gently moving the plate back and forth several times. Incubate the plate for at least 30 min at 37 °C to let the clumps attach.
    8. Replace the medium with 750 µL of PSC culture medium if many floating dead cells are observed; otherwise, add 250 µL of PSC culture medium. Place the plate at 37 °C and 5% CO2 in an incubator.
      ​NOTE: Medium replacement after 30 min is critical, especially for unstable cell lines (e.g., NHPs).
    9. Change the medium every 2-3 days until the colonies grow large enough for passaging. For medium change, follow step 4 of the protocol.
  2. Single-cell passaging
    1. Prepare the basement membrane matrix-coated culture plate, as stated in step 5.1.1, with the addition of 10 µM Y-27632 to the PSC culture medium.
      Optional: Add 10 µM Y-27632 to the cells 1-3 h prior to passaging to enhance the survival of sensitive cell lines.
    2. Aspirate the medium and wash the cells by adding 500 µL of DPBS. Remove the DPBS and add 300 µL of detachment solution to the wells.
    3. Incubate the plate at 37 °C for 5-10 min. When sufficient detachment of the cells is observed under the microscope, add 700 µL of PSC culture medium or DPBS.
    4. Pipette up and down 5-10x using a p1000 pipette until the cells are dissociated into single cells. Do not pipette too much, in order to prevent cell damage.
    5. Transfer the cell suspension to a 15 mL tube containing at least 2 mL of DPBS to dilute the detachment solution.
    6. Centrifuge the tube at 200 × g for 5 min and aspirate the solution completely, without disrupting the cell pellet.
    7. Resuspend the pellet in 500 µL of PSC culture medium supplemented with 10 µM Y-27632.
    8. Count the cells and seed 5,000-7,000 cells per basement membrane matrix-coated 12-well plate, prepared in step 5.2.1.
      NOTE: If a different cell number is needed, change to a bigger or smaller well accordingly.
    9. Incubate the plate for at least 30 min at 37 °C and 5% CO2 to let the cells attach.
    10. Replace the medium with 750 µL of PSC culture medium + 10 µM Y-27632 if many dead cells are observed; otherwise, add 250 µL + 10 µM Y-27632.
      ​NOTE: This step is critical, especially for the unstable cell lines (e.g., NHPs).
    11. Place the plate at 37 °C and 5% CO2 in an incubator.
    12. Change the medium to PSC culture medium without Y-27632 1 to 2 days after splitting, to allow the cells to display the classic colony morphology again (Figure 3B).
    13. Change the medium every 2 days until the colonies grow large enough. For medium change, follow section 4 of the protocol.
  3. Passaging of iPSCs by colony picking
    1. Prepare basement membrane matrix-coated 12-wells as stated in step 5.1.1.
    2. Aspirate the medium and wash the cells by carefully adding 500 µL of DPBS. Remove the DPBS and add 500 µL of 0.5 mM EDTA to the well.
    3. Incubate the plate at RT for 1-3 min and observe the cells under the microscope, until detachment of the colony is visible on the borders.
    4. Remove the EDTA and carefully add 500 µL of DPBS. Aspirate the liquid before slowly adding 500 µL of PSC culture medium to the well, without detaching the cells.
    5. Use a p200 pipette to pick the desired colony under the microscope, without collecting the differentiated cells. To do this, gently scratch over the colony while taking up the medium containing cells.
    6. Transfer each picked colony into one basement membrane matrix-coated well, as prepared in step 5.3.1. Dissociate the cells into small clumps using a p1000 pipette, by pipetting the cells 2-5x.
    7. Incubate the plate for 30 min at 37 °C and 5% CO2, allowing the clumps to attach.
    8. Replace the medium with 750 µL of PSC culture medium if many floating dead cells are observed; otherwise, add 250 µL of PSC culture medium.
      ​NOTE: Medium replacement after 30 min is critical, especially for the unstable cell lines (e.g., NHPs).
    9. Place the plate at 37 °C and 5% CO2 in an incubator.
    10. Change the medium every 2-3 days until the colonies grow large enough for passaging. To do this, follow section 4 of the protocol.

6. Freezing of urinary cells and iPSCs for long-term storage

NOTE: Routinely, iPSCs are frozen as clumps in cell freezing medium without counting. Pipetting should be minimal, to avoid dissociation into single cells. For urinary cells, routinely, 1.5 × 104 to 3 × 104 cells are frozen per tube, allowing the user to thaw one tube directly in one well of a 12-well plate without the need of another counting step.

  1. Prepare 5 mL of DPBS in a 15 mL tube.
  2. For the freezing of urinary cells, follow steps 2.2-2.4 of the protocol. For the freezing of iPSCs, follow steps 5.1.2-5.1.5 from the clump passaging protocol.
  3. Transfer the suspension to the 15 mL tube prepared in step 6.1. For the freezing of urinary cells, count 10 µL of the cell suspension using a hemocytometer. Centrifuge the cells for 5 min at 200 × g, and aspirate the supernatant completely.
  4. Resuspend the cell pellet in 400 µL of cell freezing medium per tube, and distribute the cells to the desired amount of cryotubes.
  5. Transfer the cryotubes immediately to -80 °C. Transfer the frozen tubes to a -150 °C freezer or liquid nitrogen 1 day after freezing at -80 °C for long-term storage.

7. Thawing of urinary cells and iPSCs

  1. For the thawing of urinary cells, prepare the desired amount of gelatin-coated 12-wells, as stated in step 1.1 of the protocol. For iPSCs, prepare the basement membrane matrix-coated 12-well plates, as stated in step 5.1.1. In both cases, do not exchange the matrix with medium.
  2. Prepare a 15 mL tube containing 4 mL of DPBS, and store it at 37 °C.
  3. Place a frozen vial of cells quickly in a 37 °C water bath for thawing, until a piece of floating ice becomes visible.
    NOTE: Wipe the cryotube with ethanol before and after incubation in the water bath to avoid contaminations.
  4. Add 500 µL of REMC medium for urinary cells, or 500 µL of PSC culture medium for iPSCs to the ice-containing suspension, and transfer the suspension immediately to the pre-warmed 15 mL tube prepared in step 7.2.
  5. Centrifuge the tube at 200 × g for 5 min, and discard the supernatant completely.
  6. For urinary cells, resuspend the pellet in 1 mL of REMC medium. For iPSCs, carefully resuspend the pellet in 750 µL of PSC culture medium. Avoid pipetting too much, in order to keep the clumps intact.
    Optional: Supplementing the medium with 10 µM Y-27632 can support the survival of iPSCs after thawing.
  7. Aspirate the matrix from the 12-well plates prepared in step 7.1, and carefully transfer the cell suspension to the well.
  8. Place the plate overnight at 37 °C and 5% CO2 in an incubator.
  9. The next day, replace the medium with PSC culture medium, without Y-27632 for iPSCs and with REMC for urinary cells.
  10. Grow the cells at 37 °C and 5% CO2 in an incubator.
  11. Change the medium every 2-3 days until the cells grow large enough for passaging. For medium change, follow section 4 of the protocol.

8. Immunocytochemistry

NOTE: Immunostaining with antibodies targeting pluripotency-related markers such as NANOG, OCT3/4, SOX2, TRA-1-60 and EpCAM is one of the most widely used validations of newly generated iPSCs. Further information about the antibodies and dilutions can be found in the Table of Materials.

  1. Plate iPSCs 1-3 days prior to use in an appropriate number of 12-well plates. Aspirate the medium, wash the cells by adding 500 µL of DPBS, and remove the DPBS. Add 400 µL of 4% paraformaldehyde (PFA) per well, and fix the cells for 15 min at RT.
  2. Remove the 4% PFA, and wash the cells 3x with DPBS. Add 400 µL of blocking buffer per well, and incubate the plate for 30 min at RT.
  3. Aspirate the blocking buffer, and add the antibodies diluted in 400 µL of antibody dilution buffer (ADB) to each well. Incubate the plate at 4 °C overnight.
  4. Remove the ADB containing the primary antibodies, and wash cells 3x with DPBS.
  5. Aspirate the DPBS, and add 400 µL of secondary antibodies diluted in ADB per well. Incubate the plate for 1 h at RT in the dark.
  6. Remove the ADB, and wash cells 3x with DPBS. Add 1 µg/mL 4',6-diamidino-2-phenylindole (DAPI) diluted in DPBS per well, and incubate for 3 min at RT.
  7. Aspirate the DAPI solution, and wash the cell 3x with DPBS. Add 500 µL of DPBS for imaging.

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Representative Results

When isolating cells from human and NHP urine, different types of cells can be identified directly after isolation. Squamous cells, as well as various smaller round cells, get excreted with the urine; female urine contains far more squamous cells than male urine (Figure 1B - Day 0; Supplementary Figure S1). After 5 days of culture in primary urine medium, the first adherent proliferating cells can be seen (Figure 1A,B - Day 5). At this point, half of the medium is replaced with REMC proliferation medium every day, until the first colonies appear. On approximately day 13, the adherent cells grow into large colonies that can be passaged (Figure 1B - Day 13). These colonies can be formed out of two morphologically distinct cell types - one having a more epithelial-like phenotype with cells growing closely attached, and the other showing a more mesenchymal-like morphology with elongated shape and higher migrating ability (Figure 1C). After the first passage, urinary cells grow as a monolayer, and can be passaged when the culture reaches 80% confluency (Figure 1B - Day 17).

Figure 1
Figure 1: Isolation and cultivation of urinary cells. (A) Workflow for establishing urinary cells from human and non-human primate samples. (B) Phase-contrast images showing the growth and colony formation of urinary cells until the first passage (p1). (C) Two different cell types isolated from urine samples can be distinguished after 2 weeks of culture. Scale bars = 250 µm (B,C). Please click here to view a larger version of this figure.

To generate iPSCs from urinary cells, a transduction with Sendai viruses introducing the Yamanaka factors OCT3/4, SOX2, KLF4, and c-MYC is used. To increase the transduction efficiency, urinary cells are incubated with the virus for 1 h in suspension, before seeding onto basement membrane matrix-coated wells (Figure 2A). Some adherent cells can be seen in the wells 1 day after transduction (Figure 2B - Day 1). After 5-10 days, these cells start to form colonies, and early morphological changes can be observed, indicating reprogramming of the cells (Figure 2B - Day 14). Many of these colonies progressively show an ESC-like morphology, and can be picked in PSC culture medium when the diameter exceeds 1 mm. After picking, the cells form colonies displaying the typical ESC morphology within approximately 4 days (Figure 2B - Days 21 & 24). When the iPSC colonies grow big enough, the cells can be passaged for the first time using the clump passaging protocol (protocol step 5.1).

This reprogramming approach has been used to generate iPSCs from humans, Gorilla gorilla (gorilla), and Pongo abelii (orangutan)15. While the probability to obtain urinary cells is quite variable, and at least two- to threefold lower for chimpanzees than for humans15, reprogramming of the obtained urinary cells was mostly successful in our experience. In general, 0.19% of the infected urinary cells give rise to colonies with ESC-like morphology, resulting in at least one colony in 87% of the reprogramming attempts15. All generated iPSCs share the same properties and show the typical ESC-like colony morphology, characterized by tightly packed cells with clearly defined edges (Figure 2C). In addition, all iPSCs display a normal karyotype, and the absence of SeV reprogramming vectors was verified by PCR after a minimum of five passages, as recommended by the SeV reprogramming kit manufacturer (data not shown)15. Immunocytochemistry is used to test the expression of pluripotency-associated proteins in the nucleus, such as NANOG, OCT3/4, and SOX2. Surface markers such as TRA-1-60, EpCAM, and SSEA4 can also be used (Figure 2D); however, SSEA4 is also expressed in the urinary cells15. Moreover, flow cytometry analysis can be performed using these surface markers to provide quantitative information on the pluripotency status of the iPSCs (method described by Ohnuki et al.48). This approach is used to show that 95.3% of the analyzed orangutan iPSCs are positive for TRA-1-60 (Figure 2E). Furthermore, the differentiation capacity of NHP iPSCs can be proven via in vitro embryoid body (EB) differentiation (method described by Geuder et al.15). As shown in Figure 2F, gorilla iPSCs are able to differentiate into endoderm (ɑ-fetoprotein), mesoderm (ɑ-smooth muscle actin), and ectoderm (β-III tubulin) lineages.

Figure 2
Figure 2: Generation and characterization of urine derived iPSCs. (A) Reprogramming workflow of urine-derived cells. (B) Phase-contrast images showing the morphological changes of gorilla urinary cells during reprogramming until the establishment of iPSC colonies. Scale bars = 500 µm. (C) Morphology of established iPSC colonies from human, gorilla, and orangutan cell cultures. Scale bars = 500 µm. (D) Immunofluorescence staining of gorilla iPSCs with pluripotency-associated markers at passage 8: NANOG, OCT3/4, SSEA4, SOX2, EpCAM, and TRA-1-60. Scale bars = 100 µm. (E) Flow cytometry analysis of orangutan iPSCs stained with anti-TRA-1-60-PE. Unstained cells were used as control. (F) Immunofluorescence analysis of endoderm (AFP), mesoderm (ɑ-SMA), and ectoderm (β-III TUB) after embryoid body differentiation of gorilla iPSCs. Nuclei were counterstained with DAPI. Scale bars = 100 µm. Abbreviations: iPSCs = induced pluripotent stem cells; PE = phycoerythrin; AFP = α-fetoprotein; ɑ-SMA = alpha smooth muscle actin; β-III TUB = beta III Tubulin; DAPI = 4',6-diamidino-2-phenylindole. Please click here to view a larger version of this figure.

When the iPSC colonies reach a diameter of roughly 2 mm, or are about to touch each other, the cells should be split. We suggest using the clump passaging protocol for routine maintenance. In this protocol, 0.5 mM EDTA is used to detach the cells for 3-5 min, depending on the colony size. During EDTA incubation, the borders of the colonies start to peel off, and visible gaps appear between the cells (Figure 3A). The EDTA is to be removed when an appropriate detachment is observed under the microscope. Prolonged incubation with EDTA leads to a higher fraction of single cells that are not capable of surviving. Next, the cells should be dissociated to clumps of similar size, as shown in Figure 3A. The cells should not be dissociated into small clumps, as this reduces survival and can lead to more differentiating cells.

When an experiment requires the seeding of a defined number of cells, the single-cell passaging protocol should be used. This protocol includes supplementing the medium with a Rock inhibitor, which enables the single cells to survive. It is expected that attached single cells show a different morphology, compared to cells growing in a colony after clump seeding (Figure 3B - Day 0). After 1-2 days of culture, the single cells start to grow into colonies again, while preserving their morphology as well as their loose cell-cell contact, if a Rock inhibitor is added to the medium (Figure 3B - Day 2). When these small colonies are observed, the Rock inhibitor should be removed from the medium, allowing the cells to display the typical ESC-like morphology again (Figure 3B - Day 4).

To judge if an iPSC colony is of high quality, several aspects need to be taken into consideration. First, it is normal that healthy colonies show no or a small amount of spontaneous differentiation; however, an increased number of differentiated cells (>10%) is indicative of poor iPSC quality. Additional characteristics of low iPSC quality are a loss of border integrity and uniformity (Figure 3C: right), as well as loose cellular packaging with visible gaps between the cells (Figure 3D: right). In contrast, high-quality iPSCs are characterized by defined borders, tight cellular packaging, and prominent nucleoli (Figure 3C, D: left images).

Figure 3
Figure 3: Primate iPSC culture. (A) Phase-contrast images showing the detachment of iPSC colonies during EDTA incubation following the clump passaging protocol and the optimal clump size after dissociation. Scale bars = 250 µm. (B) Timeline of iPSC recovery after single-cell passaging. The Rock inhibitor Y-27632 was removed from day 2 onward. Scale bars = 250 µm. (C) Left: example of a high-quality human iPSC colony with defined borders. Right: example of a poor-quality colony showing less border integrity and differentiated cells. Scale bars = 500 µm. (D) Example images for a high-quality human iPSC colony (left) in comparison to a low-quality one with loosely packed cells (right). Scale bars = 100 µm. Abbreviation: iPSCs = induced pluripotent stem cells. Please click here to view a larger version of this figure.

Supplementary Figure S1: Overview of cell types found in human urine. (A) Cells isolated from female urine. (B) Cells isolated from male urine. Scale bar = 100 µm. Please click here to download this File.

Supplementary Table S1: Buffer and media compositions used in this study. Please click here to download this File.

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Discussion

iPSCs are valuable cell types as they allow the generation of otherwise inaccessible cell types in vitro. As the starting materials for reprogramming, for example, fibroblasts are not easily available from all primate species, this paper presents a protocol for the generation of iPSCs from urine-derived cells. These cells can be obtained in a non-invasive manner, even from non-sterile primate urine samples, by supplementing the culture medium with broad-spectrum antibiotics.

Several critical steps in the protocol are worth some additional discussion. First, when isolating cells from non-sterile urine, it is important to supplement the medium with a broad-spectrum antimicrobial reagent to reduce the risk of contamination. If the growth of microbiological contamination becomes apparent despite the use of antibiotics, it is recommended to discard the entire culture and disinfect the area; in addition, in our experience, continued culturing does not produce healthy growing urinary cells. Second, when starting to reprogram cells, preparing several dishes with different numbers of SeV-transduced cells is highly recommended, to avoid the risk of detachment of all cells by overconfluency and to increase the probability of iPSC acquisition. In general, plating a higher density of SeV-transduced cells is expected to yield more reprogrammed iPSC colonies, whereas culturing the reprogramming cells for a period of ~20 days often results in overconfluency, followed by detachment of the entire well. Third, at each step that requires dissociation of iPSCs, it is critical to avoid extensive pipetting, which leads to significant cell death. Especially when the clump passaging is performed, it is important to keep the clumps at an appropriate size (Figure 3A). Clumps that are too big may lead to quick differentiation, while clumps that are too small can decrease survival.

We observed that the probability of obtaining urinary cells is quite variable between samples and aliquots, but found no evidence that smaller volumes have a lower success rate in obtaining colonies. In general, we isolated an average of 7.6 colonies from 100 mL of human urine. Assuming this average rate and a Poisson distribution, obtaining at least one colony has a probability of ~98% for 50 mL and ~30% for 5 mL of starting material. In our case, it was possible to isolate at least one colony from 5 mL of human urine in 42% of the attempts15. Based on this, the isolation of colonies can still be worth trying, even if only small volumes of urine are available. In case the reprogramming of urinary cells fails, and no iPSC colonies can be observed, plating different numbers of transfected cells can result in reprogramming of the urine-derived cells. The emerging iPSC colonies can be also identified as iPSCs by live staining using surface markers (e.g., TRA-1-60 or alkaline phosphatase [AP]; data not shown). It is known that AP activity is upregulated in pluripotent stem cells, and it can be used to transiently stain stem cells during the culture process. Another possible pitfall of the protocol is the optimal culture of NHP iPSCs. Using this protocol, we confirmed that human and NHP iPSCs maintain their pluripotent state for more than 50 passages using a commercially available medium. However, in comparison to human iPSCs, NHP iPSCs are more prone to spontaneously differentiate in our experience, potentially due to the culture conditions that were optimized for human and not for NHP cultures. In addition, there is great variability among iPSC clones in survival rates after plating, growing speed, and tendency of differentiation. Therefore, one often needs to determine the optimal splitting ratio, passage timing, and clump size for each clone individually.

We have demonstrated that even a volume of 5 mL can be sufficient to isolate urinary cells from NHPs15. However, while we found no significant differences in the number of isolated urinary cells in humans, gorillas, and orangutans, chimpanzees had a lower ratio, as we could not isolate urinary cells from a total of 87 mL. Hence, for some species, it might be necessary to collect much more urine than for others. Collecting large volumes from small primates can be especially challenging, but as the isolation of urinary cells is fairly cost-efficient, it is practical to just try it out for particular species and available volumes of urine. Unfortunately, the urine samples cannot be frozen, which limits the radius over which samples can be collected. However, we showed that a storage time of 4 h at 4 °C had no influence on the number of colonies isolated15, and a longer storage time or improving storage conditions might well be possible. We have introduced several types of quality control tests for validation. However, even if an iPSC line satisfies these criteria, clonal heterogeneity is still substantial, which has been explained by genetic background49 and epigenetic status50,51,52,53,54. To apply primate iPSCs to future comparative analysis, one should consider the heterogeneity and use enough clones to average out the clonal variation, thus avoiding misinterpretation of the result.

Nevertheless, the use of urine as a source for primary cells has a major advantage over conventional methods using, for example, fibroblasts for reprogramming, as they can be obtained completely non-invasively. In addition, this protocol allows for the generation of iPSCs from urine-derived cells with a shorter period than conventional reprogramming. With this method, colonies become visible within 5-15 days, whereas we have found that colonies from primate fibroblasts (rhesus, baboon) tend to appear later, after 20-30 days. In addition, with the use of this suspension infection method, 0.19% of virus-transduced cells gave rise to colonies with pluripotent cell-like morphology. This resulted in at least one colony in 87% of the attempts15. In comparison, the conventional transduction method of SeV with attached cells and lipofection with episomal plasmids were shown to have a significantly lower reprogramming efficiency, with 0.09% and 0.009% of the cells forming a pluripotent colony, respectively15.

In summary, this method allows for the isolation of urinary cells non-invasively and the generation of feeder-free iPSCs from nearly any human donor and many NHPs. This increases the accessibility to precious cell types that can be differentiated from these iPSCs. Moreover, this method can be used to produce patient-specific iPSCs that may be used for disease modeling or future cell-based therapies. Finally, as iPSCs can be generated from only small volumes of urine, this method can be applied to many more different primate or even mammalian species. Thus, this method can be a powerful tool for cross-species comparison, that may allow for better understanding of the evolution of human-specific traits.

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Disclosures

The authors have no conflicts of interest to disclose.

Acknowledgments

This work was supported by DFG EN 1093/5-1 (project number 458247426). M.O. was supported by JSPS Overseas Research Fellowship. All figures were created with BioRender.com. Flow cytometry was performed with the help of the Core Facility Flow Cytometry at Biomedical Center Munich. We would like to thank Makoto Shida and Tomoyo Muto from ASHBi, Kyoto University, for support of videography.

Materials

Name Company Catalog Number Comments
Accumax™ cell detachment solution (Detachment solution) Sigma-Aldrich SCR006
Amphotericin B-Solution Merck A2941-100ML
Anti-Human TRA-1-60 Mouse Antibody  Stem Cell Technologies 60064 Dilution: 1/200
Anti-Human TRA-1-60 PE-conjugated Antibody  Miltenyi Biotec 130-122-965 Dilution: 1/50
Bambanker™ (Cell freezing medium) Nippon Genetics BB01
Bovine Serum Albumin (BSA) Sigma-Aldrich A3059-100G
Cell culture multiwell plate, 12-well CELLSTAR Greiner BIO-ONE 665180
Countess™ II automated cell counter Thermo Fisher Scientific AMQAX1000
CryoKing® 1.5 mL Tubes with 2D Barcode (Cryotubes) Sued-Laborbedarf 52 95-0213 Different types of Cryotubes can be used for freezing. The 2D barcode tubes have the advantage that the sample info can be stored in a database with unique tube information.
CytoTune™ EmGFP Sendai Fluorencence Reporter (GFP Sendai virus) Thermo Fisher Scientific A16519
CytoTune™-iPS 2.0 Sendai Reprogramming Kit (Sendai virus reprogramming kit) Thermo Fisher Scientific A16518
DAPI 4',6-Diamidine-2'-phenylindole dihydrochloride Sigma-Aldrich 10236276001
DMEM High Glucose TH.Geyer L0102
DMEM/F12 w L-glutamine Fisher Scientific 15373541
Donkey anti-Mouse IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor™ 488 Thermo Fisher Scientific A-21202 Dilution: 1/500
Donkey anti-Rabbit IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor™ 594 Thermo Fisher Scientific A-21207 Dilution: 1/500
DPBS w/o Calcium w/o Magnesium TH.Geyer L0615-500
EpCAM Recombinant Polyclonal Rabbit Antibody (22 HCLC) Thermo Fisher Scientific 710524 Dilution: 1/500
Ethylenediamine tetraacetic acid (EDTA) Carl Roth CN06.3
Falcon Tube 15 mL conical bottom Greiner BIO-ONE 188271-N
Falcon Tube 50 mL conical bottom Greiner BIO-ONE 227261
Fetal Bovine Serum, qualified, heat inactivated, Brazil (FBS) Thermo Fisher Scientific 10500064
FlowJo V10.8.2 FlowJo  663441
Gelatin from porcine skin Sigma-Aldrich G1890-1KG
Geltrex™ LDEV-Free, hESC-Qualified, Reduced Growth Factor Basement Membrane Matrix Thermo Fisher Scientific A1413301
GlutaMAX™ Supplement Thermo Fisher Scientific 35050038
Heracell™ 240i CO2 incubator Fisher Scientific 16416639
Heraeus HeraSafe safety cabinet Kendro 51017905
Human EGF, premium grade Miltenyi Biotec 130-097-749
ImageJ  Fiji Version 2.9.0
MEM Non-Essential Amino Acids Solution (100X) Thermo Fisher Scientific 11140035
Microcentrifugation tube PP, 1.5 mL Nerbe Plus 04-212-1000
Microscope Nikon eclipse TE2000-S Nikon TE2000-S
Mouse anti-alpha-Fetoprotein antibody R&D Systems MAB1368 Dilution: 1/100
Mouse anti-alpha-Smooth Muscle Actin antibody R&D Systems MAB1420 Dilution: 1/100
Mouse anti-beta-III Tubulin antibody R&D Systems MAB1195 Dilution: 1/100
mTeSR™ 1 STEMCELL Technolgies 85850
Nanog (D73G4) XP Rabbit mAb  Cell Signaling Technology 4903S Dilution: 1/400
Normocure™ (Antimicrobial Reagent) Invivogen ant-noc
Oct-4 Rabbit Antibody  Cell Signaling Technology 2750S Dilution: 1/400
Paraformaldehyde (PFA) Sigma-Aldrich 441244-1KG
Penicillin-Streptomycin (10.000 U/ml) (PS) Thermo Fisher Scientific 15140122 Penicillin-Streptomycin mix contains 100 U/mL Penicillin and 100 µg/mL Streptomycin.
Recombinant Human FGF-basic PeproTech 100-18B
Recombinant Human PDGF-AB PeproTech 100-00AB
Refrigerated benchtop centrifuge SIGMA  4-16KS
Renal Epithelial Cell Basal Medium ATCC PCS-400-030
Renal Epithelial Cell Growth Kit ATCC PCS-400-040
Sox2 (L1D6A2) Mouse mAb #4900 Cell Signaling Technology 4900S Dilution: 1/400
SSEA4 (MC813) Mouse mAb NEB 4755S Dilution: 1/500
StemFit® Basic02 Nippon Genetics 3821.00 The production of this medium was discontinued, use StemFit Basic04CT for human cell lines or StemFit Basic03 for non-human primates instead.
Triton X-100  Sigma-Aldrich T8787-50ML
TrypLE™ Select Enzyme (1x), no phenol red (Dissociation enzyme) Thermo Fisher Scientific 12563011
Waterbath Precision GP 05 Thermo Fisher Scientific TSGP05
Y-27632, Dihydrochloride Salt (Rock Inhibitor) Biozol BYT-ORB153635
Antibody dilution buffer For composition see the supplementary table S1
Blocking buffer For composition see the supplementary table S1
REMC medium For composition see the supplementary table S1
Primary urine medium For composition see the supplementary table S1
PSC culture medium For composition see the supplementary table S1
PSC generation medium For composition see the supplementary table S1
Urine wash buffer For composition see the supplementary table S1

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Tags

Primate Induced Pluripotent Stem Cells Urine-derived IPSCs Generation Protocol Maintenance Protocol Non-invasive Cell Source Centrifugation Urine Wash Buffer Amphotericin Gelatin-coated Plate Basement Membrane Matrix-coated Plate
Generation and Maintenance of Primate Induced Pluripotent Stem Cells Derived from Urine
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Cite this Article

Radmer, J., Geuder, J., Edenhofer,… More

Radmer, J., Geuder, J., Edenhofer, F. C., Enard, W., Ohnuki, M. Generation and Maintenance of Primate Induced Pluripotent Stem Cells Derived from Urine. J. Vis. Exp. (197), e64922, doi:10.3791/64922 (2023).

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