The Teratoma Assay: An In Vivo Assessment of Pluripotency

Protocol
Part of the Methods in Molecular Biology book series (MIMB, volume 767)

Abstract

A teratoma is a nonmalignant tumor comprised of a disorganized mixture of cells and small foci of tissue comprised of cells from all three of the embryonic germ-layers. By definition, a cell is pluripotent if it can differentiate into cells derived from all three of the embryonic germ-layers: ectoderm, mesoderm, and endoderm. In the teratoma assay, putative pluripotent stem cells (PSCs) are implanted into an immune-compromised mouse where they may proliferate and differentiate to form a teratoma. The PSCs grow at the implantation site supported by a complex mixture of factors from the local milieu, as well as circulating factors that are vital components of normal mammalian physiology. After a predetermined time of 6–12 weeks or when the tumor has reached sufficient size, it is removed and subjected to histopathological analysis. The teratoma may be further processed by immunocytochemistry and gene expression profiling. This chapter describes methods to generate teratomas through the implantation of putative PSC lines in the SCID mouse. Implantation at the following sites is described: (1) intramuscular, (2) subcutaneous, (3) under the testis capsule, and (4) under the kidney capsule.

Key words

embryonic stem cells pluripotent stem cells pluripotency assay teratoma testis ­capsule teratoma kidney capsule teratoma 

1 Introduction

While in vitro differentiation assays and in silico gene expression arrays are useful in assessing pluripotency, the gold standard remains the teratoma assay (1, 2). This in vivo assay provides a means to assess the developmental potential of human pluripotent stem cell (hPSC) lines at a level that cannot yet be achieved using in vitro and in silico assays. A pluripotent stem cell, by definition, is a cell that can differentiate into cells derived from all three embryonic germ-layers: ectoderm, mesoderm, and endoderm. When putative PSCs are transplanted in immune-­compromised mice, they are exposed to a complex mixture of growth and extracellular matrix factors that cannot, so far, be fully replicated in a culture dish. This mixture of factors promotes the growth and differentiation of the PSCs into teratomas; begin tumors that contain a complex mixture of cells and tissues derived from all three germ-layers (review (3)). The teratoma assay is part of the standard set of quality control and basic characterization assays used in hPSC laboratories. It is performed as part of routine culture evaluation, when new embryonic stem cell (ESC) or induced pluripotent stem cell (iPSC) lines are generated, and when PSCs are expanded and banked to make working stocks (4, 5, 6). Several engraftment sites have proven useful for teratoma production; however, the graft site, number of cells implanted, and the cell preparation has been shown to influence the type of somatic cells found in the teratoma, whether the teratoma is cystic or solid tumor, and the growth rate of the teratoma (7, 8, 9, 10, 11).

To date, the largest single study assessing the functional pluripotentiality of human ESCs via teratoma assay was performed by the International Stem Cell Initiative (ISCI) under which teratomas were generated and analyzed from 15 independent hESC lines (12). Investigators implanted cells from each of these hESC lines under the testis capsule of SCID mice and evaluated a total of 37 histological slides. Most of the hESC lines in the study ­produced teratomas. Ectodermal and mesodermal tissues predominated in the teratomas. Neural tissue was most often present as immature rosettes. Mesoderm included fibroblasts, capillaries, smooth muscle, striated muscle, cartilage, bone, and fat. Endodermal tissues included gland-like structures lined with columnar or cuboidal epithelium. Interestingly, three of the cell lines produced teratomas that contained foci of undifferentiated cells that had undergone malignant transformation into embryonal carcinomas, which when the cell lines were karyotyped were found to contain aneuploid cells. Excellent examples of histological sections of hESC-derived teratomas have been published (5).

This chapter describes the production of teratomas following implantation into four different sites in the SCID mouse: intramuscular injection in the lower flank, subcutaneous injection in the lower leg, implantation under the testis capsule, and implantation under the kidney capsule. Each of these sites is effective at generating teratomas, but with varying efficiency, and each requires a different level of surgical skill. In each case, PSCs are implanted into SCID mice, the mice are monitored for 6–12 weeks, and the tumor is harvested and analyzed for the appearance of cellular derivatives from all three germ-layers by a qualified clinical pathologist. Implantation under the testis or kidney capsule is major survival surgery and requires a high level of surgical expertise not required for intramuscular or subcutaneous injections. However, these surgical implantation methods have advantages in that they require fewer cells and use fewer animals than the injection procedures as they tend to be more efficient sites for engraftment. Recent reports, however, have suggested that resuspending the cells in Matrigel™ increases the teratoma efficiency after injection and may eliminate the need for surgery to achieve high rates of engraftment (10, 11).

Whichever site is chosen for implantation, PSC cultures should be undifferentiated and actively growing (seeChapter 8). Care should be taken to ensure the test sample is representative of the entire culture.

2 Materials

2.1 Injection: Intramuscular or Subcutaneous

  1. 1.

    5 SCID-BEIGE mice/cell line or culture to be assayed (see Note 1).

     
  2. 2.

    Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F12 (DMEM/F12).

     
  3. 3.

    Sterile 1-cc syringe with 23 g, ½-in. needle.

     
  4. 4.

    Measuring calipers.

     

2.2 Implantation Under the Testis Capsule

  1. 1.

    3–5 male-SCID-BEIGE mice/cell line to be tested.

     
  2. 2.

    Dulbecco’s Modified Eagle’s Medium/Nutrient Mixture F12 (DMEM/F12).

     
  3. 3.

    Glass capillary pipettes, pulled and fire-polished on one end for implanting cells under the capsule without rupturing the membrane.

     
  4. 4.

    Small Animal Clipper with #40 blade for shaving the mouse’s abdomen prior to surgery.

     
  5. 5.

    70% ethanol.

     
  6. 6.

    Betadine solution.

     
  7. 7.

    Sterile gauze pads.

     
  8. 8.

    Sterile surgical pack that includes: fine dissecting scissors, serrefine clamp, blunt forceps, watchmaker’s forceps #5, absorbable suture size 5-0 with an attached curved size 10 needle that is triangular and pointed, surgical stapler and 9-mm wound clips.

     
  9. 9.

    Anesthetic as directed by your veterinarian, institutional animal care and use committee, and biosafety committee (see Note 2).

     
  10. 10.

    Heating pad or slide warmer.

     
  11. 11.

    Clean cage for postoperative recovery.

     
  12. 12.

    Measuring calipers.

     

2.3 Implantation Under the Kidney Capsule

  1. 1.

    All items listed in Subheading 2.2 for surgical implantation under testis capsule.

     
  2. 2.

    26-mm diameter Chalazion forceps (desmarres) optional (see Note 3).

     

2.4 Teratoma Tumor Harvest and Analysis

  1. 1.

    Sterile surgical pack containing dissecting scissors and forceps.

     
  2. 2.

    Sterile scalpel or razor blade.

     
  3. 3.

    Dulbecco’s Phosphate-Buffered Saline (D-PBS).

     
  4. 4.

    Neutral buffered formalin 10%.

     
  5. 5.

    Liquid nitrogen for flash freezing tissue if RNA isolation is to be performed.

     

3 Methods

Regarding cell cultures: Ideally, a healthy hPSC culture in the log phase of growth is harvested (see Chapter 8). This can usually be achieved by harvesting the cells 1 or 2 days before they would routinely be subcultured.

Regarding the use of mice: Procedures that involve the use of live animals require institutional approval prior to initiating the experiment. Investigators are encouraged to work with their veterinarians and skilled animal care personnel who can provide them with surgical training and expert animal handling and care, utilizing best practices developed at their institution.

3.1 Teratoma Formation via Intramuscular or Subcutaneous Injection

Teratoma formation by injection of PSCs is the easiest method. No surgery is required and tumor growth can be monitored by visual observation and palpation. However, a large number of cells are required and only 25–50% of the mice develop tumors. Nevertheless, teratoma formation by injection has been used successfully to assess the pluripotency of hESCs and iPSCs (13, 14, 15). The efficiency of tumor formation may be improved by suspending the cells in extracellular matrix components such as Matrigel™ (10).
  1. 1.

    Collect undifferentiated PSCs: Harvest cells from 1 to 6 wells of a six-well dish. Lift the cells from the dish as you would when passaging using collagenase IV or dispase (see Chapter 8). Try not to carry along the feeder cells. Wash cells twice with DMEM/F12, by resuspending the cells in 5 ml of DMEM/F12 and spinning at 200  ×  g. After the first wash, count the cells while they are being spun down for the second time. Cells may not be single cells, estimate the number of cells/small clump. Resuspend the cells to a concentration of 1–2  ×  107 cells/ml in DMEM/F12.

     
  2. 2.
    Inject hPSCs: Inject 50 μl of cell suspension either subcutaneously on the lower hind leg, near the ankle, or 50–100 μl into the thigh muscle using a sterile 1-cc syringe and a 23-g, ½-in. needle (see Note 4). Repeat the injection into each of five mice.
    1. (a)

      Inject: 0.5–1  ×  106 cells/50 μl subcutaneous injection into the lower leg

       
    2. (b)

      Inject: 3–5  ×  106/50 μl cells into the thigh muscle

       
     
  3. 3.

    Observe the animals daily. Watch for changes in appearance and behavior. Monitor the injection site for tumor growth, for about 6–12 weeks or until the predetermined experimental endpoint (see Note 5).

     
  4. 4.

    Dissect the tumor. When the tumor is palpable and about 5 mm in size, or the predetermined endpoint for tumor growth is met, euthanize the mouse and surgically remove the tumor.

     

3.2 Teratoma Formation via Implantation Under the Testis Capsule

Teratoma formation via implantation under the testis capsule has been used to assess pluripotency of hESC lines (16, 17, 18) and was the method chosen by the International Stem Cell Initiative (ISCI) to comparatively assess the pluripotency of 15 independent hESC lines (12). This engraftment site has advantages over subcutaneous and intramuscular injection sites and implantation under the kidney capsule; it does not require a large number of cells, the testis is not a vital organ, and the teratoma growth can be monitored by visual observation and palpation. Transplantation of cells under the testis capsule is a fairly straightforward operation, with the surgical setup similar to that of vasectomy.

Surgery requires preapproval, specialized training, and planning in order to ensure that the location of the surgery and method of anesthesia is in keeping with the institutional rules and regulations.
  1. 1.

    Collect undifferentiated PSCs: Using aseptic technique, manually dissect hPSC colonies into clumps of 200–400 cells. 10–15 clumps will be implanted/testis. Carefully collect the bits of colonies in a sterile 1.5-ml microfuge tube containing 1 ml of DMEM/F12. If using dispase or collagenase to dissociate the colonies, collect between 10,000 and 100,000 cells and wash the cells twice to remove the enzyme (see Note 6).

     
  2. 2.

    Prepare an appropriate surgical location. This is an approved location where survival surgery can be performed aseptically. Assemble sterile surgical instruments and supplies, as well as postoperative materials, such as clean cage and heating pad or slide warmer to aid in postoperative recovery.

     
  3. 3.

    Anesthetize the mouse: The choice of anesthesia should be determined by consulting with your local veterinarian and biosafety committee. Ideally the chosen anesthetic will keep the animal anesthetized for 20–30 min and have minimal negative effects on the animal as well as personnel (see Note 2).

    After administering the anesthetic, monitor the animal for slowed breathing and perform a reflex check by gently squeezing the rear paw and monitoring response. When the mouse is under anesthesia, it will not withdraw its paw and its breathing will be slow and shallow.

     
  4. 4.

    Place the mouse on its back on the prepared surgical surface.

     
  5. 5.

    Shave the lower abdomen.

     
  6. 6.

    Swab shaven area with 70% ethanol or Betadine solution.

     
  7. 7.

    Using aseptic technique and sterile instruments make a small incision (1–2 cm) in the lower abdomen at the height of the knees. First make an incision in the skin and then a slightly smaller incision in the wall of the abdomen.

     
  8. 8.

    Gently squeeze the scrotum to push the testis up into the abdomen.

     
  9. 9.

    Find the fat pad attached to the testis; using blunt forceps, gently pull the fat pad to remove the testis from the abdomen.

     
  10. 10.

    Using a small serrefine clamp immobilize the testis by clamping the fat pad to expose and stabilize in an accessible position for transplantation.

     
  11. 11.

    Under a dissecting microscope, carefully lift the testis capsule (membrane surrounding the testis) with a fine forceps and puncture it with the tip of a sterile 26-g needle. Then using the pulled and polished glass micropipette, inject about 25–30 μl of cell suspension under the testis capsule. Place the cells toward the back of the testis without puncturing it. This will help the cells remain inside the capsule when the testis is placed back into the abdomen.

     
  12. 12.

    Carefully release the serrefine clamp and gently push the testis back into the abdomen using a blunt forceps. The testis will descend into the scrotum on its own.

     
  13. 13.

    Suture the abdomen wall with 1–2 stitches of absorbable suture.

     
  14. 14.

    Close the skin with wound clips.

     
  15. 15.

    Place the animal into a prewarmed clean cage.

     
  16. 16.

    Observe closely until the mouse recovers from anesthesia and apply analgesics as necessary and advised by your veterinarian.

     
  17. 17.

    Place cage in the animal room.

     
  18. 18.

    Observe the animal carefully daily. Check for changes in appearance and behavior.

     
  19. 19.

    Remove wound clips, using the clip remover tool, as soon as the incision has healed, 1–2 weeks following surgery.

     
  20. 20.

    Monitor the testis for tumor growth. Depending on the number of cells implanted, the tumor is likely to be present at 6 weeks and can be grown for additional 6 weeks.

     
  21. 21.

    When the tumor is palpable, about 5 mm in size, or at the predetermined experimental endpoint, euthanize the mouse and remove the tumor for analysis.

     

3.3 Teratoma Formation via Implantation Under the Kidney Capsule

Implantation of adult and embryonic tissues under the kidney capsule has been used for many years to study tissue rejection and to obtain teratocarcinomas from early mouse embryos. Implantation under the kidney capsule has been reported to give the highest efficiency of tumor formation (10), but it requires a great deal of surgical skill since the kidney is a vital organ. This protocol was adapted from the one found in Manipulating the Mouse Embryo, Second Edition (19).
  1. 1.

    Collect undifferentiated PSCs: Using aseptic technique, manually dissect hPSC colonies into clumps of 200–400 cells. 10–15 clumps will be implanted under the capsule. Carefully collect the bits of colonies in a sterile 1.5-ml microfuge tube containing 1 ml of DMEM/F12. If using dispase or collagenase to dissociate the colonies, collect between 10,000 and 100,000 cells and wash the cells twice to remove the enzyme (see Note 6).

     
  2. 2.

    Prepare an appropriate surgical location. This is an approved location where survival surgery can be performed aseptically. Assemble sterile surgical instruments and supplies, as well as postoperative materials, such as clean cage and heating pad or slide warmer to aid in postoperative recovery.

     
  3. 3.

    Anesthetize the mouse: The choice of anesthesia should be determined by consulting with your local veterinarian and biosafety committee. Ideally the chosen anesthetic will keep the animal anesthetized for 30–45 min (see Note 2). After administering the anesthetic, monitor the animal for slowed breathing and perform a reflex check by gently squeezing the rear paw and monitoring response. When the mouse is under anesthesia, it will not withdraw its paw and its breathing will be slow and shallow.

     
  4. 4.

    Shave the abdomen of the mouse.

     
  5. 5.

    Swab with Betadine or 70% ethanol.

     
  6. 6.

    Working to the right side of the midline, make a 1–2 cm ­incision in the skin and a slightly smaller incision in the ­abdomen wall.

     
  7. 7.

    Find the fat pad that is connected to the kidney and using a blunt forceps, gently pull the kidney through the opening by the fat pad.

     
  8. 8.

    Immobilize the kidney using a Desmarres chalazion forceps. Allow the surface to dry for a few minutes.

     
  9. 9.

    Use a watchmaker’s forceps to make a small hole in the capsule membrane.

     
  10. 10.

    Moisten the capsule with a small amount of sterile PBS and using moistened forceps make a pocket underneath the capsule.

     
  11. 11.

    Insert a capillary pipette containing the cells into the pocket and as far away from the tear as possible and deposit the cells into the capsule.

     
  12. 12.

    Release the kidney from the Demarres chalazion forceps and gently put it back into the body cavity using the blunt forceps.

     
  13. 13.

    Sew body wall with one or two stitches.

     
  14. 14.

    Close the skin with wound clips.

     
  15. 15.

    Place the animal in a clean, prewarmed cage for postoperative observation.

     
  16. 16.

    Observe the animal daily.

     
  17. 17.

    Remove wound clips after wound has healed, 7–10 days following surgery.

     
  18. 18.

    Euthanize the animal at the experimental endpoint, 6–12 weeks following implantation, and remove the tumor to be processed for analysis.

     

3.4 Teratoma Harvest and Analysis

Below is a brief description of three ways to analyze teratomas. Histopathology is the standard assay and should be carried out by a qualified pathologist. When histopathology, immunocytochemistry, and gene expression analysis are applied in combination to the analysis of the teratoma; however, the teratoma assay can also be used to enhance our understanding of development in addition to its utility as a key assay of pluripotency (2, 20).
  1. (a)

    Histopathology: The tumor should be collected in D-PBS, washed three times, cut into pieces no more than 5-mm thick and fixed in 10% neutral buffered formalin. The fixed tumor is embedded in paraffin and slides are made. The tumor is sectioned (5–8 μm), fixed to slides, stained with hematoxylin and eosin, and evaluated by a pathologist (reviewed in (5)). TeratomEye, an automated assay system has been developed to identify the three representative tissue types – muscle, gut, and neural epithelia (21).

     
  2. (b)

    Immunocytochemistry: Fixed tumor is embedded in paraffin and slides are made OR the tumor may be flash frozen in liquid nitrogen and then cryo-sectioned (2, 20). The slides are then processed for immunocytochemistry and labeled with lineage and cell-type-specific antibodies. Antibody staining is used to help identify early stage differentiated tissues that do not yet have identifiable morphology and can be useful in identifying the types of tissues arising from each germ layer, but it cannot take the place of careful histological analysis. Importantly, very few antibodies are specific for a single cell type.

     
  3. (c)

    Gene Expression Analysis: Tissues are snap-frozen in liquid nitrogen and stored in sterile cryogenic vials until processed for RNA or DNA isolation. Standard protocols for RT-PCR or genome-wide microarrays are applied to the teratomas for gene expression analysis and compared to undifferentiated PSCs (6).

     

4 Notes

  1. 1.

    Mice: SCID-BEIGE (C.B-Igh-1b/GbmsTac-Prkdcscid-LystbgN7)

    Severe combined immune deficient (SCID) mice are valuable xenotransplant models and have been used for many years to study immune rejection. While several strains of immunodeficient mice have been reported to support teratoma formation from hESCs including SCID, NOD-SCID, and nude mice, the SCID-BEIGE may be a superior recipient. The SCID-BEIGE is a double mutant created by breeding C.B-17 scid to the C57BL/6-bg strain. It carries both the scid mutation which causes the lack of B and T cells and the beige mutation which causes cytotoxic T cell and macrophage defects, as well as reduced natural killer cell activity.

     
  2. 2.

    Anesthesia: There are several choices for anesthesia. Some investigators prefer isoflurane and methoxyflurane, but these require specialized scavenging systems, since they are gaseous derivatives of ether. Others prefer Nembutal®, the brand name for injectable Phenobarbital sodium solution, which is under strict FDA control, yet others prefer zoletil-50/xylazine or ketamine-xylazine. All of these can be very effective when used appropriately. 2,2,2-tribromoethanol (Avertin) is another anesthetic, one that has been used for many years and if properly made and stored is a good and safe anesthetic. Recently, it has fallen out of favor because when it is not properly stored it can break down to form dibromoacetaldehyde and hydrobromic acid, both strong irritants and it has been shown to effect the efficiency of transgenic mouse production (22). If using Avertin, protect it from heat and discard the unused solution after 2 weeks. With so many anesthetics available and new data regarding safety being made available, it is best to consult with the veterinarian at your institution to determine which anesthetic is believed to be the best one for this use given the particular laboratory setup, skill level, and regulatory issues at your institution.

     
  3. 3.

    The Chalazion forceps is a specialized surgical instrument that may facilitate the transplant of cells under the kidney capsule. It is used to hold the kidney in place while the cells are deposited in the capsule. It is designed with a solid bottom and open ring top, and has a thumb-screw mechanism to clamp the top and bottom. It was originally designed for ophthalmic procedures.

     
  4. 4.

    Keep the cells on ice until just before injection, when they are loaded into the syringe without a needle. This will limit the damage caused to the cells by the pressure generated when drawing them through a small needle.

     
  5. 5.

    The endpoint for the experiment: the experimental endpoint can be a predetermined length of time, 6 weeks, or when the teratoma has reached a certain size, 5 mm. Whatever endpoint is chosen, the health status of animals is the primary consideration. Since the animals are observed on a daily basis, it will be easy to determine if the animal is having a health issue and should be sacrificed before the predetermined endpoint. The number of cells transplanted has been shown to have an impact on how quickly the tumor forms. One wants to give the tumor enough time to develop into readily identifiable cells and tissue foci, hitting the right balance between the number of cells implanted and the time the tumor is allowed to develop is key to a good assay.

     
  6. 6.

    Implantation under the testis or kidney capsule do not require PSC colonies to be disaggregated to single cells. Small bits of colonies are ideal for these engraftment methods. Since the cells are collected in clumps, without dissociation to single cells, one may take a small aliquot of the cell-clump suspension and dissociate to single cells with trypsin, allowing one to determine the approximate number of cells transplanted.

     

Notes

Acknowledgments

Author wishes to thank many colleagues and friends, especially Philip Schwartz, Jeanne Loring, Martin Pera, and Melissa Carpenter, for many thoughtful discussions on stem cell biology.

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Copyright information

© Springer Science+Business Media, LLC 2011

Authors and Affiliations

  1. 1.Center for Department of Applied Technology DevelopmentBeckman Research Institute, City of HopeDuarteUSA

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