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. Author manuscript; available in PMC: 2021 Mar 18.
Published in final edited form as: Methods Mol Biol. 2018;1801:207–223. doi: 10.1007/978-1-4939-7902-8_17

Analysis of Thyroid Tumorigenesis in Xenograft Mouse Model

Xuguang Zhu 1, Sheue-Yann Cheng 1
PMCID: PMC7971365  NIHMSID: NIHMS1674474  PMID: 29892827

Abstract

Analysis of thyroid tumorigenesis in xenograft mouse model is important to study human thyroid cancer. Recent studies have made big strides toward understanding the molecular mechanisms by which thyroid hormone nuclear receptors (TR) act to maintain normal cellular functions in growth, differentiation, and development. Despite growing interest, the role of TR in oncogenesis remains to be fully elucidated. Two TR genes give rise to three major TR isoforms: TRα1, TRβ1, and TRβ2. These TR subtypes express in a tissue- and development-dependent manner. Research has been directed at understanding the mechanisms by which TR could mediate aberrant cellular signaling that contributes to oncogenesis, at dissecting possible distinct roles of TR isoforms in oncogenesis, and at the differential susceptibility of target tissues to the oncogenic actions of TR. This chapter gives a brief overview of the current undersatanding of known molecular oncogenic actions of TR. Here, we describe analysis of thyroid tumorigenesis used in interrogating the in vivo oncogenic actions of TR.

Keywords: Thyroid hormone receptor, Oncogenesis, Thyroid tumorigenesis

1. Introduction

Early indications that TR could be involved in oncogenic actions came from association studies. A loss in the expression of the THRB gene because of the truncation/deletion of chromosome 3p where the THRB gene is located was reported in many malignancies including lung, melanoma, breast, head and neck, renal cell, uterine cervical, ovarian, and testicular tumors [1–6]. The THRA gene locus undergoes frequent loss of heterozygosity in sporadic breast cancer, and rearrangement of the THRA gene has also been reported in leukemia, breast, and stomach cancer [7–9]. Somatic mutations of TRs have been found in human hepatocellular carcinoma [10], breast cancer [11], and pituitary tumor [12, 13]. TR mutants identified in human hepatocellular carcinoma lose T3 binding activity and transcription capacity, and exhibit dominant negativity [10].

The association of the dominant negative action with oncogenic actions of TR was supported by the findings that TRα1 is the cellular counterpart of the retroviral v-ERBA involved in the neoplastic transformation leading to acute erythroleukemia and sarcomas [14, 15]. v-ERBA is a highly mutated chicken TRα1 that does not bind T3 and loses the ability to activate gene transcription. v-ERBA competes with TR for binding to thyroid hormone response elements (TREs) and interferes with the normal transcriptional activity of liganded-TR on several promoters [16, 17]. The discovery that male transgenic mice over-expressing v-ErbA develop hepatocellular carcinomas provided the direct evidence to show the v-ERBA oncoprotein can promote neoplasia in mammals through its dominant negative activity [18]. These studies suggested that mutated TRs are involved in tumorigenesis. Studies aiming to secure unequivocal evidence to demonstrate the oncogenic of mutant TR by cell-based studies and murine mouse models are described below.

That dominant negative TRβ mutants could function as an oncogene was demonstrated by studies using genetically engineered mice. A TRβ mutant isolated from a patient, known as PV, suffering from resistance to thyroid hormone was targeted to the Thrb gene locus in mice (ThrbPV mice; [19]). ThrbPV/PV mice spontaneously develop follicular thyroid cancer as they age [20]. Using ThrbPV/PV mice, a series of studies elucidated how TRβPV could function as an oncogene [21–23]. TRβPV propels thyroid carcinogenesis by activation of tumor promoters such as cyclin D1 [24], β-catenin [25], phosphatidylinositol 3-kinase (PI3K)/AKT [26–28], pituitary tumor transforming gene (PTTG) [29, 30], and SRC/FAK [31]. TRβPV could also promote thyroid cancer progression by repression of tumor suppressors such as the peroxisome proliferator-activated receptor γ (PPARγ) [32]. These altered signaling pathways during thyroid carcinogenesis of ThrbPV/PV mice are consistent with the changes reported for the carcinogenesis in human thyroid. Molecular mechanisms by which TRβPV acts to aberrantly affect these critical cellular regulators have been elucidated. TRβPV acts at the transcriptional level via dominant negative action to suppress the tumor suppressor PPARγ [33]. TRβPV can also act via direct protein-protein interaction to affect key signaling molecules. TRβPV, by complexing with the p85 subunit of PI3K, constitutively stimulates PI3K’s activity to activate its immediate downstream AKT signaling [26, 27]. When complexing with PTTG, TRβPV stabilizes PTTG, resulting in a high accumulation of PTTG in tumor cells. Accumulated PTTG aberrantly delays mitotic progression [29]. In addition, TRβPV, by physically complexing with β-catenin, prevents proteasome degradation of β-catenin, resulting in constitutive activation of β-catenin signaling [34]. Gelsolin, an actin binding protein, is involved in controlling cell morphology, motility, growth, and apoptosis. Studies have indicated that physical association of TRβPV with gelsolin reduces its binding to actin, leading to disarrayed cytoskeletal architectures and increased cell motility, thus contributing to metastasis [35]. These findings reveal diverse molecular mechanisms by which TRβPV acts as an oncogene to promote thyroid carcinogenesis.

In addition to thyroid tumors, ThrbPV/PV mice also spontaneously develop thyroid stimulating hormone-secreting tumors (TSH-omas) [36]. The TRβPV induces over-expression of cyclin D1 to propel cell cycle progression by the activation of the cyclin D1-cyclin-dependent kinase-retinoblastoma protein-E2F pathway. The T3-bound wild-type TRβ represses cyclin D1 expression via tethering to the cyclin D1 promoter in binding to the c-AMP response element binding protein. However, this repression effect is lost in mutant TRβPV which cannot bind T3. The loss of binding thus results in constitutive activation of cyclin D1 to propel growth of the pituitary in ThrbPV/PV mice [36]. These results indicate that the oncogenic action of TRβPV is acting through indirect interaction with another transcriptional factor on the chromatin for transcription activation. At present, it is not clear whether this mechanism by which TRβPV mediates its oncogenic actions is unique in the pituitary or also operates in other target sites.

While correlative data suggested that TRβ mutations could increase the risk of mammary tumor development, unequivocal evidence was still needed. To this end, ThrbPV/PV mice were crossed with Pten+/− mice to explore how TRβPV could affect the susceptibility of Pten+/− mice in the development of mammary tumors [37]. The presence of two ThrbPV alleles markedly augments the risk of mammary hyperplasia in Pten+/− mice. TRβPV increases the activity of signal transducer and activator of transcription (STAT5) to increase cell proliferation and the expression of the STAT5 target gene encoding β-casein in the mammary gland. Cell-based studies showed that in cells expressing TRβ, T3 suppresses STAT5-mediated transcription activity and downstream target gene expression. This T3-mediated suppression is lost in cells expressing TRβPV, resulting in aberrant activation of STAT5 signaling to stimulate mammary growth [37]. Collectively, these studies showed that TRβPV can act as an oncogene in several target tissues and via diverse mechanisms to promote carcinogenesis depending on the cellular context.

Oncogenic actions of TRβ mutants have also been elucidated by using murine xenograft models. Studies described above clearly demonstrated that a TRβ mutant (TRβPV) functions as an oncogene in several target tissues of ThrbPV/PV mice. A fundamental question was whether the oncogenic activity of mutated TRβ is uniquely dependent on the PV mutated sequence (C-terminal 14 amino acid frameshift mutation; [38]). Studies were carried out using the murine xenograft model approach to address this question [39]. Murine xenograft models are used to investigate the factors involved in malignant transformation, invasion, and metastasis, as well as to examine response to therapy. In this type of model, tumor cells expressing the oncogene are transplanted into immunocompromised mice, either under the skin or into the organ type in which the tumor originated.

Using the xenograft mouse model approach, investigators assessed the region of TRβ mutants imparting oncogenic actions in four C-terminal frame-shift mutants—TRβPV, Mkar, Mdbs, and AM. The C-terminal helixes 11 and 12 are critically involved in the structural changes of the ligand binding domain upon binding of T3 [40]. The frame-shift mutated sequence of TRβPV is located in helix 12. The Mkar mutation has a T insertion at nucleotide 1590–1591 that leads to a frameshift mutation in the C-terminal 28 amino acids encompassing helix 11 and 12 [41]. The Mdbs mutation has a C insertion at nucleotide 1643–1644 that leads to a frameshift mutation in the C-terminal 10 amino acids located in helix 12 [41]. AM is a mutant that was constructed to combine the part of the mutation from Mkar (amino acids 436–446) and revert the distal amino acid sequence back to that of wild-type TRβ1 (amino acids 447–461, located in helix 11; [41]). Remarkably, these C-terminal mutants induce similar growth of tumors in mouse xenograft models [39]. These four mutants similarly interact with the p85α regulatory subunit of PI3K to aberrantly activate PI3K-AKT-mTOR and PI3K-ERK-MMP signaling to increase cell proliferation and invasiveness. Further, they also activate the PI3K-STAT3-BIM pathway to decrease apoptosis [39]. These results argue against the idea that the oncogenic activity of TRβPV is uniquely dependent on the TRβPV mutated sequence. Rather, these four mutants could favor a C-terminal conformation that interacts with the C-terminal SH2 domain of p85α to initiate activation of PI3K to relay downstream signaling to promote tumorigenesis. These studies identified three other TRβ1 mutants as oncogenes in addition to TRβPV. Further, they uncovered an “onco-domain” in the mutated C-terminal region of TRβ1.

If mutated TRβ functions as an oncogene, that would support the earlier observations in which the loss in the expression of the THRB gene because of the truncation/deletion or epigenetic silencing is associated with many malignancies [1–6]. Mouse xenograft models have provided a valuable tool to address whether re-activation of the silenced THRB gene could attenuate thyroid carcinogenesis. A positive correlation between the extent of promoter hypermethylation of the THRB gene and the progression of differentiated thyroid cancer was found in patient tissue specimens and in several human thyroid cancer cell lines [42]. When human thyroid cancer cell lines in which the THRB gene was silenced by hypermethylation were treated with demethylation agents such as 5 -aza-2 -deoxycytidine (5-aza-CdR) and zebularine, the expression of the THRB gene was reactivated concurrently with inhibition of cancer cell proliferation, migration, and tumor growth in a xenograft model. Re-expression of the THRB gene inhibited proliferation and migration of thyroid cancer cells through suppressing the activation of β-catenin signaling pathway [42]. These studies demonstrated that TRβ acts as a tumor suppressor in thyroid cancer cells.

With the use of mouse xenograft models, how functional expression of the THRB gene could act to inhibit tumorigenesis was delineated in human thyroid cancer cells (FTC-133 and FTC-236) stably expressing of TRβ [43]. Expression of the THRB gene in FTC-133 cells, versus control FTC cells without TRβ, reduced cancer cell proliferation and impeded migration of tumor cells through inhibition of the AKT-mTOR-p70 S6K pathway. TRβ expression in FTC-133 and FTC-236 led to less tumor growth in xenograft models. Importantly, new vessel formation was significantly suppressed in tumors induced by FTC cells expressing TRβ versus control FTC cells without TRβ. The decrease in vessel formation was mediated by the downregulation of vascular endothelial growth factor in FTC cells expressing TRβ. These findings indicated that TRβ acts as a tumor suppressor through downregulation of the AKT-mTOR-p70 S6K pathway and decreases vasular endothelial growth factor expression in FTC cells [43]. These studies raise the possibility that TRβ could be considered a potential therapeutic target for thyroid cancer.

Mouse xenograft models were also used to evaluate whether TRβ could act as a tumor suppressor in breast cancer MCF-7 cells, as it does in thyroid cancer cells [44]. Parental MCF-7 cells express the estrogen receptor (ER) but not TRβ. The tumor suppression activity of TRβ was evidenced by the decreased tumor growth in MCF-7 cells stably expressing TRβ (MCF-7-TRβ). Cell-based studies indicated that the estrogen (E2)-dependent growth of MCF-7 cells was inhibited by the expression of TRβ in the presence of T3. In a xenograft mouse model, large tumors rapidly developed after inoculation of MCF-7-Neo cells in athymic mice. In contrast, markedly smaller tumors (98% smaller) were found when MCF-7-TRβ cells were inoculated in athymic nude mice, indicating that TRβ inhibits the E2-dependent tumor growth of MCF-7 cells. Further detailed molecular analysis showed that TRβ acted to activate apoptosis and decrease proliferation of tumor cells, resulting in inhibition of tumor growth. The TRβ-mediated inhibition of tumor growth was elucidated via down-regulation of the JAK-STAT-cyclin D pathways. This in vivo evidence shows that TRβ could act as a tumor suppressor in breast tumorigenesis and provides new insights into the role of TR in breast cancer.

The studies described here illustrate the usefulness of mouse xenograft models to understand tumor suppressor functions of TRβ and to provide unequivocal evidence that TRβ mutants function as an oncogene. Subheadings 2–4 describe the detailed methodology for using the mouse xenograft.

2. Materials

2.1. Making Stable Cell Lines

  1. FTC-133 and FTC-236 cell lines.

  2. Complete medium for FTC-133 and FTC-236 cells: Dulbecco’s modified Eagle’s medium/Ham’s F12 (1:1) medium supplemented with 10% fetal bovine serum, 10 μg/mL bovine insulin, 1 mIU/mL bovine thyrotropin, 100 units/mL penicillin, and 100 μg/mL streptomycin.

  3. Plasmids: pFH-IRESneo and pFH-IRESneo-TRβ1 plasmids containing the G418 resistant gene (the Neo gene).

  4. G418 200 mg/mL stock solution.

  5. 6-well plates, T25 flasks, and T75 flasks.

  6. Opti-MEM medium.

  7. Lipofectamine 2000 DNA Transfection Reagent.

  8. Incubator for cell culture.

  9. Culture Hood for cell culture.

2.2. Xenograft Tumor Assay

  1. 0.05% Trypsin-EDTA.

  2. Complete cell culture medium.

  3. Trypan blue solution.

  4. Corning Matrigel basement membrane matrix. Leave Matrigel on ice overnight 1 day before recollection of the cells.

  5. 1 mL syringes, Gauge 27, length ½ inch.

  6. Digital caliper.

  7. Sorvall RT6000B refrigerated centrifuge.

2.3. Western Blot Analysis

  1. The antibodies against phosphorylated (p-)retinoblastoma (Rb), p-AKT, total AKT, p-p70 S6K, total p70 S6K, p-eIF4B, total eIF4B, p-GSK3β, total GSK3β, p-4EBP1, total 4EBP1, and glyceraldehyde 3-phosphate dehydrogenase (GAPDH).

  2. Antibodies against cyclin B1, cyclin D1, cyclin E, total Rb, and VEGF.

  3. Tissue lysis buffer: 50 mM Tris–HCl pH 8.0, 150 mM NaCl, 1 mM EDTA, 1.0% NP-40; 0.1% Triton X-100, protease inhibitors.

  4. Laemmli 2× buffer/loading buffer: 4% SDS, 10% 2-mercaptoethanol, 20% glycerol, 0.004% bromophenol blue, 0.125 M Tris–HCl, pH 6.8.

  5. Running buffer: 25 mM Tris base, 192 mM glycine, 0.1% SDS.

  6. Transfer buffer: 31 mM Tris pH 8.3, 240 mM glycine, 20% methanol, 0.04% SDS.

  7. Tris-buffered saline with Tween-20 (TBST): 25 mM Tris, pH 7.6, 137 mM NaCl, 3 mM KCl, 0.1% Tween-20.

  8. Blocking buffer: Add 5% milk to TBST buffer.

  9. Misonix XL-2000 Sonicator.

  10. Beckman Coulter DU 640 spectrophotometer.

  11. Immobilon-P Membrane, PVDF, 0.45 μm.

  12. Western Lightning® Plus-ECL Kit.

  13. Electrophoresis Power Supply.

  14. Novex XCell SureLock Mini-Cell and XCell II Blot Module, Invitrogen.

  15. Sorvall Legend Micro 17 microcentrifuge.

2.4. Immunohistochemistry Analysis

  1. Xylene, Sigma.

  2. Ethanol (100%, 95%, 75%, and 50%).

  3. Hematoxylin.

  4. Wash buffer: 1× PBS.

  5. Primary antibody against CD31 (as an example, CD31 is an endothelial marker for detecting angiogenesis).

  6. 10× phosphate buffered saline (PBS): 137 mM NaCl; 2.7 mM KCl; 4.3 mM Na2 HPO4; 1.47 mM KH2PO4, pH 7.4.

  7. 0.05% citraconic anhydride, pH 7.4.

  8. 0.3% hydrogen peroxide: To prepare, add 2 mL 30% hydrogen peroxide to 198 mL deionized water.

  9. Blocking solution: 10% normal goat serum in 1×PBS.

  10. DAB reagent: Prepare according to the manufacturer’s recommendations (Peroxidase Substrate Kit, Vector Laboratories, Inc.).

  11. Glass slide staining jars.

  12. Coverslip: Fisherbrand cover glasses: squares, size: 25 mm.

3. Methods

3.1. Making Stable Cell Lines

Many different approaches are possible for establishing stable cell lines, depending on the type of cells of interest and the constructs to be inserted into the cell genome. The protocol described below is specific for establishing cell lines via the lipofectamine transfection approach. The procedure is used to establish a cell line expressing the wild-type thyroid hormone receptor.

  1. For a stable cell line to be created, plate cells in a 6-well tissue culture plate 24 h before transfection such that the cells reach 80% confluence at the time of transfection.

  2. Transfect the cells in two wells within a 6-well plate with 2 μg of total plasmid DNA per well. Dilute 5 μL lipofectamine in 250 μL Opt-MEM and 2 μg of plasmid in 250 μL Opt-MEM. Add diluted plasmid to diluted lipofectamine reagent and incubate for 15 min at room temperature. Add the DNA- lipid complex to the cells and incubate them for 5 h.

  3. Add complete medium and culture the cells for 24 h in the incubator.

  4. Add 3 mL of media containing 2 mg/mL G418. Change media every 2–3 days. The cells with plasmid integration will survive and form colonies by 7–14 days after transfection.

  5. After single colonies form, transfer them first to a 24-well plate and then to a T25 flask or a T75 flask for further expansion of cells.

3.2. Xenograft Tumor Assay

Xenograft mouse models have been widely used to determine the in vivo activity of anti-cancer therapeutics before clinical development. Immunodeficient mice, such as athymic (nude) or severe combined immune deficient mice, are commonly used since these mouse strains exhibit very high growth rates for xenografts. The most common models are xenografts of cell lines grown subcutaneously. These xenograft tumors in nude mice can be easily monitored and measured for tumor progression. The following section describes the procedures for establishing a subcutaneous xenograft mouse model in nude mice.

3.2.1. Preparation of Tumor Cells

  1. Use the 5 × 106 cells for injection.

  2. Grow cells in complete medium and exclude any contamination.

  3. Remove medium and wash cells with 1×PBS. In a T-75 flask, add 2 mL of 0.05% trypsin-EDTA and incubate at 37 °C for 1–5 min. Disperse cells and add complete medium. Collect cells in a 50 mL conical tube.

  4. Count cells using a hemocytometer. Mix cells 1:1 with trypan blue solution.

  5. Calculate the viable cell number by using the following formula: Average counts × 10,000 × dilution factor = cell numbers/mL medium.

  6. Centrifuge at 800 rpm for 3 min. Remove the supernatant and suspend cells to achieve a concentration of 5 × 106 cells/100 μL medium. Add an equal volume of Matrigel and mix well; place on ice for injection.

3.2.2. Preparation of Mice

Nude mice should be 4–6 weeks old.

3.2.3. Preparation of the Injection

  1. Sterilize the inoculation area of the mice with an ethanol prep pad. Use a 1 mL syringe with a needle (gauge 27, length ½ in.) for inoculation.

  2. Mix cells and then draw into a syringe without a needle. Attach a needle and remove trapped bubbles.

  3. Inject 200 μL cells (5 × 106) subcutaneously into the lower right flank of the mice.

  4. Start treatment when the tumors have reached an average volume of 100 mm3.

  5. Tumor diameters are measured with a digital caliper.

  6. Tumor volume in mm3 is calculated by the formula: Volume = (π/6) × (length) × (width) × (height).

3.3. Western Blot Analysis

Western blotting is used to visualize proteins that have been separated by gel electrophoresis based on protein molecular weights. The proteins, as well as the molecular protein markers indicating protein sizes, are separated in a gel and transferred from the gel to a PVDF (polyvinylidene fluoride) membrane. The membrane with transferred proteins can then be probed by primary antibodies specific against the target of interest and visualized using secondary antibodies and detection reagents.

3.3.1. Preparation of Lysate from Tissues

  1. Dissect the xenograft tumors from nude mice as quickly as possible to prevent degradation by proteases.

  2. Place the tumor tissues in Eppendorf tubes and immerse in liquid nitrogen to snap freeze. Keep tissue samples in a freezer at −80 °C for later use.

  3. For tissue homogenization, cut an approximately 10 mg piece of tissue on dry ice. Place the tumor tissues in Eppendorf tubes and add 100 μL of ice-cold lysis buffer rapidly to the tube, homogenize with a homogenizer, and sonicate for two rounds of 10 s on ice with a 30 s interval.

  4. Centrifuge for 5 min at 13,800 × g at 4 °C in a microcentrifuge. Remove the tubes from the centrifuge and place on ice. Transfer the supernatant to a fresh tube kept on ice and discard the tubes with pellet.

3.3.2. Sample Preparation

  1. Use 1 to 2 μL tissue lysate to perform a protein quantification assay. Determine the protein concentration for each cell lysate.

  2. Use 30 μg of total protein for the analysis. Calculate how much lysate to load and add an equal volume 2× Laemmli sample buffer.

  3. Boil each cell lysate in sample buffer at 95 °C for 5 min to reduce and denature samples.

3.3.3. Loading and Running the Gel

  1. Load denatured samples into the wells of the SDS-PAGE gel, along with molecular weight marker.

  2. Run the gel for 1–2 h at 120 V until bromophenol blue is close to the bottom of the gel.

3.3.4. Transferring the Protein from the Gel to the Membrane

Cut the PVDF membrane into a size matched to the size of the gel and activate PVDF with methanol for 5 s and rinse with transfer buffer. Prepare a “filter paper-gel-PVDF membernce-filter paper” stack” (Fig. 1). Set the current at 400 mA and run the transfer for 2–3 h.

Fig. 1.

Fig. 1

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Assembly for the transfer of proteins from SDS-Polyacrylamide gel to PVDP membrane. PVDP membrane and polyacrylamide gel are sandwiched between two filter paper, Sponges and two plate electrodes

3.3.5. Antibody Staining

  1. Block the membrane for 1 h in the blocking buffer at room temperature.

  2. Incubate the membrane with diluted primary antibody recommended by the manufacturer in blocking buffer for overnight incubation at 4 °C.

  3. Wash the membrane in three washes of TBST for 5 min each.

  4. Incubate the membrane with the recommended dilution of conjugated secondary antibody in blocking buffer at room temperature for 1 h.

  5. Wash the membrane in three washes of TBST for 5 min each.

  6. For signal development, follow the kit manufacturer’s recommendations. Remove excess reagent and cover the membrane in transparent plastic wrap.

  7. Acquire image using darkroom development techniques for chemiluminescence.

3.4. Immunohistochemistry Analysis

Immunohistochemistry analysis provides the most direct approach for identifying both the cellular and subcellular distribution of protein in situ.

3.4.1. Deparaffinization/Rehydration

  1. Bake paraffin slides at 60 °C oven for 3 h.

  2. Incubate slides in three washes of xylene for 5 min each.

  3. Incubate slides in two washes of 100% ethanol for 5 min each.

  4. Incubate slides in two washes of 95% ethanol for 5 min each.

  5. Incubate slides in one wash of 75% ethanol for 5 min.

  6. Incubate slides in one wash of 50% ethanol for 5 min.

  7. Wash sections twice in 1×PBS for 5 min each.

3.4.2. Antigen Unmasking

Incubate the slides in 0.05% citraconic anhydride at 98 °C for 60 min. Cool down on bench top for 30 min.

3.4.3. Staining

  1. Wash slides in 1×PBS two times for 5 min each.

  2. Incubate slides in 10% normal goat serum in 1xPBS at room temperature for 1 h.

  3. Wash slides in 1×PBS twice for 5 min each.

  4. Incubate the slides with 100–400 μL primary antibody diluted in 10% normal goat serum in 1×PBS. Incubate overnight at 4 °C.

  5. Remove antibody solution and wash slides in 1×PBS six times for 5 min each.

  6. Add 100–400 μL horseradish peroxidase conjugated secondary antibody diluted in 10% normal goat serum. Incubate for 1 h at room temperature.

  7. Remove secondary antibody solution and wash sections six times with 1×PBS for 5 min each.

  8. Add 100–400 μL DAB to each slide and monitor staining closely. As soon as the slides are stained, immerse them in 1×PBS two times for 5 min each.

  9. Counterstain slides in hematoxylin for 1 min.

  10. Wash slides in 1×PBS two times for 5 min each.

3.4.4. Dehydrate Sections

  1. Incubate slides in 95% ethanol two times for 1 min each.

  2. Incubate in 100% ethanol two times for 1 min each.

  3. Incubate in xylene two times for 2 min each.

3.4.5. Mount Coverslips

To protect the stained tissues, pipet 12 μL of mounting medium permount (Fisher Scientific, Catalog#SP15–100) to the center of the stained tissues. Carefully place the coverslip on the mounting media. This procedure is carefully carried out so that no air bubbles are trapped under the coverslip. After the mount medium is dry, the slides may be used for subsequent observations and images recording.

4. Notes

  1. Making stable cell lines
    1. Only use healthy cells
      Passage cells 2–3 times after thawing before using them in transfection experiments. Do not allow the cells to become overgrown. Only use cells with more than 90% viability.
    2. Prepare high-quality DNA for transfection.
      Prepare DNA using an endotoxin-free protocol. Determine the DNA quality by measuring the OD 260/280 ratio, which should be between 1.7 and 1.9.
    3. Plate cells 1 day earlier.
      Seed your cells at a density such that they will be 70–90% confluent at the time of transfection.
    4. Prepare lipid DNA complexes.
      For maximum performance, prepare complexing the lipid and DNA in Opti-MEM medium.
    5. Use a positive control such as GFP or LacZ reporter plasmid to assess transfection efficiency.
      The percent of cells transfected can be determined using fluorescence microscopy. Alternatively, LacZ expression can be used to determine transfection efficiency.
    6. Choice a selective marker to establish cell lines.
      Several selection markers are available for the establishment of cell lines. If feasible, choice puromycin for fast selection.
  2. Xenograft tumor assay
    1. Only use healthy cells for subcutaneous inoculation. Do not allow the cells to become overgrown. Use 1–5 × 106 cells with more than 90% viability for inoculation.
    2. The matrigel should be thawed overnight on ice in a 2 °C–6 °C refrigerator.
    3. The cells mixed with matrigel should be kept on ice before subcutaneous inoculation.
  3. Western blot analysis
    1. Select best primary antibody.
      Primary antibody is one of the keys for good experimental results. Search and find the best primary antibody for Western blot analysis.
    2. Keep up the protein transfer efficiency.
      Use protein marker to monitor the transfer efficiency.
    3. Make sure to equilibrate membranes and gels on transfer solution.
      For PVDF membranes, first immerse PVDF membrane in 100% methanol for a few seconds, then equilibrate with transfer buffer. During incubation with antibody, the membrane should not be allowed to be dry.
    4. Blocking solution
      Appropriate blocking agent is critical for Western blot. Blocking solutions work better when supplemented with a mild detergent like Tween-20. Nonfat dry milk or BSA fraction V at working concentrations ranging between 0.5% and 5% are often used for blocking agent.
    5. Optimize your incubation time
      In many cases, a few hour incubation should be enough to visualize the protein of interest, however, overnight incubation at 4 °C will allow more sufficient time for the antigen-antibody reaction.
  4. Immunohistochemistry analysis
    1. Sufficient deparaffinization
      Due to its hydrophobicity, paraffin is insoluble in antibody solution. An insufficient deparaffinization will lead to uneven immunohistochemistry staining or enhancing background.
    2. Perform heat-induced or protease-induced epitope retrieval
      Antigens can be masked as a result of the fixation process, which makes antibody impossible to access antigen. Antigen retrieval is needed to unmask antigens for the binding of antibody.
    3. Block endogenous peroxidases
      To avoid staining artifacts, it is important to block endogenous peroxidases prior to using horseradish peroxidase (HRP) antibody conjugates.
    4. Block nonspecific binding sites
      Blocking should be performed prior to incubation with the primary antibody to prevent nonspecific antibody binding.
    5. Incubate with primary antibody
      Check on the manufacturer’s datasheet that the antibody has been tested in the specific immunohistochemical method. During incubation, make sure that no bubbles are trapped inside hybridization solution. It is important to put the slides inside a moisturized chamber to prevent the dryup of solution during overnight incubation.
    6. Incubate with DAB or other substrate solution
      In an immunoenzymatic staining, a colored precipitate is formed due to the reaction of an enzyme with its substrate. Make sure that all of slides are stained with same batch of the reagents and same length of time so that it is feasible to compare the intensity of staining among slides.
    7. Counterstain
      Counterstaining is crucial for a IHC experiment as the counterstain provides background contrast.
    8. Mount coverslip
      Mounting protects the specimen from damage while adding contrast during microscopy. It is important that no air bubbles are trapped under coverslip.

5. Perspectives

The understanding of oncogenic actions of TRβ mutants has been facilitated by the creation of mouse models in which thyroid cancer, and TSHomas spontaneously develop. Extensive elucidation of altered signaling pathways in thyroid carcinogensis in the mouse models has indicated that TRβPV acts as an oncogene via multiple molecular mechanisms. TRβPV can function by interfering with the transcription activity of TRβ by abnormal repression in the expression of tumor promoters (e.g., PPARγ). TRβPV can also act at the transcription level independent of TR, via “gain-of- function.” Importantly, TRβPV can also act via extra-nuclear sites, for example by initiating the actions via direct protein-protein interaction with key cellular regulators such as PI3K. Identification of key signaling pathways and regulators propelling thyroid carcinogenesis provides new opportunities for potential molecular targets for diagnosis and treatments.

Moreover, by means of mouse xenograft models, oncogenic action of TRβPV can be studied in cultured cell lines. Using this approach, in addition to TRβPV, the oncogenic actions of several other C-terminal TRβ mutants were also demonstrated, indicating that the oncogenic actions of C-terminal TRβ mutants are not TRβPV-sequence dependent [39]. The mouse xenograft models were also used to elucidate the underlying mechanism by which wild-type TRβ functions as a tumor suppressor in cell lines derived from different tissues [43, 44]. In view of the ease and versatility of mouse xenograft models, one question that would be of interest to ascertain is whether the dominant negative TRα1 mutants could function as an oncogene. Earlier findings demonstrated that V-erbA is a mutated TRα1, which acts in neoplasia by blocking erythroid differentiation and by altering the growth properties of fibroblasts [45]. While the v-erbB locus alone is sufficient to induce erythroleukemia and sarcoma independent of the v-erbA gene, the v-erbA by itself is not capable of independently causing transformation in either erythroid cells or fibroblasts [45, 46]. It is likely that another oncogene such as v-erbB (a mutated version of epidermal growth factor receptor; EGFR) would be needed, as in AEV-induced erythroleukemia and sarcoma, to collaborate with dominant negative TRα1 mutants to bring out the transformed phenotypes. Addressing these challenges and others that may emerge subsequently will certainly lead to recognition and appreciation of the important roles of TR in cancer biology.

Acknowledgments

We regret any reference omissions due to length limitation. We wish to thank all colleagues and collaborators who have contributed to the work described in this review. The research described in this review by the authors and their colleagues at National Cancer Institute was supported by the Intramural Research Program of the Center for Cancer Research, National Cancer Institute, National Institutes of Health.

Reference

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