Analysis of Mouse Growth Plate Development

Abstract

To investigate skeletal development, pathophysiological mechanisms of cartilage and bone disease, and eventually assess innovative treatments, the mouse is a very important resource. During embryonic development, mesenchymal condensations are formed, and cells within these mesenchymal condensations either directly differentiate into osteoblasts and give origin to intramembranous bone, or differentiate into chondrocytes and form a cartilaginous anlage. The cartilaginous anlage or fetal growth plate is then replaced with bone. This process is also called endochondral bone development, and it is responsible for the generation of most of our skeleton. In this Review, we will discuss in detail the most common in vivo and in vitro techniques our laboratory is currently using for the analysis of the mouse fetal growth plate during development.

INTRODUCTION

To investigate skeletal development, pathophysiological mechanisms of cartilage and bone disease, and eventually assess innovative treatments, animal models are an essential resource. Mouse is the most widely used animal model to study skeletal development for several reasons. Mouse models share many developmental and physiological features with humans (Karsenty and Ferron, 2012, Mouse Genome Sequencing, et al., 2002, Nguyen and Xu, 2008, Piret and Thakker, 2011). In addition, its relatively small size coupled with its reproductive abilities makes it an ideal species to study development. Lastly, mice can be genetically manipulated. During the past decades, analysis of genetically modified mice has led to numerous and major breakthroughs in the understanding of the genetic, cellular and molecular mechanisms of cartilage and bone development and homeostasis.

Development of bone is a finely orchestrated process both spatially and temporally, requiring careful coordination of multiple steps, which are controlled by a variety of molecular mechanisms and signaling pathways. Bones can be formed mainly through two distinct processes. During embryonic development, mesenchymal condensations are formed, and cells within these mesenchymal condensations either directly differentiate into osteoblasts and give origin to intramembranous bone, which forms the flat bones of the skull, or differentiate into chondrocytes and generate a cartilaginous anlage. The cartilaginous anlage or fetal growth plate is then replaced with bone. This process is also called endochondral bone development, and it is responsible for the generation of most of our skeleton (Karsenty, 2003, Kronenberg, 2003, Michigami, 2013, Zelzer and Olsen, 2003). Within the cartilaginous template, chondrocytes proliferate and produce a matrix enriched in type II, IX and XI collagens and in specific proteoglycans, mainly aggrecan (Aigner and Stove, 2003, Heinegard, 2009). Chondrocytes eventually form a columnar structure before exiting the cell cycle; they next undergo hypertrophy, and they start producing a matrix enriched in type X collagen. Terminally differentiated hypertrophic chondrocytes either die or transdifferentiate into cells of the osteoblastic lineage. Hypertrophic chondrocytes also express markers such as matrix metalloproteinase 13 (MMP-13) and vascular endothelial growth factor (VEGF) (Maes, et al., 2012b, Zelzer, et al., 2004) that are critically important to promote degradation of the cartilaginous matrix and its invasion by blood vessels, and thus pave the way to the replacement of the cartilaginous anlage with bone. Histological description of the cartilage growth plate is shown in Figure 1.

Figure 1.

Cartilage growth plate histology of a newborn mouse after H&E staining (Bar=100μm).

In this Review, we will discuss in detail the most common in vivo and in vitro techniques our laboratory is currently using to analyze the fetal growth plate during development. Basic Protocol 1 describes how to generate paraffin-embedded tissue sections; Support Protocol 1 provides details on how to properly orient samples in paraffin blocks. Basic Protocol 2 illustrates how to generate fresh frozen sections. Alternate Protocol 2 describes the preparation of frozen sections from formalin-fixed specimens. Basic Protocol 3 discusses the Alizarin Red/Alcian Blue whole mount staining, which allows the evaluation of the shape and size of the skeleton. Basic Protocols 4–10 describe the most common stainings we routinely perform on paraffin-embedded tissue sections. In particular, Basic Protocol 4 and 5 describe H&E and Safranin O stainings, which allow morphological examination of the growth plate and analysis of glycosaminoglycans (GAGs) accumulation, respectively. Basic Protocol 6 discusses in situ hybridization, a powerful technique for the identification of specific mRNA species within individual cells in tissue sections. Basic Protocol 7 describes how to perform in situ cell death detection (TUNEL assay), an established method that detects DNA fragments and thus cell death. Basic Protocol 8 gives a brief overview of a standard immunohistochemistry technique, which allows for the detection of antigens in tissue sections. Basic Protocol 9 provides a description of BrdU proliferation assay, which allows cell proliferation evaluation by directly measuring DNA synthesis through an antibody-based detection of the nucleoside analog bromo-deoxyuridine (BrdU); Support Protocol 9 provides details on how to detect BrdU with a fluorescent dye. Alternate Protocol 9 illustrates the EdU proliferation assay, which represents a quicker and easier alternative to BrdU staining. PCNA staining, a third procedure to estimate cell proliferation, is presented in Basic Protocol 10, and Support Protocol 10 provides details on how to detect PCNA with a fluorescent dye. Basic Protocol 11 provides a description of EF5 staining, which detects levels of hypoxia in the growth plate by immunofluorescence. Basic Protocol 12 and Alternate Protocol 12 detail a simple and standardized method to quantify signals on histological sections. Lastly, Basic Protocol 13 and Support Protocol 13 give a brief overview on chondrocyte isolation and adenovirus transduction for in vitro studies.

BASIC PROTOCOL 1: MOUSE EMBRYOS HARVESTING FOR PARAFFIN SECTIONS

The analysis of endochondral bone development can start as early as E11.5 when formation of mesenchymal condensation occurs. In this first section we will describe how to isolate, fix, dissect and process tissues from mouse embryos. We will also detail all the essential steps to obtain paraffin-embedded tissue sections of fetal growth plates. Paraffin sections are indeed suitable for a variety of stainings, such as in situ hybridization analysis, immunohistochemistry with selected antibodies, BrdU and EdU labeling and TUNEL assay.

1. Isolation

Materials

• Isoflurane or CO₂ chamber

• 1X sterile Phosphate Buffer Saline (PBS) (see recipe #1)

• 70% ethanol (EtOH) (see recipe #2)

• Sterile scalpels, scissors and forceps

• Ice

• 100mm Petri dishes

• Stereo microscope

Methods

1. Sacrifice the pregnant female with isoflurane or CO₂.

2. Immediately after sacrifice, pinch the abdominal skin and make a longitudinal incision (1 cm) with surgical scissors. Pull the skin apart to expose the abdominal cavity.

3. Carefully cut the peritoneum and move the intestine to better locate the uterus.

4. Remove the uterus.

5. Immediately place the uterus on ice.

6. Prepare the stereo microscope and place a Petri dish with 1X sterile PBS on the stage.

7. Quickly rinse the uterus with sterile 1X PBS.

8. Peel the soft tissue with sterile forceps and quickly remove each embryo from its amniotic sac.

9. Place the embryos in 1X sterile PBS.

10. Rinse each embryo 3 times with sterile 1X PBS in clean Petri dishes.

11. Clean the instruments with 70% EtOH between different embryos.

2. Fixation

After isolation, the embryos need to be fixed in 4% paraformaldehyde (PFA) at 4°C. This cross-linked fixative gives good accessibility and retention of mRNAs.

Materials

• 4% PFA (see recipe #3)

• 1X PBS (see recipe #1)

• 70% EtOH (see recipe #2)

• 25mL glass scintillation vials

• 100mm Petri dishes

• Forceps

Methods

1. Fill glass scintillation vials with 20–25mL 4% PFA.

2. Rinse each embryo with 1X PBS in a clean Petri dish.

3. Open the embryo’s abdomen with scissors to facilitate rapid fixation, taking care not to nick the organs or ribs.

4. Place each embryo in a glass scintillation vial.

5. Store the vials at 4°C.

6. Replace 4% PFA with fresh 4% PFA after 24 hours.

7. Replace 4% PFA with 70% EtOH the following day.

8. Store the specimens at 4°C until dissection.

   • For optimal results, dissection should be performed within 1–2 weeks.

3. Dissection

Mouse embryos can be dissected before or after fixation. However, fixation preserves tissues and gives the investigator time to properly dissect the specimens.

Materials

• Micro dissecting forceps and scissors

• 100mm Petri dishes

• 1X PBS (see recipe #1)

• 70% EtOH (see recipe #2)

• Microsette biopsy cassettes

• Plastic or glass containers

• Stereo microscope

Methods

1. Place a clean Petri dish with a small amount of 1X PBS on the stage of the stereo microscope.

2. Remove the embryo from the glass scintillation vial and place it in the Petri dish.

3. Select the forelimb or hindlimb to dissect and carefully remove it by cutting the proximal end with microdissecting scissors.

4. Peel off the thin skin layer from the limb with forceps.

5. Carefully remove the soft tissues surrounding bone and cartilage with the forceps without smashing the growth plate or the bone.

6. Place the dissected specimen in a microsette biopsy cassette and store it in 70% EtOH at 4°C until processing.

4. Processing

Processing includes the stepwise dehydration of the samples with increasing concentrations of ethanol, followed by clearing with xylene and impregnation with paraffin.

Because melted paraffin wax is hydrophobic, most of the water in a specimen must be removed before it can be infiltrated with wax. This process is commonly carried out by immersing specimens in a series of ethanol solutions of increasing concentration until pure, water-free alcohol is reached. A series of increasing concentrations is used to avoid excessive distortion of the tissue. Xylene is then used to “clear” the tissue and to allow wax infiltration. Xylene is miscible both with ethanol and wax, and therefore represents an ideal intermediate solvent. After treatment with xylene, the tissue can then be infiltrated with paraffin wax.

The appropriate temperature for the paraffin wax depends on the formulation used. We typically use paraffin with “low” melting temperature, below 60°C, because it better preserves epitope and mRNA integrity for immunohistochemistry and in situ hybridization, respectively.

Materials

• 70% EtOH (see recipe #2)

• 80% EtOH (see recipe #4)

• 95% EtOH (see recipe #5)

• 100% EtOH

• Xylene

• Paraffin (Fisherbrand Paraplast X-Tra Tissue Embedding Medium; Fisher Scientific; catalog# 23-021-401)

• Automated processor

Methods (using an automated processor)

1. Place the cassettes in the processor.

2. 70% EtOH 1 hour.

3. 80% EtOH 1 hour.

4. 95% EtOH 1 hour.

5. 95% EtOH 1 hour.

6. 100% EtOH 1 hour.

7. 100% EtOH 1 hour.

8. Xylene 1 hour.

9. Xylene 1 hour.

10. Paraffin wax at 58°C under vacuum 1 hour.

11. Paraffin wax at 58°C under vacuum 1 hour.

12. Paraffin wax at 58°C under vacuum 1 hour.

5. Embedding

After processing, the specimen needs to be inserted into a paraffin “block” which can be clamped onto a microtome for section cutting. The proper orientation of the specimen is the critical step during embedding.

Materials

• Paraffin

• Molds (plastic or metallic)

• Micro dissecting forceps

• Magnifying glass

• Embedding station

Methods

1. Place cassettes in the paraffin reservoir of the embedding station to equilibrate specimen to optimal embedding temperature (~20 minutes).

2. Fill the mold with paraffin.

3. Orient specimen in the mold (see Support Protocol 1).

4. Hold specimen in place with forceps, carefully move the mold over the cold plate.

5. Hold specimen in place until the bottom part of the paraffin hardens.

6. Place bottom of the cassette on top of the mold, and add some wax on the top.

7. Leave block on cold plate (~20 minutes) to completely harden.

8. Remove the block from the mold and remove the excess wax from the cassette.

6. Sectioning

Materials

• Paraffin blocks

• Disposable low profile blades

• Flat ice block

• Forceps

• Paintbrush

• Superfrost Plus slides

• Slide boxes

• Water bath

• Microtome

Methods

1. Orient the blade on the microtome (angle ~5°).

2. Set up 5μm of thickness.

3. Cool the paraffin block face down on the ice block.

   • Frequently cool the block on ice during sectioning (up to 15–20 minutes).

4. Place cooled block in microtome chuck.

5. Orient block for optimal cutting.

   • Orient the block in the microtome chuck to have the proximal and the distal growth plates at the same level (parallel section).

   • Always place block back in the chuck in the same direction.

6. Gently pull paraffin ribbon from cutting surface.

7. Gently detach ribbon from cutting surface with a fine paintbrush.

8. Float cut ribbon on water bath at 50–55°C.

9. Using positively charged (+) slides, pick up sections by gently placing the slides underneath the sections.

10. Store the slides at 4°C until use

    • Slides can be stored up to 1 year at 4°C.

SUPPORT PROTOCOL 1: ORIENTATION OF PARAFFIN-EMBEDDED SPECIMENS FOR GROWTH PLATE SECTIONING

Specimen orientation described below has been specifically designed to obtain complete and comparable longitudinal sections of the limbs. Physiological curvature of bony parts may interfere with the cutting and may lead to misinterpretation of the growth plate analysis.

Forelimb

Embed forelimbs, dorsal side down, and pads facing up.

Right forelimb

• Orient the scapula (or the humerus) to point to the right and the distal end of the radius and ulna to the left of the mold.

Left forelimb

• Orient the scapula (or the humerus) to point to the left and the distal end of the radius and ulna to the right of the mold.

Hindlimb

Embed hindlimbs with the inner thigh downwards.

Right hindlimb

• Orient the femur to point to the right and the paw to the left of the mold.

Left hindlimb

• Orient the femur to point to the left and the paw to the right of the mold.

Spine

Embed spine with the dorsal side facing up.

BASIC PROTOCOL 2: MOUSE EMBRYOS HARVESTING FOR FRESH FROZEN SECTIONS

Frozen sections of OCT-embedded (Optimal Cutting Temperature) specimens have inherent disadvantages when compared to their paraffin counterparts, such as poor morphology and poor resolution at higher magnifications. They are also technically more difficult to cut. Still, the use of frozen sections remains essential for the demonstration of many antigens by immunohistochemistry, for more sensitive detection of poorly expressed mRNAs by in situ hybridization analysis, and for detection of temperature-sensitive enzymatic activities such as alkaline phosphatase.

In this section we will thus describe all the necessary steps to obtain good quality fresh frozen sections.

1. Isolation

Proceed as described in Basic Protocol 1.

2. Dissection

Dissection must be performed immediately after isolation to obtain fresh frozen sections.

Proceed as described in Basic Protocol 1.

3. Embedding

Embedding must be performed immediately after dissection to obtain fresh frozen sections.

Materials

• Plastic molds

  • OCT Compound (Fisher Healthcare™ Tissue-Plus™, Fisher Scientific; catalog #4585)

• Flat block of dry ice

• Forceps

• −80°C freezer

Methods

1. Label base mold and partially fill the mold with OCT.

2. Place dissected tissue in the pre-labeled base molds.

3. Flatten the tissue at the bottom of the mold with forceps.

4. Let the specimen equilibrate in the liquid OCT for 1–2 minutes.

5. Put the mold on a flat block of dry ice.

   • Isopentane chilled with dry ice can also be used to freeze the OCT block.

6. Hold the tissue with forceps until the OCT on the bottom part of the mold solidifies.

7. Store the frozen block at −80°C until use.

   • Immediately store the frozen block at −80°C.

   • Frozen blocks can be stored up to 1 year at −80°C.

4. Sectioning

Materials

• Tissue frozen block

• High profile blade

• Forceps

• Paintbrush

• Superfrost Plus slides

• Slide box

• Cryostat

Methods

1. Equilibrate the frozen blocks in the cryostat at −20°C for 15 minutes.

2. Orient blade on the cryostat (angle ~30°).

3. Set up 10 μm as thickness.

4. Apply some OCT on the back of the frozen block.

5. Attach the cooled block to a specimen disc.

6. Place the specimen disc in the cryostat chuck.

7. Orient block for optimal cutting.

   • Orient the specimen in the plastic mold as described in Support Protocol 1.

   • Orient the block in the cryostat chuck to have the proximal and the distal growth plates at the same level.

8. Pick up 1 or 2 sections per slide.

9. Collect slides in slide box.

10. Store slide box at −80°C for long-term storage or −20°C for short-term storage.

    • Frozen slides can be stored up to 1–2 weeks at −20°C and up to 6 months at −80°C.

ALTERNATE PROTOCOL 2: MOUSE EMBRYOS HARVESTING FOR FIXED FROZEN SECTIONS

Formalin-fixed specimens can be also used to generate frozen sections. In this case the morphology of the tissue is better maintained than in the fresh frozen tissue.

1. Isolation, Fixation and Dissection

Follow these first three steps as described in Basic Protocol 1.

2. Embedding

Materials

• 30% sucrose/PBS (see recipe #6)

• 1X PBS (see recipe #1)

• OCT Compound (Fisher Healthcare™ Tissue-Plus™, Fisher Scientific; catalog #4585)

• Glass beaker

• Rocking platform

• Plastic container

• Plastic molds

• Flat block of dry ice

• Forceps

• −80°C freezer

Methods

Before starting:

• Soak the fixed tissues in 30% sucrose/PBS on a rocking platform overnight at 4°C.

1. Quickly rinse the sample in 1X PBS.

2. Proceed as described in Basic Protocol 2.

3. Sectioning

Follow this step as described in Basic Protocol 2.

BASIC PROTOCOL 3: ALIZARIN RED/ALCIAN BLUE WHOLE MOUNT STAINING

Whole-mount skeletal staining permits the evaluation of the shape and size of the skeleton. Alizarin Red/Alcian Blue staining distinguishes bone, which contains calcium, and cartilage, which is enriched in glycosaminoglycans (GAGs) (McLeod, 1980). GAGs or mucopolysaccharides are homo- or heteropolymers consisting of a repeating disaccharide unit, which in turn consists of an amino sugar (N-acetylglucosamine or N-acetylgalactosamine and a uronic sugar (glucuronic acid or iduronic acid) or galactose (Esko, et al., 2009). GAGs are negatively charged and therefore attract water. Alizarin Red is an anionic dye, it thus binds to calcium (Horobin, 2010); Alcian Blue on the other hand, is a cationic dye that strongly binds to the negatively charged GAGs (Schenk, 1981). This technique is thus widely used to analyze skeletal maturation and to detect defects in endochondral bone development, in mineralization and in skeletal patterning (Mangiavini, et al., 2014, Pfander, et al., 2004, Provot, et al., 2007, Schipani, et al., 2001). An example of Alizarin Red/Alcian Blue staining of a newborn mouse is shown in Figure 2.

Figure 2.

Whole mount Alizarin Red S/Alcian blue staining of newborn mouse. (Bar=5mm).

Materials

• 1X PBS (see recipe #1)

• 95% EtOH (see recipe #5)

• 100% acetone

• 0.3% Alcian Blue/70% EtOH stock solution (see recipe #7)

• 0.1% Alizarin Red S/95% EtOH stock solution (see recipe #8)

• Alizarin Red/Alcian Blue staining working solution (see recipe #9)

• 10% potassium hydroxide (KOH) stock solution (see recipe #10)

• 1% KOH working solution (see recipe #11)

• 20% glycerol/1% KOH (see recipe #12)

• 50% glycerol/1% KOH (see recipe #13)

• 80% glycerol/1% KOH (see recipe #14)

• 100% glycerol

• Ice

• 100mm Petri dishes

• Scissors and forceps

• 25mL glass scintillation vials

• 37°C incubator

Methods

1. Sacrifice the pregnant female and isolate the embryos as described in Basic Protocol 1.

2. Rinse each embryo with 1X PBS in a clean Petri dish.

3. Remove skin and viscera from the embryos with microdissecting forceps.

   Place each embryo in a glass scintillation vial filled with 95% ethanol for 5 days.

   • Incubation time in the solutions may vary based on the age of the specimens

4. Occasionally agitate the vials.

5. Transfer the specimens in 20–25mL 100% acetone for 2 days to remove fat.

6. Place specimen in 20–25mL Alizarin Red/Alcian Blue staining working solution at 37°C in the incubator for up to 5 days.

7. Briefly wash specimen in distilled water (dH₂O).

8. Clear the specimen in 1% KOH working solution for at least 12–48 hours; up to 4 days.

   • Change 1% KOH working solution every day.

9. Continue clearing the specimen through 20% glycerol/1% KOH for ~5 days-1 week.

10. Continue clearing the specimen through 50% glycerol/1% KOH for ~5 days-1 week.

    • Before proceeding to 50% glycerol/1% KOH, status of the specimen can be assessed by transferring the specimen into a Petri dish containing 20% glycerol/1% KOH and quickly examining it under a stereo microscope. However, prolonged exposure to this concentration of KOH in conjunction with the heat originating from the microscope bulb may cause disintegration of the specimen.

11. Complete clearing of the specimen in 80% glycerol/1% KOH for ~1–2 weeks.

12. Store the specimen in 100% glycerol at 4°C.

BASIC PROTOCOL 4: HEMATOXYLIN AND EOSIN STAINING ON THE FETAL GROWTH PLATE

Paraffin-embedded tissue sections are suitable for a variety of stains. A deparaffinization step and a rehydration step are necessary to remove the paraffin and to allow the stain to penetrate into the tissue. Hematoxylin and eosin staining (H&E staining) is one of the basic stainings in histology and it used to evaluate tissue morphology (Titford, 2005). Hematoxylin is a basic/positive stain and it binds to acidic and negatively charged substances such DNA and RNA. Hematoxylin generates a violet/blue nuclear staining. Eosin is acidic/negative stain and it binds to basic and positively charged substances such as proteins. Eosin generates a pink cytoplasmic staining. There are different kinds of hematoxylin, but Harris Hematoxylin is the most commonly used because its dark blue color provides a good contrast with the pink eosin (Levdik, 1989). H&E stain is a valuable tool to evaluate the morphology of the developing growth plate (Mangiavini, et al., 2014, Pfander, et al., 2004, Provot and Schipani, 2007, Schipani, et al., 2001). A representative image of H&E staining of a mouse forelimb is displayed in Figure 3.

Figure 3.

H&E staining of E15.5 mouse forelimb (Bar=100μm).

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene in Coplin jars

• 100% EtOH in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• dH₂O in Coplin jars

• Harris Hematoxylin in Coplin jars

• Eosin Y in Coplin jars

• Cold tap water

• Slides racks

• Xylene-based mounting medium

• Glass cover slips

• Slide warmer

Methods

Before starting:

• Dry paraffin-embedded tissue sections on the slide warmer at 60°C for at least 2–4 hours.

1. Deparaffinize, clear and rehydrate the tissue sections:

   • Xylene, 3 times for 2 minutes each.

   • 100% EtOH, 2 times for 1 minute each.

   • 95% EtOH, 1 time for 1 minute.

   • dH₂O, 2 times for 1 minute each.

     • H&E staining can also be performed on frozen sections thereby avoiding the deparaffinization and dehydration steps.

2. Stain slides with Harris Hematoxylin for 75 seconds.

   • Xylene, 100% EtOH, 95% EtOH, Harris Hematoxylin and Eosin Y solutions may be re-used for 3–4 H&E stains.

   • Harris Hematoxylin must be re-filtered before each use.

3. Gently rinse slides in cold tap water for 2 minutes.

   • Let the tap water run a few minutes in advance to cool it down.

   • The tap water flush must be gentle.

4. Incubate slides in dH₂O 2 times for 1 minute each.

5. Incubate slides in 95% EtOH for 2 minutes.

6. Stain slides with Eosin Y for 15 seconds.

7. Incubate slides in 95% EtOH 2 times for 30 seconds each.

8. Incubate slides in 100% EtOH 4 times for 1 minute each.

9. Incubate slides in xylene 3 times for 2 minutes each.

10. Coverslip with xylene-based mounting medium.

    • If bubbles form during coverslipping, gently press on the glass cover slip with forceps to remove them.

BASIC PROTOCOL 5: SAFRANIN O STAINING TO EVALUATE GAGs IN MOUSE GROWTH PLATE

GAGs represent an essential component of the chondrocyte matrix (see Basic Protocol 3 for details on GAGs structure). Safranin O is a red metachromatic cationic dye, which binds the negatively charged groups (polyanions) of the GAGs in a pH-dependent manner. Safranin O thus permits to evaluate in vivo the presence of GAGs in the growth plate (Mangiavini, et al., 2014). Figure 4 illustrates an example of Safranin O staining in a mouse forelimb.

Figure 4.

Safranin O staining of E15.5 mouse forelimb (Bar=100μm). The accumulation of GAGs is shown in red.

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene in Coplin jars

• 100% EtOH in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• dH₂O in Coplin jars

• Weigert’s Iron Hematoxylin working solution (Sigma, see recipe #15)

• 0.1% Fast Green solution (see recipe #16)

• 0.08% Safranin O solution (see recipe #17)

• 1% acetic water (see recipe #18)

• Wheaton glass staining dishes or Coplin jars

• Paper filters

• Slide racks

• Transfer pipettes

• Xylene-based mounting medium

• Glass cover slips

• Slide warmer

Methods

Before starting:

• Dry the paraffin-embedded tissue sections on the slide warmer overnight at 60°C.

1. Deparaffinize, clear and rehydrate the tissue sections:

   • Xylene, 3 times for 2 minutes each.

   • 100% EtOH, 2 times for 1 minute each.

   • 95% EtOH, 1 time for 1 minute.

   • dH₂O, 2 times for 1 minute each.

     • Use fresh xylene and EtOH.

2. Apply working Weigert’s Iron Hematoxylin on each section with the transfer pipette (~1mL per slide) and incubate for 10 minutes at RT.

   • Filter the solutions immediately before use.

3. Rinse sections in dH₂O for 1 minute at RT.

4. Wash sections in cold tap water for 10 minutes.

5. Rinse slides in dH₂O for 1 minute at RT.

6. Incubate sections in 0.1% Fast Green working solution for 10 minutes at RT.

   • Filter the solutions immediately before use.

7. Rinse slides in 1% acetic water for 10–15 seconds.

8. Incubate sections in 0.08% Safranin O working solution for 5 minutes at RT.

   • Filter the solutions immediately before use.

9. Dehydrate slides:

   • 95% EtOH, 2 times for 1 minute each.

   • 100% EtOH, 3 times for 1 minute each.

   • Xylene, 3 times for 2 minutes each.

10. Coverslip with xylene-based mounting medium.

11. Dry slides overnight at RT.

BASIC PROTOCOL 6: IN SITU HYBRIDIZATION: A METHOD TO DETECT mRNA IN VIVO

In situ hybridization is a powerful technique for identifying specific mRNA species within individual cells in tissue sections (Wilkinson and Nieto, 1993). In this Protocol, we describe a specific in situ hybridization technique in which the complementary RNA probes (riboprobes) are labeled with radioactive bases (³⁵S-UTP). During in situ hybridization, the riboprobe hybridizes to the target sequence; next, the probe excess is washed away after prior hydrolysis using RNase A. Parameters such as temperature and salt detergent concentration can be manipulated to eliminate any non-specific interaction (i.e. only exact sequence matches will remain bound). The bound probe is visualized using autoradiography and photographic emulsion (Mangiavini, et al., 2014, Pfander, et al., 2004, Provot, et al., 2007, Schipani, et al., 2001). Riboprobes can also be fluorescent- or antigen-labeled (digoxigenin or biotin) (Wilkinson, 1995). However, radioactive-in situ hybridization is very sensitive, and it allows detection of low abundance mRNAs. Representative pictures of in situ hybridization are shown in Figure 5.

Figure 5.

In situ hybridization for detection of type II Collagen (Col2a1, a), type X Collagen (Col10a1, b) and SOX9 (c,d) mRNAs in E15.5 mouse forelimbs. Bright-field images (black signal) are shown in (a–c). Darkfield image (silver/white signal) is shown in (d) (Bar=100μm).

In situ hybridization requires different steps:

• Labeling riboprobes

• Pre-hybridization treatment

• Hybridization

• Post-hybridization washing

• Dipping

• Developing and counterstaining

1. Labeling riboprobes

Materials

• 5X transcription buffer (Promega, catalog #P118B)

• 100mM dithiothreitol (DTT) (Promega, catalog #P117B)

• rATP (Promega, catalog #P113B)

• rCTP (Promega, catalog #P114B)

• rGTP (Promega, catalog #P115B)

• ³⁵S-UTP (Perkin Elmer, catalog #NEG739H001MC)

• RNase inhibitor (Promega, catalog #N211A)

• Linear DNA template of target gene

• RNA polymerase (T3, T7 or Sp6) (Fisher, catalog #BP3206-1; Promega, catalog # P207B- P108B)

• DNase I (Promega, catalog #M610A)

• 0.5M EDTA pH 8.0

• 5X NTE (see recipe #19)

• DEPC-treated water

• Surface decontaminant (RNase Away)

• Scintillation fluid (i.e. Scintiverse)

• Probequant G-50 columns (GE, catalog #28-9034-08)

• Ice

• RNase-free tubes

• Sterile filtered tips

• Glass scintillation vials

• Vortex

• Water bath at 37°C

• Bench top centrifuge

• Geiger counter

• Liquid Scintillation Counter (LSC)

Methods

Before starting:

• Clean all the work surfaces with RNase Away.

• Check water bath temperature (37°C).

1. Prepare the riboprobe following this order:

   • 14μL 5X transcription buffer
   • 12μL 100 mM dithiothreitol (DTT)
   • 21μL rATP
   • 31μL rCTP
   • 41μL rGTP
   • 56μL ³⁵S-UTP
   • 61.5μL RNase inhibitor
   • 71.5μL (1μg) linear DNA template of target gene
   • 82μL RNA polymerase (T3, T7 or Sp6)

     • Keep all the reagents on ice.

     • If more than 1 riboprobe is labeled, prepare a master mix of 5X transcription buffer, 100mM DTT, rATP, rCTP and rGTP.

     • Survey all the work surfaces, the supplies and yourself with the Geiger counter during the procedure.

2. Incubate the tubes in the water bath for 1 hour at 37°C.

3. Remove the tubes from the water bath and add 1μL of RNA polymerase (T3, T7 or Sp6).

4. Incubate the tubes in the water bath for an additional hour at 37°C.

5. Remove the tubes from the water bath and add 1μL of RNase inhibitor and 1μL of DNase I.

6. Incubate the tubes in the water bath for 15 minutes at 37°C.

7. Prepare the mix:

   • 7μL 0.5 M EDTA

   • 14μL 5X NTE

   • 38μL DEPC-treated water

8. Remove the tubes from the water bath and add the mix to the riboprobe.

9. Purify probes using mini Probequant G-50 columns:

   • Thoroughly vortex the column to fully suspend the Sephadex.

   • Remove bottom from column and loosen cap a quarter turn.

   • Place in sterile 2mL collection tube provided by the kit.

   • Spin at 735G for 1 minute.

   • Discard the collection tube with the eluted buffer.

   • Place the columns in fresh, sterile, RNase-free labeled microfuge tubes.

   • Apply the riboprobe to the middle of the column bed.

   • Spin at 735G for 2 minutes.

10. Quantify riboprobe activity using a LSC:

    • Fill 20mL glass scintillation vials with 5mL scintillation fluid.

    • Place 1μL of the labeled-probe in each vial.

    • Count for 1 minute.

    • Store riboprobes at −20°C until use.

      • For each labeled probe, the counts should be between 500,000 and 2 million Counts Per Minute (CPM).

2. Pre-Hybridization Treatment (first day)

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene

• 100% EtOH

• 95% EtOH (see recipe #5)

• 4L 1X PBS (see recipe #1)

• 600mL 4% PFA (see recipe #3)

• 1.2L dH₂O

• 600μL 10mg/mL proteinase K stock solution (see recipe #20)

• 600mL 10μg/mL proteinase K working solution (see recipe #21)

• 600mL 0.2N hydrochloric acid (HCl) (see recipe #22)

• 600mL 0.1M triethanolamine/30mM acetic anhydride (TEA/AA) pH 7.5 (see recipe #23)

• 600mL 70% EtOH (see recipe #2)

• RNase away

• Beakers

• 600mL Wheaton glass staining dishes (at least 5–6)

• Stainless steel slide racks

• 50mL Falcon tubes

• Slide warmer

Methods (for up to 50 slides)

Before starting:

• Dry selected slides on slide warmer overnight at 60°C.

• Clean all the work surfaces and all the supplies with RNase Away; then rinse them 3 times with dH₂O.

1. Deparaffinize, clear and rehydrate the tissue sections:

   • Xylene, 3 times for 5 minutes each.

   • 100% EtOH, 2 times for 1 minute each.

   • 95% EtOH, 1 time for 1 minute.

   • 1X PBS, 1 time for 1 minute.

     • Use fresh xylene and EtOH.

2. Transfer slides in a stainless steel slide rack in a Wheaton glass dish containing 1X PBS.

   • Avoid drying the sections during this step.

3. Wash slides in 1X PBS for 5 minutes.

4. Submerge slides in 4% PFA for 15 minutes (do not discard the solution after use).

   • Prepare 4% PFA 1–2 hours before starting the experiment.

5. Wash slides in 1X PBS for 5 minutes.

6. Submerge slides in 10μg/mL proteinase K for 15 minutes.

   • Carefully shake the slide rack at each passage to properly mix the solutions.

7. Submerge slides in 4% PFA for 10 minutes (re-use solution from step #4).

8. Wash slides in 1X PBS for 5 minutes.

9. Submerge slides in 0.2N HCl for 10 minutes.

10. Wash slides in 1X PBS for 5 minutes.

11. Pour the 0.1M TEA solution into a Wheaton glass dish containing 585mL dH₂O.

12. Transfer the slides into the dish.

13. Directly pour the AA solution onto the slides.

14. Gently shake the slide rack, and start the timer for 10 minutes.

15. Wash slides in 1X PBS for 5 minutes.

16. Submerge slides in 70% EtOH for 5 minutes.

17. Submerge slides in 95% EtOH for 5 minutes.

18. Dry the slides on a clean pad.

    • After the procedure, leave the slides on a clean pad for approximately 1 hour.

3. Hybridization (first day)

Materials

• UltraPure formamide (Sigma, catalog #47671)

• DEPC-treated water

• 50X Denhardt’s solution (see recipe #24)

• Hybridization solution (see recipe #25)

• 10mM sodium acetate (see recipe #26)

• 1M dithiothreitol (DTT) (see recipe #27)

• ³⁵S labeled riboprobes

• Humidified hybridization chamber

• RNase-free tubes

• RNase away

• Parafilm

• Vortex

• Bench top centrifuge

• Water bath at 85°C

• Hybridization oven at 55°C

• Geiger counter

Methods

Before starting:

• Prepare humidified hybridization chambers (cover the bottom of the chamber with Whatman paper, mix 50% formamide with 50% ddH₂O and distribute the solution on the paper).

• Prepare cover slips by cutting small pieces of parafilm. The size of the cover slips should match the size of the tissue sections.

• Warm up at RT an aliquot of hybridization solution.

• Defrost an aliquot of 1M DTT and vortex it to mix well.

• Check the temperatures of the water bath and of the hybridization oven (85°C and 55°C).

• Lay slides flat in the humidified hybridization chamber.

1. Dilute the riboprobe using these proportions:

   • 900μL hybridization solution

   • 50μL 1M DTT

   • 50μL ³⁵S labeled riboprobe

     • This amount of diluted riboprobe is sufficient for 10–12 small tissue sections.

     • Survey all the work surfaces, the supplies and yourself with the Geiger counter during the procedure.

2. Vortex well and spin down the solution.

3. Denature the solution in the water bath at 85°C for 3 minutes.

4. Add 80–100μL of the denatured riboprobe solution to the tissue section.

   • Quickly apply the denatured riboprobe directly on the tissue section.

5. Place a small piece of parafilm over each section, rolling to prevent bubbles.

   • If bubbles form under the parafilm cover slip, carefully remove them by pressing on the parafilm with a sterile tip.

6. Hybridize overnight at 55°C.

4. Post-Hybridization Washing (second day)

Materials

• 600mL 1X saline-sodium citrate (SSC) (see recipe #28)

• 900mL 4X SSC (see recipe #29)

• 1.8L 4X SSC/formamide 50%/50% (see recipe #30)

• 3L 2X SSC (see recipe #31)

• 900μL 20mg/ml RNase A (see recipe #32)

• 600mL 70% EtOH/0.1X SSC (see recipe #33)

• 600mL 80% EtOH/0.1X SSC (see recipe #34)

• 600mL 95% EtOH/0.1X SSC (see recipe #35)

• 600mL dH₂O

• 600mL 70% EtOH (see recipe #2)

• Wheaton glass staining dishes with lids (at least 10; use different dishes from the pre-hybridization treatment)

• Stainless steel slide racks (use different racks from the pre-hybridization treatment)

• Microwave

• Water bath at 52°C

• Water bath at 37°C

• Kodak Biomax MR single emulsion films (Kodak, catalog #870 1302)

• Autoradiography cassettes

• Clear tape

• Geiger counter

• Darkroom

Methods

Before starting:

• Warm water baths 52°C and 37°C.

• Warm a Wheaton glass dish containing 600mL of 2X SSC in the microwave for ~45 seconds.

• Place the glass dish in the water bath at 37°C and allow the solution to reach the temperature.

  • Prepare all incubation solutions in advance and heat them in the water baths.

  • Make sure that all the heated Wheaton glass dishes have a lid.

  • Check that the water levels in the water baths completely surround the solutions contained in the glass dishes.

  • Do not start the procedure until all the solutions have reached the correct temperature.

1. Place the slides in 1X SSC until all parafilm floats off the sections.

   • Do not peel off the parafilm to avoid damaging the sections.

2. Pick up all floating cover slips and discard them in the radioactive waste.

   • Survey all the work surfaces, the supplies and yourself with the Geiger counter during the procedure.

3. Incubate the slides in 2X SSC/formamide at 52°C in the water bath for 5 minutes.

4. Move slides to next glass dish and incubate them in 2X SSC/formamide at 52°C in the water bath for other 5 minutes.

5. Wash slides in 2X SSC at RT 2 times for 1 minute each.

6. Add 900μL of 20mg/mL RNase A to the warmed 2X SSC glass dish and then move the slides to the dish.

7. Incubate the slides for 30 minutes at 37°C.

8. Wash slides in 2X SSC at RT 2 times for 1 minute each.

9. Incubate the slides in 2x SSC/formamide at 52°C in the water bath for 5 minutes.

10. Incubate slides in 70% EtOH/0.1X SSC at RT for 3 minutes.

11. Incubate slides in 80% EtOH/0.1X SSC at RT for 3 minutes.

12. Incubate slides in 95% EtOH/0.1X SSC at RT for 3 minutes.

13. Rinse slides in dH₂O at RT for 10 seconds.

14. Incubate slides in 70% EtOH at RT for 1 minute.

15. Dry slides at RT for approximately 1 hour.

16. Place slides in an autoradiography cassette and secure their edges with clear tape.

17. In the darkroom, place a Kodak Biomax MR single emulsion film on top of the slides.

    • Only one side of the film has emulsion; make sure to place the film with the notched corner at the bottom right of the autoradiography cassette.

18. Expose slides to Kodak Biomax MR single emulsion film for a minimum of 12 hours.

    • The exposure time in the autoradiography cassette may vary based on the riboprobe. The signal intensity depends on the quality of the riboprobe, gene expression levels and quality of the sections. Testing signal intensity with a film is critical important to establish emulsion exposure time (see below). Exposure time in the autoradiography cassette may therefore vary between 12 and 72 hours.

19. Proceed to the next step if positive signal is detected on the film.

5. Dipping (third day)

Materials

• Kodak Autoradiography Emulsion NTB 2 (VWR Scientific, catalog #IB8895666)

• 40% glycerol stock solution (see recipe #36)

• 2% glycerol working solution (see recipe #37)

• Blade

• Opaque plastic bottles

• Heavy-duty aluminum foil

• 1L glass beaker

• Dip miser (Electron Microscopy Science, catalog # 70520)

• Metal stand for the dip miser (Electron Microscopy Science)

• Silica Gel Tel Tale (desiccant)

• Slides boxes (for 25 slides)

• 35mm Petri dishes

• Paper towels

• Tape

• Slide draining rack

• Microwave

• Water bath at 45°C

• Darkroom

Methods

Before starting:

• Divide Aliquot Kodak Autoradiography Emulsion (in the darkroom) into 5 equal parts with a blade and put into small, opaque bottles.

• Wrap each opaque bottle with 3 layers of aluminum foil.

• Warm the water-bath to 45°C in the darkroom.

• Warm some dH₂O in the microwave; fill the water bath halfway. Place a 1L beaker filled with 45°C dH₂O into the water bath. Leave the water bath lid open.

  • Before starting the procedure, check that the dH₂O temperature is 45°C both inside the 1L beaker and in the water bath.

• Place the dip miser and metal apparatus into the beaker in the water bath. Make sure that the water level is just below the top of the dip miser.

  • Carefully clean the dip miser before the procedure.

  • Check that the dip miser is dry to avoid the contamination of the emulsion with water.

• Place 1 aliquot of emulsion, and 20mL of 2% glycerol working solution outside the beaker in the water bath.

• Activate the desiccant in the microwave for 30 seconds.

• Add the desiccant to a 35mm Petri dish and secure the lid with tape.

• Place a blank slide, covered with tape, in a slide box between the desiccant and the slide area.

• Remove slides from autoradiography cassettes and place them in the slide box.

  • If more riboprobes are tested, divide them accordingly into different slide boxes.

• Keep the same orientation for each slide (example: frosted side on the left).

• Write on a label tape the riboprobe name, the number of slides, the date of dipping, and the date to develop; place the label on the appropriate slide box.

1. Take aluminum foil, slide draining rack(s) and slide boxes to the darkroom.

2. Turn on red light and turn off overhead light.

3. Add the emulsion aliquot to the 2% glycerol working solution:

   • Slowly pour down the edge of the 50mL Falcon tube.

   • Gently mix by inversion to prevent bubble formation.

   • Slowly pour the emulsion mixture in the dip miser and place it in the metal apparatus.

4. Remove label tape from the slide box; put it on the draining rack, leaving space for slides above the tape.

5. Remove slides from the slide box one at a time, and carefully dip each into the emulsion for 1 second holding the frosted end of the slide.

6. At the end of 1 second, partially lift the slide out of the dip miser and quickly dip it again 2 more times.

7. Tap the slide on some paper towel to remove the excess of emulsion.

8. Place the slide frosted end down in the draining rack (above the label tape).

9. Repeat the last two steps until all slides are coated with emulsion.

10. After 15 minutes (or more) replace slides in the appropriate boxes, keeping all frosted ends on the same side.

11. Replace lid and turn box on its side, oriented with frosted ends of the slides toward bottom.

12. Wrap each box with 3 layers of heavy-duty aluminum foil, maintaining the orientation of the slides in the box.

13. Remove the label tape from the draining tray and affix to the outside of the wrapped box (on the opposite side of the slides’ frosted ends).

14. Store the slides at RT for 12–24 hours, away from radiation and with frosted ends side down and labeled side up.

15. Transfer slide boxes to 4°C away from radiation and expose for the appropriate length of time.

    • Emulsion exposure time is usually 3 times longer than that of film; it can vary between 2–3 days and 10 days.

6. Developing and counterstaining (fourth day)

Materials

• 500mL Kodak Developer solution (see recipe #38)

• 500mL Kodak Fixer solution (see recipe #39)

• 600mL dH₂O

• Harris Hematoxylin in Coplin jars

• Tap water

• dH₂O in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• Eosin Y in Coplin jars

• 100% EtOH in Coplin jars

• Xylene in Coplin jars

• 1L glass beakers (2)

• Stainless steel slide racks

• 600mL Wheaton glass staining dishes (3)

• Xylene-based mounting medium

• Glass cover slips

• Kimwipes

• Windex

• Stirrer

• Darkroom

• Microscope with darkfield capacity

Methods

Before starting:

• Remove slide boxes from refrigerator and warm to RT for at least 30 minutes.

• Allow Developer and Fixer solutions to equilibrate at RT for ~15 minutes.

• Bring the glass dishes containing the solutions (in this order: Developer, Fixer and dH₂O), slide boxes and slide racks in the darkroom.

1. Unwrap slide boxes and place slides in the slide rack.

2. Incubate slides in the Developer solution for 2 minutes.

3. Incubate slides in the Fixer solution for 4 minutes.

4. Rinse slides in dH₂O (the light can be turn on from this step).

   • If more than one slide rack needs to be developed, perform a rinse in dH₂O between the developer and the fixer step, to keep the fixer solution clean.

   • After the fixer solution, slides can stay in dH₂O until ready to counterstain (up to 30 minutes).

5. Counter-stain:

   • Incubate slides in Harris Hematoxylin for 75 seconds.

   • Wash slides in cold tap water for 2 minutes.

   • Rinse slides in dH₂O 2 times for 1 minute each.

   • Place slides in 95% EtOH for 2 minutes.

   • Stain slides with Eosin Y for15 seconds.

   • Dehydrate slides with 95% EtOH 2 times for 30 seconds each.

   • Dehydrate slides with 100% EtOH 4 times for 1 minute each.

   • Incubate slides in xylene 3 times for 2 minutes each.

     • Use fresh EtOH and xylene.

     • Let the tap water run a few minutes in advance to have it cold.

     • The tap water flush needs to be gentle.

     • Carefully rinse the sections in cold tap water until they turn blue (about 2 minutes).

6. Coverslip with xylene-based mounting medium.

7. Dry slides overnight at RT.

8. Remove the emulsion from the back of the slides using Windex and Kimwipes.

   • Images can be acquired both in brightfield (the signal is black) and in darkfield (the signal is grey/white). However, the darkfield acquisition is much more sensitive and it is thus recommended for weak signals.

BASIC PROTOCOL 7: IN SITU CELL DEATH DETECTION (TUNEL ASSAY)

Terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) is an established method to detect DNA fragments and thus cell death. This staining relies on the ability of the enzyme terminal deoxynucleotidyl transferase (TdT) to incorporate labeled dUTP into free 3′-hydroxyl termini generated by the fragmentation of genomic DNA. TdT catalyzes the template-independent addition of deoxynucleotide triphosphates to the 3′-OH ends of DNA. When labeled nucleotides (dUTP) are incorporated by TdT, nuclei can be easily detected by standard immunohistochemical or immuno-fluorescent techniques (Gavrieli, et al., 1992, R., 1998). Immunohistochemical techniques can be very cumbersome and time consuming; we therefore prefer to use a direct immunofluorescent technique as described in this unit (Maes, et al., 2012b, Mangiavini, et al., 2014, Pfander, et al., 2004, Provot, et al., 2007, Schipani, et al., 2001). The Protocol described hereafter has been modified from the Roche “In Situ Cell Death detection Kit”. Figure 6 displays a representative TUNEL assay in mouse forelimbs.

Figure 6.

DAPI (a,c) and TUNEL (b,d) assay in E15.5 and P21 mouse forelimbs. Note the presence of TUNEL-positive cells in the perichondrium and primary spongiosa at E15.5. (Bar=100μm).

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene in Coplin jars

• 100% EtOH in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• 3L 1X PBS (see recipe #2)

• Permeabilization solution (see recipe #40)

• In Situ Cell Death detection Kit (Roche, catalog #11 684 817 910; see recipe #41)

• 5mg/mL DAPI stock solution (see recipe #42)

• 300nM DAPI working solution (see recipe #43)

• ProLong^® Gold Antifade Mountant (Life Technologies, catalog #P36930)

• PAP pen

• Humidified chamber

• Whatman paper

• Ice

• 250mL Wheaton glass staining dishes

• Slide racks

• Glass cover slips

• Aluminum foil

• Slide warmer

• Rocker platform

• Incubator at 37°C

• Cold room

• Microscope for fluorescence detection

Methods

Before starting:

• Dry the sections on the slide warmer overnight at 60°C.

• Prepare humidified chambers (cover the bottom of the chamber with Whatman paper and distribute 50mL 1X PBS on the paper).

• Prepare the permeabilization solution in the cold room.

  • Prepare fresh permeabilization solution in the cold room (at 4°C) and check its temperature.

1. Deparaffinize, clear and rehydrate the tissue sections:

   • Xylene, 3 times for 2 minutes each.

   • 100% EtOH, 2 times for 1 minute each.

   • 95% EtOH, 1 time for 1 minute.

   • 1X PBS, 1 time for 5 minutes.

     • Use fresh EtOH and xylene.

2. Incubate the slides in the permeabilization solution for 2 minutes on ice.

3. Wash the slides in 1X PBS 3 times for 2 minutes each on the rocking platform.

4. Dry the area around the tissue and circle it with the PAP pen.

   • Dry the area around the tissue before applying the PAP pen without excessively drying the tissue itself.

5. Lay the slides flat in the humidified chamber.

6. Apply ~50μL of TUNEL reaction mixture to each slide. Apply ~50μL of the label solution on the negative control (from now on keep the slides in the dark).

   • Include a negative control with each staining experiment.

7. Incubate the slides in the humidified chamber at 37°C for 1 hour.

8. Wash the slides in 1X PBS 3 times for 2 minutes each on the rocking platform.

   • Carefully remove the excess of 1X PBS on the tissue sections with a Kimwipe after every washing step.

9. Apply ~300μL of 300nM DAPI working solution to each slide.

   • DAPI is a potential mutagen and teratogen. Handle and dispose of it according to its biohazard nature.

10. Incubate the slides in the humidified chamber at RT for 2–5 minutes.

11. Wash the slides in 1X PBS 3 times for 2 minutes each on the rocking platform.

12. Coverslip with ProLong^® Gold Antifade Mountant.

13. Wrap the slides in aluminum foil and store them at 4°C.

    • Stained slides can be stored up to 2 weeks at 4°C or up to 6 months at −20°C (protected from the light).

14. Observe the sections with fluorescent microscopy.

    • Use an excitation wavelength in the range of 350–400nm and an emission wavelength of 450–500nm (blue) to detect DAPI-stained nuclei and an excitation wavelength in the range of 450–500nm and an emission wavelength in the range of 515–565nm (green) to detect TUNEL-positive cells.

Retrieve the fluorescent signal within 7 days.

BASIC PROTOCOL 8: IMMUNOHISTOCHEMISTRY ON FORMALIN-FIXED PARAFFIN-EMBEDDED TISSUES

Immunohistochemistry allows the detection of antigens in tissue sections by the use of specific antigen-antibody interactions that culminate in the attachment of a marker to the antigen (Ramos-Vara and Miller, 2014). For light microscopy, the marker may be a fluorescent dye (i.e. rhodamine, FITC, Hoechst) or the product of an enzyme (alkaline phosphatase, horseradish peroxidase: HRP) (Taylor CR, 2002). There are numerous immunohistochemical techniques, which may be used to localize antigens; the direct method is a one-step technique that requires a labeled antibody directly reacting with the antigen (Coons and Kaplan, 1950, Polak JM, 2003). The indirect method involves an unlabeled primary antibody that binds to the target antigen in the tissue and a labeled secondary antibody that reacts with the primary antibody (Coons, et al., 1955, Polak JM, 2003). In this latter application, the secondary antibody can be labeled with an enzyme (alkaline phosphatase or HRP), a fluorescent dye or with biotin. Biotin labeling requires an additional incubation with appropriately labeled streptavidin (Guesdon JL, 1979). Formalin-fixed paraffin-embedded tissues require a series of steps to make the epitopes available for antibody binding. These steps include classic deparaffinization and antigen retrieval. Antigen retrieval is necessary to break the crosslinks created by the formalin fixative, and involves the use of heat and pH or proteases (i.e. hyaluronidase, chondroitinase or proteinase K) (Bogen, et al., 2009). For the application of HRP-conjugated antibody it is necessary to prevent background staining caused by endogenous peroxidase, which can be blocked by treating the sections with a solution containing 3% hydrogen peroxide/methanol. Moreover, a blocking step is also necessary to avoid nonspecific binding of the first antibody. Common blocking buffers include normal serum, non-fat dry milk or Bovine Serum Albumin (BSA). For selected antibodies, it is necessary to amplify the signal. A variety of methods have been used to amplify the signal, such as the Renaissance Tyramide Signal Amplification (TSA) system (Perkin Elmer). The TSA technology uses HRP-dependent catalysis and deposition of biotin-labeled tyramide onto tissue sections, with minimal if any loss of resolution (Gross and Sizer, 1959). Deposited biotins are detected with HRP-streptavidin. In this Protocol we will describe an example of indirect immunohistochemistry involving the TSA technology (Mangiavini, et al., 2014, Pfander, et al., 2004, Provot and Schipani, 2007, Schipani, et al., 2001). This method is particularly recommended for weak signals.

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene in Coplin jars

• 100% EtOH in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• 1L 1X Tris Buffered Saline (TBS; see recipe #44)

• 250mL sodium citrate solution (see recipe #45)

• 250mL 3% hydrogen peroxide/methanol (H₂O₂/MeOH; see recipe #46)

• TSA kit (Perkin Elmer, catalog #NEL700A; see recipe #47)

• Unconjugated primary antibody

• 4L 1X Tris NaCl Tween 20 (TNT; see recipe #48)

• Biotin-conjugated secondary antibody

• DAB (3, 3′-diaminobenzidine) Peroxidase (HRP) substrate kit (Vector Labs, catalog # SK-4100; see recipe #49)

• Mayer’s Hematoxylin

• 1X PBS (see recipe #1)

• Xylene-based mounting medium

• Humidified chamber

• Plastic container for sodium citrate solution

• 250mL Wheaton glass staining dishes (4–5)

• Slide rack

• PAP pen

• Whatman paper

• Microwave

• Water bath at 95°C

• Rocking platform

• Glass cover slips

• Slide warmer

Methods

Before starting:

• Dry the paraffin-embedded tissue sections on the slide warmer overnight at 60°C.

• Prepare humidified chambers (cover the bottom of the chamber with Whatman paper and distribute 50mL 1X TBS on the paper).

  • Do not start the procedure until the sodium citrate solution has reached 95°C.

• 1Deparaffinize, clear and rehydrate the tissue sections:

  • Xylene, 3 times for 5 minutes each.

  • 100% EtOH, 2 times for 1 minute each.

  • 95% EtOH, 1 time for 1 minute.

  • 1X TBS, 1 time for 5 minutes.

    • Use fresh xylene and EtOH.

• 2Transfer the slides in the plastic container.
• 3Incubate slides in the sodium citrate solution for 10 minutes at 95°C in the water bath.
• 4Remove the container from the water bath.
• 5Let slides in the sodium citrate solution for approximately 30 minutes at RT.
• 6Incubate slides in 3% H₂O₂/MeOH for 10 minutes at RT.
• 7Wash slides in dH₂O for 5 minutes at RT.
• 8Wash slides in 1X TBS for 1 minute at RT.

  • Carefully remove the excess of 1X TBS on the tissue sections with a Kimwipe after every washing step.

• 9Circle section with PAP pen.

  • Dry the area around the tissue before applying the PAP pen without drying the tissue itself.

• 10Lay the slides flat in the humidified chamber.
• 11Apply approximately 100μL of TNB (see recipe #47) on each slide.
• 12Incubate slides in moist chamber for 30 minutes at RT.
• 13Pour off TNB and immediately apply 100μL of unconjugated primary antibody (appropriately diluted in TNB). Apply only TNB on negative controls.

  • Keep all the reagents on ice during the procedure.

  • Include a negative control with each staining experiment.

• 14Incubate slides in moist chamber overnight at 4°C.

The following day:

• 15Equilibrate slides at RT for 30 minutes.
• 16Wash slides in 1X TNT (see recipe #48) 3 times for 2 minutes each on a rocking platform.
• 17Apply 100μL of the appropriate biotinylated secondary antibody (appropriately diluted in TNB).

  • Keep all the reagents on ice during the procedure.

• 18Incubate slides in moist chamber for 30 minutes at RT.
• 19Wash slides in 1X TNT 3 times for 2 minutes each on a rocking platform.
• 20Apply 100μL of HRP-Streptavidin (HRP-SA from TSA kit) diluted 1:100 in TNB.
• 21Incubate slides in moist chamber for 30 minutes at RT.
• 22Wash slides in 1X TNT 3 times for 2 minutes each on a rocking platform.
• 23Apply 100μL of Biotynil Tyramide (from TSA kit) diluted 1:50 in amplification diluent (from TSA kit).
• 24Incubate slides in moist chamber for 5 minutes at RT.
• 25Wash slides in 1X TNT 3 times for 2 minutes each on a rocking platform.
• 26Apply 100μL of HRP-SA (from TSA kit) diluted 1:100 in TNB.
• 27Incubate slides in moist chamber for 30 minutes at RT.
• 28Wash slides in 1X TNT 3 times for 2 minutes each on a rocking platform.
• 29Apply 100μL of DAB solution on each section.

  • Protect DAB from the light and use the solution within 1 hour.

  • It is possible to add 2 drops of Nickel solution (from DAB HRP substrate kit) to the DAB to obtain a darker signal.

• 30Incubate slides in moist chamber until brown color develops.
• 31Rinse slides in dH₂O.
• 32Counterstain slides with 2 drops of Mayer’s Hematoxylin.
• 33Incubate slides in moist chamber for 3 minutes at RT.
• 34Wash slides in cold tap water for 2 minutes.
• 35Incubate slides in 1X PBS for 30 seconds.
• 36Wash slides in dH₂O for 1 minute.
• 37Dehydrate slides:

  • 95% EtOH, 2 times for 1 minute each.

  • 100% EtOH, 4 times for 1 minute.

  • Xylene, 3 times for 2 minutes each.

• 38Coverslip with xylene-based mounting medium.
• 39Dry slides overnight.

BASIC PROTOCOL 9: PROLIFERATION ASSAYS BrdU ASSAY TO EVALUATE CELL PROLIFERATION

Measuring chondrocyte ability to proliferate is crucial for the study of endochondral bone development. The most accurate method to study chondrocyte proliferation rate is by directly measuring DNA synthesis through an antibody-based detection of the nucleoside analog bromo-deoxyuridine (BrdU) (Gratzner, 1982). BrdU is a uridine derivative that can be incorporated into DNA in place of thymidine during S-phase. BrdU is then detected by a direct immunoreaction using a labeled monoclonal anti-BrdU antibody. We currently use a specific kit (Invitrogen) to perform this technique (Mangiavini, et al., 2014, Schipani, et al., 2001). This assay requires the injection of BrdU intraperitoneally in a pregnant mouse. Figure 7 illustrates an example of BrdU assay in a fetal mouse growth plate.

Figure 7.

BrdU assay in E15.5 mouse proximal epiphysis of tibia. (Bar=100μm).

1. In vivo labeling with BrdU and FdU

Materials

• 10mg/mL 5-bromo-2′-deoxyuridine (BrdU)-1.2mg/mL 5-fluoro-2′-deoxyuridine (FdU) solution (see recipe #50)

• 1X PBS (see recipe #1)

• 4% PFA (see recipe #3)

• 1mL syringes

• 27G needles

• Balance

• Ice

• Sterile 100mm Petri dishes

• Glass scintillation vials

• Sterile scissors and forceps

Methods

1. Weigh pregnant mouse.

2. Inject intraperitoneally 10mg/mL BrdU-1.2mg/mL FdU solution 2 hours before sacrificing the mouse (0.1mL of 10mg/ml BrdU-1.2mg/mL FdU stock solution per 10g of mouse).

   • BrdU is a potential mutagen and teratogen. Handle and dispose of it according to its biohazard nature.

   • FdU is used to inhibit endogenous production of thymidine that could compete with BrdU.

   • The labeling time for skeletal tissues is typically 30 to 60 min for embryos at the gestation days 10.5 (E10.5) to E13.5, 1 to 2 hours for E14.5 to E18.5 fetuses, and 2 to 4 hours for post-natal mice.

3. Deliver the embryos by cesarean section as described in Basic Protocol 1.

4. Place the embryos on ice.

   • Upon isolation, immediately place the embryos on ice to ensure BrdU incorporation stops at same time for all embryos.

5. Separate mouse embryos and extensively rinse them with sterile 1X PBS in Petri dish.

6. Rinse each embryo 3 times with sterile 1X PBS in clean Petri dishes.

7. Fix, dissect, process and section specimens as described in Basic Protocol 1.

2. BrdU detection

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene in Coplin jars

• 100% EtOH in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• 4L 1X PBS (see recipe #1)

• 3% H₂O₂/MeOH (see recipe #46)

• dH₂O

• BrdU staining kit (Invitrogen, catalog #93-3943; see recipe #51)

• DAB HRP Peroxidase substrate kit (Vector Labs; catalog #SK-4100; see recipe #49)

• PAP pen

• Xylene-based mounting medium

• 250mL Wheaton glass staining dishes (4–5)

• Slide rack

• Whatman paper

• Humidified chamber

• Glass cover slips

• Rocking platform

• Incubator at 37°C

• Slide warmer

Methods

Before starting:

• Dry the paraffin-embedded tissue sections on the slide warmer overnight at 60°C.

• Prepare humidified chambers (cover the bottom of the chamber with Whatman paper and distribute 50mL 1X PBS on the paper).

1. Deparaffinize, clear and rehydrate the tissue sections:

   • Xylene, 3 times for 5 minutes each.

   • 100% EtOH, 2 times for 1 minute each.

   • 95% EtOH, 1 time for 1 minute.

   • 1X PBS, 1 time for 5 minutes.

     • Use fresh xylene and EtOH.

2. Incubate slides in 3% H₂O₂/MeOH for 10 minutes at RT.

3. Wash slides in 1X PBS 3 times for 2 minutes each on a rocking platform.

4. Circle section with PAP pen.

   • Dry the area around the tissue before applying the PAP pen without drying the tissue itself.

5. Lay the slides flat in the humidified chamber.

6. Apply trypsin solution (from Invitrogen kit, see recipe #51) on each slide.

7. Incubate slides in moist chamber for 15 minutes at 37°C.

8. Wash slides in dH₂O 3 times for 2 minutes each on a rocking platform.

   • Carefully remove the excess of liquid on the tissue sections with a Kimwipe after every washing step.

9. Apply denaturing solution (reagent #2 from Invitrogen kit) on each slide.

10. Incubate slides in moist chamber for 30 minutes at RT.

11. Wash slides in 1X PBS 3 times for 2 minutes each on a rocking platform.

12. Apply blocking solution (reagent #3) on each slide.

13. Incubate slides in moist chamber for 30 minutes at RT.

14. Remove blocking solution and apply 2 drops of biotinylated mouse anti-BrdU (reagent #4).

15. Incubate slides in moist chamber for 2 hours at RT.

16. Wash slides in 1X PBS 3 times for 2 minutes each on a rocking platform.

17. Apply 2 drops of HRP-streptavidin solution (reagent #5).

18. Incubate slides in moist chamber for 15 minutes at RT.

19. Wash slides in 1X PBS 3 times for 2 minutes each on a rocking platform.

20. Apply 100μL of DAB HRP substrate solution on each section.

    • Protect DAB from the light and use the solution within 1 hour.

    • DAB HRP Peroxidase substrate is also provided by the Invitrogen kit; however we prefer using DAB HRP substrate kit from Vector labs as the signal is stronger.

    • It is possible to add 2 drops of Nickel solution to the DAB to obtain a darker signal.

    • Detection with a fluorescent dye is also possible (for details see Support Protocol 9).

21. Incubate slides in moist chamber for 10 minutes at RT.

22. Rinse slides in dH₂O.

23. Counterstain slides with 2 drops of Hematoxylin (reagent #7).

24. Incubate slides in moist chamber for 5 minutes at RT.

25. Wash slides in cold tap water for 2 minutes.

26. Incubate slides in 1X PBS for 30 seconds.

27. Wash slides in dH₂O for 1 minute.

28. Dehydrate slides:

    • 95% EtOH, 2 times for 1 minute each.

    • 100% EtOH, 4 times for 1 minute.

    • Xylene, 3 times for 2 minutes each.

29. Coverslip with xylene-based mounting medium.

30. Dry slides overnight.

SUPPORT PROTOCOL 9: BrdU DETECTION WITH A FLUORESCENT DYE

Additional Materials

• Streptavidin Alexa-Fluor^®-conjugated

• 5mg/mL DAPI stock solution (see recipe #42)

• 300nM DAPI working solution (see recipe #43)

• ProLong^® Gold Antifade Mountant (Life Technologies, catalog #P36930)

• Aluminum foil

• Microscope for fluorescence detection

Methods

1. Proceed as described in Basic Protocol 9 up to step #15.

2. Replace HRP-streptavidin solution (reagent #5 from Invitrogen kit) with Streptavidin Alexa-Fluor^®-conjugated (diluted 1:100 in blocking solution reagent #1 from Invitrogen kit).

3. Incubate slides in humidified chamber (protected from light) for 15 minutes at RT.

4. Wash slides in 1X PBS 3 times for 2 minutes each on a rocking platform.

5. Apply 300μL of DAPI working solution on each section.

   • DAPI is a potential mutagen and teratogen. Handle and dispose of it according to its biohazard nature.

6. Incubate slides in humidified chamber (protected from light) for 2–5 minutes at RT.

7. Wash slides in 1X PBS 3 times for 2 minutes each on a rocking platform.

8. Coverslip with ProLong^® Gold Antifade Mountant.

9. Wrap the slides in aluminum foil and store them at 4°C.

   • Stained slides can be stored up to 2 weeks at 4°C or up to 6 months at −20°C (protected from the light).

10. Observe the sections with fluorescent microscopy within 7 days.

    • Use an excitation wavelength in the range of 350–400nm and an emission wavelength of 450–500nm (blue) to detect DAPI-stained nuclei and the appropriate excitation and emission wavelengths to detect Alexa Fluor®-BrdU-positive cells.

ALTERNATE PROTOCOL 9: EdU ASSAY TO EVALUATE CELL PROLIFERATION

A major disadvantage of BrdU detection is that it requires the use of trypsin (or similar reagents), which may cause tissue damage. Moreover, BrdU staining is a time consuming procedure. More recently, an easier and quicker proliferation assay has been developed (Salic and Mitchison, 2008). It utilizes 5-ethynyl-2′-deoxyuridine (EdU), which is a nucleoside analog of thymidine and is incorporated into DNA during S-phase. Detection is based on a copper-catalyzed cycloaddition, where a stable triazole ring is formed by covalently coupling between the alkaline group present in the EdU and the Alexa-Fluor^®-conjugated azide group. This technique also requires the injection of EdU intraperitoneally in pregnant mouse. An example of EdU assay in a fetal mouse growth plate is shown in Figure 8.

Figure 8.

DAPI (a) and EdU (b) assay in E15.5 mouse proximal epiphysis of tibia. Merged picture is shown in (c). (Bar=100μm).

1. In vivo labeling with EdU

Materials

• Click-iT® EdU Alexa Fluor^® 488 Imaging Kit (Invitrogen, catalog #C10337; see recipe #52)

• 1X PBS (see recipe #1)

  1mL syringes

• 27G needles

• Balance

• Ice

• Sterile 100mm Petri dishes

• Glass scintillation vials

• Sterile scissors and forceps

Methods

1. Weigh pregnant mouse.

2. Inject intraperitoneally EdU stock solution 2 hours before sacrificing the mouse (0.1mL of 10mM EdU stock solution per 10g of mouse).

   • EdU is a potential mutagen and teratogen. Handle and dispose of it according to its biohazard nature.

3. Deliver the embryos by cesarean section as described in Basic Protocol 1.

4. Place the embryos on ice.

5. Separate mouse embryos and extensively rinse them with sterile 1X PBS.

6. Rinse each embryo 3 times with sterile 1X PBS in clean Petri dishes.

7. Clean the instruments with 70% EtOH between different embryos.

8. Fix, dissect, process and section specimens as described in Basic Protocol 1.

2. EdU detection

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene in Coplin jars

• 100% EtOH in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• 2L 1X PBS (see recipe #1)

• Click-iT® EdU Alexa Fluor^® 488 Imaging Kit (Invitrogen, catalog #C10337; see recipes #53,54,55,56,57)

• Humidified chamber

• 250mL Wheaton glass dishes

• Slide rack

• PAP pen

• ProLong^® Gold Antifade Mountant (Life Technologies, catalog #P36930)

• Glass cover slips

• Slide warmer

Methods

Before starting:

• Dry the paraffin-embedded tissue sections on the slide warmer overnight at 60°C.

• Prepare humidified chambers (cover the bottom of the chamber with Whatman paper and distribute 50mL 1X PBS on the paper).

1. Deparaffinize, clear and rehydrate the tissue sections:

   • Xylene, 3 times for 5 minutes each.

   • 100% EtOH, 2 times for 1 minute each.

   • 95% EtOH, 1 time for 1 minute.

   • 1X PBS, 1 time for 5 minutes.

     • Use fresh xylenes and EtOH.

2. Prepare the Click-iT^® reaction cocktail following this order:

   • 430μL 1X Click-iT^® reaction buffer.

   • 20μL CuSO4 (component E).

   • 1.2μL Alexa Fluor azide working solution.

   • 50μL 1X Click-iT^® EdU buffer additive working solution.

     • Prepare Click-iT® reaction cocktail in that specific order to obtain the optimal reaction.

     • Use the Click-iT® reaction cocktail within 15 minutes from preparation.

     • That volume of Click-iT® reaction cocktail can stain 10 small tissue sections.

3. Circle tissue sections with PAP pen.

   • Dry the area around the tissue before applying the Pap Pen without drying the tissue itself.

4. Lay the slides flat in the humidified chamber.

5. Apply the Click-iT^® reaction cocktail (100–200μL) to each section and incubate for 30 minutes at RT in the humidified chamber (protected from the light).

   • Perform all the assay steps in the dark.

6. Wash slides in 1X PBS 2 times for 2 minutes each (protected from the light).

   • Carefully remove the excess of PBS on the tissue sections with a Kimwipe after every washing step.

7. Apply Hoechst 33342 working solution (100–200μL) to each section and incubate for 2 minutes at RT in the humidified chamber (protected from the light).

   • 300nM DAPI working solution (see recipe #43) may be used instead of the Hoechst 33342 working solution.

   • Hoechst 33342 and DAPI are potential mutagens and teratogens. Handle and dispose of them according to their biohazard nature.

8. Wash slides in 1X PBS 2 times for 2 minutes each (protected from the light).

9. Coverslip the slides with ProLong^® Gold Antifade Mountant.

10. Wrap the slides in aluminum foil and store them at RT for 24 hours and then at 4°C.

    • Stained slides can be stored up to 2 weeks at 4°C or up to 6 months at −20°C (protected from the light).

11. Observe the sections with fluorescent microscopy.

    • Use an excitation wavelength in the range of 350–400nm and an emission wavelength of 450–500nm (blue) to detect Hoechst 33342-stained nuclei (or DAPI) and an excitation wavelength in the range of 450–500nm and an emission wavelength in the range of 515–565nm (green) to detect Alexa Fluor® 488-EdU-positive cells.

    • Retrieve the fluorescent signal within 7 days.

BASIC PROTOCOL 10: PROLIFERATING CELL NUCLEAR ANTIGEN (PCNA) ASSAY

Proliferating cell nuclear antigen (PCNA) is a cofactor of DNA polymerase delta and it is synthesized during S, G2 and M phases of mitosis (Moldovan, et al., 2007). PCNA is thus considered an antigen expressed during DNA synthesis and its detection by immunohistochemistry represents a valid readout of cell proliferation. However, PCNA immunostaining is not the gold standard method to analyze cell proliferation as it reveals cells that are proliferating at the time of tissue collection, but not cells that were proliferating within a defined period of time (Yu, et al., 1992). This assay is thus used as a second independent method to assess cell proliferation. The Protocol discussed below has been modified from “Invitrogen PCNA STAINING KIT” (Mangiavini, et al., 2014).

Materials

• 5μm paraffin-embedded tissue sections on Superfrost Plus slides

• Xylene in Coplin jars

• 100% EtOH in Coplin jars

• 95% EtOH in Coplin jars (see recipe #5)

• 4L 1X PBS (see recipe #1)

• 250mL sodium citrate solution (see recipe #45)

• 3% H₂O₂/MeOH (see recipe #46)

• PCNA STAINING KIT (Invitrogen, catalog # 93-1143)

• DAB HRP Peroxidase substrate kit (Vector Labs, catalog #SK-4100; see recipe #49)

• dH₂O

• PAP pen

• Xylene-based mounting medium

• Plastic container for sodium citrate solution

• 250mL Wheaton glass staining dishes (4–5)

• Slide rack

• Whatman paper

• Humidified chamber

• Glass cover slips

• Microwave

• Water bath at 95°C

• Rocking platform

• Slide warmer

Methods

Before starting:

• Dry the paraffin-embedded tissue sections on the slide warmer overnight at 60°C.

• Prepare humidified chambers (cover the bottom of the chamber with Whatman paper and distribute 50mL 1X PBS on the paper).

• 2Deparaffinize, clear and rehydrate the tissue sections:

  • Xylene, 3 times for 5 minutes each.

  • 100% EtOH, 2 times for 1 minute each.

  • 95% EtOH, 1 time for 1 minute.

  • 1X PBS, 1 time for 5 minutes.

    • Use fresh xylenes and EtOH.

• 3Transfer the slides in the plastic container.
• 4Incubate slides in the sodium citrate solution for 10 minutes at 95°C in the water bath.
• 5Remove the container from the water bath.
• 6Let slides in the sodium citrate solution for approximately 30 minutes at RT.
• 7Incubate slides in 3% H₂O₂/MeOH for 10 minutes at RT.
• 8Wash sections in 1X PBS 3 times for 2 minutes each on a rocking platform.
• 9Circle tissue sections with PAP pen.

  • Dry the area around the tissue before applying the Pap Pen without drying the tissue itself.

• 10Lay the slides flat in the humidified chamber.
• 11Apply 2/3 drops of blocking solution (reagent #1 from Invitrogen PCNA STAINING KIT) on each section.
• 12Incubate slides in the humidified chamber for 30 minutes at RT.
• 13Apply 2/3 drops of biotinylated mouse anti-PCNA (reagent #2) on each section.
• 14Incubate sections in the humidified chamber for 1 hour at RT.
• 15Wash sections in 1X PBS 3 times for 2 minutes each on a rocking platform.
• 16Apply 2/3 drops of HRP-Streptavidin (reagent #3) on each section.
• 17Incubate slides in the humidified chamber for 10 minutes at RT.
• 18Wash sections in 1X PBS 3 times for 2 minutes each on a rocking platform.
• 19Apply 100μL of DAB HRP substrate solution each section.

  • It is possible to add 2 drops of Nickel solution (from DAB HRP substrate kit) to the DAB to obtain a darker signal.

  • Detection with a fluorescent dye is also possible (see Support Protocol 10).

• 20Incubate slides in moist chamber for 5 minutes at RT.
• 21Rinse slides in dH₂O for 1 minute.
• 22Counterstain slides with 2 drops of Hematoxylin (reagent #5).
• 23Incubate slides in moist chamber for 2 minutes at RT.
• 24Wash slides in cold tap water for 2 minutes.
• 25Incubate slides in 1X PBS for 1 minute.
• 26Wash slides in dH₂O for 1 minute.
• 27Dehydrate slides:

  • 95% EtOH, 2 times for 1 minute each.

  • 100% EtOH, 4 times for 1 minute.

  • Xylene, 3 times for 2 minutes each.

• 28Coverslip with xylene-based mounting medium.
• 29Dry slides overnight.

SUPPORT PROTOCOL 10: PCNA DETECTION WITH A FLUORESCENT DYE

Additional Materials

• Streptavidin Alexa-Fluor^®-conjugated (diluted 1:100 in blocking solution reagent #1 from Invitrogen PCNA STAINING KIT)

• 300nM DAPI working solution (see recipe #43)

• ProLong^® Gold Antifade Mountant (Life Technologies, catalog #P36930)

• Aluminum foil

• Microscope for fluorescence detection

Methods

1. Proceed as described in Basic Protocol 10 up to step #14.

2. Apply 100–200μL of Streptavidin Alexa-Fluor^®-conjugated.

   • Apply Streptavidin Alexa-Fluor®-conjugated in the dark and protect slides from the light.

3. Incubate sections in the humidified chamber for 10 minutes at RT.

4. Apply 100–200μL of DAPI working solution.

   • DAPI is a potential mutagen and teratogen. Handle and dispose of it according to its biohazard nature.

5. Incubate sections in the humidified chamber for 2–5 minutes at RT.

6. Wash sections in 1X PBS 3 times for 2 minutes each on a rocking platform.

   • Carefully remove the excess of PBS on the tissue sections with a Kimwipe after every washing step.

7. Coverslip the slides with ProLong^® Gold Antifade Mountant.

8. Wrap the slides in tin foil and store them at RT for 24 hours and then at 4°C.

   • Stained slides can be stored up to 2 weeks at 4°C or up to 6 months at −20°C (protected from the light).

9. Observe the sections with fluorescent microscopy.

   • Use an excitation wavelength in the range of 350–400nm and an emission wavelength of 450–500nm (blue) to detect DAPI-stained nuclei and the right excitation and emission wavelength to detect positive cells based on the type of Streptavidin Alexa-Fluor®-conjugated used.

   • Retrieve the fluorescent signal within 7 days.

BASIC PROTOCOL 11: EF5 STAINING: A METHOD TO DETECT HYPOXIA IN THE DEVELOPING GROWTH PLATE

EF5 is a pentafluorinated derivative of etanidazole, a compound developed at the University of Pennsylvania by Dr. Cameron Koch and Dr. Sydney Evans (Koch, 2002). EF5 has a nitro (NO₂) group attached to the imidazole ring structure. In hypoxia, the NO₂ group is reduced to an amino group (NH₂) (Horsman, et al., 2012, Kizaka-Kondoh and Konse-Nagasawa, 2009). One of the intermediate products of this reduction is highly reactive and can bind to any cellular protein, forming adducts that can be recognized with specific antibodies (Horsman, et al., 2012, Kizaka-Kondoh and Konse-Nagasawa, 2009, Koch, 2002). EF5 is thus a marker to detect hypoxia and is used in vitro and in vivo (Horsman, et al., 2012, Kizaka-Kondoh and Konse-Nagasawa, 2009, Koch, 2002). Chondrocytes in the developing growth plate survive and differentiate in a hypoxic environment; changes in oxygen levels in cartilage are therefore critical for cartilage development (Amarilio, et al., 2007, Maes, et al., 2012a, Provot and Schipani, 2007, Provot, et al., 2007, Schipani, et al., 2001). We analyze levels of hypoxia in the growth plate by immunofluorescent detection of EF5 adducts (Provot, et al., 2007, Schipani, et al., 2001). This assay requires the intraperitoneal injection of EF5 in a pregnant mouse. A representative picture of EF5 staining in a mouse forelimb is displayed in Figure 9.

Figure 9.

EF5 staining of E15.5 mouse elbow joint. (Bar=100μm).

1. In vivo labeling with EF5

Materials

• 10mM EF5 stock solution (see recipe #59)

• 1mL syringes

• 27G needles

• Balance

• Ice

• Sterile 100mm Petri dishes

• Scintillation glass vials

• Sterile forceps

Methods

1. Weigh pregnant mouse.

2. Inject intraperitoneally 10mM EF5 stock solution 2 hours before sacrificing the mouse (0.1mlL of 10mM EF5 stock solution per 10g of mouse).

   • The EF5 powder and the antibodies need to be directly purchased from Dr. Cameron Koch (University of Pennsylvania Department of Radiation Oncology) at this site: http://hypoxia-imaging.org.

3. Deliver the embryos by cesarean section.

4. Place the embryos on ice.

5. Dissect mouse embryos and extensively rinse them with sterile 1X PBS in a Petri dish.

6. Rinse each embryo 3 times with sterile 1X PBS in clean Petri dishes.

7. Clean the instruments with 70% EtOH between different embryos.

8. Dissect the embryos and immediately embed the specimens in OCT as described in Basic Protocol 2.

2. EF5 detection

Materials

• 10μm fresh frozen tissue sections on Superfrost Plus slides

• 100% acetone

• 2L 1X PBS (see recipe #1)

• 5% mouse serum/PBS (blocking solution; see recipe #60)

• 3% BSA/PBS (antibody carrier; see recipe #61)

• Anti-EF5 conjugated antibody

• dH₂O

• 300nM DAPI working solution (see recipe #43)

• ProLong^® Gold Antifade Mountant (Life Technologies, catalog #P36930)

• PAP pen

• Whatman paper

• Humidified chamber

• Slide racks

• 250mL Wheaton glass staining dishes

• Aluminum foil

• Microscope for fluorescence detection

Methods

Before starting:

• Incubate slides in 100% acetone for 10 minutes at 4°C.

  • Place 100% acetone at 4°C few hours before starting the stain.

• Dry the fresh frozen sections overnight at RT.

• Prepare humidified chambers (cover the bottom of the chamber with Whatman paper and distribute 50mL 1X PBS on the paper).

1. Wash sections with 1X PBS 2 times for 2 minutes each.

   • Do not wash slides on a rocking platform and be gentle in moving slides from one solution to another because frozen sections tend to easily detach.

2. Circle tissue sections with PAP pen.

   • Dry the area around the tissue before applying the Pap Pen without drying the tissue itself.

3. Lay the slides flat in the humidified chamber.

4. Apply 100–200μL of 5% mouse serum/PBS blocking solution on each section.

5. Incubate slides in humidified chamber for 30 minutes at RT.

6. Dilute anti-EF5 conjugated antibody 1:25 in 3% BSA/PBS (protected from light).

7. Apply 100–200μL of the solution on each section (negative controls get only 3% BSA/PBS).

   • Include a negative control with each staining experiment.

   • Protect slides from the light after the incubation with the anti-EF5 conjugated antibody.

   • The anti-EF5 antibody may be conjugated with the following substrates: biotin, Cy3, Cy5 or Alexa 488. We recommend the use of an anti-EF5 antibody conjugated with a fluorescent dye.

8. Incubate slides in humidified chamber for 2 hours at RT in the dark.

9. Wash sections with 1X PBS 2 times for 2 minutes each in the dark.

10. Rinse in dH₂O.

11. Apply 100–200μL of 300nM DAPI working solution on each section.

    • DAPI is a potential mutagen and teratogen. Handle and dispose of it according to its biohazard nature.

12. Incubate slides in humidified chamber for 5 minutes at RT in the dark.

13. Coverslip the slides with ProLong^® Gold Antifade Mountant.

14. Wrap the slides in aluminum foil and store them at RT for 24 hours and then at 4°C.

    • Stained slides can be stored up to 2 weeks at 4°C or up to 6 months at −20°C (protected from the light).

15. Observe the sections with fluorescent microscopy.

    • Use an excitation wavelength in the range of 350–400nm and an emission wavelength of 450–500nm (blue) to detect DAPI-stained nuclei and the right excitation and emission wavelength to detect positive cells based on the antibody used.

    • Retrieve the fluorescent signal within 7 days.

BASIC PROTOCOL 12. QUANTIFICATION OF FLUORESCENT SIGNAL-LABELED CELLS USING IMAGE J

To fully understand the growth plate development it is crucial to quantitatively assess the information given by specific staining, immunostaining or labeling techniques. To quantify these signals, automatized techniques and image analysis tools have been developed. The general principle of the method consists, first, in designing a region of interest (ROI) within which the total cell number will be calculated. Then, in the defined ROI, cells exhibiting a positive signal will be counted. This allows calculating a ratio of positive cell/total cell number.

Materials

• DAPI image (nuclei quantification or total cell number)

• Fluorescent signal image (positive cell quantification)

• Image J software

• Adobe Photoshop software

Methods

Step 1: Image superimposition and region of interest design

1. At the time of the image acquisition, make sure that you save two images of the same field:

   • The first using the DAPI channel to quantify the total cell number.

   • The second using the channel corresponding to the fluorochrome used.

   • In Photoshop, open the DAPI and the fluorescent signal pictures.

2. Create a new file that is the size of your pictures (File/new…).

3. Create 3 distinct layers (Layer/new/Layer…).

4. On the first layer copy and paste the DAPI picture.

5. On the second layer copy and paste the fluorescent signal picture.

6. On the third layer define the ROI using the rectangle or ellipse tool.

7. Save the ROI designed on the fluorescent signal image as a JPEG file.

8. Change the relative order of the layers 1 and 2, the ROI will now appear on the DAPI image. Save it as a JPEG file.

Step 2: Detection of the total cell number

1. Open Image J.

2. Open the DAPI + ROI JPEG file.

3. Select the ROI using the ellipse or rectangle tool and copy it to a new file.

   • It is particularly important to appropriately and consistently define the region of interest. This is crucial because the proportion of the cell exhibiting a positive signal or its intensity may vary from one tissue to another or even between different regions of a given tissue.

4. Go to Image, Adjust, Brightness/Contrast, Auto and save.

5. Go to Process, Binary, Make Binary (this will make a binary black and white image).

6. Go to Process, Binary, Watershed (this divides two adjacent cells that were eventually identified as one).

7. Go to Analyze, Analyze particles and select the following options:

   • Size (pixel^2): 5 – Infinity, so that noise is eliminated.

   • Circularity: 0.00 – 1.00, so that different shapes from completely elongated to perfectly circular are included.

   • Show ellipses (so you can see where and which are the cells that the program identifies).

   • Check the boxes:

     • “Display results” (individual results will be displayed in a table that can be saved as an excel file).

     • “Summarize” (summarized results will be displayed in a table that can be saved as an excel file).

     • “Add to manager”. (This plots the cells that the program identifies).

Step 3: Detection of positive cells

1. Apply the sequence described in step 2 to the Fluorescent signal image.

ALTERNATE PROTOCOL 12: QUANTIFICATION OF SIGNAL-LABELED CELLS ON BRIGHTFIELD PICTURES USING IMAGE J

Quantification of signals can also be performed on brightfield images with a method similar to the one described above. When brightfield images are used, the counting of both the total cell number and the positive stained cells is achieved on the same picture.

Materials

• Brightfield image

• Image J software

Methods

Step1: Selection of the region of interest

1. Open Image J.

2. Open the new file.

3. Select the ROI using the ellipse or rectangle tool and copy it to a new file.

4. Go to Image, Adjust, Brightness/Contrast, Auto and save.

   • It is particularly important to appropriately and consistently define the region of interest. This is crucial because the proportion of the cell exhibiting a positive signal or its intensity may vary from one tissue to another or even between different regions of a given tissue.

Step2: Detection of the total cell number

1. Go to Process, Binary, Make Binary (this will make a binary black and white image).

2. Go to Process, Binary, Watershed (this divides two adjacent cells that were eventually identified as one).

3. Go to Analyze, Analyze particles and select the following options:

   • Size (pixel^2): 5 – Infinity, so that noise is eliminated.

   • Circularity: 0.00 – 1.00, so that different shapes from completely elongated to perfectly circular are included.

   • Show ellipses (so you can see where and which are the cells that the program identifies).

   • Check the boxes:

     • “Display results” (individual results will be displayed in a table that can be saved as an excel file).

     • “Summarize” (summarized results will be displayed in a table that can be saved as an excel file).

     • “Add to manager”. (This plots the cells that the program identifies).

Step3: Detection of positive cells

1. Go to Image/Adjust/Window bar (move the Window bar so that only positive cells can be visualized).

2. Go to Process, Binary, Make Binary (this will make a binary black and white image).

3. Go to Process, Binary, Watershed (this divides two adjacent cells that were eventually identified as one).

4. Go to Analyze, Analyze particles and select the following options:

   • Size (pixel^2): 5 – Infinity, so that noise is eliminated.

   • Circularity: 0.00 – 1.00, so that different shapes from completely elongated to perfectly circular are included.

   • Show ellipses (so you can see where and which are the cells that the program identifies).

   • Check the boxes:

     • “Display results” (individual results will be displayed in a table that can be saved as an excel file).

     • “Summarize” (summarized results will be displayed in a table that can be saved as an excel file).

“Add to manager”. (This plots the cells that the program identifies).

BASIC PROTOCOL 13: IN VITRO RECOMBINATION: A TOOL TO STUDY GROWTH PLATE DEVELOPMENT

The Cre/loxP strategy represents a valuable resource in mouse genetics as it allows a variety of modifications of the mouse genome (Nagy, 2000). The strategy is based on the use of Cre recombinase, which catalyzes the recombination between two recognition sites (lox-P) inserted within specific DNA segments (Hamilton and Abremski, 1984, Schipani, 2002). This system is widely applied both for in vivo and in vitro studies to analyze the role of specific genes in growth plate development (Aro, et al., 2012, Maes, et al., 2012a, Mangiavini, et al., 2014, Pfander, et al., 2004, Provot and Schipani, 2007, Schipani, et al., 2001). For in vitro studies, chondrocytes can be isolated from mouse models with DNA sequences of specific genes flanked by two loxP sites. Recombination of the two loxP sites and consequent alteration of the gene function can be achieved by in vitro transduction with adenovirus expressing Cre recombinase. Hereafter, method for mouse chondrocyte isolation and transduction will be described (Aro, et al., 2012, Mangiavini, et al., 2014, Pfander, et al., 2004, Provot, et al., 2007).

1. Chondrocyte isolation

Materials

• Mouse embryos or newborn pups

• 100mm Petri dishes

• 70% EtOH (see recipe #2)

• Hank’s Balanced Salt Solution (HBSS) (Gibco, catalog #24020)

• Trypsin-EDTA 0.25% (Gibco, catalog #25200)

• 10X collagenase type II stock solution (Worthington Biochemicals, catalog #4174)

• DMEM, high glucose, GlutaMAX^™ Supplement, pyruvate (Gibco, catalog #10569)

• Fetal bovine serum (GE healthcare, catalog #SH30070.03)

• Penicillin-Streptomycin (5,000 U/mL) (Gibco, catalog #15070)

• 70μm sterile nylon mesh cell strainer (BD Falcon, catalog #352350)

• Primaria^™ 6 Well Cell Clear Flat Bottom Surface-Modified Multiwell Culture (Corning, catalog #353846)

• 50mL tubes

• 50mL pipetteMicropipette and tipsDissection tools

• Scalpel

• Hemocytometer

• Stereo microscopeAgitated water bath or oven + spinning device

• Class II biological safety cabinet

• Centrifuge

Phase contrast microscope

Methods

1. Prepare a series of 50mL aliquots of HBSS, in sterile conditions and warm them to 37°C.

2. Sacrifice the embryos or newborn mice according to your protocol and institutional guidelines. Rinse the pups with 70% EtOH.

   • If mice have different genotypes, treat them individually and save a piece of tail for genotyping. Use a set of clean dissection tools for each pup, or thoroughly clean the instruments with 70% EtOH in between each specimen.

3. In a Petri dish filled with HBSS, dissect forelimb and hindlimbs.

4. For each limb, cut the paw under the stereo microscope; remove the skin and muscles. Cut the epiphyses of the bones with a scalpel and place them in a 50mL tube containing approximately 40mL of HBSS stored on ice.

5. Once all the pups are completed, wash the dissected epiphyses twice in HBSS in a class II biological safety cabinet.

6. Warm the 0.25% trypsin–EDTA at 37°C for 10 minutes before use.

7. Remove HBSS and treat the epiphyses with 20mL of 0.25% trypsin–EDTA at 37°C under agitation for 30–40min.

8. After the first digestion step, let the tube stand 1 minute for the epiphyses to pellet down.

9. In the meantime, dilute the 2mL aliquot of 10X collagenase type II stock solution in 18mL of warm HBSS to obtain a 1X collagenase type II working solution at 195unit/mL.

10. Using a pipette, carefully remove the 0.25% trypsin–EDTA supernatant without removing the predigested epiphyses.

11. Transfer the 1X collagenase type II working solution on the epiphyses and incubate 2 hours at 37°C under agitation.

12. Prepare the culture medium: DMEM, high glucose, GlutaMAX^™ Supplement, pyruvate, 10% fetal bovine serum, 1% Penicillin-Streptomycin and warm it at 37°C.

13. To stop the digestion, add 20mL of culture medium to the tube containing the epiphyses and homogenize the suspension.

14. Place 70μm nylon mesh cell strainer on a clean 50mL tube and filter the suspension through to remove eventual undigested particles.

15. Save 50μL of the cell suspension for cell counting and centrifuge the rest of it for 15 minutes at 250G to pellet the cells.

16. During the centrifugation, proceed to cell counting using a hemocytometer

17. Once the cells are pelleted, carefully remove the supernatant without disturbing the cell pellet and suspend the chondrocytes in culture medium.

18. Seed the cells in 6 well plate at a density of 40,000 cell/cm² (approximately 400,000 cell/well) and let them adhere onto the culture surface for 24 hours.

2. Mouse chondrocytes transduction

Materials

• Cre Recombinase adenovirus [Vector Biolabs, catalog #1045]

• β-gal/LacZ adenovirus [Vector Biolabs, catalog #1080]

• 50mL pipette

• Micropipette and tips

• Class II biological safety cabinet

Methods

1. The next day, refresh cell culture medium to remove non-adherent cells.

2. Remove the adenoviruses from the −80°C freezer and thaw them on ice.

3. Infect the chondrocytes with the adenoviruses using a MOI (multiplicity of infection) of 400 viral particles/cell.

   • To use a MOI of 400 viral particles/cell on 400,000 cells, 160 million of viral particles will be needed. Having an adenovirus preparation with a given viral titer of 1×1010 pfu/mL, 16μL will have to be added per well.

4. The day after infection refresh cell culture medium.

   • Waste that has been in contact with adenovirus has to be inactivated according to the protocol approved by the institution (bleach or autoclave treatment).

5. 48 hours to 72 hours post-infection cell culture can be stopped and cells used for the desired analysis.

SUPPORT PROTOCOL 13. 10X COLLAGENASE TYPE II STOCK SOLUTION PREPARATION

Materials

• Hank’s Balanced Salt Solution (HBSS) (Gibco, catalog #24020)

• Collagenase type II (Worthington Biochemicals, catalog #4174)

• 1mL syringe Luer-Lok^™ Tip (Becton Dickinson, catalog #309628)

• 20G needle (Becton Dickinson, catalog #305175)

• 50mL tubes

• 50mL pipetteMicropipette and tips

• 60mL syringe Luer-Lok^™ Tip (Becton Dickinson, catalog #309653)

• 18G needle (Becton Dickinson, catalog #305196)

• 0.22μm syringe-driven filter unit (Millex, catalog #SLGV004SL)

• Class II biological safety cabinet

Methods

1. In sterile conditions, pipet 50mL of HBBS from the bottle to a clean and labeled 50mL tube in a class II biological safety cabinet.

2. Assemble together the 1mL syringe and the 20G needle. Take 1mL from the 50mL HBSS aliquot and inject HBSS directly through the septum of the collagenase type II vial.

3. Vortex the vial to dissolve the enzyme.

4. Carefully remove the aluminum cap and the septum from the vial.

5. Using a 1mL micropipette and a clean tip, transfer the content of the vial to a clean and properly labeled 50mL tube.

6. Using a 1mL micropipette and a clean tip, rinse the collagenase type II vial with 1mL of HBSS from the 50mL aliquot and transfer it to the 50mL tube containing the enzyme solution.

   • Collagenase type II vial needs to be kept at 4°C to preserve the enzymatic activity.

   • Directly injecting through the septum of the vial, limits the loss of the lyophilized powder that may occur when you open the vial.

   • Collagenase type II is supplied with an activity ≥ 125 units/mg dry weight. For instance, a vial containing 100mg of lyophilized powder with an activity of 360 units/mg has to be diluted in a final volume of 18.5mL of HBSS.

7. Repeat the previous step 3 times to obtain 5mL of the enzyme solution.

8. Dilute the 5mL enzyme solution with HBSS to obtain a “10X collagenase type II stock solution” of 1950 enzymatic unit/mL.

9. Assemble together the 60mL syringe and the 18G needle. Collect the entire volume of the 10X collagenase type II stock solution. Remove the needle from the syringe and place the filter unit instead. Sterilize the 10X collagenase type II stock solution by filtration through a 0.22μm pore membrane device.

10. Divide the 10X collagenase type II stock solution in 2mL aliquots properly labeled and store them at −20°C.

    • Aliquots may be stored up to 2 years at −20°C.

REAGENTS AND SOLUTIONS

1. 1X PBS (1L)

10X PBS	100mL
dH2O	900mL

Stir the solution and sterilize it. Store up to 1 month at 4°C.

2. 70% EtOH (1L)

100% EtOH	700mL
dH2O	300mL

Store up to 6 months at RT.

3. 4% PFA (for 600mL solution)

PFA	24g
dH2O	540mL
10N sodium hydroxide (NaOH)	600μL
10X PBS	60mL
Diethylpyrocarbonate	600μL

Warm dH₂O in the microwave until it reaches 60°C (~1 minute). Place a glass beaker on heating block in a chemical hood set at 60°C and start stirring. Add 10N NaOH first and then the PFA in a fume hood. When powder goes into solution, turn off heat and add 10X PBS. Add diethylpyrocarbonate and let stir for another 10 minutes.

Check that pH is between 7.0 and 7.4 using pH strip. (Add 10N NaOH or 12N HCl if necessary).

Cool down the solution to RT before use. 4% PFA should be used within few hours.

Store for 2–3 days at 4°C. For long-term storage, 4% PFA may be divided into aliquots and stored up to 6 months at −20°C.

4. 80% EtOH (1L)

100% EtOH	800mL
dH2O	200mL

Store up to 6 months at RT.

5. 95% EtOH (1L)

100% EtOH	950mL
dH2O	50mL

Store up to 6 months at RT.

6. 30% sucrose/PBS

Sucrose	30g
1X PBS	~100mL

Mix and bring the volume up to 100 mL. Prepare fresh just before use.

7. 0.3% Alcian Blue/70%EtOH stock solution

Alcian Blue 8 GX	1.5g
70% EtOH	500mL

Thoroughly mix and filter. The solution may be made in advance and stored up to 6 months at RT.

8. 0.1% Alizarin Red S/95% EtOH stock solution

Alizarin Red S	0.5g
95% EtOH	500mL

Thoroughly mix and filter. The solution may be made in advance and stored up to 6 months at RT.

9. Alcian Blue/Alizarin Red Staining working solution

0.3% Alcian Blue stock solution	1mL
0.1% Alizarin Red stock solution	1mL
Acetic acid, glacial	1mL
70% EtOH	17mL

Make at least 20 ml for each embryo and prepare fresh just before use.

10. 10% potassium hydroxide (KOH) stock solution

KOH	100g
dH2O	~1L

Mix until KOH is dissolved and bring the volume to 1L. May be made in advance and stored up to 6 months at RT.

11. 1% KOH working solution

10% KOH	100mL
dH2O	900mL

May be made in advance and stored up to 6 months at RT.

12. 20% glycerol/1% KOH

dH2O	700mL
Glycerol	200mL
10% KOH stock solution	100mL

May be made in advance and stored up to 6 months at RT.

13. 50% glycerol/1% KOH

dH2O	400mL
Glycerol	500mL
10% KOH stock solution	100mL

May be made in advance and stored up to 6 months at RT.

14. 80% glycerol/1% KOH

dH2O	100mL
Glycerol	800mL
10% KOH stock solution	100mL

May be made in advance and stored up to 6 months at RT.

15. Weigert’s Iron Hematoxylin working solution

• Weigert’s Iron Hematoxylin Solution A (Sigma, catalog #HT107)

• Weigert’s Iron Hematoxylin Solution B (Sigma, catalog #HT109)

Mix 50% solution A with 50 % solution B. Filter the solution.

Prepare the working solution immediately before use and use within few hours.

A prepared Weigert’s Iron Hematoxylin is commercially available (Sigma). This solution also needs to be filtered before use.

16. 0.1% Fast Green solution (100mL)

Fast Green	0.1g
dH2O	100mL

Stir until all the powder is dissolved and then filter. The working solution can be stored up to 1 week at RT but needs to be re-filtered before use.

17. 0.08% Safranin O solution (100mL)

Safranin O	0.08g
dH2O	100mL

Stir until all the powder is fully dissolved and filter. The working solution can be stored up to 1 week at RT but needs to be re-filtered before use.

18. 1% Acetic water (1000mL)

Acetic acid	10mL
dH2O	990mL

Mix well. The solution may be stored up to 1 month at RT

19. 5X NTE

1M Tris pH 7.5	5mL
0.5M EDTA pH 8.0	5mL
5M NaCl	5mL
DEPC-treated water	35mL

Clean all the supplies with RNase Away. Divide into 500μL aliquots and store up to 1 year at −20°C.

20. 10mg/mL proteinase K stock solution

Proteinase K	100mg
DEPC-treated water	10mL

Suspend the powder until fully dissolved. Divide into 600μL aliquots and store up to 1 year at −20°C.

21. 10μg/mL proteinase K working solution

10 mg/mL proteinase K stock solution	600μL
1X PBS	600mL

Clean all the supplies with RNase away. Add the 10mg/mL Proteinase K stock solution directly into the glass dish immediately before use.

22. 0.2N HCl

12N HCl	10mL
dH2O	590mL

Add 12N HCl to 600 mL dH₂O directly into the glass dish immediately before use.

23. 0.1M TEA/30mM AA pH 7.5

TEA	7.8mL
AA	1.8mL
dH2O	15mL

Mix TEA with 10mL of dH₂O into a 50mL Falcon tube. Mix well by inversion.

Mix AA with 5mL of dH₂O into another 50mL Falcon tube. Mix well by inversion. Prepare both solutions immediately before use.

24. 50X Denhardt’s solution

Ficoll	5g
Polyvinylpyrrolidone	5g
Bovine Serum Albumin (BSA)	5g
DEPC-treated water	~500mL

Clean all the supplies with RNase away. Bring the volume to 500ml and suspend the powders until fully dissolved.

Divide into aliquots and store up to 1–2 years at −20°C.

25. Hybridization solution (50mL)

UltraPure formamide	25mL
1M Tris/HCl, pH 7.6	500μL
10mg/ml tRNA	1mL
50X Denhardt’s solution	1mL
50% Dextran Sulfate	10mL
5M NaCl	6mL
10% SDS	1.25mL
DEPC-treated water	5.25mL

Clean all the supplies with RNase away. Thoroughly mix the reagents. Prepare 5mL aliquots and store up to 1–2 years at −20°C.

26. 10mM sodium acetate, pH 5.2

3M sodium acetate pH 5.5	100μL
DEPC-treated water	~30mL

Clean all the supplies with RNase away. Adjust the pH to 5.2 with HCl and bring the volume to 30mL. Prepare 1mL aliquots and store up to 1–2 years at −20°C.

27. 1M dithiothreitol (DTT)

DTT	3.09g
10mM sodium acetate, pH 5.2	~20mL

Clean all the supplies with RNase away. Suspend the powder and bring the volume to 20mL. Divide into aliquots and store up to 1–2 years at −20°C.

28. 1X SSC (600mL)

20X SSC	30mL
dH2O	570mL

Prepare immediately before use.

29. 4X SSC (900mL)

20X SSC	180mL
dH2O	720mL

Stir the solution. Distribute ~600mL in each glass dish. Prepare immediately before use.

30. 4X SSC/formamide 50%/50%

Distribute 300mL of 4X SSC into 3 glass containers.

Warm one container in microwave for 2 minutes to ~85°C.

Add 300mL formamide in fume hood.

Place the container, with lid, into a 52°C water bath.

Repeat the passages for the other two containers.

Prepare immediately before use.

31. 2X SSC (3L)

20X SSC	300mL
dH2O	2700mL

Mix well and distribute 600mL in each glass dish. Prepare immediately before use.

32. 20mg/mL RNase A

RNase A	200mg
10mM Tris/15mM NaCl buffer, pH 7.6	~10mL

Mix by inversion and bring the volume to 10mL. Divide into 900μL aliquots and store up to 1 year at −20°C.

33. 70% EtOH/0.1X SSC

100% EtOH	420mL
dH2O	177mL
20X SSC	3mL

Mix well. Add 3mL of 20X SCC just before use.

34. 80% EtOH/0.1X SSC

100% EtOH	480mL
dH2O	117mL
20X SSC	3mL

Mix well. Add 3 mL of 20X SCC just before use.

35. 95% EtOH/0.1X SSC

100% EtOH	570mL
dH2O	27mL
20X SSC	3mL

Mix well. Add 3 mL of 20X SCC just before use.

36. 40% glycerol stock solution

100% Glycerol	40mL
dH2O	60mL

Mix well. May be prepared in advance and stored up to 6 months at RT.

37. 2% glycerol working solution

40% glycerol stock solution	1mL
dH2O	19mL

Mix well. Prepare right before use.

38. Kodak Developer solution

Developer powder (Kodak, catalog #146 4726)	34g
dH2O	~500mL

Mix until the powder is fully dissolved and bring the volume to 500mL. The solution should be slightly yellow. Prepare immediately before use.

39. Kodak Fixer solution

Fixer powder (Kodak, catalog #197 1746)	90g
dH2O	~500mL

Mix until the powder is fully dissolved and bring the volume to 500mL. The solution should be cloudy white. Prepare immediately before use.

40. Permeabilization solution

Triton X-100	250μL
Sodium citrate	0.25g
dH2O	250mL

Add the reagents to the dH₂O (chilled at 4°C) in a coldroom while stirring. Prepare immediately before use.

41. TUNEL reaction mixture for 10 small sections

Enzyme solution	50μL
Label solution	450μL

Keep the reagents on ice and mix them by pipetting. Prepare immediately before use.

42. 5mg/mL DAPI stock solution

14.3mM DAPI dihydrochloride, FluoroPure™	10mg
dH2O	2mL

Suspend the powder until fully dissolved. Divide into 50μL aliquots and store up to 1–2 years at −20°C.

43. 300nM DAPI working solution

Dilute DAPI stock solution 1:5000 in 1X PBS. The DAPI working solution can be stored up to 4–6 weeks at 4°C protected from the light.

44. 1X TBS

10X TBS	100mL
dH2O	900mL

Prepare immediately before use.

45. Sodium citrate solution (antigen retrieval solution)

Citrate buffer pH 6.0	12.5mL
ddH2O	237.5mL

Mix well and microwave the solution until it reaches 90–95°C (~2 minutes). Place it in a heated water bath at 95°C and let the solution equilibrate for about 30 minutes. Prepare immediately before use.

46. 3% H₂O₂/MeOH (quenching solution)

30% H2O2	25mL
MeOH	225mL

Prepare immediately before use.

47. TNB (blocking solution)

0.05g NEN block reagent (TSA kit)	0.05g
1X TBS	10mL

Heat 1X TBS on a stir plate to dissolve block reagent. Divide into 10mL aliquots and store up to 1–2 years at −20°C.

48. 1X TNT

10X TBS	400mL
dH2O	3600mL
Tween-20	2mL

Mix well and add Tween-20 while the solution is stirring. Prepare immediately before use.

49. DAB HRP substrate solution (from Vector Labs kit)

dH2O	5mL
Buffer solution	2 drops
DAB stock solution	4 drops
H2O2 solution	2 drops

Add the buffer solution to the dH₂O (protected from the light) and mix well by vortexing. Then, add the DAB stock solution and mix well by vortexing. Lastly, add the H₂O₂ solution and mix well by vortexing. Prepare the solution immediately before use.

50. 10mg/mL BrdU-1.2 mg/mL FdU solution

BrdU	100mg
FdU	12mg
10X PBS	1mL
ddH2O	~9mL

Mix BrdU and FdU in 1 mL of 10X PBS until fully dissolved. Bring the volume to 10 mL with ddH₂O. Divide into 400 μL aliquots and store up to 3–4 years at −20°C.

51. Trypsin solution (Brdu Staining kit)

Solution 1A	1 part
Solution 1B	2 parts

Mix right before use.

52. 10mM EdU stock solution (Click-iT^® EdU Alexa Fluor^® 488 Imaging Kit)

EdU (component A)	5mg
Dimethyl sulfoxide (DMSO component C)	2mL

Mix the powder until fully dissolved. 1X PBS may be used as an alternative to DMSO. Divide into 500μL aliquots and store up to 1 year at −20°C.

53. Alexa Fluor 488 azide working solution (Click-iT^® EdU Alexa Fluor^® 488 Imaging Kit)

Alexa Fluor 488 azide (component B)	1 vial
DMSO (component C)	70μL

Prepare the Alexa Fluor azide working solution in the dark and mix well. Divide into 5μL aliquots and store up to 1 year at −20°C.

54. 1X Click-iT^® reaction buffer

• 10X Click-iT® reaction buffer (component D)

• dH₂O

Dilute component D 1:10 with dH₂O. After use, store any remaining 1X Click-iT reaction buffer at 4°C protected from the light. This 1X solution is stable for up to 6 months.

55. 10X Click-iT^® EdU buffer additive stock solution

Click-iT® EdU buffer additive (component F)	400 mg
dH2O	2mL

Mix well until all the powder is fully dissolved. Divide into 50μL aliquots and store up to 1 year at −20°C. If the solution develops a brown color, it has degraded and should be discarded.

56. 1X Click-iT^® EdU buffer additive working solution

• 10X Click-iT® EdU buffer additive stock solution

• dH₂O

Dilute the 10X Click-iT^® EdU buffer additive stock solution 1:10. Prepare the working solution right before use.

57. Hoechst 33342 working solution (Click-iT^® EdU Alexa Fluor® 488 Imaging Kit)

• Hoechst 33342 (component G)

• 1X PBS

Dilute Component G 1:2000 in the dark. Prepare Hoechst 33342 working solution right before use and protect it from the light.

58. 0.9% Saline solution

Sodium Chloride (NaCl)	9g
dH2O	1000mL

Stir until all the powder goes into solution. Sterilize. Store up to 6 months at RT.

59. 10mM EF5 stock solution

EF5	3.02mg
0.9% NaCl	1mL

Sonicate the powder for approximately 4 hours at 37°C to completely dissolve it. Solution may be prepared in advance and stored up to 1 week at RT.

60. 5% mouse serum/PBS (blocking solution)

Normal Mouse Serum	0.5mL
1X PBS	9.5mL

Mix well. Divide into 400 μL aliquots and store up to 1 year at −20°C.

61. 3% BSA/PBS (antibody carrier)

Bovine Serum Albumin (BSA)	300mg
1X PBS	10mL

Mix well until the powder is fully dissolved. Prepare fresh and store at 4°C until use.

COMMENTARY

Background Information

The staining properties of Alcian Blue and Alizarin Red have been known for over a century (Schultze, 1897) but only in more recent years, a protocol has been optimized for the study of mouse growth plate (Jegalian and De Robertis, 1992).

H&E staining, together with Safranin O, has also been essential to understand the morphological changes that occur during endochondral ossification. H&E staining has been introduced in the 1800s (Böhmer, 1865, Fischer, 1875) and it has become the most widely used stain in medical diagnosis (Wissowzky, 1876). Safranin O, which was first introduced as a nuclear counterstaining in analysis of bacteria (Coico, 2005), is routinely used to detect GAGs in the cartilaginous matrix (Schmitz, et al., 2010). Both stainings are thus commonly performed to evaluate chondrocyte morphology and growth plate matrix composition.

In situ hybridization, which was first introduced in the early 1970s (Gall and Pardue, 1969), is a critical technique to study expression of chondrocyte mRNAs on histological sections in vivo. TUNEL assay, which has been first described in 1992 (Gavrieli, et al., 1992), classically detects cells that undergo death and it is commonly used for this purpose in the analysis of the growth plate.

The process of detecting antigens (e.g., proteins) in cells of a tissue section by exploiting the principle of antibodies binding specifically to antigens in biological tissues was first developed by Albert Coons (Coons AH, 1941) and has also become one of the gold standard techniques to evaluate accumulation of specific proteins in chondrocytes in vivo.

The original protocol to assess cell proliferation was developed with the use of ³H-thymidine (Hughes, et al., 1958). More recently, this protocol has been replaced with a more sensitive and nonradioactive method that uses the thymidine analog BrdU (Gratzner, 1982). BrdU analysis in growth plate development revealed that only two layers of chondrocytes are proliferating (the round proliferative and the columnar proliferative chondrocytes), while hypertrophic chondrocytes are postmitotic cells that have exited the cell cycle. More recently, EdU detection assay, which represents an easier and highly sensitive method to detect cell proliferation, was developed (Salic and Mitchison, 2008) and it is now routinely used to detect cell proliferation in the growth plate.

EF5 detection has been recently developed by Drs. Cameron Koch and Sydney Evans (Koch, 2002). EF5 is routinely used as hypoxia marker in vivo especially in cancer research (Evans, et al., 2001), and in vitro to establish levels of oxygenation of different cell types under different culture conditions (Koch, 2002). Moreover, a few years ago, use of EF5 staining revealed the presence of an inner, hypoxic region in the fetal growth plate (Schipani, et al., 2001).

Lastly, the introduction of the Cre/loxP strategy allowed the evaluation of the role of a specific gene in chondrocyte biology both in vivo and in vitro. In particular, the use of Cre recombinase expressing adenoviruses (Anton and Graham, 1995, Wang, et al., 1996) has allowed studying of the physiological functions of specific genes in primary cells in vitro.

Critical Parameters and Troubleshooting

Basic Protocol 1

An inadequate fixation or processing of the specimens may lead to alterations in the stainings. It is therefore critical to avoid over-fixation, over- or under-processing of the samples.

Over-fixation can be prevented by:

• Avoiding prolonged fixation of the skeletal tissues (do not leave specimens in 70% EtOH for more than 1 week before dissecting and processing).

Over-processing can be prevented by:

• Decreasing the time in the dehydrating solutions.

Under-processing can be prevented by:

• Using fresh xylene and ethanol.

Specimen orientation is another critical step to obtain complete and informative sections of the fetal growth plate. An incorrect orientation may occur and can be prevented by:

• Ensuring the correct placement of the specimen in the mold (right versus left specimens; see Basic Protocol 1).

• Checking that the specimen is completely flat at the bottom of the mold.

• Applying a light pressure over the entire specimen during the initial chilling of the paraffin block.

• Quickly embedding to avoid that the paraffin chills before the specimen has adhered to the base of the mold.

Moreover, it is mandatory to obtain complete longitudinal sections of embryonic mouse limbs to be able to compare and analyze the growth plates. A good expertise in sectioning is therefore essential and it requires a specific training. Common problems during sectioning are failure to form a ribbon, wrinkled sections, varied thickness of sections, and holes in the sections. Failure to form a ribbon can be caused by a dull blade, inappropriate angle of the blade, or warm paraffin block. These issues can be prevented by:

• Changing the blade.

• Decreasing the tilt of the blade.

• Cooling the paraffin block on the ice block for a longer period of time (~15–20 minutes).

Wrinkled sections may be caused by an incorrect processing, microtome vibrations, a dull blade, warm paraffin block, or water bath temperature that is to low. These issues can be avoided by:

• Performing a correct fixation and processing of the specimens.

• Checking that the microtome is on a firm table and in good working order.

• Changing the blade.

• Cooling the paraffin block on the ice block for a longer period of time (~15–20 minutes).

• Increasing water bath temperature (not higher that 56°C) or leaving the ribbon on water for a prolonged time (10–15 minutes).

Varied thickness of sections may be due to too little blade tilt or to loose or worn microtome parts. This can be solved by:

• Increasing the tilt of the blade.

• Checking that the microtome is on a firm table and in good working order.

In the context of fetal limbs, holes usually occur at the bone marrow cavity and they are caused by an excessive dehydration of this region. This problem can be prevented by:

• Cutting short ribbons (3–4 sections).

• Briefly soaking the block in the water bath (no more than 10 seconds) after each ribbon, then drying and cooling it on the ice block for 15–20 minutes.

Basic Protocol 2

Correct orientation, embedding and sectioning are also mandatory during the preparation of fresh frozen specimens. Thus, inadequate frozen sections are often the result of issues occurring during these critical steps. Incorrect orientation or embedding can be avoided by:

• Ensuring the correct placement in the mold (right versus left specimens; see Basic Protocol 1).

• Checking that the dry ice block is completely flat.

• Checking that the specimen is completely flat at the bottom of the mold.

  Applying a light pressure over the entire specimen during the initial chilling of the OCT block.

Common problems in sectioning frozen specimens are wrinkled sections, varied thickness of sections, and holes in the sections. Wrinkled sections are generally caused by a dull blade or an improper cryostat temperature. This can be solved by:

• Changing the blade.

• Increasing the cryostat temperature (up to −15°C).

Varied thickness of sections occurs if the blade is dull or if the cryostat has loose or worn parts. It can be prevented by:

• Changing the blade.

• Checking that the cryostat is in good working order and all the parts are tightly closed.

In the context of fetal limbs, holes usually occur at the level of the bone marrow cavity due to the different consistence of the bone marrow compared to the growth plate. The problem can be prevented by:

• Increasing the cryostat temperature (up to −15°C).

• Touching the bone marrow cavity with a finger to slightly increase the temperature at that site.

Basic Protocol 3

A good contrast between Alizarin Red and Alcian Blue is required to clearly define bone and cartilage. It is thus mandatory to properly fix, stain and digest the specimens to obtain an informative staining. An incomplete or weak staining may be caused by expired solutions, improper dissection or fixation of the specimen, or short incubation time in Alcian Blue/Alizarin Red Staining working solution. This can be prevented by:

• Checking expiration dates of the reagents.

• Preparing fresh solutions each time.

• Filtering the solutions immediately before use.

• Carefully removing all the skin, organs and fat.

• Prolonging the fixation time in 95% EtOH and 100% acetone (up to 5–7 days each).

• Increasing the incubation time in Alcian Blue/Alizarin Red staining working solution (up to 10 days).

Under-digestion of the specimens may also lead to an improper staining. This is usually caused by a too short incubation time in 1% KOH and it can be avoided by:

• Changing 1% KOH each day

• Prolonging the incubation time in 1% KOH (up to 5–7 days).

By contrast, over-digestion results in disintegration of the specimen or disarticulation of skeletal elements and it is caused by an excessive incubation time in 1% KOH. It can be prevented by:

• Reducing the incubation time in 1% KOH.

• Proceeding to 20% Glycerol/1% KOH as soon as the skeleton is clearly visible through the surrounding tissue.

Basic Protocol 4

H&E staining is the first assay to be performed during the study of the growth plate as it allows an immediate evaluation of the tissue morphology. An insufficient contrast between hematoxylin and eosin, due to inadequate nuclear or cytoplasmic staining, affects the analysis of the growth plate and needs to be avoided. In particular, pale nuclear staining, due to over-fixation of the specimen, inadequate incubation time in hematoxylin or use of depleted hematoxylin, can be prevented by:

• Leaving the specimen in fixative only for a limited amount of time (few days).

• Increasing the incubation time in hematoxylin.

• Changing often hematoxylin (every 3–4 stainings).

Dark nuclear staining is usually caused by excessive incubation time in hematoxylin or excessive thickness of the sections and it can be avoided by:

• Decreasing the incubation time in hematoxylin.

• Carefully checking the thickness of the sections during the cutting.

Also, if the nuclear color is not blue, either the hematoxylin is too old or the bluing step in tap water has not been properly performed. The problem can be overcome by:

• Checking the quality of the hematoxylin.

• Checking that the tap water is cold and eventually increasing the time in tap water.

Pale cytoplasmic staining may be caused by a prolonged time in the dehydrating solutions, an inadequate incubation time in eosin or by a reduced thickness of the sections. This issue can be prevented by:

• Reducing the timing in EtOH.

• Increasing the incubation time in eosin.

• Carefully checking the thickness of the sections during the cutting.

Dark cytoplasmic staining is caused by excessive incubation time in eosin or excessive thickness of the sections and it can be prevented by:

• Decreasing the incubation time in eosin.

• Carefully checking the thickness of the sections during the cutting.

Basic Protocol 5

Safranin O staining is also critical important in the evaluation of fetal growth plate. Similarly to H&E staining, a common issue during this staining is an incorrect contrast between Safranin O and Fast Green. Pale Safranin O may be due to expired reagents and solutions, contaminated solutions with other stainings (i.e H&E staining), over-fixation of the specimens, or inadequate incubation time in the solution. To avoid this issue:

• Use fresh xylene and EtOH to deparaffinize and dehydrate the sections.

• Prepare fresh solutions every time.

• Leave the specimen in fixative only for a limited amount of time (few days).

• Increase incubation time in Safranin O.

Another common problem during Safranin O staining is the formation of blue-black precipitates, which are caused by Weigert’s Hematoxylin debris and can be avoided by:

• Filtering the solutions immediately before use.

• Extensively rinsing the sections in dH₂O after the Weigert’s Hematoxylin incubation.

Basic Protocol 6

In situ hybridization is a very sensitive assay, which requires special care at each step. Particular attention in avoiding contamination with RNases is necessary during the preparation of the riboprobes. Moreover, a variety of issues may occur at different steps and they will be more thoroughly discussed below.

Labeling riboprobes

An adequate CPM is required to obtain properly labeled riboprobes. CPM lower than 500,000 means that an inadequate ³⁵S-UTP incorporation occurred due to expired reagents, incomplete linearization of the plasmid, inefficient reaction, or expired ³⁵S-UTP. To avoid this problem:

• Properly store all the reagents.

• Do not use expired reagents.

• Check the linearization of the cDNA with 1% agarose gel.

• Check the water bath temperature during the reaction.

• Avoid using ³⁵S-UTP 3–4 weeks past the reference date (³⁵S-UTP half life is 87.4 days)

CPM greater than 2,000,000 means an excess of un-incorporated ³⁵S-UTP. If this inconvenient occurs:

• Re-purify the riboprobe with mini Probequant G-50 columns.

Pre-hybridization, Hybridization and Post-Hybridization

A weak signal or an intense background represent the most common issues during these steps. Weak signals may be caused by over-fixation of the specimens or by a too high stringency of the procedure. It can be solved by:

• Leaving the specimen in fixative only for a limited amount of time (few days).

• Increasing the incubation time in proteinase K.

• Decreasing the incubation temperatures, the salt (5NaCl and SSC) or detergent (10% SDS) concentrations in the hybridization solution and in the post-hybridization step to reduce the stringency.

An intense background signal is due to nonspecific binding of the riboprobe to free amino acids. To reduce nonspecific binding:

• Increase the incubation time in 0.1M TEA/AA pH 7.5.

• Increase the incubation temperatures, the salt (5NaCl and SSC) or detergent (10% SDS) concentration in the hybridization solution and in the post-hybridization step to augment the stringency.

• Extend incubation time in RNase A (up to 90 minutes).

Dipping

An intense background signal may also derive from the use of expired emulsion or from the use of emulsion exposed to light. To avoid this issue:

• Do not use expired emulsion.

• Properly store emulsion at 4°C and protected from light (with 3 layers of aluminum foil).

• Use a red LED safety light in the darkroom and check for light leaking in the room.

• Cover the water bath display with aluminum foil to avoid exposing the emulsion to light.

Developing and counterstaining

The developing step can be altered either by the use of expired solutions or by exposure to light. The developing step thus can be easily improved by:

• Preparing fresh solutions each time.

• Avoiding light exposure during the incubation in developer and fixer solutions.

For problems in counterstaining refer to the troubleshooting section for Basic Protocol 4.

Basic Protocol 7

TUNEL assay is also a very sensitive procedure; it is thus critical to obtain an intense fluorescent signal when performing this staining. Occasionally, the signal may be weak or the fluorescent background may be too intense. Weakness in the signal may be caused by the over-fixation of the specimens or by the use of expired reagents. To obtain a better signal:

• Avoid prolonged fixation of the specimens.

• Do not proceed with the experiment until the permeabilization solution has reached 4°C.

• Do not use expired reagents.

• Properly store reagents and solutions.

• Prepare the TUNEL mixture solution immediately before use and keep it on ice until use.

A high background is caused by nonspecific binding or by an excessive dehydration of the sections. To reduce the background:

• Do not use any enzymatic antigen retrieval in addition to the permeabilization.

• Include a negative control with each staining experiment to assess the degree of background staining.

• Do not let the sections dry out during the staining procedure.

Basic Protocol 8

Immunohistochemistry is a complex procedure, which involves many critical steps. Issues may thus occur at different steps and they usually lead to the detection of a weak signal or detection of intense background. Detection of a weak signal is caused by a variety of reasons including: over-fixation of the specimens, inefficacy of the antigen retrieval, inadequate primary antibody concentration, wrong incubation time or temperature with the primary antibody and, finally, wrong choice of the primary antibody. Different adjustments in the protocol may increase the signal, such as:

• Avoiding prolonged fixation of the specimens.

• Trying a different antigen retrieval method (i.e. trypsin, proteinase K, chondroitinase or hyaluronidase).

• Carefully titrating the antibody to find the best concentration (following the manufacturers’ indications).

• Trying a different incubation time or temperature for the primary antibody (i.e. 1 hour at RT).

• Adding multiple amplification steps with tyramide and HRP-streptavidin.

• Carefully choosing the primary antibody by comparing different manufacturers and by checking the literature.

Intense background is due to nonspecific binding. Nonspecific binding may be the consequence of: endogenous peroxidase, cross-reaction of the secondary anti-mouse antibody to endogenous mouse tissue immunoglobulins and other components, excessive incubation time in DAB HRP substrate solution. To reduce nonspecific binding:

• Increase the incubation time in 3%H₂O₂/MeOH if HRP-conjugated antibody or streptavidin is used.

• Try different blocking solutions (i.e. BSA, normal mouse serum).

• Avoid using primary antibodies raised in the same species of the specimens.

• Try to use polyclonal antibodies rather than monoclonal ones when mouse tissue is analyzed.

• Decrease incubation time with the DAB HRP substrate solution.

• Include a negative control (in which primary antibody is omitted) with each staining experiment to assess the degree of background staining.

Basic Protocol 9

Both BrdU and EdU require the injection in the abdominal cavity of the pregnant female. Thus, a possible complication during these assays is an ineffective in vivo labeling, which results in the detection of a weak signal. To avoid this issue:

• Use new reagents and properly store them.

• During injection, point the needle towards diaphragm at shallow angle in the center of the abdomen to avoid uterus and guts.

• Adjust the labeling time based on the age of the embryos. The labeling time for skeletal tissues is typically 30 to 60 min for embryos at the gestation days 10.5 (E10.5) to E13.5, 1 to 2 hours for E14.5 to E18.5 fetuses, and 2 to 4 hours for post-natal mice.

A weak signal can also be determined by other complications that may be encountered performing an immunostaining (See Troubleshooting for Basic Protocol 8 for details). Moreover, during EdU assay, a high fluorescence background can be detected and it is usually caused by the dehydration of the tissue sections. To prevent this problem:

• Do not let the sections dry out during the staining procedure.

Basic Protocol 10

Problems during the PCNA staining are the common complications that may be encountered during an immunohistochemical analysis. See Troubleshooting for Basic Protocol 8 for details.

Basic Protocol 11

Similarly to BrdU and EdU, EF5 staining also requires an intraperitoneal injection in a pregnant female. An ineffective in vivo labeling can therefore cause detection of a weak signal. To avoid this issue:

• Use new reagents and properly store them. In particular, store EF5 powder protected from the light and with desiccant at RT.

• Prepare the 10mM EF5 stock solution the day before the injection.

• Thoroughly dissolve the EF5 powder with a sonicator.

• During injection, point the needle towards diaphragm at shallow angle in the center of the abdomen to avoid uterus and guts.

General immunohistochemical difficulties may also be encountered during the EF5 staining (See Troubleshooting for Basic Protocol 8 for details).

Basic Protocol 12

Background noise may interfere with the quantification of signals. It could be assimilated as positive signal and therefore interpreted as false positive during the analysis. It is thus recommended to reduce the background at minimum levels by increasing the contrast and the threshold size. Contrast can be improved using the Image J software:

• Go to Image/Adjust/Brightness/Contrast and progressively increase the contrast.

Moreover, Image J enables to reduce the background by intensifying the threshold size:

• Go to Analyze, Analyze particles and gradually increase the “Size (pixel^2)” value until background noise is completed eliminated.

Basic Protocol 13

Chondrocyte isolation must be performed in sterile conditions to avoid contamination, which may affect the cells. Moreover, yield and or cell viability may occasionally vary during the procedure. Different parameters can be adjusted to obtain a higher number of viable cells.

Incubation time:

• If the tissue is not fully digested, the incubation time may be increased. By contrast, if the tissue is fully digested but the cells are damaged, incubation time can be reduced to preserve cell viability.

Enzyme concentration:

• If the tissue is not fully digested, increase the enzyme concentration. If the enzyme is overly active and impairs cell viability, its concentration can be reduced. Another way to decrease the enzymatic activity is to add an exogenous source of protein (bovine serum albumin 0.1–0.5% w/v, fetal calf serum 1–10% v/v).

When Cre/lox-P strategy is used, recombination efficiency is another critical parameter that needs to be appropriately modulated to reach a sufficient number of transfected cells.

To improve recombination efficiency, the MOI can be adjusted. If cell viability is impaired due to a too high MOI but recombination efficiency is low, a repeated infection procedure at low MOI can be implemented. The goal is to expose the cell to a sustained infection with a viral load of lower intensity:

• Briefly, on a 3-day period, repeat daily the infection step using a MOI of 100.

Anticipated results

All the techniques described in this Review have been optimized and standardized for the evaluation of mouse fetal growth plate. Thus, we can predict the outcome of the assays performed on mouse embryos in physiological conditions. In particular, whole mount Alizarin Red/Alcian Blue staining is supposed to show the presence of both mineralized matrix and cartilage during fetal bone development. Also, we expect to detect the presence of GAGs by Safranin O staining throughout the proliferative and columnar layer of the growth plate. Moreover, common mRNA markers of proliferative (i.e. Col2a1 and aggrecan) and hypertrophic (i.e Col10a1 and SPP1) chondrocytes should be clearly identifiable by in situ hybridization. We expect to detect BrdU or EdU positive cells in the proliferative chondrocytes as well, while we do not expect positive signals in the hypertrophic layer. At the fetal developmental stage we do not normally notice a significant number of TUNEL-positive cells within the growth plate, except for a few cells at the border with the primary spongiosa, but we usually detect cell death in the perichondrium, which therefore represents a good positive internal control. Moreover, consistent with the presence of an inner hypoxic region (Schipani, et al., 2001), we should detect positive EF5 staining in the inner portion of the fetal growth plate, particularly in proximity of the articular surface and in the pre-hypertrophic region.

Time considerations

The preparation and analysis of the samples for an exhaustive study on the fetal growth plate may require up to 1 month. Some initial steps of the process, such as fixation and processing, are standardized and completed within 4 days. By contrast, dissection and embedding timing depends on the expertise of the operator and may therefore take between few minutes to one hour.

Timing required to complete the different stainings described in this Chapter varies between few hours and few weeks. In particular, whole mount Alizarin Red/Alcian Blue staining and in situ hybridization are the most time consuming techniques as they may take up to 2 or 4 weeks depending on the age of the specimens and on the mRNA species to be detected, respectively. H&E, Safranin O and TUNEL assays on the other hand can be completed within two hours. An immunohistochemical analysis may require between 5 hours and 2 days depending on the immunostaining to be performed (see Basic Protocol 8 for details). Some of the techniques presented in this Review (BrdU, EdU and EF5 assays) require an in vivo labeling step of generally 3–4 hours (time may vary depending on the age of the embryos). Detection of signal on the BrdU and EdU assay is calculated between 5–7 hours and 1 hour, respectively. EdU assay has therefore a considerable advantage on the BrdU assay. PCNA staining is generally performed in 4–6 hours. Preparation of the 10mM EF5 stock solution is the most time consuming step of the EF5 assay as the dissolution of the powder may require up to 4–6 hours. EF5 detection is then performed in 3–4 hours. Image analysis and signal quantification are also standardized procedures that should not take more than 10–15 minutes for each image. Lastly, mouse chondrocyte isolation can be performed in approximately 6 hours; however, timing may significantly vary depending on the expertise of the operator and on the number of samples.