BManual Processing

Compared with frozen tissue sections, paraffin-embedded tissues afford greater morphological detail for cell type identification. For best results, optimize infiltration and dehydration protocols for different tissue types. Automated processing of samples submitted to a histopathology laboratory can yield poor results if one automated procedure is used for all specimen types. Hogan et al.1 described a procedure for manual tissue processing. A modified version of that protocol is outlined below. For small or soft tissue specimens (e.g., mouse embryos, full-thickness mouse skin, endocrine glands), this procedure can be done in 2 to 3 days, and steps 12 through 14 can be replaced by an additional 100% paraffin incubation. For large specimens (e.g., a whole rat brain), one week might be required for proper infiltration. Steps 2 through 4 are skipped in many protocols, regardless of tissue type. Prepare etha-nolic solutions in DEPC-H2O unless indicated otherwise. Perform all steps from fixation to dehydration (100% ethanol) at 4°C, xylene infiltration at ambient tem perature, and paraffin infiltration at 5 to 10°C above the melting point of the paraffin formulation. For example, Paraplast X-tra (Oxford Labware No. 8889-503002) melts at 50 to 54°C. Set the vacuum oven at 57 to 60°C. Vacuum is necessary to degas the paraffin and accelerate infiltration. Many variations of the following protocol are successful.

1. Flow Chart for Manual Dehydration and Paraffin Embedment

1. Fix tissue in fresh 4% buffered paraformaldehyde, 2 to 24 h

2. Wash in 1 x PBS to remove excess fixative, 60 min or overnight

(PBS minimizes shrinkage artifacts due to hypotonicity)

(Tissue can be stored in 70% ethanol at 4°C or -20°C. Prolonged storage in 100% ethanol causes tissue to become brittle and difficult to cut.)

10. Xylenes, 60 min

11. Xylenes, 60 min

(Use separate xylenes for embedment, dewaxing, and dehydration.)

12. Xylenes:Paraffin (2:1), 60 min

13. Xylenes:Paraffin (1:1), 60 min

14. Xylenes:Paraffin (1:2), 60 min

15. 100% paraffin, 60 min

16. 100% paraffin, 60 min or overnight

17. Embed in molds containing fresh 100% paraffin. If necessary, remelt to orient specimens. Cool the blocks to ambient temperature and store them in Ziploc® bags at 4°C to minimize dehydration.

2. Tips for Manual Dehydration and Paraffin Embedment

Tissue processing is easily done by hand using baked glass scintillation vials, a small vacuum oven, baked glass bottles for ethanolic solutions, histology grade xylenes, disposable Tri-Pour beakers, paraffin, and embedding molds and labels. Use the same glass vials for all steps; decant one solution and replace it with the next. Infiltration (fixative, ethanolic series, and xylenes) can be accelerated by gentle rocking. Rectangular Peel-A-Way embedding molds (22 x 30 x 30 mm; Poly-sciences No. 18646B) have an advantage over conventional flat tissue cassettes (Polysciences No. 21467) in that rectangular blocks can be placed in a microtome holder in different orientations. To make a holder for these molds, cut six rectangular holes in the lid of a Sigma shipping box (6.25 x 3.5 x 2.0 inches). Two of these boxes fit easily inside a small vacuum oven (e.g., Precision No. 31468). Prepare the specimen labels in advance, in pencil. Determine how the tissue will be sectioned, and then use sterile toothpicks to orient specimens appropriately in the molds. Tissue-Tek cold plates (Sakura Finetek No. 4650) are an inexpensive alternative to an embedding center.1

c. Paraffin Tissue Sections

1. Strategies for Data Analysis

There are many advantages to covering the entire surface of the in situ hybridization slides with serially sectioned, paraffin "ribbons." First, if the same pattern of specific hybridization is observed in consecutive sections on the same slide, the signal is probably real. Second, serial sections make it likely to obtain publication-quality data in a single experiment. An air bubble or fiber may render one section unsuitable for photomicroscopy, but a clean adjacent section often exhibits the same expression pattern. Even the expression in a single cell layer can be detected in more than one 5 |im section. Third, serial sections add a third dimension (z axis) to the data set. For example, an entire embryonic day 13.5 mouse can be serially sectioned and mounted on «16 microscope slides, and specific hybridization can be traced throughout the entire embryo. Finally, nonspecific hybridization can be controlled rigidly by using serial sections to compare results for sense and antisense cRNA on adjacent sections on the same slide. Immediately before the hybridization step, use a Pap pen (Polysciences No. 21841) to make a hydrophobic barrier between the left and right halves of microscope slides containing serially sectioned tissue. Apply hybridization mixtures containing sense cRNA to one compartment and antisense cRNA to the other compartment. Alternatively, two antisense probes can be applied to adjacent sections, on the same slide. It is imperative to apply each probe to alternating sides of the microscope slides since the emulsion is always thicker at one end, giving the appearance of more developed silver grains.

2. Tips for Cutting and Collecting Tissue Sections

Freshly embedded tissue blocks are soft and cut easily. Dehydrated blocks can be rehydrated in DEPC-H2O to improve their cutting characteristics. Cut 4 to 7 |im serial sections using disposable blades to minimize RNase contamination. Charged, precleaned microscope slides (e.g., Fisherbrand Superfrost Plus, No. 12-550-15) can be used without pretreatment. Simply rinse the slides with ethanol and wipe them dry to remove dust and fibers, immediately before use. Label slides on the frosted end with pencil or solvent-resistant marker (e.g., Securline Marker II/Superfrost, Polysciences No. 21947). Tissues will stick to the charged surface if they are dried completely on a slide warmer (42 to 45°C, overnight). Use a covered slide warmer since dust is a potential source of RNase activity.

If a flotation bath is used to collect sections, use a glass flotation tray that can be baked to destroy RNases, and do not cover the bath with aluminum foil. Metal oxides cause positive chemography in nuclear track emulsions. Alternatively, float tissue sections on DEPC-H2O applied directly to the slides using a disposable 60 cc syringe and 22 |im syringe filter (Costar No. 8110). Preheat the DEPC-H2O-covered slides at least 5 min on the slide warmer to minimize trapped air bubbles beneath the tissue sections. As the sections expand, reposition them while removing excess water with a disposable 3 cc syringe and 27 gauge needle.

The following tools are useful for these procedures: sterile, cotton-tipped applicator sticks and xylenes to clean the microtome blade; razor blades to trim blocks; paintbrushes (#00 to #2), fine pointed forceps and a disposable (27 gauge) needle to manipulate the sections; ethanol in a squirt bottle and DEPC-H2O in a spray bottle to control temperature, humidity, and static electricity; sterile cotton gauze pads to clean tools; a bucket of ice and a beaker of DEPC-H2O to chill and hydrate tissue blocks. The black waxed surface on the inside of individually wrapped autoradiographic film (Kodak) is ideal for collecting ribbons and cutting them into desired lengths. For the fundamentals of tissue sectioning, consult Hogan et al.1 and a basic histology textbook.

d. In Vitro Transcription and Generation of Radiolabeled cRNA

1. Designing Templates for In Vitro Transcription

Ideally, choose a vector that has two RNA polymerase promoters so that the same plasmid can be used to generate antisense and sense (control for nonspecific hybridization) cRNA. Subclone inserts containing 100 to 500 bp of gene-specific sequence. Avoid heterologous probes. Choose a subcloning strategy that minimizes the length of the multiple cloning region between the RNA polymerase promoter and the gene-specific sequence. Select restriction enzymes to linearize the templates for in vitro transcription, avoiding those that generate 3' protruding ends (e.g., Aat II, Apa I, Ban II, Bgl I, Bsp1286 I, BstX I, Cfo I, Hae II, HgiA I, Hha I, Kpn I, Pst I, Pvu I, Sac I, Sac II, Sfi I, Sph I). If unavoidable, incubate the linearized template with T4 DNA polymerase or Klenow fragment of E. coli DNA polymerase to produce blunt ends, using conventional protocols.2

2. Purification of Linearized Templates

Prepare clean plasmid DNA (e.g., Qiagen maxi-prep kit or comparable product). Digest aliquots (10 to 30 |g) of plasmid DNA for 1 to 2 h with the appropriate restriction enzymes for synthesis of sense and antisense cRNA. Add more enzyme and incubate longer to ensure completion. Before stopping the reaction, resolve an aliquot («2 |g) of each digest on a thin agarose gel to confirm that the template is completely linearized and that it appears as a discrete band of the predicted size. An overloaded gel will reveal residual, uncut plasmid. Purify the templates using RNase-free reagents as described in detail1,2 and outlined below. It is not necessary to gel-purify the templates, but it is very important to remove all traces of phenol and salts.

1. Extract the digests with an equal volume of Tris (pH 8) saturated phe-nol:chloroform:isoamyl alcohol 25:24:1.

2. Re-extract the aqueous phase with an equal volume of chloroform.

3. Precipitate the aqueous phase by adding 1/10 volume of 3 M sodium acetate (pH 5.2) and two volumes of ice cold ethanol.

4. After centrifugation, wash the pellet with 70% ethanol, dry, and resuspend the DNA in 10 mM Tris—1 mM EDTA at a final concentration of 1 pg/|jl. Estimate recovery at 65%.

3. The In Vitro Transcription Reaction

This is a critical step where a novice often has trouble. The protocol of Hogan et al.1 generates radiolabeled cRNA of very high specific activity. A slightly modified version of that protocol is outlined below. A common misconception is that the cRNA must be full-length. While this is important, it should not be achieved by compromising specific activity. It is imperative to generate probes of the highest specific activity possible. The key is: do not add unlabeled UTP to the reaction. The concentration of 35S UTP in the reaction is much lower than that of the other rNTPs. As 35S UTP becomes incorporated into cRNA, if the concentration of this free nucleotide becomes rate-limiting, the radiolabeled cRNAs may terminate prematurely. Even if extension is incomplete, the specific activity of the cRNA will still be very high (in the absence of unlabeled UTP). Prepare two reactions: one for the sense template and one for the antisense template. Bring reagents to ambient temperature (except enzymes) and mix as indicated below:

Reagent

Amount

Final Concentration

[35S]UTP =125 |Ci

10 |il

-6.25 ||M

(New England Nuclear, NEG-039H)

5 X transcription buffer (provided with

4 |l

enzyme)

200 mM DTT

1 |l

10 mM

RNasin (Promega)

1 |l

Linear DNA template (1 |g)

1 |l

DEPC-H2O

1 |l

10 mM @ rGTP, rATP, rCTP

1 |l

500 ||M @

RNA polymerase (T3, T7, or SP6)

1 |l

Total:

20 |l

Mix the reactants by finger-tapping the tube and incubate at 37°C for 2 h. Concentrated preparations of polymerases can be used. Additional enzyme (1 pl) can be added at the midpoint. Purify the radiolabeled cRNA as outlined below and described in more detail by Hogan et al.1 Caution: Use high quality microfuge tubes to prevent radioactive contamination, and dispose of radioactive waste appropriately.

1. Digest the DNA template by adding: 1 pl RNasin

1 pl tRNA (20 mg/ml) 1 pl RNase-free DNase (RQ1, Promega) Incubate at 37°C for 15 min.

2. Extract the radiolabeled cRNA after adding:

55 pl DEPC-H2O

10 pl 3 M sodium acetate, pH 5.2

100 pl phenol:chloroform:isoamyl alcohol (25:24:1)

3. Transfer «90 pl of the aqueous phase to a sterile microfuge tube. Add the following and precipitate the cRNA at least 60 min at -20°C:

45 pl 5 M ammonium acetate 400 pl ethanol

After centrifugation, wash the pellet with 70% ethanol. Dissolve the dried cRNA in 100 |l of 200 mM DTT. Count a 2 |l aliquot and dilute to 1 x106 cpm/pl by adding deionized formamide to 50% final concentration. Add 200 mM DTT to make up the remaining volume. Recount a 2 pl aliquot. It is important that the hybridization mixtures contain sense and antisense cRNAs at the same cpm/ml. Diluting the cRNA minimizes pipeting errors. Store the cRNA at -20°C until use. Follow the guidelines of Hogan et al.1 to evaluate 35S UTP incorporation.

For rapid analysis of the integrity of the radiolabeled cRNA, resolve 1 to 3 x106 cpm of each probe on a very thin native agarose gel. Fix the gel in 10% trichloroacetic acid for 10 min. Dry the gel and expose it to autoradiographic film overnight. Although the molecular size of the 35S cRNA cannot be estimated accurately by this method, the relative migration vs. a DNA ladder is informative when compared between experiments and DNA templates. A good probe will not always be a single discrete band; however, there should not be very high-molecular-size products or a generalized smear of radioactivity.

4. Choice of Isotope

The higher energy isotope 33P UTP can be substituted for 35S UTP to shorten exposure times.3 However, owing to its shorter half-life, this advantage is lost if the 33P isotope is not used as soon as possible after synthesis. By contrast, 35S labeled cRNA can be used for up to one month with excellent results, if initially prepared from a fresh synthesis. The 33P UTP isotope is more expensive, and, theoretically, its higher energy (longer pathlength) yields less resolution.

5. Limited Hydrolysis of Radiolabeled cRNA

Hogan et al.1 described methods for limited hydrolysis to reduce the size of the radiolabeled cRNA to 100 to 200 bp. To calculate the duration of hydrolysis, the size (kb) of the radiolabeled cRNA must be known. This step can be tricky. The length of the radiolabeled cRNA should be, but is not necessarily the same as that predicted from the linearized cDNA template. When a probe is being used for the first time, size the transcripts on denaturing polyacrylamide gels, before and after hydrolysis. Failure to do so may result in the loss of radiolabeled cRNA at this step, caused by overestimation of hydrolysis time and complete hydrolysis.

This step is more problematic for long transcripts, which have a greater probability of premature termination. In practice, excellent results can be obtained without hydrolysis even with cDNA inserts >1 kb. Clearly, limited hydrolysis should be done if satisfactory results are not obtained without hydrolysis. Ideally, when using a probe for the first time, compare results ± limited hydrolysis. The simplest approach, however, is to construct templates to contain short, gene-specific inserts that do not require limited hydrolysis and that generate full-length cRNA under the reaction conditions. Full-length transcripts provide more stringent control for nonspecific hybridization because, if designed properly, the sense and antisense cRNAs will encode opposite strands of the same sequence.

E. Prehybridization

Prehybridization procedures aim to permeabilize the tissue sections and to block nonspecific binding sites. These procedures facilitate the diffusion of cRNA into the tissue and binding to its targeted mRNAs, as well as the diffusion of excess, unbound cRNA out of the tissue in the posthybridization wash. Prehybridization protocols are optimized for specific fixation methods. High background can result when different fixatives are used in conjunction with a single prehybridization protocol. Prehybridization treatments are done conveniently in baked glass Wheaton staining dishes (250 ml capacity; with glass lids and trays; Wheaton No. 900200). The glass trays (10 slots) hold 20 slides paired back-to-back or placed in a zigzag fashion. Caution: Wheaton dishes are sensitive to rapid temperature changes. They can be baked safely at 260°C with incremental temperature changes (e.g., 90°C, 180°, 260°C). It is important, that they be undisturbed until cooled to ambient temperature.

The flow chart below contains minor modifications of the prehybridization protocol of Hogan et al.1 One significant difference is the use of pronase E (Protease type XIV; Sigma No. P-5147; 112 ^g/ml final concentration) in place of proteinase K.1 The number at each step refers to a single Wheaton dish containing the specified solution (17 dishes in this scheme). "Refresh" means that fresh solution should be added before reusing the same dish. Ethanolic solutions are indicated simply by the % ethanol in DEPC-H2O. The PBS is 1 x final concentration in DEPC-H2O. "TEA/AA" indicates 0.1 M triethanolamine buffer (Sigma No. T-1377) to which 0.25% acetic anhydride (Sigma No. A-6404) is added immediately before use. If two racks (40 slides) are processed in one experiment, most of the solutions (except for some of the lower % ethanolic solutions) should be added fresh for the second rack. Discard the 100% ethanol after deparaffinization to prevent carryover of residual xylenes and paraffin.

1. Xylenes I 10 min

2. Xylenes II 10 min

4.

100% II

5 min

5.

95%

4 min

6.

85%

4 min

7.

75%

4 min

8.

65%

4 min

9.

50%

4 min

10.

30%

2 min

11.

PBS I

5 min

12.

4% buffered paraformaldehyde

20 min

13.

PBS II

5 min

13.

PBS II refresh

5 min

14.

Pronase E

7 min

13.

PBS II

5 min

12.

4% buffered paraformaldehyde

5 min

15.

DEPC-H20

few seconds

16.

TEA/AA

10 min

17.

TEA/AA

10 min

11.

PBS I

5 min

10.

30%

2 min

9.

50%

4 min

8.

65%

4 min

7.

75%

5 min

6.

85%

5 min

5.

95%

5 min

4.

100% II refresh

5 min

3.

100% I refresh

10 min

Air dry the slides 30 to 60 min before hybridization. Cover them loosely (inverted "V") with clean aluminum foil to keep out dust.

f. Hybridization 1. Cover Glasses

Hybridization should be done using cover glasses or another type of coverslip. At 50°C, evaporation is noticeable by 24 to 36 h even with cover glasses. Evaporation is pronounced without cover glasses and introduces unacceptable variability in the experimental results. Several alternatives to cover glasses are available, including commercial RNase-free plastic covers and parafilm strips.1 Cover glasses must be siliconized to reduce surface tension and trapping of air bubbles. If a PAP pen is used to divide the slides into compartments, siliconized cover glasses are easily cut to size by scoring with a diamond pencil.

Siliconize the cover glasses (24 x 60 mm; Corning No. 2935-246) well in advance. The following protocol yields excellent results without pretreatment to inhibit RNases, provided that the cover glasses are handled only with clean forceps and clean, gloved hands. First, use fine, pointed forceps to immerse the cover glasses individually in a baked glass Coplin staining jar (Wheaton No. 900520) containing Sigmacote (Sigma No. SL-2). Sigmacote should be used near a fumehood. Dry the cover glasses in a vertical position (e.g., against a test tube rack covered with clean aluminum foil). Re-dip the cover glasses in 100% ethanol. Dry them thoroughly in a vertical position (avoid dust) and repack into the original box. Sigmacote reserved for this purpose can be reused if filtered between uses to remove dust and fibers (e.g., baked glass funnel and Whatman paper).

2. The Hybridization Mixture

Prepare the hybridization mixture from frozen stocks, just before use. Prepare more than actually needed—a generous estimate is 120 to 150 |jl per microscope slide. The protocol below was adapted from Hogan et al.1

for 1 ml hybridization mixture

10 X Salts

100 ^l

Deionized formamide

400 ^l

50% dextran sulfate

200 ^l

tRNA (20 mg/ml)

10 ^l

DTT (1 M)

8 ^l

DEPC-H2O

82 ^l

Diluted probe

200 ^l

Add radiolabeled cRNA at a final concentration of 25 x106 cpm/ml of hybridization mixture. Good results can be obtained within the range 20 to 40 x106 cpm/ml, but this should be determined experimentally. After calculating the total cpm needed, calculate the volume of each radiolabeled cRNA (previously diluted to «106 cpm/pl) that must be added. This volume should be much less than the 200 |jl allotted for diluted probe above. The difference between these two volumes should be added as a mixture of deionized formamide and 200 mM DTT (50:50).

Heat the hybridization mixture containing radiolabeled cRNA at 80 to 100°C, for 2 to 3 min, before use. Vortex vigorously and place on ice. Apply the mixture with a pipet tip (0 to 200 |jl tip) held nearly parallel to the microscope slide. Use the broad, lateral surface of the pipet tip to spread the mixture evenly, as it is pushed out of the tip. Wet all of the tissue sections with the hybridization mixture (no air bubbles) before applying a cover glass.

A simple, humidified hybridization chamber can be made using plastic Tupper-ware containers.1 Alternatively, use a plastic desiccator box (Bel-Art No. H42053-0001) placed inside a forced-air oven (e.g., Gallenkamp Plus II). Fill the desiccator tray with DEPC-H2O instead of desiccant and preheat the unit (50 to 55°C). Apply hybridization mixture and cover glasses to the slides and place them directly on the removable plastic shelves in the desiccator box. Two racks of 20 slides each are easily accommodated. Equilibrate the contents of the box to the preset oven temperature (50 to 55°C) for 10 to 15 min before sealing the door for the overnight incubation («18 h). This same oven can be used to bake the cleaned glassware (260°C) at the end of the experiment.

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