Morphological recognition of apoptotic cells


1. Introduction

In this chapter we outline the basic techniques that can be used to identify apoptotic cells on the basis of morphology. All these techniques require the use of some type of microscope, and obviously, the better the microscope then the easier your job is in assessing cell morphology. In addition to the subjective assessment of cellular/nuclear appearance, the use of immuno-cytochemical techniques specifically designed for the recognition of apoptotic cells is also outlined. Finally, we discuss how to extract quantitative information from such observations, and how such quantitative data may be interpreted. Techniques for detecting the biochemical changes accompanying apoptosis are described in subsequent chapters.

1.1 Key morphological features of apoptotic cells

A seminal paper in apoptosis research was that by Kerr et al. (1). Their electron microscopic study of prednisolone-induced cell death in the kidney defined the term apoptosis and provided the standard reference for the key morphological features associated with this form of cell death. A cell undergoing apoptosis proceeds through various stages of morphological change (see Figure 1). These are shrinkage of the cell away from its neighbours, plasma membrane blebbing, cytoplasmic and nuclear condensation, non-random cleavage of chromatin, margination of chromatin in the nucleus, nuclear fragmentation, and cellular fragmentation into smaller apoptotic bodies. Cells and cell fragments are ultimately phagocytosed by neighbouring cells and 'professional' phagocytes.

fames W. Wilson and Christopher S. Potten . Normal cell

Cell shrinkage away from neighbouring cells

Plasma membrane blebbing Cytoplasmic and nuclear condensation

Margination of condensed chromatin

Margination of condensed chromatin


Nuclear and cellular I.®, fraa mentation

Nuclear and cellular I.®, fraa mentation


Figure 1. Schematic diagram of morphological changes associated with apoptosis.

2. Light and fluorescent microscopy techniques for the assessment of apoptosis

2.1 Preparation of cell or tissue samples 2.1,1 Slide preparation

The majority of protocols outlined in this chapter require the mounting of samples on glass microscope slides. In order to increase the adhesiveness of cell or tissue sections, slides need to be coated, or 'subbed'. The two most common subbing agents are gelatine and 3-aminopropyl triethoxysilane (APES). Gelatine is a good general purpose reagent, however, if microwave-based antigen retrieval is used for immunohistochemistry that is to be carried out in parallel with morphological assessment, then APES is recommended.

Protocol 1. Gelatine subbing

1. Acid-wash the slides or covers lips for 1-2 days, in 10% acetic acid/90% ethanol.

2. Rinse (x 5) in deionized water, then 95% ethanol, and finally 100% ethanol. Store in 100% ethanol.


* 10% acetic acid

3. Dissolve 0.5 g gelatin in 100 ml of deionized water (at 65°C). Allow the solution to cool to room temperature prior to filtering through Whatman number 1 filter paper. Store the gelatine at 4°C.

4. When required, re-melt the gelatine using gentle heat (i.e. in a water bath).

5. Dip slides/coverslips in liquid gelatine, in a dust-free environment (i.e. a microbiological safety cabinet).

6. Place the slides/coverslips in a rack and allow to dry vertically.

Protocol 2. APES subbing Reagents


1. Wash the slides in decon-90 (2% in water) for 5 min, and warm water for 5 min.

2. Place the slides in an acetone bath for 1 min, prior to immersion in 2% APES in acetone, for 5 min.

3. Rinse the slides in fresh acetone for 1 min, and in running deionized water for 5 min.

2.1.2 Preparation of tissue sections

Protocol 3. Using formaldehyde-fixed, paraffin wax-embedded tissue

Equipment and reagents

• chloroform


1. Wash the tissue (animal/human) in ice-cold, phosphate-buffered saline (PBS, pH 7.4), prior to fixation in 4% formaldehyde in PBS overnight, at 4°C.

2. Dehydrate the tissue through a series of graded alcohols (70% x 3,

Protocol 3. Continued

90%, 95%, 100% X 3). Allow 30-60 min for each step, depending on the size of the tissue sample.

3. Transfer the tissue to chloroform/ethanol. After 30-60 min, transfer to 100% chloroform. Change to fresh chloroform for a further 30-60 min.

4. Place the tissue in molten wax at 60°C, for 30 min. Place under partial vacuum and leave for a further 30 min. Change to fresh wax (at 60°C) and place under full vacuum for 2 h.

5. Dispense fresh wax into a mould, place on a cooling tray, and allow to begin to set.

6. Place the tissue in the required orientation within the wax and allow to set fully.

7. Section at 3-5 (im using a microtome. Expand the sections by floating on deionized water, at about 48-50°C. Pick up the sections on to APES-or gelatin-coated slides. Rack the slides and dry at 37°C overnight.

8. Store the slides in a dry, cool place. If slides are also to be used for immunohistochemistry, store at 4°C.

Protocol 4. Using frozen tissue sections

Equipment and reagents

• liquid nitrogen

• OCT compound (optimal cutting temperature: OCT)

• cryostat and cryovials


1. Wash the tissue in ice-cold PBS (pH 7.4). Remove excess moisture by blotting on a paper towel.

2. Freeze the tissue in the vapour phase of liquid nitrogen, by placing in a small petri dish and floating this on the surface of the liquid nitrogen. Store in cryovials, under liquid nitrogen, until use. For some tissues, i.e. skeletal muscle, freezing the tissue in liquid nitrogen-cooled isopentane is advised to give better preservation of tissue architecture.

3. Embed in OCT compound and section using a cryostat (tissue may be stored in cryovials at -80°C or lower, until required).

4. Pick up sections on to APES- or gelatin-coated microscope slides or coverslips.

5. Fix sections in acetone/methanol (1:1), for 3 min at -20°C, in a spark-proof freezer. Air-dry the sections for 10 min and store at -80°C until use.

2.1.3 Preparation of cell culture samples

The preparation of samples from cell cultures is relatively straightforward and less labour intensive than the preparation of tissue sections.

Protocol 5. Preparation of cells in suspension cultures Method

1. Harvest the cells from tissue culture.

2. Resuspend the cells at a density of ~1 x 106 cells/ml.

3. Load 100-200 nl of cell suspension into the reservoir of a cytospin slide, and spin on to coated slides, at 500 r.p.m. (c. 30g) for 2-3 min (Shandon Cytospin 3). Allow to air-dry.

Cells may be fixed immediately after harvesting or after spun slides have been air-dried. Common fixation techniques are 30 min in 4% paraformaldehyde in PBS (pH 7.4) at 4°C or 3 min in acetone/methanol (1:1), at-20°C.

Unfixed cells in suspension may also be mixed directly with nuclear stains, as detailed in Protocol 70.

Protocol 6. Preparation of cells from monolayer cultures


1. Either, harvest cells using trypsin/EDTA treatment and treat as in Protocol 5.

2. Or, if cells are able, grow them on glass coverslips (or slides). Coverslips can be easily removed from the tissue culture dish using a suitable implement. Cells on slides/coverslips can be fixed according to the methods in Protocol 5.

3. Alternatively, grow cells in chamber slides. Fixation of cells on chamber slides should be carried out using 100% methanol at -20°C; acetone will dissolve the plastic slide. As an alternative to chamber slides, cells can be grown In conventional culture flasks, and when assessment of morphology is required the top and side of the flask may be removed by a model-makers electric cutting tool, or by other means.

2.2 Nuclear counterstains

There are numerous stains and dyes that are employed to assess nuclear morphology. This chapter will concentrate on the methods we currently employ in our laboratory, which we feel allow accurate assessment of nuclear morphology and apoplosis. Stains and dyes arc separated according to their use in either light or fluorescent microscopy.

2.2,1 Stains for light microscopy

Haeinatoxylin and thionin blue are the two nuclear countersigns that are most frequently used in our laboratory. They are both used for conventional light microscopic examination of sections of formaldehyde-fixed, wax-cm-bedded tissue. They give a blue stain to all nuclei and have good contrast. The condensed chromatin within apoptotic cells stains particularly heavily. Mitotic nuclei also stain darkly, but can he differentiated because of their larger size and more fuzzy appearance. The chromatin masses within apoptotic cells tend to have 'sharp' borders. In the intestinal epithelium, mitotic cells tend to appear more displaced towards the centre of the crypt lumen, although this can also be true of some apoptotic cells/bodies. In our laboratory we routinely use haematoxylin and eosin staining for assessment of apoptosis in tissue sections, in parallel with whole-tissue autoradiography for studying tritiated thymidine incorporation. Thionin blue is most commonly used as a counter-stain when apoplosis is being assessed in parallel with immunoreaetivity in wax-embedded tissue sections. Examples of apoptotic cells in haematoxylin-and thionin-stained small intestinal epithelia are shown in Figure 2.

Wax-embedded tissue sections need to be hydrated prior to staining, as in Protocol 7. If the staining is carried out at the end of a procedure such as

Rehydrating After Radiation Treatment
Figure 2. H&E staining of large intestinal crypt (A) and thionin blue staining of small intestinal crypt (B), from a mouse, 4 h after exposure to 16 Gy 7-radiation. Apoptotic cells/bodies are indicated by arrow heads.

immunocytochemistry or autoradiography, the sections will already be in a hydrated state and can be used directly, as detailed in Protocol 8.

Protocol 7. Rehydration of slides Reagents


1. Warm the slides to 60°C for 10-15 min in an oven, to melt the wax.

2. Place in fresh xylene for 5 min, with constant agitation.

3. Transfer the slides to absolute alcohol for 5 min.

4. Rehydrate the slides through a graded series of alcohols: three further changes of 100%, then 95%, 90%, 70%, and 40%, with 3 min in each.

5. Finally, rinse the slides in deionized water.

Protocol 8. Haematoxylin (and eosin: H&E)



1. Place slides in Gill's x 2 haematoxylin for 3 min.

2. Rinse slides in running water for 1 min.

3. Give slides three dips in alkali water (deionized water with 4-5 drops of ammonia solution) and return to running water for a further minute. This results in the pink staining turning blue. If the blue coloration is too dark (this can be checked quickly using a microscope), it can hinder morphological assessment and the scoring of any parallel autoradiography. The stain may be lightened by placing slides in acid water (deionized water with a few drops of HCI) for a few seconds. Then rinse the slide in running water and re-examine.

4. Transfer to alcoholic eosin (0.4% eosin in 70% ethanol) for 1 min.

5. Rinse in running water for 1 min.

6. Dehydrate through a series of graded alcohols (40%, 70%, 90%, 95%, and 3-4 changes of 100% alcohol), with 5 min in each.

7. Place in xylene for 30 min, then mount slides using a permanent mount (XAM, DPX).

fames W. Wilson and Christopher S. Potten Protocol 9. Continued

8. Allow slides to dry overnight, prior to microscopic examination of sections.

Gill's haematoxylin no. 2 (product code 6765007, Shandon Inc.) is the stain of choice in our laboratory as ¡t is relatively stable and gives reproducible and uniform staining. There are other, commomly used haematoxylins, including Ehrlich's, Meyer's, and Harris's. Ehrlich's stain requires two months to ripen prior to use, although it is very stable and gives excellent morphology. Meyer's stain is prepared with chloral hydrate, and Harris's with mercuric oxide, which make them unattractive for general laboratory use. Both the latter stains go off quickly compared with Gill's. More information regarding the applications of different haematoxylin stains can be obtained in: Theory and practice of histological hechniques (ed. J.D. Bancroft and A. Stevens), p. 107. Churchill Livingstone, Edinburgh (1990).

Protocol 9. Thionin blue


• Thionin blue (4 parts solution A and 1 part •

solution B: 8 ml glacial acetic acid, 18 ml 5

solution B)

M sodium hydroxide, made up to 100 ml

• 80% methanol

with deionized water

• solution A: 0.5 g thionin acetate (Sigma, •

100% ethanol

T7029) in 100 ml methanol (filtered)


1. Place the slides in thionin blue for 10 min.

2. Transfer to 80% methanol for 10 min.

3. Give the slides 10 dips in 95% ethanol. Repeat.

4. Transfer to absolute ethanol for 10 min.

5. Treat as for Protocol 8, steps 7 and 8.

Thionin may be reused many times before staining intensity is impaired. Methanol (80%) may

also be reused, until it becomes too discoloured.

2.2.2 Stains for fluorescence microscopy

Fluorescent nuclear counterstains are most appropriate for use with cultured cell systems, and when fluorescent detection methods are being used in parallel for immunohistochemistry. The most commonly used are 4',6-diamidino-2-phenylindole (DAPI), Hoechst 33258 and 33342, acridine orange, and propidium iodide. As with the stains used for light microscopy, the tissue must be hydrated prior to staining.

Hoechst 33258 stains non-apoptotic human and murine nuclei differentially. Human nuclei have a uniform, diffuse stain, whereas murine nuclei demonstrate several small, brightly staining bodies (2). Although there are no data to suggest that apoptotic cells from different species are stained differentially, this property of the Hoechst dye can be very useful when carrying out morphological assessment of xenografted human tissues in immune-deficient mice (3, 4), as it permits a distinction to be made between mouse and human tissue. One possible caveat with the use of Hoechst dyes is that they have been shown to induce apoptosis (5). However, this is only going to be a problem in unfixed tissue, and the length of time required to induce this effect (3 h) means that it should be irrelevant in all but exceptional circumstances.

Protocol 10. Fluorescent stains


Make up stains as follows:

• DAPI: 5 mg/ml stock in methanol. Prior to use, dilute 1:10000 in PBS, pH 7.4 (Sigma D9542).

• Hoechst 33258 and 33342: 100 n.g/ml stock in PBS, pH 7.4. Prior to use, dilute 1:10 in PBS (Sigma B2883 and B2261).


1. Incubate the cells/sections with stain for 3-5 min. For cell suspensions, resuspend the cells at a density of ~2 x 10® cells/ml and mix with an equal volume of dye.

2. Wash the cells/sections twice in PBS.

3. Mount using an aqueous-based mountant, with an anti-fade additive (Vectorshield, H1000; Vector Labs Inc.).

Acridine orange has been the dye of choice for many years for researchers studying developmental cell death in Drosophila (6-8). Acridine orange is a vital dye, i.e. it is excluded by viable cells. Staining of non-fixed, whole embryos with acridine orange allows the investigator to examine the spatial and temporal aspects of cell death in developing Drosophila embryos, using confocal fluorescence microscopy. Experimental details concerning this technique can be found in refs 6-8.

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