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Biotechnic & Histochemistry

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How Romanowsky stains work and why theyremain valuable — including a proposed universalRomanowsky staining mechanism and a rationaltroubleshooting scheme

RW Horobin

To cite this article: RW Horobin (2011) How Romanowsky stains work and why they remainvaluable — including a proposed universal Romanowsky staining mechanism and a rationaltroubleshooting scheme, Biotechnic & Histochemistry, 86:1, 36-51

To link to this article: http://dx.doi.org/10.3109/10520295.2010.515491

Published online: 14 Jan 2011.

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Correspondence: Richard W Horobin, School of Life Sciences,

The University of Glasgow, University Avenue, Glasgow G12

8QQ, Scotland, UK. E-mail: RichardWHorobin@tomcroy.co.uk

© 2011 The Biological Stain Commission

Biotechnic & Histochemistry 2011, 86(1): 36–51.

How Romanowsky stains work and why they remain valuable — including a proposed universal Romanowsky staining mechanism and a rational troubleshooting scheme

RW Horobin

School of Life Sciences, The University of Glasgow, University Avenue, Glasgow G12 8QQ, Scotland, UK

Abstract

An introduction to the nomenclature and concept of “ Romanowsky stains ” is followed by a brief account of the dyes involved and especially the crucial role of azure B and of the impurity of most commercial dye lots. Technical features of standardized and traditional Romanowsky stains are outlined, e.g., number and ratio of the acidic and basic dyes used, solvent effects, staining times, and fi xation effects. The peculiar advantages of Romanowsky staining are noted, namely, the polychromasia achieved in a technically simple manner with the potential for stain intensifi cation of “ the color purple. ” Accounts are provided of a variety of physicochemically relevant topics, namely, acidic and basic dyeing, peculiarities of acidic and basic dye mixtures, consequences of differential staining rates of different cell and tissue components and of dif-ferent dyes, the chemical signifi cance of “ the color purple, ” the substrate selectivity for purple color formation and its intensifi cation in situ due to a template effect, effects of resin embed-ding and prior fi xation. Based on these physicochemical phenomena, mechanisms for the various Romanowsky staining applications are outlined including for blood, marrow and cyto-logical smears; G-bands of chromosomes; microorganisms and other single-cell entities; and par-affi n and resin tissue sections. The common factors involved in these specifi c mechanisms are pulled together to generate a “ universal ” generic mechanism for these stains. Certain generic problems of Romanowsky stains are discussed including the instability of solutions of acidic dye – basic dye mixtures, the inherent heterogeneity of polychrome methylene blue, and the result-ing problems of standardization. Finally, a rational trouble-shooting scheme is appended.

Key words: AT-rich genome , bacteria , blood fi lm , carmine , G-banded chromosome , Giemsa , Leishman , malaria parasite , marrow smear , microorganism , paraffi n section , parasite , resin section , trouble-shooting , Wright

even its expansion, “ Romanowsky – Giemsa stain, ” really is.

The present article is a non-encyclopedic, non-historical review of the state of the art concern-ing staining mechanisms of Romanowsky stains. Major current application areas of these stains are addressed, including bacteriology, cytogenetics, cytology, hematology and parasitology as well as the staining of paraffi n and resin histology sec-tions. The relation of the staining mechanisms to the peculiar advantages of Romanowsky staining is discussed and a rational, i.e., staining mechanism-based, trouble-shooting guide is provided.

36 DOI:10.3109/10520295.2010.515491

The term, “ Romanowsky stain, ” is a generic des-cription of the azure B/polychromed methylene blue – eosin stains family, whose currently popular variants include the Giemsa, Leishman and Wright stains. Readers should see the historical account by Dr. K. Krafts in this journal issue to appreci-ate how partial the term Romanowsky stain, or

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How Romanowsky stains work 37

azures A and B, and that occurring with azure A is technically unsatisfactory owing to lack of color contrast with basophilic cytoplasms as illustrated by Wittekind (1986). The acidic dye, however, does not have to be eosin Y and other halogenated fl uo-resceins such as eosin B perform satisfactorily (Wit-tekind 1983). The structures of the dyes named in this review are shown in Fig. 1. Both azure B and eosin Y currently are available commercially in a satisfactorily pure state.

Since the earliest years (see Krafts, this issue), and still today, however, nearly all laboratory work-ers have used polychromed methylene blue as a source of azure B rather than the pure compound. Polychrome methylene blue is a mixture of bluish dyes, including azure B, produced from methylene blue by oxidative treatments such as exposure to atmospheric oxygen under alkaline conditions or acidic dichromate solution as reviewed by Lillie (1977). Exact compositions, and in particular the amount of azure B present, varies among manufac-turers and lots. This has been well and repeatedly documented (e.g., Roe et al. 1940, Nerenberg and Fischer 1963, Marshall et al. 1975).

So what are the characteristics of Romanowsky staining? The most obvious is polychromy. Mini-mally, this requires just two dyes applied from a single dye bath. In addition to red and blue stained structures, a striking “ purple ” (Wittekind) or “ magenta ” (Sumner) color also occurs in sev-eral structures (the “ Romanowsky-Giemsa effect ” ). The overall outcome is termed “ Romanowsky staining. ”

Nature of the dyes in Romanowsky stains

Early investigators, using spectroscopic analy-ses, argued for the importance of azure B (Lillie 1977). This view was confi rmed using dyes derived from specifi c syntheses, followed by chromato-graphic purifi cation; for a summary, see Wittekind et al. (1978). These workers demonstrated that the only dyes necessary for complete polychrome Romanowsky staining were the basic (cationic) dye azure B and the acid (anionic) dye, eosin Y. In fact, the only basic dyes giving the color purple are

N

S NN

CH3 CH3

CH2H3C

N

S NN

CH3 CH3

CH3H

N

S NNH H

CH3H3C

N

S NNH CH3

CH3H

N

S NNH H

CH3H

N

S NNH H

HH

N

S ON

CH3

H3C

N

S ONH

H3C

sym-dimethylthionine

methyleneblue azureB

azureA

eninoihtCeruza

methylenevioletBernsthen methythionoline

Cl Cl

Cl Cl

Cl Cl

O O

Br

O

Br

BrBrCOO

O O

Br

O

Br

NO2O2NCOO

2Na 2NaeosinY eosinB

Fig. 1. Structures of dyes discussed including azure B and eosin Y. The ionic species shown are those likely to predominate

under routine Romanowsky staining conditions. Only single resonance forms are shown.

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38 Biotechnic & Histochemistry 2011, 86(1): 36–51

particular, azure B content. Despite such techni-cal variation, it is striking how many laboratories are not carrying out Romanowsky staining in the precise sense, because their published works show cells with blue, not purple, nuclei. Indeed, many workers in reputable institutions are content to use such “ non-Romanowsky Romanowsky stain-ing ” as can be seen readily by keying “ Giemsa ” into “ Google images. ” Having made these general points, some comments on technical variations are pertinent.

Dyes of standardized and traditional stains

Only azure B plus various halogenated fl uoresceins produce the Romanowsky polychromy including the color purple. Other components of polychrome methylene blue fail to either give the color purple or provide contrast between nucleus and cyto-plasm. Methylene blue itself does not give purple nuclei, but does provide sky blue basophilic cyto-plasms. Consequently, two types of standardized Romanowsky stains have been developed.

The fi rst are two-dye mixtures of azure B plus eosin Y. One such example, using azure B isothio-cyanate and eosinic acid, was proposed by an expert panel (Wittekind et al. 1976) of the International Council for Standardization in Hematology (ICSH). Another two-dye stain used the more soluble azure B chloride plus the more widely available sodium eosinate (Marshall et al. 1981). The second type is a three-dye mixture, again containing azure B and eosin Y, but with methylene blue added (Marshall et al. 1975). The rationale was that when azure B is contaminated with other oxidation products, the cytoplasmic staining becomes grayer and less con-trasted with purple nuclei; however, this is com-pensated for by the blue staining dye. Note that successful staining with many polychrome meth-ylene blue-eosin mixtures, often with low azure B content, can thus be understood as inadvertently “ impure three-dye mixtures. ”

The ratio of the acid and basic dyes is signifi cant. Work with pure dyes showed that a wide range of molar ratios, up to 16:1 azure B:eosin, gave satis-factory staining of blood and marrow fi lms (Wit-tekind 1983). Use of azure eosinate precipitates, however, with a molar ratio around 2:1, are not satisfactory. Consequently, as use of polychrome methylene blue makes precise control of the azure B-eosin ratio unachievable, it is understandable that staining recipes since Giemsa ’ s fi rst Romanowsky formulas have provided an excess of cationic dye over anionic eosin. The pure dye experiments also demonstrated that for successful histological

The polychroming processes result in demethy-lation and eventually deamination of the tetram-ethylated methylene blue parent compound. Most resulting dyes, including azure B, are expected to be cationic under Romanowsky staining conditions and all compounds retain the fused tricyclic het-erocyclic ring structure. There is some variation of hydrophilic-lipophilic character. These differences are summarized numerically in Table 1 using the parameters: electric charge (Z), largest conjugated fragment (LCF), and log P, respectively; the latter parameters were estimated as described by Horobin (2001).

Halogenated-fl uorescein acid dyes that have been used or investigated for use in Romanowsky stains typically are dianionic under Romanowsky staining conditions. The sizes of the conjugated sys-tems and lipophilicities vary slightly as summarized in Table 1. Analytical studies have shown that many commercial lots comprise mixtures of the nominal dye with incompletely halogenated impurities that are of lower lipophilicity (Marshall and Lewis 1974, Marshall 1976).

Technical features of Romanowsky staining systems

Protocols for preparing and using Romanowsky stains show considerable variation, even within a single application area. These differences probably arose from efforts to optimize staining when using dye lots that varied greatly in composition and, in

Table 1. Some physicochemical features of dyes menti-

oned in this review

Dye properties1

Dye name (CI number) Z LCF log P

Methylene blue (52015) 1� 18 �0.9Azure B (52010) 1� 18 �1.2Symmetrical dimethylthionine 1� 18 �1.6Azure A (52005) 1� 18 �1.5Azure C (52002) 1� 18 �1.9Thionine (52000) 1� 18 �2.2Methylene violet Bernsthen

(52041)

0 18 �2.8

Methylthionoline 0 18 �1.9Eosin B (45400) �2 18 �0.6Eosin Y (45380) �2 18 �0.3

Dyes typically derived from methylene blue are listed in order of

decreasing methylation/amination. Acid (anionic) dyes are listed

in order of increasing halogenation.1Z, the nominal electric charge; LCF, the largest conjugated

fragment; log P, the logarithm of the octanol–water partition

coefficient.

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How Romanowsky stains work 39

Although this does not preclude the Romanowsky effect, such preparations require longer staining times and/or higher dye concentrations.

Exploitation of characteristic features of Romanowsky staining

Romanowsky staining is characterized by polychro-masia, particularly the formation of a distinctive purple coloration in chromatin and the granules of neutrophils and platelets and, in tissue sections, collagenous connective tissue and mucin. This polychromasia is used to distinguish populations of morphologically related cells, especially for iden-tifi cation of the cells in blood and marrow smears and cytological preparations.

Another key feature is that considerable inten-sifi cation of this purple staining, without over-staining of blue and red colors, can be achieved in a technically straightforward way, e.g., by extending staining time (Boon and Drijver 1986; Plate 6) or by increasing dye concentrations. Such intensifi cation facilitates detection of small cells and cellular components near the limit of normal optical resolution. Examples include chromosome G-bands, neutrophil and platelet granules, and bacterial spores.

Not only can such polychrome staining with intensifi cation be obtained in a technically simple way (two dyes and a single staining solution), but it can be achieved on fi lms, smears, dabs etc., or paraffi n or resin sections. A variety of fi xatives can be used, although modifi cations of the stain-ing protocol may be needed. For a dramatic visual demonstration of the colored detail to be seen in Romanowsky stained resin embedded histology sections, see Wittekind (1991; Figs. 4 and 5).

Some physicochemical background

The physicochemical phenomena underlying the mechanisms of Romanowsky stains are considered here in a general way. Not all phenomena discussed apply to all Romanowsky staining methods and the staining mechanisms of particular applications are discussed later.

Acidic and basic dyeing mechanisms

A simple way to conceptualize acid and basic dye-ing is to view it as an ion-exchange process (Fig. 2). Colored anions, e.g., those of eosin Y, exchange with the mobile inorganic anions to neutralize the fi xed positive ions of proteins. Colored cations, e.g., those

application, the azure B-eosin molar ratio had to be much lower, in the range of 6 – 3:1 (Wittekind 1983).

Solvent effects

The “ Marshall ” standardized methods noted above used azure B chloride and the traditional methano-lic stock solutions for both two- and three-dye vari-ants. The ICSH (or “ Wittekind ” ) variant, however, used the less soluble isocyanate, which required the superior, albeit more potentially hazardous solvent, dimethylsulfoxide. Both solvents, when diluted with buffer in the working solutions, permit generation of the color purple. Dimethylsulfoxide, however, provides a longer working solution usable lifetime than methanol. Also, the color purple is unstable in the presence of high concentrations of the lower alcohols, methanol and ethanol, as reported at least as early as 1944 (Lillie 1944 ). This is one reason why published histological applications of the Romanowsky stain often failed to generate purple cell nuclei; they still used the traditional alcoholic dehydrating agents (e.g., Cramer et al. 1973). These problems can be avoided by using isobutanol or iso-propanol as dehydrating agents (Wittekind 1983). The color purple also is removed quickly by dilute acetic acid.

In fact, the effects of pH are marked regardless of the nature of the acid involved. The relation of pH to acidophilic – basophilic balance, and so to overall color, are well known. Erythrocytes, for example, become blue-green if the pH is too high. The occurrence of the color purple, however, also is pH sensitive. After methanolic fi xation, the color often most is intense at pH 7-8, diminishes at pH 6-5, and is completely inhibited at pH 3.5.

Staining times

Crucially, formation of the color purple follows an initial blue staining. Preparations do not merely become more intensely stained, they also become more purple as staining times are lengthened. Optimal staining times are related to specimen thickness, e.g., cell monolayers such as blood smears show purple colors sooner than tissue sections do.

Fixation

Fixation also infl uences uptake of acid and basic dyes, and formation of the color purple. Aldehyde fi xatives, e.g., for histology, increase basophilia.

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40 Biotechnic & Histochemistry 2011, 86(1): 36–51

Cl ¯

Acid dyeing of cationic biopolymer by eosin Y

Na+

Na+

Na+

Na+

Azure B

2 Na+ 2 Cl¯

eg protein at low pH

Cl¯

Cl¯

Cl¯

Eosin Y

eg DNA or RNA

Cl¯Na+

Azure B

Na+

Na+

Na+ Cl¯

Basic dyeing of anionic biopolymer by azure B

Eosin Y

Fig. 2. Acid and basic dyeing seen as ion exchange processes. Mobile counterions, shown as Na� and Clˉ, in actuality

depend on the species present in the staining solution.

of azure B, exchange with the mobile inorganic cations to neutralize the fi xed negative ions on such biopolymers as DNA, RNA or acidic glycosamino-glycans (GAGs). Such a model accounts for both affi nity, which as described is entirely an entropy driven process, and selectivity, which in this model depends only on the fi xed charges on the biopoly-mers, given that electrical neutrality of tissue and dye bath must be maintained.

Such an ion exchange picture also can explain the effects of pH on dye uptake. Consider cellular biopolymers carrying acidic and basic substitu-ents whose pKa values fall within the pH range of Romanowsky stains. Their electrical charges vary with pH, and as the charges vary, so will the ion exchange possibilities as illustrated in Fig. 3.

But dyes are not merely ions. They have large aromatic systems that provide regions of both hydrophobicity (lipophilicity) and polarizable electrons. So acid and basic dyeing is not just ion exchange, but typically involves other phenom-ena that contribute to dye – biopolymer affi nity. Consider three possibilities. First, there may be dye – biopolymer attractive forces such as those occurring between the polarizable aromatic rings of dyes and of the bases of nucleic acids. Second, the hydrophobic effects that arise in aqueous milieu when both dye and biopolymer have hydrophobic regions can come into contact. Both of these pro-cesses are involved in dye binding that involves the

insertion of cationic dyes between the base pairs of the double helix of DNA ( “ intercalation ” ). A third contribution to dye – biopolymer affi nity may be formation of dye – dye complexes such as occurs with metachromasia, e.g., the staining of mast cell granules by many basic dyes. The formation of the color purple, as described below, probably results from the formation of a dye complex of a somewhat different type.

Acid dye-basic dye mixtures

After mixing an acid (anionic) and basic (cationic) dye in a single solution, one possible outcome is the formation of an insoluble salt. All of the cat-ionic dyes present in polychrome methylene blue do precipitate from aqueous solutions upon addi-tion of eosin; the material precipitated contains the basic dye and acid dye in approximately 2:1 molar ratio, which corresponds to neutral salts. Water miscible organic solvents, such as metha-nol, dimethylsulfoxide or glycerol, reduce hydro-phobic effects between the hydrophobic species in aqueous solutions, such as the Romanowsky staining solutions, which reduces precipitation.

The aqueous solutions mixed to produce such azure eosinates contain dye at high concentrations and precipitation is rapid. Dye concentrations in staining solutions are lower and precipitation is slower. Nevertheless, precipitation of azure

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How Romanowsky stains work 41

Rate effects of biological origin

Some basophilic molecules such as acid GAGs found in the granules (lysosomes) of neutrophils are highly swollen in aqueous media, which permits rapid dye diffusion. Other basophilic biopolymers are intimately associated with proteins and it is the overall permeability of the nucleoprotein complex that controls dye entry. Thus the RNA in the dense nucleolar or cytoplasmic ribosomes stains slowly with a basic dye, whereas the DNA in chromatin stains faster (Goldstein 1963a and b).

Even within cell monolayers, the thickness of a cell infl uences stain penetration; thus, “ in general, parasites require longer staining than blood ele-ments ” (Clark 1973) — and the cells of parasites are in general larger than those of human blood. In particular, the morphology can infl uence rate of formation of the color purple. This occurs fi rst in the thinner portions of the preparation (Wittekind 1979, Horobin et al. 1989). It is intriguing that similar rate effects also have been reported in cell-free DNA fi lms of varying thickness (Friedrich et al. 1990).

When slabs of coherent biological material are used, i.e., tissue sections, the situation becomes more complex. The porosity of some acidophilic entities, typically protein or protein-rich, also has been assessed using the refractive index as a mea-sure. Relative porosity fell in the sequence: colla-gen fi bers � smooth muscle cytoplasm � pancreatic exocrine secretion granules � red blood cells (Gold-stein 1965). In keeping with a diffusion rate control model, staining rates also fall into this sequence (Horobin 1974, cited in Horobin 1982).

Rate effects technically produced

Morphology is infl uential in this case also. Thus marrow smears stain faster, both overall and in terms of how soon the color purple is seen, than do sections of bone marrow (Wittekind 1983, Wittekind et al. 1984). Moreover, as considered in more detail later, resin sections stain more slowly than paraffi n sections. By analogy, in the thicker portions of cell smears, the nuclei stain blue, not purple (Boon and Drijver 1986). Formaldehyde fi xation also infl uences (retards) the rate of formation of the color purple compared to the rate following methanolic fi xation (e.g., Wittekind 1983). As described in more detail in the “ Fixation ” section below, this is probably due to the loss of free amino groups in proteins, which reduces the rate of uptake of eosin.

Finally, note that dye molecules of different sizes tend to diffuse through specimens at differ-ent rates as demonstrated dramatically a century

eosinates onto slides or within the pipework of autostainers can be a signifi cant technical problem (Gregory and Maher 2009). The molar ratio of the basic and acid dyes in the precipitate is approxi-mately 2:1 and the presence of excess acid or, especially, basic dye tends to reduce precipitation (Lillie 1977).

Consequences of differential staining rates

Two dye binding sites with equal affi nity for a dye can nevertheless be colored differentially if they stain at different rates. In progressive staining, the site that binds dye fastest becomes selectively stained after short periods of staining. Regressive staining, which involves differentiation of “ over-stained ” preparations, also fi ts this picture. If a preparation is over-stained, then de-stained, sites slowest to de-stain retain dye longest and so become selectively colored. Note that similar outcomes could follow if some cellular targets have a higher density of bind-ing sites than other targets with the same affi nity for dye. For reasons of simplicity, however, this is not stated repeatedly.

Rate control of Romanowsky staining patterns has diverse mechanisms, and several are discussed below. For convenience, such effects are catego-rized as either “ of biological origin ” or “ technically produced. ” This topic has been discussed at length elsewhere, albeit without reference to Romanowsky stains (Horobin 1982).

Na+

Cl¯

Cl¯

Generic zwitterionic

protein in a neutral

dyebath

Na+¯OOC

+H3N

NH3+

¯OOC

HOOC

+H3N

NH3+

HOOC

¯OOC

H2N

NH2

¯OOC

Protein in an

acidic dyebath

Protein in an

alkaline dyebath

No

exchangeable

mobile ions

Exchangeable

cations, so

basophilic

Exchangeable

anions, so

acidophilic

Fig. 3. Effect of changes in pH on the ionic status of

biopolymers, thus on uptake of acid and basic dyes.

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42 Biotechnic & Histochemistry 2011, 86(1): 36–51

structure or remove the water completely such as when lower alcohols are used for dehydration. Zimmermann ’ s group carried out a detailed spec-troscopic study of a DNA, azure B, eosin Y model system, and they also favored hydrophobic effects as a major contributor to the dye-dye binding involved in formation of the color purple, although in their picture the DNA formed part of the com-plex (Friedrich et al. 1990). Because the conditions of the latter study were markedly different from those of practical Romanowsky staining and because the color purple arises at sites not containing DNA, the signifi cance of the study is not clear.

Finally, note that additional azure B-eosin complexes may exist. For example, in strongly aci-dophilic substrates, such as eosinophil granules, an initial uptake of eosin is followed by binding of azure B, which gives a “ red-brown ” coloration that might be due to a different complex (Wittekind 1991). Yet another possible case arises with the “ car-mine ” or “ red ” nuclear coloration of Romanowsky stained malarial parasites (for an historical account, see Baker 1958). Regarding the latter, it is worth not-ing that Plasmodium falciparum DNA contains the unusual feature of “ super AT-rich regions ” in its genome (Woynarowski et al. 2007), and, of course, AT-rich DNA is signifi cant in Romanowsky staining of “ banded ” chromosomes.

Substrate selectivity for formation of the

color purple

Whatever the origin of the color purple, whether it is a complex or merely an azure eosinate salt, its sub-strate-selective formation constitutes a distinct prob-lem. Why, for example, is DNA-rich chromatin purple, but not the adjacent RNA-rich nucleoli? Although such questions cannot yet be answered defi nitively, contributing possibilities can be proposed.

The color purple requires the presence of both azure B and eosin in the same region of the specimen. Therefore, both affi nities for and rates of uptake into a substrate could control selectiv-ity. Rate effects certainly play a part. Chromatin, for example, fi rst stains blue, then purple. At the stage of staining when chromatin becomes purple, basophilic cytoplasms still are blue, but if staining times are prolonged suffi ciently, these cytoplasms also become purple. With extremely extended times, many other substrates become purple as well (Horobin and Walter 1987). The causes of such rate effects are varied and are considered as appropri-ate for each of the staining applications, e.g., blood smears, G-banding, resin section staining etc., dis-cussed below.

ago by Mann (1902). Large dyes typically stain slower, perhaps a direct effect of particle size (Seki 1932), and perhaps also because tissue affi nity of large dyes is often greater due to their larger con-jugated systems, which favor dye-tissue van der Waals attractions (Horobin 1982). Taking the rela-tive molecular masses of the dyes as an indication of size, eosin Y is almost three times the size of azure B, so the latter may be expected to stain con-siderably faster.

The chemical meaning of the color purple

The color purple produced within biological speci-mens following azure B – eosin staining has an absorption maximum near 550 nm. This usually is regarded as resulting from formation of an azure B – eosin or azure B – eosin – biological substrate com-plex (Wittekind 1985, Friedrich et al. 1990, respec-tively). An alternative view, which regarded this absorption as overlapping of absorption peaks of constituent dyes (Galbraith et al. 1980), cannot be correct for many tissue sites, e.g., chromatin, mucus and neutrophil granules, which have no signifi -cant affi nity for eosin in the absence of azure B as pointed out for chromosomes by Sumner (1990). This absorption does not arise in solution, rather only in some biological matrix or at an interface such as the surface of a solution (Sumner and Evans 1973, Wittekind 1983). Examples of biological sites where the color purple arises include chromatin and gran-ules of neutrophils and platelets in blood smears and tissue sections, in the bands of G-banded chro-mosomes, and in collagenous connective tissue and mucus granules of tissue sections.

Although most investigators have regarded for-mation of the color purple as resulting from com-plex formation, the specifi c interactions between the dyes, and perhaps also the biosubstrate, still are debated. An early suggestion proposed formation of an acceptor/donor azure B-eosin charge transfer complex (Zanker 1981). This concept was adopted by Wittekind (1985) who noted the possible role of hydrogen bonding between the N – H groups of azure B and eosin for facilitating electron transfer. The latter feature would account for lack of purple staining when methylene blue is used with eosin Y and its occurrence when azure A, with two N – H groups, is used; however, the question of why less methylated dyes, e.g., thionine, do not show the Romanowsky effect remains unresolved.

Wittekind (1985) also considered that azure B and eosin are associated in part due to hydrophobic effects, which would explain the lability of the com-plex when exposed to solvents that disrupt water

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How Romanowsky stains work 43

sections than in smear preparations, however, but that certain regions stain very much slower.

This markedly non-uniform staining inhibition is due to non-uniform resin infi ltration. This in turn is due to the viscous and, even when water miscible, slightly hydrophobic character of the infi ltrating resin monomers. These properties cause resin to be largely absent from dense structures, e.g., chromo-somes, elastic fi bers, nucleoli, red blood cells, and from hydrophilic structures, e.g., collagen fi bers, GAG-rich lysosomes and mucus granules. Other regions, such as nuclear chromatin and the cyto-plasmic matrix, though not the small, dense ribo-somes, are infi ltrated well by the resin (Horobin 1983, Gerrits et al. 1990).

Staining of resin sections consequently displays evidence of site-selective staining inhibition by resin superimposed upon the rate and affi nity effects owing to the biological substrates. It may be noted that resin inhibition can arise from both a reduction of free volume available for dye diffusion and dye-polymer affi nity as when a lipophilic dye stains the resin (Horobin et al. 1992).

What this implies for Romanowsky staining is straightforward. A small, weakly hydrophilic dye such as azure B can access resin sections with only slight inhibition of entry. Access by the three times larger and weakly lipophilic eosin Y, however, is inhibited signifi cantly in resin-containing structures such as chromatin and the cytoplasmic matrix. As an example of the resulting staining outcomes, con-sider cytoplasmic basophilia, namely, the staining of ribosomal RNA. This staining is blue In smears, because the dense ribosomes impede entry of the eosin Y. In resin sections, however, because ribo-somes are dense, they are, unlike the cytoplasmic matrix, poorly infi ltrated. Because routine staining times for resin sections are necessarily longer or staining temperatures higher than for smears, eosin has relatively better access to ribosomes than to matrix. Consequently, ribosomes that are exposed on the surface of the section stain purple and this reversal of the “ normal ” staining pattern has been observed in glycol methacrylate sections of bone marrow (Horobin and Boon 1988).

Chemical and physical effects of fi xation

Changing the fi xative used could infl uence Romanowsky staining in two ways. First, chemically, e.g., aldehyde fi xatives such as formaldehyde react with tissue amino groups, which results in decreased acidophilia (Horobin 1982). This has been observed to lead to preparations with an overall bluer color-ation (Wittekind 1983). Different fi xatives, however,

Intensifi cation of the color purple

by a template effect

Almost a century after the introduction of the poly-chrome methylene blue-eosin staining system, an explanation was offered for the stain ’ s character-istic of intense purple staining of extremely small objects such as the submicrometer lysosomes of neutrophils or the tiny G-bands of chromosomes. Sumner (1980), who was concerned with the lat-ter structures, proposed that because the “ levels of staining of … nuclei and chromosomes with Giemsa and similar mixtures are far greater than those nor-mally attained with … methylene blue … perhaps Giemsa is not dyeing chromosomes in the normal sense, but is in fact being precipitated preferentially in them … the initial stages of Giemsa staining involve ionic binding of thiazines alone … A second process in staining would then be the binding of thiazines to eosin to form a precipitate, thereby free-ing the reactive groups on the DNA to bind more thiazine. ” Therefore, this involves a DNA-induced co-precipitation, which will be termed here a tem-plate effect and which proceeds without reaching equilibrium. In the context of staining blood smears and other complex tissues, one can, in fact, replace Sumner ’ s term “ DNA ” with other anionic biopoly-mers, e.g., hyaluronic acid in the lysosomes of neu-trophils and platelets. Such a generalized template effect mechanism is illustrated in Fig. 4, where the precise chemical nature of the azure B-eosin com-plex is not specifi ed.

To appreciate the shift in mechanistic perspective represented by the template intensifi cation effect, consider the views of two well known fi gures in his-tochemistry and biotechnique. John R. Baker, in his classic monograph, considered that an azure – eosin complex was unlikely to transfer from solution onto DNA; however, he had no suggestions regarding what did happen (Baker 1958). Moreover, Ralph D. Lillie, an investigator with considerable experimen-tal experience in the fi eld, argued as late as 1977 that staining of the characteristic granules of neutrophils refl ected their affi nity for an azure B – eosin compound (Lillie 1977).

Effects of resin embedding media

Resin (plastic) embedding media have an inhibitory infl uence on dye uptake when sections are stained with the resin still present. For example, staining rates of resin sections of bone marrow are slower than those of marrow smears unless the stain is heated (Horobin and Boon 1988). It is not merely that many structures stain somewhat slower in resin

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44 Biotechnic & Histochemistry 2011, 86(1): 36–51

also alter the native morphology in different ways. Thus the classic alcohol and acidic alcohol fi xations give rise to “ shattered ” preparations at a micro-scopic level, whereas aldehyde fi xation produces a more coherent structure (Horobin 1982).

Empirically, alcoholic fi xatives produce speci-mens that tend to stain faster than formalin fi xed preparations. Where two dyes of different sizes are used simultaneously, faster staining typically favors entry of the larger dye. Either of these mechanisms apparently could underlie the observed effect of for-malin fi xation, namely, that the color purple fails to occur in nuclear chromatin, which instead stains blue. If staining times are lengthened or eosin bind-ing is enhanced by lowering the pH of the dye bath (Lillie 1977), the purple staining is restored. It has been established, however, by comparing prepara-tions fi xed in alcohol followed by formaldehyde to those fi xed in formaldehyde followed by alco-hol that the effect is due to the chemical changes (Horobin and Walter 1987).

Staining mechanisms of Romanowsky and related stains

While all methods use the typical Romanowsky staining system based on azure B, usually in the form of polychrome methylene blue, and eosin Y,

the procedures used for preparing specimens, the morphology of the material so prepared, and the staining procedures vary markedly. These differ-ences are outlined below.

Specimen pretreatment

Fixation of cell smears for hematology or cytol-ogy routinely use methanol. Chromosome spreads usually are prepared using methanol-acetic acid. Tissue prepared for sectioning, whether in par-affi n or resin, usually has been fi xed in formalin, although other fi xatives occasionally have been used. Chromosome banding uses various pretreatments of the fi xed chromosome spreads prior to staining in addition, e.g., with a solution of a hot neutral aqueous buffer, a surfactant, or a protease.

Morphology of the prepared specimen

Hematological or cytological preparations are cell monolayers produced as brushings, dabs, smears, spins etc. For diagnosis of protozoal parasitic infections, thick, i.e., many cells thick, fi lms some-times are used. Cytogenetic chromosome spreads are monolayers of chromosomes. Paraffi n sections and sections cut from methylmethacrylate resin blocks are stained after removing the embedding

Large,

slow diffusing

Azure B

Na+

Na+

Na+

Eosin Y

polyanion

Cl¯

Azure B

Na+

Na+

Na+

small,

fast diffusing

Azure B

insoluble azure-eosin

complex deposited

nearpolyanion

Azure B

Eosin Y

Azure B

2 Na+ 2 Cl¯

Fig. 4. Intensifi cation of the Romanowsky effect with continued staining seen as a template effect. Note that the precise

relation of azure B to eosin is not defi ned.

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How Romanowsky stains work 45

swollen character of GAG-rich lysosomes in partially aqueous media. Failure to stain pur-ple, as with the lysosomes of monocytes, also is explained by this model, because monocyte lysosomes lack signifi cant GAG content (Fedorko and Morse 1965).

3. The easy visibility of the purple staining of tiny granules, which occurs only if the batch of Romanowsky stain used contains suffi cient azure B, is due to intensifi cation owing to a template effect.

Banding of chromosome spreads

Several distinct phenomena need explanation. Why do the chromosomes stain, and in particular, why do they stain purple? Why is it that only certain “ bands ” stain strongly? And how is it that these tiny structures are stained so intensely as to be notice-able? To address these questions, four aspects of mechanism are discussed below.

1. The fi rst step involves basic dyeing of DNA by azure B.

2. The blue staining subsequently is converted to purple as an azure B – eosin complex is formed. Note that if the chromosomes are covered with incompletely dispersed cytoplasmic material, the entry of the larger eosin ions is retarded and this step is inhibited (Sumner 1990).

3. The question of selectivity is more conten-tious. Various suggestions have been made to explain why dye binding favors certain lim-ited regions of the chromosomes, the G-bands. For example, it is known that G-bands corre-spond to AT-rich regions. Because AT base pairs interact less strongly than GC base pairs, this location would be expected to favor inser-tion of an intercalating dye such as azure B, and indeed in an azure B – eosin Y – DNA model system, the azure B was observed to be intercalated (Friedrich et al. 1990). This obvi-ously is not the only, or perhaps not even the overriding, phenomenon, however, because strongly stained G-bands occur only following various pretreatments. These are varied, e.g., proteases, detergents, hot buffer, and it is not apparent what the common features are. The subject of selectivity is still a topic of lively debate; for a recent contribution see Ushiki and Hoshi (2008). Unfortunately, the view Sumner espoused in 1990, “ we still lack a sat-isfactory hypothesis, ” remains the case today, as he confi rmed to the author recently (Sum-ner 2008, personal communication).

media using suitable solvents. Glycol methacrylate sections are stained with the resin in situ; these are the eponymous resin sections. Sections are typi-cally 0.5 – 5.0 μ m thick with the resin sections falling at the thin end of this range.

Staining conditions

Blood and marrow monolayers, e.g., smears, are stained for up to 30 min at room temperature. Stain-ing times for chromosome spreads are much shorter, while those for sections are longer. The staining of smears and sections, and in particular resin sections, has on occasion been accelerated considerably by heating the solutions, sometimes with a microwave oven (Boon and Kok 1987). Section staining some-times uses regressive procedures with differentia-tion in aqueous acetic acid or more exotic mixtures (Wittekind et al. 1991). To retain the color purple, however, contact with acetic acid differentiation solutions must be minimized, and dehydration must avoid use of the lower alcohols and instead use air drying or solvents such as butanol or isopropanol.

Mechanistic summaries of each general method-ology are given below.

Blood, bone marrow and cytological smears

Explanations for development of the full Romanowsky polychromasia will be considered below. The specifi c issues are: why do any dyes bind, and in particular, why do some substrates stain red, some blue and some purple? In addition, why are certain tiny granules in neutrophils and platelets stained so strongly as to be readily notice-able? To deal with these questions three aspects of mechanism are relevant as follows.

1. Acid and basic dyeing occurs in sites rich in polycations and polyanions, respectively. This, in the fi nal stage of the staining, gives rise to blue staining of ribosome-rich ( “ basophilic ” ) cytoplasms and nucleoli and red staining of eosinophil granules and red blood cells.

2. An additional color, purple, is produced in a limited number of relatively fast staining sites, notably chromatin and certain granules in neutrophils and platelets. Such rate-controlled selectivity involves transformation of initial azure B staining into an azure B-eosin complex as illustrated in the color plates in Boon and Drijver (1986). The faster staining is due to DNA-rich chromatin, which has a more porous structure than RNA-rich ribosomes, and to the

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46 Biotechnic & Histochemistry 2011, 86(1): 36–51

selectivity specifi c to the demonstration of microorganisms is the use of various pre-treat-ments prior to staining. Thus, if the parasite load is low, the sample size can be increased by using thick blood fi lms. In this case, how-ever, the parasites may be obscured by the mass of red blood cells, which are therefore lysed either osmotically or enzymatically.

Three additional topics requiring

brief mention

1. Romanowsky stains also are widely used with additional types of cellular monolayers, nota-bly cytological specimens, e.g., pleural fl uid smears or lymph node aspirates. Another example is the staining of spermatozoa, both of humans and many other mammals, to assess male reproductive status. Mechanistically, how-ever, such cases are analogous to the blood smears discussed above.

2. Multicellular parasites, e.g., tapeworms, also may be Romanowsky stained. As this is done within tissue sections, however, the process is mechanistically unremarkable; see below.

3. The phenomenon of the “ carmine ” or “ red pur-ple ” nuclear coloration of some Romanowsky stained parasites was discussed above.

Tissue sections: paraffin

The full range of Romanowsky staining polychro-masia, including the color purple, can be achieved with paraffi n sections. The technical requirements are that suffi cient azure B is present in the stain, that the staining time is long enough, and that excessive exposure to solvents such as acetic acid during dif-ferentiation or lower alcohols during dehydration be avoided. Given these circumstances, the staining is not particularly fi xative-dependent.

Some of the questions requiring mechanistic explanation are by now familiar. Why do the acidic and basic dyes bind, and why is the color purple generated? In particular, why do tissue elements such as collagen fi bers, cartilage matrix and some mucins stain purple? The longer staining times and use of formaldehyde as a fi xative also call for dis-cussion. The mechanistic factors needed to explain these are listed below.

1. Acid and basic dyeing occurs in sites rich in polycations and polyanions, respectively. This is the cause of the red staining of muscle cyto-plasm and of eosinophil granules and red blood cells; this accounts also for the blue

4. The visibility of the tiny purple stained G-bands is due to a template intensifi cation effect.

Staining of microorganisms and other

single cell entities

Microorganisms, whether bacterial, protozoal or fun-gal, sometimes are stained as monolayers (smears, fi lms or dabs) and sometimes within tissue sections. We are concerned here only with the mechanisms underlying selective visualization of the pathogens; the staining of other tissue elements in sections is discussed in a later section. The problems requiring mechanistic explanation are as follows. Why do the pathogens stain, and in particular, why stain pur-ple? How is it that these often tiny cells are stained suffi ciently strongly to be noticed? These questions and their implied extensions, are addressed below.

1. All the target organisms contain DNA, either in nucleoids of bacteria or in chromatin of eukaryotic cells. The fi rst step is basic dyeing of DNA by azure B.

2. This dye then interacts with eosin to yield the purple azure B – eosin complex. Note that prep-arations discussed in this section are, for vari-ous reasons, often slow staining. Thus, although bacterial cells are tiny, they usually are very dense; within a smear, the cells of parasites often are larger and thicker than white blood cells. In any event, it is common to use thick fi lms of blood for diagnosis of protozoal para-sites in parasitological samples. The effects of such rate limiting factors were discussed above. The technical consequence of this is shown by the commonplace use of diffusion-enhancing procedures such as extended staining times (even overnight for parasitic cells within paraf-fi n sections; Bartlett 2008) or heated staining solutions (for an illustration of the benefi ts of which, see Boon and Kok 1987).

3. The visibility of tiny purple stained microor-ganisms is due to the template intensifi cation effect described above.

4. Selectivity of staining calls for comment. While intense staining of morphologically discord-ant organisms is the primary feature for iden-tifi cation, the mechanisms of two additional tactics should be noted. The fi rst is differentia-tion with solvents, such as dilute acetic acid or isopropanol, of slow staining specimens subjected to extended or heated staining. The mechanism of such destaining has been discussed above. An additional source of

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How Romanowsky stains work 47

Romanowsky staining: is there a universal mechanism?

The mechanisms of the major applications of Romanowsky stains in modern biomedicine have been sketched. It is obvious that some factors such as specimen thickness, proportion of AT-bases in DNA, and occlusion by resin embedding media, have infl uence only in particular staining systems. There is no single, all-inclusive mechanism. Nev-ertheless, there are features that are important for all the staining systems described, namely, initial acidic and basic dyeing, formation of a purple azure B – eosin complex in a rate-controlled manner, and intensifi cation of the purple staining due to a tem-plate effect. These factors do constitute a “ universal mechanism ” of sorts, common to all methods as set out diagrammatically in Fig. 5.

Romanowsky staining problems and a staining mechanism-based trouble-shooting scheme

Certain problems with Romanowsky stains, namely, the unstable nature of a working Romanowsky staining solution and the heterogeneous nature of polychrome methylene blue, are inherent to the routine Romanowsky procedures and so are dis-cussed fi rst, followed by comments on the problems of stain standardization. Once again, the account given here refers back to the discussion of general physicochemical principles.

Inherent instability of Romanowsky stains

As discussed earlier, as soon as the substantially aqueous staining solutions are prepared, precipi-tation of material approximating azure- eosinates commences. The amount of precipitation tends to be minimized by factors such as minimal stand-ing time before staining, low dye concentration and low temperature, presence of excess acidic or especially basic dye, presence of larger proportions of organic solvent; dimethylsulfoxide and glycerol are particularly effective inhibitors of precipitation. Precipitation onto slides or within the pipework of autostainers can constitute a signifi cant technical problem.

Inherent dye impurity

Dye lots labeled “ azure B, ” even when purchased from reputable vendors, may contain some uncer-tain proportion of the named dye, but essentially

staining of ribosome-rich ( “ basophilic ” ) cyto-plasms, e.g., of secretory cells.

2. The color purple is seen in chromatin and the granules of neutrophils. This rate-controlled selectivity is described above and involves transformation of initial azure B staining into an azure B-eosin complex. Because the color purple is unstable in the presence of lower alcohols, dehydration through ethanol must be brief or be replaced by air drying or use of agents such as butanol.

3. Rate effects also are seen. Paraffi n sections are thicker and more coherent, especially if for-maldehyde fi xed, than methanol fi xed cell monolayers. Consequently, such sections require extended staining times: “ 90 min are recommended for obtaining the full polychro-matic staining pattern ” (Wittekind 1991). Such staining can be usefully differentiated, tradi-tionally with aqueous dilute acetic acid, but also with media such as isopropanol contain-ing tannic acid.

4. The occurrence of the color purple in tissue elements such as collagen fi bers, cartilage matrix and mucus granules also can be explained by a rate control mechanism. In par-affi n sections, probably due to the swollen character of the plentiful GAGs found at all these sites, these are relatively rapidly staining structures.

5. Inhibition, although not inevitable elimina-tion, of formation of the azure B-eosin com-plex by formaldehyde fi xation is probably a chemical effect.

Tissue sections: resin

The full range of Romanowsky staining poly-chromasia including the color purple occurs; long staining times again are required and over-staining followed by differentiation can be used; fi xation in formaldehyde is possible if suitable modifi cations to the protocols are made. So far, the phenomena and the mechanistic explanations are the same as for paraffi n sections and the details do not call for repetition.

An apparently novel phenomenon, however, the effective reverse color coding of nuclear chro-matin and basophilic cytoplasms has been reported (Horobin and Boon 1988). The appearance of blue nuclei and purple cytoplasms in lymphocytes, for example, is due to a combination of differential resin occlusion effects plus use of formaldehyde fi xation. Taken together, these phenomena reverse some of the rate effects seen in smears.

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48 Biotechnic & Histochemistry 2011, 86(1): 36–51

are relabeled polychrome methylene blue as discussed earlier. A few vendors have supplied azure B in a fairly pure state, but because this is con-siderably more expensive than azure B-containing polychrome methylene blue, such material is not likely to be used for the preparation of stains pur-chased by routine diagnostic laboratories and even “ pure ” lots can be less pure than claimed (Marshall 1979). Because the concentration of the key ingredi-ent, azure B, consequently varies markedly among lots, the intensity of the Romanowsky effect also is variable.

Standardization of Romanowsky stains

Presumably because of these phenomena, the color balance and appearance of Romanowsky stained preparations acceptable to different laboratories also is varied. This may be appreciated most sim-ply by inspecting the images of blood cells in dif-ferent hematology atlases. Preferences also have been investigated systematically by comparing the opinions of experienced working hematologists concerning the acceptability of various standard-ized formulations (Marshall et al. 1978).

Despite these problems, Romanowsky staining can be standardized to varying degrees. If using polychrome methylene blue, stains based on dye lots certifi ed by the Biological Stain Commission are recommended. Such reagents are on average no more expensive and when used with appropri-ate protocols, are more reliable. If more rigorous standardization is desired, various standard stain-ing procedures based on “ pure ” dyes have been devised, tested and the protocols published (e.g., Marshall et al. 1975, 1981, Wittekind et al. 1976).

A trouble-shooting scheme for Romanowsky

staining

In the past, laboratory workers have tended to view Romanowsky staining as capricious and diffi cult to “ get right. ” A trouble-shooting scheme can be based on the account offered above, however, and one is provided in Table 2.

Conclusions

There is a “ universal ” staining mechanism for Romanowsky stains that involves:

RED stained

eosinophil

granules &

erythrocytes

PURPLE

chromatin,

neutrophil

granules, &

platelets

BLUE

stained

lymphocyte

cytoplasms,

& nucleoli Continuous deposition

of the azure B-eosin Y

complex due to a

template effect — giving

a stain intensification

deposit of

PURPLE

complex

RED

stained

acidophilic

sites

BLUE

stained

basophilic

sites

Uptake of “second” dye by faster

staining sites only, yielding an

azure B–eosin Y complex

No short-term changes of

color in slower staining sites

Application of Romanowsky stain,

containing the

red acid (anionic) dye eosin Y, plus the

blue basic (cationic) dye azure B

Uptake of dye anions and

cations by simple acid and

basic dyeing

Fig. 5. The Romanowsky staining mechanism: the universal core processes. For illustrative purposes, the diagram

assumes that a blood smear is being stained.

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How Romanowsky stains work 49

Table 2. Trouble-shooting Romanowsky staining of blood smears and related specimens.

Problem Possible causes

A. Stain precipitates on slides or within pipework

of autostainer

1. Buffer concentration too high; reduce

2. Methanol (glycerol, dimethylsulfoxide) content too low; increase

3. Staining solution too old; replace

4. Temperature too high; keep coolerB. Overall staining too pale, but color balance

satisfactory

1. If “pure” azure B, dye content too low; due to: Impure lot of dye

or stock solution; replace Error in dilution or weighing; check

and correct Precipitation of stain; see AC. Cell nuclei blue, not purple 1. If “pure” azure B, dye content much too low, see B.

2. If “polychrome methylene blue,” azure B content of lot is low; replace3. If either dye type:

Methanol (etc.) content too high; reducepH of staining solution too low; check and correctStaining time too short; lengthenStaining temperature too low; checkEosin concentration in stock solution too low; make freshSpecimen too thick; lengthen staining time.If histological section, formalin fi xation too extended; lengthen

staining time or increase dye concentrationIf histological section, acetic acid differentiation or alcoholic

dehydration too long; reduce or change solventsOcclusion of specimen; check under the microscope for

presence of any overlying material.D. Neutrophils appear agranular in blood smears,

collagen fi bers pink, not purple in histological

sections

Any of factors listed in C

E. Neutrophils appear “toxic,” i.e., grossly enlarged

granules; general purple tinge to basophilic

cytoplasms

pH too high; check

Staining time too long; reduce

Azure B concentration too high; checkF. Erythrocytes and eosinophil granules too blue pH too high; check

Wrong buffer used; checkStaining time too long; reduce

G. Erythrocytes and eosinophil granules too

brownish orange, not pink

Some standardized stains give this appearance; rinse briefl y in

distilled waterpH too high; check and adjust

H. Basophil granules in blood smears fail to stain Fix with azure dye dissolved in methanol

1. Acidic and basic dyeing to give a polychrome effect.

2. Formation of an azure B-eosin Y complex, which provides further polychromasia, “ the Romanowsky-Giemsa effect. ”

3. Formation of this complex is rate-control-led, and exhibits a template intensifi cation effect.

4. The latter phenomenon provides the potential for stain intensifi cation.

Romanowsky stains have two advantageous features:

1. They can provide a technically straightfor-ward polychrome staining.

2. They can visualize entities at or below the limits of optical resolution under a standard microscope.

When technical problems arise with Romanowsky stains, “ rational ” trouble-shooting is possible.

Acknowledgments and dedications

I wish to acknowledge Prof. I. McGrath, Division of Integrated Biology, FBLS, University of Glasgow, for providing facilities, and P.N. Marshall, A.T. Sum-ner and Richard Hartley (Department of Chemistry, University of Glasgow) for helpful discussion and comment. I dedicate this paper to R.D. Lillie and D.W. Wittekind, two colorful giants of histotechnol-ogy. I had the benefi t and, sometimes, pleasure of meeting both men. The former told me long ago “ Young men like theories, but I think you should go back to the lab and do more experiments, Horobin, ” while the latter said to me many times “ But Dick … ” Perhaps I should have listened to them more attentively.

Declaration of interest: The authors report no con-fl icts of interest. The authors alone are responsible for the content and writing of the paper.

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50 Biotechnic & Histochemistry 2011, 86(1): 36–51

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