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Clinical staining of the ocular surface: Mechanisms and interpretations Q4
A.J. Bron a, *, 1, P. Argüeso b, 1, M. Irkec c, 1, F.V. Bright d, 1 a
Nuffield Department of Clinical Neurosciences and Nuffield Laboratory of Ophthalmology, University of Oxford, UK Schepens Eye Research Institute, Boston, MA, USA c Hacettepe University School of Medicine, Ankara, Turkey d Dept. of Chemistry, University at Buffalo, The State University of New York, Buffalo, USA b
a r t i c l e i n f o
a b s t r a c t
Article history: Received 12 August 2014 Received in revised form 8 October 2014 Accepted 8 October 2014 Available online xxx
In this article we review the mechanism of ocular surface staining. Water-soluble dyes are excluded from the normal epithelium by tight junctions and the surface glycocalyx. Shed cells can take up dye. A proportion of normal corneas, show sparse, scattered time-dependent, punctate fluorescein uptake, which, we hypothesise, is due to a graded loss of the glycocalyx barrier, permitting transcellular entry into pre-shed cells. In pathological staining, there is little evidence of ‘micropooling’ at sites of shedding and the term ‘punctate erosion’ may be a misnomer. It is more likely that the initial event involves transcellular dye entry and, in addition, diffusion across defective tight junctions. Different dye-staining characteristics probably reflect differences in molecular size and other physical properties of each dye, coupled with differences in visibility under the conditions of illumination used. This is most relevant to the rapid epithelial spread of fluorescein from sites of punctate staining, compared to the apparent confinement of dyes to staining cells with dyes such as lissamine green and rose bengal. We assume that fluorescein, with its lowest molecular weight, spreads initially by a paracellular route and then by transcellular diffusion. Solution-Induced Corneal Staining (SICS), related to the use of certain contact lens care solutions, may have a different basis, involving the non-pathological uptake of cationic preservatives, such as biguanides, into epithelial membranes and secondary binding of the fluorescein anion. It is transient and may not imply corneal toxicity. Understanding the mechanism of staining is relevant to the standardisation of grading, to monitoring disease and to the conduct of clinical trials. © 2014 Published by Elsevier Ltd.
Keywords: Punctate corneal staining Punctate keratitis Fluorescein dye Lissamine green Rose bengal Glycocalyx SICS PATH
Contents 1. 2.
3. 4. 5. 6.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anatomical and physiological factors affecting staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. General aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Tight junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Microplicae and the apical glycocalyx . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Aquaporins and gap junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The permeability of the ocular surface epithelia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Epithelial turnover and cell shedding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Epithelial metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Punctate staining of the ocular surface by instilled dyes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. Historical aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. Factors determining dye uptake and visibility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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* Corresponding author. E-mail address: [email protected] (A.J. Bron). 1 Percentage of work contributed by each author in the production of the manuscript is as follows: A.J. Bron: 50%; P. Argueso 25%; M. Ircek 10%; F.V. Bright 15%. http://dx.doi.org/10.1016/j.preteyeres.2014.10.001 1350-9462/© 2014 Published by Elsevier Ltd.
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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7.
8.
9.
Dyes in current use for staining the ocular surface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Fluorescein sodium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. Rose bengal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3. Lissamine green . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Terminology and mechanisms advanced to explain dye uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1. Terminology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2. Proposed mechanisms of punctate staining with topical dyes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.1. Micropooling at the site of epithelial cell shedding. ‘Punctate epithelial erosions’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.2. Uptake across defective epithelial tight junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.3. Transcellular uptake of dye and intercellular spread . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.4. Implications for the character of punctate staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.5. Fluorescein spread within the epithelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.6. Dye uptake following exposure to multipurpose solutions (MPS) e ‘transient hyperfluorescence’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3. Other forms of punctate staining at the ocular surface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Discussion and conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Uncited references . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction Topical dyes are used extensively to characterize ocular surface diseases and quantify their severity. The distribution of micropunctate staining may provide an etiological clue. The most frequently used dyes are disodium fluorescein, lissamine green and rose bengal, although mixtures of dyes have also been used (Norn, 1962, 1965, 1967, 1972; Toda and Tsubota 1993; Chodosh et al., 1994; Korb et al., 2008; Yoon et al., 2011), including a triple dye mixture containing alcian blue (Norn, 1964).
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There has been a longstanding debate as to the mechanism of staining by these dyes, which is relevant not only to the pathophysiology that they help to reveal, but also to the standardization of grading in clinical practice. In this review we will use the term ‘punctate staining’ in a general sense, to refer to punctate spots of dye at the surface of the epithelium, whether or not they are taken up into cells. Various possible mechanisms will be discussed, and in order to understand them it is important to consider those aspects of the corneal and conjunctival epithelia that influence dye behaviour.
Fig. 1. Transverse sections of normal human corneal epithelium. a. Semi-thin section, b. TEM, c. Microplicae, d. Wing Cells, e. Basal Cells. Key: All ¼ anterior limiting layer (Bowman's layer); Bc ¼ basal cell; ep ¼ epithelium; Sc ¼ most superficial epithelial cell; Str ¼ stroma; Wc ¼ wing cell. Courtesy of Prof Saeed Akhtar.
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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2. Anatomical and physiological factors affecting staining 2.1. General aspects
Q1
The corneal epithelium is a 5 layered structure possessing a basal layer of columnar cells, about 10 mm wide, intermediate layers of wing-shaped cells and a superficial layer of large, flat, polygonal cells, about 35 mm in diameter (Lemp and Mathers, 1989) (Fig. 1). These most superficial cells are connected to one another by intercellular tight junctions (zonulae occludentae) that encircle each cell and to an important extent obliterate the intervening paracellular space (Fig. 2). They thus retard the passage of ions and of hydrophilic molecules above a certain size, from the tears into the epithelium. Functionally, these junctions are sufficiently tight to convert this layer into an almost perfect semipermeable membrane towards sodium (Maurice, 1957), but are sufficiently leaky to allow the permeation of small hydrophilic molecules. 2.2. Tight junctions The tight junctions (Tjs) of the corneal epithelium consist of the trans-membrane proteins occludin, claudin and the junctional adhesion molecules (JAM), and the peripheral membrane proteins, ZO-1,-2 and -3 and MUPP-1 (Tsukita et al., 2001; Ban et al., 2003a, b). Other cytoplasmic proteins, such as cingulin and 7H6 antigen, are also present (Tsukita et al., 2001). Their organisation has been summarised by Ban et al. (2003a, b). Occludins and claudins contain 4 transmembrane domains, with both their N- and C-termini directed towards the cytoplasm. Occludin combines with tissuespecific members of the claudin family to form paired strands, one from each adjacent cell, that cross between the cells and close the intercellular space (Furuse et al., 2002). Occludin functions as a regulatory protein controlled by phosphorylation and has no significant structural role (Furuse et al., 1993). It may have permeability-related functions (Yu et al., 2005) and influence cell division (Wang et al., 2005).
Fig. 2. Diagram of the human corneal epithelium to show components which influence permeability. The glycocalyx of the surface epithelial cells is shown in pink (Glx). Tight junctions (Tj e in green) retard the entry of water-soluble molecules into the paracellular spaces. Functional gap junctions (Gj) are not present in the most superficial epithelial layer (layer 1) and are of limited functionality in the second layer (dotted channels). Gap junctions are fully functional in the third and deeper layers (continuous lines). Desmosomal attachments are not shown (See text for details).
3
The ZO (zonula occludens) proteins are members of the Membrane-Associated Guanylate Kinase (MAGUK) proteins and are located at membrane contact points of the Tjs. Here they form a complex which bridges between the Tj occludins and claudins and the actin cytoskeleton of the cell, which may be influenced by extracellular stimuli. The claudins constitute a family of (23 kDa) proteins and are the only junctional proteins to show tissue specificity. Yoshida et al. have summarized their role as structural components of Tjs in the corneal and the bulbar conjunctival epithelia (Yoshida et al., 2009). Despite the differences in permeability between these two epithelia they found that each epithelium expressed the same claudin subtypes, namely claudins 1, 4 and 7 throughout their thickness, although the expression of type 7 was superficial in the conjunctival epithelium. They postulated that it is the ratios of claudins and their phosphorylation status that determine the tightness of Tjs in these apical surface cells. They suggested too, that the presence of the same claudin subtypes in the subapical cell layers of these epithelia reflected an ability to rapidly assemble the components of tight junctions in preparation for the desquamation of surface cells. Although the tight junctions of the surface epithelial cells provide a barrier to the passage of ions and hydrophilic molecules, the apical plasma membranes and their associated glycocalyces are readily permeable to lipophilic molecules. The baso-lateral surfaces of the epithelial cells are highly interdigitated and separated from contiguous cells by narrow intercellular, or paracellular, spaces, bridged by numerous, scattered desmosomes. The basal cells are attached to the epithelial basal lamina by hemidesmosomes. The paracellular space exists between all the cells of the epithelium and provides a route for solute movement from the tears, extending from the level of the tight junctions anteriorly, to the epithelial basal lamina posteriorly. This space is expanded in bullous keratopathy because, with progressive endothelial failure, the hydrostatic pressure within the corneal stroma and epithelium becomes increasingly less negative and, ultimately, in advanced endothelial failure, becomes positive (Hatton et al., 2004; Bron, 2011). This point coincides with the onset of detectable epithelial edema (Miller and Benedek, 1973). The total surface area of the human conjunctiva is around 17.65 cm2 (±2.12) and that of the cornea, 1.04 cm2 ± 0.12 (Watsky et al., 1988). Its general architecture is similar to that of the cornea, but differs in some important respects. The bulbar conjunctiva exhibits a gently undulating surface and is loosely attached to the underlying sclera, which ensures that the globe is unrestricted during eye movements. The tarsal conjunctiva is firmly attached to the underlying tarsal plate. The conjunctiva is highly vascular while the cornea is avascular. Abundant goblet cells are distributed throughout the conjunctival epithelium except for two triangular, paralimbal zones in the horizontal meridian (Kessing, 1966). They are the source of the gel mucin of the tears (MUC5AC) (Argüeso and Gipson, 2001). The tight junctions of the conjunctival epithelium are more leaky than those of the corneal epithelium, possibly, as noted, because of differences in the number and type of claudin subunits that they contain (Yoshida et al., 2009). In keeping with this, the paracellular channels demonstrated by electron microscopy are much wider than those of the corneal epithelium (Bron et al., 1997; Huang et al., 1989). The conjunctival epithelium will permit the diffusion of small peptides in the region of 25 kD or more (Ottiger et al., 2009). 2.3. Microplicae and the apical glycocalyx The most superficial cells of the epithelium and conjunctiva, the apical cells, exhibit microplicae and microridges (around 150 nm in
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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height in guinea pig cornea (Nichols et al., 1983)), which project into the tears and increase the epithelial surface area (Smolin et al., 2004). Also, the superficial cells of both the cornea and conjunctiva, express a glycocalyx which is integrated with the apical plasma membrane (Nichols et al., 1983). In the human cornea the glycocalyx contains the membrane-embedded mucins, MUC1, MUC4 and MUC16, which have ecto- and endo-domains, galectin-3 and the mucin carbohydrate epitope T-antigen (Argüeso and Gipson, 2001) (Fig. 3a,b). Cell surface mucins are concentrated at the tips of the microplicae, forming a dense glycocalyx at the epithelial-tear film interface (Mantelli and Argueso, 2008). MUC4 has been demonstrated in conjunctival epithelium, with decreasing levels toward the central cornea (Inatomi et al., 1996; Pflugfelder et al., 2000). MUC16 is present in the epithelial glycocalyx of both the cornea and conjunctiva and is known to interact, through a polybasic amino acid sequence in the cytoplasmic tail, with proteins in the cytoskeleton (Blalock et al., 2007). Galectin-3 is an animal lectin defined by its affinity towards b-galactosides and the presence of one conserved carbohydrate-binding domain. Although galectin-3 exists as a monomer in solution, it can self-associate through
intermolecular interactions involving the N-terminal domain. When bound to a multivalent ligand, it can therefore mediate crosslinking of glycoproteins, such as the mucins and is thought to play this role within the glycocalyx (Fig. 4). The presence of MUC16 and galectin-3 appears to be essential for the exclusion of dyes from sheets of mature, cultured, human corneal epithelial cells (Argüeso and Sumiyoshi, 2006, Argueso et al., 2009) and probably from the intact ocular surface. The glycocalyx confers additional properties to the ocular surface, the most important of which is its intrinsic wettability (Cope et al., 1986; Liotet et al., 1987). When the glycocalyx is altered pathologically, wetting of the cornea becomes imperfect and tear stability is compromised. It has long been recognized that the hydrated, hydrophilic character of the mucins is due to extensive glycosylation of the numerous serine and threonine residues within tandem repeat regions (Argüeso and Gipson, 2001). Abnormalities in mucin glycosylation have been identified in many disorders where the stability of the tears is compromised, such as contact lens wear and dry eye (reviewed in Guzman-Aranguez and Argueso, 2010).
Fig. 3. a. Diagram showing the structural motifs within the three membrane-associated mucins expressed by the ocular surface epithelia. A common characteristic of all mucins is the presence of a variable number of tandem repeats (TR) of amino acids rich in serine and threonine that are highly O-glycosylated. Membrane-associated mucins have a transmembrane domain in the carboxy terminus that tethers the mucin into the apical cell membrane. In the cytoplasmic tail region (CT), MUC1 has tyrosine residues that can be phosphorylated (P) indicating potential signal transduction capability. In addition, the CT of MUC1 has been shown to be associated with b and g catenins, and is hypothesized to be associated with the actin cytoskeleton (Parry et al., 1990; Yamamoto et al., 1997). MUC4 has EGF-like domains and may be involved in growth regulation (from Gipson I.K. (2004). Distribution of mucins at the ocular surface. Exp Eye Res 78(3): 379-388. with permission). 3b. Electron micrograph of a corneal epithelial surface cell from a mouse, 37 days postnatal, after incubation with WGA-gold. Note the dense binding on the surface of this young cell. Aqueous uranyl acetate and Reynolds lead citrate stain. Original magnification 50,000 [from e Hazlett and Mathieu (1989). Glycoconjugates on corneal epithelial surface: effect of neuraminidase treatment. J Histochem Cytochem 37(8): 1215e1224. e with permission].
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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5
Knowledge of the connectivity of the corneal and conjunctival epithelial cells is important to an understanding of dye behaviour at the ocular surface. Studies of human and rabbit cornea using 5,(6) carboxyfluorescein (MWt. 376.32), have suggested that cells in or below the third epithelial layer are well coupled, while coupling of the surface cells is absent or deficient (Williams and Watsky, 1997, 2002). This difference is probably due to the differential expression of gap junctional proteins in the different epithelial layers Table 1 p 12: Permeability Characteristics of the Corneal and Conjunctival Epithelium. Data €ma €l€ from Ha ainen et al. (1997). Property
Fig. 4. Proposed model of galectin-mucin barrier formation on epithelial surfaces. It is conceived that two membrane-associated mucins, MUC1 and MUC16, interact with galectin-3 within the epithelial glycocalyx of individual cell membranes to provide a barrier under physiological conditions (- modified from Argueso et al. (2009). Association of cell surface mucins with galectin-3 contributes to the ocular surface epithelial barrier. J Biol Chem 284 (34): 23037e23045. e with permission).
2.4. Aquaporins and gap junctions Across the thickness of the epithelium, adjacent epithelial cells are connected to one another by water channels, which are engaged in transepithelial water transport (Candia et al., 2006) and by gap junctions which allow the exchange of small molecules and ions
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Contents lists available at ScienceDirect
Progress in Retinal and Eye Research journal homepage: www.elsevier.com/locate/prer
Clinical staining of the ocular surface: Mechanisms and interpretations Q4
A.J. Bron a, *, 1, P. Argüeso b, 1, M. Irkec c, 1, F.V. Bright d, 1 a
Nuffield Department of Clinical Neurosciences and Nuffield Laboratory of Ophthalmology, University of Oxford, UK Schepens Eye Research Institute, Boston, MA, USA c Hacettepe University School of Medicine, Ankara, Turkey d Dept. of Chemistry, University at Buffalo, The State University of New York, Buffalo, USA b
a r t i c l e i n f o
a b s t r a c t
Article history: Received 12 August 2014 Received in revised form 8 October 2014 Accepted 8 October 2014 Available online xxx
In this article we review the mechanism of ocular surface staining. Water-soluble dyes are excluded from the normal epithelium by tight junctions and the surface glycocalyx. Shed cells can take up dye. A proportion of normal corneas, show sparse, scattered time-dependent, punctate fluorescein uptake, which, we hypothesise, is due to a graded loss of the glycocalyx barrier, permitting transcellular entry into pre-shed cells. In pathological staining, there is little evidence of ‘micropooling’ at sites of shedding and the term ‘punctate erosion’ may be a misnomer. It is more likely that the initial event involves transcellular dye entry and, in addition, diffusion across defective tight junctions. Different dye-staining characteristics probably reflect differences in molecular size and other physical properties of each dye, coupled with differences in visibility under the conditions of illumination used. This is most relevant to the rapid epithelial spread of fluorescein from sites of punctate staining, compared to the apparent confinement of dyes to staining cells with dyes such as lissamine green and rose bengal. We assume that fluorescein, with its lowest molecular weight, spreads initially by a paracellular route and then by transcellular diffusion. Solution-Induced Corneal Staining (SICS), related to the use of certain contact lens care solutions, may have a different basis, involving the non-pathological uptake of cationic preservatives, such as biguanides, into epithelial membranes and secondary binding of the fluorescein anion. It is transient and may not imply corneal toxicity. Understanding the mechanism of staining is relevant to the standardisation of grading, to monitoring disease and to the conduct of clinical trials. © 2014 Published by Elsevier Ltd.
Keywords: Punctate corneal staining Punctate keratitis Fluorescein dye Lissamine green Rose bengal Glycocalyx SICS PATH
Contents 1. 2.
3. 4. 5. 6.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anatomical and physiological factors affecting staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. General aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Tight junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Microplicae and the apical glycocalyx . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Aquaporins and gap junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The permeability of the ocular surface epithelia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Epithelial turnover and cell shedding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Epithelial metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Punctate staining of the ocular surface by instilled dyes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1. Historical aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. Factors determining dye uptake and visibility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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* Corresponding author. E-mail address: [email protected] (A.J. Bron). 1 Percentage of work contributed by each author in the production of the manuscript is as follows: A.J. Bron: 50%; P. Argueso 25%; M. Ircek 10%; F.V. Bright 15%. http://dx.doi.org/10.1016/j.preteyeres.2014.10.001 1350-9462/© 2014 Published by Elsevier Ltd.
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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8.
9.
Dyes in current use for staining the ocular surface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Fluorescein sodium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. Rose bengal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3. Lissamine green . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Terminology and mechanisms advanced to explain dye uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1. Terminology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2. Proposed mechanisms of punctate staining with topical dyes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.1. Micropooling at the site of epithelial cell shedding. ‘Punctate epithelial erosions’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.2. Uptake across defective epithelial tight junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.3. Transcellular uptake of dye and intercellular spread . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.4. Implications for the character of punctate staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.5. Fluorescein spread within the epithelium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.6. Dye uptake following exposure to multipurpose solutions (MPS) e ‘transient hyperfluorescence’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3. Other forms of punctate staining at the ocular surface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Discussion and conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Uncited references . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction Topical dyes are used extensively to characterize ocular surface diseases and quantify their severity. The distribution of micropunctate staining may provide an etiological clue. The most frequently used dyes are disodium fluorescein, lissamine green and rose bengal, although mixtures of dyes have also been used (Norn, 1962, 1965, 1967, 1972; Toda and Tsubota 1993; Chodosh et al., 1994; Korb et al., 2008; Yoon et al., 2011), including a triple dye mixture containing alcian blue (Norn, 1964).
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There has been a longstanding debate as to the mechanism of staining by these dyes, which is relevant not only to the pathophysiology that they help to reveal, but also to the standardization of grading in clinical practice. In this review we will use the term ‘punctate staining’ in a general sense, to refer to punctate spots of dye at the surface of the epithelium, whether or not they are taken up into cells. Various possible mechanisms will be discussed, and in order to understand them it is important to consider those aspects of the corneal and conjunctival epithelia that influence dye behaviour.
Fig. 1. Transverse sections of normal human corneal epithelium. a. Semi-thin section, b. TEM, c. Microplicae, d. Wing Cells, e. Basal Cells. Key: All ¼ anterior limiting layer (Bowman's layer); Bc ¼ basal cell; ep ¼ epithelium; Sc ¼ most superficial epithelial cell; Str ¼ stroma; Wc ¼ wing cell. Courtesy of Prof Saeed Akhtar.
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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2. Anatomical and physiological factors affecting staining 2.1. General aspects
Q1
The corneal epithelium is a 5 layered structure possessing a basal layer of columnar cells, about 10 mm wide, intermediate layers of wing-shaped cells and a superficial layer of large, flat, polygonal cells, about 35 mm in diameter (Lemp and Mathers, 1989) (Fig. 1). These most superficial cells are connected to one another by intercellular tight junctions (zonulae occludentae) that encircle each cell and to an important extent obliterate the intervening paracellular space (Fig. 2). They thus retard the passage of ions and of hydrophilic molecules above a certain size, from the tears into the epithelium. Functionally, these junctions are sufficiently tight to convert this layer into an almost perfect semipermeable membrane towards sodium (Maurice, 1957), but are sufficiently leaky to allow the permeation of small hydrophilic molecules. 2.2. Tight junctions The tight junctions (Tjs) of the corneal epithelium consist of the trans-membrane proteins occludin, claudin and the junctional adhesion molecules (JAM), and the peripheral membrane proteins, ZO-1,-2 and -3 and MUPP-1 (Tsukita et al., 2001; Ban et al., 2003a, b). Other cytoplasmic proteins, such as cingulin and 7H6 antigen, are also present (Tsukita et al., 2001). Their organisation has been summarised by Ban et al. (2003a, b). Occludins and claudins contain 4 transmembrane domains, with both their N- and C-termini directed towards the cytoplasm. Occludin combines with tissuespecific members of the claudin family to form paired strands, one from each adjacent cell, that cross between the cells and close the intercellular space (Furuse et al., 2002). Occludin functions as a regulatory protein controlled by phosphorylation and has no significant structural role (Furuse et al., 1993). It may have permeability-related functions (Yu et al., 2005) and influence cell division (Wang et al., 2005).
Fig. 2. Diagram of the human corneal epithelium to show components which influence permeability. The glycocalyx of the surface epithelial cells is shown in pink (Glx). Tight junctions (Tj e in green) retard the entry of water-soluble molecules into the paracellular spaces. Functional gap junctions (Gj) are not present in the most superficial epithelial layer (layer 1) and are of limited functionality in the second layer (dotted channels). Gap junctions are fully functional in the third and deeper layers (continuous lines). Desmosomal attachments are not shown (See text for details).
3
The ZO (zonula occludens) proteins are members of the Membrane-Associated Guanylate Kinase (MAGUK) proteins and are located at membrane contact points of the Tjs. Here they form a complex which bridges between the Tj occludins and claudins and the actin cytoskeleton of the cell, which may be influenced by extracellular stimuli. The claudins constitute a family of (23 kDa) proteins and are the only junctional proteins to show tissue specificity. Yoshida et al. have summarized their role as structural components of Tjs in the corneal and the bulbar conjunctival epithelia (Yoshida et al., 2009). Despite the differences in permeability between these two epithelia they found that each epithelium expressed the same claudin subtypes, namely claudins 1, 4 and 7 throughout their thickness, although the expression of type 7 was superficial in the conjunctival epithelium. They postulated that it is the ratios of claudins and their phosphorylation status that determine the tightness of Tjs in these apical surface cells. They suggested too, that the presence of the same claudin subtypes in the subapical cell layers of these epithelia reflected an ability to rapidly assemble the components of tight junctions in preparation for the desquamation of surface cells. Although the tight junctions of the surface epithelial cells provide a barrier to the passage of ions and hydrophilic molecules, the apical plasma membranes and their associated glycocalyces are readily permeable to lipophilic molecules. The baso-lateral surfaces of the epithelial cells are highly interdigitated and separated from contiguous cells by narrow intercellular, or paracellular, spaces, bridged by numerous, scattered desmosomes. The basal cells are attached to the epithelial basal lamina by hemidesmosomes. The paracellular space exists between all the cells of the epithelium and provides a route for solute movement from the tears, extending from the level of the tight junctions anteriorly, to the epithelial basal lamina posteriorly. This space is expanded in bullous keratopathy because, with progressive endothelial failure, the hydrostatic pressure within the corneal stroma and epithelium becomes increasingly less negative and, ultimately, in advanced endothelial failure, becomes positive (Hatton et al., 2004; Bron, 2011). This point coincides with the onset of detectable epithelial edema (Miller and Benedek, 1973). The total surface area of the human conjunctiva is around 17.65 cm2 (±2.12) and that of the cornea, 1.04 cm2 ± 0.12 (Watsky et al., 1988). Its general architecture is similar to that of the cornea, but differs in some important respects. The bulbar conjunctiva exhibits a gently undulating surface and is loosely attached to the underlying sclera, which ensures that the globe is unrestricted during eye movements. The tarsal conjunctiva is firmly attached to the underlying tarsal plate. The conjunctiva is highly vascular while the cornea is avascular. Abundant goblet cells are distributed throughout the conjunctival epithelium except for two triangular, paralimbal zones in the horizontal meridian (Kessing, 1966). They are the source of the gel mucin of the tears (MUC5AC) (Argüeso and Gipson, 2001). The tight junctions of the conjunctival epithelium are more leaky than those of the corneal epithelium, possibly, as noted, because of differences in the number and type of claudin subunits that they contain (Yoshida et al., 2009). In keeping with this, the paracellular channels demonstrated by electron microscopy are much wider than those of the corneal epithelium (Bron et al., 1997; Huang et al., 1989). The conjunctival epithelium will permit the diffusion of small peptides in the region of 25 kD or more (Ottiger et al., 2009). 2.3. Microplicae and the apical glycocalyx The most superficial cells of the epithelium and conjunctiva, the apical cells, exhibit microplicae and microridges (around 150 nm in
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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height in guinea pig cornea (Nichols et al., 1983)), which project into the tears and increase the epithelial surface area (Smolin et al., 2004). Also, the superficial cells of both the cornea and conjunctiva, express a glycocalyx which is integrated with the apical plasma membrane (Nichols et al., 1983). In the human cornea the glycocalyx contains the membrane-embedded mucins, MUC1, MUC4 and MUC16, which have ecto- and endo-domains, galectin-3 and the mucin carbohydrate epitope T-antigen (Argüeso and Gipson, 2001) (Fig. 3a,b). Cell surface mucins are concentrated at the tips of the microplicae, forming a dense glycocalyx at the epithelial-tear film interface (Mantelli and Argueso, 2008). MUC4 has been demonstrated in conjunctival epithelium, with decreasing levels toward the central cornea (Inatomi et al., 1996; Pflugfelder et al., 2000). MUC16 is present in the epithelial glycocalyx of both the cornea and conjunctiva and is known to interact, through a polybasic amino acid sequence in the cytoplasmic tail, with proteins in the cytoskeleton (Blalock et al., 2007). Galectin-3 is an animal lectin defined by its affinity towards b-galactosides and the presence of one conserved carbohydrate-binding domain. Although galectin-3 exists as a monomer in solution, it can self-associate through
intermolecular interactions involving the N-terminal domain. When bound to a multivalent ligand, it can therefore mediate crosslinking of glycoproteins, such as the mucins and is thought to play this role within the glycocalyx (Fig. 4). The presence of MUC16 and galectin-3 appears to be essential for the exclusion of dyes from sheets of mature, cultured, human corneal epithelial cells (Argüeso and Sumiyoshi, 2006, Argueso et al., 2009) and probably from the intact ocular surface. The glycocalyx confers additional properties to the ocular surface, the most important of which is its intrinsic wettability (Cope et al., 1986; Liotet et al., 1987). When the glycocalyx is altered pathologically, wetting of the cornea becomes imperfect and tear stability is compromised. It has long been recognized that the hydrated, hydrophilic character of the mucins is due to extensive glycosylation of the numerous serine and threonine residues within tandem repeat regions (Argüeso and Gipson, 2001). Abnormalities in mucin glycosylation have been identified in many disorders where the stability of the tears is compromised, such as contact lens wear and dry eye (reviewed in Guzman-Aranguez and Argueso, 2010).
Fig. 3. a. Diagram showing the structural motifs within the three membrane-associated mucins expressed by the ocular surface epithelia. A common characteristic of all mucins is the presence of a variable number of tandem repeats (TR) of amino acids rich in serine and threonine that are highly O-glycosylated. Membrane-associated mucins have a transmembrane domain in the carboxy terminus that tethers the mucin into the apical cell membrane. In the cytoplasmic tail region (CT), MUC1 has tyrosine residues that can be phosphorylated (P) indicating potential signal transduction capability. In addition, the CT of MUC1 has been shown to be associated with b and g catenins, and is hypothesized to be associated with the actin cytoskeleton (Parry et al., 1990; Yamamoto et al., 1997). MUC4 has EGF-like domains and may be involved in growth regulation (from Gipson I.K. (2004). Distribution of mucins at the ocular surface. Exp Eye Res 78(3): 379-388. with permission). 3b. Electron micrograph of a corneal epithelial surface cell from a mouse, 37 days postnatal, after incubation with WGA-gold. Note the dense binding on the surface of this young cell. Aqueous uranyl acetate and Reynolds lead citrate stain. Original magnification 50,000 [from e Hazlett and Mathieu (1989). Glycoconjugates on corneal epithelial surface: effect of neuraminidase treatment. J Histochem Cytochem 37(8): 1215e1224. e with permission].
Please cite this article in press as: Bron, A.J., et al., Clinical staining of the ocular surface: Mechanisms and interpretations, Progress in Retinal and Eye Research (2014), http://dx.doi.org/10.1016/j.preteyeres.2014.10.001
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5
Knowledge of the connectivity of the corneal and conjunctival epithelial cells is important to an understanding of dye behaviour at the ocular surface. Studies of human and rabbit cornea using 5,(6) carboxyfluorescein (MWt. 376.32), have suggested that cells in or below the third epithelial layer are well coupled, while coupling of the surface cells is absent or deficient (Williams and Watsky, 1997, 2002). This difference is probably due to the differential expression of gap junctional proteins in the different epithelial layers Table 1 p 12: Permeability Characteristics of the Corneal and Conjunctival Epithelium. Data €ma €l€ from Ha ainen et al. (1997). Property
Fig. 4. Proposed model of galectin-mucin barrier formation on epithelial surfaces. It is conceived that two membrane-associated mucins, MUC1 and MUC16, interact with galectin-3 within the epithelial glycocalyx of individual cell membranes to provide a barrier under physiological conditions (- modified from Argueso et al. (2009). Association of cell surface mucins with galectin-3 contributes to the ocular surface epithelial barrier. J Biol Chem 284 (34): 23037e23045. e with permission).
2.4. Aquaporins and gap junctions Across the thickness of the epithelium, adjacent epithelial cells are connected to one another by water channels, which are engaged in transepithelial water transport (Candia et al., 2006) and by gap junctions which allow the exchange of small molecules and ions
![[Clinical diagnostics for the tear film and the ocular surface].](/assets/img/hold.png)
