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Electron microscopy of membrane-coating granules and a cell surface coat in keratinized and nonkeratinized human oral epithelium
Copyright © 1973 by Academic Press, Inc. All rights of reproduction in any form reserved J. ULTRASTRUCTURE RESEARCH
43, 205-219 (1973)
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Electron Microscopy of Membrane-Coating Granules and a Cell Surface Coat in Keratinized and Nonkeratinized Human Oral Epithelium A. F. HAYWARDand MARGARETEHACKEMANN1
Department of Oral Anatomy, Royal Dental Hospital of London, London, W.C.2., England Received July 26, 1972, and in revised form December 12, 1972 Membrane-coating granules occur in the cells of the keratinized epithelium of human gingiva and in the nonkeratinized buccal epithelium. During the differentiation of the epithelial cells, the cytoplasmic granules first increase in number and then decrease again as they are discharged into the intercellular spaces predominantly along the distal border of the cells. During this process, the outer membrane of each granule fuses with the cell plasma membrane. In gingival epithelium the contents of the granules stain positively with the periodic acid-thiocarbohydrazide-silver protein technique (PA-TCH) believed to indicate the presence of glycoprotein. The positive reaction is mainly observed just within the periphery of the granules. After granule fusion, a positive PA-TCH reaction appears along segments of the outer surface of the cell membrane, and it persists there until the cells are shed from the stratum corneum. The nonkeratinized buccal mucosa shows markedly less staining in the granules, closely associated with their limiting membrane. Plasma membrane staining is extensive in the superficial layers of the epithelium. One function of membrane-coating granules is in the formation of a cell surface coat, probably containing glycoprotein. The manifestation of this function varies with the degree of keratinization and possibly with other properties of the epithelium. As the keratinocytes of squamous epithelia differentiate during their passage from the basal layer to the free surface, granules appear in the cytoplasm. The granules have been given a variety of descriptive names from which may be selected "small dense granules" (29), "membrane-coating granules" (20), "keratinosomes" (38), and "cementsomes" (10). The expression membrane-coating granules has become popular and will be used in this communication even though its functional implications must be treated with reserve. 1 Present address: Department of Electron Microscopy, Clinical Research Centre, Harrow, Middlesex, England.
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The life cycle of membrane-coating granules (MCGs) is well documented for epithelia from a number of sources (5, 8, 10, 11, 18-20). They first appear in the keratinocytes of the stratum spinosum and are preferentially located near the superficial or distal membrane of the cell. When the cells reach the stratum granulosum of a keratinized epithelium or the more superficial layers of the stratum spinosum in a nonkeratinized epithelium, the MCGs decrease in number and ultimately disappear from the cytoplasm. There is evidence that during this process, their contents are discharged from the cell into the intercellular spaces (5, 8, 11, 18). The contents of MCGs in keratinized epithelium are lamellated and can therefore be identified in the intercellular spaces in the stratum granulosum and stratum corneum. The function of membrane-coating granules has not been fully elucidated. The name used here was originally coined as an expression of the concept that the granules contributed to the phenomenon of "membrane-thickening" (3). It is now known that this thickening of the plasma membrane is independent of the activity of membrane-coating granules (6, 18). Some components that may be of functional significance have been demonstrated in the granules. Hydrolytic enzymes, notably acid phosphatase (40, 41) and aryl sulfatase (25), have been shown to occur. Acid phosphatase has been found in both keratinized and nonkeratinized epithelia (33). Phospholipids have been demonstrated in MCGs of keratinized epithelia (i0, 24) and could be responsible for the lamellated structure of their contents. There have been reports of the presence of glycoproteins based on a positive reaction given by the periodic acid-silver methenamine method (12, 25), but other workers have been unable to confirm this observation (21). The MCG contents are discharged from the cell into the intercellular space, and they might contribute to glycoprotein material demonstrable within these spaces (4, 22, 39). An alternative method for glycoproteins, the periodic acid-thiocarbohydrazide-silver proteinate technique, has given differing results in keratinizing epithelia of different histological characteristics (13). Thus, in the hard palate of the rat, a positive reaction is readily observed in both MCGs and cell surface coats whereas in the soft palate of the rat no reaction is apparent. In this study the variation in the properties of MCGs and their relation to the cell surface coat has been investigated in human oral mucosa. Except for the dorsal surface of the tongue, the human oral cavity is lined with two types of epithelium. The masticatory type, including the gingival mucosa, is keratinized, though subject to variations possibly dependent on dental health (31). The lining mucosa which includes the buccal mucosa is nonkeratinized. These two epithelia present a greater divergence of structure than the examples from the rat and they also display structurally different MCGs. The membrane-coating granules of the two tissues have been examined electron microscopically using the periodic acid-thiocarbohydrazide technique (PA-TCH).
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MATERIAL AND METHODS Clinically healthy tissue was obtained by biopsy from an adult female volunteer. Samples of gingiva, from the molar ridge just distal to the third molar, and of vestibular buccal mucosa were cut into thin slices and fixed in 4% formaldehyde and 4 % glutaraldehyde in 0.16 M cacodylate buffer (pH 7.4) for 45 min at room temperature. Part of each sample was washed in buffer, dehydrated in methanol, and embedded in Araldite (CIBA-Geigy). The remainder was washed and treated with 1% osmium tetroxide in Veronal acetate buffer (pH 7.4) for 30 min before dehydration and embedding. Thin sections of osmicated tissues were cut and mounted on copper grids and electronstained with lead citrate and uranyl acetate. Sections of unosmicated tissues were mounted on gold grids and processed by the periodic acid-thiocarbohydrazide silver proteinate (PATCH) method described by Thi6ry (35) which is based on that of Seligman et al. (30). The P A - T C H method was also applied to sections of osmicated tissues which had first been treated with 3 % hydrogen peroxide for 30 min. The P A - T C H method is analogous to the periodic acid-Schiff technique (PA-S) (26) and periodic acid silver methenamine method (PA-SM) (17). It depends on the oxidation of the 1:2 glycol linkages of certain carbohydrates to aldehydes. Thiocarbohydrazide reacts with such aldehyde groups but at the same time retains its own capacity to reduce silver proteinate. Treatment of sections with silver proteinate solution releases silver at the sites of the original glycol linkages. The method is easily controlled and is less susceptible to the spontaneous "silvering" which occurs readily with PA-SM. The technique used was after Thi6ry (35) as follows: gold grids with sections attached were immersed in 1. 1 % aqueous periodic acid for 40 min followed by washing in water. 2. 0.2% thiocarbohydrazide (Taab Laboratories) in 20% acetic acid for 72 hours at room temperature (approx. 18-20°C). 3. 10% acetic acid wash. 4. 1% aqueous silver proteinate (Roques) in darkness for 30 minutes at room temperature. Most of the sections were subsequently electron-stained with lead citrate and uranyl acetate before examination with the AE1 EM6 electron microscope at an accelerating voltage of 60 kV. Some sections were examined without lead and uranyl staining using the AE1 EM6 and the Phillips 300 microscopes at accelerating voltages of 40 kV. The following controls were applied to the PA-TCH method: 1. To eliminate the presence of pre-existing aldehyde groups, for example, from the fixative, or other reducing compounds, sections were processed without periodic acid and/or without TCH. 2. To eliminate the possibility that reducing groups other than aldehydes might be produced by oxidation, a different oxidizing agent, i.e., 3 % hydrogen peroxide solution, was substituted for the periodic acid. 3. To confirm that the reaction of the T C H was with aldehyde groups, the periodic acid step was followed by specific aldehyde blocking procedures prior to the T C H reaction. A number of those used in light microscopy (26) were applied, but various problems were encountered, especially with the penetration of the reagents into Araldite sections. The efficacy of the methods was tried out on sections of striated muscle from hamster cheek
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pouch, known to contain deposits of glycogen and subjected to similar preparatory techniques. Three methods were found to be effective when used for 5 days at room temperature: (a) a saturated (0.4%) solution of dimedone (5,5-dimethyl-l,3-cyclohexanedione) in 5% acetic acid; (b) 11% aminophenol in glacial acetic acid; (c) aniline hydrochloride (9 ml aniline and 8 ml concentrated HCI made up to 100 ml with distilled water).
RESULTS
Gingiva The epithelium of the gingiva used for this study was ortho-keratinized. Sections .cut vertically through the epithelium showed the four layers of cells typical of such tissue, the stratum basale, spinosum, granulosum, and corneum. Electron micrographs showed the presence of membrane-coating granules in the cells of the middle layers of the stratum spinosum (Fig. 1). Most of them were located near the distal (i.e., superficial) cell membrane, but a few, identical granules could be seen near the opposite, proximal membrane. No association with the Golgi apparatus or other cytoplasmic component was observed. The individual MCGs (Figs. 1 and 2 a, b) were approximately 0.15 # m in diameter and were rounded or elongated. The outer membrane displayed a trilaminar or unit membrane structure with a total thickness of approximately 15 nm. Lamellations such as those illustrated are now regarded as typical of granules from a keratinized epithelium. In addition to the lamellations there was some evidence of a different pattern of material just beneath the limiting membrane of the MCGs. This took the form of an ill-defined granular or flocculent material which, in certain planes of section, had a meshlike appearance (Fig. 2a, b). This was particularly noticeable in obliquely cut granules where lamellations were not visible. When the cells had reached the stratum granulosum there was evidence of fusion of the outer membrane of the MCGs with the plasma membrane (Fig. 3). The intercellular spaces in the most superficial layers of the stratum granulosum and the adjacent layers of the stratum corneum contained lamellated material apparently identical to the contents of the MCGs of the cytoplasm.
FIG. 1. Section from the cytoplasm of a keratinocyte of the stratum spinosum of human gingiva showing the location of membrane-coating granules (MCG)near the distal cell boundary, x 78 000. FIGS. 1-3 and 11-12 are from aldehyde-fixed osmicated tissues stained with lead citrate and uranyl acetate solutions only. FI6. 2a, b. Membrane-coating granules of gingiva showing lamellated contents and meshlike peripheral material (-+). × 150 000. FIG. 3. Fusion of a single MCG with the distal plasma membrane at the boundary between stratum granulosum (SG)and stratum corneum (SC). The lamellated granule contents are in continuity with the intercellular space, x 200 000.
15--731825 J . Ultrastructure R e s e a r c h
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PA-TCH staining. Preliminary investigations showed that P A - T C H staining could not be successfully applied after osmium tetroxide had been included in tissue preparations unless the sections were treated with hydrogen peroxide. The best staining was obtained with unosmicated tissues. The lamellations of M C G contents were not preserved by aldehyde fixation alone but were visible only after osmication. Lamellations were usually poorly visualized in peroxide-treated sections. P A - T C H staining produced a silver deposit in the MCGs (Fig. 4). It was concentrated at the periphery adjacent to the limiting membrane. It was not possible to see the precise relationship of the stain to the membrane. In unosmicated material the contents of the MCGs were almost invariably replaced by an electron translucent space containing no silver stain. Occasionally "ghosts" of lamellations were observed or a few thin strands stretching across the granule. In peroxide-osmium preparations the silver deposit was more diffusely distributed through the M C G contents and though lamellations were not often seen, when they occurred (Fig. 5) they were stained with silver. From the stratum basale to the stratum spinosum, where the concentration of cytoplasmic MCGs was at its maximum, there was no silver staining associated with the plasma membrane of the cell. This observation was confirmed by examining sections which had not been stained with lead and uranyl salts in case a minor degree of staining had been obscured by the use of the heavy metals. As the number of cytoplasmic MCGs decreased and their outer membranes fused with the cell membrane (Fig. 6), staining became apparent on the outer surface of the cell membrane. Such staining was observed therefore between the most superficial layers of the stratum granulosum (Fig. 7) and throughout tLe stratum corneum (Fig. 8) including the free surface of desquamating cells. The silver deposit on the cell surface was found on short, discrete lengths of plasma membrane, and between these lengths no stain was visible. There was no staining within desmosomes. The silver was most prominent on the distal surface of the FIG. 4. Distal part of the cytoplasm of a keratinocyte from gingiva showing PA-TCH staining. Most of the stain in the membrane-coating granules is peripherally distributed, x 55 000. FIGs. 4 and 6-8 are from unosmicated tissue stained with PA-TCH and lead citrate and uranyl acetate. FIG. 5. Membrane-coating granules from osmicated aldehyde-fixed gingiva after peroxide treatment and PA-TCH staining. The upper granule shows lamellations and a meshlike appearance and is heavily stained. In this type of preparation most of the granules do not show lamellations. × 150 000. FIG. 6. The junction of a cell of stratum corneum (SC) and of stratum granulosum (SG) showing fusion of membranes of membrane-coating granules (MCG). The inner surface of their membrane stains with PA-TCH. × 92 000. FIG. 7. Intercellular and surface coat PA-TCH staining between cells of the last two layers of stratum granulosum of gingiva. The stain is predominantly associated with the distal surface of the cells. × 43 000. FI6. 8. PA-TCH staining between cells of the stratum corneum of gingiva. No staining is observed within desmosomes, x 43 000.
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FIG. 9. Membrane-coating granules (MCG) from a cell of the stratum spinosum of gingival epithelium in which treatment with periodic acid has been followed by the aldehyde blocking reagent aniline hydrochloride. Subsequent TCH and silver protein staining produces no reaction in the granules. x 57 000. FIG. 10. Part of the same preparation as Fig. 9 showing no staining reaction in the surface coat of the most superficial layer of the stratum granulosum when aldehyde blockade is introduced into the PA-TCH method, x 96 000.
cells a n d was only occasionally f o u n d o n the opposite side of the intercellular space or proximal surface of the cell. The layer of silver was separated f r o m the visible electron-dense i n n e r leaflet of the cell :membrane by a n a r r o w t r a n s l u c e n t space. The outer leaflet of the cell m e m b r a n e was n o t visible as a separate entity. Where the staining was especially dense, strands of silver-stained material stretched across the intercellular space at right angles to the cell m e m b r a n e . M e m b r a n e thickening occurs in gingiva as in other s q u a m o u s epithelia by the a p p o s i t i o n of a n electron dense substance to the cytoplasmic surface of the p l a s m a m e m b r a n e . This material was u n s t a i n e d by the P A - T C H method. Sections processed by control m e t h o d s n u m b e r e d 1-3, a, b, a n d c were u n i f o r m l y u n s t a i n e d in both the m e m b r a n e - c o a t i n g granules a n d surface coat (Figs. 9 a n d 10). The aldehyde blocking reagents also suppressed staining of k n o w n glycogen deposits in sections of muscle, b u t the time required for t h e m to do so was generally shorter
FIG. 11. Keratinocytes from middle layers of stratum spinosum of buccal mucosa showing the distribution of MCGs in the distal part of the cytoplasm. There are one or two at the opposite side of the cell. (~) M, mitochondria, x 9 000. FIG. 12. MCGs of buccal mucosa showing outer unit membrane and central electron dense core. There is an ill-defined material on the inner surface of this membrane x 140 000. FI~. 13. Fusion of MCG membrane with plasma membrane in buccal mucosa. The contents are still intact but are in continuity with the intercellular space, x 140 000.
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than that necessary to eliminate the PA-TCH reaction in granules or membrane coat. Buccal mucosa
The epithelium of the human buccal mucosa is normally nonkerafinized. The component layers of cells were therefore restricted to the stratum basale and stratum spinosum. The keratinocytes undergo a progressive change as they pass toward the surface, showing a degree of degeneration in the superficial layers of the stratum spinosum. Membrane thickening similar to that found in the gingiva occurs ill this region. Membrane-coating granules were found in the cells of the middle layers of the stratum spinosum. As in the gingiva they were distributed asymmetrically in the cytoplasm with a higher concentration along the distal cell membrane (Fig. 11). The granules were similar in size and shape to those of gingiva. There was an outer unit membrane, but the contents, instead of being lamellated, consisted of a central electron dense core (Fig. 12). Ill-defined dense material was apposed to the internal surface of the limiting membrane and was separated from the core by a translucent space traversed by thin electron-dense strands. As in gingiva, MCGs decreased in number and ultimately disappeared from the cytoplasm before membrane thickening occurred. Fusion of the granule membrane and the cell membrane was occasionally observed (Fig. 13), and the contents of the granule were then found to be in continuity with the intercellular space. Since the contents lacked distinctive lamellations, it was not possible to trace the fate of granule material beyond the stage of fusion. Silver staining. The results of PA-TCH staining in the buccal mucosa were considerably less striking than in the gingiva. The silver deposit was substantially less in amount and was easily obscured by heavy metal staining, particularly when examined at an accelerating voltage of 60 kV. The observations are based on sections which were not heavy metal-stained and were examined at 40 kV. The use of a lower accelerating voltage permits a higher contrast to be obtained on the exposed negative plate in order to compensate for the loss of contrast due to lack of heavy metal staining. There was a narrow band of silver deposit closely apposed to the inner surface of the limiting membrane of each membrane-coating granule (Fig. 14), but no stain was apparent in the granule contents. In the stratum basale and in the stratum spinosum, where the number of cytoplasmic MCGs was high, there was no silver staining of the cell membrane. As the number of granules decreased, however, the plasma membrane of the cell acquired a silver-positive layer on its outer surface (Fig. 14). The staining obscured the outer leaflet of the plasma membrane. In contrast to the discontinuous segments of staining observed in the gingiva, the
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FIG. 14. Unosmicated PA-TCH stained buccal epithelium showing silver stain over the inner surface of the MCG membrane and over the surface coat of the plasma membrane. There is no stain in the MCG contents or in the intercellular spaces (without heavy metal staining photographed at 40 kV). x 58 000.
cells of the buccal mucosa displayed a continuously stained outer surface layer. Only within the desmosomes was no staining observed. There was no silver deposit in the intercellular spaces. Control sections, processed without periodic acid treatment and/or without thiocarbohydrazide were in all cases completely unstained by the silver proteinate.
DISCUSSION This study confirms what is already known about the distribution and fate of membrane-coating granules and also that these features are the same in keratinized and nonkeratinized epithelia. The structure of the M C G s observed in the nonkeratinized epithelium is in agreement with that described by previous authors (32, 33). Particular attention is drawn to the presence of a differentiated layer just within the limiting membrane of the granules which m a y be independent of the lamellated contents found in gingival epithelium. The M C G s contain material that stains positively with the P A - T C H reaction. This result is similar to that obtained in human skin by others using the periodic acid-silver methenamine reaction (12, 25). In the present study glycogen has not been excluded as a source of staining by using diastase digestion because of a shortage of unfixed material. Preliminary experiments (unpublished) had shown that diastase digestion applied to formaldehyde-glutaraldehyde fixed tissue was not always ade-
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quate to suppress PA-TCH staining due to glycogen. Glycogen is not known to have the distribution described for this PA-TCH reactive material. The reaction has been shown by the controls used to be due to aldehyde groups produced by periodic acid oxidation and the material most likely to be responsible for Such a reaction is glycoprotein. There is one other possibility worth considering and that is that phospholipids in the membrane-coating granules (10, 24) are responsible for the staining. Although it appears to be unlikely that those phospholipids which give a positive PAS reaction (26) would occur in the sites at present under investigation, it is not easy to eliminate the possibility. Work on the epithelium of the rat hard palate (unpublished) has shown that phospholipid extraction with chloroform and methanol for 3 days has no effect on subsequent PA-TCH staining which is of a similar distribution in that tissue to human gingiva. It is proposed therefore to assume, subject to further information that the PA-TCH reaction in human oral mucosa is due to a glycoprotein. The existence of a glycoprotein surface coat on these cells is not surprising. PASpositive material has often been demonstrated in the intercellular region of squamous epithelia (4, 22, 39). Indeed it is a common finding that surfaces of cells of all types are coated with glycoprotein (27, 28). In most tissues the intracellular source of such carbohydrate-containing coats has not been demonstrated. Investigations have been concentrated on the intestinal epithelium with its prominent surface coat (15) and radiolabeled glycoprotein precursors have been traced, via the Golgi apparatus, to the cell surface (2, 16, 23). In carrying out work of this nature on the intestinal mucosa, Bennett and Leblond (2) also refer in passing to labeling of the surface coat of squamous epithelia but give no further details. From this present study and the others concerned with glycoprotein staining in squamous epithelia (12, 25) it appears that these tissues have a specific mechanism for the production of a cell surface coat and that membrane-coating granules form the main part of such a mechanism. The possibility that the granules originate in the Golgi apparatus (7, 18, 20) provides an analogy with the route taken by glycoprotein precursors in the intestinal epithelium. No evidence of glycoprotein staining was, however, found in the Golgi apparatus in this material. The PA-TCH method gives preparations with an appearance of precise localization of the carbohydrate material, and they indicate that it is particularly prominent at the periphery of the MCGs. The contents of the MCGs appear distinctly lamellated only after osmication and, since PA-TCH is applicable only to unosmicated material or tissues from which osmium has been removed, the relationship of the staining to the lamellations is uncertain. Osmicated preparations provide some evidence of a separate element of the contents at the periphery which might correspond to the stainable material. In the absence of
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osmication the lamellations might be either extracted during processing or displaced to the periphery of the MCGs. In peroxide-treated osmicated material, the lamellae are probably still present even when not visible. In such sections the reaction is certainly more diffuse though still partly peripheral, suggesting that at least some of it is due to a substance located within the lamellated material. It is possible, however, that this use of peroxide has rendered the method less specific. It has been claimed that more intense oxidation, for example, by prolonged periodic acid treatment, renders some acid mucopolysaccharides Schiff-positive (3). The behavior of the PA-TCH-stained material after the granules have been discharged from the cell suggests a very close affinity with the limiting membrane. The micrographs of fusion show that the mechanism of discharge is the incorporation of the M C G membrane into the cell membrane with release of the contents into the space. In the gingiva the staining shows no tendency to spread along the membrane but remains on short discrete segments of membrane, where each granule fused with the cell membrane. This appearance persists up to the surface of the epithelium. On the other hand, the lamellated contents are believed to disperse within the spaces and along the cell membrane (18) with loss of their ultrastructural appearance as the cells progress toward the surface. It is thought therefore that at least some of the stainable material is a membrane-linked component separate from the lamellated M C G contents. The variation in staining properties between tissues is well illustrated by human oral mucosa. It is of interest to try to correlate the differences observed with the concept of a spectrum of keratinization originally proposed by Weinmann (36) and elaborated by Alvares and Meyer (1) and others. Glycoprotein staining is pronounced in human gingival epithelium and in rat hard palate (13) which fall into a category of densely cornified epithelia (I, 14) near one extreme of the spectrum. By contrast, the buccal epithelium, which is nonkeratinized and falls at the opposite extreme, is stained lightly. The correlation of staining and degree of keratinization is not absolute since rat soft palate which falls into an intermediate category (14) does not exhibit staining (13). The epithelium of hamster cheek pouch has been examined (unpublished). It is of similar structure to rat soft palate and shows no glycoprotein staining. There is no evidence for stainable material in the contents of MCGs in human buccal mucosa, and their composition is unknown. The stained layer is closely associated with the limiting membrane. There is no discontinuity of the surface coat in buccal epithelium. It is very noticeable that the keratinocyte surface becomes highly convoluted especially in comparison with the gingiva. The membrane of the MCGs augments that of the cell, and this has been postulated as one of the functions of MCGs (5). No estimates are available of the number of granules in different tissues
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but it could be that the proportion of the cell membrane derived from M C G s in buccal epithelium is so high that most of it appears to be coated with glycoprotein. F r o m these observations, it is clear that one of the functions of M C G s is to produce a glycoprotein surface coat for keratinocytes. This function is modulated from tissue to tissue and is partly correlated with the degree of keratinization. It seems likely that there are other functions which might also be varied in a similar or converse way. The physiological properties of mucosae with which the difference might be correlated have not been investigated, but several workers have suggested a role for membrane-coating granules in determining the level of permeability or barrier function of epithelia (18, 34). The AE1 EM6 was supplied by the Welcome Foundation and facilities to use the Phillips 300 microscope were generously made available by Dr. R. Dourmashkin at the Medical Research Council, Clinical Research Centre. Technical assistance was provided by Mrs. R. Watson. REFERENCES 1. ALVARES,O. F. and MEYER, J., in SQUIER,C. A. and MEYER, J. (Eds.), Current Concepts of the Histology of Oral Mucosa. Thomas, Springfield, Illinois, 1972. 2. BENNETT,G. and LEBLOND,C. P., J. Cell Biol. 51, 875 (1972). 3. BERNrIELD,M. R. and BANERJEE,S. D., J. Cell Biol. 52, 664 (1972). 4. COHEN, L., Arch. Oral Biol. 13, 163 (1968). 5. FAR,MAN, A. I., J. Cell Biol. 21, 491, (1964). 6. - Anat. Rec. 156, 269 (1966). 7. FREI, J. V. and SI~ELOON,H., & Biophys. Biochem. CytoL 11, 719 (1961). 8. FRITmOF, L. and WERSXLL, J., J. Ultrastruct. Res. 12, 371, (1965). 9. GRUBB, C., HACKEMANN,M. and HILL, K. R., J. Ultrastruct. Res. 22, 458 (1968). 10. HASmMOTO,K., Arch. Dermatol. Forsch. 240, 349 (1971). 11. HASr~IMOTO,K., GROSS, B. G. and LEWR, W. F. J. Invest. Dermatol. 44, 119 (1965). 12. HASmMOTO,K., GROSS, B. G., NELSON, R. and L~WR, W. F., J. Invest. Dermatol. 47, 205 (1966). 13. HAYWARD,A. F. Arch. Oral Biol. 18, 67 (1973). 14. HAYWARD, A. F., HAMILTON,A. I. and HACICEMANN,M. M. A., Arch. Oral Biol. in Press (1973). 15. ITO, S., Anat. Rec. 148, 294 (1964). 16. - ibid. 151, 489 (1965). 17. MAmNOZZI, V., J. Biophys. Biochem. Cytol. 9, 121 (1961). 18. MARXINEZ,I. R. and PETERS, A., J. Anat. 130, 93 (1971). 19. MATOLTSY,A. G., J. Ultrastruct. Res. 15, 510, (1966). 20. MATOLTSY,A. G. and PARAKKAL,P. F., J. Cell Biol. 24, 297 (1965). 21. MERCER, E. H., JAHN, R. A. and MAmACH, H. I., J. Invest. Dermatol. 51, 204 (1968). 22. MEYER, J. and GERSON, S. J,~ Periodontics 2, 284 (1964). 23. NEUTRA, M. and LEBLOND,C. P. J. Cell Biol. 30, 137 (1966). 24. OLAH, I. and R6HLICH, P., Z. Zellforsch. Mikrosk. Anat. 73, 205 (1966).
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25. OLSON,R. L., NORDQUIST,R. E. and EVERETT,M. A., Arch. Klin. Exp. Dermatol. 234, 15 (1969). 26. PEARSZ,A. G. E., Histochemistry: Theoretical and Applied. Vol. 1., 3rd ed. Churchill, London, 1968. 27. RAMBOURQ,A. and LEBLOND, C. P., J. Cell BioL 32, 27 (1967). 28. RAMBOURG,A., NEUTRA, M. and LEBLOND,C. P., Anat. Rec. 154, 41 (1966). 29. SELBY,C. C., J. Invest. Dermatol. 29, 131 (1957). 30. SELIGMAN,A. M., HANKER,J. S., WASSERKRUG~H., DMOCHOWSKI,H., and KATZOFF,L., J. Histochem. Cytochem. 13, 629 (1965). 31. SICHER, H. and BHaSKAR, S. N. (Eds.), Orban's Oral Histology and Embryology, 7th ed., p. 224. Mosby, St. Louis, 1972. 32. SmVERMAN,S., J. Dent. Res. 46, 1433 (1967). 33. SILVERMAN,S. and KEARNS, G., Arch. Oral Biol. 15, 169 (1970). 34. SQUIER, C. A. and WATERHOtTSE,J. P., Nature (London) 215, 644 (1967). 35. THI~RY,J.-P., J. Microsc. (Paris) 6, 987 (1967). 36. WEINMANN,J. P., J. Dent. Res. 19, 57 (1940). 37. WEINSTOCK,M. and WILGRAM, G. F., J. Ultrastruct. Res. 30, 262 (1970). 38. WILGRAM,G. F., Arch. Dermatol. 94, 127 (1966). 39. WrSLOCKI,G. B., FAWCETT, D. W. and DEMPSEY, E. W., Anat. Rec. 110, 359 (1951). 40. WOLFF, K. and HOLUBAR, K., Arch. Klin. Exp. Derrnatol. 231, 1 (1967). 41. WOLFF, K. and SCHREINER,E., Arch. DermatoL 101, 276 (1970).
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Electron Microscopy of Membrane-Coating Granules and a Cell Surface Coat in Keratinized and Nonkeratinized Human Oral Epithelium A. F. HAYWARDand MARGARETEHACKEMANN1
Department of Oral Anatomy, Royal Dental Hospital of London, London, W.C.2., England Received July 26, 1972, and in revised form December 12, 1972 Membrane-coating granules occur in the cells of the keratinized epithelium of human gingiva and in the nonkeratinized buccal epithelium. During the differentiation of the epithelial cells, the cytoplasmic granules first increase in number and then decrease again as they are discharged into the intercellular spaces predominantly along the distal border of the cells. During this process, the outer membrane of each granule fuses with the cell plasma membrane. In gingival epithelium the contents of the granules stain positively with the periodic acid-thiocarbohydrazide-silver protein technique (PA-TCH) believed to indicate the presence of glycoprotein. The positive reaction is mainly observed just within the periphery of the granules. After granule fusion, a positive PA-TCH reaction appears along segments of the outer surface of the cell membrane, and it persists there until the cells are shed from the stratum corneum. The nonkeratinized buccal mucosa shows markedly less staining in the granules, closely associated with their limiting membrane. Plasma membrane staining is extensive in the superficial layers of the epithelium. One function of membrane-coating granules is in the formation of a cell surface coat, probably containing glycoprotein. The manifestation of this function varies with the degree of keratinization and possibly with other properties of the epithelium. As the keratinocytes of squamous epithelia differentiate during their passage from the basal layer to the free surface, granules appear in the cytoplasm. The granules have been given a variety of descriptive names from which may be selected "small dense granules" (29), "membrane-coating granules" (20), "keratinosomes" (38), and "cementsomes" (10). The expression membrane-coating granules has become popular and will be used in this communication even though its functional implications must be treated with reserve. 1 Present address: Department of Electron Microscopy, Clinical Research Centre, Harrow, Middlesex, England.
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The life cycle of membrane-coating granules (MCGs) is well documented for epithelia from a number of sources (5, 8, 10, 11, 18-20). They first appear in the keratinocytes of the stratum spinosum and are preferentially located near the superficial or distal membrane of the cell. When the cells reach the stratum granulosum of a keratinized epithelium or the more superficial layers of the stratum spinosum in a nonkeratinized epithelium, the MCGs decrease in number and ultimately disappear from the cytoplasm. There is evidence that during this process, their contents are discharged from the cell into the intercellular spaces (5, 8, 11, 18). The contents of MCGs in keratinized epithelium are lamellated and can therefore be identified in the intercellular spaces in the stratum granulosum and stratum corneum. The function of membrane-coating granules has not been fully elucidated. The name used here was originally coined as an expression of the concept that the granules contributed to the phenomenon of "membrane-thickening" (3). It is now known that this thickening of the plasma membrane is independent of the activity of membrane-coating granules (6, 18). Some components that may be of functional significance have been demonstrated in the granules. Hydrolytic enzymes, notably acid phosphatase (40, 41) and aryl sulfatase (25), have been shown to occur. Acid phosphatase has been found in both keratinized and nonkeratinized epithelia (33). Phospholipids have been demonstrated in MCGs of keratinized epithelia (i0, 24) and could be responsible for the lamellated structure of their contents. There have been reports of the presence of glycoproteins based on a positive reaction given by the periodic acid-silver methenamine method (12, 25), but other workers have been unable to confirm this observation (21). The MCG contents are discharged from the cell into the intercellular space, and they might contribute to glycoprotein material demonstrable within these spaces (4, 22, 39). An alternative method for glycoproteins, the periodic acid-thiocarbohydrazide-silver proteinate technique, has given differing results in keratinizing epithelia of different histological characteristics (13). Thus, in the hard palate of the rat, a positive reaction is readily observed in both MCGs and cell surface coats whereas in the soft palate of the rat no reaction is apparent. In this study the variation in the properties of MCGs and their relation to the cell surface coat has been investigated in human oral mucosa. Except for the dorsal surface of the tongue, the human oral cavity is lined with two types of epithelium. The masticatory type, including the gingival mucosa, is keratinized, though subject to variations possibly dependent on dental health (31). The lining mucosa which includes the buccal mucosa is nonkeratinized. These two epithelia present a greater divergence of structure than the examples from the rat and they also display structurally different MCGs. The membrane-coating granules of the two tissues have been examined electron microscopically using the periodic acid-thiocarbohydrazide technique (PA-TCH).
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MATERIAL AND METHODS Clinically healthy tissue was obtained by biopsy from an adult female volunteer. Samples of gingiva, from the molar ridge just distal to the third molar, and of vestibular buccal mucosa were cut into thin slices and fixed in 4% formaldehyde and 4 % glutaraldehyde in 0.16 M cacodylate buffer (pH 7.4) for 45 min at room temperature. Part of each sample was washed in buffer, dehydrated in methanol, and embedded in Araldite (CIBA-Geigy). The remainder was washed and treated with 1% osmium tetroxide in Veronal acetate buffer (pH 7.4) for 30 min before dehydration and embedding. Thin sections of osmicated tissues were cut and mounted on copper grids and electronstained with lead citrate and uranyl acetate. Sections of unosmicated tissues were mounted on gold grids and processed by the periodic acid-thiocarbohydrazide silver proteinate (PATCH) method described by Thi6ry (35) which is based on that of Seligman et al. (30). The P A - T C H method was also applied to sections of osmicated tissues which had first been treated with 3 % hydrogen peroxide for 30 min. The P A - T C H method is analogous to the periodic acid-Schiff technique (PA-S) (26) and periodic acid silver methenamine method (PA-SM) (17). It depends on the oxidation of the 1:2 glycol linkages of certain carbohydrates to aldehydes. Thiocarbohydrazide reacts with such aldehyde groups but at the same time retains its own capacity to reduce silver proteinate. Treatment of sections with silver proteinate solution releases silver at the sites of the original glycol linkages. The method is easily controlled and is less susceptible to the spontaneous "silvering" which occurs readily with PA-SM. The technique used was after Thi6ry (35) as follows: gold grids with sections attached were immersed in 1. 1 % aqueous periodic acid for 40 min followed by washing in water. 2. 0.2% thiocarbohydrazide (Taab Laboratories) in 20% acetic acid for 72 hours at room temperature (approx. 18-20°C). 3. 10% acetic acid wash. 4. 1% aqueous silver proteinate (Roques) in darkness for 30 minutes at room temperature. Most of the sections were subsequently electron-stained with lead citrate and uranyl acetate before examination with the AE1 EM6 electron microscope at an accelerating voltage of 60 kV. Some sections were examined without lead and uranyl staining using the AE1 EM6 and the Phillips 300 microscopes at accelerating voltages of 40 kV. The following controls were applied to the PA-TCH method: 1. To eliminate the presence of pre-existing aldehyde groups, for example, from the fixative, or other reducing compounds, sections were processed without periodic acid and/or without TCH. 2. To eliminate the possibility that reducing groups other than aldehydes might be produced by oxidation, a different oxidizing agent, i.e., 3 % hydrogen peroxide solution, was substituted for the periodic acid. 3. To confirm that the reaction of the T C H was with aldehyde groups, the periodic acid step was followed by specific aldehyde blocking procedures prior to the T C H reaction. A number of those used in light microscopy (26) were applied, but various problems were encountered, especially with the penetration of the reagents into Araldite sections. The efficacy of the methods was tried out on sections of striated muscle from hamster cheek
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pouch, known to contain deposits of glycogen and subjected to similar preparatory techniques. Three methods were found to be effective when used for 5 days at room temperature: (a) a saturated (0.4%) solution of dimedone (5,5-dimethyl-l,3-cyclohexanedione) in 5% acetic acid; (b) 11% aminophenol in glacial acetic acid; (c) aniline hydrochloride (9 ml aniline and 8 ml concentrated HCI made up to 100 ml with distilled water).
RESULTS
Gingiva The epithelium of the gingiva used for this study was ortho-keratinized. Sections .cut vertically through the epithelium showed the four layers of cells typical of such tissue, the stratum basale, spinosum, granulosum, and corneum. Electron micrographs showed the presence of membrane-coating granules in the cells of the middle layers of the stratum spinosum (Fig. 1). Most of them were located near the distal (i.e., superficial) cell membrane, but a few, identical granules could be seen near the opposite, proximal membrane. No association with the Golgi apparatus or other cytoplasmic component was observed. The individual MCGs (Figs. 1 and 2 a, b) were approximately 0.15 # m in diameter and were rounded or elongated. The outer membrane displayed a trilaminar or unit membrane structure with a total thickness of approximately 15 nm. Lamellations such as those illustrated are now regarded as typical of granules from a keratinized epithelium. In addition to the lamellations there was some evidence of a different pattern of material just beneath the limiting membrane of the MCGs. This took the form of an ill-defined granular or flocculent material which, in certain planes of section, had a meshlike appearance (Fig. 2a, b). This was particularly noticeable in obliquely cut granules where lamellations were not visible. When the cells had reached the stratum granulosum there was evidence of fusion of the outer membrane of the MCGs with the plasma membrane (Fig. 3). The intercellular spaces in the most superficial layers of the stratum granulosum and the adjacent layers of the stratum corneum contained lamellated material apparently identical to the contents of the MCGs of the cytoplasm.
FIG. 1. Section from the cytoplasm of a keratinocyte of the stratum spinosum of human gingiva showing the location of membrane-coating granules (MCG)near the distal cell boundary, x 78 000. FIGS. 1-3 and 11-12 are from aldehyde-fixed osmicated tissues stained with lead citrate and uranyl acetate solutions only. FI6. 2a, b. Membrane-coating granules of gingiva showing lamellated contents and meshlike peripheral material (-+). × 150 000. FIG. 3. Fusion of a single MCG with the distal plasma membrane at the boundary between stratum granulosum (SG)and stratum corneum (SC). The lamellated granule contents are in continuity with the intercellular space, x 200 000.
15--731825 J . Ultrastructure R e s e a r c h
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PA-TCH staining. Preliminary investigations showed that P A - T C H staining could not be successfully applied after osmium tetroxide had been included in tissue preparations unless the sections were treated with hydrogen peroxide. The best staining was obtained with unosmicated tissues. The lamellations of M C G contents were not preserved by aldehyde fixation alone but were visible only after osmication. Lamellations were usually poorly visualized in peroxide-treated sections. P A - T C H staining produced a silver deposit in the MCGs (Fig. 4). It was concentrated at the periphery adjacent to the limiting membrane. It was not possible to see the precise relationship of the stain to the membrane. In unosmicated material the contents of the MCGs were almost invariably replaced by an electron translucent space containing no silver stain. Occasionally "ghosts" of lamellations were observed or a few thin strands stretching across the granule. In peroxide-osmium preparations the silver deposit was more diffusely distributed through the M C G contents and though lamellations were not often seen, when they occurred (Fig. 5) they were stained with silver. From the stratum basale to the stratum spinosum, where the concentration of cytoplasmic MCGs was at its maximum, there was no silver staining associated with the plasma membrane of the cell. This observation was confirmed by examining sections which had not been stained with lead and uranyl salts in case a minor degree of staining had been obscured by the use of the heavy metals. As the number of cytoplasmic MCGs decreased and their outer membranes fused with the cell membrane (Fig. 6), staining became apparent on the outer surface of the cell membrane. Such staining was observed therefore between the most superficial layers of the stratum granulosum (Fig. 7) and throughout tLe stratum corneum (Fig. 8) including the free surface of desquamating cells. The silver deposit on the cell surface was found on short, discrete lengths of plasma membrane, and between these lengths no stain was visible. There was no staining within desmosomes. The silver was most prominent on the distal surface of the FIG. 4. Distal part of the cytoplasm of a keratinocyte from gingiva showing PA-TCH staining. Most of the stain in the membrane-coating granules is peripherally distributed, x 55 000. FIGs. 4 and 6-8 are from unosmicated tissue stained with PA-TCH and lead citrate and uranyl acetate. FIG. 5. Membrane-coating granules from osmicated aldehyde-fixed gingiva after peroxide treatment and PA-TCH staining. The upper granule shows lamellations and a meshlike appearance and is heavily stained. In this type of preparation most of the granules do not show lamellations. × 150 000. FIG. 6. The junction of a cell of stratum corneum (SC) and of stratum granulosum (SG) showing fusion of membranes of membrane-coating granules (MCG). The inner surface of their membrane stains with PA-TCH. × 92 000. FIG. 7. Intercellular and surface coat PA-TCH staining between cells of the last two layers of stratum granulosum of gingiva. The stain is predominantly associated with the distal surface of the cells. × 43 000. FI6. 8. PA-TCH staining between cells of the stratum corneum of gingiva. No staining is observed within desmosomes, x 43 000.
l
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FIG. 9. Membrane-coating granules (MCG) from a cell of the stratum spinosum of gingival epithelium in which treatment with periodic acid has been followed by the aldehyde blocking reagent aniline hydrochloride. Subsequent TCH and silver protein staining produces no reaction in the granules. x 57 000. FIG. 10. Part of the same preparation as Fig. 9 showing no staining reaction in the surface coat of the most superficial layer of the stratum granulosum when aldehyde blockade is introduced into the PA-TCH method, x 96 000.
cells a n d was only occasionally f o u n d o n the opposite side of the intercellular space or proximal surface of the cell. The layer of silver was separated f r o m the visible electron-dense i n n e r leaflet of the cell :membrane by a n a r r o w t r a n s l u c e n t space. The outer leaflet of the cell m e m b r a n e was n o t visible as a separate entity. Where the staining was especially dense, strands of silver-stained material stretched across the intercellular space at right angles to the cell m e m b r a n e . M e m b r a n e thickening occurs in gingiva as in other s q u a m o u s epithelia by the a p p o s i t i o n of a n electron dense substance to the cytoplasmic surface of the p l a s m a m e m b r a n e . This material was u n s t a i n e d by the P A - T C H method. Sections processed by control m e t h o d s n u m b e r e d 1-3, a, b, a n d c were u n i f o r m l y u n s t a i n e d in both the m e m b r a n e - c o a t i n g granules a n d surface coat (Figs. 9 a n d 10). The aldehyde blocking reagents also suppressed staining of k n o w n glycogen deposits in sections of muscle, b u t the time required for t h e m to do so was generally shorter
FIG. 11. Keratinocytes from middle layers of stratum spinosum of buccal mucosa showing the distribution of MCGs in the distal part of the cytoplasm. There are one or two at the opposite side of the cell. (~) M, mitochondria, x 9 000. FIG. 12. MCGs of buccal mucosa showing outer unit membrane and central electron dense core. There is an ill-defined material on the inner surface of this membrane x 140 000. FI~. 13. Fusion of MCG membrane with plasma membrane in buccal mucosa. The contents are still intact but are in continuity with the intercellular space, x 140 000.
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than that necessary to eliminate the PA-TCH reaction in granules or membrane coat. Buccal mucosa
The epithelium of the human buccal mucosa is normally nonkerafinized. The component layers of cells were therefore restricted to the stratum basale and stratum spinosum. The keratinocytes undergo a progressive change as they pass toward the surface, showing a degree of degeneration in the superficial layers of the stratum spinosum. Membrane thickening similar to that found in the gingiva occurs ill this region. Membrane-coating granules were found in the cells of the middle layers of the stratum spinosum. As in the gingiva they were distributed asymmetrically in the cytoplasm with a higher concentration along the distal cell membrane (Fig. 11). The granules were similar in size and shape to those of gingiva. There was an outer unit membrane, but the contents, instead of being lamellated, consisted of a central electron dense core (Fig. 12). Ill-defined dense material was apposed to the internal surface of the limiting membrane and was separated from the core by a translucent space traversed by thin electron-dense strands. As in gingiva, MCGs decreased in number and ultimately disappeared from the cytoplasm before membrane thickening occurred. Fusion of the granule membrane and the cell membrane was occasionally observed (Fig. 13), and the contents of the granule were then found to be in continuity with the intercellular space. Since the contents lacked distinctive lamellations, it was not possible to trace the fate of granule material beyond the stage of fusion. Silver staining. The results of PA-TCH staining in the buccal mucosa were considerably less striking than in the gingiva. The silver deposit was substantially less in amount and was easily obscured by heavy metal staining, particularly when examined at an accelerating voltage of 60 kV. The observations are based on sections which were not heavy metal-stained and were examined at 40 kV. The use of a lower accelerating voltage permits a higher contrast to be obtained on the exposed negative plate in order to compensate for the loss of contrast due to lack of heavy metal staining. There was a narrow band of silver deposit closely apposed to the inner surface of the limiting membrane of each membrane-coating granule (Fig. 14), but no stain was apparent in the granule contents. In the stratum basale and in the stratum spinosum, where the number of cytoplasmic MCGs was high, there was no silver staining of the cell membrane. As the number of granules decreased, however, the plasma membrane of the cell acquired a silver-positive layer on its outer surface (Fig. 14). The staining obscured the outer leaflet of the plasma membrane. In contrast to the discontinuous segments of staining observed in the gingiva, the
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FIG. 14. Unosmicated PA-TCH stained buccal epithelium showing silver stain over the inner surface of the MCG membrane and over the surface coat of the plasma membrane. There is no stain in the MCG contents or in the intercellular spaces (without heavy metal staining photographed at 40 kV). x 58 000.
cells of the buccal mucosa displayed a continuously stained outer surface layer. Only within the desmosomes was no staining observed. There was no silver deposit in the intercellular spaces. Control sections, processed without periodic acid treatment and/or without thiocarbohydrazide were in all cases completely unstained by the silver proteinate.
DISCUSSION This study confirms what is already known about the distribution and fate of membrane-coating granules and also that these features are the same in keratinized and nonkeratinized epithelia. The structure of the M C G s observed in the nonkeratinized epithelium is in agreement with that described by previous authors (32, 33). Particular attention is drawn to the presence of a differentiated layer just within the limiting membrane of the granules which m a y be independent of the lamellated contents found in gingival epithelium. The M C G s contain material that stains positively with the P A - T C H reaction. This result is similar to that obtained in human skin by others using the periodic acid-silver methenamine reaction (12, 25). In the present study glycogen has not been excluded as a source of staining by using diastase digestion because of a shortage of unfixed material. Preliminary experiments (unpublished) had shown that diastase digestion applied to formaldehyde-glutaraldehyde fixed tissue was not always ade-
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quate to suppress PA-TCH staining due to glycogen. Glycogen is not known to have the distribution described for this PA-TCH reactive material. The reaction has been shown by the controls used to be due to aldehyde groups produced by periodic acid oxidation and the material most likely to be responsible for Such a reaction is glycoprotein. There is one other possibility worth considering and that is that phospholipids in the membrane-coating granules (10, 24) are responsible for the staining. Although it appears to be unlikely that those phospholipids which give a positive PAS reaction (26) would occur in the sites at present under investigation, it is not easy to eliminate the possibility. Work on the epithelium of the rat hard palate (unpublished) has shown that phospholipid extraction with chloroform and methanol for 3 days has no effect on subsequent PA-TCH staining which is of a similar distribution in that tissue to human gingiva. It is proposed therefore to assume, subject to further information that the PA-TCH reaction in human oral mucosa is due to a glycoprotein. The existence of a glycoprotein surface coat on these cells is not surprising. PASpositive material has often been demonstrated in the intercellular region of squamous epithelia (4, 22, 39). Indeed it is a common finding that surfaces of cells of all types are coated with glycoprotein (27, 28). In most tissues the intracellular source of such carbohydrate-containing coats has not been demonstrated. Investigations have been concentrated on the intestinal epithelium with its prominent surface coat (15) and radiolabeled glycoprotein precursors have been traced, via the Golgi apparatus, to the cell surface (2, 16, 23). In carrying out work of this nature on the intestinal mucosa, Bennett and Leblond (2) also refer in passing to labeling of the surface coat of squamous epithelia but give no further details. From this present study and the others concerned with glycoprotein staining in squamous epithelia (12, 25) it appears that these tissues have a specific mechanism for the production of a cell surface coat and that membrane-coating granules form the main part of such a mechanism. The possibility that the granules originate in the Golgi apparatus (7, 18, 20) provides an analogy with the route taken by glycoprotein precursors in the intestinal epithelium. No evidence of glycoprotein staining was, however, found in the Golgi apparatus in this material. The PA-TCH method gives preparations with an appearance of precise localization of the carbohydrate material, and they indicate that it is particularly prominent at the periphery of the MCGs. The contents of the MCGs appear distinctly lamellated only after osmication and, since PA-TCH is applicable only to unosmicated material or tissues from which osmium has been removed, the relationship of the staining to the lamellations is uncertain. Osmicated preparations provide some evidence of a separate element of the contents at the periphery which might correspond to the stainable material. In the absence of
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osmication the lamellations might be either extracted during processing or displaced to the periphery of the MCGs. In peroxide-treated osmicated material, the lamellae are probably still present even when not visible. In such sections the reaction is certainly more diffuse though still partly peripheral, suggesting that at least some of it is due to a substance located within the lamellated material. It is possible, however, that this use of peroxide has rendered the method less specific. It has been claimed that more intense oxidation, for example, by prolonged periodic acid treatment, renders some acid mucopolysaccharides Schiff-positive (3). The behavior of the PA-TCH-stained material after the granules have been discharged from the cell suggests a very close affinity with the limiting membrane. The micrographs of fusion show that the mechanism of discharge is the incorporation of the M C G membrane into the cell membrane with release of the contents into the space. In the gingiva the staining shows no tendency to spread along the membrane but remains on short discrete segments of membrane, where each granule fused with the cell membrane. This appearance persists up to the surface of the epithelium. On the other hand, the lamellated contents are believed to disperse within the spaces and along the cell membrane (18) with loss of their ultrastructural appearance as the cells progress toward the surface. It is thought therefore that at least some of the stainable material is a membrane-linked component separate from the lamellated M C G contents. The variation in staining properties between tissues is well illustrated by human oral mucosa. It is of interest to try to correlate the differences observed with the concept of a spectrum of keratinization originally proposed by Weinmann (36) and elaborated by Alvares and Meyer (1) and others. Glycoprotein staining is pronounced in human gingival epithelium and in rat hard palate (13) which fall into a category of densely cornified epithelia (I, 14) near one extreme of the spectrum. By contrast, the buccal epithelium, which is nonkeratinized and falls at the opposite extreme, is stained lightly. The correlation of staining and degree of keratinization is not absolute since rat soft palate which falls into an intermediate category (14) does not exhibit staining (13). The epithelium of hamster cheek pouch has been examined (unpublished). It is of similar structure to rat soft palate and shows no glycoprotein staining. There is no evidence for stainable material in the contents of MCGs in human buccal mucosa, and their composition is unknown. The stained layer is closely associated with the limiting membrane. There is no discontinuity of the surface coat in buccal epithelium. It is very noticeable that the keratinocyte surface becomes highly convoluted especially in comparison with the gingiva. The membrane of the MCGs augments that of the cell, and this has been postulated as one of the functions of MCGs (5). No estimates are available of the number of granules in different tissues
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but it could be that the proportion of the cell membrane derived from M C G s in buccal epithelium is so high that most of it appears to be coated with glycoprotein. F r o m these observations, it is clear that one of the functions of M C G s is to produce a glycoprotein surface coat for keratinocytes. This function is modulated from tissue to tissue and is partly correlated with the degree of keratinization. It seems likely that there are other functions which might also be varied in a similar or converse way. The physiological properties of mucosae with which the difference might be correlated have not been investigated, but several workers have suggested a role for membrane-coating granules in determining the level of permeability or barrier function of epithelia (18, 34). The AE1 EM6 was supplied by the Welcome Foundation and facilities to use the Phillips 300 microscope were generously made available by Dr. R. Dourmashkin at the Medical Research Council, Clinical Research Centre. Technical assistance was provided by Mrs. R. Watson. REFERENCES 1. ALVARES,O. F. and MEYER, J., in SQUIER,C. A. and MEYER, J. (Eds.), Current Concepts of the Histology of Oral Mucosa. Thomas, Springfield, Illinois, 1972. 2. BENNETT,G. and LEBLOND,C. P., J. Cell Biol. 51, 875 (1972). 3. BERNrIELD,M. R. and BANERJEE,S. D., J. Cell Biol. 52, 664 (1972). 4. COHEN, L., Arch. Oral Biol. 13, 163 (1968). 5. FAR,MAN, A. I., J. Cell Biol. 21, 491, (1964). 6. - Anat. Rec. 156, 269 (1966). 7. FREI, J. V. and SI~ELOON,H., & Biophys. Biochem. CytoL 11, 719 (1961). 8. FRITmOF, L. and WERSXLL, J., J. Ultrastruct. Res. 12, 371, (1965). 9. GRUBB, C., HACKEMANN,M. and HILL, K. R., J. Ultrastruct. Res. 22, 458 (1968). 10. HASmMOTO,K., Arch. Dermatol. Forsch. 240, 349 (1971). 11. HASr~IMOTO,K., GROSS, B. G. and LEWR, W. F. J. Invest. Dermatol. 44, 119 (1965). 12. HASmMOTO,K., GROSS, B. G., NELSON, R. and L~WR, W. F., J. Invest. Dermatol. 47, 205 (1966). 13. HAYWARD,A. F. Arch. Oral Biol. 18, 67 (1973). 14. HAYWARD, A. F., HAMILTON,A. I. and HACICEMANN,M. M. A., Arch. Oral Biol. in Press (1973). 15. ITO, S., Anat. Rec. 148, 294 (1964). 16. - ibid. 151, 489 (1965). 17. MAmNOZZI, V., J. Biophys. Biochem. Cytol. 9, 121 (1961). 18. MARXINEZ,I. R. and PETERS, A., J. Anat. 130, 93 (1971). 19. MATOLTSY,A. G., J. Ultrastruct. Res. 15, 510, (1966). 20. MATOLTSY,A. G. and PARAKKAL,P. F., J. Cell Biol. 24, 297 (1965). 21. MERCER, E. H., JAHN, R. A. and MAmACH, H. I., J. Invest. Dermatol. 51, 204 (1968). 22. MEYER, J. and GERSON, S. J,~ Periodontics 2, 284 (1964). 23. NEUTRA, M. and LEBLOND,C. P. J. Cell Biol. 30, 137 (1966). 24. OLAH, I. and R6HLICH, P., Z. Zellforsch. Mikrosk. Anat. 73, 205 (1966).
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25. OLSON,R. L., NORDQUIST,R. E. and EVERETT,M. A., Arch. Klin. Exp. Dermatol. 234, 15 (1969). 26. PEARSZ,A. G. E., Histochemistry: Theoretical and Applied. Vol. 1., 3rd ed. Churchill, London, 1968. 27. RAMBOURQ,A. and LEBLOND, C. P., J. Cell BioL 32, 27 (1967). 28. RAMBOURG,A., NEUTRA, M. and LEBLOND,C. P., Anat. Rec. 154, 41 (1966). 29. SELBY,C. C., J. Invest. Dermatol. 29, 131 (1957). 30. SELIGMAN,A. M., HANKER,J. S., WASSERKRUG~H., DMOCHOWSKI,H., and KATZOFF,L., J. Histochem. Cytochem. 13, 629 (1965). 31. SICHER, H. and BHaSKAR, S. N. (Eds.), Orban's Oral Histology and Embryology, 7th ed., p. 224. Mosby, St. Louis, 1972. 32. SmVERMAN,S., J. Dent. Res. 46, 1433 (1967). 33. SILVERMAN,S. and KEARNS, G., Arch. Oral Biol. 15, 169 (1970). 34. SQUIER, C. A. and WATERHOtTSE,J. P., Nature (London) 215, 644 (1967). 35. THI~RY,J.-P., J. Microsc. (Paris) 6, 987 (1967). 36. WEINMANN,J. P., J. Dent. Res. 19, 57 (1940). 37. WEINSTOCK,M. and WILGRAM, G. F., J. Ultrastruct. Res. 30, 262 (1970). 38. WILGRAM,G. F., Arch. Dermatol. 94, 127 (1966). 39. WrSLOCKI,G. B., FAWCETT, D. W. and DEMPSEY, E. W., Anat. Rec. 110, 359 (1951). 40. WOLFF, K. and HOLUBAR, K., Arch. Klin. Exp. Derrnatol. 231, 1 (1967). 41. WOLFF, K. and SCHREINER,E., Arch. DermatoL 101, 276 (1970).