Biochimie, 68 (1986) 611-617 © Soci6t6 de Chimie biologique/Elsevier, Paris

611

Minireview

Structure of gastrointestinal mucins: searching for the Rosetta stone

Christian L. LABOISSE

Laboratoire de Biologic et Physiologic des Celhdes Digestives, U239 1NSERM, Facult~ Bichat, 16 rue Huchard, 75018 Paris, France

Introduction

Mucins are high molecular weight glycoproteins secreted by specialized epithelial ceils of the gas- trointestinal mucosa as a visco-elastic gel. It is gene- rally admitted that the function of mucins is to lubricate and to protect the epithelial lining of the gastrointestinal mucosa.

Since mucins are secreted in large amounts throughout the gastrointestinal tract, it may well be imagined that the structure of these molecules is perfectly known in 1986. This is not the case however, mainly because, due to their very high molecular weights (more than 106), they are not easily amenable to the classical analytical proced- ures of biochemistry. In the same way, the exact functions of these substances are not clearly understood.

However, recent investigations have shed some light on the structure and biosynthesis of these sub- stances. This paper will focus on recent investiga- tions aimed at puriflyng mucins and at elucidating the structure, antigenic markers and biosynthesis of gastrointestinal mucins. Particular emphasis will be given to recently available and promising in vitro model systems to study the structure and biosyn- thesis of mucins.

Isolation, purification and polymeric structures

When considering the size, macromolecular beha- vior or rheological properties of the mucus gel, it is of primary importance to isolate and purify gastro-intestinal mucins in such a way that the puri- fied macromolecules are representative of those pre- sent in situ, in the mucus gel. Therefore, in this chapter, the emphasis will be on non-degradative methods of isolation and purification of mucins. As pointed out by Allen et aL, it is essential to start

with mucus gel scraped off the surface of the mucosa, since luminal mucus can contain large amounts of proteolytically degraded glycopro- tein [1]. Solubilization of the mucous gel can be achieved by brief homogenization [2] in a salt solu- tion, like 0.2 M NaCI. Mucins extracted in such non-denaturing conditions retain their full gel- forming properties [3]. They can be also extracted by using low-shear procedures in denaturing sop vents such as guanidium chloride [4]. A major advantage of this latter procedure, advocated by Carlstedt, is the suppression of degradative enzy- mes [4] which are likely to be present in mucosal scrapings. Guanidium chloride supplemented with low molecular weight proteinase inhibitors can be used to protect the primary structure of mucins during extraction and purification [4]. This method of extraction, however, may perturb the 'native' conformation of mucins.

Mucus gel scraped from the mucosal surface is heavily contaminated with proteins and nucleic acids from mucosal ceils, together with contribu- tions from intestinal secretions and the gut micro- flora that characterize the lower regions of the digestive tract [2]. The starting point of many methods aimed at purifying mucin glycoproteins is generally gel chromatography with a large pore bed. Although mucus glycoproteins can be considerably separated by gel filtration on Sepharose 4B from the majority of lower molecular weight proteins, these mucin preparations still contain some low molecular weight, non-covalently bound, extra- neous proteins [5]. In addition, DNA can coelute with undegraded mucins in the void volume of even the most porous gels [4]. The most successful method for removing contaminating proteins as well as DNA is equilibrium centrifugation in a CsCI density gradient. This method, pioneered by Creeth [6], has been successfully used to purify pig gastric mucus [51, pig colonic mucus [7], human gastric mucus [8] and pig small-intestinal mucus [2]. An

612 C.L. Laboisse

extensive purification of mucins can be achieved by associating the method of equilibrium centrifuga- tion in a CsCI gradient with a gel filtration techni- que [2]. Mucins can be considered as pure preparations only when the absence of nucleic acids, non-covalently bound lipids, non-mucin pro- teins and proteoglycans from the extracellular matrix can be conclusively demonstrated. In parti- cular, sodium dodecyl sulphate-polyacrylamide gel electrophoresis is the most sensitive method to detect contaminating proteins or glycoproteins [9]. Typically, in non-reduced conditions, mucins do not enter polyacrylamide gels [9, 10], even those containing very low acrylamide concentrations, and they barely enter agarose-polyacrylamide compo- site gels [11]. In contrast, contaminating proteins o r glycoproteins can be easily detected as bands of lower molecular weight in such gels.

A pure mucin preparation, even from one indi- vidual, exhibits some degree of microheterogeneity with respect to size, density and charge. This micro- heterogeneity, also referred to as polydispersity [12], reflects a continuity of molecular structure centered around a mean size or compositional state [13]. The microheterogeneity of mucins is explained by the natural variation in the composi- tion of oligosaccharide chains [13].

Molecular weight determinations, performed on mucins from different sources, show that purified mucins display MrS ranging from 2 × 106 for pig gastric mucins [14] to 15× 106 for pig colonic mucins [7]. There are considerable variations in the molecular weight of mucins from different regions of the gastrointestinal tract of the same species. Reports on the M r of the same type of mucin by different investigators show variations [4]. In par- ticular, Carlsdtedt points out that the MrS of mucins extracted in denaturing solvents, in the pre- sence of proteinase inhibitors, are generally higher than those obtained by homogenization in solvents compatible with protease activity [4].

Analysis of the protein and carbohydrate com- position of human mucins shows that the protein content is between 12 and 17070 [15, 16] by weight of the molecule. Mucin molecules consist of pro- tein cores with numerous carbohydrate side chains.

Many studies have been devoted to the elucida- tion of the polymeric structure of mucins. In par- ticular, with regard to the question of whether the mucin molecules isolated from the gastrointestinal tract or from other origins are polymericstruc- tures whose subunits are linked by disulphide brid- ges, there is considerable controversy [17-20,26]. The presence of subunits linked by S-S bridges in gas- tric mucins was first recognized by Snary et al. [14].

Further studies have shown that, with thiol reduc- tion, human gastric [8], pig gastric [14, 21,22], pig small-intestinal [23] and pig colonic [7] mucins undergo depolymerization, liberating subunits with molecular weights ranging from 250 000 to 720 000. In addition, during reduction of some mucins, non- glycosylated peptides are released, with an appa- rent M r on SDS-polyacrylamide gels of 70000 for pig gastric mucins [9], 90 000 for pig small-intestinal mucin [23] and 118 000 for rat [11] and human [24] small intestinal mucin. Although there is no firm evidence as to their precise role, it is generally assu- med that these peptides account for the linking of mucin monomers within the 'native' polymeric mucin [1]. The validity of this general scheme (i.e. polymeric structure maintained by disulfide brid- ges) has been questioned on the basis of experiments showing the absence of release of subunits upon reduction of human [15, 19] and rat [25] small-intes- tinal mucins and of ovine submaxillary mucins [20]. In addition, in the case 0f ovine submaxillary mucins [20] there was no cysteine residue at all within the peptidic core of the molecule. Recently, however, it has been shown by Fahim et aL [11] and Mantle et al. [24] that the absence of the ' l ink ' peptide found in the early experiments of reduction of intestinal mucins was probably due to incomplete reduction of the mucin. With regard to the sub- maxillary mucins, it may be that their structure is very different from that of the mucins of the gas- trointestinal tract.

In addition to these experiments of thiol reduc- tion, studies of proteolytic attack of mucus glyco- proteins by papain, pronase or pepsin have played an important role in the understanding of the struc- ture of these substances [1,2,24]. These studies have led to the concept of two regions of proteins in these glycoproteins, a heavily glycosylated region which is protected from the proteolytic attack and a non-glycosylated (or ' naked' region) which is sus- ceptible to the protease digestion. The protein frac- tion of pig gastric mucin which is susceptible to proteolytic attack represents about 35°70 by weight of the total protein content of the molecule [5, 27]. In particular, the 70 000 dalton link protein found in pig gastric mucin is lost upon proteolysis by papain [9], confirming that it is not protected by glycoconjugates.

The careful analysis of the structure of pig gas- tric mucins by Allen and his group has led him to propose a model for their polymeric structure: according to this, four highly glycosylated subunits are joined together by the central 70 000 dalton pro- tein. Since the arrangement of the glycosylated subunits around the link peptide is like that of the

Structure o f gastrointestinal nntcins 613

vanes of a windmill around their axis, this model is also referred to as the 'windmill model'. Other models have been also proposed to explain the poly- meric structure of mucins. In their ' flexible thread model', Carlstedt et aL propose [4] that the subunits, each of them containing naked and glycosylated regions, are attached end to end by disulfide bridges [28]. This model was first postu- lated to accomodate hydrodynamically for cervi- cal mucins [29], but was later extended to encompass data for pig gastric mucins as well as mucins from chronic bronchitic sputum [30]. Electron-microscopic examination of pig gastric, cervical and respiratory mucins by Carlstdet and Sheehan [31], as well as the findings of Slayter et aL [32] on respiratory mucins, suggest that all these mucins behave as flexible extended threads. In his model, Carlstedt points out that naked (protease- sensitive) regions of the mucins are stabilized by intramolecular disulphide bonds. Such a model pre- dicts that the ' l ink' proteins, as defined by Allen, are not present in mucins. How then can the release of low molecular weight proteins upon reduction of mucins be explained ? According to Carlstedt's hypothesis, some proteases present during extrac- tion or handling of the mucins would 'nick the naked' region but they would be unable to destroy the polymeric structure, since the naked peptides would be still maintained cohesive by the intramo- lecular disulphide bonds. However, upon reduc- tion, these peptides would be freed and would be interpreted as ' l ink' peptides [4].

Finally, the nature of the gel-forming noncova- lent interactions between constituent glycoprotein molecules is unknown, but recent evidence for gas- tric mucus [33] suggests that such interactions occur between the glycosylated regions of the glycopro- tein, possibly by stable interdigitation of the car- bohydrate chains.

Carbohydrate chains of mucins

To study the carbohydrate side chains, it is not necessary to isolate the mucins in their 'native' state, and proteolysis is a rapid and efficient way to obtain relatively pure mucin glycoproteins. A major advance in the study of the structure of the carbohydrate side chains has come from the dis- covery that the linkage of glycoconjugates of mucins to the protein core (O-glycosidic linkage) could be specifically cleaved in alkaline aqueous solution with protection of reducing groups by borohydride reduction (a process termed beta- elimination) [34].

Mucins contain up to 85°70 carbohydrate which is arranged in oligosaccharide side chains of varying length and degree of branching, covalently linked to the protein backbone. These oligosaccharides fall into two broad groups, namely acidic and neutral. Of particular interest are the acidic oligosacchari- des: mucins from many mammalian sources con- tain sialic acid residues and sulphate groups, and these substituents are responsible for the polyanion- ic character of the macromolecules.

The carbohydrate side chains can be considered to have three structurally distinct domains, namely a core region incorporating the linkage to protein, a backbone region which may be linear or bran- ched, and a peripheral chain terminating region [35]. Each of these domains is endowed with parti- cular antigens. These carbohydrate antigens are now well defined by means of monoclonal antibo- dies [36] :

Core region. The oligosaccharide chains of mucins are joined to the protein core by a linkage between N-acetylgalactosamine (GalNac) and the oxygen of serine or threonine (referred to as an O-glycosidic linkage). This type of linkage is rather unusual, but it is not specific for mucin glycoproteins. It is also observed in a variety of other glycoprotein molecu- les, such as the proteoglycans of connective tissue, some hormones [37], the CD8 antigen at the surface of T lymphocytes [38], or the ZP3 glyc0protein pre- sent in the mouse egg's zona pellucida [39].

Biosynthesis of the side chain can be arrested after addition of the first carbohydrate. Alterna- tively, the core structure can elongate through addi- tion of other carbohydrates. In particular, a galactose residue joined by a 1--3 linkage to GalNac constitutes the T blood group antigen T (T from 'Thomsen-Friedenreich'). This structure reacts with peanut lectin [40]. T antigen is the pre- cursor of the blood group antigens M and N, and is found in human gastric glycoproteins only in ' non- secretors' and in meconium extracts, regardless of secretor status [41]. This antigen is cryptic in goblet cell mucins of normal human colonic mucosa of the adult [41,42]. It can be revealed only after neuraminidase treatment of the mucins, as shown by immunofluorescence, using PNA and fluo- rescein-coupled anti-PNA antibodies on paraffin sections of human colonic mucosa [421.

Backbone regions. These are made up of repeat disaccharide sequences of galactose and N- acetylglucosamine, and are endowed with antigens i, I and IMa [43]. These antigens were first recog- nized as developmentally regulated antigens of

614 C.L. Laboisse

human erythrocytes [44], and are now known also to be carbohydrate structures of human gastric glycoproteins. In the majority of the ' secretor ' population, antigen IMa is not found in gastric glycoproteins, because the antigenicity of this struc- ture is normally masked by blood group antigens A, B or H. On the other hand, IMa is found in glycoprotein extracts from human gastric mucosa of 25°7o of 'non-secretor ' persons [41,45] and in meconium extracts, regardless of the receptor status. Ii determinants are never found in human colonic glycoproteins [41].

The peripheral regions. They specify ABH, Lewis a and Lewis b antigens. In human gastric glyco- proteins, the expression of ABH antigens is found only in extracts o f ' secretor' persons [45]. The epi- thelium of the large intestine of the fetus contains blood group antigens A, B and H throughout its entire length. However, at parturition the blood group substances of the left colon rapidly disap- pear, leaving the epithelium of the adult left colon devoid of blood group antigens [46-49]. Recently, several reports were concerned with the demons- tration of novel blood related carbohydrate speci- ficities which are useful to define gastrointestinal developmental antigens: SSEA-I antigen, also desi- gnated as Le x antigen, which was first observed at the 8-cell stage of the mouse embryo, is found also in the intestinal meconium of the human fetus [50]. The sialylated blood group antigen Le a, defined by hybridoma antibody 19.9 [51], is a normal com- ponent of glycoproteins isolated from gastric mucosae of 'non-secretors ' . [36]. In a thorough immunohistologic study, Raux et al. [52] found this antigen to be associated to the gastrointestinal mucins of human fetuses and newborns.

Taken together, all these studies point to the great diversity of the antigenic markers of mucin glyco- conjugates, which are related to blood group anti- gens. It is clear also that there are marked differences in the expression of these antigens throughout the gastrointestinal tract. Finally, recent studies have stressed the importance of these anti- gens as onco-developmental markers [36].

Protein of mucus glycoproteins

Numerous studies on the amino acid composition of mucins have revealed a characteristically high serine and threonine content of the protein back- bone of the glycoproteins [5, 7, 9, 11, 15, 53, 54]. Threonine and serine are the amino acids involved in the linkage o f the carbohydrate side chains to

Table I. Repartition of M antigens in the gastrointesti- nal tract of the human adult.

Antigen Localization in the gastrointestinal tract

Stomach Small intestine Colon

Duod. Jej. lie. Prox. Dist.

MI + + + . . . . . M3 - + + + + + + + + + + + + + + + M3C . . . . + + + + + + M3SI - + + + + + + + + + ++ +

Data summarized from [58, 63, 64]. Semi-quantative assessment: + + + : most of the mucous cells are stained; + + : 50°70 or less of the mucous cells are stained; + : only rare mucous cells are stained; - : negative staining.

the protein core by N-acetylgalactosamine. Protein loss on digestion of the colonic-mucus glycoproteins is characterized by a considerable loss of content of all amino acids except threonine, serine and pro- line [7]. These results demonstrate clearly that the peptide region rich in serine, threonine and proline is heavily glycosylated and resistant to proteolysis. The other ' naked ' regions, which have a less dis- tinctive amino acid composition, are susceptible to hydrolysis by proteases [7]. Amino acid analyses of the link peptides have been also performed [9, 11]. The 70 000 dalton ' l ink' peptide o f gastric mucins is rich in glycine and glutamic acid, with substantial amounts of cystein [9]. Recently, Wes- ley et aL [55] have addressed the issue of whether peptide differences could exist between neutral and acidic human intestinal mucins. Their results show that acidic and neutral mucins exhibit differences in amino acid composition and that these differen- ces cannot be assigned either to the 118 000 dalton link peptide or to the naked peptide regions in the mucin. Rather, the differences appear to be local- ized in the highly glycosylated region. These results suggest therefore that the oligosaccharide compo- sition of small intestinal and colonic mucin depends on the transcriptional control or specific core pep- tides [551.

Deglycosylation of mucins can be achieved by means of specific enzymes [20] or treatment with trifluoromethanesulfonic acid [56] which cleaves the O-glycosydic bonds. These deglycosylated mucins, termed apomucins, serve as acceptors o f glycosy- lation in vitro, to determine in particular the mini- mum size requirements of protein core for efficient glycosylation. Hill et al. reported the purification and sequence analysis of tryptic peptides represen- ting 106 of the 650 residues in ovine submaxillary

Structure o f gastrointestinal nntcins 615

apomucin [57]. They found no salient features in the sequences adjacent to serine and threonine resi- dues that might prescribe a substrate acceptor re- quirement for incorporation of GalNAc into O-glycosidic linkage. However, there appeared to be a minimum size requirement for glycosylation.

Antigens of muein glycoproteins

It has been conclusively demonstrated that it is pos- sible to raise polyclonal antibodies against gastro- intestinal mucins, which, after proper absorption, specifically recognize the mucins and do not react with the mucin-associated blood group antigens [58-64]. More specifically, it has been possible to raise antibodies specific for gastric mucins, refer- red to as MI antigens by Bara et al. [58]. Other anti- bodies were made specific for the human small intestinal mucins [60] and others specific for large intestinal mucins [58-60]. Zweibaum et aL [65] have identified a polymorphic determinant system in colonic secretions (WZ) having no correlation with blood group ABH and Lewis determinants. Finally, it is clear from several studies that mucins differ in their antigenic composition throughout the gastrointestinal tract. This is exemplified by the work of Bara et aL [58] together with that of Nardelli et aL [63, 64], who defined four antigenic markers (Table I): MI, associated with human gastric fucomucins, and M3, M3SI and M3C res- tricted to intestinal, small-intestinal and colonic goblet cells, respectively. Antibodies to mucins have been useful in developing specific radioimmuno- assays for rat [66] and human [61, 67] goblet cell mucins. Recently, antibodies to mucins were used to analyze the antigenic determinants of human small-intestinal goblet cell mucin [24]. In parti- cular, it was demonstrated that antibodies pre- pared against the whole mucin did not label the ' link' peptide, as shown by immunoblot analysis of the mucin subunits separated on polyacrylamide gels under reducing conditions.

Mucin biosynthesis

their intracellular topology. The initial glycosyla- tion reaction in the synthesis of O-linked glyco- proteins involves the direct transfer of N- acetylgalactosamine from uridine diphosphate (UDP)-GalNAc to the polypeptide by a U D P - GalNAc: polypeptide transferase and does not require oligosaccharide preassembly nor lipid inter- mediates [68]. Apart from the Asn-GlcNAc lin- kage which occurs only in the presence of a parti- cular peptide sequence, there is apparently no pri- mary amino acid sequence requirement for the polypeptide: GalNAc transferase [57]. The subcel- lular sites of oligosaccharide initiation have been investigated by four general approaches [69]: (1) radioautography following injection of radioactive precursors into living animals, (2) subcellular frac- tionation of tissues from animals previously injected with radioactive precursors, (3) glycosyltransferase

assays of subcellular fractions, and (4) electron- microscopic localization of GalNAc residues in cel- lular compartments of intestinal goblet cells using colloidal gold-coupled lectins as probes [70].

Taken together, these data point clearly to the Golgi apparatus as the subcellular compartment where O-glycosylation events first occur [69, 70]. Once the first sugar has been incorporated into the peptide chain, subsequent elongation can occur via stepwise addition of monosaccharides from sugar nucleotide substrates to the growing oligosaccha- ride acceptor. Each step is catalyzed b y a glycosyl- transferase which is specific not only for the glycosyl donor but also for the acceptor molecule. Recent investigations indicate that the enzymes involved in the synthesis of O-linked oligosaccha- rides occupy separate compartments within the Golgi apparatus [71]. It is therefore tempting to spe- culate that oligosaccharide chain elongation of mucins occurs in different regions of the Golgi apparatus as the glycoproteins move from the ' cis' side towards the' trans' side [70]. Finally, it should be pointed out that, in addition to the glycosyla- tion steps, mucus glycoproteins undergo several post-translati0nal modifications, which include such steps as sulfation and acylation [72].

The biosynthesis of glycoproteins is particulary well known in the case of those with N-glycosidic lin- kages. It involves assembly of a lipid-linked oligo- saccharide, transfer of the oligosaccharide en bloc to the nascent peptide chain, processing of the oli- gosaccharide chains by glycosidases and addition of terminal sugars by glycosyltransferases. Much less information is available on the assembly of the oligosaccharide chains of O-linked glycoproteins or

Conclusion: New tools to study mucin struc- ture and biosynthesis

Although considerable attention has been focused on defining the structure and the biosynthesis of mucins, some important questions remain unans- wered. For example, what is the exact polymeric structure of mucin glycoproteins ? How do the ini-

616 C.L . Laboisse

tial steps of mucin biosynthesis proceed? Is the pep- tidic core of mucins synthesized as a single peptide, or is it produced from the transcription of differ- ent mRNAs?

It can be suggested that some of these problems will be addressed in the near future by using cell culture systems. For example, short-term cultures of hamster tracheal epithelial cells have proved use- ful in studying the mucus glycoproteins [73] of the respiratory tract. However, there are still only a few reports on the successful culture of normal epithe- lial cells of the gastrointestinal tract [74]. Interes- tingly, Rattner et aL have recently reported a method for maintaining guinea pig gastric mucous cell monolayers in short-term culture [75]. The ideal model system for the study o f mucins would be a long-term culture of pure mucus-secreting cells, which would have several advantages. First, it would be a source of mucins with constant characteristics, allowing reproducible experiments. Secondly, the mucins secreted in the culture medium would be free of any bacterial contamina- tion. Thirdly, since the mucous cells would be maintained in a relatively defined environment, it would be possible to test the effects of changes in the environmental conditions on mucin syn- thesis. However, the establishment of long-term cultures of epithelial cells from the gastrointestinal tract is often accompanied by the loss o f their differentiated functions. To circumvent this pro- blem, some investigators turned their attention toward gastrointestinal carcinoma cell lines, and they found that some of these cell lines are useful model systems to study differentiated functions o f the gastrointestinal tract [76-78]. In particular, it has been possible to establish several stably differentiated clonal cell lines after treatment of human colonic adenocarcinoma HT29 cells with sodium butyrate [78]. Among the clones isolated so far, several consist of homogeneous populations of polarized mucus-secreting cells. These clonal cell lines retain their capacity to secrete mucins in long-term culture under standard culture condi- tions as well as in serum-free medium. The mucins produced by these clonal ceils can be characterized by using either biochemical or immunological methods [79]. In other words, these lines are cons- tant sources of mucins with defined characteristics. Therefore, with such model systems it will probably be possible to study in detail the initial steps of mucin synthesis by using standard analytical methods such as ' pu l se -chase ' labelling of the intracellular proteins, followed by immunopre- cipitation with specific antibodies of the mucin glycoproteins.

References

l Allen A., Bell A., Mantle M. & Pearson J.P. (1982) in: Mucus in Health and Disease - 11. A dvances in Experimental Medicine and Biology, Vol. 144 (Chantler E.N., Elder J.B. & Elstein M., eds.), Ple- num Press, New York. pp. 115-133

2 Mantle M. & Allen A. (1981) Biochem. J. 195,267-275

3 Allen A. (1981) in: Physiology o f the Gastrointesti- nal Tract (Johnson L.R. ed.), Raven Press, New York. pp. 617-639

4 Carlstedt I., Sheehan J.K., Corfield A.P. & Gallag- her J.T. (1985) Essays Biochem. 20, 40-76

5 Starkey B.J., Snary D. & Allen A. (1974) Biochem. J. 141,633-639

6 Creeth J.M. & Denborough M.A. (1970) Biochem. J. 117, 879-891

7 Marshall T. & Alien A. (1978) Biochem. J. 173, 569-578

8 Pearson J.P., Allen A. & Venables C.W. (1980) Gas- troenterology 78, 709-715

9 Pearson J.P., Allen A. & Parry S. (1981) Biochem. J. 197, 155-162

I0 Gold D.V., Shochat D. & Miller F. (1981) J. BioL Chem. 256, 6354-6358

11 Fahim R.E.F., Forstner G.G. & Forstner J.F. (1983) Biochem. J. 209, 117-124

12 Gibbons R.A. (1963) Nature (London) 200, 665-666 13 Wesley A.W., Forstner J.F. & Forstner G. (1983)

Carbohydrate Res. 115, 151-163 14 Snary D., Allen A. & Pain R.H. (1970) Biochem.

Biophys. Res. Commun. 40, 844-851 15 Jabbal I., Kells D.I.C., Forstner G. & Forstner J.F.

(1976) Can. J. Biochem. 54, 707-716 16 Schrager J. & Oates M.D.G. (1971) Digestion 4,

1-12 17 Le Treut A., Lamblin G., Houdret N., Degand P.

& Roussel P. (1981) Biochimie 63,425-434 18 Slayter H.S., Lamblin G., Le Treut A., Galabert C.,

Houdret N., Degand P. & Roussel P. (1984)Eur. J. Biochem. 142, 209-218

19 Forstner J.F., Jabbal I., Qureshi R., Kells D.I.C. & Forstner G.G. (1979) Biochem. J. 181,725-732

20 Hill H.D., Reynolds J.A. & Hill R.L. (1977) J. BioL Chem. 252, 3791-3798

21 Allen A. (1978) Br. Med. Bull. 34, 28-33 22 Allen A. & Snary D. (1972) Gut 13,666-672 23 Mantle M., Mantle D., Allen A. (1981) Biochem. J.

195, 277-285 24 Mantle M., Forstner G.G. & Forstner J.J. (1984) Bio-

chem. J. 217, 159-167 25 Forstner J.F., Jabbal I. & Forstner G.G. (1973) Can.

J. Biochem. 51, 1154-1166 26 Shogren R.L., Jamieson A.M., Blackwell J. & Jen-

toft N. (1984) J. Biol. Chem. 259, 14657-14662 27 Scawen M. & Allen A. (1977) Biochem. J. 163,

363-368 28 Carlstedt I. & Sheehan J.K. (1984) Ciba Found.

Symp. 109, 137-156

Struc ture o f gastrointest inal muc ins 617

29 Carlstedt I., Lindgren H. & Sheehan J.K. (1983) Bio- chem. J. 213, 427-435

30 Carlstedt I. & Sheehan J.K. (1984) Biochem. Soc. Trans. 12, 615-617

31 Carlstedt I. & Sheehan J.K. (1985) in: Oral Interfa- cial Reactions o f Bone, Soft Tissue and Saliva (Glantz P.O., Leach S.A. & Ericson T., eds.), IRL Press, Oxford, pp. 97-105

32 Slayter H.S., Lamblin G., Le Treut A., Galabert C., Houdret N., Degand P. & Roussel P. (1984) Eur. J. Biochem. 142, 209-218

33 Bell A.E., Allen A., Morris E. & Ross-Murphy S. (1984) Trans. Biochem. Soc. 12, 648

34 Carlson D.M. (1968) J. Biol. Chem. 243,616-626 35 Hounsell E.F. & Feizi T. (1982) Med. BioL 60,

227-236 36 Feizi T. (1985) Nature (London) 314, 53-57 37 Cole L.A., Birken S. & Perini F. (1985) Biochem.

Biophys. Res. Commun. 126, 333-339 38 Snow P.M., Keizer G., Coligan J.E. & Terhorst C.

(1984) J. lmmunoL 133, 2058-2066 39 Florman H.M. & Wassarman P.M. (1985) Cell 41,

313-324 40 Lotan R., Skutelsky E., Danon D. & Sharon N.

(1975) J. BioL Chem. 250, 8518-8523 41 Picard J.K. & Feizi T. (1983) MoL bnmunoL 20,

1215-1220 42 Cooper H.S. (1982) Lab. Invest. 47, 383-390 43 Hounsell E.F. & Feizi T. (1982) Med. BioL 60,

227-236 44 Marsh W.L. (1961) Br. J. HaematoL 7, 200-209 45 Picard J., Edward D.W. & Feizi T. (1978) J. Clin.

Lab. ImmunoL 1, 119-128 46 Szulman A.E. (1960) J. Exp. Med. 111,785-807 47 Szulman A.E. (1962) J. Exp. Med. 115,977-1005 48 Szulman A.E. (1964) J. Exp. Med. 119, 503-516 49 Yuan M., Itzkowitz S.H., Palekar A., Shamsuddin

A.M., Phelps P.C., Trump B.F. & Kim Y.S. (1985) Cancer Res. 45, 4499-4511

50 Gooi H.C., Feizi T., Kapadia A., Knowles B.B., Solter D. & Evans M.J. (1981) Nature (London) 292, 156-158

51 Koprowski H., Steplewski Z., Mitchell K., Herlyn D. & Fulner P. (1979) Somat. Cell. Genet. 5, 957-972

52 Raux H., Labbe F., Fondaneche M.C., Koprowski H. & Burtin P. (1983) Int. J. Cancer 32, 315-319

53 La Mont J.T. & Ventola A.S. (1980) Biochim. Biophys. Acta 626, 234-243

54 Tettamanti G. & Pigman (1968) Arch. Biochem. Biophys. 124, 41-50

55 Wesley A., Mantle M., Man D., Qureshi R., Forst- ner G. & Forstner J. (1985) J. BioL Chem. 260, 7955 -7959

56 Edge A.S., Faltynek C.R., Hof L., Reichert L.E. Jr. & Weber P. (1981) A n a l Biochem. 118, 131-137

57 Hill H.D. Jr., Schwyzer M., Steinman H.M. & Hill R.L. (1977) J. Biol. Chem. 252, 3799-3804

58 Bara J., Loisillier F. & Burtin P. (1980) Br. J. Can- cer 41,209-221

59 Gold D. (1981) Cancer Res. 41,767-772 60 Ma J., De Boer W.G.R.M. & Nayman J. (1982) Can-

cer 49, 1664-1667 61 Qureshi R., Forstner G.G. & Forstner J.F. (1979) J.

Clin. Invest. 64, 1149-1156 62 Pant K.D., Dahlman H.L. & Goldenberg D.M.

(1977) bnmunoL Commun. 6, 411-421 63 Nardelli J., Bara J., Rosa B. & Burtin P. (1983) J.

Histochem. Cytochem. 31, 366-375 64 Nardelli J., Loridon-Rosa B., Bara J. & Burtin P.

(1984) Cancer Res. 44, 4157-4163 65 Zweibaum A., Oriol R., Dausset J., Marcelli-Barge

A., Ropartz C. & Lanset S. (1975) Tissue Antigens 6, 121-128

66 Forstner J.F., Ofosu F. & Forstner G.G. (1977) Anal. Biochem. 83,657-665

67 Roomi N., Laburthe M., Fleming N., Crowther R. & Forstner J. (1984) Am. J. Physiol. 247 (Gastroin- test. Liver PhysioL 10), GI40-GI48

68 Babczinski P. (1980) FEBS Lett. 1 i7, 207-211 69 Schachter H. & Williams D. (1982) in: Mucus in

Health and Disease - 1I. Advances in Experimen- tal Medicine and Biology, Vol. 144 (Chantler E.N., Elder J.B. & Elstein M., eds.), Plenum Press, New York, pp. 3-28

70 Roth J. (1984) J. Cell Biol. 98, 399-406 71 Elhammer A. &Kornfeld S. (1984)./. Cell. BioL 98,

327-331 72 Slomiany B.L., Takagi A., Liau Y.H., Jozwiak Z.

& Slomiany A. (1984) J. Biol. Chem. 259, 11997-12000

73 Kim K.C., Rearick J.I., Nettesheim P. & Jetten A.M. (1985) J. BioL Chem. 260, 4021-4027

74 Moyer M.P. (1983) Proc. Soc. Exp. BioL Med. 174, 12-15

75 Rattner D.W., Ito S., Rutten M.J. & Silen W. (1985) In Vitro 21,453-462

76 Pinto M., Appay M.D., Simon-Assman P., Cheva- lier G., Dracopoli N., Fogh J. & Zweibaum A. (1982) BioL Cell. 44, 193-196

77 Pinto M., Robine-Leon S., Appay M.D., Kedinger M., Triadou N., Dussaulx E., Lacroix B., Simon- Assman P., Haffen K., Fogh J. & Zweibaum A. (1983) Biol. Cell. 47, 323-330

78 Augeron C. & Laboisse C.L: (1984) CancerRes. 44, 3961-3969

79 Augeron C., Maoret J.J., Laboisse C.L. & Grasset E. (1986) in: 25 Years o f Research on the Brush Bor- der Membrane and Sodium-Coupled Transport (Alvarado F. & van Os C.H., eds.), INSERM Symposium Series, Vol. 26, Elsevier, Amsterdam (in press)