Chitin in protistan organisms

Europ.J.Protistol. 29,1-18 (1993) February 19, 1993

Review

Chitin in Protistan Organisms Distribution, Synthesis and Deposition Maria Mulisch University of KOln, Department of Zoology, Experimental Morphology, K61n

Contents I Introduction II Structure and Chemical Composition of Chitin III Methods for the Identification and Localization of Chitin IV Distribution of Chitin among the Protists V Chitin Synthesis and Chitin Deposition VI Chitin Degrading Enzymes VII Conclusion Acknowledgements References I Introduction Chitin, the polymer of N-acetyl-D-glucosamine (GlcNAc), is one of the most abundant macromolecular biological products on earth. It is present in the cell walls of fungi and in the exoskeleton of almost all invertebrates (except sponges, most anthozoa, scyphozoa, and echinoderms), and it is absent in vertebrates and most autotrophic organisms [55,56,84]. Chitin biomass production can be very high: in eutrophic freshwaters, it can reach values of 20 glm-2/year-1 [57]. Because of the great amount of chitin available, and because of the physical (mechanical strength, elasticity), chemical (biodegradability, low chemical reactivity, insolubility in water) and biological properties (e.g., positive effects on wound healing), chitin has become the base for many industrial and medical products (for review see [84]). Chitin has at least two functions in most organisms: it protects cells and organisms against mechanical or chemical stress from the environment, and it also supports and determines their shape. The synthesis, deposition, modification and degradation of chitin are therefore major features of morphogenesis. As chitin formation occurs in the life cycle of many fungi, nematodes, and arthropods parasitic on plants or vertebrates which do not produce chitin, there is an increasing interest in the understanding of chitin synthesis and degradation, in order to aid the design of therapeutical drugs, fungicides or insecticides. © 1993 by Gustav Fischer Verlag, Stuttgart

During the last years, knowledge on chitin metabolism has tremendously increased (for reviews see, e.g., [22, 116]). However, most interest on chitin formation has been focussed on fungi (for reviews see [8, 22]), less on arthropods [18], but relatively little is known about chitin synthesis in protists. Although supporting chitinous structures are not uncommon in unicellular eukaryotes, reports of chitin are rare, and comprehensive studies are totally lacking. Nevertheless, protists provide interesting models for the study of chitin metabolism, as chitin synthesis is localized at defined sites of separate cells, and at certain often inducible - stages of their life cycle. In the following review, I will therefore try to summarize the information available about the distribution and formation of chitin in protists of the last 1 - 2 decades. Because the clear identification of chitin is an absolute prerequisite for investigations on the pathways of chitin synthesis and deposition, relevant techniques and methods will be briefly and critically reviewed. The results will be compared with those on chitin formation in fungi and arthropods. This may lead to a better understanding of chitin structure and deposition in protists. In many organisms, the genesis, shape and chemical composition of chitinous structures do not only depend on synthesis, but also on degradation and modification of chitin [39]. Therefore, chitin degradation is an essential aspect of chitin metabolism. Some information about chitinases of protists will be mentioned and critically discussed in the last section of the article. II Structure and Chemical composition of Chitin Chitin is a linear polysaccharide of ~(1-4)-linked N-acetyl-D-glucosamine units (Fig. 1). It may also contain glucosamine units in different proportions [85]. Mainly deacetylated chitin is called chitosan. The chain lengths vary: in fungi, each chain may consist of 100 (Saccharomyces) - 8000 (Scylla serratia) units [22]. The polymers assemble laterally to form microfibrils by strong hydrogen bonds between the> N -H-group of one chain with the> C =O-group of a neighbouring chain. Extracellular microfi0932-4739/93/0029-0001 $3.5 0/0

2 . M. Mulisch

cellulose [71]. In exoskeletons of anim als, chitin-prot~in associations are predominant that may have a crystalline order [12]. The proteinous matrix may be hardened ~y deposition of calcium carbo~ate ~nd phosp,hate .as In crustace an cuticles, or by tanrung WIth phenolic derivates as in insect cuticles [84, 85]. In protist chitinous structures proteins (mainly glycopro teins) and mucopol ysaccharides have been detected (see Table 1).

o

ill Methods for the Identification and Localization of Chitin Fig. 1. Repeating unit of chitin (modified from [22]).

bril assembly is inhibited by several dyes that occupy these groups thus preventing the formation of the hydrogen bonds , such as Calcofluor white [46]. Three crystallographic forms of chitin can be distinguished, which differ in the relative orientation of t~e polymer chains (Fig. 2). Their distrib~tion, at least In animals, is not related to taxonomy, as different forms may occur in one organism [84]. In a-chitin, the most abundant form in fungi and arthropods, adjacent chains are o,riented antiparallel [84]. ~-Chitin consists of parallel chains and occurs, for example in the Loligo pen [84], pogonophore tubes [11], in the spines of centric diatoms [47], and .i~ t~e lorica of the ciliate Eufolliculina uhligi [81]. ~-ChItIn IS highly crystalline and considerably ~wells in wa,ter [84]. In y-chitin, which has been detected In a fe~ anImal,s [84], two of thre e adjacent chains are parallel while the third one is oriented in the opposite direction. The microfibrils in insect cuticles have a diameter of about 2,5-3 nm [84]. In crustacea and fungi, they are wider (diameter 20-25 nm) [84]. This size is comparable to that (15-20 nm) of the chitin fibrils in the lorica of the ciliate Eufolliculina uhligi [81] (Figs. 12-13 ), whereas the ~- chitin in centric diatom s form s microfibrils up to 30 nm that may assemble to ribbons with a diameter of 50 .nm [50]. The crystalline chitin is relatively stable against treatments with hot alkal i (up to 20 % Na OH or KOH ), , , organic solvents, or dilut e acids [8~] ... In most supporting structures , chitin IS associated With other substances. In fungal cell walls, these are essentially other pol ysaccharides, mainly c~itosan a n~ ,glucan, ~­ glucan being possibly covalently linked to chlt~n. [~2, 24]. The cell wall of the zoosporic fungus Rhizidiomyces apopbysatus is reported to contain chitin as well as

a-chitin

it

it~

~-chitin

y-chitin

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Fig. 2. Ori entation of the polymer chains in the three cristallographic forms of chitin.

Many histochemical methods, mainly used in the past, are not exclusively specific for chitin [84]. This refers also to the staining with Calcofluor white .(Cellofluor) .(for method see [101]) that binds to ~-1,4-llnked and mixed linked polysaccharid es (including cellulose), and for the chlor-iodine-zinc method [47]. X-ray diffraction and IR-spectro scopy are the most reliable methods for the determination of chitin [84]. How ever, only few investigators have used these t~ch­ niques for the analysis of extracellular pro~ucts ~f pr?tI sts. Chitin has been demon strated by x-ray diffraction In the lorica of the chrysoflagellate Poterioochromon~s stipit~ta [47], and in the cyst walls of the amoeboid protists Entamoeba invadens [3] and Chaos illinoisensis [111]. The x-ray diffraction pattern s of fibrils from spines ?f ce?~ric diatoms [50] and from the lorica of the heterotrich clh.a~e Eufolliculina uhligi [81] (Fig. 3) resemble that of ~-chltln (for reviews see [48, 49]). Another approach is to identify chitin by the action of chitinase. After extraction with hot alkali, Bussers and Jeun iaux [18,20,54-56] have tr~ated a ,n.umber ,of protecting structures of protists, ma~nl y of c.tl~ates, With purified chitinases (commercially available chitinases may contain cont aminations of other enzymes!). The degradation of the structures indicates their chitin nature . Chitinases hydrolyze the polysaccharid e to oligosaccharides, after longer incubati on times these may be further degrad ed to GlcNAc (see below). The digest!on products can be determined by several methods (for reviews see, e.g., [22, 84]). Chitinases work more ef~ectiv7ly ~n ?a scent (single) chitin chains than on crystalhne microfibril s [22], and they may fail to attack fibrils masked by other substances. Therefore, extraction of the material by organic solvents (usually chlo~~form/m ethanol~ and by hot alkali or protease before chitinase treatment IS recommended. The chitinase techniqu e may be also applied on ultrathin sections [109]. Ho wever, as chitin f~b rils (in contra~t to chito san [93]) do not stain in conventional electron microscopical preparation s, they may just beco~e in.visible if the embedding matrix is removed by contammatmg enzymes of the chitinase. The digestion of the chitinous structures must be verified by careful controls (e.g., by staining of the chitina se treated mat erial with lectin-gold complexes). More reliable methods for the ultr astructural localization of chitin involve treatments of the sections with gold-coupled chitin-binding proteins. These may include

Chitin in Protists . 3 Table 1. Distribution of chitin in protists Species

Classification

Giardia lamblia Thelahonia * Rhizidiomyces apopbysatus

Further Systematic Specification

Conformation

Localization

Other components

Ref.

Diplomonada Microsporidia Hypochytriomycota

cyst wall cyst wall cell wall

glycoproteins ? cellulose

[70, 89, 130] [30] [34]

Chytridiomycota Plasmodiophoromycota Rhizopoda Testacealobosia Rhizopoda (?) Rhizopoda Granuloreticulosea Chlorophyta Chlorophyta Chlorophyta Bacillariophyta Bacillariophyta Chrysophyta

cell wall cell wall ? ? ? ? ? ? ? B-chitin B-chitin a-chitin?

test cyst wall cyst wall shell cell wall cell wall cell wall spines spmes stalk

Folliculinopsis producta Parafolliculina violacea Eufolliculina uhligi

Oomycetes Bicosoecida ,. Choanoflagellates Choanoflagellates Ciliophora Ciliophora Ciliophora

Salpingoecidae* Codonosigidae' Heterotrichia Heterotrichia Heterotrichia

? ? ? ? ? B-chitin

cell wall lorica theca theca lorica lorica lorica

Blepharisma spp. Phacodinium metschnikoffi Climacostomum virens Fabrea salina

Ciliophora Ciliophora Ciliophora Ciliophora

Heterotrichia Heterotrichia Heterotrichia Heterotrichia

cyst cyst cyst cyst

Bursaria truncatella Oxytricha bifaria Euplotes spp.

Ciliophora Ciliophora Ciliophora

Colpodea Stichotrichida Hypotrichida

cyst wall cyst wall cyst wall

Cothurina sp. Telotrochidium henneguyi Hyalophysa chattonii Nassulopsis lagenula Furgasonia blochmanni Pseudomicrothorax spp.

Ciliophora Ciliophora Ciliophora Ciliophora Ciliophora Ciliophora

Peritrichia Peritrichia Apostomatida Nassophorea Nassophorea Nassophorea

lorica cyst wall cyst wall cyst wall cyst wall cyst wall

Nassula spp.

Ciliophora

Nassophorea

cyst wall

Plasmodiophora brassicae Plagiopyxa sp.' Entamoeba spp. Chaos illinoisensis Allogromia sp. * Ulva lactuca " Valonia ventricosa * Pitophora oedogonia Thalassiosira spp. Cyclotella spp. Poterioochromonas stipitata

wall wall wall wall

[75] [9J ? ? ? ? cellulose cellulose ?

iron ? ? proteins ? pigment, proteins, acid mucopolysaccharides, anorganic material pigment, proteins? ? ? mucopolysaccharides, proteins silica, proteins proteins mucopolysaccharides, proteins [21] ? ? proteins ?

?

glycoproteins, proteins, mucopolysaccharides proteins, acid mucopolysaccharides

[84] [1,5] [84, 111] [84] [84] [84J [93] [50,135] [50, 135] [47-49] [7] [31] [16] [16] [21J [20] [78, 80, 81] [20,79] [20] [20] [18,20] [18,20] [109] [20,41] [20] [20] [61,62] [20J [20] [18,20,79J [18,20J

* methods for identification of chitin not described (a) chitinase, (b) lectins binding to GlcNAc-residues, such as wheat germ agglutinin (WGA) [1,41,61, 79, 123] (Figs. 14-16) or tomato lectin (ToL) [130], or (c) antibodies against chitin oligomers [89] and polymers [127, 128]. Succinylated WGA is more specific for chitin than WGA, because the latter binds also to sialic acid residues [130]. The label can be enhanced by two-step procedures, for example by first a treatment of sections with lectin and then with a colloidal gold-coupled antilectin [94]. Although WGA binds to chitin in sections of conventionally fixed and embedded material [79], it may

show a higher and more specific affinity for chitin structures fixed without osmium tetroxide and embedded in hydrophilic media such as LR white or Lowicryl [96]. For the light microscopical detection of chitin, the chitin-binding proteins can be coupled to fluorescent dyes (e.g., FITC), and applied to fixed or unfixed, whole-mount (Figs. 5 -7) or sectioned material [126]. All cytochemical methods need a number of careful controls. Usually, these include competitive inhibition of the binding reaction by chitin oligomers (especially triacetylchitotriose [96]) and alternative staining with lectins specific for other sugars.

4 . M. Mulisch The results are more reliable, however, if additionally the material is preincubated in chitinase. Chitin in biological sections can also be detected by diffraction contrast electron microscopy [37]. IV Distribution of Chitin among the Protists Reviewing the occurrence of chitin in extracellular structures of protists is difficult. In the past, the term chitin was often used with little care about the chemical composition of the polymer. Chitin was mainly defined by morphological criteria and histological staining properties, and on the base of these observations, many structures were described as 'chitinous' or 'of chitinous nature', but this is not a satisfactory means of identifying chitin. On the other hand, some cells may produce only small amounts of chitin which cannot be detected by insensitive methods. The following summary is therefore provisional and includes mostly those which satisfy contemporary criteria. Chitin has been detected in spines, stalks, loricae and cysts of protists of diverse taxonomic groups (summarized in Table 1). Primitive groups. Molecular and structural information indicate that the most primitive protists were amitochondriate [92, 117]. Collectively these have been referred to as the Archezoa [25] or the Hypochondria [92]. The primitively amitochondriate protists include: the Microsporidia, pelobionts, retortamonads, diplomonads, oxymonads, trichomonads, hyperrnastigids and possibly Entamoeba. In the spore wall of several members of the Microsporidia, 8 - 9 nm-fibrils have been identified that are suspected to represent chitin [64, 65, 124]. Investigations by fluorescence microscopy on Thelahonia spores support this hypothesis [30]. Detailed information is available about the composition of the cyst wall of a member of the diplomonads. Giardia lamblia is a parasite of the human intestine where it causes diarrhea. The disease is transmitted by infective cysts which are excreted in the feces of infected hosts into the external environment. The resistent cyst walls contain chitin as a major structural component [130]. This has been confirmed by chitinase digestion, lectin-binding studies [129, 130] and immunocytochemistry using chitin antibodies [89]. Amoeboid protists. Many different evolutionary lineages of protists have given rise to amoeboid organisms the relatively primitive Heterolobosea having produced acrasid slime molds and vahlkampfiid amoeba, whereas Acanthamoeba appears to have its origin very late in eukaryotic evolution and in proximity to the green algae [117]. We do not know how many times amoeboidy has appeared, but a rational recategorization of the amoeba must await further work. The capacity to synthesize cellulose is widespread among amoeboid protists, and is evident in Heterolobosea (Schizopyrenus, Naegleria, and acrasid slime molds), the Euamoebae (Acanthamoeba and Hartmanella), and in the Eumycetozoea ([15,36, 102, 120, 133], see also [13] for references). Molecular studies indicate that these groups are not closely related. The formation of chitinous cysts

has been recorded only rarely (see Table 1). It Occurs in members of the rhizopoda, and it has been reported also from a foraminifer, Allogromia sp. [84]. The organicshells of other granuloreticulosea are described as complexes of protein and mucopolysaccharides ('tectin') [63]. Most detailed studies deal with the demonstration [2, 3], localization [1] and synthesis [5, 27] of chitin in the cyst wall of two species of the parasitic, amitochondriate Entamoeba. The x-ray diffraction pattern of the alkali-resistent fibrils [3] and their specific labelling by WGA-gold [1] clearly indicate chitin as a major structural component. Flagellated protists. The flagellated protists are here regarded as those taxa previously assigned to the flagellated algae and to the heterotrophic flagellates. These groups are now recognized as polyphyletic and include organisms that have appeared at many different times during eukaryotic evolution. Of the algal protists, many groups regarded as algae (red algae, green algae, etc.) produce cellulose or glycoproteinous walls (for reviews see [69, 126]). An exception is provided by some members of the Chlorophyta [84,93] which produce chitin material. Chitin formation is also rare among heterotrophic flagellates. In the choanoflagellate families Salpingoecidae and Codonosigidae, the thecae are reported to contain either chitin or cellulose or mucopolysaccharides [16]. Chitin has been detected in the recently 'recognized stramenopiles [91] (see Table 1), a group which 'includes chrysophytes, diatoms, brown algae, oomycetes, some heterotrophic protists, and other rdlated.organisms.Only some of the groups previously assigned to the chrysophytes appear to be able to synthesize chitin.Others,suchas Dinobryon, produce cellulose [43]. In 'Poterioochromonas, the wine-glass-shaped lorica contains a ;framework of helically arranged chitin fibrils. Their ultrastructure, composition and formation have been carefully .analyzed by x-ray microanalysis, electron microscopical and biochemical methods [46, 47, 126]. The diatoms form another part of the stramenopiles and chitin has been reported here [45,50]. The centric diatoms produce long, stiff appendages to reduce the sinking velocity. The appendages of Cyclotella and Thalassiosira consist of the highly crystalline ~-chitin (for review see [49]). Heterotrophic stramenopiles in which chitin formation has been reported include the bicosoecids [31], and the oomycetcs [9]. The chemical composition of the wall scales of labyrinthulids is not known [100]. The Thraustochytriidae produce scales of highly sulfated galactan, but no chitin [26]. Ciliates. This group is much more homogeneous than the other ones mentioned [68]. A number of ciliate cysts and loricae appear to contain chitin (see Table 1). Its presence has been demonstrated by chitinase digestion in many species especially of heterotrichs and peritrichs, in Bursaria, and in the nassophorean Nassula and Pseudomicrothorax, whereas it appears to be absent in hypotrichs (except Euplotes), in Colpoda, Bresslaua, Woodruffia, Didinium, and tintinnids, respectively [18,20]. In the litostome ciliate Didinium, polysaccharides of unknown nature have been identified [108]. The absence of chitin in

Chitin in Protists . 5 tint innid loricae argues aga inst a s uggested relat ion ship between tintinn ids and the chiti nozoa [71], fossile, cystlike stru ctures from certa in sedime nts. Lat er published cyto chemical and x-ray analyses mainly confirm this pattern. of chitin distri buti on (for references see below) : Chitin has not been found among colpodid ciliates (except Bursaria): it is not present in the sorocarp of Sorogena stoianovitchae [14], an d the cyst walls of Colpoda [11 9] and Grossgloekneria acuta. The extracellular structures of Sorogena [14J and Colpoda [10, 73, 110J appear mainly to consist o f glycoproteins with GlcNAc residues. Cysts or loricae of heterotrichs so far analyzed contain chitin. The loricae of folliculinids (Fig. 3) consist of 20 nm fibrils of ~-chitin which are embedded in a matrix of pigment s, proteins and mucop olysaccharides [81]. Th e cyst walls of Stentor, Blepharisma, Climacostomum an d Fabrea appea r ultrastructurally similar (1 06], the thr ee latter have been shown to be chitino us by a treatm ent with chitinase [18, 20]. Chitin occurs also in aposte me ciliates. Hyalophysa ehattoni forms two structura lly different cysts during its life cycle. Bot h are reported to contain chitin [61,62]. The phoreric cyst is sur rounded by two wa ll layers, the inner appears to be chitino us, the outer layer, as well as an attachment peduncle, are made of protein [62]. Th e wa ll of the reprod uctive cyst lacks the pro tein parts an d is much th icker [61].

Th ere are conflicting results as to the presence of chitin a mong hypotrich ciliates. Whereas Bussers and [euni aux [20] did not observe degrad ati on of the cysts of Oxytricha sp. by purified chitinase, Rosati et aI. (109J report ed the disapp ear ance of cystic layers of Oxy tricha bitaria after a combined pr orease-chitinase treatment on ultrathin section s. Unfort unately, Rosat i et al. did not cont rol the digestion of the chitin by, for example, postlab elling of the sections with WGA- gold (see section on meth od s above). Furth er studies may decide, if (all?) oxytrichids are able to synthesize chitin. Chitin has been more clearly demo nstra ted in the cyst of Euplotes muscicola [4 1, 123]. In all ot her hypotrich ciliates ana lyzed, the cysts apparently contai n no chitin and only glycoproteins, often with GIcNA c-rich carbohydrate mo ieties [42, 74]. Chitin distribut ion may therefore reflect the phylogenetic distance between Euplotes and other hypotrichs (e.g., [112]). The situa tion is also not clear among the peritrichs. Chitinase digested the lor ica of Cothurnia sp. [20], but histochemical sta ining reactions failed to detect chitin in the loricae of Platycola decumbens [131] and Thuricola [olliculata (38]. As in the case of the hypotrichs, more careful studies are needed to confir m the presence or abse nce of chitino us stru ctu res in this group. Th e ciliates are now clustered with Apicomplexa and dinoflagellates, and referred to as alveolates [134]. Th e spo re wa lls of memb ers of the Apicomplexa apparentl y do not contai n chitin [127]. Dinoflagellat es do no t synthesize chitin either, but cellulose [71).

Figs. 3-7. Chitin in the ciliate Eu(olliculina uhligi. - Fig. 3. SEM micrograph ofthe bottle-shaped lorica which is fixed to the substrate by a fibrillar basal plate (arrow). - Figs. 4- 5. Light micrographs of a swarmer after triggering of exocytosis by alcian blue. The extruded material which forms a cysr-like layer (arrow) around the eel! has binding sites to FITC-coupled WGA (Fig. 5). Figs. 6-7. Fluorescence micrographs of loricae stained byWGA-FITC. - Fig. 6. Optical section through the lorica. - Fig. 7. Fibrillar basal plate. Bar = 100 urn.

6 . M. Mulisch

Other groups. Additionally to the groups mentioned above, chitin has been detected in organisms with uncertain relationship to the protists, such as the Myxozoa which are discussed to be related to the Cnidaria (e.g., [29]). The polar capsules of the Myxozoa are surrounded by a thick extracellular wall composed of two layers. The inner, transparent one is reported to be resistent to alkaline hydrolysis [66]. Chytridiomycetes share many features including chitinous walls with the fungi [7]. This refers also to the plasmodiophorids [75]. Pneumocystis was formerly regarded as member of the protists. Pneumocystis carinii is a cause of human pneumonia infecting especially the lungs of patients suffering from AIDS. Polyclonal antisera against chitin oligomers and polymers recognized cysts and trophozoites in the host tissue [127]. Both, the presence of chitin in trophozoites [127] and recent rRNA sequence analysis confirm the phylogenetic association of Pneumocystis with the fungi [32, 132]. The account of chitin distribution is certainly far from being complete, as only relatively few detailed studies on this subject have been published. Additionally, research interest has focussed on some groups, especially on human pathogens. The knowledge of the chemistry and development of their protective layers offers the opportunity to design strategies to interrupt their life cycle and to control the spread of the disease. The little information available suggests that chitin synthesis might be an ancient feature of eukaryotes [54] as it has been reported in the two groups emerging from the base of the eukaryotic radiation. However, it is expressed

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only in certain groups. The capacity to synthesize chitin appears to be lost in most autotrophic organisms, probably because they don't have enough spare nitrogen to build in their extracellular material. Plenty of nitrogen is provided by proteins of the food of heterotrophic organisms [84]. The supply with nitrogen therefore might be one limiting factor for the expression of chitin synthesis. The chemical composition of protective layers also might be modified as a strategy against enzymatic attacks of predators, and it is possibly useful for protists living in soil where external chitinolytic enzymes (for example secreted by fungi) may accumulate. The structure and chemical composition of protistan extracellular structures - like in algae and fungi [84] probably reflects phylogenetic relationships (compare [20,28, 72, 103]). However, the chitin distribution is heterogeneous even among the ciliates, where most species (unfortunately only of certain groups) were investigated (Fig. 8). Chitin synthesis therefore appears to be not a character to discriminate eukaryotic groups, but it may help to confirm relationships.

V Chitin Synthesis and Chitin Deposition

Chitin Synthesis in Fungi and Arthropods Chitin synthesis is best understood in fungi, especially in the yeast Saccharomyces. Like many protists, these organisms are easy to cultivate in large numbers, chitin can be

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Chitin in Protists . 7 formed by all cells at defined stages and at defined sites, and therefore the pathway of chitin synthesis is accessible to biochemical and genetical analyses. As a number of reviews [8,22,24] and a tremendous amount of original papers are available on this subject, I will here just give a short summary of recent results and models, knowledge of which is necessary for the discussion on chitin formation in protists. In fungi, the chitin synthetases are bound to the plasma membrane, to which they are said to be transported by small vesicles (diameter less than 100 nm), the so-called chitosomes [8, 58]. Within the membrane, the inactive zymogen (= chitin synthase) may be activated by proteolytic enzymes (in vitro) or probably by dephosphorylation (in vivo) [22]. The pathway of chitin synthesis is depicted in Fig. 9 [84]. Chitin synthesis appears to be a transmem-

Glucose ATP hexokinase v ADP

" Glucose-6-phosphate plucose phosphate Isomerase

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glucosamine acid

glutamine-fructose 6-phospha e aminotransferase

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phosphoacetyl glusos· amine mutase

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N-Acetylglucosamine-1-phosphate - UTP UDP·N·acetylgluCos· amine pyrophosphorylase "

l> pyrop hosphate

UDP-N-Acetylglucosamine chitin synthetase (chitin doacetylase) - c· UDP

® Fig. 9. Pathway of chitin synthesis (redrawn from [118]).

brane event: UDP-GlcNAc approaches the plasma membrane from the cytoplasm, and, by the help of chitin synthetase, adds the amino sugar to the growing polysaccharide chain that is simultaneously extruded [22] (Fig. 10). No external energy is needed for this process [22]. If the synthetases are tightly packed in the plasma membrane, the closely spaced chitin chains may crystallize outside the cell directly into microfibrils; otherwise single, separate chains may appear that would be susceptible to enzymatic modifications (e.g., deacetylation to chitosan) [8] (Fig. 10). In Saccharomyces cerevisiae, three chitin synthases (Chs 1, Chs 2, Chs 3) have been characterized, and their structural (CHS 1, CHS 2) and regulating genes (CAL 1) have been cloned and sequenced [17,114,122]. Fungal chitin synthetase can be competitively inhibited by polyoxins (produced by Streptomyces cacaoi) and nikkomycins (produced by Streptomyces tentae), that both are analogues of the substrate UDP-GlcNAc [22,23]. Chitin synthesis in arthropods is less understood (for review see [118]). This is certainly due to the methodological difficulties dealing with a multicellular system. During the molting period many cells are involved not only in forming a complex, multi-layered cuticle of different components (one of which is chitin), but also in the degradation of part of the old cuticle at the same time. Some progress has been made by studying insect cell cultures which secrete only small amounts of chitinous material (e.g., [60,67,99]). Horst [51] solubilized and partly purified a chitin synthetase from the larvae of the crustacean Artemia. The studies from Horst and others suggest that chitin synthesis in insects and crustacea begins in the rER with a protein which becomes glycosylated, and to which a chitin chain is added in the Golgi apparatus (via the chitin synthetase) [51,52,59,60,99]. The resulting chitop rote in is thought to be extruded by exocytosis [51]. Chitin synthesis in arthropods appears to be a two-step process via lipidlinked intermediates, thus resembling the synthesis of glycoproteins [52]. This hypothesis is further supported by studies with protein synthesis inhibitors and inhibitors of N-glycosylation (e.g., tunicamycin), which both inhibit chitin formation in arthropods [52,99]. Synthesis in vivo can be also specifically inhibited by insecticides of the benzoylphenylurea type (e.g., difluobenzuron) and, to a lesser extent, by inhibitors of fungal chitin synthetase (e.g., polyoxins) [22]. The mechanism of the action of benzoylphenylurea type inhibitors is still unclear. Studies on activation and inhibition of chitin synthetases in various animals suggest that, although there might be an ancestral route of chitin synthesis, the fine regulation varies among the species [118]. A number of different chitoproteins from arthropods have been isolated and biochemically characterized [51,60,99]. Typically, these are unusually resistent to proteases, probably due to the protection by the chitin moieties [60]. Kramerov et al. [60] propose that the chitoproteins of Drosophila consist of a polypeptide backbone to which the carbohydrate chains are linked by O-glycosidic bonds (Fig. 11).

8 . M. Mulisch

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Chitin Synthesis in Protists Only a few rep orts deal with the pathway of chitin synthesis and deposition in protists (for references see below) . Th ese are mai nly cyto logical studies, sometimes combined with cytochemical investigat ion s. In vitro studies on the chitin synthetases are published only from Entamoebainvadens. Many aspects of the in vivo pathway of chi tin formation have been recorded by cytological, biochemical and immuncytochemical investigations of the ciliate Eufolliculina uhligi.

Cyto logica l studies indica te th at in mos t protists so far analyze d, the chitin synthases are localized in the plasma memb ran e (for reviews see [48, 49]). In the centric diatom Thalassiosira, the chitin spines grow fro m conica l invagina tio ns of the plasma memb rane which have different iated coa t regions on both faces [44, 45]. In the plasma membrane of the cytoplasmic tail of the chryso phyte Poterioochromonas stipitata, aggrega tes of intramembrane particles (IM Ps) have been ob served by freeze-fracturing that, however, are not the chitin synt hases [48]. Herth [46] proposes that the chitin synthetases are moved by the

Chitin in Protists . 9

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forces of chain elongation, cytoplasmic microtubules channeling these movements and contributing to extracellular microfibril orientation (see [46] and [49] for details). Difluobenzuron, an inhibitor of insect chitin synthesis, had no effect on chitin formation in these protists.

In describing cyst wall formation in the apostome ciliate

Hyalophysa chattoni, Landers [61,62] ruled out the

involvement of cytoplasmic vesicles or extrusomes in chitin formation. No exocytosis has been observed during secretion of the cyst wall in the hypotrich ciliate Euplotes muscicola [41]. Therefore, in these ciliates, the chitin synthetases are probably localized in the plasma membrane. In Entamoeba invadens, the cyst is formed inside the trophozoite [4]. A particulate chitin synthetase and a soluble chitin synthetase have been recently characterized in encysting E. invadens, both being developmentally regulated [27]. Like in fungi, they catalyze the stepwise addition of GlcNAc from UDP-GlcNAc. In contrast to the situation in fungi, none of the amoebal chitin synthases was activated by proteolytic enzymes. Synthesis in vitro was neither inhibited by polyoxin D nor by nikkomycin [27], although both fungal chitin synthesis inhibitors block cyst wall formation in E. invadens in vivo [5]. Tunicamycin had no effect on chitin formation, suggesting that chitin synthesis in Entamoeba more closely resembles that in fungi than that in arthropods [5]. In some chitin synthesizing protists (Blepharisma [105,107], Eufolliculina [48], Giardia [33, 104]), membrane-bound precursors are found to be involved in wall formation. In the heterotrich ciliate Eufolliculina uhligi, the chitin fibrils of the lorica (Figs. 12 -13) are deposited by a motile, non-feeding swarmer [78] which is produced by binary fission of the trophont [83]. The cortex of the

13 Figs. 12-13. Chitin fibrils of Eufolliculina uhligi. - Fig. 12. SEM micrograph of the surface of a newly formed lorica. - Fig. 13. The alkali-extracted chitin fibrils show typical kinking-sites (arrowheads). In places, they split into microfibrils (arrows) (from [81]). Bar = 0.1 urn.

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Figs. 14-16. Ultrathin sections through the cortex of the lorica secreting Eufolliculina uhligi, postlabelled by WGA-gold. Only few gold particles mark material inside cortical vesicles (arrowheads). - Fig. 14. Formaldehyde/glutaraldehyde-fixed, Lowicryl-embedded cell. The lorica (L) is heavily labelled. - Figs. 15-16. Ruthenium red/OsOa-fixed, Epon-embedded cell at the onset of lorica (L) formation. - Fig. 15. Polysaccharide material is extruded by exocytosis of cortical vesicles.- Fig. 16. Fibrillar, WGA-binding material (arrows), often attached to cilia (C), appears at a certain distance of the cell. Bar = 1 urn,

Chitin in Protists . 11 swarmer contains numerous vesicles that attach to the cytoplasmic face of the plasma membrane [82]. The vesicles contain heterogeneous material [77 ]. During lorica formation, the vesicles release their contents to the outside via exoc ytosis (Fig. 15). Swarmers which previousl y ha ve been triggered to extrude the vesicle content by a treatment with the cationic dye alcian blue, and which subsequently are induced to undergo metamorphosis [76] do not build a lorica. Thi s indicates that the lorica is made exclu sively from exoc ytosed material. As the sites of exocytosis correspond to the sites of chitin dep osition, the extrusive vesicles in the swarmer were suspected [81] to contain chitin in a polymeric state. This is confirmed by biochemical studies of 3H-GlcNAc-labeled swarmers in which a great amount of the incorporated radioactivity is bound to macromolecular, alkali-resistent material. The material is digestible with chitinase. The artifici al triggering of extrusion of the vesicle material does not lead to lorica form ation nor to fibril assembl y (Figs. 4, 5). Binding studies with colloidal gold-coupled WGA on ultrathin sections of Lowicr yl-embedd ed swarmers before and dur ing lorica formation (Figs. 14-1 6), and with FITCconjugated WGA on the extruded vesicle contents (Fig. 5) indicate that the chitin chains inside the vesicles are somehow masked. Hence, they might be further proces sed during lorica formation - before or after extrusion. The cortical vesicles appear to be produced during the trophic

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Figs. 17-18. Blotted, SDS-gel -electrophoretically separatedproteins of swarmers of Eufolliculina uhligi incubated in anti-chitin (Fig. 17) and in WGA (Fig. 18). - Fig. 17. The antibody against polymeric chitin recognizes two distinct protein bands (arrowheads). - Fig. 18. The same bands (beside others) appear after incubation of the blotted proteins in WGA. mw = molecular weight (x 10'3).

stage of E. uhligi from the rER and the Golgi apparatus [77]. Thi s resembles the situation in arthropods [51] where chitin synthesis is thought to be link ed to the glycoprotein pathway [51, 52, 59, 60, 99]. We have therefore analyzed the SDS-gel-electrophoretically separated proteins of the swa rmers by immunoblotting ana lysis using a pol yclonal antibody against polymeric chitin. Th e antibody has been demonstrated to detect chitin in extracellular structures of diverse organisms [35, 51, 127, 128 ]. Thi s antibody binds specifically to certain proteins of the swarmer which have also binding sites for WGA (Figs. 17, 18 ). The observations strongly suggest that the swarmers contain chitin polymers which are covalently linked to protein. Chitin formation in E. uhligi (and possibly in other heterotrich ciliates) therefore appears to resemble the more complicated two-step process in arthropods [48]. E. uhligi works out to be a promising system for the study of chitin formation, as early (synthesis of pr ecursors) and late steps (deposition of chitin and fibril formation) appear to occur in different types of cells, and thu s can be anal yzed separately. In addition, chitin deposition is inducible [76], which offers the opportunity of biochemical investigations of defined stages. Th e indications for the existence of (at least) two different pathwa ys of chitin synthesis among the protists and even among the ciliates - suggest that certain step s can be easily modified . The data are not yet sufficient to follow the phylogenetic route(s) of the mechanisms of chitin assembl y. The y make clear that proti sts pro vide interesting models for the study of chitin synthesis, and the y will certa inly help to understand ch itin form ation in higher taxa such as arthropods.

VI Chitin Degrading Enzymes The ability to degrade chitin is more widely distributed am on g the prokaryotic and eukaryotic organisms than the ab ility to synthesize chitin [84]. For exa mple, chitinolytic enzymes are expressed in all insect ivorous animals (including verte brates) [84], and in man y higher plants [86]. In chitin-pro ducing organisms, these enzymes are often involved in morphogenesis, such as growth and division of fungal hyphae [24,40] or moulting in arthropods [118 ]. Chitinolytic enzymes should therefore also be present among the protists where they might be used for feeding (in mycophagou s species, para sites of arthropods, etc. ) or might play a role in morphogenetic processes such as excysta tion. Three enzymatic system s are kno wn to degrade the GlcN Ac polymer: lysozymes, exo chitinases and endochitinases (= ~-hexasominida ses ) (for review see [22,84]). Lysozyrnes (endo-~-N-acetylmuraminida ses) degrade the glycan of the bacterial cell walls, but a number of them (e.g., egg white lysozyme) ma y also hydrolyze chitin, thu s overlapping in their specificiti es with chitinases, Exochitinases attack the chitin chain s from the free ends, liberating single aminosugar units, whereas endochitinases act within the chains and produce oligomers of random lengths (Fig. 10). In a strict sense, on ly exo chitinases are regarded

12 . M. Mulisch

as chitinases [22]. The enzymes can be distinguished by using defined substrates (e.g., [6]) or inhibitors. Endochitinases of fungi, arthropods and the amoeboid protist Entamoeba invadens are competitively inhibited by the azo-oligosaccharide Allosamidin [95, 125]. Chitinases may also act as transglycosylases involved in cross-linking chitin to other wall components [39]. As the chitinolytic enzymes differ in their final products, their characterization is usually based on biochemical studies. Crude or purified extracts of cells are incubated with an appropriate, labelled substrate, and the degradation products are then photometrically or chromatographically determined (see [22] for details). The sensitivity of recently developed tests is very high (see e.g., [53]). There are methods for the demonstration of chitinases among electrophoretically separated proteins [90]. Despite the interesting and promising aspects that are offered by analyzing the degradation of chitin and the enzymatic systems involved, only a few serious studies have been carried out on the properties, localization and regulation of chitinases in protists [53,113, 121, 125]. Many earlier results are still doubtful because insensitive methods were used. Also, in non-axenically grown cultures contaminations of the preparation by chitin degrading enzymes from other organisms, especially from bacteria, are always a problem and can be only excluded by careful controls. The following example illustrates this problem. The mycophagous ciliate Grossglockneria acuta (Fig. 19) attacks fungal hyphae by perforating the hyphal

G

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wall and ingesting the cytoplasmic content [97,98]. The perforation appears strictly localized at the site where the oral apparatus of the ciliate contacts the hypha (Fig. 22), thus resembling the situation during feeding by mycophagous amoeba [87,88]. This observation suggests [97, 98] that chitinolytic enzymes act at the oral apparatus of the ciliate (Fig. 21). Exocytosis of small vesicles occurs in this area during the attack (Figs. 22 - 24). This suggests that the chitinases are stored in the cell and locally secreted in response to an unknown signal. The presence of active chitinases can be demonstrated in feeding but not in starved cells. Controls and blotting analysis of the chitinbinding proteins of the ciliate with a polyclonal antibody against plant (potato) chitinases have shown that the extract was highly contaminated by chitinases that were probably secreted into the medium by the host Mucor mucedo (Fig. 20). As the chitinase activity of Grossglockneria acuta is suspected to be quite low (the cell wall of Mucor contains much chitosan [115]) and may be present only during a short period and only in contact with the fungus, a reliable identification/characterization of the ciliate enzymes has not yet proved possible. Methods for the detection of the enzymes in single cells need to be developed at least for investigations on chitinases of strictly mycophagous organisms. A promising approach would be the production of appropriate antibodies which could be used for the demonstration and localization of chitinolytic enzymes at the ultrastructural level. Combined biochemical and immunocytochemical studies could also help to clarify the role of chitinolytic enzymes in the formation and degradation of chitin containing extracellular structures of protist organisms.

VII Conclusion

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Figs. 19-20. Grossglockneria acuta. - Fig. 19. Line drawing of the cell. OA = oral apparatus. Bar = 10 urn, - Fig. 20. Blot of chitin affinity purified, SDS-gel-electrophoretically separated proteins of Grossglockneria acuta (G) and of Mucor mucedo culture medium (M), probed with an antibody against potato chitinase. The marked proteins of G. acuta have similar molecular weights (arrowheads) as anti-chitinase-binding proteins of the control. mw = molecular weight (x 10-3 ).

Problems in reviewing chitin distribution arise by the different use of the term" chitin" in older and new reports. In the past chitin was defined mainly by histological staining properties. These are not reliable with the result that there are a number of conflicting observations related to chitin in particular groups. In such cases, restudy by modern techniques will be required. The definition of chitin has now become more exact, chitin being a polymer of mainly GlcNAc-units, which may be covalently linked to proteins. Therefore, today chitin can be (and should be) clearly demonstrated by a number of sensitive tools and methods such as immunodetection, chitinase-digestion or x-ray diffraction. The pattern of distribution of chitin among the protist groups, and its detection in extracellular structures of organisms near the base of eukaryotic evolution suggests that the expression of chitin synthesis might not be a helpful feature for the confirmation of phylogenetic relationships. However, the way chitin is synthesized and assembled may be discriminatory. Despite the great abundance and biological importance of chitin, the knowledge of its synthesis and deposition is still poor, especially with respect to the protists. The studies on fungi and animals indicate at present two

Chitin in Protists . 13

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Figs. 21-24. Ultrathin sections through the oral area of Grossglockneria acuta. The arrows point to vesicles which are suspected to contain chitinolytic enzymes. - Fig. 21. Oral apparatus (OA) of a non-feeding cell. - Fig. 22. G. acuta attacking the wall (W) of a Mucor hypha (M) (The living cell was preembedded in soft agarose to keep it fixed to the hypha during the fixation procedure). Figs. 23- 24. Attacking cell (Fig. 23) and feeding cell (Fig. 24) detached from the host. - Fig. 23. Membranous stacks (arrowheads) appear to accumulate beneath the oral area. - Fig. 24. They might provide membrane material for the growing food vacuole (FV). Bar = 1 11m.

different path wa ys of chitin for ma tion : (1) a one-step process (found in fun gi) where chitin synthetases in th e plasma membrane directly assemble and extrude the polysaccharid e, and (2) a two-step pro cess (found in arthropods) that invol ves the production of chitoproteins

via the Golgi apparat us. As chitin synthesis appears to be an anci ent feature of euka ryotic org anisms, but differentl y regulated among the species, it is not surprising to find features of both pathwa ys expressed amon g the phylogenetically old and diverse proti sts. It is as yet unclear wh eth er

14 . M. Mulisch

even more routes of chitin form ation exist among the unicellular organi sms. In this respect it would be int eresting to search for chitoproteins in cysts of various species th at are reported to consist of glycoproteins with GlcNA cgroups. Chitoproteins might be detected in cells that use membrane-bound pre cur sors for the form ation of their chitin structures. We have also to find out if the different path ways of chitin synthesis are related to phylogeny, or if the mod es of chitin formation are easily changeable and thu s may have developed independently in different groups (e.g., in arthropods and in ciliates). From the current state of knowledge, the latter app ears to be more probable.

8 Bart nicki-C arcia S. (1989): Th e bioch emical cytology of chitin and chitos an synt hesis in fungi. In: Skjak-Brsek G.,

9

10

11

Acknowledgements Man y th an ks to my hu sband, Dr. U. Ma rkma nn-M ulisch. I wo uld like to thank Pro f. Dr . W. Herth who stimulated my interest in the problem of chitin formation in protists. I tha nk also Dr . D. J. Patt erson , Prof. Dr. K. Spindler and Pro f. Dr. S. Berking for valua ble discussion s and many helpful suggestions, M. Hartmann , who did th e frustrating sectioning of rhe oral appa ratus of Grossglockneria acuta, J. Jacobi for dra wing Fig. 15, and G. Schermuly for help with the lirerature resear ch. The anti bo dy against pot ato chitinase was a gift of Dr. E. Kombrinck, MPI fur Zu chtu ngsfor schun g, Koln, Dr. M. N . Ho rst, Me rcer University, Macon , USA, provided the antibody aga inst polymeric chitin chito pro tein. My wo rk was suppo rted by the Deut sche For schun gsgemeinschaft (M u 728/2- 1 and Mu

728/4 - 1).

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87 -100. 19 Bussers J.-c., Hoesdor ff M., Bolome M., Greco N. et Goffinet G. (1986): L'enk ystement du cilie hypotriche Euplotes muscicola. Proti stologica, 22, 457 -460. 20 Bussers J.-c. et Jeuniaux C. (1974): Recherche de la chitine dan s les productions rnetapl asmatiq ue de quelques cilies, Protistol ogica, 10, 43 -46. 21 Bussers J.-c., Voss-Fouca rt M. F. and Bou chez-Decloux N. (1977): Ultrastructu re and chemical compos itio n of the lor ica of Folliculinopsis producta (Ciliata Heterotrichida). Abstr. Int. Congr. Prot ozool., 50, 358 (abstract). 22 Ca bib E. (1987): Th e synthesis and degrad ation of chitin. Adv. Enzymol., 59, 59 -10 1. 23 Ca bib E. (1991): Different ial inhibi tion of chitin synrherases 1 and 2 from Saccharomyces cerevisiae by polyoxin D and nikk omycins. Ant imicrob. Agent s Chemo ther., 35,

170- 173. 24 Cabib E., Roberts R. and Bowers B. (1982): Synthesis of th e yeast cell wall and its regulation . Ann. Rev. Biochem., 51, 763 - 793. 25 Cavalier-Smith T. (1991): Cell diversification in heterotrophi c flagellates. In: Patterson D. J. and Larsen

J. (eds.): Th e

Chitin in Protists . 15

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32

33

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41

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Key words: Protists - Chitin - Chitin synthesis - Chitin deposition - Chitin degradation - Chitinase Maria Mulisch, Universitat Koln, Zoologisches Institut, 1. Lehrstuhl, Weyertal 119, 5000 Koln 41, Germany