Bacteria and digestive enzymes in the alimentary tract of Radix peregra (Gastropoda, Lymnaeidae)

The extracellular digestive enzymes cellobiase and chitobiase were quantitatively determined in the esophagus, stomach, and intestine of Radix peregru. Although both enzymes had comparable Michaelis constants between 16 and 32 PM liter-l, the enzyme activities were significantly higher for cellobiase (2 nanoKatals [nKat] from the whole gut tract) than for chitobiase (0.3 nKat). As became evident after a treatment with antibiotics, chitobiase was provided by bacteria and cellobiase was synthesized in the digestive glands of the animals. A lysozyme assay indicated that R. peregra was able to digest bacterial food. Cellulose and chitin are two of the most abundant structural polysaccharides in nature. The ability of animals to digest this ubiquitous food resource is rather limited and often dependent on symbiotic microorganisms (see Vonk and Western 1984). Freshwater pulmonates are reported to have both cellulolytic (Skoog 1978) and chitinolytic enzymes (Jeuniaux 1955). However, quantitative data on enzyme activities are scarce, and although bacteria were found in the gut tract of pulmonates (Reavell 1980), their possible role in hydrolyzing these refractory substances is not yet clear (Charrier and Lesel 1991). This study quantitatively describes the determination of celluloseand chitin-degrading enzymes and the contribution of bacteria in the activity of these enzymes in a freshwater pulmonate snail. Traditionally, analyses of digestive enzymes were made in whole-body homogenates of the animals under investigation. However, this makes it impossible to differentiate between true digestive enzymes and the enzymes of other catabolic processes (Hoeger and Mommsen 1984). In pulmonates, intracellular digestion is of minor importance and extracellular digestion is predominant in the gut tract (Salwini-Plawen 1988). Therefore, I chose the pulmonate snail Radix peregra for these studies, which digests cellulose and chitin (Owen 1966; Skoog 1978) and also harbors more bacteria in its digestive tract than do other species (Reavell 1980). Animals were grown in the laboratory on an unlimited diet of lettuce, Tetramin fish flakes, and Chlamydomonas reinhardtii (Chlorophyceae) cells. These foods were chosen because lettuce provides cellulose, and Tetramin, which contains ground arthropod exoskeletons (Tetra Werke pers. comm.), contributes chitin to the snail’s diet. Two enzymes, cellobiase (EC 3.2.1.21) and chitobiase (EC 3.2.1.30), were assayed in the isolated extracellular contents of the esophagus (Es), stomach (St), and intestine (In). This subdivision of the snail digestive tract follows Carriker’s (1946) description of the closely related species Lymnaea stagnalis appressa. The esophagus included the buccal cavity, proesophagus, and postesophagus. The stomach contained the crop, the gizzard, and the pylorus (with hepatic vestibule, cecum, and atrium). The intestinal section included the prointestine, intestine, and rectum. The enzyme analysis was made with chromogenic substrate analogs (methylumbelliferone substrates). By starting the enzyme analysis with different substrate concentrations (from 3 to 250 PM) and following the increase in fluorescence per unit time at the beginning or the enzymatic reactions, Michaelis-Menten curves were obtained. A calibration curve, set up with different concentrations of pure methylumbelliferone, served to transform these fluorescences and calculate enzyme kinetics (K, and V,,, values). Boiled (10 min) enzyme extracts were used as controls so that any background fluorescence present did not interfere with the enzyme analysis. A detailed description of the dissection of the animals, preparation of enzyme extracts, enzyme analysis, and calculation of enzyme kinetics are given elsewhere (Brendelberger 1997). In addition to the laboratory-reared animals, cellobiase and chitobiase were also analyzed in field-collected snails for comparison. These wild animals were collected in the Rhine River near Cologne and were measured (but not fed) within 24 h after collection. To determine the possible role of microorganisms in providing hydrolytic enzymes in the gut of R. peregra, a group of animals was treated with antibiotics. To the aerated tapwater, in which the animals were reared, the following antibiotics and antimycotic were mixed together and then added at the indicated final concentrations: penicillin (100,000 units liter-l), dihydrostreptomycin (10 mg liter-l), and amphotericin B (2.5 mg liter-l). In the case of penicillin (active against Gram-positive bacteria) and amphotericin B (active against fungi), the concentrations indicated are those generally recommended by laboratory manuals for sterile cultures (e.g. Lindel and Bauer 1983). Therefore, I assumed that both Gram-positive bacteria and fungi were quantitatively eliminated by this treatment, which lasted 2 weeks, by changing the rearing water and the antibiotics/antimycotic every third day. Dihydrostreptomycin, which is active against both Gram-positive and Gram-negative bacteria, was applied at only 10% of the recommended concentration, because at nominal concentrations the snails became inactive and died within a few hours. Any microorganisms surviving were thought to be Gram-negative bacteria. The snails were considered active and unaffected by the antibiotic treatments when the amount of feces per animal per day did not decrease by > 10% compared to the untreated control. Animals treated with antibiotics were also subjected to enzyme analysis in esophagus, stomach, and intestine as described above. The effect of antibiotic treatment on bacterial abundance was checked by epifluorescence counts of microorganisms after DAPI staining (Porter and Feig 1980). At least 200