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Applied and Environmental Microbiology, October 2002, p. 5113-5122, Vol. 68, No. 10
0099-2240/02/$04.00+0 DOI: 10.1128/AEM.68.10.5113-5122.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.
Bart Deplancke,3 H. Rex Gaskins,3,4 Michael A. Apicella,5 and Margaret J. McFall-Ngai1*
Pacific Biomedical Research Center, Kewalo Marine Laboratory, University of Hawaii, Honolulu, Hawaii 96813,1 Department of Zoology, University of Hawaii at Manoa, Honolulu, Hawaii 96822,2 Division of Nutritional Sciences,3 Departments of Animal Sciences and Veterinary Pathobiology, University of Illinois, Urbana-Champaign, Illinois 61801,4 Department of Microbiology, University of Iowa, Iowa City, Iowa 522425
Received 20 March 2002/ Accepted 2 July 2002
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ABSTRACT |
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INTRODUCTION |
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In the present study we sought to (i) determine when and how mucus secretion is initiated in juvenile E. scolopes, (ii) define what host cells are responsible for secretion of the mucus that forms the matrix in which symbionts aggregate prior to colonization, and (iii) define the impact of the colonization of the light organ by V. fischeri on the secretory activity of mucus-producing host tissues.
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MATERIALS AND METHODS |
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Unless otherwise indicated, bacterial strains were grown at 25°C in a seawater-based tryptone medium (3) to the mid-logarithmic phase, and cell density was estimated spectrophotometrically by determining optical density at 600 nm. Squid were placed individually in 5 ml of either filter-sterilized seawater (FSSW) or unfiltered natural seawater (USW), which contained around 106 bacteria/ml (14) but did not harbor V. fischeri in sufficient quantities to colonize the host. Juvenile E. scolopes placed in either FSSW or USW did not form a symbiotic association. Test bacterial strains were added to FSSW or USW to obtain a final density of approximately 103, 104, 105, or 106 test bacteria/ml of seawater. One of the following six bacterial strains was used for each inoculation experiment: the wild-type strain V. fischeri ES114; two nonmotile strains, NM200 and NF201, which are derivatives of ES114 (19); Vibrio parahaemolyticus KNH1 (35); Bacillus cereus 43-25 (grown in Luria-Bertani medium) (15), a marine Bacillus sp. strain; and heat-killed, tetramethyl rhodamine isocyanate-labeled Staphylococcus aureus (Molecular Probes, Eugene, Oreg.). The vibrio strains and B. cereus carried a plasmid containing a gene that encodes a green fluorescent protein (GFP) (35). Hatchling squid were also exposed to suspensions of various types of sterile particles, including 1-, 2-, or 10-µm-diameter fluorescent polystyrene beads (Molecular Probes) at a concentration of 106 beads/ml, silicon dioxide particles at a concentration of 1 mg/ml, or carmine particles at a concentration of 0.1 mg/ml, in FSSW.
Microscopy.
For light microscopic analyses of mucin histochemistry, juvenile host animals or light organs from adult squid were fixed in 20 ml of Carnoy’s fixative (50% ethanol, 40% glacial acetic acid, 10% chloroform) at 4°C for 2 h. Samples were then dehydrated by washing them two times (10 min each time) in 100% ethanol at 4°C and then two times (10 min each time) in xylene at room temperature. Samples were then embedded in paraffin, sectioned with a Microm HM340E microtome, mounted on glass slides, and stored at 4°C before histochemical analysis.
To stain for various mucin types, sections were deparaffinized by using two changes of xylene, followed by washes with 100 and 70% ethanol before rehydration in deionized water. Sections were stained with either (i) high-iron diamine (HID) to visualize sulfomucins and alcian blue (AB) (pH 2.5) to visualize sialomucins (43) or (ii) periodic acid-Schiff stain (PAS) to visualize neutral mucins and AB (pH 2.5) to visualize sialomucins (6). Briefly, for HID-AB staining, sections were incubated in a solution containing 0.24% dimethyl meta-phenylenediamine dihydrochloride, 0.04% dimethyl para-phenylenediamine dihydrochloride, and 1.7% FeCl3 for 16 h at 25°C with agitation during the first hour. The sections were then rinsed in running tap water (RTW) and stained with a 1% AB solution for 10 min before they were rinsed in RTW. Stained samples were dehydrated, cleared in xylene, and then mounted with Permount. For PAS-AB staining, sections were rinsed in 3% acetic acid for 3 min and then incubated for 30 min with a 1% AB solution. Samples were rinsed in RTW, placed in 0.5% periodic acid for 10 min, rinsed in RTW, placed in Schiff's reagent for 10 min, reduced in 0.5% sodium metabisulfite, rinsed in RTW for 10 min, dehydrated, cleared in xylene, mounted with Permount, and viewed with a compound microscope.
To prepare specimens for confocal microscopy following incubation with bacterial strains or particles, hatchling squid were either stained with a fluorochrome for laser-scanning confocal microscopy (LSM) or left unstained for differential interference contrast microscopy. For fluorescence staining, animals were placed for 30 min in FSSW containing 1 µM CellTracker Orange (Molecular Probes), a fluorochrome that labels the cytoplasm of viable eukaryotic cells. The animals were rinsed with several changes of FSSW and then anesthetized in either a 1:1 solution of 80 mM MgCl2 and FSSW or a 2% solution of ethanol in FSSW. The animals were placed ventral side up in a depression well slide, and the mantle and funnel of each animal were dissected away to expose the underlying light organ. Fluorescently labeled animals and bacteria were simultaneously viewed by LSM with a Zeiss LSM 510 confocal microscope. Bacterial aggregations in unstained animals were viewed by using LSM and differential interference contrast microscopy.
To visualize mucus secretions by LSM, animals were incubated with fluorescently labeled wheat germ agglutinin (WGA) (Oregon Green or tetramethyl rhodamine; Molecular Probes), a lectin that specifically binds to N-acetylneuraminic acid (sialic acid) residues, which occur in the mucus secretions of the E. scolopes light organ (35). The animals were incubated with WGA at a final concentration of 10 µg/ml of FSSW for 30 min, rinsed two times (1 min each time) in FSSW, dissected, and viewed by LSM. Fluorescein isothiocyanate-labeled Ulex europaeus agglutinin (Sigma-Aldrich, St. Louis, Mo.), a lectin that labels L-fucose residues, which do not occur in the mucus secretions of the juvenile light organ (35), was used as a control to assess the degree of nonspecific binding of lectins to host mucus. To determine whether sialated residues that might be associated with the surfaces of V. fischeri cells could be the source of sialic acid residues in the crypts, culture-grown V. fischeri cells and symbionts isolated from adult light organs (34) were stained with WGA.
Treatment with PGN and LPS.
Peptidoglycan (PGN) from either V. fischeri (17) or S. aureus (Fluka, Milwaukee, Wis.) was treated with a 1% (wt/wt) solution of lysozyme in 0.1 M sodium acetate buffer (pH 5) for 10 min to digest the PGN into smaller fragments. The lysozyme was then inactivated by boiling the solution for 5 min. A control containing lysozyme without PGN was prepared in a similar manner. The treated PGN at a final concentration between 100 and 300 µg/ml or a corresponding equal volume of the lysozyme control was then added to 1 ml of FSSW, and hatchling squid were added. The animals were incubated for 2 to 3 h in these solutions and then prepared for lectin staining for mucus and viewed by LSM.
In experiments in which animals were exposed to PGN-conjugated beads, a 2% aqueous suspension of 2-µm-diameter, fluorescent, carboxylate-modified polystyrene beads (Molecular Probes) was added (50:50) to a 1-mg/ml solution of lysozyme-treated PGN from S. aureus in 50 mM 2-(N-morpholino)ethanesulfonic acid (MES) buffer (pH 6.0). A set of negative control beads was similarly treated, but no PGN was added. After a 15-min incubation at room temperature, 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (pH 6.5) was added to a concentration of 4 mg/ml to initiate covalent coupling of PGN to the beads. The 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide-treated suspensions were incubated overnight with shaking, and the coupling reaction was then quenched for 30 min by adding glycine to a final concentration of 100 mM. The beads were then washed 10 times in 50 mM sodium phosphate buffer with 0.1 M NaCl (pH 7.4) and then filtered through 10- and 3-µm-pore-size filters (Nuclepore, Pleasanton, Calif.) with repeated washing to remove coated beads larger than 3 µm. The beads were then washed again five times in FSSW before use. Both PGN-linked and control beads were stained with fluorescent WGA to confirm that PGN had been successfully bound to the beads. Both sets of beads were incubated for 3 h in FSSW and centrifuged, and the supernatants were collected and filtered through a 0.22-µm-pore-size filter (Millipore, Bedford, Mass.). The beads (106 beads/ml) with or without PGN or filtered supernatants were added to hatchling squid, and after a 3-h incubation the animals were stained for mucus detection as described above.
Hatchling squid were exposed to LPS from either V. fischeri (1), V. cholerae, or Escherichia coli (Sigma Chemical Co.) or to purified lipid A from V. fischeri (1) at concentrations of 10 and 100 ng/ml and 1 µg/ml in FSSW as previously described (18). These concentrations were chosen because they effectively promote mucus secretion in other animal systems (2, 25, 26, 49) and induce developmental changes in E. scolopes (18). The suspensions of both LPS and lipid A were sonicated for 30 s before they were added to the squid.
Experimental manipulation of colonization events.
To characterize the early events leading to mucus secretion and aggregation formation, the interaction of host tissues with bacteria or bacterium-sized particles was analyzed during the first 2 h after hatching. Freshly hatched squid were exposed to bacterial strains or 1-, 2-, or 10-µm-diameter fluorescent polystyrene beads (Molecular Probes), all at a concentration of 106 cells or beads per ml of FSSW. The animals were viewed by LSM between time zero and 1 h at 5-min intervals following exposure and then at 1.5 and 2 h, and the presence of any bacteria or beads attached to the ciliated surfaces of the light organ or within the ducts or crypt spaces was noted.
A series of experiments was conducted to determine whether colonization of the light organ affected mucus shedding and aggregation formation. To determine whether the ability to secrete mucus and form aggregations persists following early colonization of the crypts by V. fischeri, hatchling squid were continually exposed to wild-type V. fischeri cells for the first 24 h, and the ability of V. fischeri to form aggregations was observed by LSM at 2-h intervals. Then, to determine whether the host continued to aggregate newly available bacteria from the environment after prolonged colonization of the crypts (i.e., after 24 h), hatchling squid were first infected with unlabeled wild-type V. fischeri cells, and successful colonization was detected by monitoring luminescence with a TD-20/20 luminometer (Turner, Sunnyvale, Calif.). Animals that were symbiotic for 24, 48, 72, and 96 h were then exposed to GFP-labeled V. fischeri cells and viewed by LSM after 3 to 4 h of incubation to determine whether they remained capable of aggregating symbionts outside the organ after colonization of the crypts for specific times. The ability of animals maintained as aposymbiotic organisms (i.e., animals exposed only to USW) to form aggregations with added V. fischeri cells was also assessed at each of these time points.
In experiments in which light organs were cured of their symbionts following colonization, animals were continuously exposed to chloramphenicol at a final concentration of 10 µg/ml, and the decline in the symbiont population was monitored by monitoring luminescence as previously described (13). To determine whether cured E. scolopes juveniles are capable of forming bacterial aggregations, the chloramphenicol-treated animals were then inoculated with GFP-labeled V. fischeri and viewed by LSM after 3 to 4 h of incubation to determine whether aggregations had formed. In all experiments, aposymbiotic, symbiotic, and cured animals were stained with WGA to detect host mucus.
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RESULTS |
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1- or 5-µm-diameter, living or dead, gram-negative or -positive bacteria or to 1-, 2-, or 10-µm-diameter particles, neither bacteria nor particles were observed in the crypt spaces during the first 20 to 25 min after hatching. Between approximately 30 and 60 min following exposure, up to 90% of the animals exposed to 1-µm-diameter gram-negative and -positive bacteria or to 1-µm-diameter particles had between one and three bacterial cells or particles in the crypt spaces, as determined by LSM (Table 1). The 5-µm-diameter bacteria and 2- and 10-µm-diameter beads were never observed in the crypt spaces. These data indicate that passive early entry into host crypt spaces is not limited by particle type or viability but by size.
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Unlike the V. fischeri cells that later colonize the host, the motile strains of both wild-type V. fischeri and V. parahaemolyticus were no longer visibly motile after they entered the crypts during this early permissive period. When the light organs were plated, no CFU resulted, although culture-grown strains of bacteria exposed to the animals have a plating efficiency of 100% (38), suggesting that either the transport process itself or the conditions of the hatchling crypt space compromised the viability of the cells. Furthermore, the ability of the bacteria or particles to enter the crypts was restricted to a limited time frame. Although bacteria and 1-µm-diameter beads entered the crypts during the first hour after exposure, neither bacteria nor beads were observed in the crypt spaces of squid 2 h after inoculation (n = 20). These data indicate that the bacteria or particles that entered during the initial phase were cleared or degraded by the host in some manner.
Taken together, these data demonstrate that while colonization eventually occurs as the result of migration of motile strains of aggregated V. fischeri into the crypts between 2 and 5 h after inoculation (35), the light organ crypts are temporarily permissive to entry by other bacteria or particles that are 1 µm in diameter. However, these early recruits are cleared, and V. fischeri cells that may initially enter do not appear to contribute to the population of cells that eventually colonizes the host.
Depletion of mucus stores from the ciliated epithelial appendages of the hatchling host light organ.
Mucus secretion for aggregation formation, which normally commences 1 to 2 h following hatching, occurred principally from the cells of the ciliated epithelial appendages on the surface of the juvenile light organ. Although the entire ciliated epithelial fields of the juvenile light organ were coated with sialomucin (Fig. 2A and B), only the cells of the appendages of newly hatched squid had the histological appearance suggestive of mucus secretion, i.e., vivid staining of mucins localized to the apical regions of the cells (Fig. 2C). Distinct blue staining, indicative of sialomucins, and pink staining, indicative of neutral mucins, were observed in these intracellular stores (Fig. 2C). Sialomucin appeared to be the dominant mucin type found embedded in the densely ciliated cell surfaces, and neutral mucin was detected at relatively low levels in these extracellular environments (Fig. 2B and C). In animals maintained in FSSW, a coat of sialomucin covered the appendages, but little shedding of the mucus was observed (Fig. 2A and B). HID-AB staining showed that few sulfated mucins occurred anywhere except in the reflector tissue of the light organ, and this staining also confirmed that sialomucin covered the ciliated fields (data not shown).
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Because exposure to both gram-positive and gram-negative bacteria and their products led to mucus secretion, the ability of PGN, a common cellular component of both bacterial types, to induce host mucus secretion was tested. In addition, the abilities of the gram-negative bacterial cell wall molecule LPS and lipid A, the endotoxic component of LPS, to induce host mucus secretion were also tested; these molecules are known inducers of animal cell mucus secretion (12, 25). Animals exposed to V. fischeri or S. aureus PGN that had been lysozyme treated and heat inactivated exhibited mucus secretion from the ciliated appendages (Fig. 2D and Table 2). Heat-inactivated lysozyme by itself did not cause detectable mucus secretion. Neither LPS or lipid A from V. fischeri nor LPS from V. cholerae or E. coli caused detectable mucus secretion from host ciliated cells (Table 2). These data indicate that bacterial PGN, but not LPS or lipid A, is capable of inducing the release of host mucus from intracellular stores, although they do not prove that PGN is the only bacterial product that causes mucus secretion from these host cells.
Link between early permissiveness and mucus shedding.
A series of experiments was performed to determine whether early permissiveness of the light organ is required for mucus shedding and whether the eventual loss of permissiveness is due to the mucus shedding. Because the size of PGN-conjugated, 2-µm-diameter beads precluded their entrance into the crypt spaces (Table 1), these beads were used to determine their ability to cause host mucus shedding from outside the crypts. Exposure of the animals to these beads caused mucus shedding from the light organ surface (Table 2), while supernatants derived from incubation of the PGN-conjugated beads in FSSW for the same period (i.e., 4 h) did not cause mucus shedding in five of six animals tested. Since it is difficult to render the animals free of all contaminating material in FSSW, we feel that a response in a low number of animals (one of six animals in this instance) is not unexpected. These findings confirmed that the PGN on the beads was stably linked, providing few small molecules that might be shed from the beads that would interact with the crypt cells. Beads exposed to linkage agents in the absence of PGN caused mucus shedding in only one of six animals tested, confirming that the linkage treatment of the beads is relatively ineffective at causing this effect in the host. These data suggest that (i) host receptors for the induction of mucus secretion occur on the outside of the light organ (i.e., outside the crypts) and (ii) the early permissiveness of the organ to bacterial cells is not essential for the induction of mucus secretion.
Experiments were performed to determine whether the loss of the early nonspecific permissiveness of the light organ, which normally occurs 1 to 2 h following hatching, is a hard-wired, time-dependent, developmental change or the result of induced mucus secretion. To determine whether the block involves a developmental change, freshly hatched animals were maintained in FSSW for 3 h and then were exposed to GFP-labeled V. parahaemolyticus cells. Within 30 to 60 min after inoculation (3.5 to 4.0 h after hatching), these nonspecific bacterial cells were still able to enter the crypt spaces (Table 1), and they were cleared from the host crypts by 2 h after exposure (5 h after hatching). These data suggest that the block of permissiveness is not merely a time-dependent developmental process.
To determine whether the induced mucus secretion, which normally occurs concomitant with the block of permissiveness at 1 to 2 h following hatching, is responsible for the block, fresh hatchling squid were exposed to PGN from S. aureus for 3 h and then inoculated with GFP-labeled V. parahaemolyticus. Copious mucus secretion was observed, and 30% of the animals viewed (n = 10) contained between one and three nonspecific cells, levels which are lower than the levels of permissiveness observed before mucus secretion (Table 1). These data suggest that mucus secretion is, at most, only partly responsible for the block of early light organ permissiveness and that other factors or activities are required to achieve a complete block.
Taken together, these studies of the relatedness of early permissiveness and induction of mucus secretion suggest that although these events occur in close sequence, they are relatively independent. However, mucus secretion likely contributes to the cessation of nonspecific entry.
Developmental changes in mucus secretion and symbiont aggregation.
A series of experiments were conducted to determine whether host mucus secretion and bacterial aggregation patterns change after colonization of the light organ. The appendages of 72-h aposymbiotic juveniles maintained in USW and stained with fluorescently labeled WGA contained a markedly thicker covering of sialated mucus than the appendages of symbiotic juveniles after 48 h of colonization (Fig. 3A and B). Aposymbiotic juveniles that were maintained in V. fischeri-free seawater were able to secrete mucus and aggregate bacteria when they were exposed for 3 to 4 h to V. fischeri at any time over the first 96 h (Fig. 3C). In symbiotic animals, the ability to secrete mucus and aggregate V. fischeri persisted up to 24 h after colonization (Fig. 3D). However, by 48 h after successful colonization with V. fischeri, little host mucus shedding was detected (Fig. 3B), and GFP-labeled V. fischeri that was in the environment of the host was no longer detected in aggregations outside the light organ (Fig. 3E). When a colonized host light organ was cured with the antibiotic chloramphenicol, mucus shedding from the ciliated fields was again detected (Fig. 3F). Concomitantly, the host's ability to aggregate V. fischeri was restored (Fig. 3G), although the number of cells amassed was lower (10%) than the number of cells detected in animals that had been maintained as aposymbiotic organisms over same time frame and then exposed to V. fischeri (Fig. 3C and G). The lower numbers of symbiont cells in the aggregations were most likely due to the fact that because morphogenesis had been induced by colonization (33), fewer cells were present in the ciliated epithelium that secreted mucus. These results indicate that the presence of the symbiont in the crypts inhibits the ability of the host to aggregate bacteria further after the association is established, although this host response can be reinitiated by eliminating the symbiont. Thus, symbiont suppression of host mucus secretion from the ciliated appendages and bacterial aggregation requires persistent interaction with intact V. fischeri cells in the crypt spaces, which are several tissue layers away from the ciliated fields of the juvenile light organ.
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DISCUSSION |
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In the host's natural environment, the light organ contains only strains of V. fischeri (30). However, the finding that hatchling crypts are transiently open to 1-µm-diameter bacteria and particles indicates that the factors regulating exclusive entry of V. fischeri do not fully function during early stages of the symbiosis. Because the early entrants, including V. fischeri, were cleared from the crypt spaces, the crypts might be considered hyperspecific during this period (i.e., they allow entry of symbiotic and nonsymbiotic bacteria and particles but inhibit subsequent persistence of these bacteria and particles). The process of removal remains to be described but may involve a harsh chemical environment in the crypt spaces (42, 47; Visick and Ruby, Abstr. 97th Gen. Meet. Am. Soc. Microbiol.) or the activity of the crypt cells. A previous study of the E. scolopes light organ showed that host blood cells or hemocytes, which have activities similar to those of vertebrate macrophage cells, are found in the crypt space environment (34). To what extent these macrophages remove early bacterial invaders from the light organ crypts remains to be determined. Whatever the mechanism(s), the results of this study and previous studies (30) suggest that once the early entrants into the organ are cleared, V. fischeri is the sole occupant of the crypt space throughout the life of the host animal.
Production of the mucus in which symbionts form a biofilm.
The present results suggest that during embryogenesis the host develops a nascent light organ that is biochemically and biophysically ready to interact with the environment to harvest symbionts. Upon exposure of a hatchling animal to the natural seawater environment, the ciliary activity of the superficial light organ epithelium commences and mucus is shed by the cells. In the context of the squid-vibrio symbiosis, these findings reveal another role for the ciliated epithelial fields in the promotion of colonization of host tissues by the symbionts. Thus, these tissues are responsible not only for currents and entrainment of V. fischeri-containing seawater and mucus during aggregate formation (35) but also for production and delivery of the mucus.
Under the experimental conditions of the present study, the induction of mucus secretion was relatively nonspecific; i.e., mucus secretion was induced by both gram-negative and gram-positive bacteria, as well as by PGN, although it was not induced by nonbiological particles that were the same size as the bacteria (Table 2). Previous studies of the colonization process have indicated that only gram-negative bacteria aggregate in the mucus (35) and that only V. fischeri sets up permanent residence in the crypt spaces (24). Taken together, these data indicate that the colonization of the E. scolopes light organ is a process of ever-increasing specificity.
PGN was the molecule found in both gram-negative and gram-positive bacteria that caused host mucus secretion in these experiments. Although the PGN of gram-positive bacteria is exposed, PGN is buried beneath the outer membrane of gram-negative bacteria. Thus, for the PGN of gram-negative bacteria to be active in the induction of mucus secretion, cells must either lyse or bleb their surfaces to expose PGN. Studies of V. fischeri have shown that it actively blebs its cell surface (Apicella, unpublished data), and similar activity has been noted in other environmental and pathogenic gram-negative bacteria (9, 21, 28, 36). Studies of the activity of PGN in other systems have demonstrated that it is a potent activator of the eukaryotic immune system and leads to the production of numerous inflammatory mediators (31), although we are not aware of any other studies implicating PGN in the induction of mucus secretion.
The induction of mucus secretion by PGN-conjugated beads suggests that the receptors for PGN are outside the light organ. The nature of these receptors remains to be determined, but several recent studies of animal receptors of bacterial products provide some intriguing candidate molecules. PGN is a known ligand for certain Toll receptors, where its binding induces the activity of the evolutionarily conserved NF-
B pathway (39, 48). In addition, recently, a new class of PGN receptors has been described, whose pathway of activation has not yet been elucidated (22, 26). Similar PGN recognition proteins in E. scolopes may be involved in the binding of PGN that could promote or induce, either directly or indirectly, the shedding of mucus.
The data obtained from histological and lectin staining demonstrate that sialated mucus is shed from the ciliated epithelial appendages in response to bacteria. The apparent decrease in these intracellular mucus stores, along with the continued mucus secretion in animals that were maintained in USW but were not exposed to V. fischeri, indicates that there is a high rate of turnover of mucus within the ciliated appendages. The ability to dispense and alter internal mucus stores quickly is not unprecedented in E. scolopes. A previous study of mucus secretory cells found in the mantle epidermis of E. scolopes demonstrated that this squid has the ability to secrete and chemically modify its mucus secretions from this tissue almost instantaneously (40).
Because symbiotic animals retain the ability to shed mucus and aggregate bacteria over the first 24 to 48 h, the host may be continually sampling the environment using mucus secretions and the ciliated fields until it is successfully colonized by V. fischeri. By 48 h after colonization, host mucus shedding decreases, and further aggregation formation is inhibited. These data suggest that V. fischeri is able to inhibit the ability of the host to aggregate bacteria by suppressing mucus secretion after colonization, because curing of the light organ resulted in renewed mucus shedding and subsequent bacterial aggregation, although at lower levels than the levels in aposymbiotic animals of the same age (Fig. 3). A previous study showed that 12 h of exposure to V. fischeri cells leads to irreversible apoptosis and regression of the ciliated epithelial fields (13). Therefore, the inability of cured animals to aggregate bacteria and secrete mucus at the same levels as aposymbiotic animals may be due to fewer secretory cells being present.
Whereas colonization of the crypt spaces by V. fischeri inhibits mucus secretion by the cells of the superficial epithelium, an increase in the sialated mucin level was observed in the crypts in response to colonization. Previous studies of symbiont-induced changes in the host have shown that direct interactions with V. fischeri cause the crypt epithelial cells to swell (33) and induce increases in the microvillar densities of these cells (23). Thus, the present study adds to our knowledge concerning the effects that the symbionts have on the host cells with which they directly interact. The function of the mucus in the crypts (e.g., as a nutrient source for the bacteria or as part of a type of host immune response) and whether mucus secretion is coupled to these developmental events remain to be determined.
The mechanism by which V. fischeri changes the secretory activity in the two epithelia of the symbiotic organ, the remote superficial ciliated epithelium and the directly associated crypt epithelium, is also unresolved. One possibility is that the symbionts remotely change the density or activity of host receptors for bacterial signaling molecules. Once the PGN receptors are identified in E. scolopes, whether mucus secretion changes due to alterations in the density of these receptors can be investigated. Another possibility is that the symbionts influence the expression of mucin-encoding genes directly in either or both fields of cells. Bacterium-induced changes in mucus production have been reported previously for both cooperative and pathogenic associations, but most often the bacteria have a direct effect on the host cells (5, 10). A previous study of LPS from the human stomach commensal bacterium Helicobacter pylori showed that while exposure to the LPS of this bacterium initially induced mucus secretion, after prolonged exposure it actually led to concentration-dependent inhibition of secretion (41). Intact, viable H. pylori cells cause similar effects and have also been shown to suppress expression of the mucin-encoding genes in cultured human gastric cells (4, 32). Although V. fischeri LPS does not appear to induce mucus secretion in E. scolopes, it may have an indirect suppressive effect on host cells of the superficial epithelium once the symbiont colonizes the light organ. Other pathogenic bacteria, such as P. aeruginosa, have been reported to cause increases in mucus production by epithelia in the lungs of patients suffering from cystic fibrosis (11), similar to the observed mucus induction of the squid crypt epithelia after colonization by V. fischeri.
The present study demonstrated that a bacterium in a beneficial association is able to alter host mucus production in a variety of ways. An initial nonspecific response of superficial tissues to environmental bacteria and/or their products sets the stage for harvesting of the specific partner. Once colonization by this partner has occurred, the activity of the partner changes mucus production in both remote tissues and the tissues with which they directly interact. Thus, the squid-vibrio symbiosis may be a useful model for studying the role of microbially derived factors in altering mucin synthesis and secretion during the interactions of bacteria with their animal hosts.
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ACKNOWLEDGMENTS |
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This research was funded by a grant from the W. M. Keck Foundation (to M.M.-N. and M. A. A.), by NIH grant RR12294 (to E. G. Ruby and M.M.-N.), by NSF grant IBN 9904601 (to M.M.-N. and E. G. Ruby), and by USDA NRI grant 98-35206-6429 (to H.R.G.).
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FOOTNOTES |
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Present address: Stanford University, Hopkins Marine Station, Pacific Grove, CA 93950.
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