Research Zebra Mussel Destruction by a Lake Michigan Sponge: Populations, in Vivo 31P Nuclear Magnetic Resonance, and Phospholipid Profiling T I M O T H Y A . E A R L Y * ,† A N D T H O M A S G L O N E K †,‡ Aquatic Research Institute, Inc., 2001 East 135th Street, East Chicago, Indiana 46312, and MR Laboratory, Midwestern University, 5300 South Ellis Avenue, Chicago, Illinois 60615
The sponge Eunapius fragilis (Leidy), that has become abundant in southwestern Lake Michigan, overgrows zebra mussels, Dreissena polymorpha (Pallas). The overgrown mussels die. Population dynamics show that in the presence of zebra mussels, E. fragilis is the dominant species (61% of sponges), with 90% of these sponges being associated directly to the mussels. In vivo 31P NMR spectroscopy performed on overgrown mussels reveals that mussel ATP is depleted in a manner indicative of a tissue in anoxia (Figure 2). 31P NMR phospholipid profiles reveal that the phospholipids of such mussels have been degraded to their lyso-phospholipid forms. Phospholipid degradation may be the direct result of the lysosomal chemical action of the sponge or a secondary effect resulting from sponge-induced death by anoxia followed by microbial action on the weakened energy-depleted mussels.
Introduction The sphaeroform sponge, Eunapius fragilis (Leidy) (1), is increasing in numbers in the southwestern quadrant of Lake Michigan, where epizoic growth of these sponges on live zebra mussels, Dreissena polymorpha (Pallas) (2), has been reported (3). Encapsulated mussels are killed and apparently digested. Predation of mollusks by sponges is a rare or unknown phenomenon, depending upon the point of view, and until recently (4), the concept of the sponge as predator was generally not accepted. Luxuriant sponge overgrowth, however, has been noted to visibly interfere with the opening of valves of unionid species (5), often with lethal effects (6). To gain insight at the biochemical level into how E. fragilis may be destroying zebra mussels that it overgrows, techniques of in vivo 31P NMR spectroscopy (7-9) and phospholipid profiling (10, 11) were used to evaluate zebra mussel high-energy phosphates and membrane phospholipids. In vivo 31P NMR profiling of living tissues provides quantitative information from which a direct assessment of organism energetics may be derived (7, 8). Phospholipid profiling provides a quantitative distribution of the membrane phospholipids that are integral to the regulation of cellular function * Corresponding author telephone: (219)391-8518; fax: (708)3861980; e-mail:
[email protected]. † Aquatic Research Institute, Inc. ‡ Midwestern University. 10.1021/es980874+ CCC: $18.00 Published on Web 04/28/1999
1999 American Chemical Society
and that provide the barrier through which extracellular signals are transduced. Both the qualitative and quantitative features of the phospholipid profile can be influenced by environmental factors (12). Setting. First introduced into the Great Lakes at Lake St. Clair (1986) (2), zebra mussels spread rapidly and extended to near coastal southern Lake Michigan by 1991 (13). At that time, water visibility in Lake Michigan was only 2-3 m, and sponges were rare. A fingerform sponge, Spongilla lacustris (Linneaus) (1), today found in great abundance, numbered only 10-15 per m2 in their most preferred habitats. A sphaeroform sponge, Eunapius fragilis (1, 14), also observed in great abundance today, was never observed. In the spring of 1992, zebra mussels had colonized virtually all suitable substrates within the near-shore waters of southern Lake Michigan (1000-10 000/m2). Also in 1992, the first specimens of the sphaeroform sponge were observed within the Hammond Marina harbor, and these were observed as having overgrown zebra mussels. Lake water visibility had improved to 12-18 m, and zebra mussel population densities continued to increase. By 1993, the sponge populations had increased to over 100/m2 in their preferred habitats. It was observed that in areas where high sponge populations existed, zebra mussel populations were relatively low (15). Where sponges were not found, heavy zebra mussel druses had formed. Divers reporting from various locations within the southern basin of Lake Michigan noted that sponges were now found in great abundance where they had never been seen before. A report was issued noting the relationships among the sponges and the zebra mussels (15). By spring 1994, some sponge populations had increased to as high as 1000/m2 (16). Within the matrix of individual sponges, closed mussels were found along with empty but closed mussel valves, both held in the closed position by sponge overgrowth. Spongeovergrown, but still living, mussels had discarded their byssus. No sponge growth was observed within closed valves, however. It appeared that these freshwater sponges may be predating zebra mussels, either directly through chemical digestive action or indirectly by oxygen or nutrient deprivation followed by secondary bacterial decomposition, with the sponges feeding on the bacteria (1). This unusual activity for a sponge (4) warranted further investigation. In addition to sponge and mussel populations, 31P NMR in vivo spectroscopy (7-9) was used to test whether zebra mussels were being energy-deprived and subjected to a digestive process. In vivo 31P spectroscopy quantifies phosphates, such as ATP and sugar phosphates, in intact living mussels, providing an index (17) of energy metabolism. 31P phospholipid profiling of mussel extracts yields a quantitative analysis of from 15 to 24 generic phospholipid classes, which include the phospholipids, such as phosphatidylcholine (PC) and sphingomyelin (SM), along with their degradation products, such as lysophosphatidylcholine (LPC) and sphingosylphosphorylcholine (lyso SM) (11, 18). These data can be used to assess biochemical processes involving the cellular membrane (11).
Experimental Section Field Site. The field site was the ship wreck of the Material Service Barge, located 5.6 Km due North of the Hammond Marina, outside the Calumet Harbor breakwall in the open waters of Lake Michigan in 11 m of water (Loran C, 50202/ 33426.6) (19). This site is a typical Lake Michigan open-water VOL. 33, NO. 12, 1999 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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TABLE 1. Lake Michigan Sponge Population Dynamics Sponge Populations Per Grid (% of Animals Detected) sponge animal E. fragilis E. muelleri S. lacustris
description of substrate within the grid area
grids containing zebra mussels grids without zebra mussels
61
19
7
4
72
24
Proportion of Sponge Species Associated with Zebra Mussels % of mussel-associated 90 27 25 sponge population % of sponge community 89 9 2
habitat in which populations of sponges and zebra mussels are represented. Population Dynamics. Aufwuchs (20) population dynamics of the site’s three common sponges, Ephaditia muelleri, Spongilla lacustris, and Eunapias fragilis, were made employing scuba and a grid to count species at designated locations (n ) 116) within the field site. Harvesting and Preparation. Samples of sponge-overgrown zebra mussels were taken from vertical substrate and the undersides of overhanging surfaces at depths of 6 m. Zebra mussel samples completely free of sponges were taken from appropriate locations in the immediate vicinity. The specimens, in 20 L of Lake Michigan water, were transported directly to the MR Laboratory. Juvenile zebra mussels (1-6 mm) were carefully separated from druses. Juvenile zebra mussels overgrown by sponge were gently separated from the sponge matrix, and their valves cleaned of adhering
sponge material; most of these mussels remained closed throughout the experimentation. 31P Nuclear Magnetic Resonance (NMR). For in vivo analysis (7-9, 21, 22), a collection of 200, 1-4 mm live juvenile mussels were placed into a 10 mm NMR sample tube containing Lake Michigan water and analyzed immediately. Following the NMR analysis, the mussels were found to be alive. Sponge-free mussels introduced into Lake Michigan water following the NMR measurement opened and began filtering water; most sponge-overgrown mussels, with sponge removed, continued to hold their valves closed. The in vitro phospholipid analysis (10, 11, 18, 23, 24) was conducted on each of six phospholipid samples per group (sponge-free, sponge-overgrown). Juvenile mussels (5 g), washed free of debris, were homogenized in a glass blender containing 100 mL of chloroform-methanol (2/1, v/v). This extract, having only one liquid phase, was then filtered into a separatory funnel. In the modified (24) Folch et al. (25) procedure employed, the aqueous backwash solution (20 mL) contained 0.2 M potassium (ethylenedinitrilo)tetraacetic acid (KEDTA) adjusted to pH 6.0. Following shaking of the separatory flask to scrub the extract of cations other than potassium, the two liquid phases were allowed to separate for 24 h, and the recovered chloroform phase was evaporated at 37 °C using a rotary evaporator. The analytical medium for 31P NMR phospholipid analysis was a hydrated chloroform-methanol NMR reagent specifically designed for the quantification (10, 11, 18, 24) of phospholipids by 31P NMR. The reagent has two parts: Reagent A is composed of chloroform containing 5% benzened6 (for spectrometer field-frequency stabilization) and, for
FIGURE 1. E. fragilis overgrowing zebra mussels. Note that only the siphons of one large mussel remain free of sponge overgrowth. 1958
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FIGURE 2. In vivo 31P NMR phosphatic metabolite spectral profiles (7, 8) of healthy, sponge-free zebra mussels (top) and spongeovergrown zebra mussels (bottom). Live zebra mussels were placed into the NMR sample tube containing Lake Michigan water: PME, phosphomonoesters, including the sugar phosphates; Pi, inorganic orthophosphate (spectral pH value 6.96); PArg, phosphoarginine; ATP, adenosine triphosphate with γ-, r-, and β-phosphate groups; ADP, adenosine diphosphate with β- and r-groups. The low signalto-noise ratio in the bottom spectrum reflects the depleted and emaciated condition of sponge-overgrown zebra mussels.
TABLE 2. Zebra Mussel in Vivo 31P NMR Low- and High-Energy Phosphate Profiles 31P
phosphatea PME Pi
NMR chemical shift (ppm)
phosphate concn (mol % of total detected phosphorus) spongespongefree overgrown
Low-Energy Phosphates 8-2 19.2 1.50 33.9
High-Energy Phosphates -3.31 3.5 γ, -5.58 36.0 R, -10.50 β, -19.09 ADPb R, -10.50 4.7 β, -19.09 nucleoside-12- -14 1.8 diphosphosugars dinucleotide -12 0.9 phosphorusb PArg ATP
40.9 32.4
6.7 16.9
3.1
a Phosphate abbreviations: PME, phosphomonoesters; Pi, inorganic orthophosphate; PArg, phosphoarginine; ATP, adenosine triphosphate; ADP, adenosine diphosphate; nucleosidediphosphosugars, uridine diphosphoglucose, etc., cytidine diphosphocholine, etc.; dinucleotides, nicotinamide adenine dinucleotide, etc. b Calculated by difference (9, 22).
quantification, an appropriate amount of trimethyl phosphate; reagent B is composed of a 4/1 (v/v) solution of methanol and aqueous 0.2 M CsEDTA, pH 6.0. Up to 100 mg of lipid extract was dissolved in reagent A (2 mL) and transferred to a 10 mm NMR sample tube. Reagent B (1 mL) was added, the admixture was stirred thoroughly, and the sample was placed into the spectrometer for analysis.
FIGURE 3. 31P NMR spectral phospholipid profiles (10, 11, 24) of healthy, sponge-free zebra mussels (top) and sponge-overgrown zebra mussels (bottom): LPG, lysophosphatidylglycerol; LPA, lysophosphatidic acid; LDPG, lysodiphosphatidylglycerol; GPLAS, glycerol plasmalogen; PG, phosphatidylglycerol; LEPLAS, lyso-ethanolamine plasmalogen; LPE, lysophosphatidylethanolamine; LPS, lysophosphatidylserine; PA, phosphatidic acid; DPG, diphosphatidylglycerol; DHSM, dihydrosphingomyelin; EPLAS, ethanolamine plasmalogen; PE, phosphatidylethanolamine; PS, phosphatidylserine; SM, sphingomyelin; U, uncharacterized phospholipid; LAAPC, lysoalkylacylphosphatidylcholine; LPC, lysophosphatidylcholine; PI, phosphatidylinositol; AAPC, alkylacylphosphatidylcholine; PC, phosphatidylcholine. In addition, zebra mussels contain phosphonophospholipids. The phosphono-phospholipid resonance band is displaced 20 ppm downfield from their corresponding phosphatephospholipid analogues and are not shown in the figure. The zebra mussels of this study exhibited three phosphono-phospholipid signals: LDAG-AEP, lyso-diacylglyceryl-(2-aminoethyl)phosphonate; DAG-AEP, diacylglyceryl-(2-aminoethyl)phosphonate; and CER-CP, ceramide-(2-tetramethylammonium-ethyl)phosphonate. The NMR spectrometer used in this investigation was a heteronuclear GE 500 NB system, operating at 202.4 MHz for 31P. Chemical-shift data are reported relative to the usual standard of 85% phosphoric acid (7, 9, 21, 26); however, the primary internal reference was the R-group resonance from adenosine triphosphate (ATP), at -10.5 ppm, for the in vivo determinations (9, 21) and phosphatidylcholine (PC) at -0.84 ppm (10, 11) for phospholipid determinations. Spectrometer conditions used in the analysis of live zebra mussels are given along with those used for phospholipids (as parentheticals): pulse sequence, one pulse; pulse width, 18 µs, which corresponds to a 45° spin-flip angle; sweep width 7042 (5000) VOL. 33, NO. 12, 1999 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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TABLE 3. Zebra Mussel 31P NMR Phospholipid Profilesa phospholipid
31P NMR chemical shift (ppm)
LDAG-AEP DAG-AEP CER-CP LPG LPA LDPG GPLAS PG LEPLAS LPE LPS PA DPG DHSM EPLAS PE PS SM U LAAPC LPC PI AAPC PC
21.60 21.25 20.42 0.89 0.79 0.66 0.62 0.54 0.48 0.43 0.40 0.34 0.18 0.11 0.06 0.03 -0.01 -0.09 -0.19 -0.23 -0.29 -0.35 -0.78 -0.84
phospholipid concn (mol % ( SD) sponge-free sponge- overgrown 16.60 ( 0.15 2.43 ( 0.19 0.64 ( 0.12 0.54 ( 0.10 0.058 ( 0.115 0.268 ( 0.033 0.350 ( 0.119 4.04 ( 0.12 1.46 ( 0.29 25.61 ( 0.21 6.42 ( 0.31 4.66 ( 0.40 1.78 ( 0.11
4.07 ( 0.07 11.68 (0.42 19.41 ( 0.15
0.130 ( 0.102 5.67 ( 0.11 1.44 ( 0.11 0.048 ( 0.026 1.30 ( 0.22 0.188 ( 0.017 0.27 ( 0.13 1.03 ( 0.13 0.480 ( 0.036 1.060 ( 0.088 1.085 ( 0.049 0.225 ( 0.114 3.95 ( 0.24 1.71 ( 0.19 21.58 ( 0.43 11.11 ( 0.39 2.88 ( 0.39 8.86 ( 0.09 0.648 ( 0.068 2.760 ( 0.053 1.150 ( 0.059 3.29 ( 0.12 17.80 ( 0.40 11.35 ( 0.26
2-tail probability 0.000 0.000 0.002 0.015 0.000 0.000 0.181 0.513 0.195 0.000 0.000 0.001 0.000
0.000 0.000 0.000
a Phospholipid abbreviations: LDAG-AEP, lyso-diacylglyceryl-(2-aminoethyl)phosphonate; DAG-AEP, diacylglyceryl-(2-aminoethyl)phosphonate; CER-CP, ceramide-(2-tetramethylammoniumethyl)phosphonate; LPG, lysophosphatidylglycerol; LPA, lysophosphatidic acid; LDPG, lysodiphosphatidylglycerol; GPLAS, glycerol plasmalogen; PG, phosphatidylglycerol; LEPLAS, lyso-ethanolamine plasmalogen; LPE, lysophosphatidylethanolamine; LPS, lysophosphatidylserine; PA, phosphatidic acid; DPG, diphosphatidylglycerol; DHSM, dihydrosphingomyelin; EPLAS, ethanolamine plasmalogen; PE, phosphatidylethanolamine; PS, phosphatidylserine; SM, sphingomyelin; U, uncharacterized phospholipid; LAAPC, lysoalkylacylphosphatidylcholine; LPC, lysophosphatidylcholine; PI, phosphatidylinositol; AAPC, alkylacylphosphatidylcholine; PC, phosphatidylcholine (11, 18).
Hz; acquisition delay, 71 (100) µs; free-induction decay size, 4096 (16 384) channels; interpulse delay, 500 µs; acquisition time, 290 (1640) ms; number of acquisitions, 9300 (10 000). Additionally, a computer-generated exponential filter timeconstant introducing 50 (0.6) Hz line broadening was applied to reduce background noise. Data reductions, including chemical-shift measurements (9), spectral curve analysis (27), when required, signal-area integrations, and relative metabolite mole-percentages (28) were computed. Proton broadband decoupling also was applied routinely (0.25 W). Statistics. Tests of statistical differences were made using the two-tailed Student’s t-test for independent samples in a program that also computes the F value used to test homogeneity of variance and its probability (29).
Results Population Dynamics. Aufwuchs (20) population dynamics of the field site’s three common sponges, E. fragilis, E. muelleri, and S. lacustris, noting the number of instances where each sponge made contact with zebra mussels, is presented in Table 1. Comparing average sponge populations within the grid area, in the presence of zebra mussels, E. fragilis appeared as the dominant species, representing 61% of the total sponge community (by numbers of animals per grid). E. muelleri and S. lacustris represented 19% and 7%, respectively. Within grids containing no zebra mussels, E. fragilis represented only 4% of sponges present, with E. muelleri being the dominant sponge (72%) and S. lacustris representing 24%. Regardless of whether zebra mussels were present, the ratio of E. muelleri to S. lacustris was the same (mussels present, 73%/27%; absent, 75%/25%), with only the proportion of E. fragilis being dependent upon mussel population density. Comparing sponge species that were associated directly to the mussels (by attachment, or contact, or overgrowth), it was found that 90% of the E. fragilis population was 1960
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associated with mussels, whereas only 27% of E. muelleri and 25% of S. lacustris were described by similar association. When comparing only those sponges attached to, or in contact with, zebra mussels, E. fragilis represented 89% of the community, whereas E. muelleri and S. lacustris represented 9% and 2%, respectively. This figure for E. fragilis reflects the same proportion within the total sponge community as that within its own population (about 90%). Further, zebra mussel contact by E. fragilis may be characterized almost exclusively as overgrowth leading to complete mussel encapsulation. In contrast, contact by E. muelleri and S. lacustris may be characterized, as peripheral contact resulting from sponge growth within a confined space. In no case were zebra mussels found within the matrix of E. muelleri or S. lacustris. An example of E. fragilis overgrowing zebra mussels is shown in Figure 1. In Vivo 31P NMR Spectroscopy. In vivo 31P NMR spectra of sponge-free and sponge-overgrown zebra mussels are presented in Figure 2, along with relative signal area quantifications in Table 2. Sponge-free zebra mussels exhibit the three characteristic signals of ATP in high amount. Also detectable are the high-energy phosphates, phosphoarginine (7, 30) and the nucleoside diphosphosugars centered at -11.5 ppm. The low-energy phosphate monoester (PME) band contains the sugar phosphates of glycolysis and inorganic orthophosphate (Pi) and are present in low relative amounts. Such a spectrum (22, 31) is typical of healthy well-oxygenated tissues. In contrast, the spectrum from sponge-overgrown zebra mussels is depleted in ATP to the point where this essential phosphate is nearly undetectable. Adenosine diphosphate, a product of ATP degradation, represents essentially all of the remaining high-energy phosphate component. The lowenergy phosphate bands of PME and Pi account for the bulk of the detectable phosphate. Such a spectrum is characteristic of highly stressed and/or dying tissue (8). The elevated PME
band in such a spectrum is most characteristic of oxygendeprivation, where the tissue has resorted to anaerobic glycolytic metabolism for ATP production. Phospholipid Profiles. Zebra mussel 31P NMR, phospholipid profile spectra are presented in Figure 3 and the quantitative data in Table 3. There are marked qualitative differences between the profiles from sponge-free and sponge-overgrown zebra mussels, most notably in the spectral profile from sponge-overgrown mussels, phosphatidylcholine and phosphatidylserine have been reduced and a series of lysophospholipid degradation products have been generated. These lyso-derivatives (LPG, LPA, LDPG, LGPLAS, LEPLAS, LPE, LPS, LAAPC, and LPC) are created through the action of a phospholipase-A2 activity acting on the parent phospholipids (PG, PA, DPG, GPLAS, EPLAS, PE, PS, AAPC, and PC). Such lysosomal activity is among the first steps in a digestive process. Another is the removal of the phospholipid polar headgroup through a phospholipase-D activity. The phospholipid that most prominently displays this activity is phosphatidylcholine. This phospholipid is decreased, while the deacylated product LPA is increased. Since it is apparent from signal-to-noise ratios that the total quantity of extracted phospholipid per unit zebra mussel is reduced, a phospholipase-C activity, which removes the phosphate group from the glycerol moiety rendering the lipid residue undetectable by 31P NMR, also must be present. Further, the lysophosphatidic acid spectral signal is rather broad and exhibits finestructure, indicating the presence of a group of LPA derivatives. These derivatives most probably are created by the actions of both a phospholipase A1 activity, which deacylates phospholipids at the glycerol C1-position, and a phospholipase A2 activity.
to colonize native unionid species (5). The exotic D. polymorpha, however, offers E. fragilis several benefits not found with native Unionidae: it is comparatively immobile, it does not borrow, and it occupies substrates more suitable to sponges. It is the zebra mussel’s size, however, that offers the greatest and probably the key benefit. In comparison, the much smaller size enables the sponge to overgrow the zebra mussel in a relatively short time, a factor critical to the seasonally limited growth cycle of sponges in Lake Michigan. With encapsulation, the mussel soon dies. Thus incorporated within the matrix of the sponge, the normal cellular migration typically associated with sponge feeding takes place. This process, which may or may not involve the participation of bacteria, provides the sponge with an ample internal food supply. Whether these activities of the freshwater sponge E. fragilis may represent a mechanism that can, or should, be viewed as a natural control for zebra mussels is beyond the scope of this paper. Its identification as a native species exerting significant impact on a local level, however, provides the opportunity for serious, long-term investigation in similar environments or on a more widespread basis. The sponge acting as predator has been documented recently for a marine sponge (4). This report presents evidence that freshwater sponges in the Great Lakes also may exhibit similar behavior.
Discussion
(1) Frost, T. F. In Ecology and Classification of North American Freshwater Invertebrates; Thorp, J. H., Covich, A. P., Eds.; Academic Press: New York, 1991; pp 95-124. (2) Zebra Mussels, Biology, Impacts, and Control; Nalepa, T. F., Schloesser, D. W., Eds.; Lewis Publishers: Boca Raton, FL, 1993. (3) Lauer, T. E.; Spacie, A. J. Great Lakes Res. 1996, 22, 77-82. (4) Vacelet, J.; Boury-Esnault, N. Nature 1995, 373, 333-335. (5) Ricciardi, A.; Reiswig, H. M. Can. J. Zool. 1993, 71, 665-681. (6) Ricciardi, A.; Snyder, F. L.; Kelch, D. O.; Reiswig, H. M. Can. J. Fish Aquat. Sci. 1995, 52, 2695-2703. (7) Burt, T. C.; Glonek, T.; Ba´ra´ny, M. J. Biol. Chem. 1976, 251, 2584-2591. (8) Burt, C. T.; Glonek, T.; Ba´ra´ny, M. Science 1977, 195, 145-149. (9) Ba´ra´ny, M.; Glonek, T. In Methods in Enzymology; Frederiksen, D. L., Cunninghan, L. W., Eds.; Academic Press: New York, 1982; Vol. 85B, pp 624-676. (10) Meneses, P.; Glonek, T. J. Lipid Res. 1988, 29, 679-689. (11) Glonek, T.; Merchant, T. E. In Advances in Lipid Methodology; Christie, W. W., Ed.; The Oily Press, Ltd.: West Ferry, 1995; pp 1-47. (12) Early, T. A.; Kundrat, J. T.; Schorp, T.; Glonek, T. Comp. Biochem. Physiol. 1996, 114B, 77-89. (13) Ring, W. Chicago Tribune 1991, 105 (Monday, April 15, Sec2), 1. (14) Penney, J. T.; Racek, A. A. In U.S. Natural Museum; Smithsonian Institution Press: Washington, DC, 1968; Bull. No. 272. (15) Goettel, R. The Helm (Illinois-Indiana Sea Grant Program) 1993, 10, 3. (16) Lauer, T. E., personal communication. (17) Greiner, J. V.; Kopp, S. J.; Glonek, T. Arch. Ophthalmol. 1984, 102, 770-771. (18) Glonek, T. In P-31 NMR Spectral Properties in Compound Characterization and Structural Analysis; Quin, L. D., Verkade, J. G., Eds.; VCH Publishers: New York, 1994; pp 283-294. (19) NOAA. Calumet and Indiana Harbors, 19th ed.; 1990; Chart # 14929. (20) Ruttner, F. Fundamentals of Limnology, 3rd ed.; University of Toronto Press: Toronto, Canada, 1974; pp 183-191. (21) Kopp, S. J.; Glonek, T.; Greiner, J. V. Science 1982, 215, 16221625. (22) Glonek, T.; Kopp, S. J. Magn. Reson. Imag. 1985, 3, 359-376. (23) Meneses, P.; Para, P.; Glonek, T. J. Lipid Res. 1989, 30, 458-461.
The described interactions between E. fragilis and zebra mussels result in mussel cellular energy deprivation, death, and digestion. The most probable mechanism appears to be anoxia, caused by the growing sponge interdicting the mussel’s oxygenated water supply. At some point in the process, mussel valves are closed and apparently held that way by the growing sponge matrix. The missing byssus on large adult sponge-overgrown mussels indicates an attempt on the part of the mussel to relocate to a more favorable habitat, which is one of the actions the adult mussel may take under stress. The sponge overgrowth is tenacious, however. Deprived of oxygen and nutrition, the mussel’s energy reserves are depleted. Eventually, this results in the animal’s inability to maintain its ATP reserves. Collapse of the ATP reserve (8) initiates a chain of chemical processes resulting in runaway chemical hydrolysis and animal death signaled by dissolution of the animal’s cellular membranes that may be actively promoted by the encapsulating sponge. Cellular membranes support the chemical machinery of the cell. When these are disrupted, the cell can no longer function. Disruption of cell membranes occurs following hydrolysis of the component phospholipids to their lyso derivatives. The lyso-phospholipids are efficient detergents and extremely toxic. Their accumulation results in cellular dissolution. The relevance of these sponge-zebra mussel interactions finds its place in consideration of those mechanisms that, individually or collectively, represent components contributing toward the dynamic equilibrium within a system. This is particularly important in cases wherein certain organisms, already adapted to exploit similar native species, can expand their niche to incorporate a newly introduced (exotic) species. The apparent predation of D. polymorpha by sponges may not describe an entirely new phenomenon but rather a variation of normal adaptive behavior. E. fragilis is known
Acknowledgments The authors thank Lake Michigan Charters, Inc., for their support, and the John G. Shedd Aquarium for their cooperation in the identification of the sponge species.
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Received for review August 25, 1998. Revised manuscript received March 19, 1999. Accepted March 22, 1999. ES980874+
