Metabolic changes induced in oysters (Crassostrea virginica) by the parasitism of Boonea impressa (Gastropoda: Pyramidellidae)

Camp. Biochem. Physiol. Vol. 90A, No. 2, pp. 279-290, 1988

0300-9629/88 $3.00 + 0.00

0 1988 Pergamon Press plc

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METABOLIC

CHANGES

(CRASSOSTREA

INDUCED

IN OYSTERS

VIRGINICA) BY THE OF BOONEA IMPRESSA

PARASITISM (GASTROPODA:

PYRAMIDELLIDAE)

M. E. WHITE,* E. N. POWELL,S. M. RAY,~ E. A. WILSONand C. E. zASTROW$ Department of Oceanography, Texas A&M University, College Station, Texas 77843 and tDepartment of Marine Biology, Texas A&M University at Galveston, Galveston, Texas 77550, USA (Received 21 October 1987) Abstract-l.

The ectoparasitic snail Boonea (= Odostomiu) impressa is an important parasite of oysters. Although results varied among tissues and levels of parasitism, all oysters parasitized by B. impressa exhibited alterations in biochemical composition after 1 month. 2. Few changes were observed in small (24 cm) or large (68 cm) oysters parasitized by 5 or 10 snails, respectively. In contrast, carbohydrate concentration decreased in small oysters parasitized by 15 or 25 snails and increased in large oysters parasitized by 30 snails. 3. Decreased carbohydrate content probably resulted from direct removal of assimiliated carbon by B. irnpressu. Increased carbohydrate concentration may be due to the combined impact of B. irnpressawith infections by the protozoan Perkinsus (= Dermocystidium) marinus. P. marinus infection increased in intensity with snail parasitism. Increased lipid content in the mantle tissue of large oysters and increased taurine content in most oysters and oyster tissues could be similarly explained. 4. Overall, mantle tissue was the most severely affected, possibly because snail parasitism produces direct injury to this tissue as well as the systemic impact caused by removal of assimilated carbon.

community dynamics. In communities where the host is the dominant species, as Crassostrea virginica is on Previous studies examining marine parasitism docuoyster reefs, reduced health of the host may substanmented both the occurrence and abundance of paratially influence the entire community. The purpose of sites and indicated the importance of these organisms this study was to examine the effect of an ectoparasite in marine communities (e.g. Elder, 1979; Cake and (Boonea impressa) on the health and biochemical Menzel, 1980; Eiras, 1986). Marine parasites can composition of its host (C. virginica). Like other change the host’s growth rate (White et al., 1984), parasitic organisms, B. impressa directly removes reproduction (Reaka, 1978; Beck, 1980; Bell and nutrients from the host. B. impressu accomplishes this Stancyk, 1983) and chance of mortality (Galleni et by piercing the mantle tissue of the oyster using a al., 1980). Only a few studies, however, have exam- stylet at the end of a long proboscis, after which the ined the impact of marine parasites on the health and host’s body fluids are sucked by means of a buccal biochemical composition of their hosts (e.g. Ford, pump (Fretter and Graham, 1949; Allen, 1958; Maas, 1986a, b). Most of these studies concentrated on the 1965; White et al., 1984). B. impressa is both ubiquieffects of parasitism on the free amino acid pool tous and abundant on oyster reefs from Massachu(Feng et al., 1970; Richards, 1969; Kasschau, 1975; setts to the Gulf of Mexico, with as many as 100 Viva& and Cuq, 1981; Soniat and Koenig, 1982). parasites per oyster observed (Hopkins, 1956; Despite the paucity of marine documentation, re- Robertson, 1978; Larsen, 1985). White et al. (1984, search on terrestrial organisms indicates that, in 1987) documented reduced growth and an increased many cases, lipid, carbohydrate and protein com- prevalence of disease in oysters parasitized by B. position also should be altered substantially by paraimpressa, confirming the deleterious impact of this sitism (Brockelman, 1978; von Brand, 1979; Dahlectoparasite on oyster populations. These effects can man and Vinson, 1980; Thompson and Binder, 1984). be ascribed to a reduction in that portion of the A knowledge of how parasites affect growth, health oyster’s assimilated energy available for growth and and reproduction of their hosts is necessary to undermaintenance (Ward and Langdon, 1986; White et al., stand the factors governing host population and in press). That is, B. impressu may induce a state of mild to severe starvation in actively feeding oysters. To assess the impact of B. impressa on oyster nutri*Present address: National Center for Atmospheric tion, changes in total soluble protein, lipid and Research,5 Environmental and Societal Impacts Group, carbohydrate concentration and the free amino acid PO Box 3000, Boulder, Colorado 80307-7000, USA. pool (FAA) were examined in parasitized and un&The National Center for Atmospheric Research is sponparasitized oysters. All of these measures are comsored by the National Sciencd Foundation. _ monly used to monitor stress and health in bivalves IPresent address: University of Maryland, CEES, Chesapeake Biological Laboratory, Solomons, MD 20688 (e.g. Jeffries, 1972; Riley, 1980; Soniat and Ray, 1985). The results reported here indicate that B. USA. INTRODUCTION

279

M. E.

280

WHITE et al.

impressa significantly alters the metabolism of C. uirginica at abundances normally found in the field.

Table I. The number of snails placed on each oyster in experiments reported herein Number of snails per oyster

MATERIALS

AND METHODS

No snails

Laboratory procedures

Experiments were conducted using the flowing sea-water system at the Port Aransas Marine Laboratory. Large (68 cm) and small (24 cm) oysters were obtained from Confederate Reef near Galveston, Texas and Mud Island and Big Slough Reefs on Harbor Island, near Port Aransas, Texas, respectively. Each experiment consisted of a control and two experimental groups. Experimental groups were exposed to low or high levels of parasites covering a range of abundances similar to those observed by us on Texas reefs (Table 1). A set of 15 oysters, with each oyster in an individual container, was used for each of the three groups. Oysters were placed in PVC containers covered with coarse cloth and rubber bands to prevent snail immigration or emigration. Sea-water was fed by gravity into each container from a flowing sea-water system. Two sets of experiments were conducted: in the autumn, both large and small oysters were used; in the summer, only small oysters were used. At the beginning of the summer experiment, 10 oysters were killed as pre-controls so that any biochemical alteration due to collection or laboratory stress could be assessed (e.g. Koenig et al., 1981; Kendall et al., 1984; Haux et al., 1985). Individual underwater weights were recorded weekly for each oyster, according to the method of Andrews (1961). Concurrently, containers were cleaned of any deposited sediment and, if a snail was dead or missing, it was replaced. At the end of 4 weeks, all oysters were killed. Immediately, all tissues to be assayed (Table 2) were dissected out and placed in vials on dry ice. In addition, a small section of mantle tissue was removed for analysis of Perkinsus (= Dermocystidium) marinus using the thioglycollate method of Ray (1966). The intensity of P. marinus infection was assessed on the S-point scale commonly used to quantify infection intensity (Mackin, 1962). Biochemical analysis

Prior to analysis, selected tissues were lyophilized and weighed. Subsequently, tissues were homogenized at 4°C. The tissue homogenate was analysed spectrophotometrically for soluble protein by the Lowry method, as modified by Peterson (1977). The tissue homogenate was brought up to 10% trichloroacetic acid (TCA) with 50% TCA in the cold. Samples were spun, the pellet resuspended in SDS-NaOH for 30 min (Powell et al., 1982) spun again and the supernatant analysed. Bovine serum albumin was used as the standard. Lipids were extracted from the tissue homogenate using chloroform/methanol and measured gravimetrically (Folch et al., 1957). Total carbohydrate was determined in both the water phase and pellet obtained from the chloroform/methanol extraction of the tissue homogenate by the spectrophotometric method of Hewitt (1958). Samples were evaporated and rehydrated with 5% sulphuric acid to remove chloroform which interfered with the assay. Sucrose standards were used.

Low levels of snails

Autumn experiment Small Oysters Large Oysters

0 0

5 IO

15 30

Summer experiment Small Oysters

0

IO

25

Free amino acid (FAA) analyses were performed on a Dionex amino acid analyser using a lithium citrate buffer system and o-phthalaldehyde (OPA) as the detecting compound. Ninhydrin also was used for detection of amino acids in the total tissue of small oysters in the autumn because OPA fails to detect proline. No significant changes were observed in proline in the samples; hence OPA was used as the sole detecting agent for the remaining analyses. Norleucine and a-amino butyric acid were added as internal standards. The latter was not present naturally in sufficient quantities to compromise the analysis. Traces of chloroform in the water phase of the initial chloroform/methanol extraction interfered with the analysis; hence the water phase was extracted a second time using only chloroform. Samples were centrifuged in the cold and the water phase from the second extraction analysed for free amino acids. Recoveries (initial chloroform/methanol extraction through final analysis) were >95% for all amino acids examined, including the relatively unstable glutamine and hypotaurine, as assessed by standards. Hypotaurine was identified as compared to the coeluting urea (see Amende and Pierce, 1978) by (a) its relative reactivity to OPA and ninhydrin and (b) successful oxidation with iodine to taurine. Statistical analysis concentrated on taurine, hypotaurine, glycine, alanine, glutamic acid, aspartic acid, serine, threonine and glutamine which made up the bulk of the FAA pool that could be readily determined using OPA. Condition index was calculated as the ratio between shell-less tissue dry weight and mantle cavity volume (see Lawrence and Scott, 1982). Statistical analyses

A multivariate analysis of covariance (MANCOVA) was conducted to test for significant changes in biochemical composition. Parallelism was maintained as described by Smith and Coull (1987). Least squares means were used to examine the change in specific components at different levels of parasitism. Parasite level and the covariates dry tissue weight, mantle cavity volume and Perkinsus level were the independent variables in the model. In one case-the mantle tissue of small oystersdry weight covaried with the level of parasitism. Consequently, dry weight was deleted from the model and the remaining dependent variables were normalized to tissue dry weight. Although B. impressa can affect the intensity of infection by P. marinus in oysters (see White et al., 1987), P. marinus itself has a major impact on

Table 2. Tissues examined for biochemical analyses in the summer and autumn exoeriments Tissues examined

Autumn

Oyster group

Mantle

Gill

Adductor

Small Large

X

X

X

Small

X

X

Total

experimenl

Summer experimenr X indicates tissues examined.

High levels of snails

X

281

Oysters parasitized by Boonea biochemical composition (Soniat and Koenig, 1982). Accordingly, the intensity of infection by P. murinus was used as an independent variable. Cumulative weight gain over the entire experiment was examined as a dependent variable, along with the biochemical components. A detailed discussion of weekly weight gain and factors influencing shell deposition has been reported by White ef al. (in press). The MANCOVA was also used to examine each component of the FAA pool. In addition, the FAA pool was examined in several other ways: (a) the total pool, (b) the total pool less taurine and hypotaurine, (c) the total pool less taurine, hypotaurine, glycine and alanine, (d) each amino acid as a percent of the total pool and (e) the ratios, taurine/hypotaurine, aspartic acid/alanine, and taurine/ glycine. Taurine and hypotaurine are the only amino acids examined which are not found in protein. Therefore, it is important to examine changes in the pool exclusive of these amino acids. Examination of changes in the minor constituents of the FAA pool was facilitated by deleting taurine, hypotaurine, glycine and alanine from the analysis. Significant changes in FAA concentration can be due to generalized increases or decreases in the pool or specific changes in individual FAA. Which of the two is important can be assessed using the percentage composition of each FAA in the pool (Powell et al., 1984). The ratios, taurine/glycine and alanine/aspartic acid have been used to assess bivalve health and aerobic/anaerobic state, respectively (e.g. Jeffries, 1972; Collicutt and Hochachka, 1977). The taurine/hypotaurine ratio has not been used previously. Weight gain was not available for samples from precontrol oysters. Thus, rather than a MANCOVA, a Duncan’s Multiple Range test was used for statistical analyses that included the pre-controls. RESULTS

E&et of Boonea impressa on carbohydrate, protein and FAA concentration

lipid,

Changes in biochemical composition of oysters varied with parasite concentration (Figs 1-6) from relatively small alterations in oysters with few para-

sites (five for small and 10 for large oysters) to

substantial changes in oysters with many parasites (15 or 25 parasites for small oysters and 30 for large oysters). Carbohydrate concentration was affected more than lipid or protein concentration. In the experiment, carbohydrate decreased summer significantly in mantle tissue of small oysters parasitized by 25 snails (Table 3; Fig. 1). Carbohydrate also decreased, though not significantly, in the gill tissues of these oysters (Table 4; Fig. 2). In the autumn experiment, total carbohydrate decreased significantly in small oysters parasitized by 15 snails, but not by five snails (Table 5; Fig. 3). In contrast, significant increases in carbohydrate content were observed in the adductor and mantle tissues of large oysters parasitized by 30 snails (Tables 6 and 7; Figs 45). Likewise, gill tissue of highly parasitized, large oysters had an increased concentration of carbohydrate, although not significant (Table 8; Fig. 6). No significant changes occurred in protein concentration Lipid concentration changed in any tissue. significantly (increased) only in the mantle tissue of large oysters with 30 parasites (Fig. 4). Condition index was not significantly affected. The concentration of total free amino acids decreased significantly in the adductor muscle of large oysters. No other significant changes occurred in the total pool. Effect of Boonea impressa on individual amino aciak Most changes in individual amino acids occurred in the main contributors to the free amino acid pool, i.e. taurine, hypotaurine, glycine and alanine. Taurine concentration increased significantly in the mantle and gill tissues of parasitized, small oysters (summer experiment) and in the total tissue of small oysters (autumn experiment) (Figs l-3). No significant changes occurred in large oysters. In contrast, hypotaurine concentration changed significantly (decreased) only in gill tissue of large oysters. Glycine and alanine only showed significant changes in the

Mantle Tissue of Small

TotalFree

Oysters

Amino Acids (--)

TotalFreeAminoAcids(-) Total Free Amino Acids

Carbohydrate

Co;tol

COll$Ol

10 s,n,,ils

25 Snails

10 Snails

25 Snails

Fig. 1. Least squares means significance levels @lotted as I-P) from the MANCOVA of mantle tissue in small oysters. Total free amino acids include the entire pool examined, total free amino acids (-) does not include taurine and hypotaurine, and total free amino acids ( - - ) does not include taurine, hypotaurine, glycine and alanine.

M. E. WHITEet al.

282

Gill Tissue of Small Oyslers

Shell Weinht

Gain

Total Free Amino Acids i-4 Total Free Amino Acids (4 Total Free Amino Aci

Carbohydrate C0rl~0l 25 Snails

Control vs IO Snails

10 Snails

*lanine~~~~f

25 gaits

Control

Control

25 s”,“a,ts IO SZ3ik

10 Snails 25 Sviails

Fig. 2. Least squares means significance levels (plotted as 1-P) from the MANCOVA of gill tissue in small oysters. Total free amino acids-as for Fig. 1.

autumn experiment. Both decreased signi~~antly in the adductor muscle of large highly-parasitized oysters and glycine decreased significantly in the total tissue of small oysters parasitized by five snails. Although not always significant, alanine concentration also decreased in all three tissues (gill, mantle, adductor muscle) of highly parasitized, large oysters. Amino acids in smaller concentrations changed significantly only at high parasite levels and in the autumn experiment. Threonine decreased in the mantle tissues of large oysters, and glutamic acid decreased and glutamine increased in the adductor tissue of large oysters. Serine increased in small oysters.

The FAA pool can respond to stress in one of two ways: individual amino acids can change in concentration relative to others or the concentration of all amino acids can increase or decrease proportionally. The latter typically indicates a higher level of stress than the former (Powell et al., 1982, 1984). Significant changes in the FAA pool in oysters parasitized by B. irnpressa were primarily due to changes in the relative concentrations of individual FAA. The relative concentrations of the individual amino acids changed significantly in the total tissue and mantle tissue of small oysters (P = 0.01 and 0.007, respectively) and the adductor tissue of large oysters (P = 0.05). In small oysters, an increase in taurine relative to the

TotalTissue of Small Oysters

TotalFreeAmlnoAcids(--) Total Free Amino Acids (-) Total Free Amino Acids

/wz# A

/ /%?-c

Protein

/‘I

Lipid Carbohydrate

/ Control

Control

5 Snails

15fYrYails

5 Zaiis

15 X&its

Aspartic Acid Alanine ,:,

I@ R

/

/’ w,m

v,.e

060 Control

036 ,’ 0% , Control 5 Snails

15G%ls

5 &Is

-

15 Eails

Fig. 3. Least squares means significance levels (plotted as I-P) from the MANCOVA of total tissue in small oysters From the autumn experiment. Total free amino acids-as foe Fig. I.

Oysters parasitized by Boonea

283

Gill Tissue 01 Large Oysters

Total Free Amino Acids (--) lotalFr~AminoAcids(-)

TotalFree Amino

P I

i’

Acids

30 SvnSails 10 SvnHils 30 6ils

Fig. 4. Least squares means significance levels (plotted as 1-P) from the MANCOVA of gilI tissue in large oysters. Total free amino acids-as for Fig. I.

Adduclor Tissue of Large Oysters Threonifle

TotalFreeAminoAcids(--1 lotalFreeAminoAcids(-)

TotalFree

Amino Acids

_Lz

/ /tz?___

&?./_g3

f

1

/

Protein 64, Lipid Carbohydrate /A=

/

Control

Control

10 Snails

!OsY~ails 10 Lails

3OkZiails

Fig. 5. Least squares means significance levels (plotted as 1-P) from the MANCOVA of adductor tissue in large oysters. Total free amino acids-as for Fig. 1.

Mange Tissue of Large Oysfers

Shell

TotalFreeAminoAcids(TotalFreeAminoAcids( Total Free Amino

Carbclhydrate

@(l-P)<090 30 Snails 10 Snails 30

@i

i

Co$rol 30Snails

-P)>O.sQ

10 psails

10 Snails SOSnails

Fig. 6. Least squares means significance levels (plotted as I-P) from the MANCOVA of mantle tissue in large oysters. Total free amino acids-as for Fig. 1.

7

284

M. E. WHITE et al. Table 3. Mean and standard deviation for biochemical components in the mantle tissue of small oysters in the summer experiment

Variable -Total free amino acids FAA (-) FAA (- -) Aspartic acid Threonine Serine Glutamic acid Glycine Alanine Taurine Hypotaurine Glutamine Lipid Protein Carbohydrate Shell weight gain Mantle cavity volume Perkinsus level

Tissue dry weight

Oysters with no snails (n=ll) SD Mean -~-750 299 121 41 50 17 15 5 2 3 10 9 22 10 33 12 38 IS 211 87 352 223 0 0 78 24 334 58 64 32 -0.04 0.20 2.94 0.82 1.09 I .64 21.87 7.86

Oysters with 10 snails per oyster (n = IO) Mean SD

Oysters with 25 snails per oyster (n = 3) Mean SD

Pm-control oysters (no snails) (n = IO) Mean SD

799 106 44 16

207 47 18 7

1

1

793 72 106 18 45 4 13 4 5 6 6 2 20 6 32 7 29 I6 338 28 348 60 0 0 98 19 332 54 31 6 0.00 0.14 2.95 0.49 0.50 0.87 14.90 1.03

113 148 58 20 3 5 29 42 48 335 290 0 48 338 41 N/A N/A N/A 23.99

8 3 19 8 27 10 34 20 324 46 378 147 0 0 93 27 354 38 66 25 -0.02 0.08 3.10 0.48 1.00 I .75 20.98 4.05

185 43 28 8 4

t 25 14 17 96 104 0 29 42 19 N/A N/A N/A 4.7

Amino acids are expressed in pmol/g dry weight; carbohydrate, lipid and protein, in mg/g dry weight; volume, in ml; shell weight gain, in g; dry weight, in mg. Perkinsus level is the mean intensity of infection by P. marinus assessed on the 5-point scale of Mackin (1962). FAA (-) is the free amino acid pool minus taurine and hypotaurine. FAA f - - ) is the free amino acid pool minus taurine, hypotaurine, glycine and alanine. Values of weight gain and shell volume are for the entire animal, P. marinus, for the mantle tissue.

remaining FAA was the primary cause. In large oysters, a reduction in pool size was produced by disproportionate losses of taurine, glycine and alanine. The ratio of taurine to giycine was significantly affected by parasitism only in the mantle tissue of small oysters (P = 0.04). Differences were most pronounced between control oysters and oysters with 10 parasites. The ratio of taurine to hypotaurine was

only influenced significantly by parasitism in the gill tissue of small oysters (P = 0.10). Parasitism also affected the ratio of aspartic acid to alanine in the total tissue of small oysters (P = 0.10). In the summer experiment, with the exception of the FAAs, the biochemical components of the precontrol oysters rarely differed significantly from the controls (Table 9). FAA, however, frequently differed

Table 4. Mean and standard deviation for biochemical components in the gill tissue of small oysters in the summer experiment

Variable Total free amino acids FAA (-) FAA (--‘-) Aspartic acid Threonine Serine

Glutamic acid Glycine Alanine Taurine Hypotaurine Glutamine Lipid Protein Carbohydrate Shell weight gain Mantle cavity volume Perkinsus level Tissue dry weight

Oysters with no snails in = 11) SD Mean s39 346 140 63 65 34 23 I2 2 2 11 11 30 14 39 23 34 I5 336 102 363 296 0 0 110 23 474 92 72 27 0.02 0.03 2.96 0.78 I .oo 1.59 20.69 6.5

Oysters with 10 snails per oyster (Jr = 9) Mean SD

Oysters with 25 snails per oyster (n = 3) Me& SD

245 771 1019 123 32 209 97 53 I3 21 6 37 1 0 2 7 4 11 24 8 4s 55 32 15 56 37 16 332 56 559 326 211 251 0 0 0 109 14 143 428 58 473 55 76 37 -0.02 0.08 0.01 3.10 0.48 2.95 1.oo 1.75 0.50 21.65 7.19 16.16

340 201 84 31 2 9 44 54 65 193 78 0 100 24 27 0.14 0.49 0.87 1.29

Pre-control oysters {no snaits) (n = IO) Mean SD 591 145 48 21

217 43 14 9

1

I

6 21 58 50 223 256 0 95 357 58 N/A N/A N/A 35.47

3 7 32 I6 58 167 0 21 126 30 N/A N/A N/A 11.69

See Table 3 footnote for explanation of units and terms used. Values of weight shell volume are for the entire animal, P. marinus, for the mantle tiSsUe.

gain and

285

by Boonea

Oysters parasitized

Table 5. Mean and standard deviation for biochemical components in the total tissue of small oysters in the autumn experiment

Variable Total free amino acids FAA (-) FAA (- -) Aspartic acid Threonine Serine Glutamic acid Glycine Alanine Taurine Hypotaurine Glutamine Lipid Protein Carbohydrate Shell weight gain Mantle cavity volume Perkinsus level Tissue dry weight

Oysters with no snails (n = 8) Mean SD 568 288 68 25 3 4 32 106 II5 188 92 3 88 63 103 0.04 2.00 0.06 142.67

Oysters with 5 snails per oyster (n = 7) Mean SD

694 215 266 17 69 I8 24 IO 2 I 4 I I 31 9 9 82 44 44 II5 31 40 281 134 67 I48 99 19 8 8 4 98 IO 16 67 7 I2 99 34 39 0.05 0.06 0.04 0.013 2.21 0.65 0.18 0.00 0.00 39.20 144.41 44.57

93 61 15 7 2

Oysters with I5 snails per oyster (n = 8) Mean SD 605 214 64 24 3 5 29 I08 IO1 220 III 5 95 68 58 0.00 I.91 0.06 142.43

82 58 8 IO I I 6 47 29 50 43 7 22 I8 22 0.02 0.48 0.18 66.41

See Table 3 for exnlanation of units and terms used. Values for P. marinusare for the mantle. tissue.

significantly. Taurine was lower in the gill tissue of pre-controls compared with other groups and alanine was higher (Table 10). Glycine was higher in the mantle tissue of pre-controls (Table 10). Thus, oysters did experience some laboratory stress during the course of the experiment, but most effects were minor. Eflects of P. marinus

for P. marks, we analysed only these data. marinus significantly influenced the concentrations

of glycine, taurine, serine and aspartic acid in large oysters (P = 0.03, 0.05, 0.03 and 0.05, respectively). In addition, P. marinus affected the ratio of aspartic acid to alanine (P = 0.04). DISCUSSION

on biochemical composition

P. marinus was reported to alter the amino acid composition of the mantle tissue of C. uirginica (Soniat and Koenig, 1982). It did so in our experiments as well. Because only the mantle was assayed

Concentrations of carbohydrate, lipid, protein and FAA in control oysters agreed with those previously determined for Crassostrea gigas and Crassostrea uirginica (Sidwell et al., 1979; Soniat and Ray, 1985;

Table 6. Mean and standard deviation for biochemical components in the mantle tissue of large oysters in the autumn experiment

Variable Total free amino acids FAA (-) FAA (- -) Aspartic acid Threonine Serine Glutamic acid Glycine Alanine Taurine Hypotaurine Glutamine Lipid Protein Carbohydrate Shell weight gain Mantle cavity volume Perkinsus level Tissue dry weight

Oysters with no snails Me;t = 10) SD 606 272 98 36 4 20 33 62 126 244 126 4 94 II4 I45 -0.45 24.97 4.15 78.02

P.

I80 83 41 I9 3 35 I4 32 61 107 50 4 :“7 35 0.28 4.02 I.16 21.04

Oysters with IO snails per oyster (n = IO) Mean SD

Oysters with 30 snails per oyster Me(ann=lo) SD

548 I51 229 80 75 28 28 7 3 2 6 2 33 21 66 I9 88 45 200 I05 123 31 4 5 86 25 80 25 I40 48 -0.68 0.39 22.16 6.27 4.17 1.06 88.81 19.66

676 268 255 86 80 29 35 20 2 2 9 7 28 8 77 34 97 48 287 I25 134 65 5 6 I28 53 84 38 I88 I13 -0.64 0.55 23.71 6.76 4.3 I .29 78.65 27.18

See Table 3 footnote for explanation of units and terms used. Values for shell volume and weight gain are for the entire animal.

M. E. WHITE et al.

286

Table 7. Mean and standard deviation for biochemical components in the adductor tissue of large oysters in the fall experiment

Variable Total free amino acids FAA (-) FAA f- -) Aspartic acid Threonine Serine Glutamic acid Giycine Alanine Taurine Hypotaurine Glutamine Lipid Protein Carbohydrate Shell weight gain Mantle cavity volume Perkinsus level Tissue dry weight

oysters with no snails (n = 10) Mean SD .507 234 302 115 54 20 26 14

Oysters with IO snails per oyster (n = 10) Mean SD 502 296 54 24

179 101 16 II

I 3 23 117 131 III 93

I 37 82 54 -0.45 24.91 4.15 274.63

:. 7 45 61 149 43 2 8 30 22 0.28 4.02 1.16 129.16

: 20 147 105 86 122 3 41 107 44 -0.68 22.16 4.17 210.19

: 6 60 31 49 47 6 7 67 14 0.39 6.27 I .06 82.95

Oysters with 30 snails per oyster (n = IO) Mean SD 384 237 51 24

217 192 28 18

I

1

4

3

;: 88 56 90 S 45 86 69 -0.64 23.71 4.30 219.98

7: 64 42 56 8 I1 30 25 0.55 6.76 1.29 69.20

See Table 3 footnote for explanation of units and terms used. Values for mantle cavity volume and weight gain are for the entire animal, P. marinus, for the mantle tissue.

Table 8. Mean and standard deviation for biochemical components in the gill tissue of large oysters in the fall experiment Oysters with no snails Meg=

‘) SD

Total free amino acids FAA f-) FAA (- -) Aspartic acid

Variable

855 331 86 42

342 163 38 23

Serine Threonine Glutamic acid Glycine Alanine Taurine Hypotaurine Glutamine Lipid Protein Carbohydrate Shell weight gain Mantle cavity volume Perkinsus level Tissue dry weight

72 3I 31 IO 120 52 125 79 372 130 152 80 4 4 110 13 160 :; 73 -0.57 0.30 25.19 3.78 3.90 I .47 131.48 24.88

Oysters with 10 snails per oyster Meg = 3, SD 740 282 14 31 51 32 97 112 298 160 4 103 184 81 -0.94 20.61 3.00 148.18

Oysters with 30 snails per oyster Mzn = 6) SD

173 78 29 I9

561 244 64 25

0 3;

62 3;:

21 47 64 4 6 55 8 0.86 8.63 0.71 70.29

126 59 24 16

82 223 94 5 104 140 91 -0.54 23.86 3.91 142,68

: 11 26 31 69 49 4 12 46 37 0.45 8.20

1.43 37.54

See Table 3 footnote for explanation of units and terms used. Values for mantle cavity volume and weight gain are for the entire animal, P. marinus, for the mantle tissue.

Table 9. Results of Duncan’s Multiple Range tests examining differences in protein, lipid and carbohydrate among pre-control, control and experimental levels in the summer experiment Mantle tissue Number of snails per oyster .O Pre-control

Carbohydrate concentration -~A A B AB

Protein concentration . . ~._ A A A A

~Carbohydrate Lipid concentration concentration ..-__-... AB A A A A A A B

concentration

Gill tissue Protein concentration

Different letters indicate significant differences among groups (01= 0.05) within columns only.

A AB A B

Lipid concentration A A A A

Oysters parasitized by Boonea Table 10. Results of Duncan’s Multiple Range tests examining differences in amino acid concentrations among precontrol, control and experimental levels in the summer experiment Number of snails per __ oyster --. 0 10 25 Pre-control

Mantle tissue .---___ Taurine Glycine B AR A AB

AB B AB A

Gill tissue -... Alanine Taurine B AB AB A

Different letters indicate significant differences groups (a = 0.05) within columns only.

B B A C among

Soniat and Koenig, 1982). Boonea impressa significantly influenced carbohydrate, lipid and amino acid composition, particularly in the mantle tissue and in small oysters, but alterations occurred in al1 tissues and oyster sizes examined. At equivalent densities, B. impressa will remove a higher proportion of the energy available for growth in small oysters (White et al., in press); hence the accentuated effect of snail parasitism on small oysters. B. impressa feeds directly on the mantle tissue, thus the more pronounced effects of snail parasitism on the biochemical composition of the mantle may be a direct result of combining a localized inff ammatory reaction associated with wound healing {e.g. Cheng, 1967) with a systemic effect produced by removal of a portion of the oyster’s assimilated carbon. Carbohydrate decreased in small oysters parasitized by B. impressa, but increased in large (6-8 cm) oysters. In oysters, carbohydrate, especially the important storage compound, glycogen, generally increases during the winter and early spring and then decreases as lipid is produced during gonadal maturation (Soniat and Ray, 1985). Spawning begins in late spring and continues through to early autumn (Soniat and Ray, 1985). During this period, carbohydrate reserves are lower than the rest of the year (Lee and Pepper, 1956; Sidwell et at., 1979). Accordingly, the reduction in carbohydrate in parasitized oysters from the summer experiment probably resulted from a decrease in the amount of assimilated carbon available for growth and maintenance when stored reserves were already low. Probably, a combination of direct feeding by the snail and the snail’s ability to disrupt the normal feeding behavior of the host (Ward and Langdon, 1986) were involved. Explaining the increase in carbohydrate content in gill and adductor tissue of large oysters is more complex. Large oysters differed from small ones in several important respects. Being larger, the amount of total assimilated carbon consumed by the snails was lower even though more snails were used (see White et al., in press). Therefore, a direct reduction of carbohydrate reserves by snail feeding might not be expected. Additionally, percent infection and infection intensity of P. marinus was significantly higher in large oysters (White et al., 1987). The effect of P. m~ri~us on oyster metabolism is poorly known, but many endoparasites affect their host’s carbohydrate metabolism (e.g. Mengebier and Wood, 1969; Hopkins, 19.57; Mackin, 1962). Rates of gluconeogensis are frequently modified (Thompson and Binder, 1984; Thompson, 1986). In our experiments, feeding by B. impressa significantly increased the

287

severity of infection by P. mar&s (White et al., 1987), thus increasing any interaction of P. marinus with host metabolism. One interaction might be an increase in gluconeogenesis. Although speculative, it is interesting that the only oysters in which carbohydrate decreased (small oysters) were those with a low prevatence of P. marinus. Carbohydrate composition should be affected more by snail parasitism than lipid or protein composition because carbohydrate is the main storage product of oysters (Galtsoff, 1964) whereas the latter two include important structural components and are relatively resistant to changes in assimilated carbon (e.g. Fisher and Newell, 1986). Soluble protein did not change significantly in any of the six experiments. The only significant change in lipid was an increase in mantle tissue of large oysters in the autumn. Mackin (1962) described an increase in lipid content during infection by P. marine. The intensity of infection by P. ma&us increased significantly in large oysters parasitized by 30 snails during the autumn experiment. Changes in the amino acid pool often occur when organisms are stressed (Jeffries, 1972; Powell et al., 1982; Heavers and Hammen, 1985). Typically, either the entire free amino acid pool is more-or-less uniformly reduced, or alterations in the relative proportions of the FAA in the pool occur. Both effects have been found in parasitized invertebrates (Feng at al., 1970; Watts, 1971; Trede and Becker, 1982). Parasitism by B. impressa had two pronounced impacts on the FAA pool: (1) the presence of B. impressa produced changes in the relative concentrations of certain amino acids; (2) increasing severity of P. marks infection produced by B. impressa augmented the effect of P. marinus on the FAA pool. Responses of individual amino acids to parasitism varied considerably. Frequently the concentration of a specific amino acid declined in one tissue and/or oyster group and increased in another. Some definite patterns were observed, however. Taurine concentration normally increased significantly in parasitized small oysters. Taurine is closely associated with osmotic regulation (Florkin, 1966; Feng et al., 1970). Feng et al. (1970) postulated that increased taurine in parasitized organisms could be attributed to maintenance of osmotic balance necessitated by depletion of the other components of the host’s FAA pool. FAA are important in the osmotic balance of oysters (Hammen, 1969) and the remaining components of the FAA pool of small oysters did generally decrease in our experiments. Per~insus mar&us aIso influenced the structure of the FAA pool in oysters. In the mantle tissue of large oysters, the severity of infection by P. ma&us significantly influenced the concentrations of taurine, glycine, serine and aspartic acid. Increased taurine concentration and changes in glycine and aspartic acid concentration with increased severity of infection by P. marinus were also observed by Soniat and Koenig (1982). Because B. impressa intensified P. marinus infection in our experiments, part of the effect of P. marinus that we observed could ultimately be attributed to the snail. Particularly in large oysters, the FAA pool exclusive of taurine and hypotaurine generally decreased.

M. E. Wnm

288

Reductions occurring in the FAA pool may result from the breakdown of amino acids for energy or direct removal by the snail. metabolism of some non-essential amino acids is closely linked to carbohydrate metabolism (Heavers and Hammen, 1985; Gabbott, 1976). For instance, an increase in carbohydrate content in large oysters in the fall coincided with a decrease in alanine concentration. Glycine and glutamic acid also decreased. Although most significant differences in amino acids occurred between control and highly parasitized oysters, in a few instances the less heavily parasitized group of oysters had lower amino acid concentrations than either controls or the highly parasitized group. Possibly, some breakdown of protein occurred in the highly parasitized oysters to replenish the FAA pool (e.g. Riley, 1980). Ratios between amino acids changed little. The taurine to glycine ratio, used as an indicator of stress in marine invertebrates (Jeffries, 1972), changed significantly in the mantle of highly parasitized, small oysters. Unlike other cases where a change in glycine was the primary cause (e.g. Roesijadi, 1979; Roesijadi and Anderson 1979), an increase in taurine concentration produced the effect here. Had valve closure produced by parasitism been of sufficiently long duration to induce anoxia, a change in the aspartic acid to alanine ratio might have been observed (e.g. Collicutt and Hochachka, 1977; Powell et af., 1982), but none was observed. Hypotaurine, which has rarely been reported in invertebrates because it coelutes with urea in many separatory procedures (Amende and Pierce, i978), was present in high concentrations in these oysters. The only presently known function of hypotaurine is as an intermediate in taurine formation (Rosa and Stipanuk, 1985; Jacobsen and Smith, 1968). The concentrations we observed, however, seem extremely high for a simple intermediate.

et al.

commonly encountered in the field. In comparison to the relatively minor impact on biochemical composition, White et ai. (in press) observed significant changes in growth rate, fecundity and disease (P. mar&us) severity at this level of parasitism. Except when abundances are very high (40 or more snails per oyster), the primary impact of snail feeding on all but the smallest oysters is to reduce the oyster’s net productivity (White et al., in press); hence impacts on growth and fecundity are expected. On the other hand, changes in lipid, protein and carbohydrate content usually require a negative energy balance (e.g. starvation-Swift and Ahmed, 1983; Riley, 1980). Consequently, changes in biochemical composition should have been minor in our experiments. This was the observed result. A~ordingly, reduced growth and fecundity and increased disease intensity should be the primary effects of snail parasitism in field populations, except in the smallest oysters. Oysters 30mm or less in size, fed upon by even a few snails, may have a negative energy balance, however. In these oysters, significant biochemical effects and even mortality can be expected as a result of snail feeding. Acknowledgements-We thank C. Kitting for assistance with labomtory experiments and A. Tirpak for help during

field collection. Thanks also to A. Craia, D. Davies, C. Fox, A. Logan and M. Meyers who zded with the laboratory experiments. The University of Texas generously provided us with space and use of the facilities at the Marine institute in Port Aransas. We appreciate helpful comments made by J. Parrack and Drs D. Owens and M. Sweet, on earlier versions of the manuscript. We thank R. Covington who typed the manuscript and tables. This research was funded bv institutional erants NA83AA-DO0061 and NA85AA-‘D-FG128 to Teias A&M University by the National Sea Grant Program, National Oceanic and Atmospheric Administration, US Department of Commerce to E.P. and S.R., and a Sea Grant Marine Fellowship and a

Postdoctoral Fellowship from the National Center for Atmospheric Research to M.W. We appreciate this support.

CONCLUSIONS

Changes in the ~ncentrations of lipid, carbohydrate and the FAA pool were observed in oysters parasitized by Boonea impressa. Most of these changes can be explained by (1) a direct impact of B. impressa on the mantle tissue, (2) a systemic effect produced by a reduction of the assimilated carbon available to the oyster and (3) the interaction of this snail with the protozoan parasite, Perkinsus marinus. Even large oysters parasitized by a seemingly insignificant number of B. impressa suffered significant changes in their normal metabolic balance. Ford (1986a, b) and Soniat and Koenig (1982) also recorded significant metabolic changes in Hap1osForid~um ne~son~-parasitized oysters and P. rn~r~us-parasitized oysters, respectively. The impact on small oysters was more pronounced, especially in the mantle tissue where B. impressa feeds. In large oysters, most of the impact could be ascribed to the influence of B. impressa on P. marinus infection, whereas, in small oysters, a direct reduction in the availability of assimilated carbon was more important. The lower of the two parasite levels used in these experiments (e.g. five for small oysters) is more

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