Printer Friendly

Characterization of phenoloxidase from Crassostrea virginica hemocytes and the effect of Perkinsus marinus on phenoloxidase activity in the hemolymph of Crassostrea virginica and Geukensia demissa.

ABSTRACT The enzyme phenoloxidase is believed to be a component of internal defense in invertebrates. We detected phenoloxidase activity in the membrane fraction of hemocytes from the eastern oyster, Crassostrea virginica. The activity is associated with a protein with a molecular weight of 133 kDa. That the enzyme is a phenoloxidase and not a peroxidase was shown by inhibition of the activity by diethyldithiocarbamic acid. The phenoloxidase displayed a preference for diphenol substrates and was inactivated by incubation at temperatures above 70[degrees]C. The activity was highest in a pH range of 6.0-7.5. The oyster pathogen P, marinus causes serious disease in the commercial oyster, Crassostrea virginica, but not the mussel Geukensia demissa. We measured the effect of P. marinus cells on the phenoloxidase activity in hemolymph from C. virginica and G. demissa over a 6-h time course. The presence of P. marinus significantly suppressed the phenoloxidase activity of both species at the 2-h time interval. The phenoloxidase activity seemed to increase at the 4-hr and 6-h time intervals. These data suggest that a transient inhibition of host phenoloxidase may play a role in the study of P. marinus infection.

KEY WORDS: bivalves, phenoloxidase, internal defense, hemocytes, molluscs, biochemistry, Crassostrea virginica, Perkinsus, Geukensia demissa


Although the Gulf of Mexico leads all other regions of the United States in oyster production, there has been a steady decline in the total annual harvest (Suppan 2000). A major reason for the decline in oyster production in the United States has been the spread of Perkinsus marinus Levine. P. marinus, a parasite of the eastern oyster, Crassostrea virginica Gmelin, was first reported in the Gulf of Mexico (Mackin et al. 1950) but is now prevalent in C. virginica populations along the Atlantic coast from Maine to Florida and in the Gulf of Mexico from Florida to Mexico (Andrews & Hewatt 1957, Mackin 1962, Burreson et al. 1994). P. marinus is the causative agent of Dermo, a lethal disease of the eastern oyster (Andrews 1988). In the Gulf of Mexico, the market-size component of oyster populations generally suffers about 50% yearly mortality because of P. marinus (Mackin 1961, Mackin 1962, Hofstetter 1977). Typically, over 60% of the animals in nearly all oyster populations in the Gulf of Mexico are infected with P. marinus (Craig et al. 1989, Wilson et al. 1990). In the initial stages of infection, P. marinus cells are readily recognized and phagocytized by the hemocytes of the oyster; however, the hemocytes have little or no ability to destroy the ingested parasites, and may actually be instrumental in spreading the disease throughout the oyster (Perkins, 1976). As the infection progresses, hemocytes are lysed (Mackin 1951, Perkins 1976), releasing the pathogens.

Humoral factors such as reactive oxygen intermediates, lysozyme, hemagglutinins and phenoloxidase have been implicated as possible defense mechanisms in oysters (Anderson 1996) and P. marinus infections have been shown to affect the humoral and cellular host defense mechanisms of bivalve molluscs. For example, the chemiluminescent response of oyster hemocytes to zymosan or Escherichia coli, which corresponds to the amount of reactive oxygen intermediates produced by the hemocytes, is higher in hemocytes from infected oysters (Anderson et al. 1992, Anderson et al. 1995). However, exposure of hemocytes to P. marinus does not increase the chemiluminescent response (La Peyre 1993, La Peyre et al. 1995b, Volety & Chu 1995).

Phenoloxidase has been detected in the hemolymph of a wide variety of invertebrate and vertebrate species (Smith & Soderhall 1991). This enzyme converts phenolic substrates, such as L-dopa and 4-methylcatechol, to melanin. Phenoloxidase has been found in the hemocytes and hemolymph of various bivalve molluscs (Coles & Pipe 1994, Renwrantz et al. 1996, Asokan et al. 1997, Caraballal et al. 1997, Lopez et al. 1997, Deaton et al. 1999), but the role of phenoloxidase in bivalve host defense mechanisms has not yet been elucidated. We have examined the substrate specificity, effects of inhibitors, and the effects of pH and temperature on the activity of the phenoloxidase of the oyster C. virginica. In addition, the effect of P. marinus on the activity of phenoloxidase in the hemolymph of 2 bivalves, C. virginica and Geukensia demissa Sowerby was studied. The former species is susceptible to infection by P. marinus, whereas the latter is not.



Specimens of the eastern oyster, Crassostrea virginica, were purchased from Pearl Reef Oyster Company in Perry, Louisiana; specimens of the Atlantic ribbed mussel, Geukensia demissa, were collected from a salt marsh near Cocodrie, Louisiana. The specimens were maintained in recirculating, filtered seawater (15 [per thousand]) at room temperature (23[degrees]C). The animals were not fed and were used within 2 wk of collection.

Preparation of Hemocyte Extracts

A hole was punched through the shell of an oyster, and a syringe containing 2 mL of cold hemocyte antiaggregate solution (HAAS; 0.1 M phosphate buffer with 150 mg EDTA and 100 mg sodium chloride, pH 7.6) was inserted into the adductor muscle sinus. The hemolymph (usually 2 mL) was withdrawn and mixed with the HAAS by inversion of the syringe. Usually, the hemolymph of 3 or more oysters was pooled. The diluted hemolymph was centrifuged at x750g at 4[degrees]C for 10 min. The plasma was collected and placed on ice; the cellular pellet was resuspended in 1 mL HAAS, and centrifuged again. The supernatant was removed and discarded. Without resuspending the cellular pellet, 200 [micro]L of homogenization buffer (20 mM Tris, 5 mM Ca[Cl.sub.2], pH 6.5) were added to the tube. The mixture was homogenized on ice for 1 min. An additional 800 [micro]L of homogenization buffer was then added, and the homogenate was centrifuged at x16,000g for 20 min at 4[degrees]C. The supernatant (hemocyte lysate supernatant or HLS) was collected and placed on ice. The pellet (hemocyte membranes) was resuspended in 200 [micro]L of de-ionized water, vortexed, centrifuged at x16,000g for 10 min at 4[degrees]C, and the supernatant was discarded. This procedure was repeated twice, and the resulting membrane preparation was resuspended in 200 [micro]L of de-ionized water and placed on ice.

Detection of Phenoloxidase Activity on Polyacrylamide Gels

The method of Nellaiappan and Vinayagam (1986) was used, with some variations in the experimental technique, to detect phenoloxidase activity in polyacrylamide gels. The protein components of the plasma, HLS, and hemocyte membranes were separated on 7.5% sodium dodecylsulfide (SDS) polyacrylamide gels by electrophoresis (Bio-Rad Mini-PROTEAN II). After electrophoresis, the gels were washed 3 times for 10 min each in 50 mM phosphate buffer (pH 7.6). Next, the gel was incubated in a mixture consisting of a 4:1 (v/v) ratio of substrate solution (25 mg 4-methylcatechol dissolved in 50 mM phosphate buffer, pH 7.6) to 3-methyl-2-benzothiazolinone hydrazone hydrochloride (MBTH) (0.3%) dissolved in ethanol. The gel was allowed to develop for 1 h at room temperature and then washed for 10 min in de-ionized water three times. Finally, the gel was washed in a solution of 7% acetic acid and dried. The molecular weight of the band in the membrane fraction that displayed phenoloxidase activity was estimated by comparison with the migration of standard protein molecular weight markers (Life Technologies BenchMark prestained protein ladder).

To determine whether the enzyme activity was caused by a phenoloxidase or a peroxidase, oyster pheloxidase PAGE gels were assayed for activity in the presence of phenoloxidase inhibitors. The gels were incubated for 10 min in phosphate buffer containing either 5, 10 or 15 mM concentrations of diethylthiocarbamate (DETC). We also incubated gels in buffer containing (5-15 mM) phenylthiourea or tropolone.

To examine the substrate specificity of the enzyme, polyacrylamide gels were loaded with the hemocyte membrane preparation, subjected to electrophoresis, and then incubated in various monophenolic, diphenolic and triphenolic substrates for 60 min at room temperature. The substrates tested included the monophenols L-tyrosine, tyrosine methyl ester, and tyramine; the diphenols 4-methylcatechol, dopamine, 3,3'-diaminobenzidene (DAB), L-3,4-dihydroxyphenylalanine (L-DOPA) and hydroquinone; and the triphenol pyrogallol. The gels were then washed twice for 5 min each in phosphate buffer and then incubated in the 4-methylcatechol/MBTH reaction mixture.

The optimal pH range of enzymatic activity was determined using 3,3-dimethylglutaric acid (DMG); aliquots of 50 mM DMG were adjusted to pH ranging from 3.5-7.5 in increments of 0.5 pH units with HCl or NaOH. Gets were washed in de-ionized water for 5 min following electrophoresis. The gels were then preincubated for 10 min in buffers of the desired pH, and the buffer was discarded. Gels were then incubated for 30 min in the 10 mM 4-methylcatechol substrate solution (prepared in DMG buffer of the desired pH) and MBTH solution. Finally, the gels were rinsed, fixed and dried as previously described.

The effect of temperature on the functional integrity of the enzyme was studied. Prior to electrophoresis, aliquots of the hemocyte membrane preparation were incubated at temperatures of 20, 30, 40, 50, 60, 70, 80, 90 and 100[degrees]C for 10 min. Each aliquot was separated by SDS-PAGE; the gels were then incubated in the substrate 4-methyl-catechol and MBTH solutions and processed as above.

Sample Preparation--Hemolymph Incubation with P. marinus

Hemolymph was collected from the Atlantic ribbed mussel and the eastern oyster by prying the valves apart and inserting a syringe with a 22-gauge needle into the adductor muscle sinus. Adequate volumes of hemolymph could be obtained only from the larger animals, so in most cases, samples of hemolymph were obtained by pooling hemolymph from 2-3 individuals. Two-milliliter aliquots of each hemolymph sample were placed in each of two glass test tubes (13 x 100 mm), and 20 [micro]L of a culture of Perkinsus marinus (ATCC number 50,509 [DBNJ-1]; 1 x [10.sup.5] cells [mL.sup.-1]) were added to one of the test tubes. The hemolymph in the other test tube was mixed with 20 [micro]L of isosmotic seawater to serve as a control. The test tubes were incubated at room temperature, and at intervals of 0, 1, 2, 4 and 6 h, a 200-[micro]L sample of each test tube was assayed for phenoloxidase activity using a microplate spectrophotometric assay.

Spectrophotmetric Phenoloxidase Assay

Phenoloxidase activity in C. virginica and G. demissa hemolymph was measured using a colorimetric assay (Pye 1974) modified for 96 well microplates. The assay mixture consisted of 50[micro]L Tris chloride buffer (50 mM, pH 7.5), 100 [micro]L hydroxyproline-benzoyl ester (20 mM), and 10 [micro]L 4-methylcatechol (100 mM). A 200 [micro]L aliquot of hemolymph was added to the assay mixture and after 30 min of incubation at room temperature, the absorbance of the wells at 590 nm was read with a microplate reader (Tecan). Test tubes containing 2 mL seawater or seawater mixed with 20 uL of the P. marinus culture were also assayed to measure auto-oxidation of the 4-methyl-catechol substrate. In all cases, the enzyme-catalyzed color production was considerably higher than that because of auto-oxidation of the substrates. The data were reported as the difference in absorbance between paired samples of hemolymph and hemolymph mixed with P. marinus. The protein content of the hemolymph was measured using Bradford reagent with bovine serum albumin as a standard. Student t-test was used to evaluate differences between the means of enzyme activity at the various time points.


Phenoloxidase activity is present in the hemocyte membranes; the activity corresponds to a protein band with a molecular weight of 133 kD (Fig. 1A). We also detected a less intense band of activity corresponding to a molecular weight of about 250 kD. The phenoloxidase activity in the plasma is below the detection limits of the gel assay we used. Only the 135 kD protein band from hemocyte membrane preparations was assayed in the remainder of our gel experiments.

To determine the effectiveness of tropolone, PTU and DETC as specific inhibitors, we did preliminary experiments using PAGE gels loaded with mushroom tyrosinase or horseradish peroxidase (Sigma Chemical Co.) and assayed with the same substrate solution we used for the oyster enzyme. The mushroom tyrosinase (a phenoloxidase) separated into four bands of activity on the PAGE gels. DETC abolished the activity of all four bands but PTU inhibited only two of them. Neither PTU nor DETC inhibited the activity of horseradish peroxidase. DETC completely abolishes the activity of the oyster enzyme (Fig. 1B). PTU depresses, but does not completely inhibit the activity of phenoloxidase in the hemocyte membranes (data not shown).


Tropolone inhibits both the monophenolase and dihydroxyphenolase activity of mushroom tyrosinase (Kahn and Adrawis 1985a), but in the presence of hydrogen peroxide, it can serve as a substrate for horseradish peroxidase (Kahn & Andrawis 1985b). We tested tropolone on our mushroom tyrosinase PAGE gels and found it to be only partially effective in blocking the oxidation of 4-methylcatechol. Tropolone is a more effective inhibitor of mushroom tyrosinase when L-DOPA or dopamine instead of 4-methylcatechol is the substrate (Kahn & Andrawis 1985a). Tropolone had no effect on mushroom tyrosinase or C. virginica phenoloxidase.

The phenoloxidase is active on all of the diphenolic substrates but is not active with monophenol or triphenol substrates (Table 1). The phenoloxidase activity was detected on the gels only within a pH range of 6.0-7.5 (Fig. 1C). The activity was abolished by incubation of the protein at temperatures [greater than or equal to] 70[degrees]C prior to electrophoresis, incubation at lower temperatures had no effect on the activity (Fig. 1D).

The microplate assay detected phenoloxidase in the whole hemolymph of C. virginica and G. demissa. The variability of phenoloxidase activity among individuals of the same species is large, but a typical hemolymph sample had an activity of 50-200 x [10.sup.-3] absorbance units/min/mg protein. There were no consistent differences in the phenoloxidase activity of the hemolymph from the two species.

The effect of P. marinus on the phenoloxidase activity in hemolymph from C. virginica and G. demissa is shown in Figures 2 and 3, respectively. In both species, phenoloxidase activity is significantly decreased at the 2 h time interval after exposure to P. marinus. The phenoloxidase activity at the 1 and 6 h time intervals is not different from that at the beginning of the experiment in either species.



The membrane fraction of oyster hemocytes contains at least two proteins that display phenoloxidase activity. Phenoloxidase has been detected in the hemocytes of the marine mussels Mytilus galloprovincialis Lamarck, Mytilus edulis L. Perna viridis L., and the clam Ruditapes decussates Chiamenti (Coles & Pipe 1994, Renwrantz et al. 1996, Asokan et al. 1997, Caraballal et al. 1997, Lopez et al. 1997). The phenoloxidase activity that we measured in the hemolymph of G. demissa and C. virginica in this study is comparable to activities found in the blood of other species of marine molluscs measured with the same assay system (Deaton et al. 1999).

The molecular weight of phenoloxidase from a variety of molluscs has been reported. Two phenoloxidase proteins, of 381 kD and 316 kD, were isolated from the hemocyte lysate supernatant (HLS) of Mytilus edulis (Renwrantz et al. 1996). A 70 kD phenoloxidase was isolated from the periostracum of Modiolus demissus (= Geukensia demissa) (Waite & Wilbur 1976). Phenoloxidase in the foot gland of Mytilus edulis was reported to have a molecular mass of 260 kD (Maruyama et al. 1991). Phenoloxidase detected in the ejected ink of cephalopods has molecular masses of 205 kD, 125 kD and 135 kD for Octopus vulgaris Cuvier, Sepia officinalis L. and Loligo vulgaris Lamarck, respectively (Prota et al. 1981). These proteins are about the same size as the bands of activity we detected in C. virginica hemocytes. The molecular weights of prophenoloxidase from arthropods range from 71-80 kD (Soderhall et al. 1994). For example, the deduced amino acid sequence of a prophenoloxidase cloned from cDNA from the crayfish Pacifastacus leniusculus represents a protein of 80 kD (Aspan et al. 1995). In summary, the phenoloxidases of most molluscs are larger proteins than those typical of arthropods.

Phenoloxidase activity has been associated with cell membranes in several species. In the insect Ceratitis capitata Wiedemann, prophenoloxidase has been detected on the hemocyte surface (Charalambidis et al. 1996). Membrane bound phenoloxidase activity has been reported in the trematode Fasciola gigantica Cobbold and implicated in defense mechanisms (Nellaiappan et al. 1989). Phenoloxidase has also been reported to be associated with the cell membranes of plants (Escribano et al. 1997a and Escribano et al. 1997b). Our results suggest that most of the phenoloxidase activity in molluscan hemocytes is associated with the plasma membrane.

Phenoloxidases and perioxidases oxidize some of the same phenolic substrates including catecholamine; inhibitors can be used to differentiate the two enzymes. The results we obtained with tropolone, PTU, and DETC are consistent with the conclusion that the enzyme from C. virginica hemocytes is a phenoloxidase and not a peroxidase.

The oyster enzyme shows a strong specificity for o-diphenol substrates, such as 4-methylcatechol, dopamine, and L-DOPA. Phenoloxidases with this characteristic are commonly referred to as having catecholase activity. The rate of development of colored product on the gels suggests that the oyster phenoloxidase most readily oxidized 4-methylcatechol, followed by dopamine and L-DOPA. This is similar to the substrate preferences exhibited by the phenoloxidase of F. gigantica, which most effectively oxidizes 4-methylcatechol, followed by catechol and dopamine (Nellaiappan et al. 1989). The p-diphenol substrate (laccase activity), hydroquinone, is slowly oxidized (about one hour to see color in gel) by the enzyme from C. virginica. The enzyme from F. gigantica also has low activity with hydroquinone as a substrate (Nellaiappan et al, 1989). Furthermore, the membrane bound enzyme from the eastern oyster is unable to oxidize most monophenols (cresolase activity) (Table 1). If the incubation time was extended beyond one hour, there was some activity with the monophenol tyrosine methyl ester as substrate. In plants, cresolase activity may be diminished or abolished during enzyme preparation. In addition, prolonged incubations can result in the auto-oxidation of the substrate and condensation of the resulting quinone with tannable proteins to give a brown coloration, thereby producing a false positive pseudo-enzyme reaction (Nellaiappan & Vinayakam 1986). The loss of enzyme activity after incubation of the oyster hemocyte proteins at high temperatures suggests that the color development we observed in our gels is enzymatic and not because of auto-oxidation of the substrate. Monophenols are oxidized at moderate rates by the membrane bound phenoloxidase in F. gigantica (Nellaiappan et al. 1989). In conclusion, the hemocyte phenoloxidase of C. virginica displays strong catecholase, and possibly some laccase, activity.

The phenoloxidase from C. virginica has maximal activity in the physiologic pH range. The optimum pH of phenoloxidase from M. edulis is 7.0 (Renwrantz et al. 1996). A pH optimum of 6.8 was reported for both HLS and membrane bound phenoloxidase from F. gigantica (Nellaiappan et al. 1989). The pH optimum of phenoloxidase from the periostracum of M. demissus is 8.0-8.5 (Waite & Wilbur 1976). This latter enzyme may be different from that found in the hemocytes. Oysters that are highly infected with P. marinus undergo an acidosis during emersion that is larger than that of uninfected animals (Dwyer & Burnett 1996). Whereas it is possible that the effect of P. marinus infection on acid-base balance interferes with the defensive mechanisms of the oysters, phenoloxidase would still be functional at the hemolymph pH values reported by Dwyer and Burnett (1996).

To infect an oyster, P. marinus cells must overcome the defensive mechanisms of prospective host. The plasma from some bivalves has been shown to decrease the proliferation of P. marinus in culture (Gauthier & Vasta 2002). Bivalve hemolymph contains defensive molecules such as lysosomal enzymes and antimicrobial peptides, and the hemocytes produce reactive oxygen intermediates (Anderson 1996, Mitta et al. 1999). Whereas the etiology of Dermo infection is not clearly understood, it may include entry by an overwhelming number of P. marinus cells, rapid dissemination throughout the host via the circulation, a high rate of pathogen proliferation in host cells, and depression of the host defense response (La Peyre 1996, Anderson 1999). P. marinus cells produce a number of products that decrease the motility, lysozyme activity, and hemagglutinin titers of oyster hemocytes (Garreis 1996). For example, P. marinus cells secrete multiple serine proteases (La Peyre et al. 1995a) that may inactivate the defensive mechanisms of the oyster (Garreis 1996). Products secreted by P. marinus may be involved in the decreases in the phenoloxidase activity of both C. virginica and G. demissa hemolymph that we observed after 2 h of incubation. Protease inhibitors are present in C. virginica plasma, and concentrations vary among individuals, suggesting that these molecules may be a defensive mechanism against proteases secreted by pathogens (Faisal et al. 1998). In arthropods, inactive prophenoloxidase is activated by plasma proteases (Aspan et al. 1995). If the activation mechanism in bivalves is similar, the release of protease inhibitors by oyster hemocytes in response to proteases produced by P. marinus ceils may play a role in the suppression of phenoloxidase activity in the oyster hemolymph. Whatever the mechanism, transient inhibition of phenoloxidase and possibly other host defense mechanisms may provide time for spread of pathogen cells throughout a newly infected host animal.

In summary, the biochemical characteristics of phenoloxidase from oyster hemocytes is largely similar to that of other molluscs and invertebrates. Our results suggest that P. marinus affects phenoloxidase activity in the eastern oyster, C. virginica, and the Atlantic ribbed mussel, G. demissa. Further studies are needed to determine the role of phenoloxidase in host defense mechanisms in bivalve molluscs and the significance of P. marinus suppression of phenoloxidase in the establishment and progression of the infection in susceptible species.


This work was supported by a University of Louisiana at Lafayette Doctoral Fellowship (PC J). This is contribution number 326 from the Tallahassee, Sopchoppy and Gulf Coast Marine Biological Association.


Anderson, R. 1996. Interactions of Perkinsus marinus with humoral factors and hemocytes of Crassostrea virginica. J. Shellfish Res. 15:127-134.

Anderson, R. 1999. Perkinsus marinus secretory products modulate superoxide anion production by oyster (Crassostrea virginica) hemocytes. Fish Shellfish Immunol. 9:51-60.

Anderson, R., K. Paynter & E. Burreson. 1992. Increased reactive oxygen intermediate production by hemocytes withdrawn from Crassostrea virginica infected with Perkinsus marinus. Biol Bull. 183:476-481.

Anderson, R., E. Burreson & K. Paynter. 1995. Defense responses of hemocytes withdrawn from Crassostrea virginica infected with Perkinsus marinus. J. Invert. Pathol. 66:82-89.

Andrews, J. 1988. Epizootiology of the disease caused by the oyster pathogen Perkinsus marinus and its effect on the oyster industry. Am. Fish. Soc. Spec. Publ. 18:47-63.

Andrews, J. & W. Hewatt. 1957. Oyster mortality studies in Virginia, II. The fungus disease caused by Dermocystidium marinum on oysters of Chesapeake Bay. Ecol. Monogr. 27:1-25.

Asokan, R., M. Arumugam & P. Mullainadhan. 1997. Activation of prophenol-oxidase in the plasma and haemocytes of the marine mussel Perna viridis Linnaeus. Dev. Comp. Immunol. 21:1-12.

Aspan, A., T. Huang, L. Cerenius & K. Soderhall. 1995. cDNA cloning of prophenoloxidase from the freshwater crayfish Pacifastacus leniusculus and its activation. Proc. Natl. Acad. Sci. USA 92:939-943.

Burreson, E., R. Alvarez, V. Martinez & L. Macedo. 1994. Perkinsus marinus (Apicomplexa) as a potential source of oyster Crassostrea virginica mortality in coastal lagoons of Tabasco, Mexico. Dis. Aquat. Org. 20:77-82.

Caraballal, M. J., C. Lopez, C. Azevedo & A. Villalba. 1997. Enzymes involved in defense functions of hemocytes of mussel Mytilus galloprovincialis. J. Invert. Pathol. 70:96-105.

Charalambidis, N., L. Foukas, C. Zervas & V. Marmaras. 1996. Hemocyte surface phenoloxidase (PO) and immune response to lipopolysaccharide (LPS) in Ceratitis capitata. Insect Biochem. Mol. Biol. 26:867-874.

Coles, J. & R. K. Pipe. 1994. Phenoloxidase activity in the haemolymph and haemocytes of the marine mussel Mytilus edulis. Fish Shellfish Immunol. 4:337-352.

Craig, A, E. Powell, R. Fay & J. Brooks. 1989. Distribution of Perkinsus marinus in Gulf coast oyster populations. Estuaries 12:82-91.

Deaton, L., P. Jordan & J. Dankert. 1999. Phenoloxidase activity in the hemolymph of bivalve molluscs. J. Shellfish Res. 18:223-226.

Dwyer, J. and L. Burnett. 1996. Acid-base status of the oyster Crassostrea virginica in response to air exposure and to infections by Perkinsus marinus. Biol. Bull 190:139-147.

Escribano, J., J. Cabanes, S. Chazarra & F. Garcia-Carmona. 1997a. Characterization of monophenoloxidase activity of table beet polyphenol oxidase. Determination of kinetic parameters on the tyramine/ dopamine pair. Journal Agriculture and Food Chemistry 45:4209-4214.

Escribano, J., J. Cabanes & F. Garcia-Carmona. 1997b. Characterization of latent polyphenol oxidase in table beet: Effect of sodium dodecyl sulphate. Journal of the Science of Food and Agriculture 73:34-38.

Faisal, M., E. McIntyre, K. Adham, B. Tall, M. Kothary & J. La Peyre. 1998. Evidence for the presence of protease inhibitors in eastern (Crassostrea virginica) and Pacific (Crassostrea gigas) oysters. Comp. Biochem. Physiol. 121B:161-168.

Garreis, K., J. La Peyre & M. Faisal. 1996. The effects of Perkinsus marinus extracellular proteases on oyster defense parameters in vitro. Fish Shellfish Immunol. 6:581-597.

Hofstetter, R. 1990. The Texas oyster fishery. Texas Parks Wildlife Department Bulletin 40. pp. 1-21.

Kahn, V. & A. Andrawis. 1985a. Inhibition of mushroom tyrosinase by tropolone. Phytochemistry 24:905-908.

Kahn, V. & A. Andrawis. 1985b. Tropolone as a substrate for horseradish peroxidase. Phytochemistry 24:909-913.

La Peyre, J. 1996. Propagation and in vitro studies of Perkinsus marinus. J. Shellfish Res. 15:89-101.

La Peyre, J., D. Schafhauser, E. Rizkalla & M. Faisal. 1995a. Production of serine proteases by the oyster pathogen Perkinsus marinus (Apicomplexa) in vitro. J. Euk. Microbiol. 40:304-310.

La Peyre, J., F. Chu & W. Vogelbein. 1995b. In vitro interaction of Perkinsus marinus merozoites with eastern and Pacific oyster hemocytes. Dev. Comp. Immunol. 19:291-304.

Leippe, M. & L. Renwrantz. 1988. Release of cytotoxic and agglutination molecules by Mytilus hemocytes. Dev. Comp. Immunol. 12:297-308.

Lopez, C., M.J. Caraballal, C. Azevedo & A. Villalba. 1997. Enzyme characterization of the circulating haemocytes of the carpet shell clam, Ruditapes decussates (Mollusca:bivalvia). Fish Shellfish Immunol. 7: 595-608.

Mackin, J. 1951. Histopathology of infection of Crassostrea virginica (Gmelin) by Dermocystidium marinum Mackin, Owen and Collier. Bull Mar. Sci. Gulf Caribbean 1:72-87.

Mackin, J. 1962. Oyster disease caused by Dermocystidium marinum and other microorganisms in Louisiana. Texas Inst. Mar. Sci. Publication 7:132-299.

Mackin, J., H. Owen & A. Collier. 1950. Preliminary note on the occurrence of a new protozoan parasite, Dermocystidium marinum n. sp. In: Crassostrea virginica (Gmelin). Science 111:328-329.

Maruyama, N., H. Etoh, K. Sakata & K. Ina. 1991. Studies on phenoloxidase from Mytilus edulis associated with adhesion. Agric Biol. Chem. 55:2887-2889.

Mitta, G., F. Vandenbulcke, R. Hubert & P. Roch. 1999. Mussel defensins are synthesized and processed in granulocytes then released into the plasma after bacterial challenge. J. Cell Sci. 112:4233-4242.

Nellaiappan, K., A. Devasundari & S. Dhandayuthapanl. 1989. Properties of phenol oxidase in Fasicola gigantica. Parasitology 99:403-407.

Nellaiappan, K. & A. Vinayagam. 1986. A rapid method for detection of tyrosinase activity in electrophoresis. Stain Technology 61:269-272.

Perkins, F. 1976. Dermocystidium marinum infections in oysters. Mar. Fish. Rev. 38:19-21.

Prota, G., J. Ortonne, C. Voulot, C. Khatchadourian, G. Nardi & A. Palumbo. 1981. Occurrence and properties of tyrosinase in ejected ink of cephalopods. Comp. Biochem. Physiol. 68B:415-419.

Pye, A. 1974. Microbial activation of prophenoloxidase from immune insect larvae. Nature 251:610-613.

Renwrantz, L., W. Schmalmack, R. Redel, B. Friebel & Schneeweiss. 1996. Conversion of phenoloxidase and peroxidase indicators in individual haemocytes of Mytilus edulis specimens and isolation of phenoloxidase from haemocyte extract. Comp. Physiol. B 165:647-658.

Smith, V. J. & K, Soderhall. 1991. A comparison of phenoloxidase activity in the blood of marine invertebrates. Dev. Comp. Immunol. 15:251-261.

Soderhall, K., L. Cerenius & M. W. Johansson, 1994. The prophenoloxidase activating system and its role in invertebrate defense. Annals of the New York Academy of Sciences 712:155-161.

Suppan, J. 2000. The Gulf coast oyster industry program: an initiative to address industry's research needs; J. Shellfish. Res. 19:397-400,

Volety, A. & F. Chu. 1995. Suppression of chemiluminescence of eastern oyster (Crassostrea virginica) hemocytes by the protozoan parasite Perkinsus marinus. Dev. Comp. Immunol. 19:135-142.

Waite, J. H. & K. M. Wilbur. 1976. Phenoloxidase in the periostracum of the marine bivalve Modiolus demissus Dillwyn. J. Exp. Zool. 195:359-368.

Wilson, E., E. Powell, M. Craig, T. Wade & J. Brooks. 1990. The distribution of Perkinsus marinus in the Gulf coast oyster: its relationship with temperature, reproduction, and pollutant body burden. Int. Rev. Gesamten Hydrobiol. 75:533-550.


Biology Department University of Southwestern Louisiana at Lafayette Lafayette, Louisiana 70504

Corresponding author. Email:
Substrate specificity of phenoloxidase from
Crassostrea virginica hemocytes.

Substrate Type Enzyme Activity

L-tyrosine monophenol -
L-tyrosine methyl ester monophenol -
Tyramine monophenol -
4-methylcatechol o-diphenol +
L-DOPA o-diphenol +
Dopamine o-diphenol +
DAB p-diphenol +
Hydroquinone p-diphenol +
Pyrogallol triphenol -

L-DOPA, L-3,4-dihydroxyphenylalanine, DAB, 3,3'-diaminobenzidene
COPYRIGHT 2005 National Shellfisheries Association, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2005, Gale Group. All rights reserved. Gale Group is a Thomson Corporation Company.

Article Details
Printer friendly Cite/link Email Feedback
Author:Deaton, Lewis E.
Publication:Journal of Shellfish Research
Geographic Code:1USA
Date:Aug 1, 2005
Previous Article:Fisherman choice and incidental catch: size frequency of oyster landings in the New Jersey oyster fishery.
Next Article:A comparison of the effectiveness of sandstone and limestone as cultch for oysters, Crassostrea virginica.

Related Articles
Microscopic observations of larval Ostrea circumpicta (bivalve: ostreidae) in brood chambers.
Reproductive cycle and mortality of the Japanese oyster Crassostrea gigas cultured in Bahia Falsa, Baja California, Mexico.
Parasitic and symbiotic fauna in oysters (Crassostrea virginica) collected from the Caloosahatchee River and Estuary in Florida.
Effects of polynuclear aromatic hydrocarbons on hemocyte characteristics of the Pacific oyster, Crassostrea gigas.
Immunomodulation of Crassostrea gigas and Crassostrea virginica cellular defense mechanisms by Perkinsus marinus.
Morphological, structural and functional characteristics of the hemocytes of the oyster, Crassostrea ariakensis.
Diseases of pearl oysters and other molluscs: a Western Australian perspective.
Effect of homogenate from different oyster species on Perkinsus marinus proliferation and subtilisin gene transcription.
The limits of morphometric features for the identification of black-lip pearl oyster (Pinctada margaritifera) larvae.
Adaptation of ray's fluid thioglycollate medium assay to detect and quantify planktonic stages of Perkinsus spp. parasites.

Terms of use | Privacy policy | Copyright © 2021 Farlex, Inc. | Feedback | For webmasters |