Comparative Biochemistry and Physiology, Part A 145 (2006) 114 – 122 www.elsevier.com/locate/cbpa
The salivary gland and salivary enzymes of the giant waterbugs (Heteroptera; Belostomatidae) C.C. Swart a,⁎, L.E. Deaton b , B.E. Felgenhauer b a b
Biology/Neuroscience, Trinity College, 300 Summit Street, Hartford, CT 06106, USA Department of Biology, University of Louisiana at Lafayette, Lafayette, LA 70504, USA
Received 21 November 2005; received in revised form 2 May 2006; accepted 20 May 2006 Available online 26 May 2006
Abstract The giant waterbugs are predators that utilize extra-oral digestion and are known to capture a wide variety of prey. Herein we describe the differences in salivary enzyme composition between large and small species of giant waterbug (Lethocerus uhleri, Lethocerinae and Belostoma lutarium, Belostomatinae, respectively). The saliva of L. uhleri contains 3 proteolytic enzymes and no amylase, while the salivary gland of B. lutarium produces 2 proteolytic enzymes and amylase. This fundamental difference in salivary enzyme composition correlates with the difference in diet preference between the Lethocerinae and Belostomatinae. Furthermore, we describe the ultrastructure of the salivary gland complex of B. lutarium and present data on the division of labor with respect to compartmentalization of enzyme production. Proteolytic enzymes are produced in the accessory salivary gland and amylase is produced in the main salivary gland lobe. This is the first reported evidence of protease production in the accessory salivary gland in the Heteroptera. © 2006 Elsevier Inc. All rights reserved. Keywords: Belostoma; Lethocerus; Extra-oral digestion; Proteases; Accessory salivary gland; Digestive physiology; Amylase; Prey choice
1. Introduction The True Bugs or Heteroptera all practice extra-oral digestion and may be predaceous, phytophagous, granivorous, ectoparasitic, or combinations of several modes (Cohen, 1995; Gillespie and McGregor, 2000; Boyd et al., 2002; Boyd, 2003). Various Heteroptera are considered to be beneficial (predators), or pests (herbivores), or merely anomalous (zoophytophagous) depending mainly on their feeding habits (Schaefer and Panizzi, 2000). Inherent in all of this research of Heteropteran digestive physiology is the idea that the diversity of the feeding habits includes not only a diversity of morphology and food sources, but also a bewildering diversity of digestive enzyme production. Significant differences in the composition and physiological mode of action of terrestrial heteropteran saliva has been described (Cohen, 1990, 1993, 1995; Terra and Ferreira, 1994; Boyd et al., 2002; Boyd, 2003). Based on the observation of
⁎ Corresponding author. E-mail address:
[email protected] (C.C. Swart). 1095-6433/$ - see front matter © 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpa.2006.05.010
mixed diets Boyd and colleagues (Boyd et al., 2002; Boyd, 2003) tested for the presence of carbohydrases versus proteases and feeding bristle complexity in the Mirid bugs Deraeocoris nebulosus and Deraeocoris nigritulus. Their results indicate that morphology and physiology of the digestive system is predictive of prey choice. Other work indicates that plants may be exploited mainly as a source of moisture and not primarily for nutrition (Gillespie and McGregor, 2000). There is even evidence that some omnivorous terrestrial Heteroptera can alter the content of their saliva based on immediate diet (Habibi et al., 2002). The semi-aquatic Heteroptera are considered exclusively predaceous; however, to date we are aware of only a single study of the salivary enzymes of an aquatic Heteropteran (Rees and Offord, 1969). The Belostomatidae, known collectively as the giant waterbugs, are potentially important determinants of the ecological community structure due to their broad range of prey choices and frequent high population density (Mori and Ohba, 2004; Swart and Taylor, 2004). This broad range of prey utilization indicates a very robust digestive system capable of handling numerous prey types, exploiting diverse nutrient molecules and
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employing complex search behaviors. Because all Heteroptera practice extra-oral digestion, this robustness must be mediated by the suite of enzymes present in the salivary product and the function of the ingestive organs (Cohen, 1995). Belostomatidae is divided into two sub-families; the large bodied, egg-guarding Lethocerinae and the small bodied eggbrooding Belostomatinae. Members of the Lethocerinae have been reported to feed on prey as large as small birds, snakes, and adult anurans, but more commonly feed on small fish, aquatic and terrestrial invertebrates, and anuran larvae (Torre Bueno, 1906; Babbitt and Jordan, 1996; Hirai and Hidaka, 2002; Mori and Ohba, 2004; Ohba and Nakasuji, 2006). The Belostomatinae exploit a wide range of prey types including anuran larvae, snails, and terrestrial and aquatic invertebrates (Torre Bueno, 1906; Cullen, 1969; Gonsoulin, 1973; Chase, 1999; Mori and Ohba, 2004; Swart and Taylor, 2004; Ohba and Nakasuji, 2006). Fish are notably absent from the diets of these smaller belostomatids. Both groups commonly co-occur in the same habitats and both groups exhibit a strong tendency toward cannibalism or intra-familial predation. Herein, we present data on the salivary enzymes of B. lutarium and L. uhleri and ultrastructure of the salivary gland of B. lutarium. We verify the presence of myoid cells in the alveolar components of the salivary gland and describe relevant ultrastructural detail. We test previous explanations of the division of labor of the four components of the salivary gland. We present qualitative data indicating the presence of up to 9 enzymes in the saliva of B. lutarium and 11 enzymes in L. uhleri. We also present data on the temperature and pH response curves of the proteases and amylase of B. lutarium and L. uhleri.
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Potassium Dichromate. Secondary fixative was rinsed out with 2 changes of 0.1 M Sodium Cacodylate buffer followed by an overnight rinse in running tap water. Specimens were dehydrated through a series of increasing strengths of ethanol to 100%, transferred to 100% acetone and subsequently infiltrated with a graded series of Spurr's resin. To prevent digestion of membrane components by the native salivary proteases during fixation a 0.01 M concentration of the protease inhibitor Paramethyl sulfonyl fluoride was added to the dissection fluid, Trumps fixative, and sodium cacodylate rinses. Glands in 100% Spurrs were placed in molds and heated at 70 °C for 16 h to polymerize the Spurr's resin (Spurr, 1969). Thin sections were produced using a Sorvall MT-3 ultramicrotome and represented all areas of the gland including main, lateral, and accessory lobes, and the storage sacs. Sections were placed on copper grids, stained with 5% uranyl acetate and counterstained with Reynolds Lead Citrate. Grids with stained sections were viewed and photographed in a Hitachi 6000 Transmission Electron Microscope.
2. Materials and methods
2.2.2. Fluorescence microscopy Glands were dissected as above then fixed for 30 min in 0.1 M phosphate buffer (pH = 7.0) containing 2% paraformaldehyde at 4 °C and 100 μL Triton-X100. Specimens were then rinsed 3 times in cold 0.1 M phosphate buffered saline (300 mOsmol). The whole gland was stained at room temperature in 1 μg/mL FITC-Phalloidin (Bio-Rad Inc.) in PBS for 30 min in the dark. The staining solution was rinsed out with 3 changes of PBS. Specimens were placed on slides with para-phenylene diamine as an anit-fade agent and coverslipped. Specimens were illuminated with a Mercury vapor lamp and viewed with a Nikon Y-FL fluorescent microscope. Images were captured from a SONY DKC-ST5 digital video camera.
2.1. Animals
2.3. Physiology
Lethocerus uhleri Montadon (Belostomatidae; Lethocerinae) and Belostoma lutarium Stål (Belostomatidae; Belostomatinae) are found in the same habitats and are common throughout southern Louisiana (Gonsoulin, 1973; Henry and Froeschner, 1988). Animals were collected from three sites in south central Louisiana as described in Swart and Taylor (2004) and Swart and Felgenhauer (2003). Representative samples of B. lutarium and L. uhleri are deposited at the Louisiana State Arthropod Museum, Baton Rouge, LA. Species were identified using the key characters of Gonsoulin (1973).
2.3.1. Qualitative analysis of enzymes Gland extracts from B. lutarium and L. uhleri were assayed for a variety of enzymes using a qualitative, colorometric enzyme assay system (apiZYM, bioMérieux Inc.). Glands from 10 B. lutarium were combined and ground in 1 mL of 150 mM Tris–HCl (pH 7.4.) Two individuals of L. uhleri were used to prepare a 1 mL solution. Both samples were centrifuged for 2 min at 10,000×g to remove cellular debris. The apiZYM assay (BioMérieux, Lyon, France) consists of a test strip composed of 20 cupules. Each cupule contains a synthetic substrate specific for a particular enzyme. The test can detect 19 enzymes, with the final cupule used as a negative control. First, 50 μL of the salivary gland extract was added to each cupule of the test strip. After inoculation with the gland extract, the test strip was incubated in the dark at 37 °C for 4 h. Then, one drop of reagent A (Tris-hydroxymethyl-aminomethane, HCl, sodium lauryl sulfate, and water) and one drop of reagent B (Fast Blue BB, 2-methoxyethanol) was added to each cupule. After addition of the two reagents, the test strip was incubated at room temperature for 5 min to allow staining of the product of substrate digestion. Then the test strip was placed under a
2.2. Morphology 2.2.1. TEM of salivary gland Animals were chilled in ice water, decapitated, and dissected under a dissecting microscope in 150 mM Tris–HCl (pH = 7.3). Whole glands including the main, lateral, and accessory lobes plus the storage sacs were placed in chilled Trump's Fixative (EMS inc.) and stored at 4 °C overnight. After two 30 min rinses in 0.1 M Sodium Cacodylate buffer (pH = 7.3) specimens were secondarily fixed for 1 h in 1% Osmium Textroxide with 0.1%
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1000 W incandescent light bulb for 10 min to eliminate any unbound dye (Fast Blue BB). Finally, the test strip was read by comparing the color produced in each cupule to a standard color chart. Color development was graded on a qualitative 0 to 5+ scale.
2.3.2. Protease activity Homogenates of salivary glands produced many non-proteolytic protein bands on electrophoretic gels due to tissue contamination and prevented accurate determination of salivary protein loads in both electrophoretic gels and spectrophotometric
Fig. 1. TEM and fluorescence imaging of salivary gland complex of B. lutarium. (A) The junction of two adjacent alveoli in the main gland lobe of B. lutarium. The two alveoli are both connected to a myoepithelial cell indicated by the black arrow. The upper cell is filled with a flocculent proteinaceous material, presumably salivary enzymes (f) while the lower cell lumen (l) is empty. (B) A myoepithelial cell (mec) adhering to an alveoli of the lateral gland lobe. Note the active Golgi complex in the gland cell (g). (C) The cuticular material (c) in the center of the image lines the luminal side (l) of the accessory salivary gland. (D) A large cell type lining the tubular accessory salivary gland is shown with active heterochromatin and endoplasmic reticulum (er). (E) The accessory salivary lobe when stained with FITC-Phalloidin stain for F-actin shows dispersed staining with no concentrated pattern. (F) When alveoli of the main or accessory gland lobes or the storage sacs are stained for F-actin with FITC-Phalloidin, the myoepithelial cells demonstrate strong affinity.
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assays. Therefore, pure saliva was used when possible. Saliva was collected by injecting bugs with 0.1 mL of 2% pilocarpine (a muscarinic agonist) in 300 mOsm NaCl (Miles, 1972). Within a few minutes, the pilocarpine induces the bugs to eject saliva through their proboscis allowing collection with a capillary tube. Pure saliva was ejected from the capillary tube into microcentrifuge tubes on ice containing 150 mM Tris–HCl (pH 7.5). Specimens were then frozen until used. A Bradfords assay modified for use in a microplate reader was used to quantify the amount of protein present in each tube of saliva to allow equal electrophoretic gel loadings when necessary (Bradford, 1976). Molecular weight of the protease enzymes was determined by incorporating casein into polyacrylamide gels modified from Heussen and Dowdle (1980). In an attempt to identify the action of various protease inhibitors on each protease saliva samples were treated with protease inhibitors prior to electrophoresis. Several lanes were loaded on each gel and after electrophoresis lanes were separated with a razor blade, washed in 2.5% Triton X100 for 30–60 min to remove SDS, then incubated at 37 °C for 3–5 h in 0.15 M Tris–HCl at a pH of 7.5. After incubation the reaction was stopped and gels stained in Coomassie blue. Stained gels were destained if necessary and photographed with a digital camera. Molecular weight was determined in all experiments by comparison to a molecular weight standard (Bio-Rad 161-0318). Digestion of azocasein and azoalbumin was tested by incubating coarse ground gland extract from B. lutarium and L. uhleri with a known quantity of protein content (Bradfords assay) in 0.5 mL of a 2 mg/mL solution of the substrate in Tris– Ringer buffer (150 mM Tris–HCl, 120 mM NaCl, 5 mM KCl) for 5–6 h. Three temperature treatments were used, 20 °C, 30 °C, and 37 °C and 5 pH treatments ranging from 7.0 to 9.0 in increments of 0.5 pH units. The reactions were stopped with the addition of 100 μL of 80% trichloro-acetic acid. Absorbance of the supernatant (200 μL) from experimental treatments and a negative control was read at 340 nm in a microplate photometer (Tecan). The action of two protease inhibitors on axocasein and azoalbumin digestion were tested; TLCK (Nα-tosyl-L-lysine chloromethyl ketone, Sigma 90182; applied at 50 μg/mL in 150 mM Tris–HCl, pH 6.0), and TPCK (N-p-tosyl-L-lysine ethyl ester–HCl, Sigma T4376, used at 50 μg/mL in 150 mM Tris–HCl, pH 7.5). 2.3.3. Amylase activity A microplate spectrophotometric technique for determination of α-amylase was adapted from Rinderknecht et al. (1967) using potato starch covalently linked to Remazol Brilliant Blue R (starch azure, Sigma S7629). Five microliters of saliva sample from B. lutarium and L. uhleri were incubated with 200 μL of 2% RBB-Starch in 0.02 M sodium phosphate buffer. Three temperature and 5 pH levels were studied as described in the protease spectrophotometric experiment above. 2.3.4. Division of labor of salivary gland For recovery of whole salivary glands animals were chilled in ice water, decapitated, and dissected in a Petri dish of chilled 150 mM Tris–HCl (pH 7.5) under a dissecting microscope.
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Whole glands were divided into main, lateral, storage sacs, and accessory lobes with microdissecting scissors and placed in microcentrifuge tubes containing 250 μL of 150 mM–Tris HCl at pH 7.5 on ice. An initial qualitative assay was used to determine activity of the protease and amylase enzymes. For proteases 0.5% agar gels containing 3% casein were produced as described in Cohen (1990). A 200 μL plastic pipette tip was cut with a razor blade to produce a lumen of approximately 1 mm. This tool was then used to cut 1 mm wells in the gel for injection of 10 μL salivary gland homogenate samples. After overnight incubation, gels were flooded with 3% acetic acid which turned the gel milk white in areas where casein had not been digested and clear in areas where active proteases were found. Positive (alpha-chymotrypsin, Sigma C7762) and negative (di H20) controls were performed. To determine the molecular weights of the proteases and amylase like enzymes 7.5 μL of each saliva sample (pilocarpine induced) was mixed with an equal volume of sample buffer and separated by SDSPAGE in gels containing either casein or potato starch. To examine amylase activity, a 4% agarose gel made with Tris phosphate buffer (pH 8.5) was produced containing 2 mg potato starch (modified from Cohen, 1990). Wells were punched in the gel as described above and 10 μL of salivary gland homogenate samples were loaded, incubated for 3–5 h and developed with a 2 mM iodine solution containing 2% potassium iodide. Cleared areas indicated amylase activity while blue-black areas indicated no amylase activity. Positive (alpha-amylase, Sigma A2771) and negative controls (di H2O) were performed. As a third test for amylase activity, the above SDS-PAGE procedure using casein as a substrate was modified by the substitution of starch for casein. The procedure was the same except after incubation the gel was developed with the iodine solution. Once stained, the gel was photographed with a digital camera and subsequently stained with Coomassie blue to Table 1 ApiZYM enzyme assay results yielding positive reactions from salivary gland extracts of B. lutarium and L. uhleri and their relative strength of reaction Enzyme
B. lutarium
Alkaline phosphatase Esterase Esterase lipase Trypsin α-Chymotrypsin Acid phosphatase Naphthol-AS-BI-phosphohydrolase α-Glucosidase N-acetyl-β-glucosaminidase β-Glucosidase Leucine arylamidase Valine arylamidase Cystine arylamidase α-Galactosidase β-Galactosidase α-Mannosidase α-Fucosidase
2+ 3+ 3+ 1+ 1+ 3+ 5+ 1+ 4+ – – – – – – – –
L. uhleri
Reaction strength 5+ 3+ 1+ 5+ 1+ 5+ 5+ 1+ 5+ 1+ 1+ – – – – – –
All experimental specimens were in 150 mM Tris at pH 7.4. Fifty microliters of saliva from each species were used. Experiment was run in duplicate.
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determine the protein banding pattern and estimate molecular weight of active bands.
adjacent central canal. The cuboidal glandular cells contain numerous active mitochondria and large volumes of smooth endoplasmic reticulum (Fig. 1A).
3. Results 3.1. Gland morphology 3.1.1. Main and lateral gland Both main and lateral glands consist of a cluster of grape-like alveoli each attached to a central lumen which empties anteriorly into the storage sacs. The main gland is larger in terms of both number and size of alveoli than the lateral gland. The alveoli are composed of simple cuboidal epithelia forming a sac with a hollow central lumen (Fig. 1A, B and D). The lumen holds the product of the glandular cells and empties into the
3.1.2. Accessory gland The accessory salivary gland is an elongate single-cell layered sock-like structure composed of two cell types. The distal half of the gland is composed primarily of large cuboidal cells with a few smaller cuboidal cells. The proximate half of the gland is composed of mainly the smaller cuboidal cells with a few dispersed large cuboidal cells. The lumen of the proximal portion of the gland is protected with a thin layer of cuticle while the distal portion has no endocuticular lining but instead is lined with a thick microvillus brush border. The interface between the distal and proximal portions is guarded by thick
Fig. 2. Azocasein and azoalbumen digestion by saliva of L. uhleri and B. lutarium. (A) L. uhleri saliva (150 mM Tris–HCl) digestion of 2% azocasein and azoalbumin. All differences due to temperature are statistically different (n = 3, SS = 2.16 × 10− 7, F = 103.523, P < 0.001). (B) L. uhleri saliva digestion of azocasein at various pH values with either TLCK (50 μg/mL in pH 6.0), TPCK (50 μg/mL, pH 7.5), or no protease inhibitors. (C) B. lutarium saliva digestion of azocasein and azoalbumin. All effects due to temperature are significant (n = 3, SS = 1.02 × 10− 7). (D) B. lutarium saliva (with either TLCK, TPCK, or no protease inhibitors) digestion of azocasein at various pH values. (Abs U/pg/h = spectrophotometric absorbance units per unit of azocasein or azoalbumin per picogram of salivary protein per hour of incubation time).
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3.2.4. SDS-PAGE for protease activity L. uhleri exhibited three bands of protease activity on casein impregnated SDS-PAGE while B. lutarium exhibited two active bands. Lethocerus exhibited proteases with molecular masses of approximately 30, 34, and 36 kDa while B. lutarium showed activity bands at 36 and 40 kDa (Fig. 4). 3.2.5. SDS-PAGE for amylase activity Starch impregnated agarose gels indicate a positive response for amylase from salivary gland homogenates of B. lutarium and no response from L. uhleri. Results from polyacrylamide gels agreed with those from the starch gels. B. lutarium saliva exhibited a single strong band of activity with a molecular weight of approximately 50 kDa (Fig. 5). Fig. 3. Digestion of starch as a measure of amylase activity in B. lutarium and L. uhleri. All reactions were in 200 μL of 2% RBB-Starch in 0.02 M sodium phosphate buffer.
ring of connective tissue at the origin of the endocuticular lining (Fig. 1B and F). 3.1.3. Myoepithelial cells The main and lateral glands and the storage sacs but not the accessory gland are wrapped with myoepithelial cells. These muscle cells are attached to the surface of the alveoli by a thin layer of connective tissue which covers the muscle cell and attaches to the cell membrane. Fluorescent staining for actin and transmission electron microscopy of thin sections indicates the presence of striated muscle tissue (Fig. 1A, C and E). 3.2. Physiology 3.2.1. Qualitative enzyme analysis The apiZYM enzyme assay yielded a positive reaction for nine enzymes in the salivary gland extract from B. lutarium and 11 enzymes from L. uhleri (Table 1). Lethocerus tested positive for an additional protease enzyme (leucine arylamidase) and the carbohydrase β-glucosidase.
3.2.6. Division of labor within the salivary gland Casein impregnated polyacrylamide gels indicate protease activity in the accessory gland and the storage sacs in both species tested but no protease activity in the main or lateral gland lobes. Starch impregnated polyacrylamide gels indicate a single strong band of amylase activity in the main gland of B. lutarium (at 50 kDa) but no activity from any other gland component and no activity from L. uhleri (Fig. 5). 4. Discussion The salivary glands among the four sub-orders of the Hemiptera including seven infraorders of Heteroptera are quite diverse morphologically and physiologically (Cobben, 1978; Schuh and Slater, 1995). This variety is not surprising given the numerous feeding habits represented in these 70,000+ species including, zoophagous, phytophagous, zoophytophagous, granivorous, and parasitic (haematophagous) (Goodchild, 1966; Miles, 1972; Cohen, 1990; McGavin, 1992; Terra and Ferreira, 1994). Although the Belostomatinae and Lethocerinae are very closely related, utilize similar habitats, hunting strategies (lie-in-
3.2.2. Azocasein and azoalbumin protease assay Both B. lutarium and L. uhleri exhibited proteolytic activity with azocasein and azoalbumin as substrates; the activity increased with temperature and pH (Fig. 2). Protease activity of both species was reduced by the protease inhibitors PMSF, TLCK, and TPCK (Fig. 2B and D). However, although activity was reduced, significant proteolytic activity remained in the presence of either TLCK or TPCK independently. Using TPCK and TLCK concurrently eliminated most protease activity in saliva of B. lutarium but did not significantly increase the inhibition of activity in L. uhleri saliva when compared to TLCK or TPCK alone (not shown). 3.2.3. Starch azure amylase assay B. lutarium exhibited positive amylase activity but L. uhleri did not. Amylase activity in B. lutarium peaked at a pH of 7.5 and declined above and below this value (Fig. 3).
Fig. 4. Protease activity in saliva of L. uhleri (A) and B. lutarium (B) in SDSPAGE with 5 mg casein incorporated in each gel. mw = molecular weight markers. Masses of molecular weight markers indicated in thousands of kDa on left margin.
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Fig. 5. Casein (5 mg) impregnated SDS-PAGE of the four parts of the salivary gland complex of B. lutarium (A) and L. uhleri (B). mw = molecular weight, main = main lobe, lat = lateral lobe, acc = accessory gland, stor = storage sac. (C) Comparison of activity of main salivary gland lobe of B. lutarium (BM) and L. uhleri (LM) on a starch impregnated (5 mg) SDS-PAGE stained with iodine. (D) After the iodine has been photo-catalyzed, the same gel is shown stained with Coomassie blue.
wait predators), and reproductive modes (paternal care), we show that their suites of digestive enzymes (in accord with their prey choices) are distinctly different. 4.1. Morphology Gland morphology indicates that the multi-part salivary gland of Belostomatids contains three productive parts, the main gland, lateral gland, and the accessory gland. The acinar type main and lateral gland components are emptied with the aid of myoepithelial cells. The mechanism for emptying the accessory gland probably resembles that of similar salivary structures in other insects involving a bank of muscles in the thorax (see Periplaneta sp. Baumann et al., 2002). The accessory gland is protected from autodigestion by a cuticular lining of the anterior portion of its lumen and by a cuticular lining of the salivary duct at the junction of the lateral gland duct and anteriorly through the afferent salivary duct. The lumenal surface of the lateral gland, where proteases are secreted, is protected by a thick microvillus border (Yadav, 1992; Swart and Felgenhauer, 2003). These morphological features of the gland, the microvillus border and cuticular layer, compliment the division of labor experiments indicating the production of proteases in the accessory gland. 4.2. Physiology B. lutarium produces several salivary enzymes including two proteases and an amylase. The salivary gland is divided into three functional components, the main gland which produces
amylase (among others), the accessory gland which produces the proteases and is protected by a thin layer of cuticle, and a small lateral gland. Previous work with Heteroptera indicated that the accessory gland did not produce an enzymatic product but served only to alter the pH of the saliva and activate enzymes produced in the other gland components (Baptist, 1941; Miles, 1972). Although Baptist's (1941) work on 20 species of diverse Heteroptera and more recent work on Lethocerus indicus by Yadav (1992) indicate the presence of secretory epithelia and a microvillus border, no previous work has been able to identify active enzymes from the accessory gland of any Heteropteran. Our evidence indicates that the accessory gland is very much active in producing protease enzymes. Many of the numerous works on Heteroptera salivary enzymes have bypassed the problem of the site of production of particular enzymes by using whole gland extracts or pilocarpine induced saliva which includes the products of all three productive components of the salivary gland (Cohen, 1989, 1990, 1993). One early work suggested the production of proteases in the accessory gland in the terrestrial heteropteran, Oncopeltus fasciatus (Bronskill et al., 1958) but further work on this species failed to confirm these previous results (Miles, 1967, 1972). We hope our evidence for protease production in the accessory salivary gland will encourage others to pursue further studies of division of labor among salivary gland components in other Heteroptera. Protease activity increases with increasing pH and temperature in agreement with other Heteroptera while the amylase has a peak activity at a pH of around 7.5. The larger species L. uhleri produces many of the same salivary enzymes as B.
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lutarium. However, it produces three proteases and does not exhibit salivary amylase activity. Previous workers have found trypsin-like endopeptidases in salivary glands of the terrestrial heteroptera; Zelus renardii (2 peaks at approximately 21 and 27 kDa, Cohen, 1993; Cohen and Wheeler, 1998), D. nebulosus (Boyd et al., 2002), Lygus lineolaris, and Lygus hesperus (Agustí and Cohen, 2000). Chymotrypsin-like activity has been found in the terrestrial Creontiades dilutus (Colebatch et al., 2001) and D. nebulosus (Boyd et al., 2002). Three proteolytic proteins have been demonstrated in Platymeris rhadamanthus (Edwards, 1961). The only semi-aquatic heteropteran for which data is available, Lethocerus cordofanus, demonstrates as many as five different proteolytic enzymes (Rees and Offord, 1969). Amylase activity has been found in the predatory, or partially predatory terrestrial heteroptera; Geocoris punctipes, Orius insidiosus (Zeng and Cohen, 2000a,b, Zeng et al., 2002), Z. renardii (Cohen, 1993), Deraeocoris pulchellus (Hori, 1972). We are aware of no previous data indicating amylase activity among the semi-aquatic Heteroptera. Protease inhibition by TLCK, TPCK, and PMSF indicates that the two proteases produced by B. lutarium belong to the trypsin and alpha-chymotrypsin families. The three proteases found in L. uhleri include trypsin-like and alpha-chymotrypsinlike enzymes; however, molecular weight information indicates that one of these is significantly different from its counterpart in B. lutarium. The proteolytic activity of L. uhleri saliva is significantly stronger than that of B. lutarium (Table 1) even when corrected for the amount of salivary protein present in the assay (Fig. 2). Whether this represents the mass effect of the third proteolytic enzyme activity in L. uhleri or kinetically different enzyme activities between the two species is unknown. The third protease produced by L. uhleri is not inhibited by TLCK, TPCK, and PMSF. Although we did not test a specific substrate for this third enzyme the qualitative analysis indicates that it acts as the endopeptidase; leucine arylamidase. 4.3. Ecological implications B. lutarium exploits a wide variety of small invertebrates including snails, dragonfly larvae, adult and larval Coleoptera, but do not take small fishes (Mori and Ohba, 2004; Ohba and Nakasuji, 2006; Swart and Taylor, 2004; Saha and Raut, 1992). There have been no reports of belostomatids being zoophytophagous like some terrestrial Heteroptera (e.g., Miridae) and this is not likely. One intriguing explanation for the presence of amylase in belostomatids is that they are able to exploit the plant material already ingested by their prey. L. uhleri is much larger and known to exploit small fish and occasionally other vertebrates (Torre Bueno, 1906; Gonsoulin, 1973; Smith, 1974; Kehr and Schnack, 1991; Saha and Raut, 1992; Chase, 1999; Mori and Ohba, 2004; Ohba and Nakasuji, 2006). The additional proteolytic activity in Lethocerinae may serve to either subdue larger prey faster (amphibia, fishes, snakes) or allow faster ingestion of protein rich meals. The complexity of the salivary enzymes, the functional morphology of the ingestive organs, and ecological implications of belostomatid biology deserve further attention.
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Acknowledgements We are indebted to G. Meche, and V. R. Townsend, Jr., for support in field collection and discussion of ideas. Much of this work would not have been possible without the advice and material support of Dr. T. Pesacreta and open access to the equipment at the University of Louisiana at Lafayette Microscopy Center. CCS was supported by Louisiana Board of Regents Doctoral Fellowship grant LEQSF(1998-03)-GF-28 through R. G. Jaeger. Further support was provided by funding from the Graduate Student Association of The University of Louisiana at Lafayette to CCS, and NSF grant IBN-9807948 to J. Spring and B.E.F.
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