Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects

Summary

Although many insects in the suborder Heteroptera (Insecta: Hemiptera) are venomous, their venom composition and the functions of their venom toxins are mostly unknown. This protocol describes methods to harvest heteropteran venoms for further characterization, using electrostimulation, harassment, and gland dissection.

Abstract

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, and the majority of extant heteropterans retain this trophic strategy. Some heteropterans have transitioned to feeding on vertebrate blood (such as the kissing bugs, Triatominae; and bed bugs, Cimicidae) while others have reverted to feeding on plants (most Pentatomomorpha). However, with the exception of saliva used by kissing bugs to facilitate blood-feeding, little is known about heteropteran venoms compared to the venoms of spiders, scorpions and snakes.

One obstacle to the characterization of heteropteran venom toxins is the structure and function of the venom/labial glands, which are both morphologically complex and perform multiple biological roles (defense, prey capture, and extra-oral digestion). In this article, we describe three methods we have successfully used to collect heteropteran venoms. First, we present electrostimulation as a convenient way to collect venom that is often lethal when injected into prey animals, and which obviates contamination by glandular tissue. Second, we show that gentle harassment of animals is sufficient to produce venom extrusion from the proboscis and/or venom spitting in some groups of heteropterans. Third, we describe methods to harvest venom toxins by dissection of anaesthetized animals to obtain the venom glands. This method is complementary to other methods, as it may allow harvesting of toxins from taxa in which electrostimulation and harassment are ineffective. These protocols will enable researchers to harvest toxins from heteropteran insects for structure-function characterization and possible applications in medicine and agriculture.

Introduction

Heteropteran venoms are potently bioactive substances¹. For example, the venom/saliva secretions of blood-feeding Heteroptera such as kissing bugs (Triatominae) and bed bugs (Cimicidae) facilitates feeding by disrupting hemostasis². Toxins in these venoms target multiple pathways including coagulation, platelet aggregation and vasoconstriction, as well as the pain and itch pathways. Venoms from most other heteropteran species are adapted to facilitate predation rather than blood-feeding. Their venoms cause paralysis, death and tissue liquefaction when injected into invertebrates^3,4. When injected into vertebrates, their venom may also have drastic effects. For example, injection of venom from the assassin bug Holotrichius innesi into vertebrates causes pain, muscle paralysis and hemorrhage; mice envenomated by this bug die quickly due to respiratory paralysis⁵.

Transcriptomic and proteomic studies have revealed the protein composition of some heteropteran venoms. Venoms of predaceous species are rich in proteases, other enzymes, and peptides and proteins of unknown structure and function^6,7,8. Kissing bug venom is rich in the triabin protein family, whose members profoundly affect coagulation, platelet aggregation, and vasoconstriction^2,9. However, it is not known which toxins underlie most bioactivities of venom. For example, venom of the kissing bug Triatoma infestans has been reported to be analgesic and inhibit sodium channels¹⁰, but the components responsible remain to be elucidated. Likewise, it is not known what component(s) of assassin bug venom cause paralysis or pain. A prerequisite for identifying the toxins responsible for particular venom bioactivities, and for characterizing the structure and function of novel venom toxins, is obtaining venom.

Venom has been obtained from heteropterans by electrostimulation^{5,6,7,8,11,12,13}, provocation of defensive responses^4,8, mechanically squeezing the thorax^12,14,15,16, dissecting out venom glands^{8,17,18,19,20,21,22}, and application of agonists of the muscarinic acetylcholine receptor²³. Judging the potential advantages and disadvantages of any method is complicated by the morphology of heteropteran venom glands, which consist of a main gland with two separate lumens, the anterior main gland (AMG) and posterior main gland (PMG), as well as an associated accessory gland (AG). These different gland compartments produce different protein secretions, which may be specialized for different biological functions including prey capture, defense and extra-oral digestion^8,17. In peiratine and ectrichodiine assassin bugs, the AMG has been associated with prey capture and the PMG with extra-oral digestion¹⁷. However, in the harpactorine bug Pristhesancus plagipennis the PMG is specialized for prey capture and digestion whereas the AMG is hypothesized to secrete defensive venom⁸. The AG has been described as having little secretory function in assassin bugs⁸ or as a major site of protease storage in giant water bugs²³. Clearly, further work is required to clarify the function of each gland compartment among various heteropteran subgroups, and to determine the function of most venom toxins. In this report we describe protocols for harvesting venom toxins from heteropterans toward this goal.

Protocol

This protocol complies with The University of Queensland's policy set out in Responsible Care and Use of Animals in Teaching and Research (PPL 4.20.11) as well as the National Health and Medical Research Council's Australian code for the care and use of animals for scientific purposes (8^th Edition 2013).

Caution: Take care not to be envenomated when handling assassin bugs. Take care to protect the eyes when handling species that spit venom defensively. Take care throughout not to injure the experimental animals. This includes monitoring of pressure on restraints such as rubber bands and ensuring that the proboscis is not broken.

NOTE: Optionally, anaesthetize animals by exposure to CO₂ for 0.5-2 min or cooling to 4-10 °C prior to venom harvesting in Aims 1-3 to facilitate safe transfer and restraint. Anaesthetization is not strictly required but may facilitate safe restraint of agile or strong specimens. However, animals must be awake to allow venom harvesting. Keep downstream applications in mind when deciding whether or not to add protease inhibitors.

1. Harvesting Venom Toxins by Electrostimulation

1. Obtain live specimens from which to harvest toxins.
2. Use pre-prepared plastic tweezers with positive and negative electrodes mounted on either tip. Connect electrified tweezers to an electrostimulator or a source of constant voltage that allows adjustment of the voltage.
   1. For small (~10 mm) and large (~25 mm) assassin bugs, use peak voltages of 15 and 25 V respectively.
   2. For larger heteropterans such as giant water bugs, use up to 40 V.
3. Restrain live bugs by strapping them to a platform, using a rubber band over the thorax.
4. Place the tip of the proboscis in a suitable collecting tip. For assassin bugs, use a P200 pipette tip. For giant water bugs, cut the extremity off a P200 tip to increase the size of the aperture.
   1. Gently lift the proboscis with a closed pair of clean tweezers and push the open aperture of the collection tip over the end of the proboscis.
   2. If desired, uptake ~5 µL ultrapure water prior to placing the proboscis into the collecting tip. This reduces losses of venom remaining inside of the tip, though the harvested venom will be diluted.
5. Apply electrostimulation. Dip the stimulating electrodes in conductive gel, such as 2.5 M NaCl/50% glycerol. Apply the electrodes to the thorax. For belostomatids, apply the two electrodes to the dorsal posterior surface of the head.
6. Store venom to prevent autodegradation. After venom is extruded, quickly transfer it to a tube at -20 °C or -60 °C, or a tube containing protease inhibitor cocktail.
7. Repeat steps 1.5 and 1.6 until sufficient venom is acquired or no further venom is forthcoming.

2. Harvesting of Venom Toxins by Harassment

1. Prepare animals for venom harvesting and place the tip of the proboscis in a collection vestibule, as described in subsections 1.1 and 1.3-1.4.
2. If venom is extruded spontaneously, go to step 2.3. If not, harass the animals by gently touching it on the legs, abdomen and antennae with tweezers until venom is produced.
3. Quickly transfer venom to a tube at -20 °C or -60 °C, or a tube containing protease inhibitor cocktail, if desired.

3. Harvesting of Venom Toxins by Harassment from Venom "Spitting" Species

1. Anaesthetize, or partly anaesthetize, the insect before removing it from its enclosure to prevent any premature defensive spitting.
2. Provoke venom spitting behavior. Contain and reposition the insect using the deep lid of a standard 90 x 16 mm Petri dish. Hold the lid slightly posterior and 1-4 cm above the insect to prevent flight. Most insects will spit several times, often in quick succession. Ensure all venom is collected on the underside of the dish.
3. Collect the venom on the underside of the Petri dish by rinsing with 10 µL of ultrapure water. Quickly transfer it to a tube at -20 °C or -60 °C, or a tube containing protease inhibitor cocktail.

4. Harvesting Venom Toxins by Gland Dissection

1. Sacrifice animals. Heavily anaesthetize or kill animals using >5 min exposure to CO₂. Pipe pure CO₂ directly into the air-holes of the animal's housing enclosure.
2. Pin insect to dissection tray. For assassin bugs, dissect through the ventral surface (4.3). For giant water bugs, dissect through the dorsal surface (4.4).
3. Ventral dissection
   1. Insert three pins into the posterior abdomen to hold the insect down without puncturing the venom glands.
   2. Cut a short midline incision in the ventral surface of the abdomen using a miniature scalpel. Use miniature scissors to extend the midline incision anteriorly to the head, taking care to cut the exoskeleton only and not damage internal structures.
   3. To expose the internal structures, make multiple lateral cuts extending from the midline incision to the side of the insect. Then, pin back each flap of ventral exoskeleton to reveal internal structures.
   4. For large assassin bugs, make four lateral incisions, in the mid-abdomen, anterior abdomen, between first and second legs, and in front of the first leg.
4. Dorsal dissection
   1. Remove the wings near the base. Insert three pins into the posterior abdomen to hold the insect down without puncturing the venom glands.
   2. Cut a midline incision from the head to the abdomen using miniature scissors and scalpel, taking care to cut the exoskeleton only and not damage internal structures.
   3. Force apart the two halves of the insect. Place several pins laterally along the length of the insect to leave the internal cavity exposed.
   4. Remove the flight muscles using tweezers.
5. Flood the dissection tray. Add PBS until the bug is submerged to allow internal structures float up and be more easily visualized.
6. Using tweezers and micro-scissors, carefully remove connective and nervous tissue and trachea. The venom glands appear as elongated, translucent structures extending along each side of the alimentary canal.
   1. Identify the main gland by its characteristic appearance, with anterior and posterior lobes and two ducts meeting at the hilus.
   2. If desired, identify the accessory gland by tracing the duct from the hilus. Free the main gland by cutting the two ducts emanating from the hilus.
7. Harvest the desired gland lumens. Transfer the gland to a microcentrifuge on ice containing 30 µL of PBS or PBS plus protease inhibitor cocktail. Lance the glands with a clean sharp pin.
   1. Vortex for 10 s and centrifuge (1 min, 5,000 × g, 4 °C) to empty the gland lumens. Remove the glandular tissue using tweezers.
8. Clarify the toxin extract. Centrifuge (5 min, 17,000 × g, 4°C) to remove any solid particles, retaining the supernatant and discarding the pellet. Store at -20°C or -60 °C to prevent autoproteolytic degradation.

Results

Some heteropteran species, such as the harpactorine P. plagipennis and the reduviine Platymeris rhadamanthus, reliably yield large quantities (5-20 µL) of venom in response to electrostimulation (Table 1). In general, most peiratine, reduviine, and harpactorine bugs yield venom in response to this method. Among stenopodaine bugs, electrostimulation elicited venom from Oncocephalus sp. but not Thodelmus sp. The holoptiline and emesine bu...

Discussion

The most critical step in harvesting assassin bug venom is selecting the appropriate method depending on the purposes of the study. Each of the three methods presented for harvesting heteropteran venoms has advantages and disadvantages depending on downstream applications.

Inducing bugs to expel venom from the proboscis (Protocols 1-3) avoids contamination of venom by glandular tissues. In addition, these methods are non-lethal and can be repeated many times over the course of a bug's life. El...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Electostimulator	Grass Technologies	S48 Square Pulse Stimulator	Electrostimulator allowing pulsed electrostimulation
Featherlight tweezers	Australian Entomological Supplies	E122B	For handling live venomous insects
Protease inhibitor cocktail	Sigma	4693124001	For preventing autoproteolytic digestion of venom
Dissection equipment	Australian Entomological Supplies	E152Micro	For fine dissections
Insect pins	Australian Entomological Supplies	E162	For fine dissections