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Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects
Summary April 21st, 2018
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.
Transcript
Assassin bugs are a group of insects that use venom to paralyze and pre-digest their prey. However, despite animal venoms being of interest as novel medicines, insecticides, and scientific tools, the toxins that they produce are almost entirely uncharacterized. One reason for this is that assassin bugs are less well-known in comparison to iconic venomous arthropods such as spiders and scorpions.
Another reason is that until recently, little information has been available about how to harvest their toxins. In this video, we will present methods that will enable researchers to successfully harvest venom toxins from assassin bugs, and also related insects such as giant fish-killing water bugs. Protocol one describes how to harvest venom by electrostimulation.
First, collect suitable insects. Adults and large nymphs are the easiest to work with. Using bugs two to seven days after feeding is ideal, to ensure the insects have accumulated venom stocks but do not starve.
Prepare a pair of forceps with electrodes at the tips, and connect them to an electricity supply. Ideally, this supply should be an electrostimulator capable of delivering five millisecond pulses at five Hertz. Use peak voltages between 15 and 25 volts for small or large bugs, respectively.
If an electrostimulator is not available, use constant voltages of five to 12 volts. Restrain the insect using rubber bands and a foam platform. Place the proboscis of the insect into a collecting tube.
A P200 pipette tube is ideal for most assassin bugs. Optionally, a small amount of water can be added to the tip before harvesting, to ensure maximum recovery of venom. Apply conductive gel to the electrodes, switch on the power supply, and apply the electrified tweezers to the insect.
Experiment with a range of contact points while monitoring venom expulsion from the proboscis. Store the venom at freezing temperatures to avoid autoproteolytic digestion. Repeat the electrostimulation with a new pipette tube until no more venom is forthcoming.
Protocol two describes how to harvest venom by provoking a defensive reaction. Prepare and restrain a bug as described above, inserting the proboscis into a collecting tip. Gently harass the bug to provoke a defensive reaction, without injuring the animal.
Sometimes, simply restraining the animal or inserting the proboscis into the collection tube can provoke a defensive reaction accompanied by venom expulsion from the proboscis. If not, gently touch the animal on the thorax, legs, abdomen, and especially the antennae, to elicit venom. In protocol three, in the written form of this article, we also describe how to harvest venom from species that spit venom defensively.
Protocol four describes how to harvest venom directly from the venom glands by dissection. Anesthetize animals by exposure to carbon dioxide for 10 minutes. Insert three pins into the posterior abdomen to hold the insect down without puncturing the venom glands.
Cut a short midline incision in the ventral surface of the abdomen using a miniature scalpel. Use miniature scissors to extend the midline in the anterior direction to the head, taking care to cut the exoskeleton only, and not damage internal structures. To expose the internal structures, make multiple lateral cuts extending from the midline incision to the side of the insect.
The ventral exoskeleton between each two cuts can then be pinned back to reveal the internal structures. Flood the dissection tray using a saline solution until the bug is submerged, allowing the internal structures to float up and be more easily visualized. The venom glands will appear as translucent elongated structures extending on each side of the alimentary canal.
Using tweezers and microscissors, carefully remove connective and nervous tissue and trachea, trying not to rupture the alimentary canal or the venom glands. The main gland is recognizable due to its characteristic structure, with anterior and posterior lobes, and two ducts meeting at the hilus. For assassin bugs, and perhaps other heteropterans, almost all toxins are stored in the lumens of the two lobes of the main gland.
Therefore, after freeing the main gland from trachea and connective tissue, the two ducts leading from the hilus can be cut, and the main venom gland collected. If the accessory gland is also desired, it is usually closely opposed to the gut, and can be identified unambiguously by tracing the ducts from the hilus. In any case, take care not to break the glands prematurely and lose the venom into the surrounding saline.
If the lobes of the main gland are to be separated, do this quickly after removal from the saline, and immediately proceed to harvest the contents of the gland lumens, as described in steps 4.6 and 4.7 of the written manuscript. Results will vary depending on the exact subprotocol and species of insect used. The first and second subprotocols presented in this video showed how to harvest venom toxins by electrostimulation and harassment, respectively.
Both of these methods typically yield a concentrated liquid containing approximately 50 to 250 milligrams a mil of protein, and over 100 protein-based components, including enzymes, pore-forming toxins, and peptides. The principal advantage of both of these protocols is that they are quick, non-lethal, and venom is uncontaminated by glandular tissue. The greatest disadvantage for harvesting by electrostimulation or harassment is that the glandular source of the venom obtained is unknown, and the protein content of venom harvested may differ, depending on the protocol.
Also, electrostimulation and harassment are not effective for all predatory species of heteroptera. In the last subprotocol, we showed how to dissect the engorged venom glands to harvest toxins directly. This technique also yields a cocktail of diverse proteins, although in this case the venom toxins may show some contamination with glandular tissue.
However, an advantage of harvesting toxins from dissected glands is that the glandular compartment of origin is known exactly. In conclusion, we encourage researchers to select a harvesting method appropriate to their research question. For example, investigations into the biology of venom use within a species will benefit from employing multiple harvesting techniques.
For bioprospecting studies, a single method for harvesting venoms from multiple species may be more economical, depending on the specific experimental design. We hope this video and the accompanying written article will facilitate researchers to harvest venom from assassin bugs and other heteropteran insects, which is a prerequisite to understanding its composition, bioactivity, and evolution.
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