Name: the ingestion of fluorescent, magnetic nanoparticles for determining fluid-uptake abilities in insects
Uploaded: 2017-12-20T15:00:00
Description: Watch this Scientific Journal Video about The Ingestion of Fluorescent, Magnetic Nanoparticles for Determining Fluid-uptake Abilities in Insects at JoVE.com

This work was supported by National Science Foundation (NSF) grant no. IOS 1354956. We thank Dr. Andrew D. Warren (McGuire Center for Lepidoptera and Biodiversity, Florida Museum of Natural History, University of Florida) for permission to use the butterfly images.

1. Insect Species, Preparation of Solutions and Feeding Station Setup
NOTE: Cabbage butterflies (Pieris rapae L., Pieridae) are selected as the species of representative Lepidoptera because they have been used in previous studies of fluid-uptake abilities and mouthpart morphology22,23. House flies (Musca domestica L., Muscidae) and blue bottle flies (Calliphora vomitoria L., Calliphoridae) are used because they...

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Insect mouthpart functionality is historically inferred from studies of gross morphology (e.g., lepidopteran proboscis functionality related to a drinking straw25,26); however, recent studies that incorporate experimental evidence have resulted in a paradigm shift in our understanding of the complexities of insect mouthparts and structure-function relationships2,12,13...

Many insect groups have mouthparts (proboscises) adapted for feeding on fluids, such as nectar, rotting fruit, sap flows (e.g. Diptera1, Lepidoptera2, Hymenoptera3), xylem (Hemiptera4), tears (Lepidoptera5), and blood (Phthiraptera6, Siphonaptera7, Diptera7, Hemiptera8, Lepidoptera9). The ability of insects to feed on fluids is relevant to ecosystem health (e.g. pollination10), disease transmission4,11, biodiversification2,12, and studies of convergent evolution13. Despite the wide variety of food sources, a theme among some fluid-feeding insects is the ability to acquire small amounts of fluids, which could be confined to micro- or nano-sized droplets, liquid films, or porous surfaces.
Given the extensive diversity of fluid-feeding insects (more than 20% of all animal species14,15) and their ability to feed on a variety of food sources, understanding their feeding behaviors and fluid-uptake mechanisms is important in many fields. Insect mouthpart functionality, for instance, has played a role in the development of biomimetic technology, e.g., microfluidic devices that can perform tasks such as the acquisition of small amounts of fluids using methods similar to those employed by insects16. A fundamental problem in the studies of fluid uptake mechanisms, however, is determining not only how insects feed on fluids, but acquiring experimental evidence that supports the mechanism. Solely using behavior (e.g., probing with the proboscis12,17) as an indicator for feeding is insufficient because it does not confirm the successful uptake of fluids, nor does it provide a means to determine the route that fluids travel as they pass through the insect. In addition, performing experiments with small quantities of fluids better represents natural feeding scenarios where fluids are a limiting resource2,12.
X-ray phase contrast imaging was used with the Monarch butterfly (Danaus plexippus L.) to assess how butterflies feed on small amounts of fluids from porous surfaces12. Monarch butterflies use capillary action via spaces between cuticular projections (dorsal legulae) along the proboscis to bring fluids confined to small pores into the food canal. The incoming fluids form a film on the food canal wall that grows and collapses into a liquid bridge by Plateau instability12,18, which is then transported to the butterfly&#39;s gut by action of the sucking pump in the head. Although x-ray phase contrast imaging is an optimal tool for visualizing fluid flow inside insects12,19,20,21, the technique is not readily available and a more convenient method is needed for rapid assessment of an insect&#39;s ability to uptake fluids and ingest them.
To determine if the feeding mechanism for D. plexippus applies to other Lepidoptera and also to flies (Diptera) (both groups feed on liquids from porous surfaces), Lehnert et al.13 applied a technique for assessing an insect&#39;s ability to feed on small amounts of fluids from porous surfaces, which is reported in detail here. Although the protocol outlined here is for studies that use wetted and porous surfaces, the methodology can be altered for other studies, such as those addressing pool-feeding mechanisms. In addition, the applications extend to other fields, including microfluidics and bioinspired technology.

<table>
	<tbody>
		<tr>
			<td>20% sucrose solution</td>
			<td>Domino Sugar</td>
			<td></td>
			<td>Sugar needed to produce the sucrose solution with dH2O</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS)</td>
			<td>Sigma-Aldrich</td>
			<td>P5493</td>
			<td>10X concentration diluted to 1X in dH2O for insect dissections</td>
		</tr>
		<tr>
			<td>Single depression concave slide</td>
			<td>AmScope</td>
			<td>BS-C6</td>
			<td>Slide is necessary for feeding stage setup</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>NY6004700 (60 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>NY4104700 (41 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>NY3004700 (30 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>NY2004700 (20 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>NY1104700 (11 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>TCTP04700 (10 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>TETP04700 (8 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>TMTP04700 (5 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>EMD Millipore</td>
			<td>RTTP04700 (1 &#181;m)</td>
			<td>Nylon net filters and isopore filters needed to produce a porous surface for insect feeding</td>
		</tr>
		<tr>
			<td>Iris microdissecting scissors</td>
			<td>Carolina Biological Supply Company</td>
			<td>623555</td>
			<td>Scissors used for dissections</td>
		</tr>
		<tr>
			<td>Insect pins (#1)</td>
			<td>Bioquip Products</td>
			<td>1208B1</td>
			<td>Pins used during dissections and feeding trials</td>
		</tr>
		<tr>
			<td>Extra-fine point dissecting forceps</td>
			<td>Carolina Biological Supply Company</td>
			<td>624684</td>
			<td>Dissecting equipment</td>
		</tr>
		<tr>
			<td>Leica M205 C Stereoscope</td>
			<td>Leica Microsystems</td>
			<td>M205 C</td>
			<td>Stereoscope used for dissections</td>
		</tr>
		<tr>
			<td>Inverted confocal microscope</td>
			<td>Olympus</td>
			<td>IX81</td>
			<td>Fluorescent microscope used to detect magnetic nanoparticles</td>
		</tr>
		<tr>
			<td>Fisherbrand PTFE Disposable Stir Bar</td>
			<td>Fisherscientific</td>
			<td>S68067</td>
			<td>Magnet used to detect nanoparticles</td>
		</tr>
		<tr>
			<td>Kimtech Science Kimwipes</td>
			<td>Kimberly-Clark Professional</td>
			<td>34155</td>
			<td>Tissues used to secure insects during feeding trials</td>
		</tr>
		<tr>
			<td>House fly (Musca domestica) pupae</td>
			<td>Mantisplace.com</td>
			<td></td>
			<td>insects for experiments</td>
		</tr>
		<tr>
			<td>Blue bottle fly (Calliphora vomitoria) pupae</td>
			<td>Mantisplace.com</td>
			<td></td>
			<td>insects for experiments</td>
		</tr>
		<tr>
			<td>Cabbage butterfly (Pieris rapae) larvae</td>
			<td>Carolina Biological Supply Company</td>
			<td>144102</td>
			<td>insects for experiments</td>
		</tr>
		<tr>
			<td>Finnpipette F1&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>4641080N</td>
			<td>micropipette for dispensing liquids</td>
		</tr>
		<tr>
			<td>Finntip 250 pipette tips</td>
			<td>ThermoFisher Scientific</td>
			<td>9400250</td>
			<td>micropipette tips</td>
		</tr>
		<tr>
			<td>Microscope Glass cover slides (=coverslips) (24 x 24 mm)</td>
			<td>AmScope</td>
			<td>CS-S24-100</td>
			<td>coverslips for viewing the insect&#39;s crop on confocal microscope</td>
		</tr>
	</tbody>
</table>

the ingestion of fluorescent, magnetic nanoparticles for determining fluid-uptake abilities in insects

Fluid-feeding insects ingest a variety of liquids, which are present in the environment as pools, films, or confined to small pores. Studies of liquid acquisition require assessing mouthpart structure and function relationships; however, fluid uptake mechanisms are historically inferred from observations of structural architecture, sometimes unaccompanied with experimental evidence. Here, we report a novel method for assessing fluid-uptake abilities with butterflies (Lepidoptera) and flies (Diptera) using small amounts of liquids. Insects are fed with a 20% sucrose solution mixed with fluorescent, magnetic nanoparticles from filter papers of specific pore sizes. The crop (internal structure used for storing fluids) is removed from the insect and placed on a confocal microscope. A magnet is waved by the crop to determine the presence of nanoparticles, which indicate if the insects are able to ingest fluids. This methodology is used to reveal a widespread feeding mechanism (capillary action and liquid bridge formation) that is potentially shared among Lepidoptera and Diptera when feeding from porous surfaces. In addition, this method can be used for studies of feeding mechanisms among a variety of fluid-feeding insects, including those important in disease transmission and biomimetics, and potentially other studies that involve nano- or micro-sized conduits where liquid transport requires verification.

The study of patterns in fluid-uptake abilities among fluid-feeding insects requires determination of when feeding occurs. The protocol outlined here is used to test the limiting pore size hypothesis among Lepidoptera and Diptera13. The limiting pore size hypothesis states that fluid-feeding insects cannot feed from liquid-filled pores if the pore size diameter is smaller than the diameter of the feeding conduits12. Incoming fluids from the ...

Watch this Scientific Journal Video about The Ingestion of Fluorescent, Magnetic Nanoparticles for Determining Fluid-uptake Abilities in Insects at JoVE.com

The Ingestion of Fluorescent, Magnetic Nanoparticles for Determining Fluid-uptake Abilities in Insects

Fluid-feeding insects have the ability to acquire minute quantities of liquids from porous surfaces. This protocol describes a method to directly determine the ability for insects to ingest liquids from porous surfaces using feeding solutions with fluorescent, magnetic nanoparticles.

Fluid-feeding insects ingest a variety of liquids, which are present in the environment as pools, films, or confined to small pores. Studies of liquid ...

The overall goal of this procedure is to assess physiological parameters of the feeding in fluid-feeding insects using a sucrose solution mixed with fluorescent magnetic nanoparticles. This method can help answer key questions in the entomology field such as structure function relationships of insect mouth parts and insect feeding mechanisms. The main advantage of this technique is that it allows us to determine if insects are able to ingest fluids that are present either as pools or as liquid films.This demonstration utilizes blue bottle flies because their fluid uptake abilities and mouth part morphology are well documented. House flies, cabbage butterflies, and other fluid feeding insects could be used. Obtain fly pupae and place them in an environmental chamber with 18 hours of light per day and at 23 degrees Celsius.After adults emerge from their pupae do not feed them before the experiment. For the experimental food prepare 20 percent sucrose solution with 0.5 milligrams nanoparticles per milliliter. Then, prepare a feeding station consisting of a manipulator, a clamp and a feeding stage.On the feeding stage, hold the food with a concave slide. Also, prepare various nylon net filters and membrane filters of different pore sizes to cover the concave slide. Begin with wrapping the insect's bodies, legs and wings into a folded tissue paper.Leave only the head and mouth parts exposed. While doing this, keep the insect secured by pinching it's wings together. Now, secure the wings wrapped in tissue paper between two microscope slides.Then clamp the two slides and secure the clamp to a manipulator. Then, secure the insect above the feeding stage using the manipulator. Next, deposit 30 microliters of feeding solution into the middle of the concave slide and cover the solution with a porous piece of filter paper.The droplet will spread along the filter paper filling the pores. Next, position the insect so the distal regions of the mouth parts can make contact with the wetted filter paper such that only the flat part of the slide can be reached and the insect cannot feed from the concave region. Now, use an insect pin to extend the mouth parts onto the filter paper and allow the insect to feed for 45 seconds.If the insect does not express an interest in feeding hold the mouthparts to the filter paper for the fill 45 seconds using the pin. This completes the feeding protocol. For the dissection, load a 50 milliliter watch glass with enough PBS to cover the insect's body.Set this up along with all the dissecting tools at the microscope. Now, remove the insect from the tissue paper keeping its wings closed. Then, remove the head, legs and wings using spring micro-dissecting scissors and place the thorax and abdomen into the PBS.To dissect the gut, first take hold of the distal abdomen cuticle and cut the cuticle in an anterior direction along the lateral side from the posterior end to the thorax. Take special care to ensure that only the cuticle is cut and that the alimentary canal is not damaged. Next, remove the abdominal cuticle, fat body, and other structures with the assistance of insect pins until only the thorax and alimentary canal remain in the watch glass.Now, identify the crop. It's a sack-like structure, located near the juncture of the thorax and abdomen. If needed, make additional incisions into the thorax to reveal the crop.Then, cut away the remaining thorax leaving only the alimentary canal and the crop. To test for the presence of ingested nanoparticles begin by using a fine-point dissecting forceps to transfer the crop to a cover slip for imaging. Use care to prevent rupture of the crop.Immediately image the crop on an inverted confocal microscope at 20 times magnification using the CY3 channel or phase contrast. To identify ingested nanoparticles, wave a magnetic stir bar about 10 millimeters from the crop. Make each back and forth motion take approximately one second.While waving the magnet, slowly move the operating stage to scan for nanoparticle movement inside the nearly transparent crop. If the nanoparticles are present in the crop they will wave in unison with the magnetic stir bar. An insect feeding on solution from a porous surface must create a stable liquid bridge via plateau instability to overcome the capillary pressure that keeps the liquid in the pores.It is predicted that the size of an insect's feeding conduit will determine the smallest pore size from which they can feed. After insects were dissected to assess if nanoparticles were ingested, their mouth parts were imaged to measure the size of the food canal conduit. For flies, the diameter of the pseudotrachea and the oral opening were measured.Analysis of the ingested nanoparticles suggests a close relationship between the distal mouth part conduit size, and the smallest pore size from which an insect can feed. This relationship is not seen with the proximal mouth parts. Consequently, fluid uptake for dipterans would first occur in the pseudotrachea rather than the oral opening and capillary action must have an important role in feeding.After watching this video, you should have a good understanding of how to feed insects a nanoparticle solution, dissect them to isolate their crop, and then inspect the crop for the nanoparticles. While attempting this procedure, it is important to be careful during the dissections. If the crop is ruptured, it can spill the liquid contents and nanoparticles thus ruining your sample.

Research

JoVE Journal

Biology

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