We would like to thank Dr. Josh Benoit for his assistance with bioinformatics Hailey Tobler for her help in raising Sunburst Diving Beetles and Tamara Pace for editorial assistance. This research was supported by the National Science Foundation under grants IOS-1456757 and IOS-1856341 to EKB and IOS1557936 to YT.

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The authors have nothing to disclose.

Our goal is that this compilation of methods will make RNAi widely available, especially as this tool remains a powerful synergistic technique to CRISPR/Cas9-based gene editing, with the advantage that it can be applied to the desired developmental stages of studied organisms. To exemplify this strength, we injected dsRNA into embryos and into different larval stages. Injections into eggs affected the development of embryos (Figure 2), injections into the second larval stage had apparent effects on the third larval stage (Figure 4 and Figure 6), and injections into the third larval stage showed effects in the adults (Figure 5). While the exact timing has to be established experimentally, generally, injections take effect within a few days. The success of this process can be affected by the length of the dsRNA sequence. Here, we presented examples using a little over 200 bp to more than 800 bp. As a general rule, sequences between 100 and 600 bp are preferred to limit off-target effects, but sequences up to 1000 bp yield good results22. One question in regard to RNAi is the duration of knockdown that can be achieved through this technique. Since the phenotypes were terminal at each stage, we cannot comment on this issue based on our presented results. However, it has previously been noted that RNAi effects are generally relatively long lived, and that higher concentrations lead to longer lasting knockdowns20.
One limitation of this technique is that it works better for some organisms than for others, and there seems to be no direct way of predicting how well it will work a priori. Nevertheless, it has been found to work well for a large range of different organisms. Within arthropods, this includes arachnids26, crustaceans27, and a variety of insects, with particularly high success rates in beetles28. A further complication is that differences in phenotype severity often occur between individuals despite the application of the same amount of dsRNA. As illustrated in Figure 2B, variation can even occur within an individual. In our RNAi studies targeting different genes involved in T. marmoratus larval eye development, we have frequently found that some eyes are affected more severely than others. This phenomenon may be related to the relatively dense tissue of the eye cluster, with the dsRNA better able to reach some of the units.
For the successful execution of RNAi experiments, it is critical that several parameters are optimized for the target gene. For example, the concentration of the dsRNA and the length of the targeted gene can strongly influence the outcome20. Another critical parameter is how the injections are executed, as this process can greatly influence the survival rate. For embryos, we achieved the best results by targeting the center of the embryo. A well-laid-out plate allows for the injection of 100 or more embryos in a single session. For larvae, it is critical to insert the injection needle between the segments. These injections require more dsRNA, and based on larvae availability, our injection sets here typically only consisted of a few animals at a time. For all injections, it is critical to prevent air from entering the organism.
In some cases, the feedback loops of a gene regulatory network and genetic redundancy can influence the penetrance of RNAi phenotypes, despite consistent knockdowns. This seems to be the case for our behavioral observations of larvae with highly successful knockdowns of a prominent lens protein, Lens318. Although we verified the high efficiency of these knockdowns through qPCR, considerable variation was observed in the associated phenotypes. This result highlights the necessity of properly quantifying RNAi knockdowns (for details on options see22). If there is no clear a priori expectation in regard to the resulting phenotypes, a good way of controlling for the off-target effects of RNAi is to target the same gene with two non-overlapping sequences of dsRNA and to evaluate the results for common phenotypes.
In contrast to gene-editing techniques, RNAi is also a powerful tool for studying lethal genes, and there are two ways to do so. For example, if one is interested in the functional contribution of a gene where loss-of-function early in development is known to be lethal, a functional investigation of such a gene can be achieved by simply allowing the animal to develop normally and then knocking down the gene via RNAi later in development (i.e., in the adult). Alternatively, a gene where complete loss-of-function is known to be lethal can be investigated through a partial knockdown, which can be achieved by injecting a range of dsRNA concentrations. Some of our results show knockdowns of lac2, which are known to be lethal if the cuticle in insects becomes overly soft24. Even the lac2 RNAi beetle depicted in Figure 5 would be unlikely to survive outside laboratory conditions. Another lethal gene is cut, which codes for a transcription factor that is fundamental for cell-fate specification in various organ systems in arthropods and has been linked to glia development in the Drosophila visual system29. Based on our experience with cut RNAi in T. marmoratus embryos, we can evoke informative eye phenotypes in embryos that are able to complete their embryonic eye development (unpublished observations). Here, higher dosages appear to lead to higher lethality rates, while lower doses result in observable and informative phenotypes.
Our protocol not only lists the necessary steps for a researcher to pursue RNAi experiments on T. marmoratus, as illustrated, but also is generally applicable to other organisms, especially aquatic organisms. Among aquatic organisms, there are already several examples within crustaceans such as the water fleas Daphnia30 and shrimp (for a recent review, see reference31). There are ample opportunities among aquatic insects, as they have been estimated to comprise about 6% of all insect diversity, with likely more than 200,000 species32. Furthermore, RNAi has already been performed on water striders that tend to inhabit the surface of aquatic environments33. If no genome is present, then a transcriptome can be assembled de novo. As long as this process reveals contigs of a few hundred nucleotides, dsRNA against specific genes can be designed. Our protocol for immobilizing insects in agarose will likely also be useful for other procedures, especially for soft, malleable, and aquatic organisms. Taken together, RNAi remains a powerful technique for manipulating gene expression in a diverse group of organisms, even when no other molecular and genetic tools are available.

The question of how specific genes contribute to the evolution of diverse traits is an exciting topic in biology. Over the last few decades, much progress has been made in regard to dissecting the genetic underpinnings of developmental processes in a few model organisms, such as the nematode Caenorhabditis elegans, the fruit fly Drosophila melanogaster, and the house mouse Mus musculus1. More recently, the invention of powerful gene-editing techniques such as clustered regularly interspaced short palindromic repeats (CRISPR)/Cas92 has provided the ability to change the genetic code of non-model organisms (for examples see3,4). As a result, there has been a surge in genetic studies on a variety of organisms that had not previously been approached through molecular techniques. Considering the enormous diversity of our animal kingdom, with many interesting traits or trait variances that are only represented in specific species, this progress has made it an exciting time for evolutionary-developmental biology (&#8220;evo-devo&#8221;) related work. However, genome-editing techniques that are available to non-model organisms are relatively restricted in regard to the developmental time points to which they can be applied, making it challenging to discern the temporal properties related to the role that specific genes play in any trait. In addition, transgenic techniques are often limited to genes that are nonessential for survival (i.e., whose knockout does not result in lethality). Therefore, while gene-editing techniques have started to become popular, there remains a need for effective techniques that are applicable to a variety of different organisms at specific developmental time points and facilitate partial knockdowns (rather than complete loss-of-function). Here, we draw attention to RNA interference (RNAi), a somewhat dated yet powerful gene knockdown technique5 that is particularly valuable as a synergistic approach to gene editing. Specifically, we developed procedures that allow for the application of RNAi to aquatic diving beetles as an example that illustrates the implementation of this technique, from the acquisition of the necessary molecular sequences to the successful injection of double-stranded RNA (dsRNA) into eggs and larvae.
RNAi-based gene knockdown leverages an innate defense mechanism of organisms, in which dsRNA molecules facilitate the silencing of invading nucleic acid sequences, such as viruses and transposons6. In brief, dsRNA is taken up into the cell, where it is cut into 20&#8211;25 nucleotide pieces by the Dicer enzyme. These pieces then activate the formation of the RNA-induced silencing complex (RISC), which inhibits the targeted mRNA by binding to it at specific sites using the guide strand. This process ultimately leads to mRNA degradation and hence interferes with the translation of mRNA into the respective protein6. The RNAi-based gene knockdown technique presented here therefore relies on the injection of dsRNA. For animal models, this technique was originally developed in C. elegans7 and D. melanogaster8 but has since emerged as a powerful functional genetic tool in non-model organisms9,10. Owing to its highly effective nature in some insects, RNAi can even be applied in pest management11.
As a research tool, RNAi has been used to examine how key molecular-developmental pathways function in nontraditional insect models. For example, RNAi in the flour beetle Tribolium&#160;castaneum has been instrumental in determining how deeply conserved genes contribute to specific traits in that beetle, as exemplified for the development of specifically shaped wings12,13,14 and eyes15,16. The techniques that underlie the manipulations in T. castaneum have been well described17 and rely on the ability to immobilize relatively dry eggs and larvae on a sticky surface. Such immobilization however is not possible for the wet developmental forms of aquatic organisms such as the Sunburst Diving Beetle Thermonectus marmoratus. As is the case for many nontraditional model organisms, it lacks an annotated genome. To manipulate gene expression in any organism without a genome, a reasonable and cost-efficient first step is to generate transcriptomes and identify the putative nucleotide sequences of the expressed genes of interest based on sequence similarity with related but more established model organisms, in this case, primarily Tribolium (Coleoptera) and Drosophila genes.
Here, to demonstrate how RNAi can be used on an aquatic organism, we first discuss protocols and software for RNA extraction and transcriptome generation and assemblage, which allow for the identification of specific targeted gene sequences. We then summarize the necessary steps for synthesizing gene-specific dsRNA. Subsequently, we illustrate how eggs can be injected in an aquatic environment and demonstrate incubation protocols for culturing developing embryos. In addition, we show how agarose gel can be used to completely immobilize larvae during the injection process, a technique that is generally useful during various procedures and could be applied to a variety of arthropods. To demonstrate how RNAi can be applied to different developmental stages, we include an example in which we silenced the eye pigmentation gene white in embryos. In addition, we describe an example in which the tanning gene laccase2(lac2) was silenced during both the second larval instar (to affect larvae of the third larval instar) and the third larval instar (to affect adults). Finally, we demonstrate that the injection of a lower concentration of dsRNA leads to partial knockdown, which shows that this technique can also be applied to genes where loss-of-function is known to be lethal.

<table>
	<tbody>
		<tr>
			<td>1. RNA isolation and de novo transcriptome assembly</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>liquid nitrogen</td>
			<td>University Stockroom</td>
			<td></td>
			<td>Typically locally available at research institutions</td>
		</tr>
		<tr>
			<td>pipette tips</td>
			<td>Fisher Scientific</td>
			<td>size dependent</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>RNeasy lipid tissue mini kit (RNA isolation kit)</td>
			<td>Qiagen</td>
			<td>74804</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>spectrophotometer (NnanoDrop)</td>
			<td>Thermo Fisher</td>
			<td>9836674</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1.2. De novo transcriptome assembly.</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BUSCO.v3</td>
			<td>https://busco.ezlab.org/</td>
			<td></td>
			<td>Any freely available software for assessing trasncriptome completeness can be used.</td>
		</tr>
		<tr>
			<td>CLC Genomics workbench</td>
			<td>Qiagen</td>
			<td>832021</td>
			<td>Other equivalent software packages are also available</td>
		</tr>
		<tr>
			<td>Galaxy workbench</td>
			<td>https://usegalaxy.org/</td>
			<td></td>
			<td>An open source online transcriptome assembly &#38; annotation pipeline</td>
		</tr>
		<tr>
			<td>NCBI BLASTx</td>
			<td>https://blast.ncbi.nlm.nih.gov/Blast.cgi?LINK_LOC=blasthome&#38;PAGE_TYPE=BlastSearch&#38;PROGRAM=blastx</td>
			<td></td>
			<td>An open source online alignment and annotation pipeline</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>2. cDNA synthesis, Cloning &#38; Miniprep isolation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ampicillin sodium salt</td>
			<td>Gibco</td>
			<td>11593-027</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>glycerol</td>
			<td>Fisher Scientific</td>
			<td>G33-500</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>LB broth</td>
			<td>Fisher Scientific</td>
			<td>BP1426-500</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>Omniscript RT (reverse trasncription) kit</td>
			<td>Qiagen</td>
			<td>205111</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>One Shot&#8482; TOP10 Chemically Competent E. coli</td>
			<td>Invitrogen</td>
			<td>C404010</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Fisher Scientific</td>
			<td>FB0875713</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>PureLink Quick Plasmid Miniprep Kit</td>
			<td>ThermoFischer</td>
			<td>771471</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>QIAquick PCR purification kit</td>
			<td>Qiagen</td>
			<td>28104</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>shaker-incubator</td>
			<td>Labnet</td>
			<td>211DS</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>spectrophotometer (NnanoDrop)</td>
			<td>Thermo Fisher</td>
			<td>9836674</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>thermal cycler for PCR</td>
			<td>BioRad</td>
			<td>T100</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>TOPO TA Cloning kit</td>
			<td>Invitrogen</td>
			<td>1845069</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>X-Gal Solution</td>
			<td>Thermo Scientific</td>
			<td>R0941</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>2.2 PCR amplification &#38; in vitro dsRNA synthesis</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Fisher Scientific</td>
			<td>accuSpin Micro 17R</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>ethanol</td>
			<td>Fisher Scientific</td>
			<td>A4094</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>FastTaq DNA Polymerase, dNTPack</td>
			<td>Roche</td>
			<td>13873432</td>
			<td>This kit contains all the reagents necessary for a PCR</td>
		</tr>
		<tr>
			<td>MEGAclear Transcriiption clean up kit</td>
			<td>ThermoFischer</td>
			<td>AM1908</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>MEGAscript T7 Transcription kit</td>
			<td>ThermoFischer</td>
			<td>AM1334</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>nuclease-free water</td>
			<td>Fisher Scientific</td>
			<td>AM9932</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>QIAquick PCR purification kit</td>
			<td>Qiagen</td>
			<td>28104</td>
			<td>Detailed specific protocol is provided with the kit</td>
		</tr>
		<tr>
			<td>sodium acetate</td>
			<td>Fisher Scientific</td>
			<td>BP333-500</td>
			<td>Make 3M working solution</td>
		</tr>
		<tr>
			<td>Spectrophotometer (NnanoDrop)</td>
			<td>Thermo Fisher</td>
			<td>9836674</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>3. Collecting and preparing early stage T. marmoratus embryos for dsRNA injections</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>agarose</td>
			<td>Fisher Scientific</td>
			<td>9012-36-6</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>distilled water</td>
			<td>Fisher Scientific</td>
			<td>9180</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>forceps (Dumon #4 Biology)</td>
			<td>Fine Science Tools</td>
			<td>11242-40</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>glass cavity dish (3 well-dish)</td>
			<td>Fisher Scientific</td>
			<td>50-243-43</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>microwave</td>
			<td>Welbilt</td>
			<td>turn-table</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>natural hair paintbrush</td>
			<td>Amazon</td>
			<td></td>
			<td>Any fine brush will do</td>
		</tr>
		<tr>
			<td>P1000 micro-pipetter</td>
			<td>Gilson</td>
			<td>F123602</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Fisher Scientific</td>
			<td>FB0875713</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>stereomicroscope</td>
			<td>Microsocope Central</td>
			<td>10446293</td>
			<td>Any good stereomicroscope will work</td>
		</tr>
		<tr>
			<td>transfer pipettes</td>
			<td>Fisher Scientific</td>
			<td>21-200-109</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4. dsRNA micro-injections in early stage T. marmoratus embryos</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>digital camera</td>
			<td>Edmund optics (Qimaging)</td>
			<td>Retiga 2000R</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>ethanol</td>
			<td>Fisher Scientific</td>
			<td>A4094</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>food dye</td>
			<td>Kroger</td>
			<td></td>
			<td>Any available food dye should work fine</td>
		</tr>
		<tr>
			<td>humidity chamber</td>
			<td></td>
			<td></td>
			<td>take any plastic box with a lid, sterilize it with 70% ethanol and let it dry</td>
		</tr>
		<tr>
			<td>incubator</td>
			<td>Labline</td>
			<td>203</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>injection buffer</td>
			<td></td>
			<td></td>
			<td>Prepared for 1 mL following reference #22 : Mix 10 &#181;L of 0.1 M sodium phosphate buffer, 100 &#181;L of 0.5 M potassium chloride solution, 100 &#181;L of food dye and 790 &#181;L of double-distilled water. Store in 4 &#176;C.</td>
		</tr>
		<tr>
			<td>intracellular Microinjection Systems (Picospritzer)</td>
			<td>Parker</td>
			<td>052-0500-900</td>
			<td>Currently the model III is available, but older models work also</td>
		</tr>
		<tr>
			<td>micro needle holder</td>
			<td>A-M Systems</td>
			<td>672441</td>
			<td>Other products would work equally well</td>
		</tr>
		<tr>
			<td>microinjection needles (1.2 mm x 0.68 mm, 4 inches)</td>
			<td>A-M Systems</td>
			<td>603000</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>micromanipulator</td>
			<td>Drummond Scientific Company</td>
			<td>3-000-024-R</td>
			<td>Any quality micromanipulator will work</td>
		</tr>
		<tr>
			<td>monosodium phosphate</td>
			<td>Fischer scientific</td>
			<td>7558-80-7</td>
			<td>To make 10 mL of 1 M working solution: Add 1.2 g of monosodium phosphate powder to 10 mL of Double distilled water and mix until clear solution is obtained</td>
		</tr>
		<tr>
			<td>P-1000 Micropipette Puller</td>
			<td>Sutter Instrument</td>
			<td>P-1000</td>
			<td>Puller settings:Heat 575, Pull 60, Velocity 75 Delay 110, Pressure 700.</td>
		</tr>
		<tr>
			<td>P10 micro-pipetter</td>
			<td>Gilson</td>
			<td>F144802</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>P1000 micro-pipetter</td>
			<td>Gilson</td>
			<td>F123602</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>potassium chloride</td>
			<td>Fischer scientific</td>
			<td>7447-40-7</td>
			<td>To make 100 mL of 0.5 M potassium chloride solution: Add 3.73 g of potassium chloride crystals to 100 mL of double distilled water and mix until clear solution is obtained.</td>
		</tr>
		<tr>
			<td>sodium phosphate buffer (0.1M)</td>
			<td></td>
			<td></td>
			<td>To make 10 mL of 0.1 M of this buffer: Mix 8.5 mL of 1 M sodium phosphate dibasic solution with 1.5 mL of 1 M monosodium phosphate solution. Check the pH with a pH meter and adjust accordingly to pH 7.6 at room temprature.</td>
		</tr>
		<tr>
			<td>sodium phosphate dibasic</td>
			<td>Fischer scientific</td>
			<td>7558-79-4</td>
			<td>To make 10 mL of 1 M working solution: Add 1.42 g of sodium phosphate dibasic powder to 10 mL of Double distilled water and mix until clear solution is obtained</td>
		</tr>
		<tr>
			<td>stereomicroscope</td>
			<td>Microsocope Central</td>
			<td>10446293</td>
			<td>Any good stereomicroscope will work</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>5. Preparing T. marmoratus larvae for dsRNA injections</td>
		</tr>
		<tr>
			<td>agarose</td>
			<td>Fisher Scientific</td>
			<td>9012-36-6</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>forceps (Dumon #4 Biology)</td>
			<td>Fine Science Tools</td>
			<td>11242-40</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Fisher Scientific</td>
			<td>FB0875713</td>
			<td>Products from other vendors would work equally well</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>6. dsRNA micro-injections in T. marmoratus larvae</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>injection buffer (10x)</td>
			<td></td>
			<td></td>
			<td>See section 4</td>
		</tr>
		<tr>
			<td>microinjection needles (1.2 mm x 0.68 mm, 4 inches)</td>
			<td>A-M Systems</td>
			<td>603000</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>microinjection syringe</td>
			<td>A-M Systems</td>
			<td>603000</td>
			<td>Other models would work equally well</td>
		</tr>
		<tr>
			<td>micro needle holder</td>
			<td>A-M Systems</td>
			<td>672441</td>
			<td>Other products would work equally well</td>
		</tr>
		<tr>
			<td>P-1000 Micropipette Puller</td>
			<td>Sutter Instrument</td>
			<td>P-1000</td>
			<td>Puller settings:Heat 575, Pull 60, Velocity 75 Delay 110, Pressure 700.</td>
		</tr>
		<tr>
			<td>stereomicroscope</td>
			<td>Microsocope Central</td>
			<td>10446293</td>
			<td>Any good stereomicroscope will work</td>
		</tr>
	</tbody>
</table>

rna interference in aquatic beetles as a powerful tool for manipulating gene expression at specific developmental time points

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is applicable to a wide variety of organisms and is highly efficient in the species-rich order Coleoptera (beetles). Here, we summarize the necessary steps for developing this technique in a novel organism and illustrate its application to the different developmental stages of the aquatic diving beetle Thermonectus marmoratus. Target gene sequences can be obtained cost-effectively through the assembly of transcriptomes against a close relative with known genomics or de novo. Candidate gene cloning utilizes a specific cloning vector (the pCR4-TOPO plasmid), which allows the synthesis of double-stranded RNA (dsRNA) for any gene with the use of a single common primer. The synthesized dsRNA can be injected into either embryos for early developmental processes or larvae for later developmental processes. We then illustrate how RNAi can be injected into aquatic larvae using immobilization in agarose. To demonstrate the technique, we provide several examples of RNAi experiments, generating specific knockdowns with predicted phenotypes. Specifically, RNAi for the tanning gene laccase2 leads to cuticle lightening in both larvae and adults, and RNAi for the eye pigmentation gene white produces a lightening/lack of pigmentation in eye tubes. In addition, the knockdown of a key lens protein leads to larvae with optical deficiencies and a reduced ability to hunt prey. Combined, these results exemplify the power of RNAi as a tool for investigating both morphological patterning and behavioral traits in organisms with only transcriptomic databases.

Using the protocol described above, we knocked down three different genes, namely, white, laccase2 (lac2), and Lens3 (Table 1), at a variety of different developmental stages of the Sunburst Diving Beetle T. marmoratus. We performed RNAi in T. marmoratus by injecting dsRNA at a very early stage during embryogenesis (Figure 1A). As some of the embryos do not survive the process and turn necrotic (Figure 1B), they need to be removed to keep the remaining embryos healthy. Exemplified here are the injections of dsRNA against the white gene. This gene is well known in Drosophila as one of three ATP-binding cassette (ABC) transporters involved in the uptake and storage of the precursors of eye pigment23. Accordingly, its loss-of-function results in an unpigmented, white-eye phenotype. Our results show that the injections of dsRNA targeting the orthologous white gene in T. marmoratus embryos leads to the loss of eye pigmentation in newly emerging larvae. In this case, wild type larvae are characterized by heavily pigmented eyes, and the RNAi knockdown of white leads to various levels of reduction and even complete elimination of eye pigment. Overall, we observed at least some reduction in eye coloration in 34% of surviving embryos (n = 35). Figure 2A compares a control individual and an individual with slightly lighter eye color. Figure 2B illustrates a more severe knockdown in an individual, in which the more ventral eyes (Eyes 2&#8211;5) of the cluster are completely unpigmented, whereas the dorsal eyes still show some pigmentation. These differences highlight how the efficiency of the knockdown can vary regionally, which is possibly related to differences in the efficacy of dsRNA penetration in dense tissue. Another individual shows essentially complete pigment loss in all eyes (Figure 2C).
To investigate how well RNAi works at the larval stage of T. marmoratus, we injected dsRNA targeting the tannic gene lac2 into second instar larvae a few days before they were due to molt into third instars (Figure 3) and evaluated the effect on the cuticular coloration of third instar larvae. Lac2 is a type of phenoloxidase that oxidatively conjugates proteins to make them insoluble, harder, and darker. Knockdown in the flour beetle T. castaneum has been shown to lead to lighter colored individuals in low doses but is considered lethal in high doses24. Figure 4 illustrates that this treatment also leads to lighter colored Sunburst Diving Beetle larvae. Specifically, in this experiment, 75% of the surviving injected larvae (n = 12) had reduced pigmentation (compared to 0% in the control group). Figure 4A shows an individual with relatively mild depigmentation, whereas Figure 4C illustrates the head of a T. marmoratus larva in which the dark coloration of the cuticle is nearly absent. Depigmentation was particularly evident for the central dark patterning that is typical for these larvae, whereas this pattern remained clearly visible in a control-injected individual (Figure 4B). In addition, a lightening of the tail trachea was observed, as depicted in Figure 4D. In the case where lac2 dsRNA was injected into third instar larvae, lighter adult individuals were obtained (Figure 5). Note that the wings of the knockdown beetle are somewhat deformed, likely due to its unusual softness.
In addition to altering morphological traits, it is possible to use RNAi to target genes that affect behavior. To demonstrate this, we performed RNAi against a key lens protein coding gene, Lens318, and injected it into second instar larvae to affect the optical properties of third instar larval eyes. Any effects on the lens observed here are likely because the eyes of T. marmoratus larvae undergo major eye growth at this transition, which also involves major optical changes of the lens25. RNAi knockdown in this experiment was highly efficient. Verification through qPCR showed a 100% success rate; out of 13 tested individuals, 12 were knocked down to less than 10% of the expression level of control individuals, with the remaining individual having an expression level of 17% of the control level (unpublished observation). At the phenotypic level, only some individuals were severely handicapped or incapable of prey capture, as is illustrated in Figure 6 for an individual that repeatedly approached its prey from a very close distance but consistently overshot it.
Table 1: Primer sequences and amplicons for white, lac2, and Lens3 proteins. <a href="https://cloudflare.jove.com/files/ftp_upload/61477/Table1.xlsx" target="_blank">Please click here to download this table.</a>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61477/61477fig01.jpg" /> 
Figure 1: Illustration of embryo injections.&#160;(A) Dechorionated embryos lined up on an agarose plate and injected near their center using a microinjection needle filled with dsRNA and food-dye-containing injection buffer. The scale bar represents 500 &#956;m. (B) An individual with a more severe knockdown. Injected embryos are kept in a humidity chamber and monitored daily to score phenotypes and remove dead individuals (which turn brown). The scale bar represents 5 mm. <a href="https://www.jove.com/files/ftp_upload/61477/61477fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61477/61477fig02.jpg" /> 
Figure 2: Example of fully developed embryos that were injected with dsRNA against white.&#160;(A) Comparison of an individual with a reduction in eye pigmentation (left) and a control-injected individual (right). E1&#8211;E6 refer to Eyes 1&#8211;6. (B) An individual with a more severe knockdown phenotype, which illustrates that, at times, some of the eyes within the cluster are more severely affected by the knockdown than others. (C) Individual with a nearly complete loss of eye pigmentation. The scale bar represents 200 &#956;m; Panels B and C are represented at the same scale. <a href="https://www.jove.com/files/ftp_upload/61477/61477fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61477/61477fig03.jpg" /> 
Figure 3: Illustration of larval injections.&#160;(A) Several larvae immobilized by embedding in 2% agarose with their tail spiracles left clear of any agarose. (B) Microelectrode containing the injection solution placed so that its tip can penetrate the fine membrane between two segments. (C) Blue injection dye visible at the injection site after the injection. Scale bars represent 1 cm. <a href="https://www.jove.com/files/ftp_upload/61477/61477fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61477/61477fig04.jpg" /> 
Figure 4: RNAi for laccase2 applied to second instar larvae resulting in reduced cuticle coloration in third instar larvae.&#160;(A) Relatively mild loss of coloration in a lac2 RNAi individual (bottom) when compared with a control-injected individual (top). The scale bar represents 5 mm. (B) Head of a control individual showing the characteristic dark colored pattern at the center of the head. (C) Relatively severe knockdown of lac2 leading to a nearly complete loss of central head coloration. (D) Loss of coloration in the major tail cuticle of a lac2 RNAi individual (bottom) when compared with a control-injected individual (top). Scale bars in B&#8210;D represent 1 mm. <a href="https://www.jove.com/files/ftp_upload/61477/61477fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61477/61477fig05.jpg" /> 
Figure 5: RNAi for laccase2 applied to a third instar larva resulting in reduced cuticle coloration in an adult.&#160;The knockdown individual (left) is also characterized by very soft elytra when compared with the control (right). The scale bar represents 5 mm. <a href="https://www.jove.com/files/ftp_upload/61477/61477fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61477/61477fig06.jpg" /> 
Figure 6: Knockdown of a major lens protein leading to deficiencies in prey capture.&#160;(A&#8211;C). Three examples wherein a Sunburst Diving Beetle larva, in which optical deficits were induced through RNAi, was unable to capture prey (mosquito larvae). Representative still images were selected from a video recording of characteristic prey captures. (D) Control larva catching prey. All scale bars represent 2 cm. <a href="https://www.jove.com/files/ftp_upload/61477/61477fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points at JoVE.com

RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points

RNA interference is a widely applicable, powerful technique for manipulating gene expression at specific developmental stages. Here, we describe the necessary steps for implementing this technique in the aquatic diving beetle Thermonectus marmoratus, from the acquisition of gene sequences to the knockdown of genes that affect structure or behavior.

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is ...

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7.5K Views.  University of Cincinnati. RNA interference is a widely applicable, powerful technique for manipulating gene expression at specific developmental stages. Here, we describe the necessary steps for implementing this technique in the aquatic diving beetle Thermonectus marmoratus, from the acquisition of gene sequences to the knockdown of genes that affect structure or behavior.

Video: RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points

Transient Gene Expression in Tobacco using Gibson Assembly and the Gene Gun

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately using complex regulatory strategies including differential promoters and/or signal sequences. Metabolic engineers and synthetic biologists interested in targeting enzymes to a particular organelle are faced with a challenge: For a protein that is to be localized to more than one organelle, the engineer must clone the same gene multiple times. This work presents a solution to this strategy: harnessing alternative splicing of mRNA. This technology takes advantage of established chloroplast and peroxisome targeting sequences and combines them into a single mRNA that is alternatively spliced. Some splice variants are sent to the chloroplast, some to the peroxisome, and some to the cytosol. Here the system is designed for multiple-organelle targeting with alternative splicing. In this work, GFP was expected to be expressed in the chloroplast, cytosol, and peroxisome by a series of rationally designed 5&#8217; mRNA tags. These tags have the potential to reduce the amount of cloning required when heterologous genes need to be expressed in multiple subcellular organelles. The constructs were designed in previous work11, and were cloned using Gibson assembly, a ligation independent cloning method that does not require restriction enzymes. The resultant plasmids were introduced into Nicotiana benthamiana epidermal leaf cells with a modified Gene Gun protocol. Finally, transformed leaves were observed with confocal microscopy.

This work describes a novel method for selectively targeting subcellular organelles in plants, assayed using the BioRad Gene Gun.

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately ...

Experimental Protocol for Manipulating Plant-induced Soil Heterogeneity

Coexistence theory has often treated environmental heterogeneity as being independent of the community composition; however biotic feedbacks such as plant-soil feedbacks (PSF) have large effects on plant performance, and create environmental heterogeneity that depends on the community composition. Understanding the importance of PSF for plant community assembly necessitates understanding of the role of heterogeneity in PSF, in addition to mean PSF effects. Here, we describe a protocol for manipulating plant-induced soil heterogeneity. Two example experiments are presented: (1) a field experiment with a 6-patch grid of soils to measure plant population responses and (2) a greenhouse experiment with 2-patch soils to measure individual plant responses. Soils can be collected from the zone of root influence (soils from the rhizosphere and directly adjacent to the rhizosphere) of plants in the field from conspecific and heterospecific plant species. Replicate collections are used to avoid pseudoreplicating soil samples. These soils are then placed into separate patches for heterogeneous treatments or mixed for a homogenized treatment. Care should be taken to ensure that heterogeneous and homogenized treatments experience the same degree of soil disturbance. Plants can then be placed in these soil treatments to determine the effect of plant-induced soil heterogeneity on plant performance. We demonstrate that plant-induced heterogeneity results in different outcomes than predicted by traditional coexistence models, perhaps because of the dynamic nature of these feedbacks. Theory that incorporates environmental heterogeneity influenced by the assembling community and additional empirical work is needed to determine when heterogeneity intrinsic to the assembling community will result in different assembly outcomes compared with heterogeneity extrinsic to the community composition.

Understanding the role of environmental heterogeneity in species coexistence has typically focused on types of heterogeneity that are extrinsic to the community&#8217;s species composition. We provide novel detailed methods for creating soil heterogeneity treatments using soils subject to plant-soil feedback conditioning, or heterogeneity intrinsic to the community composition.

Coexistence theory has often treated environmental heterogeneity as being independent of the community composition; however biotic feedbacks such as ...

An Aquatic Microbial Metaproteomics Workflow: From Cells to Tryptic Peptides Suitable for Tandem Mass Spectrometry-based Analysis

Meta-omic technologies such as metagenomics, metatranscriptomics and metaproteomics can aid in the understanding of microbial community structure and metabolism. Although powerful, metagenomics alone can only elucidate functional potential. On the other hand, metaproteomics enables the description of the expressed in situ metabolism and function of a community. Here we describe a protocol for cell lysis, protein and DNA isolation, as well as peptide digestion and extraction from marine microbial cells collected on a cartridge filter unit (such as the Sterivex filter unit) and preserved in an RNA stabilization solution (like RNAlater). In mass spectrometry-based proteomics studies, the identification of peptides and proteins is performed by comparing peptide tandem mass spectra to a database of translated nucleotide sequences. Including the metagenome of a sample in the search database increases the number of peptides and proteins that can be identified from the mass spectra. Hence, in this protocol DNA is isolated from the same filter, which can be used subsequently for metagenomic analysis.

This protocol is for the extraction and concentration of protein and DNA from microbial biomass collected from seawater, followed by the generation of tryptic peptides suitable for tandem mass spectrometry-based proteomic analysis.

Meta-omic technologies such as metagenomics, metatranscriptomics and metaproteomics can aid in the understanding of microbial community structure and ...

Characterization and Application of Passive Samplers for Monitoring of Pesticides in Water

Five different water passive samplers were calibrated under laboratory conditions for measurement of 124 legacy and current used pesticides. This study provides a protocol for the passive sampler preparation, calibration, extraction method and instrumental analysis. Sampling rates (RS) and passive sampler-water partition coefficients (KPW) were calculated for silicone rubber, polar organic chemical integrative sampler POCIS-A, POCIS-B, SDB-RPS and C18 disk. The uptake of the selected compounds depended on their physicochemical properties, i.e., silicone rubber showed a better uptake for more hydrophobic compounds (log octanol-water partition coefficient (KOW) &#62; 5.3), whereas POCIS-A, POCIS-B and SDB-RPS disk were more suitable for hydrophilic compounds (log KOW &#60; 0.70).

A protocol about the characterization and application of five different passive sampling devices is presented.

Five different water passive samplers were calibrated under laboratory conditions for measurement of 124 legacy and current used pesticides. This study ...

Alternative Method of Removing Otoliths from Sturgeon

Extracting the otoliths (ear bones) from fish that have very thick skulls can be difficult and very time consuming. The common practice of making a transverse vertical incision on the top of the skull with a hand or electrical saw may damage the otolith if not performed correctly. Sturgeons (Acipenseridae) are one family in particular that have a very large and thick skull. A new laboratory method entering the brain cavity from the ventral side of the fish to expose the otoliths was easier than other otolith extraction methods found in the literature. Methods reviewed in the literature are designed for the field and are more efficient at processing large quantities of fish quickly. However, this new technique was designed to be more suited for a laboratory setting when time is not pressing and successful extraction from each specimen is critical. The success of finding and removing otoliths using this technique is very high and does not compromise the structure in any manner. This alternative technique is applicable to other similar fish species for extracting the otoliths.

The goal of the protocol is to show an effective method to extract the otoliths from sturgeon carcasses.

Extracting the otoliths (ear bones) from fish that have very thick skulls can be difficult and very time consuming. The common practice of making a ...

A Flow-through Exposure System for Evaluating Suspended Sediments Effects on Aquatic Life

This paper describes the Fish Larvae and Egg Exposure System (FLEES). The flow-through exposure system is used to investigate the effects of suspended sediment on various aquatic species and life stages in the laboratory by using pumps and automating delivery of sediment and water to simulate suspension of sediment. FLEES data are used to develop exposure-response curves between the effects on aquatic organisms and suspended sediment concentrations at the desired exposure duration. The effects data are used to evaluate management practices used to reduce the interactions between aquatic organisms and anthropogenic causes of suspended sediments. The FLEES is capable of generating total suspended solids (TSS) concentrations as low as 30 to as high as 800 mg/L, making this system an ideal choice for evaluating the effects of TSS resulting from many activities including simulating low ambient levels of TSS to evaluating sources of suspended sediments from dredging operations, vessel traffic, freshets, and storms.

A robust and flexible flow-through exposure system designed to maintain sediment in suspension is presented. The system is used to investigate the effects of suspended sediment on various aquatic species and life stages in the laboratory.

This paper describes the Fish Larvae and Egg Exposure System (FLEES). The flow-through exposure system is used to investigate the effects of suspended ...

Double-stranded RNA Oral Delivery Methods to Induce RNA Interference in Phloem and Plant-sap-feeding Hemipteran Insects

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran insects account for a number of economically substantial pests of plants that cause damage to crops by feeding on phloem sap. The brown marmorated stink bug (BMSB), Halyomorpha halys (Heteroptera: Pentatomidae) and the Asian citrus psyllid (ACP), Diaphorina citri Kuwayama (Hemiptera: Liviidae) are hemipteran insect pests introduced in North America, where they are an invasive agricultural pest of high-value specialty, row, and staple crops and citrus fruits, as well as a nuisance pest when they aggregate indoors. Insecticide resistance in many species has led to the development of alternate methods of pest management strategies. Double-stranded RNA (dsRNA)-mediated RNA interference (RNAi) is a gene silencing mechanism for functional genomic studies that has potential applications as a tool for the management of insect pests. Exogenously synthesized dsRNA or small interfering RNA (siRNA) can trigger highly efficient gene silencing through the degradation of endogenous RNA, which is homologous to that presented. Effective and environmental use of RNAi as molecular biopesticides for biocontrol of hemipteran insects requires the in vivo delivery of dsRNAs through feeding. Here we demonstrate methods for delivery of dsRNA to insects: loading of dsRNA into green beans by immersion, and absorbing of gene-specific dsRNA with oral delivery through ingestion. We have also outlined non-transgenic plant delivery approaches using foliar sprays, root drench, trunk injections as well as clay granules, all of which may be essential for sustained release of dsRNA. Efficient delivery by orally ingested dsRNA was confirmed as an effective dosage to induce a significant decrease in expression of targeted genes, such as juvenile hormone acid O-methyltransferase (JHAMT) and vitellogenin (Vg). These innovative methods represent strategies for delivery of dsRNA to use in crop protection and overcome environmental challenges for pest management.

This article demonstrates novel techniques developed for oral delivery of double-stranded RNA (dsRNA) through the vascular tissues of plants for RNA interference (RNAi) in phloem sap feeding insects.

Phloem and plant sap feeding insects invade the integrity of crops and fruits to retrieve nutrients, in the process damaging food crops. Hemipteran ...

Characterization of Aquatic Biofilms with Flow Cytometry

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond quickly to environmental changes and can be thus used as indicators of water quality. Currently, biofilm assessment is mostly based on integrative and functional endpoints, such as photosynthetic or respiratory activity, which do not provide information on the biofilm community structure. Flow cytometry and computational visualization offer an alternative, sensitive, and easy-to-use method for assessment of the community composition, particularly of the photoautotrophic part of freshwater biofilms. It requires only basic sample preparation, after which the entire sample is run through the flow cytometer. The single-cell optical and fluorescent information is used for computational visualization and biological interpretation. Its main advantages over other methods are the speed of analysis and the high-information-content nature. Flow cytometry provides information on several cellular and biofilm traits in a single measurement: particle size, density, pigment content, abiotic content in the biofilm, and coarse taxonomic information. However, it does not provide information on biofilm composition on the species level. We see high potential in the use of the method for environmental monitoring of aquatic ecosystems and as an initial biofilm evaluation step that informs downstream detailed investigations by complementary and more detailed methods.

Flow cytometry in combination with visual clustering offers an easy-to-use and fast method for studying aquatic biofilms. It can be used for biofilm characterization, detection of changes in biofilm community structure, and detection of abiotic particles embedded in the biofilm.

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond ...

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly relevant as a model for the studies of osmoregulatory mechanisms because of its extreme tolerance to low salinity. It is also widely used as a model of settling biology, in particular in relation to antifouling research. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae for studies. A protocol for the all-year-round culturing of B. improvisus has been developed and a detailed description of all steps in the production line is outlined (i.e., the establishment of adult cultures on panels, the collection and rearing of barnacle larvae, and the administration of feed for adults and larvae). The description also provides guidance on troubleshooting and discusses critical parameters (e.g., the removal of contamination, the production of high-quality feed, the manpower needed, and the importance of high-quality seawater). Each batch from the culturing system maximally yields roughly 12,000 nauplii and can deliver four batches in a week, so up to almost 50,000 larvae per week can be produced. The method used to culture B. improvisus is, probably, to a large extent also applicable to other marine invertebrates with free-swimminglarvae. Protocols are presented for the dissection of various tissues from adults as well as the production of high-quality RNA for studies on gene expression. It is also described how cultured adults and reared cyprids can be utilized in a wide array of experimental designs for examining gene expression in relation to external factors. The use of cultured barnacles in gene expression is illustrated with studies of possible osmoregulatory roles of Na+/K+ ATPase and aquaporins.

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

The barnacle Balanus (Amphibalanus) improvisus is a model for studying osmoregulation and antifouling. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae. Here, a protocol for the all-year-round culturing of B. improvisus is described, including the production of larvae. The use of cultured barnacles in gene expression studies is illustrated.

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly ...

Experimental Protocol for Examining Behavioral Response Profiles in Larval Fish: Application to the Neuro-stimulant Caffeine

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and toxicological studies. Using automated digital tracking platforms, recent efforts in neuropharmacology are leveraging larval fish locomotor behaviors to identify potential therapeutic targets for novel small molecules. Similar to these efforts, research in the environmental sciences and comparative pharmacology and toxicology is examining various behaviors of fish models as diagnostic tools in tiered evaluation of contaminants and real-time monitoring of surface waters for contaminant threats. Whereas the zebrafish is a popular larval fish model in the biomedical sciences, the fathead minnow is a common larval fish model in ecotoxicology. Unfortunately, fathead minnow larvae have received considerably less attention in behavioral studies. Here, we develop and demonstrate a behavioral profile protocol using caffeine as a model neurostimulant. Though photomotor responses of fathead minnows were occasionally affected by caffeine, zebrafish&#160;were markedly more sensitive for photomotor and locomotor endpoints, which responded at environmentally relevant levels. Future studies are needed to understand comparative behavioral sensitivity differences among fish with age and time of day, and to determine whether similar behavioral effects would occur in nature and be indicative of adverse outcomes at the individual or population levels of biological organization.

Here, we present a protocol to examine larval zebrafish and fathead minnow locomotor activities and photomotor responses (PMR) using an automated tracking software. When incorporated in common toxicology bioassays, analyses of these behaviors provide a diagnostic tool to examine chemical bioactivity. This protocol is described using caffeine, a model neurostimulant.

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and ...