The authors are supported by projects ZeEBRA (R&#38;I-2019-018), a&#160;Technology Development Programme (TDP) fund, STRONG (R&#38;I-2024-007L), a&#160;TDPLite fund, NASDAC (SINO-MALTA-2022-08), a Science and Technology Cooperation grant, and DEMONSTRATE (R&#38;I-2018-007A), a Go2Market grant, all funded by the XjenzaMalta for and on behalf of the Foundation for Science and Technology. Figure 1 was created using BioRender.

Zebrafish were handled according to EU Directive 2010/63/EU. All experiments were approved by the Faculty Research Ethics Committee (FREC) and the Joint FREC Animal Research Sectoral Subcommittee (JFARSS) of the University of Malta. A flowchart of the proposed protocol can be found in Figure 1.
1. Egg production, collection, and maintenance
<ol>
	<li>Set up the fish in breeding tanks the day before, at least an hour after the last feed, with dividers in place separating the males and females. For better egg production results, use a higher ratio of females to males. Set the fish up in a room equipped with a 14 h:10 h light/dark cycle and at 27-28 &#176;C ambient temperature. 
	NOTE: A 1 L breeding tank can hold three adult fish, whereas a 1.7 L breeding tank can hold five adult fish.</li>
	<li>The following morning, following the onset of light, replace the water in the breeding tanks and remove the dividers. In case the breeding tank is not readily sloped, place it in a sloping position to ensure that shallow water is available to promote mating. In case of slow egg production and mating activity, replace the water in the breeding tanks every 45 min. Leave the fish to mate for up to 3 h or as required.</li>
	<li>Once mating is complete, gently squeeze any females that did not produce eggs to prevent long-term egg binding. Place the fish back into their respective tanks.</li>
	<li>Collect any eggs in the breeding tanks using a mesh strainer, rinse with embryo water (E3), and place in a Petri dish with E3. To prepare E3, mix&#160;16.67 mL of 60x E3 (300 mM NaCl; 10 mM KCl; 20 mM CaCl2.2H2O; 20 mM MgSO4.7H2O) with&#160;1 mL of 0.01% methylene blue and 982 mL of deionized water. Remove any unfertilized eggs or debris. Make sure the Petri dishes are not crowded, with 50 eggs per 100 mm x 15 mm Petri dish and 120 eggs per 150 mm x 15 mm Petri dish.</li>
	<li>Keep the eggs in an incubator set to 28.5 &#176;C and equipped with a 14 h:10 h day/night cycle. At 1 dpf, refresh the E3 and remove non-viable eggs to limit fungal growth.</li>
</ol>
2. Test plate preparation
<ol>
	<li>At 3 dpf, transfer one healthy-looking, hatched embryo to each well in a 96-well plate using a 1,000 &#181;L micropipette set to a 100 &#181;L volume. 
	NOTE: Non-hatched embryos should not be used, as this may result in variable compound uptake. Using embryos that appear healthy also limits bias during testing. The size of the pipette tip allows for easy collection of the zebrafish embryo without causing any physical damage. In addition, a micropipette allows for better control of the volume being dispensed into the well as opposed to a transfer pipette.</li>
	<li>In a 12-well plate, prepare the desired concentration(s) for each test chemical in separate wells (here, Bosutinib at a concentration range from 100 &#181;M to 6.25 &#181;M). In each well, add the necessary volume of test chemical using micropipettes and make up to 2.4 mL using E3. 
	NOTE: A literature search should be carried out to determine the adequate concentrations to use for the relevant test chemicals. If no information is available, several concentrations can be trialed until a suitable concentration range is determined. A minimum of five concentrations should be used, with the highest concentration causing 100% mortality and the lowest concentration causing no observable effect7.</li>
	<li>Prepare a vehicle control depending on the solvent used to dissolve the stock test chemical. 
	NOTE: Many test compounds are not water soluble and for this reason, they need to be dissolved in specific solvents (&#34;vehicles&#34;) such as ethanol or dimethyl sulfoxide (DMSO) to generate an aqueous solution. DMSO should preferably not exceed a concentration of 1% when used as a vehicle control, which is the concentration found to be safe in zebrafish14.</li>
	<li>Remove any E3 solution present in the 96-well plate using a 200 &#181;L micropipette and replace it with 100 &#181;L of the corresponding medium being tested. Be sure to test a total of 24 larvae per concentration. Use a single or 8-channel 200 &#181;L micropipette to transfer the test chemicals to the corresponding test plate wells. Include controls (e.g., E3, chemical vehicle, positive controls, etc). 
	NOTE: The plate setup accommodates up to four different test chemicals or four different concentrations of the same chemical. An example of the test plates set up for the investigation of five concentrations of the same chemical (e.g., Bosutinib) accompanied by controls is shown in Figure 1.</li>
	<li>At 24 h post exposure, perform morphological scoring (as described in section 3) and a halfmedium change, removing 50 &#181;L of medium and replacing it with freshly prepared medium. Remove and record any dead larvae in the process. Keep the larvae in an incubator set to 28.5 &#176;C and equipped with a 14 h:10 h day/night cycle. Repeat the process until 7 dpf.</li>
</ol>
3. Morphological assessment
<ol>
	<li>Using a stereo light microscope, observe the individual test larvae at an appropriate magnification. 
	NOTE: Examples of morphological defects that can be assessed are provided in Table 1.</li>
	<li>Have the morphology assessed by two independent scorers, with a third one being consulted in case of disagreements between the two scorers.</li>
	<li>Score the morphology using a binary method (i.e., present/absent), where &#39;absent&#39; indicates normal morphology (scored as &#39;0&#39;) and &#39;present&#39; indicates the presence of morphological abnormality (scored as &#39;1&#39;). Figure 2 shows examples of zebrafish at 5 dpf after 2 days of drug exposure, including an unaffected zebrafish (Figure 2A), a zebrafish with mild pericardial edema (Figure 2B), severe pericardial edema with yolk hemorrhage and yolk extension (Figure 2C), delayed development with reduced pigmentation, swim bladder and total length (Figure 2D), swollen yolk and a kinked caudal fin (Figure 2E), absent caudal fin (Figure 2F), curved notochord (Figure 2G), truncation (Figure 2H) and death with necrosis (Figure 2I). Score all abnormal phenotypes as &#39;1&#39; and the unaffected zebrafish as &#39;0&#39;.</li>
</ol>
4. LC50 determination
<ol>
	<li>Following morphology assessment, plot the cumulative percentage mortality at 7 dpf against concentration. An example is shown in Figure 3. Use the concentration curve to estimate the concentration that causes 50% mortality.</li>
	<li>Once the LC50 concentration is estimated, repeat the experiment described above by exposing 24 replicate larvae in a 96-well plate to this single estimated LC50 concentration.</li>
</ol>
5. Technical replicates
<ol>
	<li>Repeat the entire procedure using the selected five concentrations of the same chemical at least two more times on two separate occasions to ensure that the results are reproducible.</li>
</ol>

<ol>
	<li>Teame, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+zebrafish+(Danio+rerio)+as+biomedical+models.">The use of zebrafish (Danio rerio) as biomedical models.</a> Anim Front. 9 (3), 68-77 (2019).</li><li>Howe, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+zebrafish+reference+genome+sequence+and+its+relationship+to+the+human+genome.">The zebrafish reference genome sequence and its relationship to the human genome.</a> Nature. 496 (7446), 498-503 (2013).</li><li>Patton, E. E., Zon, L. I., Langenau, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+disease+models+in+drug+discovery:+From+preclinical+modelling+to+clinical+trials.">Zebrafish disease models in drug discovery: From preclinical modelling to clinical trials.</a> Nat Rev Drug Discov. 20 (8), 611-628 (2021).</li><li>Clift, D., Richendrfer, H., Thorn, R. J., Colwill, R. M., Creton, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+analysis+of+behavior+in+zebrafish+larvae:+Effects+of+feeding.">High-throughput analysis of behavior in zebrafish larvae: Effects of feeding.</a> Zebrafish. 11 (5), 455-461 (2014).</li><li>Keenan, S. R., Currie, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+developmental+phases+of+zebrafish+myogenesis.">The developmental phases of zebrafish myogenesis.</a> J Dev Biol. 7 (2), 12 (2019).</li><li>Tonelli, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish:+A+resourceful+vertebrate+model+to+investigate+skeletal+disorders.">Zebrafish: A resourceful vertebrate model to investigate skeletal disorders.</a> Front Endocrinol (Lausanne). 11, 489 (2020).</li><li>OECD. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Test no. 236: Fish embryo acute toxicity (FET). , (2013).</li><li>Chen, J. R., Lai, Y. H., Tsai, J. J., Hsiao, C. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+fluorescent+staining+platform+for+drug-screening+and+mechanism-analysis+in+zebrafish+for+bone+mineralization.">Live fluorescent staining platform for drug-screening and mechanism-analysis in zebrafish for bone mineralization.</a> Molecules. 22 (12), 2068 (2017).</li><li>He, H., Wang, C., Tang, Q., Yang, F., Xu, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Possible+mechanisms+of+prednisolone-induced+osteoporosis+in+zebrafish+larva.">Possible mechanisms of prednisolone-induced osteoporosis in zebrafish larva.</a> Biomed Pharmacother. 101, 981-987 (2018).</li><li>Huang, H. X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+bone+transgenic+zebrafish+in+anti-osteoporosis+chemical+screening.">Application of bone transgenic zebrafish in anti-osteoporosis chemical screening.</a> Animal Model Exp Med. 1 (1), 53-61 (2018).</li><li>Haney, M. G., Moore, L. H., Blackburn, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drug+screening+of+primary+patient+derived+tumor+xenografts+in+zebrafish.">Drug screening of primary patient derived tumor xenografts in zebrafish.</a> J Vis Exp. (158), e60996 (2020).</li><li>Nishimura, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+zebrafish+in+systems+toxicology+for+developmental+toxicity+testing.">Using zebrafish in systems toxicology for developmental toxicity testing.</a> Congenit Anom (Kyoto). 56 (1), 18-27 (2016).</li><li>Hayot, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluating+toxicity+of+chemicals+using+a+zebrafish+vibration+startle+response+screening+system.">Evaluating toxicity of chemicals using a zebrafish vibration startle response screening system.</a> J Vis Exp. (203), e66153 (2024).</li><li>Hoyberghs, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DMSO+concentrations+up+to+1%+are+safe+to+be+used+in+the+zebrafish+embryo+developmental+toxicity+assay.">DMSO concentrations up to 1% are safe to be used in the zebrafish embryo developmental toxicity assay.</a> Front Toxicol. 3, 804033 (2021).</li><li>Lipton, J. H., Br&#252;mmendorf, T. H., Sweet, K., Apperley, J. F., Cortes, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Practical+considerations+in+the+management+of+patients+treated+with+bosutinib+for+chronic+myeloid+leukemia.">Practical considerations in the management of patients treated with bosutinib for chronic myeloid leukemia.</a> Ann Hematol. 103 (9), 3429-3442 (2024).</li></ol>

The authors declare no conflicts of interest.

Zebrafish have become an important model for drug screening. When trying to determine a methodology to carry out toxicity testing, we found several contrasting methods from different laboratories8,9,10,11,12,13. The methods described use larvae at different ages, variable experimental timelines, different rates of medium changes, and different morphology scoring systems. Through trial and error, we established the described methodology, which has provided reliable and reproducible results.
Choosing a plate size 
Initially, experiments were performed in a 24-well plate to ensure that the larvae had enough space to move and grow. The plates were arranged with 10 larvae per well, two wells per concentration, and 1 mL of drug solution per well. This arrangement did not provide reproducible results, leading to the inference that the larvae were not absorbing equal amounts of drug from the well. Trials were then carried out using a 96-well and 48-well plate with one larva per well, which showed that larvae in 96-well plates displayed no issues with growth. Hence, experiments were continued in a 96-well plate utilizing less space and drug volumes.
Daily medium change 
Since zebrafish are absorbing the drug from the medium, we ascertained that a degree of medium change is needed to keep a consistent concentration of the drug in the medium. Moreover, a medium change removes potential waste material accumulated in the well, which can provide a toxic environment and lead to biased results. A trial was carried out with Bosutinib, where three plates were set up for the same drug, one without a daily medium change, one with a daily half medium change, and one with a full daily medium change. While the plate with no medium change exhibited increased mortality rates, the plates with half and full medium changes resulted in the same outcome by 7 dpf. Thus, a daily half medium change was chosen. This choice ensures less consumption of the expensive and limited test material, which could be required for further downstream experiments.
Exposure duration 
The age of test larvae used and drug exposure duration selected in this protocol was driven by our phenotype of interest being bone mineralization. Since bone formation commences at ~4 dpf, we chose to start with zebrafish at 3 dpf to determine whether test chemicals affected the onset of bone development. Subsequently, we ended the drug exposure studies at 7 dpf since a high degree of mortality was observed from 8 dpf onwards in view of the lack of feeding that might bias the results. Nevertheless, timeframes may be adjusted depending on the nature of the study and phenotype of interest.
Limitations 
This methodology has some limitations. Transferring 3 dpf zebrafish to multi-well test plates is time-consuming. Removing the medium from the wells without picking up the larvae is equally laborious. The medium needs to be removed from each well slowly using a single-channel pipette, ensuring that the larvae are not damaged in the process. Moreover, while a daily half medium change replenishes the test compound and ensures the removal of waste material from the well, it does not guarantee that a stable drug concentration can be maintained in the well throughout the 5 days of exposure. While high-performance liquid chromatography analysis could give further insight into the actual amount of drug being absorbed by each larva, the test&#160;would no longer qualify as a high-throughput screening method.
In conclusion, this methodology aims to target the gap in toxicity testing using zebrafish beyond 4 dpf, taking steps towards a standardized protocol applicable to wild-type, mutant, and transgenic zebrafish lines, and that is on par with the FET at the embryonic stage. This opens avenues for other downstream phenotyping assays that might not be possible at earlier stages.

Zebrafish (Danio rerio) have recently emerged as a powerful vertebrate model organism, becoming increasingly&#160;popular for studies to advance biomedical research1. Around 70% of human protein-coding genes have at least one zebrafish ortholog, with ~82% of human disease-causing genes having at least one orthologous gene in zebrafish2. The genetic conservation, coupled with similar basic organ patterning and morphology, make zebrafish an interesting model for human disease research,allowing for cross-species comparisons at the molecular level1,2,3, which is crucial for drug development.
Zebrafish develop rapidly with the tail bud forming at 10 h post fertilization (hpf) and the first somites appearing at 16 hpf. The embryos emerge from the chorion at ~2-3 days post fertilization (dpf). By 4 dpf, their digestive tract is fully developed and by 5 dpf, their swim bladder has inflated, allowing them to swim around freely and hunt for food. Despite the ability to feed by 5 dpf, larvae that are unfed can survive solely on yolk-derived nutrients until at least 7 dpf, circumventing the need for external feeding that may cause contamination and interfere with drug screening, toxicology, and behavioral testing4. By 7 dpf, larvae also develop a complete body plan with several rudimentary organ systems, including the heart and vasculature, muscle, and bone5,6. Zebrafish embryos are also fertilized ex vivo, developing externally, and are optically transparent in the early life stages, making them highly accessible for gene editing and screening of small molecules3. Additionally, their small size and high fecundity rate allow for the inclusion of more replicates, which increases statistical power, as well as for the use of multi-well plates, which saves on drug volumes and increases throughput.
Screening of novel small molecules in an animal model is important to determine the drugs&#39; ability to produce both beneficial or adverse effects, thus identifying which compounds are viable for further testing and others that might require modification to alter their activity. As part of the drug screening process, the lethal concentration 50 (LC50) of the target compound/s needs to be identified. This provides a working concentration that causes death in 50% of test animals as part of the preliminary toxicity testing.
A standardized protocol exists for toxicity testing in zebrafish embryos up to 96 h post fertilization (hpf), namely the Fish Embryo Acute Toxicity (FET) test (TG236)7. In the FET test, newly fertilized eggs are exposed to five concentrations of a chemical for up to 96 h and observed for signs of lethality, namely (i) egg coagulation, (ii) lack of somite formation, (iii) lack of tail bud detachment from the yolk sac, and (iv) lack of a heartbeat. The test allows for the determination of acute toxicity and the LC50. Furthermore, it also advises on test chamber sizes to use for screening (24-well plates), egg distribution in the test chamber, and housing conditions, amongst others, ensuring valid and reproducible results7.
While the FET test is suitable for toxicity testing at the embryonic stage, it does not go beyond 4 dpf. For phenotypes that start to develop at around this time point (e.g., mineralization of the notochord6), drug screening must proceed further than 4 dpf to enable the morphological assessment of such tissue. Moreover, we recommend that drug exposure commences at least a day before the development of the phenotype of interest. To our knowledge, no standardized protocol similar to the FET exists for drug screening in early-stage zebrafish larvae, specifically in zebrafish aged from 3 to 7 dpf. While several studies have been published describing drug testing in zebrafish larvae beyond 4 dpf, there is a lack of consistency in the methodology utilized, including the size of the test chambers, number of biological and technical replicates, rate of media changes, exposure durations, and morphological defects assessed8,9,10,11,12,13. For this reason, we developed a validated protocol for standardized toxicity testing followed by visual observation of morphological changes in zebrafish larvae at 3-7 dpf. The proposed protocol can subsequently be utilized for other downstream phenotyping assays, such as behavioral analysis, gene expression profiling, and histomorphometry.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5 mL microcentrifuge tube</td>
			<td>Starlab</td>
			<td>E1415-1510</td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#181;L micropipette (P10)</td>
			<td>Gilson</td>
			<td>F144802&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>10/20 &#181;L pipette tips</td>
			<td>Starlab</td>
			<td>S1110-3810</td>
			<td></td>
		</tr>
		<tr>
			<td>1000 &#181;L micropipette (P1000)</td>
			<td>Gilson</td>
			<td>F123602&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>1000 &#181;L pipette tips</td>
			<td>Starlab</td>
			<td>S1111-6001</td>
			<td></td>
		</tr>
		<tr>
			<td>12-well plate</td>
			<td>Starlab</td>
			<td>CC7672-7512</td>
			<td>Sterile, single-wrapped, non-treated, with lid</td>
		</tr>
		<tr>
			<td>20 &#181;L micropipette (P20)</td>
			<td>Gilson</td>
			<td>F123600&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#181;L micropipette (P200)</td>
			<td>Gilson</td>
			<td>F123601&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#181;L multi-channel pipette&#160;</td>
			<td>Gilson</td>
			<td>F81024</td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#181;L pipette tips</td>
			<td>Starlab</td>
			<td>S1113-1006</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well plate</td>
			<td>Starlab</td>
			<td>CC7672-7596</td>
			<td>Sterile, single-wrapped, flat-bottom, non-treated, with lid</td>
		</tr>
		<tr>
			<td>Breeding tanks with inside grid sloping bottom, tank dividers and lids</td>
			<td>Tecniplast</td>
			<td>ZB17BTISLOP, ZB17BTE,&#160; ZB17BTL, ZB17BTD</td>
			<td>Sloped inner tank</td>
		</tr>
		<tr>
			<td>Mesh strainer</td>
			<td>/</td>
			<td>/</td>
			<td></td>
		</tr>
		<tr>
			<td>Nitrile gloves</td>
			<td>Mercator</td>
			<td>/</td>
			<td>/</td>
		</tr>
		<tr>
			<td>Petri-dish&#160;</td>
			<td>Starlab</td>
			<td>CC7672-3394/CC7672-3359</td>
			<td>100 x 20 mm or 60 x 15 mm</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Leica M205 FCA Fluorescence stereo microscope</td>
			<td>Leica</td>
			<td>10450826</td>
			<td></td>
		</tr>
		<tr>
			<td>Peltier-cooled incubator including 2 shelves with light module cold white</td>
			<td>Memmert</td>
			<td>IPP110ecoplus, T8</td>
			<td>Equipped with day/night cycle</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl Sulfoxide</td>
			<td>Biochem Chemopharma</td>
			<td>504341000</td>
			<td>To prepare a 1% solution in E3</td>
		</tr>
		<tr>
			<td>Calcium chlorided dihydrate</td>
			<td>Biochem Chemopharma</td>
			<td>303080500</td>
			<td>To prepare E3</td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate</td>
			<td>Biochem Chemopharma</td>
			<td>313060500</td>
			<td>To prepare E3</td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>Biochem Chemopharma</td>
			<td>316030500</td>
			<td>To prepare E3</td>
		</tr>
		<tr>
			<td>SCREEN-WELL&#160;Wnt Pathway library (Bosutinib)</td>
			<td>Enzo</td>
			<td>BML-2838-0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Fisher Scientific</td>
			<td>S/3161/53</td>
			<td>To prepare E3</td>
		</tr>
	</tbody>
</table>

small molecule screening and toxicity testing in early-stage zebrafish larvae

Zebrafish have become a prominent model organism for human translational research, toxicity testing of small molecules and environmental pollutants, therapeutic drug discovery, and other biomedical research applications. The presented protocol aims to provide a clear guide for drug screening and toxicity testing in early-stage zebrafish larvae (aged 3-7 days post fertilization, dpf). Several methodologies describing toxicity testing in zebrafish larvae have been published, albeit lacking a standardized protocol for exposure durations, rate of media changes, and morphology scoring, amongst others. We experimentally determined the ideal larvae ages for conducting the experiments, the plate sizes that work best for those ages, the rate of media changes that provide reliable results while limiting wastage&#160;of test compound, and a morphology scoring system that is easily quantifiable in a statistical manner. The manuscript describes in detail a protocol for toxicity testing in zebrafish larvae aged between&#160;3 to 7 dpf, aiming to target the gap in drug toxicity testing and taking the first step towards a standardized method similar to the Fish Embryo Acute Toxicity test.

The abnormalities can be looked at individually or cumulatively over the experimental timeline until 7 dpf. The maximum morphology score per abnormality can be used to calculate morphological abnormalities as a percentage of the number of larvae used. For example, if on day 7 the pericardial edema morphology scores are at 19 and 20 for scorers 1 and 2, respectively, out of a possible score of 24, meaning 19 or 20 larvae out of 24 display pericardial edema, the percentage score can be calculated as:
<img alt="Equation 1" src="/files/ftp_upload/67759/67759eq1.jpg" />
Supplementary Table S1 shows the morphology scoring results (of both scorers) at 4 dpf following toxicity testing for the compound Bosutinib, a tyrosine kinase inhibitor indicated for the treatment of chronic myeloid leukemia15. The same morphology scoring is repeated for exposed fish aged from 5 dpf to 7 dpf. The compound was tested using a concentration range from 100 &#181;M to 6.25 &#181;M. Three technical replicates were conducted at different time points, and the average cumulative mortality rate at 7 dpf was used to determine LC50, the concentration at which 50% of the larvae survive. The LC50 can be determined at any exposure time point. However, we recommend calculating this at 7 dpf.
The cumulative average mortality at 7 dpf was used to generate an LC50 curve, which can be seen in Figure 3. For Bosutinib, the LC50 concentration was estimated to be 37.95 &#181;M.
NOTE: This estimated LC50 concentration strictly applies to this model system and can then be used for further downstream phenotyping assays.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67759/67759fig1.jpg" /> 
Figure 1: Schematic of the proposed drug screening in larvae at 3-7 dpf. The 96-well test plates are set up for the investigation of five concentrations ranging from 100 &#181;M to 6.25 &#181;M for the compound Bosutinib. Abbreviation: dpf = days post fertilization. <a href="https://www.jove.com/files/ftp_upload/67759/67759fig1large.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/67759/67759fig2.jpg" /> 
Figure 2: Representative morphological defects observed in zebrafish larvae at 5 dpf exposed to various chemical concentrations (at 2 days post exposure). (A) Control (unaffected) zebrafish, (B) Zebrafish larva showing mild pericardial edema, (C) Zebrafish with severe pericardial edema and yolk hemorrhage, (D) Zebrafish with developmental delay, reduced pigmentation, small swim bladder, (E) Zebrafish with swollen yolk and a kinked caudal fin, (F) Zebrafish with absence of caudal fin, (G) Zebrafish with curved notochord,&#160;(H) Zebrafish with truncated tail, and (I) Dead zebrafish with necrosis. Defects are noted with red arrowheads. Scale bars = 1 mm. <a href="https://www.jove.com/files/ftp_upload/67759/67759fig2large.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/67759/67759fig3.jpg" /> 
Figure 3: LC50 curve showing concentration against % mortality. The five concentrations tested were 100 &#181;M, 50 &#181;M, 25 &#181;M, 12.5 &#181;M, and 6.25 &#181;M, with % mortality being 100%, 100%, 29.2%, 12.5%, and 12.5%, respectively. <a href="https://www.jove.com/files/ftp_upload/67759/67759fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Morphological Defects</td>
			<td>Description</td>
		</tr>
		<tr>
			<td>Pericardial edema</td>
			<td>Fluid buildup around the heart with a stretched heart and malfunctioning of the blood circulation</td>
		</tr>
		<tr>
			<td>Heart beats (per minute)</td>
			<td>Reduced number of beats per minute&#160;</td>
		</tr>
		<tr>
			<td>Yolk</td>
			<td>Presence or absence of yolk edema or hemorrhage that can occur simultaneously with cardiac defects&#160;</td>
		</tr>
		<tr>
			<td>Yolk extension</td>
			<td>Yolk which is swollen, thin or absent&#160;</td>
		</tr>
		<tr>
			<td>Curved notochord</td>
			<td>Presence of a kinked or undulating notochord, and/or curved tail&#160;</td>
		</tr>
		<tr>
			<td>Swim bladder</td>
			<td>Presence, absence or partial development of the swim bladder</td>
		</tr>
		<tr>
			<td>Pigmentation</td>
			<td>Less or no melanocytes compared to the untreated fish of the same age</td>
		</tr>
		<tr>
			<td>Necrosis</td>
			<td>Presence (mild or severe) or absence of necrosis in any part of the body</td>
		</tr>
		<tr>
			<td>Delayed development</td>
			<td>Delayed development or stalled compared to wild-type controls of the same age.&#160;</td>
		</tr>
		<tr>
			<td>Response to touch</td>
			<td>Delayed or no response following a startle (e.g., using a pipette tip)</td>
		</tr>
		<tr>
			<td>Truncation</td>
			<td>Absence of posterior zebrafish body (i.e. formation of the tail)</td>
		</tr>
		<tr>
			<td>Fins</td>
			<td>Presence or absence of the caudal fin that may be accompanied by thin yolk extension</td>
		</tr>
		<tr>
			<td>Reduced body axis</td>
			<td>Fish that appear normal but when measured are shorter in length</td>
		</tr>
	</tbody>
</table>
Table 1: A list of visual morphological defects that can be observed following drug exposure studies.
Supplementary Table S1: Representative morphological results for Bosutinib at 4 dpf (1 day post exposure).&#160;<a href="https://www.jove.com/files/ftp_upload/67759/SupplementalTable_1.xlsx" target="_blank">Please click here to download this table.</a>

This protocol describes a drug screening procedure in zebrafish larvae 3-7 days post fertilization followed by morphological assessment.

Small Molecule Screening and Toxicity Testing in Early-stage Zebrafish Larvae

Zebrafish have become a prominent model organism for human translational research, toxicity testing of small molecules and environmental pollutants, ...

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JoVE Journal

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504 Views.  University of Malta. This protocol describes a drug screening procedure in zebrafish larvae 3-7 days post fertilization followed by morphological assessment. 

Video: Small Molecule Screening and Toxicity Testing in Early-stage Zebrafish Larvae

Placing Growth Factor-Coated Beads on Early Stage Chicken Embryos

The neural tube expresses many proteins in specific spatiotemporal patterns during development. These proteins have been shown to be critical for cell fate determination, cell migration, and formation of neural circuits. Neuronal induction and patterning involve bone morphogenetic protein (BMP), sonic hedgehog (SHH), fibroblast growth factor (FGF), among others. In particular, the expression pattern of Fgf8 is in close proximity to regions expressing BMP4 and SHH. This expression pattern is consistent with developmental interactions that facilitate patterning in the telencephalon.

Here we provide a visual demonstration of a method in which an in ovo preparation can be used to test the effects of Fgfs in the formation of the forebrain. Beads are coated with protein and placed in the developing neural tube to provide sustained exposure. Because the procedure uses small, carefully placed beads, it is minimally invasive and allows several beads to be placed within a single neural tube. Moreover, the method allows for continued development so that embryos can be analyzed at a more mature stage to detect changes in anatomy and in neural patterning. This simple but useful protocol allows for real time imaging. It provides a means to make spatially and temporally limited changes to endogenous protein levels.

A variety of growth factors and proteins interact to induce cells to take on different cell fates during development. Here we demonstrate the use of an in ovo preparation to address possible interactions between different proteins in development by placing beads on E2.5 chick embryos.

The neural tube expresses many proteins in specific spatiotemporal patterns during development.   These proteins have been shown to be critical for cell ...

Large Scale Zebrafish-Based In vivo Small Molecule Screen

Given their small embryo size, rapid development, transparency, fecundity, and numerous molecular, morphological and physiological similarities to mammals, zebrafish has emerged as a powerful in vivo platform for phenotype-based drug screens and chemical genetic analysis. Here, we demonstrate a simple, practical method for large-scale screening of small molecules using zebrafish embryos.

Large Scale Zebrafish-Based In vivo Small Molecule Screen

Zebrafish has emerged as a powerful in vivo platform for phenotype-based drug screens and chemical genetic analysis. Here, we demonstrate a simple, practical method for large-scale screening of small molecules using zebrafish embryos.

Given their small embryo size, rapid development, transparency, fecundity, and numerous molecular, morphological and physiological similarities to ...

Efficient Derivation of Human Cardiac Precursors and Cardiomyocytes from Pluripotent Human Embryonic Stem Cells with Small Molecule Induction

To date, the lack of a suitable human cardiac cell source has been the major setback in regenerating the human myocardium, either by cell-based transplantation or by cardiac tissue engineering1-3. Cardiomyocytes become terminally-differentiated soon after birth and lose their ability to proliferate. There is no evidence that stem/progenitor cells derived from other sources, such as the bone marrow or the cord blood, are able to give rise to the contractile heart muscle cells following transplantation into the heart1-3. The need to regenerate or repair the damaged heart muscle has not been met by adult stem cell therapy, either endogenous or via cell delivery1-3. The genetically stable human embryonic stem cells (hESCs) have unlimited expansion ability and unrestricted plasticity, proffering a pluripotent reservoir for in vitro derivation of large supplies of human somatic cells that are restricted to the lineage in need of repair and regeneration4,5. Due to the prevalence of cardiovascular disease worldwide and acute shortage of donor organs, there is intense interest in developing hESC-based therapies as an alternative approach. However, how to channel the wide differentiation potential of pluripotent hESCs efficiently and predictably to a desired phenotype has been a major challenge for both developmental study and clinical translation. Conventional approaches rely on multi-lineage inclination of pluripotent cells through spontaneous germ layer differentiation, resulting in inefficient and uncontrollable lineage-commitment that is often followed by phenotypic heterogeneity and instability, hence, a high risk of tumorigenicity6-8 (see a schematic in Fig. 1A). In addition, undefined foreign/animal biological supplements and/or feeders that have typically been used for the isolation, expansion, and differentiation of hESCs may make direct use of such cell-specialized grafts in patients problematic9-11. To overcome these obstacles, we have resolved the elements of a defined culture system necessary and sufficient for sustaining the epiblast pluripotence of hESCs, serving as a platform for de novo derivation of clinically-suitable hESCs and effectively directing such hESCs uniformly towards clinically-relevant lineages by small molecules12 (see a schematic in Fig. 1B). After screening a variety of small molecules and growth factors, we found that such defined conditions rendered nicotinamide (NAM) sufficient to induce the specification of cardiomesoderm direct from pluripotent hESCs that further progressed to cardioblasts that generated human beating cardiomyocytes with high efficiency (Fig. 2). We defined conditions for induction of cardioblasts direct from pluripotent hESCs without an intervening multi-lineage embryoid body stage, enabling well-controlled efficient derivation of a large supply of human cardiac cells across the spectrum of developmental stages for cell-based therapeutics.

We have established a protocol for induction of cardioblasts direct from pluripotent human embryonic stem cells maintained under defined conditions with small molecules, which enables derivation of a large supply of human cardiac progenitors and functional cardiomyocytes for cardiovascular repair.

To date, the lack of a suitable human cardiac cell source has been the major setback in regenerating the human myocardium, either by cell-based ...

High-throughput Screening for Small-molecule Modulators of Inward Rectifier Potassium Channels

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, atrial fibrillation, and pain1,2. For the most part, however, progress toward understanding their therapeutic potential or even basic physiological functions has been slowed by the lack of good pharmacological tools. Indeed, the molecular pharmacology of the inward rectifier family has lagged far behind that of the S4 superfamily of voltage-gated potassium (Kv) channels, for which a number of nanomolar-affinity and highly selective peptide toxin modulators have been discovered3. The bee venom toxin tertiapin and its derivatives are potent inhibitors of Kir1.1 and Kir3 channels4,5, but peptides are of limited use therapeutically as well as experimentally due to their antigenic properties and poor bioavailability, metabolic stability and tissue penetrance. The development of potent and selective small-molecule probes with improved pharmacological properties will be a key to fully understanding the physiology and therapeutic potential of Kir channels.
The Molecular Libraries Probes Production Center Network (MLPCN) supported by the National Institutes of Health (NIH) Common Fund has created opportunities for academic scientists to initiate probe discovery campaigns for molecular targets and signaling pathways in need of better pharmacology6. The MLPCN provides researchers access to industry-scale screening centers and medicinal chemistry and informatics support to develop small-molecule probes to elucidate the function of genes and gene networks. The critical step in gaining entry to the MLPCN is the development of a robust target- or pathway-specific assay that is amenable for high-throughput screening (HTS).
Here, we describe how to develop a fluorescence-based thallium (Tl+) flux assay of Kir channel function for high-throughput compound screening7,8,9,10.The assay is based on the permeability of the K+ channel pore to the K+ congener Tl+. A commercially available fluorescent Tl+ reporter dye is used to detect transmembrane flux of Tl+ through the pore. There are at least three commercially available dyes that are suitable for Tl+ flux assays: BTC, FluoZin-2, and FluxOR7,8. This protocol describes assay development using FluoZin-2. Although originally developed and marketed as a zinc indicator, FluoZin-2 exhibits a robust and dose-dependent increase in fluorescence emission upon Tl+ binding. We began working with FluoZin-2 before FluxOR was available7,8 and have continued to do so9,10. However, the steps in assay development are essentially identical for all three dyes, and users should determine which dye is most appropriate for their specific needs. We also discuss the assay's performance benchmarks that must be reached to be considered for entry to the MLPCN. Since Tl+ readily permeates most K+ channels, the assay should be adaptable to most K+ channel targets.

Methods for developing and validating a quantitative fluorescence assay for measuring the activity of inward rectifier potassium (Kir) channels for high-throughput compound screening is presented.

Specific members of the inward rectifier potassium (Kir) channel family are postulated drug targets for a variety of disorders, including hypertension, ...

Microgavage of Zebrafish Larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

Efficient and Rapid Isolation of Early-stage Embryos from Arabidopsis thaliana Seeds

In flowering plants, the embryo develops within a nourishing tissue - the endosperm - surrounded by the maternal seed integuments (or seed coat). As a consequence, the isolation of plant embryos at early stages (1 cell to globular stage) is technically challenging due to their relative inaccessibility. Efficient manual dissection at early stages is strongly impaired by the small size of young Arabidopsis seeds and the adhesiveness of the embryo to the surrounding tissues. Here, we describe a method that allows the efficient isolation of young Arabidopsis embryos, yielding up to 40 embryos in 1 hr to 4 hr, depending on the downstream application. Embryos are released into isolation buffer by slightly crushing 250-750 seeds with a plastic pestle in an Eppendorf tube. A glass microcapillary attached to either a standard laboratory pipette (via a rubber tube) or a hydraulically controlled microinjector is used to collect embryos from droplets placed on a multi-well slide on an inverted light microscope. The technical skills required are simple and easily transferable, and the basic setup does not require costly equipment. Collected embryos are suitable for a variety of downstream applications such as RT-PCR, RNA sequencing, DNA methylation analyses, fluorescence in situ hybridization (FISH), immunostaining, and reporter gene assays.

Efficient and Rapid Isolation of Early-stage Embryos from Arabidopsis thaliana Seeds

We report an efficient and simple method to isolate embryos at early stages of development from Arabidopsis thaliana seeds. Up to 40 embryos can be isolated in 1 hr to 4 hr, depending on the downstream application. The procedure is suitable for transcriptome, DNA methylation, reporter gene expression, immunostaining and fluorescence in situ hybridization analyses.

In flowering plants, the embryo develops within a nourishing tissue - the endosperm - surrounded by the maternal seed integuments (or seed coat). As a ...

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional measures of muscle health. In this video article, we demonstrate a method to measure force generation during contraction of zebrafish larval trunk muscle. Force measurements are accomplished by placing an anesthetized larva into a chamber filled with a salt solution. The anterior end of the larva is tied to a force transducer and the posterior end of the larva is tied to a length controller. An isometric twitch contraction is elicited by electric field stimulation and the force response is recorded for analysis. Force generation during contraction provides a measure of overall muscle health and specifically provides a measure of muscle function. Although we describe this technique for use with wild-type larvae, this method can be used with genetically modified larvae or with larvae treated with drugs or toxicants, to characterize muscle disease models and evaluate treatments, or to study muscle development, injury, or chemical toxicity.

Force measurements can be used to demonstrate changes in muscle function due to development, injury, disease, treatment or chemical toxicity. In this video, we demonstrate a method to measure force during a maximal contraction of zebrafish larval trunk muscle.

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional ...

A Manual Small Molecule Screen Approaching High-throughput Using Zebrafish Embryos

Zebrafish have become a widely used model organism to investigate the mechanisms that underlie developmental biology and to study human disease pathology due to their considerable degree of genetic conservation with humans. Chemical genetics entails testing the effect that small molecules have on a biological process and is becoming a popular translational research method to identify therapeutic compounds. Zebrafish are specifically appealing to use for chemical genetics because of their ability to produce large clutches of transparent embryos, which are externally fertilized. Furthermore, zebrafish embryos can be easily drug treated by the simple addition of a compound to the embryo media. Using whole-mount in situ hybridization (WISH), mRNA expression can be clearly visualized within zebrafish embryos. Together, using chemical genetics and WISH, the zebrafish becomes a potent whole organism context in which to determine the cellular and physiological effects of small molecules. Innovative advances have been made in technologies that utilize machine-based screening procedures, however for many labs such options are not accessible or remain cost-prohibitive. The protocol described here explains how to execute a manual high-throughput chemical genetic screen that requires basic resources and can be accomplished by a single individual or small team in an efficient period of time. Thus, this protocol provides a feasible strategy that can be implemented by research groups to perform chemical genetics in zebrafish, which can be useful for gaining fundamental insights into developmental processes, disease mechanisms, and to identify novel compounds and signaling pathways that have medically relevant applications.

The zebrafish is an excellent experimental organism to study vertebrate developmental processes and model human disease. Here, we describe a protocol on how to perform a manual high-throughput chemical screen in zebrafish embryos with a whole-mount in situ hybridization (WISH) read-out.

Zebrafish have become a widely used model organism to investigate the mechanisms that underlie developmental biology and to study human disease pathology ...

Environmental Screening of Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa in Zebrafish Systems

Health monitoring systems are developed and used in zebrafish research facilities because pathogens of Danio rerio such as Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa have the potential to impair animal welfare and research. The fish are typically analyzed post mortem to detect microbes. The use of sentinels is a suggested way to improve the sensitivity of the surveillance and to reduce the number of animals to sample. The setting of a pre-filtration sentinel tank out of a recirculating system is described. The technique is developed to prevent water pollution and to represent the fish population by a careful selection of age, gender, and strains. In order to use the minimum number of animals, techniques to screen the environment are also detailed. Polymerase Chain Reaction (PCR) on surface sump swabs is used to significantly improve the detection of some prevalent and pathogenic mycobacterial species such as Mycobacterium fortuitum, Mycobacterium haemophilum, and Mycobacterium chelonae. Another environmental method consists of processing the sludge at the bottom of a holding tank or sump to look for P. tomentosa eggs. This is a cheap and fast technique that can be applied in quarantine where a breeding device is submerged into the holding tank of imported animals. Finally, PCR is applied to the sludge sample and A. hydrophila is detected at the sump&#39;s bottom and surface. Generally, these environmental screening techniques applied to these specific pathogens have led to an increased sensitivity compared to the testing of pre-filtration sentinels.

Environmental Screening of Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa in Zebrafish Systems

This protocol describes the use of sump swabs and sludge analysis of zebrafish systems, which leads to increased detection compared to the sole use of sentinels to detect pathogens such as Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa. A system to monitor P. tomentosa eggs in quarantine is also proposed.

Health monitoring systems are developed and used in zebrafish research facilities because pathogens of Danio rerio such as Aeromonas hydrophila, ...

Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such as Zika, dengue and malaria. Assays for rapid, high-throughput analyses of unformulated novel chemistries against mosquito larvae and adults are presented. We describe protocols for single point-dose and dose response assays to evaluate the toxicity of small molecule chemistries to the Aedes aegypti vector of Zika, dengue and yellow fever,&#160;the malaria vector, Anopheles gambiae and the northern house mosquito, Culex quinquefasciatus, on contact and via ingestion. As an example, we evaluated the toxicity of amitriptyline, a small molecule antagonist of G protein-coupled receptors, via larval, adult topical and adult blood-feeding assay. The protocols provide a starting point to investigate insecticide potential. Results are discussed in the context of additional experiments to explore product applications and mechanisms for delivery.

Protocols are described for assessing the toxicity of chemistries to immature and adult mosquitoes for development as new classes of larvicides, adulticides and endectocides. The protocols enable high-throughput testing of multiple chemistries at single-point dose and subsequent evaluation via dose response assay to determine toxicity on contact or via ingestion.

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such ...