The authors thank Hanayo Arimoto (US Navy), Dana Johnson, Lucy Li, and Roxie White (USDA-ARS) for help with data acquisition and analysis. We also thank Drs. Pia Olafson (USDA-ARS) and Ke Wu (University of Florida) for critical review of the manuscript and Niklaus Hostettler (University of Florida) for filming the microscope footage. Funding was provided by the USDA, the WII-141C project of the Navy and Marine Corps Public Health Center, and the Deployed War Fighters Protection Program. The funders had no role in design or direction of the study or development of the manuscript.

1. Insect Preparation
<ol>
	<li>Anesthetize mosquitoes by chilling them at 4 &#176;C for several hours or by 2 min exposure to CO2 and then holding at 4 &#176;C prior to injections. Cold anesthetize flies at 4 &#176;C 2&#8210;3 h prior to injections. Ensure mosquitoes and flies are mated if oviposition is a response variable. 
	NOTE: The mosquito anesthetization method depends upon the species and strain, e.g., Culex recover much quicker from cold anesthetization than Aedes aegypti and therefore CO2 knockdown is preferable.</li>
	<li>On ice, carefully stage mosquitoes or flies by using soft forceps to lay them ventrally exposed on microscope slides ~0.5 cm apart. Standard slides can hold two rows of 6 mosquitoes each or 6&#8211;7 house flies. Leave a 1&#8211;1.5 cm space at the left or right end of the slide clear of mosquitoes or flies to aid slide handling.</li>
	<li>Keep slides of staged insects at 4 &#176;C in large Petri dishes until ready for injection. 
	NOTE: Drilling a hole through the Petri dish lid will help prevent insects from being disturbed by air displacement when capping. Placing slides on a couple of glass capillaries secured to the dish bottom will ease removal from Petri dish.</li>
</ol>
2. Insect Injection
<ol>
	<li>Prepare dsRNA as described by Estep et al.5 and Sanscrainte et al.9. If free of contaminants, dsRNA can be quantified by using a spectrophotometer to measure the absorbance at 260 nm and calculating RNA concentration in &#956;g/mL as A260 &#215; dilution factor &#215; 40.</li>
	<li>Set up dissecting microscope and microinjector over a chill table or shallow ice bucket to perform injections on the cold-anesthetized mosquitoes and flies (see step 1.1&#8211;1.3 and Figure 1).</li>
	<li>Pull glass capillaries to a fine tip with needle puller (settings: heater = 15 units, solenoid = 4 A). 
	NOTE: See Table of Materials for manufacturer&#8217;s details.</li>
	<li>Place the capillary needle into a microinjector as per the manufacturer&#8217;s instructions and break needle tip by pinching with a pair of forceps to produce a sharp point. 
	NOTE: Suggested needle opening sizes are ~150 &#181;m for mosquito and ~250 &#181;m for housefly.</li>
	<li>Set microinjector for desired delivery volume. Meniscus movement from ~50 nL aliquots is easily observable and recommended for mosquito microinjections. If available, enable fast injection settings. Rinse the needle by filling and expelling nuclease-free water 3 times.</li>
	<li>Prepare solutions for injection at a concentration such that 100&#8210;150 nL will provide desired dose for mosquitoes and 500 nL will provide desired dose for house flies. 
	NOTE: Suggested initial doses: 1 &#181;g for each mosquito and 5 &#181;g for each housefly5,9.</li>
	<li>Optionally for mosquitoes, add Rhodamine B (to aid in visualization) to solutions at a final concentration of 3.0 &#181;g/mL. 
	NOTE: The dye will be clearly visible by its pink color after injection through the nearly clear cuticle on the ventral surface at the abdomen/thorax intersection.</li>
	<li>Pipette 3&#8211;4 &#181;L of solution to inject onto a clean surface (such as a piece of paraffin film) and draw into the glass needle without sucking in any air. Depress the inject button repeatedly until liquid begins to dispense from the needle and gently wipe droplets off with a delicate task wiper. 
	NOTE: While some models of microinjectors require backfilling the syringe, the injector referenced in the Table of Materials does not.</li>
	<li>Using an ultra-fine point marker, draw hash marks ~1 mm apart starting from the liquid meniscus to the needle shank. This will aid in visualizing the meniscus movement and ensure that the solution has entered the mosquito during injection.</li>
	<li>Set the slide of mosquitoes or flies (as prepared in steps 1.2&#8211;1.3) under the needle on the chill table or ice (Figure 1). Ensure that field of view in the microscope is wide enough to see the solution meniscus in the needle.</li>
	<li>For mosquitoes, align the needle with the middle one-third of the mesokatepisternum (Figure 2A), and while bracing the mosquito against the needle with forceps placed on the opposite side, gently puncture the cuticle with the needle tip using the microinjector micrometer.</li>
	<li>For house flies, align the needle with the mesopleuron (Figure 2B) and, while bracing the fly against the needle, gently puncture the cuticle with the needle tip.</li>
	<li>Gently slide the needle into the mosquito or fly body until the tip has passed through the midline. 
	NOTE: Inserting the needle too shallow often results in the injected liquid beading out of the insect (see step 2.15).</li>
	<li>While watching for movement of the liquid meniscus in the needle, depress the inject button until the desired amount of liquid has been injected. For mosquitoes, if set to 50.6 nL aliquots, press 2X for ~100 nL or 3X for ~150 nL. For house flies, depress the inject button until ~500 nL has been injected (i.e., with 69 nL aliquots press 7X for 483 nL).</li>
	<li>If the meniscus does not move, slowly slide the mosquito or fly off the needle while watching for meniscus movement. If a portion of the injected solution beads out of the cuticle upon needle removal or if the meniscus fails to move, discard the insect as this injection was not successful.</li>
	<li>Transfer successfully injected mosquitoes in groups of 10-15 or house flies in groups of 5 to clear 3.5 oz holding cups. Cover with tulle or netting and recover at room temperature.</li>
	<li>After mosquitoes and flies have recovered (1 &#8211; 2 h), invert each holding cup over a cotton ball soaked with 10% sucrose solution. 
	NOTE: The inversion of the cup on top of the sucrose cotton aids in survival by providing easier access to a sugar meal10.</li>
	<li>Rinse needle as in step 2.5 between injection solutions. When done injecting, discard used needles in an appropriate sharps container.</li>
</ol>
3. Aedes aegypti mortality and oviposition bioassay
<ol>
	<li>For three days after injection, continue to provide mosquitoes with 10% sucrose and record the number of visibly dead mosquitoes.</li>
	<li>Fill a &#8804;12-inch artificial membrane (such as collagen sausage casing) with fresh blood (i.e., bovine for Aedes aegypti) and heat to 60&#160;&#176;C in hot water bath.</li>
	<li>Briefly dry the membrane by rolling on a paper towel and lay across the tulle caps of the 3.5 oz clear holding cups. To enhance feeding, spike blood with 1 mM ATP after heating as a phagostimulant or by handle the warmed blood sausage with clean bare hands to leave human volatiles on the casing surface.</li>
	<li>After blood feeding, replace the cotton ball soaked with 10% sucrose on top of the holding cups and allow mosquitoes to rest for ~24 h before transferring to oviposition cups.</li>
	<li>Construct oviposition cups by filling clear 3.5 oz bioassay cups with ~30 mL deionized H2O and placing a piece of seed germination paper (~4 cm wide and 5 cm long with ridges running vertically) at the bottom of the cup and against the side (Figure 3A). Cover with tulle or netting and cut a small slit (~1 cm) in the tulle or netting cap.</li>
	<li>At 24 h post-blood feeding, cut a small slit (~1 cm) in the tulle cap of the holding cup and transfer only individual females that successfully fed &#8212; with a visible blood bolus in abdomen &#8212; by gentle mouth aspiration into oviposition cups (1 female/oviposition cup). Seal the small cut in the oviposition cap with a 10% sucrose saturated cotton ball.</li>
	<li>Hold cups at ~27 &#176;C for 5&#8210;7 days to allow mosquitoes to fully oviposit while tracking daily mortality. Count eggs under a dissecting scope.</li>
</ol>
4. Musca domestica mortality and oviposition bioassay
<ol>
	<li>For three days after injection, continue to provide the flies with 10% sucrose and record the number of visibly dead flies.</li>
	<li>Three days after injection, anesthetize flies by briefly placing a polyethylene tube dispensing CO2 over the holding cups until all flies are motionless at bottom of the cups.</li>
	<li>Transfer flies to a clean cage comprised of a 4 L plastic jar with a stockinette sleeve covering the opening (10 cm, Figure 3B). Provide flies with water and a mixture of granulated sucrose, powdered milk, and dried egg yolk in an 8:8:1 ratio (by volume) for four days, recording and removing dead flies daily.</li>
	<li>Prepare dry ingredients for fresh fly larval rearing medium by mixing 75% wheat bran with 25% pelleted livestock feed by weight. Add water to reach 62% moisture (until a drop of water can barely be squeezed out).</li>
	<li>Prepare a 2.5 cm diameter ball from a 1:1 mixture of the above fresh larval media and &#8220;used&#8221; fly larval medium (medium in which flies have previously pupated). Wrap the ball in a square of black cotton cloth, squeeze lightly until medium liquid seeps through, then use rubber bands to hold the cloth in place around the ball.</li>
	<li>Put the ball in a 60 mL cup and place it in the fly cage for 5 h (Figure 3B).</li>
	<li>Rinse eggs off the oviposition ball and ensure that the eggs are removed from under folds in the cloth. Shake eggs to disrupt clusters and transfer them to a graduated 20 mL centrifuge tube with a transfer pipet.</li>
	<li>Wait until the eggs settle and note the volume of the settled eggs in the graduated tube. Add sufficient water to bring the volume up to 20X the volume of settled eggs. Record the total volume of water plus eggs.</li>
	<li>While mixing the water and egg suspension with either a magnetic stir bar or vigorous stirring, use a 1 mL pipet tip (with the end of the tip cut off) to dispense 0.5 mL of the water and egg suspension onto a piece of pre-moistened black cloth in a series of lines. Count the eggs under a dissecting microscope.</li>
	<li>Repeat step 4.9 two more times. If one of the counts is a severe outlier (i.e., differs from the other two counts by &#62;35%), make a fourth count and disregard the outlier.</li>
	<li>Calculate the mean number of eggs per sample and multiply this value by 2 to get the number of eggs/mL of the suspension. Multiply the eggs/mL value obtained by the volume of the egg suspension from step 4.8. This is the estimate for the total number of eggs laid.</li>
</ol>

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R., 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=Gene+silencing+in+adult+Aedes+aegypti+mosquitoes+through+oral+delivery+of+double-stranded+RNA.">Gene silencing in adult Aedes aegypti mosquitoes through oral delivery of double-stranded RNA.</a> Journal of Applied Entomology. 136 (10), 741-748 (2012).</li><li>Estep, A. S., Sanscrainte, N. D., Becnel, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DsRNA-mediated+targeting+of+ribosomal+transcripts+RPS6+and+RPL26+induces+long-lasting+and+significant+reductions+in+fecundity+of+the+vector+Aedes+aegypti.">DsRNA-mediated targeting of ribosomal transcripts RPS6 and RPL26 induces long-lasting and significant reductions in fecundity of the vector Aedes aegypti.</a> Journal of Insect Physiology. 90, 17-26 (2016).</li><li>Wen, D., 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=N-glycosylation+of+Viral+E+Protein+Is+the+Determinant+for+Vector+Midgut+Invasion+by+Flaviviruses.">N-glycosylation of Viral E Protein Is the Determinant for Vector Midgut Invasion by Flaviviruses.</a> mBio. 9 (1), (2018).</li><li>Liu, Y., Cheng, G., Colpitts, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Techniques+for+Experimental+Infection+of+Mosquitoes+with+West+Nile+Virus.">Techniques for Experimental Infection of Mosquitoes with West Nile Virus.</a> Methods in Molecular Biology. 1435, 151-163 (2016).</li><li>Puglise, J. 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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Defects+in+coatomer+protein+I+(COPI)+transport+cause+blood+feeding-induced+mortality+in+Yellow+Fever+mosquitoes.">Defects in coatomer protein I (COPI) transport cause blood feeding-induced mortality in Yellow Fever mosquitoes.</a> Proceedings of the National Academy of Sciences. 108 (24), E211-E217 (2011).</li><li>Drake, L. L., Price, D. P., Aguirre, S. E., Hansen, I. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNAi-mediated+gene+knockdown+and+in+vivo+diuresis+assay+in+adult+female+Aedes+aegypti+mosquitoes.">RNAi-mediated gene knockdown and in vivo diuresis assay in adult female Aedes aegypti mosquitoes.</a> Journal of Visualized Experiments: JoVE. 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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+interference+directed+against+ribosomal+protein+S3a+suggests+a+link+between+this+gene+and+arrested+ovarian+development+during+adult+diapause+in+Culex+pipiens.">RNA interference directed against ribosomal protein S3a suggests a link between this gene and arrested ovarian development during adult diapause in Culex pipiens.</a> Insect molecular biology. 19 (1), 27-33 (2010).</li><li>Atyame, C. M., 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=Comparison+of+irradiation+and+Wolbachia+based+approaches+for+sterile-male+strategies+targeting+Aedes+albopictus.">Comparison of irradiation and Wolbachia based approaches for sterile-male strategies targeting Aedes albopictus.</a> PLoS one. 11 (1), e0146834 (2016).</li><li>Becnel, J. J., Floore, T. 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All authors are employees or contractors of the United States Department of Agriculture or the Department of Defense. This manuscript was produced during the course of official duties and is by law in the public domain. The views expressed represent the opinion of the authors and do not represent an official position or endorsement by the US government. Mention of a commercial product is not a recommendation for use.

Microinjection is a valuable laboratory technique to ensure delivery of dsRNA or other biorationals (i.e., pesticides, viruses, microsporidia). While many laboratories perform microinjection, the exact amount injected is often unclear due to technical limitations of the delivery system where delivered volume is not directly measured11. Delivered volume and concentration are critical parameters that allow calculation of standard toxicological measures such as EC50 or IC50 or to define minimal effective doses5,16. This is especially important in functional studies using RNAi in adult insects, where delivery by feeding does not always result in systemic exposure to the desired particles and the actual dose crossing the midgut is unclear5.
While the microinjection procedure can be initially challenging, rapid improvements in speed and delivery accuracy are achieved with practice and patience. Mastering the injection procedure itself is a time investment, but, once accomplished, the procedure produces repeatable results and injury mortality of &#60;3%. The ratio of successful injections will increase as proficiency increases allowing injection of 300 mosquitoes or flies per day.
Many of the challenges of microinjection are due to the needle being used. Determining the proper capillary needle break point and tip angle is a challenging aspect of the procedure and requires some trial and error to determine the most effective opening size for a given species. The fine needle most useful for mosquito microinjection (~150 &#181;m) is too small to quickly deliver the larger volume injected into flies while conversely, the larger needle opening that works on flies (~250 &#181;m) causes extensive injury damage to the smaller mosquitoes and unacceptable injection mortality. A needle broken with a blunt tip is often difficult to get through the cuticle without tissue damage and increased mortality. Insect scales or tissue pieces can clog needles over the course of an experiment, so it is often helpful to have several needles pulled if replacement is required. We have found that it is easier to replace a difficult needle rather than spend time to try to clear a jammed or fouled needle.
Observing the injection solution entering the insect is critical so that unsuccessful injections can be removed (see steps 2.14-2.15). Rhodamine B addition to injection solutions provides a clear visual to ensure that cold-staged insects receive proper dosing and is especially helpful while practicing (see step 2.7).
The injection and oviposition bioassay methods presented here have successfully induced gene knockdown and measurable phenotypic effects in adult Ae. aegypti and M. domestica when using dsRNAs designed against species-specific ribosomal transcripts (Figure 4 and Figure 5A). Additionally, the ability of this method to accurately deliver microvolumes to individuals allows for dose response curves to be generated for small amounts of material (as low as 50 ng; Figure 5B). This is especially useful for methods such as screening siRNAs or dsRNAs, as producing large quantities of these molecules can be cost-prohibitive.
This method can be modified to inject biological agents (viruses, bacteria, microsporidia) or chemical pesticides, after ensuring that any delivery buffers are innocuous by injection. As delivery volume is limited by the size of the insect, producing concentrated testing solutions is of importance.
To adequately evaluate dsRNA efficacy, it is necessary to track oviposition as lethal phenotypes may only manifest in progeny. As presented here, holding female Ae. aegypti separately after blooding allows for individual clutch size determination and further testing on the same individuals can be done downstream of oviposition (i.e., gene expression studies, additional gonotrophic cycles, tissue specific knockdown) to correlate reproductive output with other measures such as gene expression. As M. domestica generally oviposit in a group response, inciting isolated gravid flies to lay is very labor intensive and not practical for large numbers of individuals17. Therefore, a method to determine mean clutch size is presented.
The microinjection method presented here provides consistent systemic delivery to mosquitoes and house flies. Coupled with mortality and oviposition tracking bioassays, these versatile tools can be used to assess the transcriptional and phenotypic effects of small RNA molecules, biological agents, and traditional pesticides where oral delivery is not viable.

Delivering small biomolecules, such as nucleic acids, to adult dipterans has proven to be challenging in both Culicidae and Muscidae. Oral uptake of dsRNAs has been reported to produce sterility (when targeting genes exclusively expressed in testes of adults) and mortality (when targeting SNF7 and a steroid receptor coactivator) in larval Aedes aegypti1,2. Mortality has also been observed when targeting HSP70 in larval Musca domestica3. Such phenotypic effects, however, have not resulted after feeding dsRNA to adult Ae. aegypti in sugar meals4,5.
Microinjection has been used to circumvent the midgut when introducing pathogens or nucleic acids, thereby inducing a systemic response5,6,7,8,9,10. Several microinjection techniques exist, involving equipment ranging from in-house produced apparatuses that require visual measurement of injection volume to microprocessor-controlled injectors that allow for automated volume delivery as low as 2.3 nL5,9,10,11. RNA interference (RNAi) triggers targeting ribosomal mRNAs, disrupt ovarian development in arthropods as diverse as the cattle tick Rhipicephalus microplus, the mosquitoes Aedes aegypti and Culex pipiens, and the house fly Musca domestica5,9,12,13. In these studies, monitoring the disruption of oviposition was essential for determining the efficacy of the inoculant as the phenotype may manifest as termination or reduction of progeny. This is a form of multigenerational lethal phenotype that is a critical and desired effect in many of the non-traditional biocontrol methods like Wolbachia-infected males introduction (sexual incompatibility) and RNAi induced sterility5,9,14. Tracking both mortality and fecundity is necessary for the characterizing and development of highly specific biorationals (naturally occurring pathogens and/or natural derivatives)15.
This protocol presents detailed microinjection techniques for both adult mosquitoes and house flies; a process that is often not well described in the literature. In addition, oviposition bioassay methodologies to adequately evaluate dsRNA effects on adult dipterans are described. These protocols were developed specifically for Aedes aegypti and Musca domestica but can be modified for other species.

<table>
	<tbody>
		<tr>
			<td>needle pulling</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>vertical pipette puller</td>
			<td>Kopf</td>
			<td>720</td>
			<td>settings: heater = 15 units, solenoid = 4 amps</td>
		</tr>
		<tr>
			<td>glass capillaries, 3.5&#34; long, ID = 0.530 mm &#177; 25 &#956;m, OD 1.14 mm</td>
			<td>World Precision Instruments</td>
			<td>504949</td>
			<td>capillaries for pulling glass needles</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>microinjector station</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nanoliter 2010</td>
			<td>World Precision Instruments</td>
			<td>NANOLITER2010</td>
			<td>microinjector</td>
		</tr>
		<tr>
			<td>manual micromanipulator</td>
			<td>World Precision Instruments</td>
			<td>KITE-L</td>
			<td>left hand KITE manipulator</td>
		</tr>
		<tr>
			<td>magnetic stand</td>
			<td>World Precision Instruments</td>
			<td>M9</td>
			<td>holds micromanipulator</td>
		</tr>
		<tr>
			<td>precision stereo zoom binocular microscope on boom stand</td>
			<td>World Precision Instruments</td>
			<td>PZMIII-BS</td>
			<td>dissecting scope for microinjections</td>
		</tr>
		<tr>
			<td>1.5X objective</td>
			<td>World Precision Instruments</td>
			<td>501377</td>
			<td>objective for microscope</td>
		</tr>
		<tr>
			<td>light LED ring</td>
			<td>World Precision Instruments</td>
			<td>504134</td>
			<td>light ring for injection microscope</td>
		</tr>
		<tr>
			<td>laboratory chill table</td>
			<td>BioQuip Products</td>
			<td>1431</td>
			<td>chill table for microinjections</td>
		</tr>
		<tr>
			<td>microscope slides</td>
			<td>Fisher Scientific</td>
			<td>12-544-1</td>
			<td>for staging insects while injections</td>
		</tr>
		<tr>
			<td>Rhodamine B, 98+%</td>
			<td>Acros Organics</td>
			<td>AC296571000</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>mosquito oviposition bioassays</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>large Petri dish</td>
			<td>Fisher Scientific</td>
			<td>FB0875714</td>
			<td>for holding staging slides</td>
		</tr>
		<tr>
			<td>plastic cup - 3 1/2 oz.</td>
			<td>Dart Container Corporation</td>
			<td>TK35</td>
			<td>mosquito oviposition bioassay cups</td>
		</tr>
		<tr>
			<td>matte tulle fabric</td>
			<td>Joanne Fabrics</td>
			<td>1103068</td>
			<td>caps for oviposition bioassays</td>
		</tr>
		<tr>
			<td>blood source</td>
			<td>locally acquired</td>
			<td></td>
			<td>typically bovine or live chickens</td>
		</tr>
		<tr>
			<td>1 x 30mm clear edible collagen casing</td>
			<td>Butcher and Packer</td>
			<td>30D02-05</td>
			<td></td>
		</tr>
		<tr>
			<td>heavy weight seed germination paper</td>
			<td>Anchor Paper Co</td>
			<td>SD7615L</td>
			<td></td>
		</tr>
		<tr>
			<td>oral aspirator with HEPA filter</td>
			<td>John W. Hock Company</td>
			<td>612</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>house fly oviposition bioassays</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4 L plastic food storage canister</td>
			<td>Walmart</td>
			<td>555115143</td>
			<td></td>
		</tr>
		<tr>
			<td>10-inch stockinette sleeve</td>
			<td>Medonthego.com</td>
			<td>FS15001H</td>
			<td></td>
		</tr>
		<tr>
			<td>wheat bran</td>
			<td>locally acquired</td>
			<td></td>
			<td>typically found at feed stores</td>
		</tr>
		<tr>
			<td>Calf Manna performance supplement</td>
			<td>ValleyVetSupply.com</td>
			<td>16731</td>
			<td>pelleted livestock feed</td>
		</tr>
		<tr>
			<td>dried egg yolk</td>
			<td>BulkFoods.com</td>
			<td>40506</td>
			<td></td>
		</tr>
		<tr>
			<td>black cotton cloth</td>
			<td>locally acquired</td>
			<td></td>
			<td>typically in craft supplies section</td>
		</tr>
		<tr>
			<td>60 mL cup</td>
			<td>Dart Container Corporation</td>
			<td>P200-N</td>
			<td></td>
		</tr>
	</tbody>
</table>

reproducible dsrna microinjection and oviposition bioassay in mosquitoes and house flies

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are delivered using a non-quantitative method. Accurate delivery of known quantities of nucleic acids or other chemicals is critical to measure the efficacy of the compound being tested and to allow reliable comparison between compounds.
Here we provide a reproducible, quantitative microinjection protocol that ensures accurate delivery of specific doses of dsRNA, reducing the mortality typically induced by injection injury. These modifications include the addition of Rhodamine B, a graduated injection needle, and an improved recovery method borrowed from Isoe and Collins. This method allows calculation of dose responses and facilitates comparisons between compounds. Versions of this method have been successfully used on three genera of mosquitoes as well as house flies to assess the reduction in fecundity resulting from gene silencing of ribosomal RNA transcripts.
This protocol provides strategies to reduce several challenges of small insect microinjection. Together, mechanical delivery of dsRNAs accompanied by visual verification, identification of effective locations for delivery, and inclusion of a post-injection recovery period ensure accurate dosing and low injury mortality. This protocol also describes an oviposition bioassay for uniform determination of effects on fecundity.

Microinjection of dsRNA in mosquitoes
This microinjection method has been used in our laboratory to evaluate gene expression, mortality and oviposition response for over 80 dsRNA triggers across several mosquito genera (Aedes, Anopheles, Culex). Injecting females from the colony strain of Ae. aegypti (ORL1952) with 1 &#181;g of dsRNA derived from the ribosomal transcripts RPS6 and RPL26 (see Estep et al.5 for dsRNA production methods and sequence data) showed significant reduction in relative expression (RE) across multiple oviposition cycles when compared to mosquitoes injected with a control dsGFP. Aedes aegypti RE measured following the 2nd gonotrophic cycle showed significant reduction of RPS6 expression (P &#60; 0.05) in mosquitoes injected with dsRPS6 as determined by a Student&#39;s t-test between the dsGFP control of the same group (Figure 4)5.
Fecundity was assessed from individual females using the oviposition bioassay described here. After 3 days post-injection, mosquitoes were blood fed, transferred into individual oviposition containers, and allowed to lay to completion. Average clutch sizes for the first gonotrophic cycle were significantly reduced for both dsRPS6 treated Ae. aegypti (n = 102 females, 1.3 &#177; 0.8 (mean &#177; SE) eggs) and dsRPL26 treated (n = 86 females, 4.0 &#177; 1.1 eggs) when compared to dsGFP injections (n = 79 females, 49.3 &#177; 3.5 eggs, Figure 5A). Clutch assessment after a second gonotrophic cycle also showed significantly reduced oviposition for both dsRNA treated groups, but with reduced effect size. Group sizes of Ae. aegypti were variable and thus means separation was performed by Dunn&#39;s test5. Twenty-four-hour mortality was normally 3% or less.
A clear dose effect was observed when injecting dsRPS6 from 1 &#181;g to 50 ng in female Ae. aegypti, and clutch size varied significantly when compared to 1 &#181;g/female dsGFP injected controls in all but the 25 ng/female dsRPS6 dose (Figure 5B).
Microinjection of dsRNA in house flies
Colony Musca domestica (ORL normal) were injected with 5 &#181;g of dsRNA constructs designed against the house fly RPS6 and RPL26 transcripts9. As was observed in Ae. aegypti, significant reduction of both specific transcript expression (RPS6 and RPL26) and reduction in clutch size was observed (Figure 4 and Figure 5A). Significant reduction in RE was determined by Student&#39;s t-test between the dsGFP control of the same group (P &#60; 0.05) and the Holm-Sidak test was used to determine significance between clutch sizes for different M. domestica treatment groups. Ovarian dissections showed reduced oogenesis in dsRPS6 and dsRPL26 treatments, while dsGFP fed flies had normal vitellogenin deposition as rated on the Tyndale-Biscoe scale9.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58650/58650fig1.jpg" /> 
Figure 1: Microinjection setup for delivering microvolumes to adult mosquitoes and house flies. Injections are performed on cold-anesthetized insects staged on microscope slides over a chill table. <a href="https://www.jove.com/files/ftp_upload/58650/58650fig1large.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/58650/58650fig2.jpg" /> 
Figure 2: Microinjection sites for Ae. aegypti and M. domestica. (A) Adult Ae. aegypti are injected in the middle one-third of the mesokatepisternum. (B) Adult M. domestica are injected in the mesopleuron. Arrows indicate sites of injection. <a href="https://www.jove.com/files/ftp_upload/58650/58650fig2large.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/58650/58650fig3.jpg" /> 
Figure 3: Oviposition bioassay setup. (A) Ae. aegypti oviposition assay cups with seed germination paper as an oviposition substrate, capped with tulle and 10% sucrose-soaked cotton. (B) Adult female M. domestica in an oviposition assay container with food (A), water (B), and larval media (C). <a href="https://www.jove.com/files/ftp_upload/58650/58650fig3large.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/58650/58650fig4.jpg" /> 
Figure 4: Relative expression of RPS6 in Ae. aegypti and M. domestica injected with species-specific dsRNAs. Aedes aegypti and M. domestica were injected with 1 and 5 &#181;g respectively of either control dsRNA (dsGFP) or dsRNA designed against RPS6 or RPL26 transcripts from each insect. Significant reduction (P &#60; 0.05) of RPS6 expression was observed in mosquitoes and flies injected with dsRPS6 and by flies injected with dsRPL26 (indicated by asterisks). Error bars represent mean &#177; SE and number at column base indicates number of individuals examined. This figure has been modified from Estep et al.5 and Sanscrainte et al.9. <a href="https://www.jove.com/files/ftp_upload/58650/58650fig4large.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/58650/58650fig5.jpg" /> 
Figure 5: Average clutch size of Ae. aegypti and M. domestica injected with species-specific dsRNAs. (A) Egg deposition was significantly reduced (P &#60; 0.05) in both Ae. aegypti and M. domestica after injection of 1 and 5 &#181;g respectively of either control dsRNA (dsGFP) or dsRNA designed against RPS6 or RPL26 transcripts from each insect. Asterisks indicate significant differences from the dsGFP control of the same group. Error bars represent mean &#177; SE and number at column base indicates number of individuals examined. This figure has been modified from Estep et al.5 and Sanscrainte et al.9. (B) Dose curve of Ae. aegypti injected with dsRPS6 from 50 ng/female to 1000 ng/female shows significant reductions in fecundity in comparison to dsGFP injected cohorts (50, 100, and 1000 ng/female). Error bars represent mean &#177; 95% CI from 36-81 individual organisms per dose. Points with letters represent significant difference between groups (P &#60; 0.05) identified by ANOVA. This figure has been modified from Estep et al.5. <a href="https://www.jove.com/files/ftp_upload/58650/58650fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies at JoVE.com

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

This protocol describes a microinjection methodology that we have standardized and used for several years to deliver specific quantities of nucleic acids directly to the hemolymph of mosquitoes and house flies. This protocol results in minimal injection mortality and allows dose correlated measurements of fecundity.

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are ...

reproducible-dsrna-microinjection-oviposition-bioassay-mosquitoes

Research

JoVE Journal

Environment

8.7K Views.  USDA/ARS Center for Medical, Agricultural, and Veterinary Entomology. This protocol describes a microinjection methodology that we have standardized and used for several years to deliver specific quantities of nucleic acids directly to the hemolymph of mosquitoes and house flies. This protocol results in minimal injection mortality and allows dose correlated measurements of fecundity.

Video: Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Optimize Flue Gas Settings to Promote Microalgae Growth in Photobioreactors via Computer Simulations

Flue gas from power plants can promote algal cultivation and reduce greenhouse gas emissions1. Microalgae not only capture solar energy more efficiently than plants3, but also synthesize advanced biofuels2-4. Generally, atmospheric CO2 is not a sufficient source for supporting maximal algal growth5. On the other hand, the high concentrations of CO2 in industrial exhaust gases have adverse effects on algal physiology. Consequently, both cultivation conditions (such as nutrients and light) and the control of the flue gas flow into the photo-bioreactors are important to develop an efficient &ldquo;flue gas to algae&rdquo; system. Researchers have proposed different photobioreactor configurations4,6 and cultivation strategies7,8 with flue gas. Here, we present a protocol that demonstrates how to use models to predict the microalgal growth in response to flue gas settings. We perform both experimental illustration and model simulations to determine the favorable conditions for algal growth with flue gas. We develop a Monod-based model coupled with mass transfer and light intensity equations to simulate the microalgal growth in a homogenous photo-bioreactor. The model simulation compares algal growth and flue gas consumptions under different flue-gas settings. The model illustrates: 1) how algal growth is influenced by different volumetric mass transfer coefficients of CO2; 2) how we can find optimal CO2 concentration for algal growth via the dynamic optimization approach (DOA); 3) how we can design a rectangular on-off flue gas pulse to promote algal biomass growth and to reduce the usage of flue gas. On the experimental side, we present a protocol for growing Chlorella under the flue gas (generated by natural gas combustion). The experimental results qualitatively validate the model predictions that the high frequency flue gas pulses can significantly improve algal cultivation.

Flue gas from power plants is a cheap CO2 source for algal growth. We have built prototype &#34;flue gas to algal cultivation&#34; systems and described how to scale up the algal cultivation process. We have demonstrated the use of a mass-transfer bio-reaction model to simulate and to design the optimal operation of flue gas for the growth of Chlorella sp. in algal photo-bioreactors.

Flue gas from power plants can promote algal cultivation and reduce greenhouse gas emissions1. Microalgae not only capture solar energy more efficiently ...

Detection of Foodborne Bacterial Pathogens from Individual Filth Flies

There is unanimous consensus that insects are important vectors of foodborne pathogens. However, linking insects as vectors of the pathogen causing a particular foodborne illness outbreak has been challenging. This is because insects are not being aseptically collected as part of an environmental sampling program during foodborne outbreak investigations and because there is not a standardized method to detect foodborne bacteria from individual insects. To take a step towards solving this problem, we adapted a protocol from a commercially available PCR-based system that detects foodborne pathogens from food and environmental samples, to detect foodborne pathogens from individual flies.Using this standardized protocol, we surveyed 100 wild-caught flies for the presence of Cronobacter spp., Salmonella enterica, and Listeria monocytogenes and demonstrated that it was possible to detect and further isolate these pathogens from the body surface and the alimentary canal of a single fly. Twenty-two percent of the alimentary canals and 8% of the body surfaces from collected wild flies were positive for at least one of the three foodborne pathogens. The prevalence of Cronobacter spp. on either body part of the flies was statistically higher (19%) than the prevalence of S. enterica (7%) and L.monocytogenes (4%). No false positives were observed when detecting S. enterica and L. monocytogenes using this PCR-based system because pure bacterial cultures were obtained from all PCR-positive results. However, pure Cronobacter colonies were not obtained from about 50% of PCR-positive samples, suggesting that the PCR-based detection system for this pathogen cross-reacts with other Enterobacteriaceae present among the highly complex microbiota carried by wild flies. The standardized protocol presented here will allow laboratories to detect bacterial foodborne pathogens from aseptically collected insects, thereby giving public health officials another line of evidence to find out how the food was contaminated when performing foodborne outbreak investigations.

A PCR-based protocol was adapted to detect Cronobacter spp., Salmonella enterica, and Listeria monocytogenes from body surfaces and alimentary canals of individual wild-caught flies. The goal of this protocol is to detect and isolate bacterial pathogens from individual insects collected as part of an environmental sampling program during foodborne outbreak investigations.

There is unanimous consensus that insects are important vectors of foodborne pathogens. However, linking insects as vectors of the pathogen causing a ...

A Fish-feeding Laboratory Bioassay to Assess the Antipredatory Activity of Secondary Metabolites from the Tissues of Marine Organisms

Marine chemical ecology is a young discipline, having emerged from the collaboration of natural products chemists and marine ecologists in the 1980s with the goal of examining the ecological functions of secondary metabolites from the tissues of marine organisms. The result has been a progression of protocols that have increasingly refined the ecological relevance of the experimental approach. Here we present the most up-to-date version of a fish-feeding laboratory bioassay that enables investigators to assess the antipredatory activity of secondary metabolites from the tissues of marine organisms. Organic metabolites of all polarities are exhaustively extracted from the tissue of the target organism and reconstituted at natural concentrations in a nutritionally appropriate food matrix. Experimental food pellets are presented to a generalist predator in laboratory feeding assays to assess the antipredatory activity of the extract. The procedure described herein uses the bluehead, Thalassoma bifasciatum, to test the palatability of Caribbean marine invertebrates; however, the design may be readily adapted to other systems. Results obtained using this laboratory assay are an important prelude to field experiments that rely on the feeding responses of a full complement of potential predators. Additionally, this bioassay can be used to direct the isolation of feeding-deterrent metabolites through bioassay-guided fractionation. This feeding bioassay has advanced our understanding of the factors that control the distribution and abundance of marine invertebrates on Caribbean coral reefs and may inform investigations in diverse fields of inquiry, including pharmacology, biotechnology, and evolutionary ecology.

This bioassay employs a model predatory fish to assess the presence of feeding-deterrent metabolites from organic extracts of the tissues of marine organisms at natural concentrations using a nutritionally comparable food matrix.

Marine chemical ecology is a young discipline, having emerged from the collaboration of natural products chemists and marine ecologists in the 1980s with ...

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated (WATER-PAM) Fluorometry

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be subsampled throughout the experiment1&#8211;7. Subsampling of large volumes can be problematic for several reasons: 1) it causes variation in the total volume and the surface area:volume ratio of the culture during the experiment; 2) pseudo-replication (i.e., replicate samples from the same treatment flask8) is often employed rather than true replicates (i.e., sampling from replicate treatments); 3) the duration of the experiment is limited by the total volume; and 4) axenic cultures or the usual bacterial microbiota are difficult to maintain during long-term experiments as contamination commonly occurs during subsampling.
The use of microtiter plates enables 1 ml culture volumes to be used for each replicate, with up to 48 separate treatments within a 12.65 x 8.5 x 2.2 cm plate, thereby decreasing the experimental volume and allowing for extensive replication without subsampling any treatment. Additionally, this technique can be modified to fit a variety of experimental formats including: bacterial-algal co-cultures, algal physiology tests, and toxin screening9&#8211;11. Individual wells with an alga, bacterium and/or co-cultures can be sampled for numerous laboratory procedures including, but not limited to: WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry, microscopy, bacterial colony forming unit (cfu) counts and flow cytometry. The combination of the microtiter plate format and WATER-PAM fluorometry allows for multiple rapid measurements of photochemical yield and other photochemical parameters with low variability between samples, high reproducibility and avoids the many pitfalls of subsampling a carboy or conical flask over the course of an experiment.

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated WATER-PAM Fluorometry

The goal of this procedure is to demonstrate the reproducibility and adaptability of using a microtiter plate format for microalgal screening. This rapid screen combines WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry to measure photosynthetic yield as an indicator of Photosystem II (PSII) health with small volume bacterial-algal co-cultures.

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be ...

The Plant Infection Test: Spray and Wound-Mediated Inoculation with the Plant Pathogen Magnaporthe Grisea

Plants possess a powerful system to defend themselves against potential threats by pathogenic fungi. For agriculturally important plants, however, current measures to combat such pathogens have proved too conservative and, thus, not sufficiently effective, and they can potentially pose environmental risks. Therefore, it is extremely necessary to identify host-resistance factors to assist in controlling plant diseases naturally through the identification of resistant germplasm, the isolation and characterization of resistance genes, and the molecular breeding of resistant cultivars. In this regard, there is need to establish an accurate, rapid, and large-scale inoculation method to breed and develop plant resistance genes. The rice blast fungal pathogen Magnaporthe grisea causes severe disease symptoms and yield losses. Recently, M. grisea has emerged as a model organism for studying the mechanisms of plant-fungal pathogen interactions. Hence, we report the development of a plant virulence test method that is specific for M. grisea. This method provides for both spray inoculation with a conidial suspension and wounding inoculation with mycelium cubes or droplets of conidial suspension. The key step of the wounding inoculation method for detached rice leaves is to make wounds on plant leaves, which avoids any interference caused by host penetration resistance. This spray/wounding protocol contributes to the rapid, accurate, and large-scale screening of the pathotypes of M. grisea isolates. This integrated and systematic plant infection method will serve as an excellent starting point for gaining a broad perspective of issues in plant pathology.

The Plant Infection Test: Spray and Wound-Mediated Inoculation with the Plant Pathogen Magnaporthe Grisea

Here, we present a protocol to test plant virulence with the plant pathogen Magnaporthe grisea. This report will contribute to the large-scale screening of the pathotypes of fungi isolates and serve as an excellent starting point for understanding the resistant mechanisms of plants during molecular breeding.

Plants possess a powerful system to defend themselves against potential threats by pathogenic fungi. For agriculturally important plants, however, current ...

Isolation and Analysis of Microbial Communities in Soil, Rhizosphere, and Roots in Perennial Grass Experiments

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The purpose of this manuscript is to provide detail on how to rapidly sample soil, rhizosphere, and endosphere of replicated field trials and analyze changes that may occur in the microbial communities due to sample type, treatment, and plant genotype. The experiment used to demonstrate these methods consists of replicated field plots containing two, pure, warm-season grasses (Panicum virgatum and Andropogon gerardii) and a low-diversity grass mixture (A. gerardii, Sorghastrum nutans, and Bouteloua curtipendula). Briefly, plants are excavated, a variety of roots are cut and placed in phosphate buffer, and then shaken to collect the rhizosphere. Roots are brought to the laboratory on ice and surface sterilized with bleach and ethanol (EtOH). The rhizosphere is filtered and concentrated by centrifugation. Excavated soil from around the root ball is placed into plastic bags and brought to the lab where a small amount of soil is taken for DNA extractions. DNA is extracted from roots, soil, and rhizosphere and then amplified with primers for the V4 region of the 16S rRNA gene. Amplicons are sequenced, then analyzed with open access bioinformatics tools. These methods allow researchers to test how the microbial community diversity and composition varies due to sample type, treatment, and plant genotype. Using these methods along with statistical models, the representative results demonstrate there are significant differences in microbial communities of roots, rhizosphere, and soil. Methods presented here provide a complete set of steps for how to collect field samples, isolate, extract, quantify, amplify, and sequence DNA, and analyze microbial community diversity and composition in replicated field trials.

Excavation of plant roots from the field as well as processing of samples into endosphere, rhizosphere, and soil are described in detail, including DNA extraction and data analysis methods. This paper is designed to enable other laboratories to use these techniques for the study of soil, endosphere, and rhizosphere microbiomes.

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The ...

Visualizing Efficacy of Pesticides Against Disease Vector Mosquitoes in the Field

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform across ecological zones. To best protect public and veterinary health from these insects, the environmental limitations of pesticides need to be investigated to inform effective use of the most appropriate pesticide formulations and techniques. We have developed a research program to evaluate combinations of pesticides, pesticide application equipment, and application techniques in hot-arid desert, hot-humid tropical, warm and cool temperate, and urban locations to derive pesticide use guidelines specific to target insect and environment. To these ends we designed a system of protocols to support efficient, cost-effective, portable, and standardized evaluation of a diverse range of pesticides and equipment across multiple environments. At the core of these protocols is the use of an array of small cages with colony-reared sentinel mosquitoes (adults and immatures) and sand flies (adults), strategically arranged in natural habitats and exposed to pesticide spray. Spatial and temporal patterns of pesticide efficacy are derived from percent mortality in sentinel cages, then mapped and visualized in a geographic information system. Maps of sentinel mortality data may be statistically compared to evaluate relative efficacy of a pesticide across multiple environments, or to study multiple pesticides in a single environment. Protocols may be modified to accommodate a variety of scenarios, including, for example, the vertical orientation of sentinels in canopy habitats or simultaneous testing of ground and aerial application methods.

The efficacy of public health pesticides targeting nuisance and disease-vector insects is not uniform across different ecological zones. Here we present a system of techniques using captive vector insects as sentinels for pesticide efficacy to derive electronic maps supporting the standard evaluation of pesticides across multiple environments.

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform ...

Colletotrichum fioriniae Development in Water and Chloroform-based Blueberry and Cranberry Floral Extracts

To accurately monitor the phenology of the bloom period and the temporal dynamics of floral chemical cues on fungal fruit rotting pathogens, floral extraction methods and coverslip bioassays were developed utilizing Colletotrichum fioriniae. In blueberry and cranberry, this pathogen is optimally controlled by applying fungicides during the bloom period because of the role flowers play in the initial stages of infection. The protocol detailed here describes how floral extracts (FE) were obtained using water-, chloroform-, and field rainwater-based methods for later use in corresponding glass coverslip bioassays. Each FE served to provide a different set of information: response of C. fioriniae to mobilized floral chemical cues in water (water-based), pathogen response to flower and fruit surface waxes (chloroform-based), and field-based monitoring of collected floral rainwater, moving in vitro observations to an agricultural setting. The FE is broadly described as either water- or chloroform-based, with an appropriate bioassay described to compensate for the inherent differences between these two materials. Rainwater that had run off flowers was collected in unique devices for each crop, alluding to the flexibility and application of this approach for other crop systems. The bioassays are quick, inexpensive, simple, and provide the ability to generate spatiotemporal and site-specific information about the presence of stimulatory floral compounds from various sources. This information will ultimately better inform disease management strategies, as FE decrease the time needed for infection to occur, thus providing insight into changing risks for pathogen infection over the growing season.

Colletotrichum fioriniae Development in Water and Chloroform-based Blueberry and Cranberry Floral Extracts

Here, bioassays designed to monitor the development of a fungal pathogen, Colletotrichum fioriniae, in the presence of blueberry or cranberry floral extracts on glass coverslips are described. Water-, chloroform-, and field rainwater- based floral extraction techniques are detailed as well as insight into how this information can be applied.

To accurately monitor the phenology of the bloom period and the temporal dynamics of floral chemical cues on fungal fruit rotting pathogens, floral ...

Leaf Area Index Estimation Using Three Distinct Methods in Pure Deciduous Stands

Accurate estimations of leaf area index (LAI), defined as half of the total leaf surface area per unit of horizontal ground surface area, are crucial for describing the vegetation structure in the fields of ecology, forestry, and agriculture. Therefore, procedures of three commercially used methods (litter traps, needle technique, and a plant canopy analyzer) for performing LAI estimation were presented step-by-step. Specific methodological approaches were compared, and their current advantages, controversies, challenges, and future perspectives were discussed in this protocol. Litter traps are usually deemed as the reference level. Both the needle technique and the plant canopy analyzer (e.g., LAI-2000) frequently underestimate LAI values in comparison with the reference. The needle technique is easy to use in deciduous stands where the litter completely decomposes each year (e.g., oak and beech stands). However, calibration based on litter traps or direct destructive methods is necessary. The plant canopy analyzer is a commonly used device for performing LAI estimation in ecology, forestry, and agriculture, but is subject to potential error due to foliage clumping and the contribution of woody elements in the field of view (FOV) of the sensor. Eliminating these potential error sources was discussed. The plant canopy analyzer is a very suitable device for performing LAI estimations at the high spatial level, observing a seasonal LAI dynamic, and for long-term monitoring of LAI.

An accurate estimation of leaf area index (LAI) is crucial for many models of material and energy fluxes within plant ecosystems and between an ecosystem and the atmospheric boundary layer. Therefore, three methods (litter traps, needle technique, and PCA) for taking precise LAI measurements were in the presented protocol.

Accurate estimations of leaf area index (LAI), defined as half of the total leaf surface area per unit of horizontal ground surface area, are crucial for ...

Cortisol Extraction from Sturgeon Fin and Jawbone Matrices

The aims of this study were to develop a technique for the extraction of cortisol from sturgeon fins using two washing solvents (water and isopropanol) and quantify any differences in fin cortisol levels among three main sturgeon species. Fins were harvested from 19 sacrificed sturgeons including seven beluga (Huso huso), seven Siberian (Acipenser baerii), and five sevruga (A. stellatus). The sturgeons were raised in Iranian farms for 2 years (2017-2018), and cortisol extraction analysis was conducted in South Korea (January-February 2019). Jawbones from five H. huso were also used for cortisol extraction. Data were analyzed using the general linear model (GLM) procedure in the SAS environment. The intra- and inter-assay coefficients of variation were 14.15 and 7.70, respectively. Briefly, the cortisol extraction technique involved washing the samples (300 &#177; 10 mg) with 3 mL of solvent (ultrapure water and isopropanol) twice, rotation at 80&#160;rpm for 2.5 min, air-drying the washed samples at room temperature (22-28 &#176;C) for 7 days, further drying the samples using a bead beater at 50 Hz for 32 min and grinding them into powder, applying 1.5 mL methanol to the dried powder (75 &#177; 5 mg), and slow rotation (40 rpm) for 18 h at room temperature with continuous mixing. Following extraction, samples were centrifuged (9,500 x g for 10 min), and 1 mL supernatant was transferred into a new microcentrifuge tube (1.5 mL), incubated at 38 &#176;C to evaporate the methanol, and analyzed via enzyme-linked immunosorbent assay (ELISA). No differences were observed in fin cortisol levels among species or in fin and jawbone cortisol levels between washing solvents. The results of this study demonstrate that the sturgeon jawbone matrix is a promising alternative stress indicator to solid matrices.

In this study, we present a protocol for cortisol extraction from the fin and jawbone of sturgeon species. Fin and jawbone cortisol levels were further examined by comparing two washing solvents followed by ELISA assays. This study piloted the feasibility of jawbone cortisol as a novel stress indicator.

The aims of this study were to develop a technique for the extraction of cortisol from sturgeon fins using two washing solvents (water and isopropanol) ...