This study was funded by the National Environment Agency (NEA), Singapore. We thank Mr Chew Ming Fai, Deputy Chief Executive Officer (Public Health), NEA, for his approval to publish the study, and A/Prof Ng Lee Ching, Group Director (Environmental Health Institute Group), NEA, for her support in this study. We also thank Dr Shuzhen Sim and Dr Denise Tan for proofreading the manuscript.

1. Rearing of mosquitoes
<ol>
	<li>Conduct all mosquito rearing and the male mating competitiveness assay under standard insectary conditions of 27 &#177; 1 &#176;C and 75-80% relative humidity, with a photoperiod of 12 h:12 h light: dark cycles.</li>
	<li>Designate the two sets of competing males as Set A and Set B for easy reference in the methodology described in this paper. Rear the mosquitoes under standardized conditions to ensure a fair comparison of their fitness during the assay. Rear the mosquitoes at a larval density of 500 larvae in 2 L of water and feed them with ground fish food powder ad libitum. 
	NOTE: For the generation of the representative results, Set A and Set B were the inbred and outcross males of the Wolbachia-infected Ae. Aegypti, respectively.</li>
	<li>Sex male and female mosquitoes at the pupal stage, and contain them separately in cages (see the Table of Materials) of dimensions W 32.5 cm x D 32.5 cm x H 32.5 cm, with mesh size of 150 x 150 and 160 &#181;m aperture. Maintain all adult mosquitoes with 10% sucrose solution.</li>
</ol>
2. Preparation of male and female mosquitoes
<ol>
	<li>Sex the male and female mosquitoes at the pupal stage according to their size differences (male pupae being smaller than female pupae) (Figure 1).</li>
	<li>For each set of mosquitoes (Set A or Set B), transfer 100 male pupae each into a prelabelled cage for sucrose feeding or RhB-sucrose feeding.</li>
	<li>Place the female pupae in small batches of 40-50 per cage. Upon the emergence of the imagoes, check the cages for the presence of male mosquitoes. 
	&#8203;NOTE: Adult male mosquitoes are smaller than females and have bushier and hairier antennae (Figure 2). Use only virgin female mosquitoes for the mating competitiveness assays. The use of pre-inseminated females will render all resulting data invalid. Thus, extreme care must be taken during sexing at the pupae stage. Do not use the female mosquitoes from a cage that has been contaminated with male mosquitoes. Extra cages of females should be prepared.</li>
</ol>
3. Preparation of 0.2% RhB - sucrose solution
NOTE: RhB is a green powder in dry form and reddish-pink in solution. Standard personal protective equipment (PPE: laboratory protection gown, nitrile gloves, and eye protection) shall be worn when handling this chemical. To avoid inhalation, weigh the RhB powder in a fume hood.
<ol>
	<li>To prepare a 0.2% w/v RhB-sucrose solution, dissolve 200 mg of RhB powder for every 100 mL of 10% w/v/ sucrose solution. Mix well to ensure all the powder is dissolved. 
	NOTE: As RhB is light-sensitive, use amber bottles, or wrap clear bottles completely with aluminum foil.</li>
</ol>
4. Feeding of male mosquitoes
NOTE: Data from RhB-sucrose feeding optimization is presented in Supplemental Material, section 1.
<ol>
	<li>Prepare 20 sugar feeder bottles with a wick. Add 10 mL of 10% sucrose in 10 feeder bottles and 10 mL of 0.2% RhB-sucrose solution in the other 10 feeder bottles (use amber bottles, or wrap the bottles in aluminum foil).</li>
	<li>Place the feeder bottles in the respective male cages (5 bottles per cage) prepared in steps 2.2 and 2.3. Allow the male mosquitoes to feed for three days before the mating experiment.</li>
</ol>
5. Checking for RhB fluorescence in male mosquitoes
<ol>
	<li>Aspirate RhB-sucrose-fed male mosquitoes, and observe them under a fluorescence stereo microscope to ensure that all the RhB-sucrose-fed male mosquitoes have been successfully marked with RhB.</li>
	<li>Turn on the mercury burner lamp and the stereo microscope. Allow the light source of the mercury burner lamp to stabilize for 10 min. Set the fluorescence filters for red fluorescence protein 1 (RFP1) (excitation wavelength 540 nm, emission wavelength 625 nm).</li>
	<li>Aspirate a small number of mosquitoes (four or five) at a time into the glass tube of the oral aspirator. Through the glass tube, observe the body of the male mosquitoes under the fluorescence stereo microscope. Exclude male mosquitoes not marked with RhB from the experiment. 
	NOTE: The abdomen of male mosquitoes marked with RhB will appear pink under white light (Figure 3A) and glow bright orange under fluorescent light (Figure 3B).</li>
	<li>Transfer Set A male mosquitoes into 12 paper cups secured with netting (mesh size 150 x 150, 160 &#181;m aperture); 6 cups each with 10 sucrose-fed male mosquitoes and the other 6 cups each with 10 RhB-sucrose-fed male mosquitoes. Repeat this step for Set B male mosquitoes.</li>
</ol>
6. Mating competitiveness assay
<ol>
	<li>Set up 12 W 60 cm x D 60 cm x H 60 cm cages with mesh size 44 x 32, 650 &#181;m aperture for the mating assay. In each of the six cages, include 10 Set A (RhB-marked) male mosquitoes, 10 Set B (unmarked) male mosquitoes, and 10 virgin wild-type female mosquitoes. In the other six cages, include 10 Set A (unmarked) male mosquitoes, 10 Set B (RhB-marked) male mosquitoes, and 10 virgin wild-type female mosquitoes. Label these cages to clearly distinguish between the two mating combinations. 
	NOTE: Based on experience, a 60 cm x 60 cm x 60 cm cage was used for the mating assay as a smaller cage may encourage mixed mating.</li>
	<li>Place the respective cups of males prepared in step 5.4 into the mating cages according to the label in step 6.1. Remove the netting and gently tap the cup to jostle the males out of the cup. Carefully remove the paper cup and the netting from the cage to ensure no mosquitoes escape from the cage. Allow the male mosquitoes to acclimate in the mating cage for at least an hour.</li>
	<li>Using an oral aspirator, transfer virgin wild-type female mosquitoes into 12 paper cups, with each cup containing 10 mosquitoes.</li>
	<li>After the acclimation period for the male mosquitoes, transfer one cup of females into every mating cage and remove the netting. Gently jostle the cup to encourage any remaining female mosquitoes out of the cup. Carefully remove the paper cup and the netting from the cage to ensure no mosquitoes escape from the cage.</li>
	<li>Allow mating to take place for 3 h. 
	NOTE: Recommended mating duration was determined through prior observations of wild-type Ae. aegypti. In experiments involving 10 females and 20 males being held in a 60 cm x 60 cm x 60 cm cage, 90% female insemination was achieved in 3 h (Supplemental Material, section 2). Do not disturb the cage during this period as agitation could lead to interrupted and mixed mating. Mixed mating (where the female has mated with both marked and unmarked males) results in a bias towards RhB-marked males as it is difficult to distinguish unmarked sperms from RhB-marked sperms under a fluorescence microscope.</li>
	<li>To terminate the mating experiment, remove all the mosquitoes from each cage using a mechanical aspirator. Cold anesthetize the mosquitoes on ice for at least 5 min. When the mosquitoes are fully anesthetized, gently pick up the female mosquitoes and house them in a separate paper cup secured with netting (mesh size 150 x 150, 160 &#181;m aperture). Label the paper cup by transferring the respective label from the mating cage onto the paper cup. 
	NOTE: It is possible to pause the experiment at this point and maintain the females with 10% sucrose solution. The RhB-marked seminal fluid will remain stable inside the female spermathecae for at least a week. It is best to keep the females alive before dissection as dead and desiccated specimens are difficult to dissect.</li>
	<li>To score the female spermathecae, cold anesthetize the female mosquitoes on ice for at least 5 min before dissection under a stereo microscope (Video 1). Examine the spermathecae under a compound light microscope (magnification 100x) for their insemination status (Figure 4). For inseminated individuals, determine whether the spermathecae contain RhB-marked seminal fluid by examining them under the fluorescence stereo microscope equipped with an RFP1 filter and a camera imaging system. 
	&#8203;NOTE: When using a fluorescence stereo microscope with an attached imaging system, it is recommended to utilize a prolonged exposure time (5 s) to increase the detection sensitivity. If the female mosquito has mated with a marked male, her spermathecae will fluoresce bright orange (Figure 5A). However, if the female mosquito has mated with an unmarked male, her inseminated spermathecae will not fluoresce (Figure 5B).</li>
</ol>
7. Disposal of RhB waste
<ol>
	<li>Treat aqueous RhB waste with activated carbon25 before discharging it as general waste-water. Dispose of solid RhB waste (mosquitoes marked with RhB, paper towels, and wicks soaked with RhB) as chemical waste. Don standard PPE when handling RhB waste.</li>
</ol>

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The authors declare that they have no competing financial interests.

Marking is commonly used in entomological research to study insect population dynamics, dispersal, behavior, and mating biology26. In SIT and IIT programs, marking is carried out to differentiate the release strain from the field population to study their dispersal and optimize the release ratio. The marking methods used include genetic marking27,28, incorporating isotopes in larval food29,30, fluorescent dust31, and dye32. For the suppression of mosquito populations using SIT or IIT, where male mating fitness is a critical component, fluorescent dyes have been used as markers to study mosquito mating biology18,19.
Conventionally, assessment of male mating competitiveness of the release strain has been assessed using female fertility assays. However, this assay is time-consuming and labor-intensive due to downstream experimental processes post-mating (Supplemental Figure S1). These processes include blood-feeding of the females, egg collection, hatching of the eggs, and enumeration of the proportion of hatched eggs to determine egg viability. On average, this assay requires 30 man-hours and two weeks of experimental work (starting from setting up of the competitiveness assay cages) to the final determination of male mating competitiveness.
his paper presents the use of a fluorescent dye, RhB, (fed as 0.2% RhB&#8211;sucrose to the mosquitoes, Supplemental Figure S2) to directly measure mating interactions between females and RhB- marked males. While this protocol requires a fluorescence stereo microscope, it obviates the need to perform the time-consuming experimental procedures mentioned above. On average, this RhB-based assay requires approximately 10 man-hours and about a day to obtain data equivalent to that from female fertility assays. This This &#62;90% time savings allows researchers researchers to perform multiple experimental replicates, providing a more robust validation of male mating fitness. In addition, this assay can be used to compare the mating competitiveness between two sterile or Wolbachia-infected&#160; mosquitoes lines.
This type of comparison is not possible with conventional female fertility assays, as females would yield non-viable eggs upon mating with both such lines.&#160; Notwithstanding, any mixed mating in the experiment will result in bias toward the marked population as it is difficult to identify unmarked sperms in female spermathecae that contain seminal fluid from both RhB-marked and unmarked males. A similar conclusion was made in a study assessing mating competitiveness of Anopheles gambiae using RhB18, whereby a greater proportion of females in the mating assay was found to be mated by marked males. As polyandry is more likely to occur in females that had previously engaged in an interrupted mating33, the probability of this occurring was reduced in this study by using fewer mosquitoes (20 males to 10 females) in a larger cage volume (0.216 m3) in these experiments.
The results showed no bias toward the RhB-marked population, indicating that mixed mating was limited. In summary, incorporation of RhB to mark males in a mating competitiveness assay is an economical and rapid way of evaluating male mating fitness. This method also allows for the direct comparison of mating competitiveness between males exposed to different doses of irradiation, reared in different rearing regimes, or those infected with different strains of Wolbachia, making it a valuable tool for the evaluation of male mating fitness for any male release-based mosquito population suppression program.

The rearing and release of sterile or incompatible males for suppression of Aedes mosquito populations is currently being evaluated in the field as a novel tool to prevent outbreaks of dengue and other Aedes-borne disease1. Male-release suppression strategies that are currently in&#160; field trials&#160; include the use of the genetic method2, irradiation (Sterile Insect Technique, SIT)3, endosymbiotic bacteria Wolbachia (Insect Incompatible Technique, IIT)4, or a combination of the latter two techniques5,6. The success of these approaches is largely dependent on the released males&#8217; ability to outcompete wild-type males and seek females to secure copulation. Otherwise, sterility cannot be induced in the target population.&#160;
In a classical SIT program, for example, male mating fitness may be affected by factors such as irradiation dose7,8,9, mass rearing protocol and the extent of inbreeding in the colony10,11,12,13,14. Furthermore, studies on the mating competitiveness may provide important knowledge on mosquito mating behavior which could be used to inform vector control strategies.&#160; &#160;
In SIT and IIT, the mating competitiveness of male mosquitoes is typically assessed by allowing both wild-type and release strain to compete for&#160; wild-type females in a cage8,11,15,16. Females are then blood-fed and their eggs hatched to determine viability. Females that lay non-viable eggs or eggs with low hatch rate are assumed to have mated with release strain males, while females that produce viable eggs are assumed to have mated with wild-type males. Mating competitiveness is then calculated with the Fried Index17. Unfortunately, this method is resource-intensive and time-consuming, and the overall Fried Index can be influenced by external confounding factors affecting egg viability such as&#160; poor egg handling and over-desiccation can result in a low hatch rate in the compatibility cross that may then lead to an artificially low Fried Index.&#160;
&#160;Moreover, this method does not allow for the direct comparison of mating competitiveness between Aedes mosquitoes that are infected with different strains of Wolbachia or that are exposed to different doses of irradiation. Hence, a more direct method is required to address these challenges. Recent studies18,19 have demonstrated the effectiveness of using the fluorescent dye, RhB, to mark male mosquitoes&#8217; seminal fluid. Marked seminal fluid is transferred and stored in the female mosquitoes&#8217; spermathecae upon successful mating, allowing direct measurement of female mating interaction with marked males. Rhodamine B is a thiol-reactive fluorone dye commonly used as a biomarker for ecological and behavioral research studies in animals including insects20. For mosquito studies, RhB is introduced by feeding with sugar or honey water containing dissolved RhB powder18,19,21,22,23,24. Upon uptake, the RhB dye binds to proteins, staining body tissue with a reddish-pink stain that fluoresces bright orange under a fluorescent light source.&#160;
The strong fluorescence signal and stability of the marking, coupled with its ability to stain insect seminal fluids, allows for the monitoring of the transfer of marked seminal fluid from the labelled male to the sperm storage organs of the female insect for mating studies18,19,21,24. The use of RhB in a male mating competitiveness assay not only allows direct measurement of the mating interaction of females with either marked and unmarked males, but the results can also be obtained within 24 h as it obviates the process of determining the egg viability, which typically requires about 10 to 14 days. Furthermore, this method overcomes the potential loss of data when the female mosquitoes do not blood-feed or die before ovipositioning. This is particularly crucial because in semi-field trials, where female mosquitoes are prone to damage and death during post-mating collection using a backpack or mechanical aspirator. To address the current limitations of using female fertility to we present an alternative method that uses RhB staining to directly measure male mosquito mating competitiveness. The method simplifies the workflow, shortened&#160; experimental time from about two weeks to one day, allowing for more experimental replicates to be carried out and allows comparison between two release strains. This protocol will be suitable for laboratories that are embarking on male release-based mosquito population suppression programs, and may be used for routine&#160; quality control and strain evaluation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Compound light microscope</td>
			<td>Olympus</td>
			<td>CX23</td>
			<td>To score for spermathecae insemination</td>
		</tr>
		<tr>
			<td>Dissection forceps</td>
			<td>Bioquip</td>
			<td>Rubis forceps (4524)</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence stereo-light microscope with RFP1 filter</td>
			<td>Olympus</td>
			<td>SZX16</td>
			<td>To check for Rhodamine B fluorescence signal</td>
		</tr>
		<tr>
			<td rowspan="2">Mosquito cages</td>
			<td rowspan="2">Bugdorm</td>
			<td>4F3030</td>
			<td>W 32.5 cm x D 32.5 cm x H 32.5 cm; mesh size of 150 x 150; 160 &#181;m aperture For holding of male and female adult mosquitoes prior to mating assay</td>
		</tr>
		<tr>
			<td>6M610</td>
			<td>W 60 cm x D 60 cm x H 60 cm; mesh size of 44 x 32; 650 &#181;m aperture 
			For mating competitiveness assay</td>
		</tr>
		<tr>
			<td>Mosquito netting</td>
			<td></td>
			<td></td>
			<td>150 x 150, 160 &#181;m aperture</td>
		</tr>
		<tr>
			<td>Rhodamine B</td>
			<td>Sigma Aldrich</td>
			<td>R6626</td>
			<td>&#8805;95% (HPLC)</td>
		</tr>
		<tr>
			<td>Stereo-light microscope</td>
			<td>Olympus</td>
			<td>SZ61</td>
			<td>For spermathecae dissection</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>MP Biomedicals</td>
			<td>SKU 029047138</td>
			<td>Food grade</td>
		</tr>
		<tr>
			<td>TetraMin tropical flakes</td>
			<td>Tetra</td>
			<td>77101</td>
			<td>Fish food for feeding larvae</td>
		</tr>
	</tbody>
</table>

using the fluorescent dye, rhodamine b, to study mating competitiveness in male aedes aegypti mosquitoes

The success of sterile or incompatible insect technique-based population suppression programs depends on the ability of released males to compete for wild-type females and induce sterility in the target population. Hence, laboratory assessment of male mating competitiveness is essential for evaluating the release strain&#8217;s fitness before field release. Conventionally, such an assay is performed by determining the proportion of viable eggs produced by the females after being simultaneously exposed to two sets of males (wild-type and release strains) for copulation. However, this process is time-consuming and laborious due to the need to first blood-feed the females for egg production and then hatch and enumerate the hatched eggs to determine egg viability.&#160;
Moreover, this method cannot discern the degree of competitiveness between two sterile or Wolbachia-infected mosquito lines as wild-type female mosquitoes will only produce non-viable eggs upon mating with both. To circumvent these limitations, this paper describes a more direct method of measuring male mosquito mating competitiveness in laboratory settings using the fluorescent dye, rhodamine B (RhB), which can be used to mark males by feeding them in sucrose solution containing RhB. After the mating assay, the presence of fluorescing sperms in the spermathecae of a female can be used to determine her mating partner. This method is cost-effective, reduces the experimental time by 90% and allows comparison of mating fitness between two sterile or Wolbachia-infected lines.&#160;

wAlbB-SG is a localized (Singapore) Ae. aegypti line stably infected with the wAlbB strain of Wolbachia. Using the protocol described in this paper, we evaluated the male mating competitiveness of an inbred and an outcrossed line of wAlbB-SG to determine if inbreeding results in a loss in male mating fitness. The inbred line had been maintained for 11 generations in the insectary, while the outcrossed line was generated by backcrossing the females with wild-type male Ae. aegypti. Males from the inbred and outcrossed lines were competed against each other for mating with wild-type wild-type female Ae. aegypti. The mating competitiveness assay was conducted in triplicate.&#160;&#160;
The results indicated that RhB did not affect the fitness of the males as the data for female insemination was not biased towards or against mating with RhB-sucrose-fed males (Table 1 and Figure 6) As RhB does not affect the mating fitness of the males, we proceed to analyse the data based on the percentage of inseminated females mated by either the inbred or outcrossed line (Table 2 and Figure 7). The result across the experimental triplicates were consistent; there was a significantly higher percentage of females mated with the outcross males than with the inbred males in all three replicates (P &#8804; 0.05, Mann-Whitney U-test). These results suggest a potential loss in male mating fitness after several generations of inbreeding in the laboratory.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62432/62432fig01.jpg" /> 
Figure 1: Lateral view of male (left) and female (right) Aedes aegypti pupae. Under the same rearing conditions, Ae. aegypti can be sexed at the pupal stage according to size; males are significantly smaller than females. <a href="https://www.jove.com/files/ftp_upload/62432/62432fig01large.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/62432/62432fig02.jpg" /> 
Figure 2: Differentiation of male (left) and female (right) Aedes aegypti adults. Adult male mosquitoes (left) have bushier and hairier antennae than the adult female; the red arrows indicate the antennae. <a href="https://www.jove.com/files/ftp_upload/62432/62432fig02large.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/62432/62432fig03.jpg" /> 
Figure 3: Rhodamine B marking of male mosquito. (A) Light microscopy; (B) fluorescence microscopy. The mosquito on the left is unmarked (fed with 10% w/v sucrose), while the one on the right is marked (fed with 0.2% RhB-sucrose). Marked mosquitoes have a visible pink abdomen under white light (the mosquito on the right in A), which fluoresces bright orange under fluorescence microscopy (B). Scale bars = 5 mm. Abbreviation: RhB = Rhodamine B. <a href="https://www.jove.com/files/ftp_upload/62432/62432fig03large.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/62432/62432fig04.jpg" /> 
Figure 4: Inseminated and non-inseminated female spermathecae under a compound light microscope (100x magnification). The insemination status of a female mosquito can be determined by observing its spermathecae under a compound light microscope. An inseminated female mosquito will contain at least one filled spermatheca while all three spermathecae of a non-inseminated female mosquito will be empty. Thread-like, motile sperm will be visible in a filled spermatheca under a compound light microscope. Scale bar = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62432/62432fig04large.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/62432/62432fig05.jpg" /> 
Figure 5: Female mosquito spermathecae inseminated with seminal fluids under a fluorescence stereo microscope. (A) RhB-marked and (B) unmarked spermathecae inseminated with RhB-marked seminal fluids will fluoresce bright orange under fluorescence microscopy. Scale bars = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62432/62432fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62432/62432fig06.jpg" /> 
Figure 6: Number of wild-type females inseminated by either the inbred or outcross males in the experimental triplicates with reciprocal marking. (A) The inbred males were marked with RhB while the outcross males were unmarked. (B) The outcross males were marked with RhB while the inbred males were unmarked. A higher number of females was observed to have mated with outcross males regardless of their marking status. <a href="https://www.jove.com/files/ftp_upload/62432/62432fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/62432/62432fig07.jpg" /> 
Figure 7: Proportion of inseminated females mated with inbred or outcross males in the 3 experimental replicates. For each experimental replicate, there is a significantly higher percentage of females mated with the outcross males (P &#8804; 0.05, Mann-Whitney U-test). <a href="https://www.jove.com/files/ftp_upload/62432/62432fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Dissection of female Aedes aegypti for spermathecae under a light stereo microscope.&#160;<a href="https://www.jove.com/files/ftp_upload/62432/Video 1.mp4" target="_blank">Please click here to download this Video.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>&#9792; x&#160; Inbred (RhB) &#9794; a</td>
			<td>&#9792; x&#160; Outcross (unmarked) &#9794; b</td>
			<td>&#9792; x&#160; Inbred (unmarked) &#9794; c</td>
			<td>&#9792; x&#160; Outcross (RhB) &#9794; d</td>
			<td>Overall insemination rate 
			(a+b+c+d/120)</td>
		</tr>
		<tr>
			<td>Replicate 1</td>
			<td>11</td>
			<td>35</td>
			<td>7</td>
			<td>40</td>
			<td>77.5% (93/120)</td>
		</tr>
		<tr>
			<td>Replicate 2</td>
			<td>6</td>
			<td>29</td>
			<td>8</td>
			<td>31</td>
			<td>61.7% (74/120)</td>
		</tr>
		<tr>
			<td>Replicate 3</td>
			<td>6</td>
			<td>36</td>
			<td>6</td>
			<td>33</td>
			<td>67.5% (81/120)</td>
		</tr>
	</tbody>
</table>
Table 1: Number of females mated with RhB-marked and unmarked wAlbB-Sg Aedes aegypti inbred and outcross males. A total number of 120 females were used in each replicate.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td colspan="2">Percent inseminated females</td>
		</tr>
		<tr>
			<td></td>
			<td>Inbred males</td>
			<td>Outcross males</td>
		</tr>
		<tr>
			<td>Replicate 1</td>
			<td>19% (18/93)</td>
			<td>81% (75/93)</td>
		</tr>
		<tr>
			<td>Replicate 2</td>
			<td>19% (14/74)</td>
			<td>81% (60/74)</td>
		</tr>
		<tr>
			<td>Replicate 3</td>
			<td>15% (12/81)</td>
			<td>85% (69/81)</td>
		</tr>
	</tbody>
</table>
Table 2: Percentage of inseminated females mated with wAlbB-Sg Aedes aegypti inbred and outcross males.
Supplemental Figure S1: Comparison of workflow for RhB-based and conventional mating competitiveness assay. In comparison to the conventional mating competitiveness assay, the simplified and shortened workflow for RhB-based mating competitiveness assay significantly reduces the experimental duration.&#160;<a href="https://www.jove.com/files/ftp_upload/62432/Figure S1.tif" target="_blank">Please click here to download this File.</a>
Supplemental Figure S2: Kaplan Meier survival curves of male adult Aedes aegypti during and after feeding with 0.2% and 0.4% rhodamine B-sucrose feeding. Percent survival of (A) male wild-type and (B) wAlbB-Sg Ae. aegypti during and after three days of feeding on 0.2% and 0.4% RhB-sucrose, compared to controls that were fed with sucrose only.&#160;<a href="https://www.jove.com/files/ftp_upload/62432/Figure S2.tif" target="_blank">Please click here to download this File.</a>
Supplemental Table S1: Insemination rate of females in a W 60 cm x D 60 cm x H 60 cm cage (ratio of 10 females to 20 males) at 1-, 2-, and 3-h time points.&#160;<a href="https://www.jove.com/files/ftp_upload/62432/Table S1.xlsx" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes at JoVE.com

Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

Here, we present a protocol for studying mating competitiveness of male Aedes aegypti using fluorescent dye as a marker. Female mosquitoes are exposed to both marked and unmarked males for copulation. Post mating, their spermathecae are examined under a fluorescence microscope to determine their mating partner.

Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

The success of sterile or incompatible insect technique-based population suppression programs depends on the ability of released males to compete for ...

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4.4K Views.  National Environmental Agency. Here, we present a protocol for studying mating competitiveness of male Aedes aegypti using fluorescent dye as a marker. Female mosquitoes are exposed to both marked and unmarked males for copulation. Post mating, their spermathecae are examined under a fluorescence microscope to determine their mating partner. 

Video: Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

Testing Visual Sensitivity to the Speed and Direction of Motion in Lizards

Testing visual sensitivity in any species provides basic information regarding behaviour, evolution, and ecology. However, testing specific features of the visual system provide more empirical evidence for functional applications. Investigation into the sensory system provides information about the sensory capacity, learning and memory ability, and establishes known baseline behaviour in which to gauge deviations (Burghardt, 1977). However, unlike mammalian or avian systems, testing for learning and memory in a reptile species is difficult. Furthermore, using an operant paradigm as a psychophysical measure of sensory ability is likewise as difficult. Historically, reptilian species have responded poorly to conditioning trials because of issues related to motivation, physiology, metabolism, and basic biological characteristics. Here, I demonstrate an operant paradigm used a novel model lizard species, the Jacky dragon (Amphibolurus muricatus) and describe how to test peripheral sensitivity to salient speed and motion characteristics. This method uses an innovative approach to assessing learning and sensory capacity in lizards. I employ the use of random-dot kinematograms (RDKs) to measure sensitivity to speed, and manipulate the level of signal strength by changing the proportion of dots moving in a coherent direction. RDKs do not represent a biologically meaningful stimulus, engages the visual system, and is a classic psychophysical tool used to measure sensitivity in humans and other animals. Here, RDKs are displayed to lizards using three video playback systems. Lizards are to select the direction (left or right) in which they perceive dots to be moving. Selection of the appropriate direction is reinforced by biologically important prey stimuli, simulated by computer-animated invertebrates.

Testing visual sensitivity in lizards using an operant conditioning paradigm that employs video playback of random-dot kinematograms and computer-generated invertebrates.

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RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

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This RNAi-mediated gene knockdown combined with an in vivo diuresis assay was developed by the Hansen lab to study the effects of RNAi-mediated knockdown of aquaporin genes on Aedes aegypti mosquito diuresis1. 
The protocol is setup in two parts: the first demonstration illustrates how to construct a simple mosquito injection device and how to prepare and inject dsRNA into the thorax of mosquitoes for RNAi-mediated gene knockdown. The second demonstration illustrates how to determine excretion rates in mosquitoes using an in vivo bioassay.

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

In this protocol we combine RNAi-mediated gene silencing with an in vivo diuresis assay to study the effects knockdown of genes of interest has on mosquito fluid excretion.

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion ...

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The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

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Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of these disease vectors. We have recently identified a bitter sensing neuron in the labellum of female Aedes aegypti that responds to DEET and other repellents, as well as bitter quinine, through direct electrophysiological investigation. These gustatory receptor neuron responses prompted our sequencing of total mRNA from both male and female labella and tarsi samples to elucidate the putative chemoreception genes expressed in these contact chemoreception tissues. Samples of tarsi were divided into pro-, meso- and metathoracic subtypes for both sexes. We then validated our dataset by conducting qRT-PCR on the same tissue samples and used statistical methods to compare results between the two methods. Studies addressing molecular function may now target specific genes to determine those involved in repellent perception by mosquitoes. These receptor pathways may be used to screen novel repellents towards disruption of host-seeking behavior to curb the spread of harmful viruses.

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Using two methods to estimate gene expression in the major gustatory appendages of Aedes aegypti, we have identified the set of genes putatively underlying the neuronal responses to bitter and repulsive compounds, as determined by electrophysiological examination.

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of ...

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In our example experiment, we demonstrate the protocol through the excision and culture of fat bodies from the yellow fever mosquito, Aedes aegypti, a principal vector of arboviruses including dengue, chikungunya, and Zika. RNA from fat bodies cultured under a physiological condition known to upregulate yolk proteins versus the control were subject to RNA-Seq analysis to demonstrate the potential utility of this procedure for investigations of gene expression.

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The insect fat body plays a central role in insect metabolism and nutrient storage, mirroring functions of the liver and fat tissue in vertebrates. Insect ...

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, chikungunya and Zika. Wolbachia-infected mosquitoes can be released into the field to either reduce population sizes through incompatible matings or to transform populations with mosquitoes that are refractory to virus transmission. For these strategies to succeed, the mosquitoes released into the field from the laboratory must be competitive with native mosquitoes. However, maintaining mosquitoes in the laboratory can result in inbreeding, genetic drift and laboratory adaptation which can reduce their fitness in the field and may confound the results of experiments. To test the suitability of different Wolbachia infections for deployment in the field, it is necessary to maintain mosquitoes in a controlled laboratory environment across multiple generations. We describe a simple protocol for maintaining Ae. aegypti mosquitoes in the laboratory, which is suitable for both Wolbachia-infected and wild-type mosquitoes. The methods minimize laboratory adaptation and implement outcrossing to increase the relevance of experiments to field mosquitoes. Additionally, colonies are maintained under optimal conditions to maximize their fitness for open field releases.

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes infected with Wolbachia are being released into natural populations to suppress the transmission of arboviruses. We describe methods to rear Ae. aegypti with Wolbachia infections in the laboratory for experiments and field release, taking precautions to minimize laboratory adaptation and selection.

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, ...

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 ...

Effect of Fluorescent Proteins on Fusion Partners Using Polyglutamine Toxicity Assays in Yeast

For the investigation of protein localization and trafficking using live cell imaging, researchers often rely on fusing their protein of interest to a fluorescent reporter. The constantly evolving list of genetically encoded fluorescent proteins (FPs) presents users with several alternatives when it comes to fluorescent fusion design. Each FP has specific optical and biophysical properties that can affect the biochemical, cellular, and functional properties of the resulting fluorescent fusions. For instance, several FPs tend to form nonspecific oligomers that are susceptible to impede on the function of the fusion partner. Unfortunately, only a few methods exist to test the impact of FPs on the behavior of the fluorescent reporter. Here, we describe a simple method that enables the rapid assessment of the impact of FPs using polyglutamine (polyQ) toxicity assays in the budding yeast Saccharomyces cerevisiae. PolyQ-expanded huntingtin proteins are associated with the onset of Huntington&#39;s disease (HD), where the expanded huntingtin aggregates into toxic oligomers and inclusion bodies. The aggregation and toxicity of polyQ expansions in yeast are highly dependent on the sequences flanking the polyQ region, including the presence of fluorescent tags, thus providing an ideal experimental platform to study the impact of FPs on the behavior of their fusion partner.

This article describes protocols to assess the effect of fluorescent proteins on the aggregation and toxicity of misfolded polyglutamine expansion for the rapid evaluation of a newly uncharacterized fluorescent protein in the context of fluorescent reporters.

For the investigation of protein localization and trafficking using live cell imaging, researchers often rely on fusing their protein of interest to a ...

Improved Fecundity and Fertility Assay for Aedes aegypti using 24 Well Tissue Culture Plates (EAgaL Plates)

Mosquitoes represent a significant public health problem as vectors of various pathogens. For those studies that require an assessment of mosquito fitness parameters, in particular egg production and hatch rates at the individual level, conventional methods have put a substantial burden on investigators due to high labor intensity and laboratory space requirements. Described is a simple method using 24 well tissue culture plate with agarose in each well and digital imaging of each well to determine egg numbers and hatch rates at an individual level with substantially reduced time and space requirements.

Improved Fecundity and Fertility Assay for Aedes aegypti using 24 Well Tissue Culture Plates EAgaL Plates

Described is a time and space-saving method to count eggs and determine hatch rates of individual mosquitoes using 24 well tissue culture plates, which can substantially increase the scale and speed of fecundity and fertility assays.

Mosquitoes represent a significant public health problem as vectors of various pathogens. For those studies that require an assessment of mosquito fitness ...