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><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>Mosquito cages</td><td>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 &amp;micro;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 &amp;micro;m aperture&lt;br/&gt; For mating competitiveness assay</td><td></td><td></td></tr><tr><td>Mosquito netting</td><td></td><td></td><td>150 x 150, 160 &amp;micro;m aperture</td></tr><tr><td>Rhodamine B</td><td>Sigma Aldrich</td><td>R6626</td><td>&amp;ge;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 ...

using-fluorescent-dye-rhodamine-b-to-study-mating-competitiveness

Research

JoVE Journal

Biology

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

Mass Production of Genetically Modified Aedes aegypti for Field Releases in Brazil

New techniques and methods are being sought to try to win the battle against mosquitoes. Recent advances in molecular techniques have led to the development of new and innovative methods of mosquito control based around the Sterile Insect Technique (SIT)1-3. A control method known as RIDL&#160;(Release of Insects carrying a Dominant Lethal)4, is based around SIT, but uses genetic methods to remove the need for radiation-sterilization5-8. A RIDL&#160;strain of Ae. aegypti was successfully tested in the field in Grand Cayman9,10; further field use is planned or in progress in other countries around the world.
Mass rearing of insects has been established in several insect species and to levels of billions a week. However, in mosquitoes, rearing has generally been performed on a much smaller scale, with most large scale rearing being performed in the 1970s and 80s. For a RIDL&#160;program it is desirable to release as few females as possible as they bite and transmit disease. In a mass rearing program&#160;there are several stages to produce the males to be released: egg production, rearing eggs until pupation, and then sorting males from females before release. These males are then used for a RIDL&#160;control program,&#160;released as either pupae or adults11,12.
To suppress a mosquito population using RIDL&#160;a large number of high quality male adults need to be reared13,14. The following describes the methods for the mass rearing of OX513A, a RIDL&#160;strain of Ae. aegypti 8, for release and covers the techniques required for the production of eggs and mass rearing RIDL&#160;males for a control program.

Mass Production of Genetically Modified Aedes aegypti for Field Releases in Brazil

To achieve population suppression of Aedes aegypti using the RIDL&#174; (Release of Insects carrying a Dominant Lethal) system, large numbers of male mosquitoes need to be released. This requires the use of mass rearing techniques and technology to provide reliable systems to obtain the maximum number of high quality male mosquitoes.

New techniques and methods are being sought to try to win the battle against mosquitoes. Recent advances in molecular techniques have led to the ...

Environment

A Method to Test the Effect of Environmental Cues on Mating Behavior in Drosophila melanogaster

An individual's sexual drive is influenced by genotype, experience and environmental conditions. How these factors interact to modulate sexual behaviors remains poorly understood. In Drosophila melanogaster, environmental cues, such as food availability, affect mating activity offering a tractable system to investigate the mechanisms modulating sexual behavior. In D. melanogaster, environmental cues are often sensed via the chemosensory gustatory and olfactory systems. Here, we present a method to test the effect of environmental chemical cues on mating behavior. The assay consists of a small mating arena containing food medium and a mating couple. The mating frequency for each couple is continuously monitored for 24 h. Here we present the applicability of this assay to test environmental compounds from an external source through a pressurized air system as well as manipulation of the environmental components directly in the mating arena. The use of a pressurized air system is especially useful to test the effect of very volatile compounds, while manipulating components directly in the mating arena can be of value to ascertain a compound's presence. This assay can be adapted to answer questions about the influence of genetic and environmental cues on mating behavior and fecundity as well as other male and female reproductive behaviors.

A Method to Test the Effect of Environmental Cues on Mating Behavior in Drosophila melanogaster

We demonstrate an assay to analyze the environmental and genetic cues that influence mating behavior in the fruit fly Drosophila melanogaster.

An individual's sexual drive is influenced by genotype, experience and environmental conditions. How these factors interact to modulate sexual behaviors ...

Behavior

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.

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 ...

Methods to Assay Drosophila Behavior

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular disease and various neurological diseases1. We have optimized simple and robust behavioral assays for determining larval locomotion, adult climbing ability (RING assay), and courtship behaviors of Drosophila. These behavioral assays are widely applicable for studying the role of genetic and environmental factors on fly behavior. Larval crawling ability can be reliably used for determining early stage changes in the crawling abilities of Drosophila larvae and also for examining effect of drugs or human disease genes (in transgenic flies) on their locomotion. The larval crawling assay becomes more applicable if expression or abolition of a gene causes lethality in pupal or adult stages, as these flies do not survive to adulthood where they otherwise could be assessed. This basic assay can also be used in conjunction with bright light or stress to examine additional behavioral responses in Drosophila larvae. Courtship behavior has been widely used to investigate genetic basis of sexual behavior, and can also be used to examine activity and coordination, as well as learning and memory. Drosophila courtship behavior involves the exchange of various sensory stimuli including visual, auditory, and chemosensory signals between males and females that lead to a complex series of well characterized motor behaviors culminating in successful copulation. Traditional adult climbing assays (negative geotaxis) are tedious, labor intensive, and time consuming, with significant variation between different trials2-4. The rapid iterative negative geotaxis (RING) assay5 has many advantages over more widely employed protocols, providing a reproducible, sensitive, and high throughput approach to quantify adult locomotor and negative geotaxis behaviors. In the RING assay, several genotypes or drug treatments can be tested simultaneously using large number of animals, with the high-throughput approach making it more amenable for screening experiments.

Methods to Assay Drosophila Behavior

Drosophila melanogaster is a genetically and behaviorally tractable model system that has been used to understand the molecular and cellular basis of many important biological processes for over a century 1. Drosophila has been well exploited to gain insights into the genetic basis of fly behavior.

Drosophila melanogaster, the fruit fly, has been used to study molecular mechanisms of a wide range of human diseases such as cancer, cardiovascular ...

Neuroscience

Measuring Sperm Guidance and Motility within the Caenorhabditis elegans Hermaphrodite Reproductive Tract

Successful fertilization is fundamental to sexual reproduction, yet little is known about the mechanisms that guide sperm to oocytes within the female reproductive tract. While in vitro studies suggest that sperm of internally fertilizing animals can respond to various cues from their surroundings, the inability to visualize their behavior inside the female reproductive tract creates a challenge for understanding sperm migration and mobility in its native environment. Here, we describe a method using C. elegans that overcomes this limitation and takes advantage of their transparent epidermis. C. elegans males stained with a mitochondrial&#160; dye are mated with adult hermaphrodites, which act as modified females, and deposit fluorescently labeled sperm into the hermaphrodite uterus. The migration and motility of the labeled sperm can then be directly tracked using an epi-fluorescence microscope in a live hermaphrodite. In wild-type animals, approximately 90% of the labeled sperm crawl through the uterus and reach the fertilization site, or spermatheca. Images of the uterus can be taken 1 h after mating to assess the distribution of the sperm within the uterus and the percentage of sperm that have reached the spermatheca. Alternatively, time-lapse images can be taken immediately after mating to assess sperm speed, directional velocity and reversal frequency. This method can be combined with other genetic and molecular tools available for the C.elegans to identify novel genetic and molecular mechanisms that are important in regulating sperm guidance and motility within the female reproductive tract.

Measuring Sperm Guidance and Motility within the Caenorhabditis elegans Hermaphrodite Reproductive Tract

Sperm must successfully navigate through the oviduct to fertilize an oocyte. Here, we describe an assay for measuring sperm migration within the C. elegans hermaphrodite uterus. This assay can provide quantitative data on sperm distribution within the uterus after mating, as well as on speed, directional velocity, and reversal frequency.

Successful fertilization is fundamental to sexual reproduction, yet little is known about the mechanisms that guide sperm to oocytes within the female ...

Developmental Biology

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy and cheap methods to distinguish between individuals in a trial. A standard technique for this is CO2 anaesthesia and then labelling or wing clipping each fly. However, this is invasive and has been shown to affect behavior. Other techniques have used coloration to identify flies. This article presents a simple and non-invasive method for labelling Drosophila that allows them to be individually identified within experiments, using food coloring. This method is used in trials where two males compete to mate with a female. Dyeing allowed quick and easy identification. There was, however, some difference in the strength of the coloration across the three species tested. Data is presented showing the dye has a lower impact on mating behavior than CO2 in Drosophila melanogaster. The impact of CO2 anaesthesia is shown to depend on the species of Drosophila, with D. pseudoobscura and D. subobscura showing no impact, whereas D. melanogaster males had reduced mating success. The dye method presented is applicable to a wide range of experimental designs.

Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

This protocol describes a simple, cost effective way to individually identify Drosophila or other insects. Demonstration data investigating mating success across three species of Drosophila show that this method is comparable or better than the use of CO2 anaesthesia.

Mating experiments using Drosophila have contributed greatly to the understanding of sexual selection and behavior. Experiments often require simple, easy ...

Measuring and Altering Mating Drive in Male Drosophila melanogaster

Despite decades of investigation, the neuronal and molecular bases of motivational states remain mysterious. We have recently developed a novel, reductionist, and scalable system for in-depth investigation of motivation using the mating drive of male Drosophila&#160;melanogaster (Drosophila), the methods for which we detail here. The behavioral paradigm centers on the finding that male mating drive decreases alongside fertility over the course of repeated copulations and recovers over ~3 d. In this system, the powerful neurogenetic tools available in the fly converge with the genetic accessibility and putative wiring diagram available for sexual behavior. This convergence allows rapid isolation and interrogation of small neuronal populations with specific motivational functions. Here we detail the design and execution of the satiety assay that is used to measure and alter courtship motivation in the male fly. Using this assay, we also demonstrate that low male mating drive can be overcome by stimulating dopaminergic neurons. The satiety assay is simple, affordable, and robust to influences of genetic background. We expect the satiety assay to generate many new insights into the neurobiology of motivational states.

Measuring and Altering Mating Drive in Male Drosophila melanogaster

This article describes a behavioral assay that uses male mating drive in Drosophila melanogaster to study motivation. Using this method, researchers can utilize advanced fly neurogenetic techniques to uncover the genetic, molecular, and cellular mechanisms that underlie this motivation.

Despite decades of investigation, the neuronal and molecular bases of motivational states remain mysterious. We have recently developed a novel, ...

Assessing Differences in Sperm Competitive Ability in Drosophila

Competition among conspecific males for fertilizing the ova is one of the mechanisms of sexual selection, i.e. selection that operates on maximizing the number of successful mating events rather than on maximizing survival and viability 1. Sperm competition represents the competition between males after copulating with the same female 2, in which their sperm are coincidental in time and space. This phenomenon has been reported in multiple species of plants and animals 3. For example, wild-caught D. melanogaster females usually contain sperm from 2-3 males 4. The sperm are stored in specialized organs with limited storage capacity, which might lead to the direct competition of the sperm from different males 2,5.
Comparing sperm competitive ability of different males of interest (experimental male types) has been performed through controlled double-mating experiments in the laboratory 6,7. Briefly, a single female is exposed to two different males consecutively, one experimental male and one cross-mating reference male. The same mating scheme is then followed using other experimental male types thus facilitating the indirect comparison of the competitive ability of their sperm through a common reference. The fraction of individuals fathered by the experimental and reference males is identified using markers, which allows one to estimate sperm competitive ability using simple mathematical expressions 7,8. In addition, sperm competitive ability can be estimated in two different scenarios depending on whether the experimental male is second or first to mate (offense and defense assay, respectively) 9, which is assumed to be reflective of different competence attributes.
Here, we describe an approach that helps to interrogate the role of different genetic factors that putatively underlie the phenomenon of sperm competitive ability in D. melanogaster.

Assessing Differences in Sperm Competitive Ability in Drosophila

Differential sperm competitive ability among Drosophila males with distinct genotypes can be ascertained through double-mating experiments. Each of these experiments involves one of the males of interest and a reference male. Readily identifiable markers in the progeny allow inference of the fraction of individuals fathered by each male.

Competition among conspecific males for fertilizing the ova is one of the mechanisms of sexual selection, i.e. selection that operates on maximizing the ...

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 ...

Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes

Transgenic mosquitoes often display fitness costs compared to their wild-type counterparts. In this regard, fitness cost studies involve collecting life parameter data from genetically modified mosquitoes and comparing them to mosquitoes lacking transgenes from the same genetic background. This manuscript illustrates how to measure common life history traits in the mosquito Aedes aegypti, including fecundity, wing size and shape, fertility, sex ratio, viability, development times, male contribution, and adult longevity. These parameters were chosen because they reflect reproductive success, are simple to measure, and are commonly reported in the literature. The representative results quantify fitness costs associated with either a gene knock-out or a single insertion of a gene drive element. Standardizing how life parameter data are collected is important because such data may be used to compare the health of transgenic mosquitoes generated across studies or to model the transgene fixation rate in a simulated wild-type mosquito population. Although this protocol is specific for transgenic Aedes aegypti, the protocol may also be used for other mosquito species or other experimental treatment conditions, with the caveat that certain biological contexts may require special adaptations.

Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes

The present protocol describes how to measure common life parameter data in Aedes aegypti mosquitoes, including fecundity, wing size, fertility, sex ratio, viability, development times, male contribution, and adult longevity. These measurements can be used to assess the fitness of transgenic mosquitoes.

Transgenic mosquitoes often display fitness costs compared to their wild-type counterparts. In this regard, fitness cost studies involve collecting life ...

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

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Dyeing Insects for Behavioral Assays: the Mating Behavior of Anesthetized Drosophila

Measuring and Altering Mating Drive in Male Drosophila melanogaster

Assessing Differences in Sperm Competitive Ability in Drosophila

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

Quantifying Fitness Costs in Transgenic Aedes aegypti Mosquitoes