We wish to thank Spencer Blankenship for help with mosquito rearing. We also thank James N. Radl and M. Dominique Magistrado for helpful feedback on the manuscript.

1. Day 0
<ol>
	<li>Hatch Ae. aegypti eggs infected with E. aedis by placing in larval rearing tray with 1 L deionized (DI) water. Add 50 mg of fish food. 
	NOTE: At the time of publication, a laboratory strain of E. aedis is only available from laboratories actively researching the parasite, as E. aedis is not amenable to long-term storage and infected eggs are not currently stored in repositories. Researchers interested in working with E. aedis can contact the corresponding author to request infected eggs. 
	NOTE:&#160;Hatching large numbers of infected eggs is generally not necessary; ten E. aedis infected Ae. aegypti larvae are sufficient to dose &#8805; 1000 healthy larvae. 
	NOTE:&#160;For all parts of this protocol, we housed mosquitoes at the following conditions: 14 h/10 h light/ dark cycle, 27 &#176;C temperature and 80% relative humidity.</li>
</ol>
2. Day 1
<ol>
	<li>After hatching, reduce density of larvae to ~100 larvae per tray, making new trays as necessary (also with 1 L DI water).</li>
	<li>Add a piece of dry cat food to each tray. Replenish food when depleted, but do not provide an excess of food. One piece of cat food (~200 mg) every three days is sufficient. 
	NOTE: Adjust food amount depending on the specific rearing conditions (i.e., reduce food if water becomes turbid or larvae are dying, increase food if larvae are severely delayed in development). Other feeding regimens and/or rearing conditions than those suggested here can be used but adjustments to timing of this standard protocol may be needed.</li>
</ol>
3. Days 4&#8210;5
<ol>
	<li>When infected larvae are 3rd &#8211; 4th instars, hatch healthy/uninfected Ae. aegypti eggs in a new tray.</li>
	<li>Rear at densities such that healthy Ae. aegypti reach 2nd &#8211; 3rd instar in 48&#8211;72 h. In our hands, this can be achieved using densities of 200&#8210;300 larvae per 1 L of water with ad libitum access to food. Hatching batches of healthy eggs over multiple days can guarantee that larvae are at the correct stage when needed.</li>
</ol>
4. Days 7&#8210;8: Horizontal transmission
NOTE: Dosing of healthy larvae with E. aedis cannot be performed until uninucleate spores are at high numbers in infected larvae (1 x 104 &#8211; 1 x 106 per larva). This occurs late in the 4th instar stage (Figure 2).
<ol>
	<li>Harvest and quantify uninucleate spores.
	<ol>
		<li>Use a transfer pipette (one may have to cut the tip to a wider diameter) to move 10 infected larvae to a 1.5 mL microcentrifuge tube.</li>
		<li>Remove breeding water with a transfer&#160;pipet and wash once by adding ~1 mL of clean DI water. Remove the wash water with a pipet, add 500 &#181;L of clean DI water to the 10 larvae, and homogenize using a pestle and mechanical homogenizer.</li>
		<li>Quantify spores using a hemocytometer at 400x magnification. 
		NOTE: Uninucleate spores can be identified by their distinct pyriform shape (i.e., pear shape; Figure 2A).</li>
	</ol>
	</li>
	<li>Dose healthy Ae. aegypti larvae with E. aedis.
	<ol>
		<li>Make fresh larval food slurry by mixing 1.2 g of liver powder, 0.8 g of brewer&#8217;s yeast and 100 mL of water. 
		NOTE: Food does not need to be fresh if it is autoclaved and stored at 4 &#176;C until use.</li>
		<li>Transfer 100 2nd &#8211; 3rd instar healthy Ae. aegypti larvae into 150 mL beakers or specimen cups.</li>
		<li>Dose each beaker of 100 larvae with 5 x 104 &#8211; 1 x 105 spores.</li>
		<li>Add 2 mL of larval food slurry and DI water to a final volume of 100 mL.</li>
	</ol>
	</li>
	<li>After 12&#8211;24 h of exposure, transfer exposed larvae into rearing trays and rear to adulthood following a standard rearing protocol16.</li>
</ol>
5. Monitoring and vertical transmission
<ol>
	<li>Monitor dosed larvae for pupation and transfer pupae as they develop to an emergence cup in a cage. Sugar feed adults ad libitum (as per16,17). Eclosing adults will be infected by E. aedis.</li>
	<li>Blood feed adults (as per16,17) and collect eggs. Vertical transmission of E. aedis occurs at this step. 
	NOTE: If additional blood meals are provided as soon as oviposition is complete, females can lay at least one additional clutch of eggs before adults suffer (often sudden) high levels of mortality.&#160;</li>
	<li>Use these Ae. aegypti eggs infected with E. aedis to continue propagation starting with step 1 of this protocol. 
	NOTE: Eggs can be stored for 2 &#8211; 3 months under appropriate conditions16.</li>
	<li>Clean all materials that came in contact with E. aedis with 10% bleach and autoclaving (if possible) to prevent contamination.</li>
</ol>

<ol>
	<li>Yellow fever. World Health Organization Available from: <a target="_blank" href="https://www.who.int/en/news-room/fact-sheets/detail/yellow-fever">https://www.who.int/en/news-room/fact-sheets/detail/yellow-fever</a> (2019)</li><li>Dengue and severe dengue. World Health Organization Available from: <a target="_blank" href="https://www.who.int/en/news-room/fact-sheets/detail/dengue-and-severe-dengue">https://www.who.int/en/news-room/fact-sheets/detail/dengue-and-severe-dengue</a> (2020)</li><li>Weaver, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prediction+and+prevention+of+urban+arbovirus+epidemics+:+A+challenge+for+the+global+virology+community.">Prediction and prevention of urban arbovirus epidemics : A challenge for the global virology community.</a> Antiviral Research. 156, 80-84 (2018).</li><li>Rather, I. A., Parray, H. A., Lone, J. B., Paek, W. K., Lim, J., Bajpai, V. K., Park, Y. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prevention+and+Control+Strategies+to+Counter+Dengue+Virus+Infection.">Prevention and Control Strategies to Counter Dengue Virus Infection.</a> Frontiers In Cellular and Infection Microbiology. 7, 336 (2017).</li><li>Becnel, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Edhazardia+aedis+(Microsporidia:+Amblysporidae)+as+a+biocontrol+agent+of+Aedes+aegypti+(Diptera:+Culicidae).">Edhazardia aedis (Microsporidia: Amblysporidae) as a biocontrol agent of Aedes aegypti (Diptera: Culicidae).</a> Proceedings and abstracts, Vth International Colloquium on Invertebrate Pathology and Microbial Control. , 20-24 (1990).</li><li>Becnel, J. J., Johnson, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impact+of+Edhazardia+aedis+(Microsporidia:+Culicosporidae)+on+a+seminatural+population+of+Aedes+aegypti+(Diptera:+Culicidae).">Impact of Edhazardia aedis (Microsporidia: Culicosporidae) on a seminatural population of Aedes aegypti (Diptera: Culicidae).</a> Biological Control. 18 (1), 39-48 (2000).</li><li>Becnel, J. J., Sprague, V., Fukuda, T., Hazard, E. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+Edhazardia+aedis+(Kudo,+1930)+N.+G.,+N.+Comb.+(Microsporida:+Amblyosporidae)+in+the+mosquito+Aedes+aegypti+(L.)+(Diptera:+Culicidae).">Development of Edhazardia aedis (Kudo, 1930) N. G., N. Comb. (Microsporida: Amblyosporidae) in the mosquito Aedes aegypti (L.) (Diptera: Culicidae).</a> Journal of Protozoology. 36, 119-130 (1989).</li><li>Hembree, S. C., Ryan, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Observations+on+the+vertical+transmission+of+a+new+microsporidian+pathogen+of+Aedes+aegypti+from+Thailand.">Observations on the vertical transmission of a new microsporidian pathogen of Aedes aegypti from Thailand.</a> Mosquito News. 42, 49-54 (1982).</li><li>Johnson, M. A., Becnel, J. J., Undeen, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+sporulation+sequence+in+Edhazardia+aedis+(Microsporidia:+Culicosporidae),+a+parasite+of+the+mosquito+Aedes+aegypti+(Diptera:+Culicidae).">A new sporulation sequence in Edhazardia aedis (Microsporidia: Culicosporidae), a parasite of the mosquito Aedes aegypti (Diptera: Culicidae).</a> Journal of Invertebrate Pathology. 70 (1), 69-75 (1997).</li><li>Becnel, J. J., Garcia, J. J., Johnson, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Edhazardia+aedis+(Microspora:+Culicosporidae)+effects+on+the+reproductive+capacity+of+Aedes+aegypti+(Diptera:+Culicidae).">Edhazardia aedis (Microspora: Culicosporidae) effects on the reproductive capacity of Aedes aegypti (Diptera: Culicidae).</a> Journal of Medical Entomology. 32 (4), 549-553 (1995).</li><li>Becnel, J. J., Johnson, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosquito+host+range+and+specificity+of+Edhazardia+aedis+(Microspora:+Culicosporidae).">Mosquito host range and specificity of Edhazardia aedis (Microspora: Culicosporidae).</a> Journal of the American Mosquito Control Association. 9 (3), 269-274 (1993).</li><li>Andreadis, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Host+range+tests+with+Edhazardia+aedis+(Microsporida:+Culicosporidae)+against+northern+Nearctic+mosquitoes.">Host range tests with Edhazardia aedis (Microsporida: Culicosporidae) against northern Nearctic mosquitoes.</a> Journal of Invertebrate Pathology. 64 (1), 46-51 (1994).</li><li>Becnel, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Safety+of+Edhazardia+aedis+(Microspora:+Amblyosporidae)+for+nontarget+aquatic+organisms.">Safety of Edhazardia aedis (Microspora: Amblyosporidae) for nontarget aquatic organisms.</a> Journal of the American Mosquito Control Association. 8 (3), 256-260 (1992).</li><li>Undeen, A. H., Becnel, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Longevity+and+germination+of+Edhazardia+aedis+(Microspora:+Amblyosporidae)+spores.">Longevity and germination of Edhazardia aedis (Microspora: Amblyosporidae) spores.</a> Biocontrol Science and Technology. 2, 247-256 (1992).</li><li>Massaro, P., Sobecki, R., Behling, C., Criswell, V., Zha, T., Devenzengo, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T.">T.</a> Automated mass rearing system for insect larvae. , (2018).</li><li>Methods in Aedes Research. BEI Resources Available from: <a target="_blank" href="https://www.beiresources.org/Portals/2/VectorResources/Methods_20in_20Aedes_20Research_202016.pdf">https://www.beiresources.org/Portals/2/VectorResources/Methods_20in_20Aedes_20Research_202016.pdf</a> (2016)</li><li>Methods in Anopheles Research. BEI Resources Available from: <a target="_blank" href="https://www.beiresources.org/portals/2/MR4/MR4_Publications/Methods_20in_20Anopheles_20Research_202014/2014MethodsinAnophelesResearchManualFullVersionv2tso.pdf">https://www.beiresources.org/portals/2/MR4/MR4_Publications/Methods_20in_20Anopheles_20Research_202014/2014MethodsinAnophelesResearchManualFullVersionv2tso.pdf</a> (2014)</li><li>Desjardins, C. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contrasting+host-pathogen+interactions+and+genome+evolution+in+two+generalist+and+specialist+microsporidian+pathogens+of+mosquitoes.">Contrasting host-pathogen interactions and genome evolution in two generalist and specialist microsporidian pathogens of mosquitoes.</a> Nature Communications. 6 (1), 1-12 (2015).</li><li>Hembree, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preliminary+Report+of+some+mosquito+pathogens+from+Thailand.">Preliminary Report of some mosquito pathogens from Thailand.</a> Mosquito News. 39 (3), 575-582 (1979).</li><li>Solter, L. F., Becnel, J. J., V&#225;vra, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Research+methods+for+entomopathogenic+microsporidia+and+other+protists.">Research methods for entomopathogenic microsporidia and other protists.</a> Manual of Techniques in Invertebrate Pathology. , 329-371 (2012).</li><li>Becnel, J. J., Undeen, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+temperature+on+developmental+parameters+of+the+parasite/host+System+Edhazardia+aedis+(Microsporidia:+Amblyosporidae)+and+Aedes+aegypti+(Diptera:+Culicidae).">Influence of temperature on developmental parameters of the parasite/host System Edhazardia aedis (Microsporidia: Amblyosporidae) and Aedes aegypti (Diptera: Culicidae).</a> Journal of Invertebrate Pathology. 60, 299-303 (1992).</li></ol>

The authors have nothing to disclose.

We present here the method originally described in Hembree and Ryan, 19828 for rearing E. aedis microsporidia in Ae. aegypti mosquitoes. The strain of E. aedis used in this study was derived from the original field collection by Stephen Hembree in Thailand in 197919. The method capitalizes on horizontal transmission, which naturally occurs in the transmission cycle of E. aedis7, to propagate the parasite in a controlled manner. This method can be challenging to newcomers who are not familiar with spore appearance, symptoms of infection in larvae, or the coordination required to successfully complete the multi-stage rearing/dosing protocol. Our hope is that the visual aids that accompany this protocol will reduce barriers to entry for researchers who wish to culture E. aedis.
We propagated E. aedis in Ae. aegypti as described above and quantified the success of parasitism in the filial generation. Briefly, we hatched E. aedis infected Ae. aegypti eggs, reared them to 4th instar, and collected uninucleate E. aedis spores from the infected larvae. We then horizontally infected healthy larvae with these spores via oral ingestion, and reared the horizontally infected larvae to adulthood. We blood fed the infected adults (parental generation) and collected eggs (filial generation), which we hypothesized would be vertically infected with the E. aedis parasite. We hatched eggs from the filial generation and collected and homogenized a subset of the larvae when they were 4th instars. We quantified the percent of larvae that were infected with E. aedis and the total spore count in all infected individuals. We found that the vast majority (96%) of individuals were infected and the mean spore load of infected larvae was ~105. We conclude that our rearing protocol resulted in highly successful propagation of E. aedis in Ae. aegypti mosquitoes.
There are multiple aspects of this protocol that may be particularly challenging for the uninitiated user. We offer below some additional information that may be of assistance. For questions regarding general mosquito rearing, a complete guide to Ae. aegypti colony maintenance is beyond the scope of this protocol. However, many common questions can be addressed by resources from the Biodefense and Emerging Infections Research Resources Repository16,17 including egg hatching, general dietary needs, housing and environmental conditions, and blood feeding. Regarding the timeline of infection, larvae hatched from infected eggs do not show signs of infection until late in the 4th instar stage. Uninucleate spores appear rapidly, over the course of 1&#8211;2 days. Larvae may appear virtually uninfected at 6 days post-hatching but highly infected by day 7 or 8 post-hatching. Additionally, it can be challenging to visualize spores in homogenized samples because there are many other microbes present in whole mosquito homogenates, including other eukaryotic single-celled organisms (e.g., yeast) of a similar size as the E. aedis uninucleate spores. The distinctive shape of E. aedis spores (Figure 2A) is a highly reliable method for identification and will help differentiate E. aedis from other microbes in the homogenate. Though it is not necessary for identification or quantification, if spore purification is desired, it can be achieved via colloidal silica density gradient centrifugation which will allow for separation of E. aedis spores from other contaminating elements in the homogenate. This process is described in detail in Solter et al.20.
Temperature and diet used in rearing practices commonly differ between laboratories, but variations will likely still yield successful parasite propagation. Minor differences in larval food type do not interfere with successful infection, though we did not explicitly test different food types in this protocol. The effect of temperature on infection has been tested and E. aedis infection was found to be robust at a wide range of temperatures21. Maximum spore production occurred at 30.8 &#176;C but was still robust at rearing temperatures as low as 20 &#176;C. Spore count was reduced dramatically at higher rearing temperatures (36 &#176;C), therefore these temperatures should be avoided for this protocol.
Contamination is always a concern when working with parasites. E. aedis is a successful parasite of Ae. aegypti and must therefore be kept separate from uninfected laboratory colonies to prevent contamination. We recommend storage of infected mosquitoes in a separate incubator if possible. We also recommended that materials used for microsporidia work (e.g., larval trays, transfer pipets, cages, egg collection cups) are designated for microsporidia work and not used more broadly throughout the insectary. All rearing materials should be sterilized with 10% bleach after use and autoclaving can be used to supplement bleach sterilization.

Aedes aegypti is the mosquito vector of multiple arboviruses (e.g., dengue, Zika, yellow fever) that together are estimated to account for hundreds of millions of disease cases each year and more than 30,000 deaths1,2. Treatment for diseases caused by these pathogens is limited to supportive care and it is likely that additional arboviruses will emerge in the future3. Control of the mosquito vector is therefore of primary importance, as it effectively prevents transmission of current and emerging pathogens4. Traditionally, vector control strategies primarily utilize chemical insecticides, but resistance to many commonly used insecticides has driven the demand for novel methods of vector control. One potential agent that has been explored for its biocontrol properties against Ae. aegypti is the parasite Edhazardia aedis5,6.
E. aedis, first identified as Nosema aedis by Kudo in 1930, is a microsporidian parasite of Ae. aegypti mosquitoes7. The development and reproduction of E. aedis is relatively complex and its life cycle can proceed in multiple ways7,8,9. One common developmental cycle is described in depth in Becnel et al., 19897 and is utilized for laboratory propagation (Figure 1)8. Briefly, the cycle begins when Ae. aegypti eggs vertically infected with E. aedis hatch into infected larvae which develop uninucleate spores in the fat body, and usually die as larvae or pupae. Uninucleate spores released from dead larvae contaminate the habitat and are ingested by healthy Ae. aegypti larvae. These spores germinate primarily in the digestive tract, infecting digestive tissue of the exposed larvae, resulting in horizontal transmission. Horizontally infected larvae develop into adults (parental generation) where binucleate spores are formed. In the female, these binucleate spores invade the reproductive tract and their associated sporoplasm infects developing egg cells. These eggs then hatch into infected larvae (filial generation), resulting in vertical transmission of the parasite and continuation of the cycle as described above.
Multiple studies have investigated the potential of E. aedis for biocontrol. Infection with E. aedis has been demonstrated to result in diminished reproductive capacity of Ae. aegypti females10. Further, in a semi-field experiment, inundative release of E. aedis resulted in the total eradication of a test Ae. aegypti population kept within a screened enclosure6. While able to undergo some stages of development in a diverse set of mosquito species, E. aedis is only vertically transmitted in Ae. aegypti, indicating a high degree of host specificity11,12. Likewise, in a laboratory assessment of the potential environmental risk associated with E. aedis, the microsporidian parasite failed to infect non-target aquatic fauna, including predators that ingested Ae. aegypti larvae infected with E. aedis13. These results highlight the potential for E. aedis to be used in biological control strategies targeting natural Ae. aegypti populations.
Despite the fact that E. aedis shows promise for use in vector control, there are challenges to culturing and deploying it on a broad scale. E. aedis spores lose infectivity in less than one day at cold temperatures (i.e., 5 &#176;C). Even at warmer temperatures (i.e., 25 &#176;C), spores rapidly lose infectivity over the course of three weeks14. Additionally, E. aedis must be cultured in live Ae. aegypti mosquitoes and controlled dosing of healthy larval mosquitoes is necessary to ensure completion of the life cycle and to prevent collapse of the population used for culture8. The requirement of in vivo culturing presents a challenge; however, recent advances in mosquito mass rearing and robotics (e.g., Massaro et al.15) could allow for large-scale generation of E. aedis spores. We anticipate that visualization of this methodology will increase accessibility to the E. aedis rearing protocol and allow more researchers to investigate the basic biology and applied potential of this system. We also anticipate that it will facilitate increased collaborations with engineers, roboticists, and the broader technology sector, which may serve to improve mass rearing of E. aedis.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61574/61574fig01v2.jpg" /> 
Figure 1: E. aedis propagation in Ae. aegypti.&#160;Propagation of E. aedis begins with hatching E. aedis infected eggs. Infected larvae are reared to 4th instar, E. aedis spores are isolated from those larvae, and the spores are used to orally infect healthy 2nd/3rd instar larvae reared from an uninfected clutch of eggs (horizontal transmission). These orally infected larvae are then reared to adulthood (parental generation) and lay eggs infected with E. aedis (vertical transmission). Infected eggs (filial generation) are then hatched to continue the infection cycle and parasite culture. <a href="https://www.jove.com/files/ftp_upload/61574/61574fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table>
	<tbody>
		<tr>
			<td>120 mL Specimen cup</td>
			<td>McKesson</td>
			<td>911759</td>
			<td>Inexpensive alternative to beaker</td>
		</tr>
		<tr>
			<td>150 mL beakers</td>
			<td>VWR</td>
			<td>10754-950</td>
			<td>For larval dosing</td>
		</tr>
		<tr>
			<td>2 oz round glass bottle</td>
			<td>VWR</td>
			<td>10862-502</td>
			<td>Bottle for 10% sucrose in adult cages</td>
		</tr>
		<tr>
			<td>3 oz. emergence cup</td>
			<td>Henry-Schein</td>
			<td>1201502</td>
			<td>For transfer of pupae to cage</td>
		</tr>
		<tr>
			<td>Adult mosquito cages</td>
			<td>Bioquip</td>
			<td>1462 or 1450ASV</td>
			<td>For adult housing</td>
		</tr>
		<tr>
			<td>Autoclave</td>
			<td></td>
			<td></td>
			<td>For sterilization</td>
		</tr>
		<tr>
			<td>Bleach</td>
			<td></td>
			<td></td>
			<td>For sterilization</td>
		</tr>
		<tr>
			<td>Brewer&#8217;s yeast</td>
			<td>Solgar</td>
			<td></td>
			<td>For feeding larvae during dosing</td>
		</tr>
		<tr>
			<td>Controlled rearing chamber</td>
			<td>Tritech</td>
			<td>DT2-MP-47L</td>
			<td>Inexpensive small rearing chamber</td>
		</tr>
		<tr>
			<td>Cotton roll</td>
			<td>VWR</td>
			<td>470161-446</td>
			<td>Wick for sugar bottles</td>
		</tr>
		<tr>
			<td>Defibrinated rabbit blood</td>
			<td>Fisher</td>
			<td>50863762</td>
			<td>For blood feeding adults</td>
		</tr>
		<tr>
			<td>Disodium ATP, crystalline</td>
			<td>Sigma-Aldrich</td>
			<td>A26209-5G</td>
			<td>For blood feeding adults</td>
		</tr>
		<tr>
			<td>Dry cat food</td>
			<td>9Lives</td>
			<td>Indoor Complete</td>
			<td>For general larval rearing</td>
		</tr>
		<tr>
			<td>Fish food flakes</td>
			<td>TetraMin</td>
			<td></td>
			<td>For general larval rearing</td>
		</tr>
		<tr>
			<td>Hemocytometer</td>
			<td>Fisher</td>
			<td>267110</td>
			<td>For counting spores</td>
		</tr>
		<tr>
			<td>Homogenizer/mixer motor</td>
			<td>VWR</td>
			<td>47747-370</td>
			<td>For homogenizing infected larvae</td>
		</tr>
		<tr>
			<td>Larval rearing trays</td>
			<td>Sterillite</td>
			<td>1961</td>
			<td>Overall dimensions are 11&#34; x 6 5/8&#34; x 2 3/4&#34;</td>
		</tr>
		<tr>
			<td>Liver powder</td>
			<td>NOW foods</td>
			<td>2450</td>
			<td>For feeding larvae during dosing</td>
		</tr>
		<tr>
			<td>Pipette 1 - 10&#181;L</td>
			<td>VWR</td>
			<td>89079-962</td>
			<td>For larval dosing</td>
		</tr>
		<tr>
			<td>Pipette 100 - 1000&#181;L</td>
			<td>VWR</td>
			<td>89079-974</td>
			<td>For food during larval dosing</td>
		</tr>
		<tr>
			<td>Pipette tips 1 - 10&#181;L</td>
			<td>VWR</td>
			<td>10017-042</td>
			<td>For larval dosing</td>
		</tr>
		<tr>
			<td>Pipette tips 100 - 1000&#181;L</td>
			<td>VWR</td>
			<td>10017-048</td>
			<td>For food during larval dosing</td>
		</tr>
		<tr>
			<td>Plastic pestles</td>
			<td>VWR</td>
			<td>89093-446</td>
			<td>For homogenizing infected larvae</td>
		</tr>
		<tr>
			<td>Sucrose, crystalline</td>
			<td>Life Technologies</td>
			<td>15503022</td>
			<td>For adult feeding</td>
		</tr>
		<tr>
			<td>Transfer pipet</td>
			<td>VWR</td>
			<td>414004-033</td>
			<td>For larval transfer, must trim ends</td>
		</tr>
	</tbody>
</table>

propagation of the microsporidian parasite edhazardia aedis in aedes aegypti mosquitoes

Edhazardia aedis is a microsporidian parasite of Aedes aegypti mosquitoes, a disease vector that transmits multiple arboviruses which cause millions of disease cases each year. E. aedis causes mortality and reduced reproductive fitness in the mosquito vector and has been explored for its potential as a biocontrol agent. The protocol we present for culturing E. aedis is based on its natural infection cycle, which involves both horizontal and vertical transmission at different life stages of the mosquito host. Ae. aegypti mosquitoes are exposed to spores in the larval stage. These infected larvae then mature into adults and transmit the parasite vertically to their offspring. Infected offspring are then used as a source of spores for future horizontal transmission. Culturing E. aedis can be challenging to the uninitiated given the complexities of the parasite&#8217;s life cycle, and this protocol provides detailed guidance and visual aids for clarification.

E. aedis infected Ae. aegypti Liverpool (LVP1b12) eggs were hatched as described in the protocol above. In the 4th instar stage, visual signs of infection could be observed, including white spore cysts throughout the fat bodies of infected larvae (an example of this phenotype is shown in Figure 2B). Uninucleate spores were harvested from 4th instar larvae by homogenizing 10 larvae in 500 &#181;L DI water. These spores were pyriform (pear shaped) and readily visible at 400x (Figure 2A). Using a hemocytometer, a spore count of 4.05 x 103 spores/&#181;L was calculated. One hundred healthy Ae. aegypti larvae were then horizontally infected with ~50,000 spores in 100 mL water for a final dose of ~500 spores/larva. Larvae were reared to adulthood (parental generation) and blood fed using defibrinated rabbit blood plus 1% (v/v) 100 mM adenosine triphosphate. Vertically infected eggs were collected (filial generation) and hatched to continue E. aedis propagation and to quantify infection success.
At seven days post-hatching, 25 filial generation larvae were transferred into individual 1.5 mL microcentrifuge tubes and washed once with DI water. Individual larvae were homogenized in 250 &#181;L of DI water and infection status and E. aedis loads were assessed using a hemocytometer. Vertical infection rate of E. aedis in the filial generation was found to be 96% and the mean spore load of infected individuals at seven days post hatching was 3.31 x 105 (Range: 3.25 x 104 &#8211; 1.47 x 106; Figure 3).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61574/61574fig02v2.jpg" /> 
Figure 2: Visualization of E. aedis infection in Ae. aegypti mosquitoes.&#160;(A) E. aedis uninucleate pyriform spores. Ten E. aedis infected 4th instar larvae were homogenized in 500 &#181;L of DI water approximately seven days post hatching. 10 &#181;L of the homogenate was loaded onto a hemocytometer and viewed at 400X. Red arrows indicate representative uninucleate E. aedis spores. (B) E. aedis infected 4th instar larvae develop distinctive white spore cysts throughout their fat body18. They also commonly have malformed and distended abdominal segments. <a href="https://www.jove.com/files/ftp_upload/61574/61574fig02large.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/61574/61574fig03.jpg" /> 
Figure 3: Culture protocol leads to effective E. aedis infection in filial generation.&#160;Ae. aegypti larvae (n = 25) from the filial generation were homogenized individually in 250 &#181;L DI water and 10 &#181;L of the homogenate was loaded onto a hemocytometer. Presence of uninucleate spores indicated a positive infection and spores were quantified for all positive samples. (A) Prevalence of infection among filial larvae. Grey corresponds to uninfected larvae, and black to infected. Numbers displayed on each segment give the absolute count of individuals in each group. (B) Spore load for each infected individual. Black dots represent the log10 transformed uninucleate spore count for each larva. <a href="https://www.jove.com/files/ftp_upload/61574/61574fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes at JoVE.com

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

A protocol to culture the microsporidian parasite Edhazardia aedis. The parasite is passaged from one generation of Aedes aegypti mosquitoes to the next via horizontal transfer at the larval stage followed by vertical transmission at the adult stage. Live sporoplasms survive long-term in infected eggs.

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

Edhazardia aedis is a microsporidian parasite of Aedes aegypti mosquitoes, a disease vector that transmits multiple arboviruses which cause millions of ...

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Research

JoVE Journal

Immunology and Infection

4.8K Views.  The Ohio State University. A protocol to culture the microsporidian parasite Edhazardia aedis. The parasite is passaged from one generation of Aedes aegypti mosquitoes to the next via horizontal transfer at the larval stage followed by vertical transmission at the adult stage. Live sporoplasms survive long-term in infected eggs.

Video: Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito (Aedes aegypti) and Tick (Rhipicephalus microplus) G Protein-coupled Receptors

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes. The majority of them belong to the superfamily of G protein-coupled receptors (GPCRs). We have focused on characterizing arthropod kinin receptors from the tick and mosquito. Arthropod kinins are multifunctional neuropeptides with myotropic, diuretic, and neurotransmitter function. Here, a method for systematic analyses of structure-activity relationships of insect kinins on two heterologous kinin receptor-expressing systems is described. We provide important information relevant to the development of biostable kinin analogs with the potential to disrupt the diuretic, myotropic, and/or digestive processes in ticks and mosquitoes.
The kinin receptors from the southern cattle tick, Boophilus microplus (Canestrini), and the mosquito Aedes aegypti (Linnaeus), were stably expressed in the mammalian cell line CHO-K1. Functional analyses of these receptors were completed using a calcium bioluminescence plate assay that measures intracellular bioluminescence to determine cytoplasmic calcium levels upon peptide application to these recombinant cells. This method takes advantage of the aequorin protein, a photoprotein isolated from luminescent jellyfish. We transiently transfected the aequorin plasmid (mtAEQ/pcDNA1) in cell lines that stably expressed the kinin receptors. These cells were then treated with the cofactor coelenterazine, which complexes with intracellular aequorin. This bond breaks in the presence of calcium, emitting luminescence levels indicative of the calcium concentration. As the kinin receptor signals through the release of intracellular calcium, the intensity of the signal is related to the potency of the peptide.
This protocol is a synthesis of several previously described protocols with modifications; it presents step-by-step instructions for the stable expression of GPCRs in a mammalian cell line through functional plate assays (Staubly et al., 2002 and Stables et al., 1997). Using this methodology, we were able to establish stable cell lines expressing the mosquito and the tick kinin receptors, compare the potency of three mosquito kinins, identify critical amino acid positions for the ligand-receptor interaction, and perform semi-throughput screening of a peptide library. Because insect kinins are susceptible to fast enzymatic degradation by endogenous peptidases, they are severely limited in use as tools for pest control or endocrinological studies. Therefore, we also tested kinin analogs containing amino isobutyric acid (Aib) to enhance their potency and biostability. This peptidase-resistant analog represents an important lead in the development of biostable insect kinin analogs and may aid in the development of neuropeptide-based arthropod control strategies.

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito Aedes aegypti and Tick Rhipicephalus microplus G Protein-coupled Receptors

This protocol provides instructions for clonal-cell line selection and a calcium bioluminescence assay to analyze the structure-activity relationships of synthesized arthropod neuropeptides on their cognate GPCRs. This assay can be used for receptor deorphanization and structure-activity relationship studies for synthetic analog design and peptide/drug-lead discovery.

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes.  The ...

A Protocol for Collecting and Staining Hemocytes from the Yellow Fever Mosquito Aedes aegypti

Mosquitoes are vectors for a number of disease-causing pathogens such as the yellow fever virus, malaria parasites and filarial worms. Laboratories are investigating anti-pathogen components of the innate immune system in disease vector species in the hopes of generating transgenic mosquitoes that are refractory to such pathogens1, 2. The innate immune system of mosquitoes consists of several lines of defense 3. Pathogens that manage to escape the barrier imposed by the epithelium-lined mosquito midgut 4 enter the hemolymph and encounter circulating hemocytes, important cellular components that encapsulate and engulf pathogens 5, 6. Researchers have not found evidence for hematopoietic tissues in mosquitoes and current evidence suggests that the number of hemocytes is fixed at adult emergence and numbers may actually decline as the mosquito ages 7. The ability to properly collect and identify hemocytes from medically important insects is an essential step for studies in cellular immunity. However, the small size of mosquitoes and the limited volume of hemolymph pose a challenge to collecting immune cells. 
Two established methods for collecting mosquito hemocytes include expulsion of hemolymph from a cut proboscis 8, and volume displacement (perfusion), in which saline is injected into the membranous necklike region between the head and thorax (i.e., cervix) and the perfused hemolymph is collected from a torn opening in a distal region of the abdomen 9, 10. These techniques, however, are limited by low recovery of hemocytes and possible contamination by fat body cells, respectively 11. More recently a method referred to as high injection/recovery improved recovery of immunocytes by use of anticoagulant buffers while reducing levels of contaminating scales and internal tissues 11. While that method allows for an improved method of collecting and maintaining hemocytes for primary culture, it entails a number of injection and collecting steps that are not necessary if the downstream goal is to collect, fix and stain hemocytes for diagnostics. Here, we demonstrate our method of collecting mosquito hemolymph that combines the simplicity of perfusion, using anticoagulant buffers in place of saline solution, with the accuracy of high injection techniques to isolate clean preparations of hemocytes in Aedes mosquitoes.

A Protocol for Collecting and Staining Hemocytes from the Yellow Fever Mosquito Aedes aegypti

A simplified yet accurate method to collect and stain mosquito hemocytes is described. Our method combines the simplicity of perfusion with the accuracy of high injection techniques to isolate clean preparations of hemocytes in Aedes mosquitoes. This method facilitates studies requiring knowledge of the types of hemocytes and their abundance.

 Mosquitoes are vectors for a number  of disease-causing pathogens such as the yellow fever virus, malaria parasites  and filarial worms. Laboratories are ...

Detection of Infectious Virus from Field-collected Mosquitoes by Vero Cell Culture Assay

Mosquitoes transmit a number of distinct viruses including important human pathogens such as West Nile virus, dengue virus, and chickungunya virus. Many of these viruses have intensified in their endemic ranges and expanded to new territories, necessitating effective surveillance and control programs to respond to these threats. One strategy to monitor virus activity involves collecting large numbers of mosquitoes from endemic sites and testing them for viral infection. In this article, we describe how to handle, process, and screen field-collected mosquitoes for infectious virus by Vero cell culture assay. Mosquitoes are sorted by trap location and species, and grouped into pools containing &le;50 individuals. Pooled specimens are homogenized in buffered saline using a mixer-mill and the aqueous phase is inoculated onto confluent Vero cell cultures (Clone E6). Cell cultures are monitored for cytopathic effect from days 3-7 post-inoculation and any viruses grown in cell culture are identified by the appropriate diagnostic assays. By utilizing this approach, we have isolated 9 different viruses from mosquitoes collected in Connecticut, USA, and among these, 5 are known to cause human disease. Three of these viruses (West Nile virus, Potosi virus, and La Crosse virus) represent new records for North America or the New England region since 1999. The ability to detect a wide diversity of viruses is critical to monitoring both established and newly emerging viruses in the mosquito population.

We describe a method to process and screen field-collected mosquitoes for a diversity of viruses by Vero cell culture assay. By employing this technique, we have detected 9 different viruses from 4 taxonomic families in mosquitoes collected in Connecticut.

Mosquitoes transmit a number of distinct viruses including important human pathogens such as West Nile virus, dengue virus, and chickungunya virus. Many ...

Intravital Microscopy of the Spleen: Quantitative Analysis of Parasite Mobility and Blood Flow

The advent of intravital microscopy in experimental rodent malaria models has allowed major advances to the knowledge of parasite-host interactions 1,2. Thus, in vivo imaging of malaria parasites during pre-erythrocytic stages have revealed the active entrance of parasites into skin lymph nodes 3, the complete development of the parasite in the skin 4, and the formation of a hepatocyte-derived merosome to assure migration and release of merozoites into the blood stream 5. Moreover, the development of individual parasites in erythrocytes has been recently documented using 4D imaging and challenged our current view on protein export in malaria 6. Thus, intravital imaging has radically changed our view on key events in Plasmodium development. Unfortunately, studies of the dynamic passage of malaria parasites through the spleen, a major lymphoid organ exquisitely adapted to clear infected red blood cells are lacking due to technical constraints.
Using the murine model of malaria Plasmodium yoelii in Balb/c mice, we have implemented intravital imaging of the spleen and reported a differential remodeling of it and adherence of parasitized red blood cells (pRBCs) to barrier cells of fibroblastic origin in the red pulp during infection with the non-lethal parasite line P.yoelii 17X as opposed to infections with the P.yoelii 17XL lethal parasite line 7. To reach these conclusions, a specific methodology using ImageJ free software was developed to enable characterization of the fast three-dimensional movement of single-pRBCs. Results obtained with this protocol allow determining velocity, directionality and residence time of parasites in the spleen, all parameters addressing adherence in vivo. In addition, we report the methodology for blood flow quantification using intravital microscopy and the use of different colouring agents to gain insight into the complex microcirculatory structure of the spleen. 
Ethics statement
All the animal studies were performed at the animal facilities of University of Barcelona in accordance with guidelines and protocols approved by the Ethics Committee for Animal Experimentation of the University of Barcelona CEEA-UB (Protocol No DMAH: 5429). Female Balb/c mice of 6-8 weeks of age were obtained from Charles River Laboratories.

We show the method for performing intravital microscopy of the spleen using GFP transgenic malaria parasites and the quantification of parasite mobility and blood flow within this organ.

The advent of intravital microscopy in experimental rodent malaria models has allowed major advances to the knowledge of parasite-host interactions 1,2. ...

Parasite Induced Genetically Driven Autoimmune Chagas Heart Disease in the Chicken Model

The Trypanosoma cruzi acute infections acquired in infancy and childhood seem asymptomatic, but approximately one third of the chronically infected cases show Chagas disease up to three decades or later. Autoimmunity and parasite persistence are competing theories to explain the pathogenesis of Chagas disease 1, 2. To separate roles played by parasite persistence and autoimmunity in Chagas disease we inoculate the T. cruzi in the air chamber of fertilized eggs. The mature chicken immune system is a tight biological barrier against T. cruzi and the infection is eradicated upon development of its immune system by the end of the first week of growth 3. The chicks are parasite-free at hatching, but they retain integrated parasite mitochondrial kinetoplast DNA (kDNA) minicircle within their genome that are transferred to their progeny. Documentation of the kDNA minicircle integration in the chicken genome was obtained by a targeted prime TAIL-PCR, Southern hybridizations, cloning, and sequencing 3, 4. The kDNA minicircle integrations rupture open reading frames for transcription and immune system factors, phosphatase (GTPase), adenylate cyclase and phosphorylases (PKC, NF-Kappa B activator, PI-3K) associated with cell physiology, growth, and differentiation 3, 5-7, and other gene functions. Severe myocarditis due to rejection of target heart fibers by effectors cytotoxic lymphocytes is seen in the kDNA mutated chickens, showing an inflammatory cardiomyopathy similar to that seen in human Chagas disease. Notably, heart failure and skeletal muscle weakness are present in adult chickens with kDNA rupture of the dystrophin gene in chromosome 1 8. Similar genotipic alterations are associated with tissue destruction carried out by effectors CD45+, CD8&gamma;&delta;+, CD8&alpha; lymphocytes. Thus this protozoan infection can induce genetically driven autoimmune disease.

The inoculation of Trypanosoma cruzi in fertile eggs prior to incubation renders the parasite kDNA minicircle integration in embryo cells genome. Crossbreeding reveals the vertical transfer of the mutations to progeny. The kDNA integrates into coding regions at several chromosomes and the chickens die with an inflammatory autoimmune heart disease.

The Trypanosoma cruzi acute infections acquired in infancy and childhood seem asymptomatic, but approximately one third of the chronically infected cases ...

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. One important application of this method is the development of high-quality physical maps useful for improving the genome assemblies for various organisms. The natural banding pattern of polytene and mitotic chromosomes provides guidance for the precise ordering and orientation of the genomic supercontigs. Among the three mosquito genera, namely Anopheles, Aedes, and Culex, a well-established chromosome-based mapping technique has been developed only for Anopheles, whose members possess readable polytene chromosomes 1. As a result of genome mapping efforts, 88% of the An. gambiae genome has been placed to precise chromosome positions 2,3 . Two other mosquito genera, Aedes and Culex, have poorly polytenized chromosomes because of significant overrepresentation of transposable elements in their genomes 4, 5, 6. Only 31 and 9% of the genomic supercontings have been assigned without order or orientation to chromosomes of Ae. aegypti 7 and Cx. quinquefasciatus 8, respectively. Mitotic chromosome preparation for these two species had previously been limited to brain ganglia and cell lines. However, chromosome slides prepared from the brain ganglia of mosquitoes usually contain low numbers of metaphase plates 9. Also, although a FISH technique has been developed for mitotic chromosomes from a cell line of Ae. aegypti 10, the accumulation of multiple chromosomal rearrangements in cell line chromosomes 11 makes them useless for genome mapping. Here we describe a simple, robust technique for obtaining high-quality mitotic chromosome preparations from imaginal discs (IDs) of 4th instar larvae which can be used for all three genera of mosquitoes. A standard FISH protocol 12 is optimized for using BAC clones of genomic DNA as a probe on mitotic chromosomes of Ae. aegypti and Cx. quinquefasciatus, and for utilizing an intergenic spacer (IGS) region of ribosomal DNA (rDNA) as a probe on An. gambiae chromosomes. In addition to physical mapping, the developed technique can be applied to population cytogenetics and chromosome taxonomy/systematics of mosquitoes and other insect groups.

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Among the three mosquito genera, namely Anopheles, Aedes, and Culex, physical genome mapping techniques were established only for Anopheles, whose members possess readable polytene chromosomes. For the genera of Aedes and Culex, however, cytogenetic mapping remains challenging because of the poor quality of polytene chromosomes. Here we present a universal protocol for obtaining high-quality preparations of mitotic chromosomes and an optimized FISH protocol for all three genera of mosquitoes.

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. ...

2D and 3D Chromosome Painting in Malaria Mosquitoes

Fluorescent in situ hybridization (FISH) of whole arm chromosome probes is a robust technique for mapping genomic regions of interest, detecting chromosomal rearrangements, and studying three-dimensional (3D) organization of chromosomes in the cell nucleus. The advent of laser capture microdissection (LCM) and whole genome amplification (WGA) allows obtaining large quantities of DNA from single cells. The increased sensitivity of WGA kits prompted us to develop chromosome paints and to use them for exploring chromosome organization and evolution in non-model organisms. Here, we present a simple method for isolating and amplifying the euchromatic segments of single polytene chromosome arms from ovarian nurse cells of the African malaria mosquito Anopheles gambiae. This procedure provides an efficient platform for obtaining chromosome paints, while reducing the overall risk of introducing foreign DNA to the sample. The use of WGA allows for several rounds of re-amplification, resulting in high quantities of DNA that can be utilized for multiple experiments, including 2D and 3D FISH. We demonstrated that the developed chromosome paints can be successfully used to establish the correspondence between euchromatic portions of polytene and mitotic chromosome arms in An. gambiae. Overall, the union of LCM and single-chromosome WGA provides an efficient tool for creating significant amounts of target DNA for future cytogenetic and genomic studies.

Chromosome painting is a useful method for studying organization of the cell nucleus and evolution of the karyotype. Here, we demonstrate an approach to isolate and amplify specific regions of interest from single polytene chromosomes that are subsequently used for two- and three-dimensional fluorescent in situ hybridization (FISH).

Fluorescent in situ hybridization (FISH) of whole arm chromosome probes is a robust technique for mapping genomic regions of interest, detecting ...

Influenza Virus Propagation in Embryonated Chicken Eggs

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous emergence of new influenza virus strains due to mutation and re-assortment complicates the control of the virus and necessitates the permanent development of novel drugs and vaccines. The laboratory-based study of influenza requires a reliable and cost-effective method for the propagation of the virus. Here, a comprehensive protocol is provided for influenza A virus propagation in fertile chicken eggs, which consistently yields high titer viral stocks. In brief, serum pathogen-free (SPF) fertilized chicken eggs are incubated at 37 &#176;C and 55-60% humidity for 10 &#8211; 11 days. Over this period, embryo development can be easily monitored using an egg candler. Virus inoculation is carried out by injection of virus stock into the allantoic cavity using a needle. After 2 days of incubation at 37 &#176;C, the eggs are chilled for at least 4 hr at 4 &#176;C. The eggshell above the air sac and the chorioallantoic membrane are then carefully opened, and the allantoic fluid containing the virus is harvested. The fluid is cleared from debris by centrifugation, aliquoted and transferred to -80 &#176;C for long-term storage. The large amount (5-10 ml of virus-containing fluid per egg) and high virus titer which is usually achieved with this protocol has made the usage of eggs for virus preparation our favorable method, in particular for in vitro studies which require large quantities of virus in which high dosages of the same virus stock are needed.

Fertile chicken eggs are widely used to produce large amounts of human influenza A virus as they provide a convenient and cost-effective system to prove high yields of virus.

Influenza infection is associated with about 36,000 deaths and more than 200,000 hospitalizations every year in the United States. The continuous ...

Measurement of Pulse Propagation Velocity, Distensibility and Strain in an Abdominal Aortic Aneurysm Mouse Model

An abdominal aortic aneurysm (AAA) is defined as a localized dilation of the abdominal aorta that exceeds the maximal intraluminal diameter (MILD) by 1.5 times of its original size. Clinical and experimental studies have shown that small aneurysms may rupture, while a subpopulation of large aneurysms may remain stable. Thus, in addition to the measurement of intraluminal diameter of the aorta, knowledge of structural traits of the vessel wall may provide important information to assess the stability of the AAA. Aortic stiffening has recently emerged as a reliable tool to determine early changes in the vascular wall. Pulse propagation velocity (PPV) along with the distensibility and radial strain are highly useful ultrasound-based methods relevant for assessing aortic stiffness. The primary purpose of this protocol is to provide a comprehensive technique for the use of ultrasound imaging system to acquire images and analyze the structural and functional properties of the aorta as determined by MILD, PPV, distensibility and radial strain.

This manuscript describes a detailed protocol for using high frequency ultrasound imaging to measure luminal diameter, pulse propagation velocity, distensibility and radial strain on a mouse model of abdominal aortic aneurysm.

An abdominal aortic aneurysm (AAA) is defined as a localized dilation of the abdominal aorta that exceeds the maximal intraluminal diameter (MILD) by 1.5 ...

In Vivo Infection with Leishmania amazonensis to Evaluate Parasite Virulence in Mice

Leishmania spp. are protozoan parasites that cause leishmaniases, diseases that present a wide spectrum of clinical manifestations from cutaneous to visceral lesions. Currently, 12 million people are estimated to be infected with Leishmania worldwide and over 1 billion people live at the risk of infection. Leishmania amazonensis is endemic in Central and South America and usually leads to the cutaneous form of the disease, which can be directly visualized in an animal model. Therefore, L. amazonensis strains are good models for cutaneous leishmaniasis studies because they are also easily cultivated in vitro. C57BL/6 mice mimic the L. amazonensis-driven disease progression observed in humans and are considered one of the best mice strains model for cutaneous leishmaniasis. In the vertebrate host, these parasites inhabit macrophages despite the defense mechanisms of these cells. Several studies use in vitro macrophage infection assays to evaluate the parasite infectivity under different conditions. However, the in vitro approach is limited to an isolated cell system that disregards the organism&#39;s response. Here, we compile an in vivo murine infection method that provides a systemic physiological overview of the host-parasite interaction. The detailed protocol for the in vivo infection of C57BL/6 mice with L. amazonensis comprises parasite differentiation into infective amastigotes, mice footpad cutaneous inoculation, lesion development, and parasite load determination. We propose this well-established method as the most adequate method for physiological studies of the host immune and metabolic responses to cutaneous leishmaniasis.

In Vivo Infection with Leishmania amazonensis to Evaluate Parasite Virulence in Mice

Here, we present a compiled protocol to evaluate the cutaneous infection of mice with Leishmania amazonensis. This is a reliable method for studying parasite virulence, allowing a systemic view of the vertebrate host response to the infection.

Leishmania spp. are protozoan parasites that cause leishmaniases, diseases that present a wide spectrum of clinical manifestations from cutaneous to ...

Propagation of the Microsporidian Parasite Edhazardia aedis in Aedes aegypti Mosquitoes

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A Calcium Bioluminescence Assay for Functional Analysis of Mosquito (Aedes aegypti) and Tick (Rhipicephalus microplus) G Protein-coupled Receptors

A Protocol for Collecting and Staining Hemocytes from the Yellow Fever Mosquito Aedes aegypti

Detection of Infectious Virus from Field-collected Mosquitoes by Vero Cell Culture Assay

Intravital Microscopy of the Spleen: Quantitative Analysis of Parasite Mobility and Blood Flow

Parasite Induced Genetically Driven Autoimmune Chagas Heart Disease in the Chicken Model

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

2D and 3D Chromosome Painting in Malaria Mosquitoes

Influenza Virus Propagation in Embryonated Chicken Eggs

Measurement of Pulse Propagation Velocity, Distensibility and Strain in an Abdominal Aortic Aneurysm Mouse Model

In Vivo Infection with Leishmania amazonensis to Evaluate Parasite Virulence in Mice