We would like to thank the Johns Hopkins Malaria Research Institute for access to and rearing of An. gambiae larvae.

1. Preparation of solutions and tools
<ol>
	<li>Preparation of dissection solution
	<ol>
		<li>To prepare dissecting solution, add 2.5 mL of 100% ethanol to 7.5 mL of distilled H2O in a 15 plastic tube. Invert the tube 3 times to mix. 
		NOTE: This solution can be stored at room temperature for several weeks.</li>
	</ol>
	</li>
	<li>Preparation of 10x phosphate-buffered saline (PBS) stock
	<ol>
		<li>To prepare 10x PBS stock, add 17.8 g Na2HPO4&#8226; 2H2O, 2.4 g KH2PO4, 80 g NaCl, and 2 g KCl to 800 mL of deionized water. Mix with a stir bar on a stir plate until the solids have fully dissolved. Adjust the final volume to 1 L with purified water. 
		NOTE: This solution can be stored at room temperature for several months.</li>
	</ol>
	</li>
	<li>Preparation of 1x PBS
	<ol>
		<li>Add 10 mL of 10x PBS to 90 mL of H2O in a sterile glass bottle. Close the lid tightly and invert 3x to mix. 
		NOTE: The solution can be stored at 4 &#176;C for several weeks.</li>
	</ol>
	</li>
	<li>Preparation of fixative.
	<ol>
		<li>Add 600 &#181;L of methanol to 200 &#181;L of glacial acetic acid. Make the solution fresh each time.</li>
	</ol>
	</li>
	<li>Construction of putty plate (a tissue culture dish filled with silicone rubber)
	<ol>
		<li>Mix the components (1:50) of epoxy (1 g) and water (50 mL) and pour the mixture into a Petri dish. Wait for the components to dry before using. 
		NOTE: Once constructed, the putty plate can be used for years.</li>
	</ol>
	</li>
</ol>
2. Gland dissection (Figure 1A)
<ol>
	<li>Collect late-stage L4 larvae (~10 days post hatch; Figure 2) from feeding trays using a plastic transfer pipette. 
	NOTE: See this helpful online manual for standard mosquito husbandry and larval culture13.
	<ol>
		<li>Place a putty plate on the stage of a dissecting microscope and transfer larvae onto the putty plate. 
		NOTE: When aliquoting larvae for initial dissection, it is helpful to transfer ~10 larvae onto the slide at one time using a plastic transfer pipette.</li>
	</ol>
	</li>
	<li>Place a drop of dissection solution (1:3 EtOH:H2O) onto the putty plate, separate from the 10 larvae.</li>
	<li>Use a plastic transfer pipette or disposable glass pipette to place one L4 larva into the 25% EtOH drop.</li>
	<li>Take forceps (#5), one in each hand, and with the non-dominant hand, grip the head of the larva. Using the dominant hand, grasp the larva with forceps just below the head and gently pull with minimal constant force, such that the head detaches from the rest of the body and the glands remain attached to the head. Discard the body portion of the larva carcass. 
	NOTE: When dissecting, set a paper towel nearby to collect the larvae carcasses.</li>
	<li>Collect the head/glands into 1 mL of 1x PBS in a 1.5 mL microcentrifuge tube. Use 1 mL of PBS for ~40 dissected glands with their attached heads. Wait for the heads/SGs to sink to the bottom of the buffer.</li>
</ol>
3. Fixation for antibody staining (Figure 1B)
<ol>
	<li>Drain the PBS starting from the top of the microcentrifuge tube using a glass transfer pipette, avoiding the sticky mosquito tissues and removing as much of the PBS solution as possible without damaging the tissues. Replace the solution with 800 &#181;L of a 3:1 mixture of methanol to glacial acetic acid. Place the tube at 4 &#176;C overnight (12-24 h recommended; 19 h preferred).</li>
	<li>The following day, drain the solution and replace it with 1 mL of cold 100% acetone. Leave for 90 s.</li>
	<li>Remove the acetone and gently rinse the tissues three times with 1 mL of 1x PBS each time. 
	NOTE: The samples should float slowly up with the flow, then fall back to the bottom of the tube by the time each PBS addition is complete.</li>
</ol>
4. Immunostaining (Figure 1B)
<ol>
	<li>Add primary antibody (e.g., Rab11) at the appropriate dilution (1:100) in 200 &#181;L of the total volume of 1x PBS. Swirl the tube contents gently with a pipette tip. Incubate overnight at 4 &#176;C.</li>
	<li>Remove the primary antibody solution and wash three times with 1 mL of 1x PBS (gently pipetting the solution in and out of the pipette).</li>
	<li>Add secondary antibody (Alex Fluor 488 Goat anti-Rabbit) at the appropriate dilution (1:200) in 200 &#181;L of total volume. Swirl the tube contents gently with a pipette tip. Incubate at room temperature for 90 min.</li>
	<li>Add dyes, such as Nile Red (lipids, 5 &#181;L of 1 &#181;g/&#181;L) and/or Hoechst (DNA, 3 &#181;L of 1 &#181;g/&#181;L), into 200 &#181;L of PBS at this stage. Incubate at room temperature for an additional 60 min. Wash gently three times with 1x PBS.</li>
</ol>
5. Mounting stained glands for microscopy (Figure 1C)
<ol>
	<li>Pipette 200 &#181;L of 100% glycerol onto a microscope slide. 
	NOTE: As glycerol is viscous, leave the pipette tip in the liquid until it reaches the appropriate volume. No tissue shrinking was observed when going straight into 100% glycerol. However, researchers can go through 30% and 50% glycerol washes in the tubes before moving the samples into 100% glycerol.</li>
	<li>Transfer the stained heads (up to 20 per slide) with the attached glands (along with other internal structures) to the microscope slide using a soft brush (Figure 3A). Spread the samples out so that they are evenly distributed along the glass slide. 
	NOTE: When depositing glands on a cover slide with a brush, forceps can be used to gently orient the SGs so that they are all oriented in the same direction.</li>
	<li>Viewing under a dissecting microscope, separate the larval head from the glands using two pairs of forceps and carefully pulling the two tissues in opposite directions. 
	NOTE: As it is challenging to completely isolate the SGs, simply remove the heads, leaving the SGs and associated internal structures. Even if the SGs remain associated with other tissues, they are easily recognized when stained with Hoechst and other markers (Figure 3B,C).</li>
	<li>Remove and discard the heads and repeat the separation process for each SG. 
	NOTE: When removing heads, set a paper towel nearby to collect them.</li>
	<li>Gently place a 1.5 mm thick coverslip on the top (avoiding air bubbles) and seal with clear nail polish.</li>
	<li>Store at 4 &#176;C in a light-proof container. 
	NOTE: Prepared slides of stained glands can be kept at 4 &#176;C in the dark for up to six months to one year without any notable changes in quality. If glycerol starts to leak as the months go by, gently lift the coverslip and pipette in more glycerol to fill holes.</li>
</ol>

<ol><li>World malaria report 2020: 20 years of global progress and challenges. World Health Organization. , Available from: 
 <a target="_blank" href="https://apps.who.int/iris/handle/10665/337660">https://apps.who.int/iris/handle/10665/337660</a> (2020).</li>
<li>Feachem, R. G. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Malaria+eradication+within+a+generation%3A+ambitious%2C+achievable%2C+and+necessary%5BTitle%5D)+AND+%22Lancet%22%5BJournal%5D)">Malaria eradication within a generation: ambitious, achievable, and necessary.</a> Lancet. 394 (10203), 1056-1112 (2019).</li>
<li>Adolfi, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Efficient+population+modification+gene-drive+rescue+system+in+the+malaria+mosquito+Anopheles+stephensi%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Efficient population modification gene-drive rescue system in the malaria mosquito Anopheles stephensi.</a> Nature Communications. 11, 5553(2020).</li>
<li>Kyrou, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+CRISPR-Cas9+gene+drive+targeting+doublesex+causes+complete+population+suppression+in+caged+Anopheles+gambiae+mosquitoes%5BTitle%5D)+AND+%22Nature+Biotechnology%22%5BJournal%5D)">A CRISPR-Cas9 gene drive targeting doublesex causes complete population suppression in caged Anopheles gambiae mosquitoes.</a> Nature Biotechnology. 36 (11), 1062-1066 (2018).</li>
<li>Rostami, M. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((CRISPR%2FCas9+gene+drive+technology+to+control+transmission+of+vector-borne+parasitic+infections%5BTitle%5D)+AND+%22Parasite+Immunology%22%5BJournal%5D)">CRISPR/Cas9 gene drive technology to control transmission of vector-borne parasitic infections.</a> Parasite Immunology. 42 (9), 12762(2020).</li>
<li>Bhatt, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Coverage+and+system+efficiencies+of+insecticide-treated+nets+in+Africa+from+2000+to+2017%5BTitle%5D)+AND+%22Elife%22%5BJournal%5D)">Coverage and system efficiencies of insecticide-treated nets in Africa from 2000 to 2017.</a> Elife. 4, 09672(2015).</li>
<li>Mellink, J. J., Vanden Bovenkamp, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functional+aspects+of+mosquito+salivation+in+blood+feeding+of+Aedes+aegypti%5BTitle%5D)+AND+%22Mosquito+News%22%5BJournal%5D)">Functional aspects of mosquito salivation in blood feeding of Aedes aegypti.</a> Mosquito News. 41 (1), 115(1981).</li>
<li>Ribeiro, J. M., Rossignol, P. A., Spielman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Role+of+mosquito+saliva+in+blood+vessel+location%5BTitle%5D)+AND+%22Journal+of+Experimental+Biology%22%5BJournal%5D)">Role of mosquito saliva in blood vessel location.</a> Journal of Experimental Biology. 108, 1-7 (1984).</li>
<li>Yamamoto, D. S., Sumitani, M., Kasashima, K., Sezutsu, H., Matsuoka, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Inhibition+of+malaria+infection+in+transgenic+Anopheline+mosquitoes+lacking+salivary+gland+cells%5BTitle%5D)+AND+%22PLoS+Pathogens%22%5BJournal%5D)">Inhibition of malaria infection in transgenic Anopheline mosquitoes lacking salivary gland cells.</a> PLoS Pathogens. 12 (9), 1005872(2016).</li>
<li>Rishikesh, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Morphology+and+development+of+the+salivary+glands+and+their+chromosomes+in+the+larvae+of+Anopheles+stephensi+sensu+stricto%5BTitle%5D)+AND+%22Bulletin+of+the+World+Health+Organization%22%5BJournal%5D)">Morphology and development of the salivary glands and their chromosomes in the larvae of Anopheles stephensi sensu stricto.</a> Bulletin of the World Health Organization. 20 (1), 47-61 (1959).</li>
<li>Jensen, D. V., Jones, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+development+of+the+salivary+glands+in+Anopheles+albimanus+Wiedemann+%28Diptera%2C+Culicidae%29%5BTitle%5D)+AND+%22Annals+of+the+Entomological+Society+of+America%22%5BJournal%5D)">The development of the salivary glands in Anopheles albimanus Wiedemann (Diptera, Culicidae).</a> Annals of the Entomological Society of America. 50 (5), 40824(1957).</li>
<li>Chiu, M., Trigg, B., Taracena, M., Wells, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Diverse+cellular+morphologies+during+lumen+maturation+in+Anopheles+gambiae+larval+salivary+glands%5BTitle%5D)+AND+%22Insect+Molecular+Biology%22%5BJournal%5D)">Diverse cellular morphologies during lumen maturation in Anopheles gambiae larval salivary glands.</a> Insect Molecular Biology. 30 (2), 210-230 (2021).</li>
<li>Benedict, M. Q. MR4. Methods in Anopheles research. Center for Disease Control. , Available from: 
 <a target="_blank" href="https://www.beiresources.org/Portals/2/ VectorResources/2016%20Methods%20in%20Anopheles%20Research%20full%20manual.pdf">https://www.beiresources.org/Portals/2/ VectorResources/2016%20Methods%20in%20Anopheles%20Research%20full%20manual.pdf</a> (2015).</li>
<li>Wells, M. B., Andrew, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Salivary+gland+cellular+architecture+in+the+Asian+malaria+vector+mosquito+Anopheles+stephensi%5BTitle%5D)+AND+%22Parasites+%26+Vectors%22%5BJournal%5D)">Salivary gland cellular architecture in the Asian malaria vector mosquito Anopheles stephensi.</a> Parasites & Vectors. 8, 617(2015).</li>
<li>Wells, M. B., Villamor, J., Andrew, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Salivary+gland+maturation+and+duct+formation+in+the+African+malaria+mosquito+Anopheles+gambiae%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Salivary gland maturation and duct formation in the African malaria mosquito Anopheles gambiae.</a> Scientific Reports. 7 (1), 601(2017).</li>
<li>Wells, M. B., Andrew, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Anopheles+salivary+gland+architecture+shapes+plasmodium+sporozoite+availability+for+transmission%5BTitle%5D)+AND+%22mBio%22%5BJournal%5D)">Anopheles salivary gland architecture shapes plasmodium sporozoite availability for transmission.</a> mBio. 10 (4), 01238(2019).</li>
<li>Clements, A. N. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aClements%2C+AN+%22The+physiology+of+mosquitoes%22">The physiology of mosquitoes</a>. , Pergamon Press. Paris. (1963).</li>
<li>Imms, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((On+the+larval+and+pupal+stages+of+Anopheles+maculipennis%2C+meigen%5BTitle%5D)+AND+%22Parasitology%22%5BJournal%5D)">On the larval and pupal stages of Anopheles maculipennis, meigen.</a> Parasitology. 1 (2), 103-133 (1908).</li>
<li>Favia, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Bacteria+of+the+genus+Asaia+stably+associate+with+Anopheles+stephensi%2C+an+Asian+malarial+mosquito+vector%5BTitle%5D)+AND+%22Proceedings+of+the+National+Academy+of+Sciences+of+the+United+States+of+America%22%5BJournal%5D)">Bacteria of the genus Asaia stably associate with Anopheles stephensi, an Asian malarial mosquito vector.</a> Proceedings of the National Academy of Sciences of the United States of America. 104 (21), 9047-9051 (2007).</li>
<li>Neira Oviedo, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+salivary+transcriptome+of+Anopheles+gambiae+%28Diptera%3A+Culicidae%29+larvae%3A+A+microarray-based+analysis%5BTitle%5D)+AND+%22Insect+Biochemistry+and+Molecular+Biology%22%5BJournal%5D)">The salivary transcriptome of Anopheles gambiae (Diptera: Culicidae) larvae: A microarray-based analysis.</a> Insect Biochemistry and Molecular Biology. 39 (5-6), 382-394 (2009).</li>
<li>Linser, P. J., Smith, K. E., Seron, T. J., Neira Oviedo, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Carbonic+anhydrases+and+anion+transport+in+mosquito+midgut+pH+regulation%5BTitle%5D)+AND+%22Journal+of+Experimental+Biology%22%5BJournal%5D)">Carbonic anhydrases and anion transport in mosquito midgut pH regulation.</a> Journal of Experimental Biology. 212 (11), 1662-1671 (2009).</li>
<li>Linser, P. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Slc4-like+anion+transporters+of+the+larval+mosquito+alimentary+canal%5BTitle%5D)+AND+%22Journal+of+Insect+Physiology%22%5BJournal%5D)">Slc4-like anion transporters of the larval mosquito alimentary canal.</a> Journal of Insect Physiology. 58 (4), 551-562 (2012).</li>
</ol>

The authors have no conflicts of interest to disclose.

The protocol described herein was adapted from a Drosophila SG dissection protocol and an adult mosquito dissection protocol14,15,16. However, most markers did not penetrate the basement membrane (data not shown) when using the adult dissection and SG staining methods. Adaptations of the adult protocol included dissecting the glands in a 25% EtOH solution, washing the glands with a combination of MeOH and glacial acetic acid, an...

Malaria is a major public health threat causing almost 230 million infections and an estimated 409,000 deaths in 20191. The majority of deaths are in sub-Saharan Africa and are caused by the parasite Plasmodium falciparum, whose insect vector is Anopheles gambiae, the subject of this video demonstration. Although the numbers indicate a significant drop in annual death rate since the turn of the century (&#62;300,000 fewer annual deaths), the promising decreases in disease rates observed from 2000 to 2015 are tapering, suggesting the need for new approaches to limiting disease transmission2. Among promising additional strategies for controlling and possibly eliminating malaria is targeting mosquito vector capacity using CRISPR/Cas9-based gene-editing and gene-drive3,4,5. Indeed, it is the targeting of the mosquito vector (through the expanded use of long-lasting insecticide-treated bed nets) that has had the greatest impact on reducing disease transmission6.
Female mosquitoes acquire Plasmodium gametocytes from an infected human during a blood meal. Following fertilization, maturation, midgut epithelium traversal, population expansion, and hemocoel navigation in their obligate mosquito hosts, hundreds to tens of thousands of Plasmodium sporozoites invade the mosquito SGs and fill the secretory cavities of the constituent secretory cells. Once inside the secretory cavities, the parasites have direct access to the salivary duct and are thus poised for transmission to a new vertebrate host upon the next blood meal. Because SGs are critical for the transmission of malaria-causing sporozoites to their human hosts, and laboratory studies suggest that SGs are not essential for blood-feeding, mosquito survival, or fecundity7,8,9, they represent an ideal target for transmission-blocking measures. Adult mosquito SGs form as an elaboration of &#34;duct bud&#34; remnants in the larval SGs that persist beyond early pupal SG histolysis10, making the larval SG an ideal target for interventions to limit adult-stage disease transmission.
Characterizing the larval stage of SG development can help develop not only a better understanding of its morphology and functional adaptations but can also aid in assessing new interventions that target this organ through gene editing of key SG regulators. Because all previous studies of larval salivary gland architecture predate immunostaining and modern imaging techniques10,11, we have developed a protocol for isolating and staining salivary glands with a variety of antibodies and cell markers12. This video demonstrates this approach to the extraction, fixation, and staining of larval SGs from Anopheles gambiae L4 larvae for confocal imaging.

<table><tbody><tr><td>&amp;nbsp;KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Millipore Sigma</td><td>P9791</td><td></td></tr><tr><td>&amp;nbsp;Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; &amp;bull; 2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Millipore Sigma</td><td>71643</td><td></td></tr><tr><td>&amp;nbsp;NaCl</td><td>Millipore Sigma</td><td>S7653</td><td></td></tr><tr><td>Acetone</td><td>Millipore Sigma</td><td>179124</td><td></td></tr><tr><td>Brush with soft bristles</td><td>Amazon (SN NJDF) Detail Paint Brush Set</td><td>B08LH63D89</td><td></td></tr><tr><td>Cover slips (22 x 50 mm)</td><td>VWR</td><td>48393-195</td><td></td></tr><tr><td>DAPI (DNA)</td><td>ThermoFisher Scientific</td><td>D1306</td><td></td></tr><tr><td>Ethyl alcohol 200 proof</td><td>Millipore Sigma</td><td>EX0276</td><td></td></tr><tr><td>Gilson Pipetman P200 Pipette</td><td>Gilson</td><td>P200</td><td></td></tr><tr><td>Glacial Acetic Acid</td><td>Sigma Aldrich</td><td>695092</td><td></td></tr><tr><td>Jewelers forceps, Dumont No. 5</td><td>Millipore Sigma</td><td>F6521</td><td></td></tr><tr><td>KCl</td><td>Millipore Sigma</td><td>58221</td><td></td></tr><tr><td>Methanol</td><td>Millipore Sigma</td><td>1414209</td><td></td></tr><tr><td>Nail polish</td><td>Amazon (Sally Hansen)</td><td>B08148YH9M</td><td></td></tr><tr><td>Nile Red (lipid)</td><td>ThermoFisher Scientific</td><td>N1142</td><td></td></tr><tr><td>Paper towels/wipes</td><td>ULINE</td><td>S-7128</td><td></td></tr><tr><td>Petri plate (to make putty plate)</td><td>ThermoFisher Scientific</td><td>FB0875712</td><td></td></tr><tr><td>Pipette Tips</td><td>Gilson</td><td>Tips E200ST</td><td></td></tr><tr><td>Plastic Transfer Pipette</td><td>Fisher Scientific</td><td>S304671</td><td></td></tr><tr><td>Primary antibodies (e.g., Crb, Rab11)</td><td>Developmental Studies Hybridoma Bank (DSHB); Andrew Lab</td><td>Mouse anti-Crb (Cq4) or Rabbit anti-Rab11</td><td></td></tr><tr><td>Secondary antibodies with fluorescent tags (e.g., Alexa Fluor 488 Goat-anti Rabbit)</td><td>ThermoFisher Scientific</td><td>A11008</td><td></td></tr><tr><td>Silicone resin and curing agent for putty plate</td><td>Dow Chemicals - Ximeter Silicone</td><td>PMX-200</td><td></td></tr><tr><td>Slides, frosted on one end for labelling</td><td>VWR&amp;nbsp; 20 X 50 mm</td><td>48393-195</td><td></td></tr><tr><td>Wheat Germ Agglutinin</td><td>ThermoFisher Scientific</td><td>W834</td><td></td></tr></tbody></table>

dissection and immunostaining of larval salivary glands from anopheles gambiae mosquitoes

Mosquito salivary glands (SGs) are a requisite gateway organ for the transmission of insect-borne pathogens. Disease-causing agents, including viruses and the Plasmodium parasites that cause malaria, accumulate in the secretory cavities of SG cells. Here, they are poised for transmission to their vertebrate hosts during a subsequent blood meal. As adult glands form as an elaboration of larval SG duct bud remnants that persist beyond early pupal SG histolysis, the larval SG is an ideal target for interventions that limit disease transmission. Understanding larval SG development can help develop a better understanding of its morphology and functional adaptations and aid in the assessment of new interventions that target this organ. This video protocol demonstrates an efficient technique for isolating, fixing, and staining larval SGs from Anopheles gambiae mosquitoes. Glands dissected from larvae in a 25% ethanol solution are fixed in a methanol-glacial acetic acid mixture, followed by a cold acetone wash. After a few rinses in phosphate-buffered saline (PBS), SGs can be stained with a broad array of marker dyes and/or antisera against SG-expressed proteins. This method for larval SG isolation could also be used to collect tissue for in situ hybridization analysis, other transcriptomic applications, and proteomic studies.

Salivary glands are relatively easy to dissect from all stage 4 larvae. Male and female larvae can be distinguished at the late L4 larval stage by a red stripe along the dorsal thorax of females but not males (Figure 2). We also observe that antennal morphology is much more elaborate in male than in female L4 larvae (Figure 2), similar to the differences observed in this structure in adult mosquitoes. Along with the considerable overall growth during the L4 stag...

Watch this Scientific Journal Video about Dissection and Immunostaining of Larval Salivary Glands from Anopheles gambiae Mosquitoes at JoVE.com

Dissection and Immunostaining of Larval Salivary Glands from Anopheles gambiae Mosquitoes

The adult mosquito salivary gland (SG) is required for the transmission of all mosquito-borne pathogens to their human hosts, including viruses and parasites. This video demonstrates efficient isolation of the SGs from the larval (L4) stage Anopheles gambiae mosquitoes and preparation of the L4 SGs for further analysis.

Dissection and Immunostaining of Larval Salivary Glands from Anopheles gambiae Mosquitoes

Mosquito salivary glands (SGs) are a requisite gateway organ for the transmission of insect-borne pathogens. Disease-causing agents, including viruses and ...

dissection-immunostaining-larval-salivary-glands-from-anopheles

Research

JoVE Journal

Biology

4.3K Views.  The Johns Hopkins University School of Medicine. The adult mosquito salivary gland (SG) is required for the transmission of all mosquito-borne pathogens to their human hosts, including viruses and parasites. This video demonstrates efficient isolation of the SGs from the larval (L4) stage Anopheles gambiae mosquitoes and preparation of the L4 SGs for further analysis.

Video: Dissection and Immunostaining of Larval Salivary Glands from Anopheles gambiae Mosquitoes

Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes

The mosquito midgut and salivary glands are key entry and exit points for pathogens such as Plasmodium parasites and Dengue viruses. This video protocol demonstrates dissection techniques for removal of the midgut and salivary glands from Aedes aegypti mosquitoes. 



The mosquito midgut and salivary glands are key entry and exit points for vector pathogens like Plasmodium falciparum and the dengue virus. This video demonstrates the dissection techniques for removing the midgut and salivary glands from Aedes aegypti mosquitoes.

The mosquito midgut and salivary glands are key entry and exit points for pathogens such as Plasmodium parasites and Dengue viruses. This video protocol ...

Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

Mosquitoes are vectors for a diverse set of pathogens including arboviruses, protozoan parasites and nematodes. Investigation of transcripts and gene regulators that are expressed in tissues in which the mosquito host and pathogen interact, and in organs involved in reproduction are of great interest for strategies to reduce mosquito-borne disease transmission and disrupt egg development. A number of tools have been employed to study and validate the temporal and tissue-specific regulation of gene expression. Here, we describe protocols that have been developed to obtain spatial information, which enhances our understanding of where specific genes are expressed and their products accumulate. The protocol described has been used to validate expression and determine accumulation patterns of transcripts in tissues related to mosquito-borne pathogen transmission, such as female salivary glands, as well as subcellular compartments of ovaries and embryos, which relate to mosquito reproduction and development. 
 The following procedures represent an optimized methodology that improves the efficiency of various steps in the protocol without loss of target-specific hybridization signals. Guidelines for RNA probe preparation, dissection of soft tissues and the general procedure for fixation and hybridization are described in Part A, while steps specific for the collection, fixation, pre-hybridization and hybridization of mosquito embryos are detailed in Part B.

Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

Temporal and spatial gene expression analyses have a crucial role in functional genomics. Whole-mount hybridization in situ is useful for determining the localization of transcripts within tissues and subcellular compartments. Here we outline a hybridization in situ protocol with modifications for specific target tissues in mosquitoes.

Mosquitoes are vectors for a diverse set of pathogens including arboviruses, protozoan parasites and nematodes.  Investigation of transcripts and gene ...

Immunology and Infection

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. ...

Using the GAL4-UAS System for Functional Genetics in Anopheles gambiae

The bipartite GAL4-UAS system is a versatile and powerful tool for functional genetic analysis. The essence of the system is to cross transgenic 'driver' lines that express the yeast transcription factor GAL4 in a tissue specific manner, with transgenic 'responder' lines carrying a candidate gene/RNA interference construct whose expression is controlled by Upstream Activation Sequences (UAS) that bind GAL4. In the ensuing progeny, the gene or silencing construct is thus expressed in a prescribed spatiotemporal manner, enabling the resultant phenotypes to be assayed and gene function inferred. The binary system enables flexibility in experimental approaches to screen phenotypes generated by transgene expression in multiple tissue-specific patterns, even if severe fitness costs are induced. We have adapted this system for Anopheles gambiae, the principal malaria vector in Africa.
In this article, we provide some of the common procedures used during GAL4-UAS analysis. We describe the An. gambiae GAL4-UAS lines already generated, as well as the cloning of new responder constructs for upregulation and RNAi knockdown. We specify a step by step guide for sexing of mosquito pupae to establish genetic crosses, that also includes screening progeny to follow inheritance of fluorescent gene markers that tag the driver and responder insertions. We also present a protocol for clearing An. gambiae embryos to study embryonic development. Finally, we introduce potential adaptions of the method to generate driver lines through CRISPR/Cas9 insertion of GAL4 downstream of target genes.

Using the GAL4-UAS System for Functional Genetics in Anopheles gambiae

The bipartite GAL4-UAS system is a versatile tool for modification of gene expression in a controlled spatiotemporal manner which permits functional genetic analysis in Anopheles gambiae. The procedures described for using this system are a semi-standardized cloning strategy, sexing and screening of pupae for fluorescent protein markers and embryo fixation.

The bipartite GAL4-UAS system is a versatile and powerful tool for functional genetic analysis. The essence of the system is to cross transgenic 'driver' ...

Genetics

Phenotypic Analysis of Rodent Malaria Parasite Asexual and Sexual Blood Stages and Mosquito Stages

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. However, effective malaria parasite targets for vaccine and chemotherapy development are still limited. This is largely due to the unavailability of relevant and practical in vivo infection models for human Plasmodium species, most notably for P.&#160;falciparum and P. vivax. Therefore, rodent malaria species have been extensively used as practical alternative in vivo models for malaria vaccine, drug targeting, immune response, and functional characterization studies of conserved Plasmodiumspp. genes. Indeed, rodent malaria models have proven to be invaluable, especially for exploring mosquito transmission and liver stage biology, and were indispensable for immunological studies. However, there are discrepancies in the methods used to evaluate the phenotypes of transgenic and wild-type asexual and sexual blood-stage parasites.&#160;Examples of these discrepancies are the choice of an intravenous vs. intraperitoneal infection of rodents with blood-stage parasites and the evaluation of male gamete exflagellation. Herein, we detail standardized experimental methods to evaluate the phenotypes of asexual and sexual blood stages in transgenic parasites expressing reporter-gene or wild-type rodent malaria parasite species. We also detail the methods to evaluate the phenotypes of malaria parasite mosquito stages (gametes, ookinetes, oocysts, and sporozoites) inside Anopheles mosquito vectors. These methods are detailed and simplified here for the lethal and non-lethal strains of P. berghei and P. yoelii but can also be applied with some adjustments to P. chabaudi and P. vinckei rodent malaria species.

Due to the striking similarities of the life cycle and biology of rodent malaria parasites to human malaria parasites, rodent malaria models have become indispensable for malaria research. Herein, we standardized some of the most important techniques used in the phenotypic analysis of wild-type and transgenic rodent malaria species.

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. ...

Fluorescence In Situ Hybridization to Study Mosquito Spermatogenesis

Whole-Mount Fluorescence In Situ Hybridization to Study Spermatogenesis in the Anopheles Mosquito

Spermatogenesis is a complex biological process during which diploid cells undergo successive mitotic and meiotic division followed by large structural changes to form haploid spermatozoa. Besides the biological aspect, studying spermatogenesis is of paramount importance for understanding and developing genetic technologies such as gene drive and synthetic sex ratio distorters, which, by altering Mendelian inheritance and the sperm sex ratio, respectively, could be used to control pest insect populations. These technologies have proven to be very promising in lab settings and could potentially be used to control wild populations of Anopheles mosquitoes, which are vectors of malaria. Due to the simplicity of the testis anatomy and their medical importance, Anopheles gambiae, a major malaria vector in sub-Saharan Africa, represents a good cytological model for studying spermatogenesis. This protocol describes how whole-mount fluorescence in situ hybridization (WFISH) can be used to study the dramatic changes in cell nuclear structure through spermatogenesis using fluorescent probes that specifically stain the X and Y chromosomes. FISH usually requires the disruption of the reproductive organs to expose mitotic or meiotic chromosomes and allow the staining of specific genomic regions with fluorescent probes. WFISH enables the preservation of the native cytological structure of the testis, coupled with a good level of signal detection from fluorescent probes targeting repetitive DNA sequences. This allows researchers to follow changes in the chromosomal behavior of cells undergoing meiosis along the structure of the organ, where each phase of the process can clearly be distinguished. This technique could be particularly useful for studying chromosome meiotic pairing and investigating the cytological phenotypes associated with, for example, synthetic sex ratio distorters, hybrid male sterility, and the knock-out of genes involved in spermatogenesis.

Author Spotlight: Whole-Mount Fluorescence In Situ Hybridization to Study Spermatogenesis in the Anopheles Mosquito  

Given their simple anatomy, Anopheles testes offer a good cytological model for studying spermatogenesis. This protocol describes whole-mount fluorescence in situ hybridization, a technique used to investigate this biological process, as well as the phenotype of transgenic strains harboring mutations in the genes involved in sperm production.

Spermatogenesis is a complex biological process during which diploid cells undergo successive mitotic and meiotic division followed by large structural ...

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Malaria remains one of the most important public health problems, causing significant morbidity and mortality. Malaria is a mosquito borne disease transmitted through an infectious bite from the female Anopheles mosquito. Malaria control will eventually rely on a multitude of approaches, which includes ways to block transmission to, through and from mosquitoes. To study mosquito stages of malaria parasites in the laboratory, we have optimized a protocol to culture highly infectious Plasmodium falciparum gametocytes, a parasite stage required for transmission from the human host to the mosquito vector. P. falciparum gametocytes mature through five morphologically distinct steps, which takes approximately 1-2 weeks. Gametocyte culture described in this protocol is completed in 15 days and are infectious to mosquitoes from days 15-18. These protocols were developed to maintain a continuous cycle of infection competent gametocytes and to maintain uninterrupted supply of mosquito stages of the parasite. Here, we describe the methodology of gametocyte culture and how to infect mosquitoes with these parasites using glass membrane feeders.

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Detailed investigations on mosquito stages of malaria parasites are critical to design effective transmission blocking strategies. This protocol demonstrates how to effectively culture infectious gametocytes and then feed these gametocytes to mosquitoes to generate mosquito stages of P. falciparum.

Malaria remains one of the most important public health problems, causing significant morbidity and mortality. Malaria is a mosquito borne disease ...

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) administration has been accomplished primarily via injection, which requires significant time and is not suitable for field applications. To overcome these limitations, here we present a more efficient method for robust activation of RNAi by oral delivery of dsRNA to adult Anopheles gambiae. Long dsRNAs were produced in Escherichia coli strain HT115 (DE3), and a concentrated suspension of heat-killed dsRNA-containing bacteria in 10% sucrose was offered on cotton balls ad-libitum to adult mosquitoes. Cotton balls were replaced every 2 days for the duration of the treatment. Use of this method to target doublesex (a gene involved in sex differentiation) or fork head (which encodes a salivary gland transcription factor) resulted in reduced target gene expression and/or protein immunofluorescence signal, as measured by quantitative Real-Time PCR (qRT-PCR) or fluorescence confocal microscopy, respectively. Defects in salivary gland morphology were also observed. This highly flexible, user-friendly, low-cost, time-efficient method of dsRNA delivery could be broadly applicable to target genes important for insect vector physiology and beyond.

Effective Oral RNA Interference RNAi Administration to Adult Anopheles gambiae Mosquitoes

The oral administration of dsRNA produced by bacteria, a delivery method for RNA interference (RNAi) that is routinely used in Caenorhabditis elegans, was successfully applied here to adult mosquitoes. Our method allows for robust reverse genetics studies and transmission-blocking vector studies without the use of injection.

RNA interference has been a heavily utilized tool for reverse genetic analysis for two decades. In adult mosquitoes, double-stranded RNA (dsRNA) ...

High-Precision Mosquito Organ Isolation for Microbiome Analyses

Dissection of Mosquito Ovaries, Midgut, and Salivary Glands for Microbiome Analyses at the Organ Level

The global burden of mosquito-transmitted diseases, including malaria, dengue, West Nile, Zika, Usutu, and yellow fever, continues to increase, posing a significant public health threat. With the rise of insecticide resistance and the absence of effective vaccines, new strategies are emerging that focus on the mosquito's microbiota. Nevertheless, the majority of symbionts remain resistant to cultivation. Characterizing the diversity and function of bacterial genomes in mosquito specimens, therefore, relies on metagenomics and subsequent assembly and binning strategies. The obtention and analysis of Metagenome-Assembled Genomes (MAGs) from separated organs can notably provide key information about the specific role of mosquito-associated microbes in the ovaries (the reproductive organs), the midgut (key for food digestion and immunity), or the salivary glands (essential for the transmission of vector-borne diseases as pathogens must colonize them to enter the saliva and reach the bloodstream during a blood meal). These newly reconstructed genomes can then pave the way for the development of novel vector biocontrol strategies. To this aim, it is required to isolate mosquito organs while avoiding cross-contamination between them or with microorganisms present in other mosquito organs. Here, we describe an optimized and contamination-free dissection protocol for studying mosquito microbiome at the organ level.

Author Spotlight: Optimizing Mosquito Organ Dissection for Studying Symbionts and Vector Microbiomes

Microbial communities in mosquitoes hold great promise for vector biocontrol strategies. Most symbionts are uncultivable, requiring metagenomic analyses. We describe a method to dissect female mosquitoes and separate ovaries, midgut, and salivary glands preventing cross-contamination, facilitating microbiome studies at the organ level, and enhancing understanding of microorganisms' roles in mosquito biology.

The global burden of mosquito-transmitted diseases, including malaria, dengue, West Nile, Zika, Usutu, and yellow fever, continues to increase, posing a ...

Larval Ring Gland Dissection: Isolation of Developmental Endocrine Organs from Drosophila

Source: Imura, E., et al. <a href="https://www.jove.com/video/55519/" target="_blank">Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila melanogaster</a>. J. Vis. Exp. (2017).
This video describes the Drosophila ring gland, a complex endocrine structure important for biosynthesis and secretion of ecdysteroids and other hormones. The featured protocol demonstrates its dissection, fixation, and immunostaining.

Source: Imura, E., et al. Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila ...

Dissection and Immunostaining of Larval Salivary Glands from Anopheles gambiae Mosquitoes

Summary

Explore More Videos

Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes

Hybridization in situ of Salivary Glands, Ovaries, and Embryos of Vector Mosquitoes

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Using the GAL4-UAS System for Functional Genetics in Anopheles gambiae

Phenotypic Analysis of Rodent Malaria Parasite Asexual and Sexual Blood Stages and Mosquito Stages

Whole-Mount Fluorescence In Situ Hybridization to Study Spermatogenesis in the Anopheles Mosquito

Plasmodium falciparum Gametocyte Culture and Mosquito Infection Through Artificial Membrane Feeding

Effective Oral RNA Interference (RNAi) Administration to Adult Anopheles gambiae Mosquitoes

Dissection of Mosquito Ovaries, Midgut, and Salivary Glands for Microbiome Analyses at the Organ Level

Larval Ring Gland Dissection: Isolation of Developmental Endocrine Organs from Drosophila