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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Malaria+eradication+within+a+generation:+ambitious,+achievable,+and+necessary.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+population+modification+gene-drive+rescue+system+in+the+malaria+mosquito+Anopheles+stephensi.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+CRISPR-Cas9+gene+drive+targeting+doublesex+causes+complete+population+suppression+in+caged+Anopheles+gambiae+mosquitoes.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9+gene+drive+technology+to+control+transmission+of+vector-borne+parasitic+infections.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coverage+and+system+efficiencies+of+insecticide-treated+nets+in+Africa+from+2000+to+2017.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+aspects+of+mosquito+salivation+in+blood+feeding+of+Aedes+aegypti.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+mosquito+saliva+in+blood+vessel+location.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibition+of+malaria+infection+in+transgenic+Anopheline+mosquitoes+lacking+salivary+gland+cells.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphology+and+development+of+the+salivary+glands+and+their+chromosomes+in+the+larvae+of+Anopheles+stephensi+sensu+stricto.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+of+the+salivary+glands+in+Anopheles+albimanus+Wiedemann+(Diptera,+Culicidae).">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diverse+cellular+morphologies+during+lumen+maturation+in+Anopheles+gambiae+larval+salivary+glands.">Diverse cellular morphologies during lumen maturation in Anopheles gambiae larval salivary glands.</a> Insect Molecular Biology. 30 (2), 210-230 (2021).</li><li>MR4. Methods in Anopheles research. Center for Disease Control Available from: <a target="_blank" href="https://www.beiresources.org/Portals/2/">https://www.beiresources.org/Portals/2/</a> (2015)</li><li>Wells, M. B., Andrew, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Salivary+gland+cellular+architecture+in+the+Asian+malaria+vector+mosquito+Anopheles+stephensi.">Salivary gland cellular architecture in the Asian malaria vector mosquito Anopheles stephensi.</a> Parasites &amp; Vectors. 8, 617 (2015).</li><li>Wells, M. B., Villamor, J., Andrew, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Salivary+gland+maturation+and+duct+formation+in+the+African+malaria+mosquito+Anopheles+gambiae.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anopheles+salivary+gland+architecture+shapes+plasmodium+sporozoite+availability+for+transmission.">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://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The physiology of mosquitoes. , (1963).</li><li>Imms, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=On+the+larval+and+pupal+stages+of+Anopheles+maculipennis,+meigen.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacteria+of+the+genus+Asaia+stably+associate+with+Anopheles+stephensi,+an+Asian+malarial+mosquito+vector.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+salivary+transcriptome+of+Anopheles+gambiae+(Diptera:+Culicidae)+larvae:+A+microarray-based+analysis.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Carbonic+anhydrases+and+anion+transport+in+mosquito+midgut+pH+regulation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Slc4-like+anion+transporters+of+the+larval+mosquito+alimentary+canal.">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, and having a 90 s acetone wash. In the original adult protocol, adult SGs were dissected in 1x PBS14,15,16. Dissecting in a 25% EtOH solution helped preserve the SGs and prevented damage during the staining periods. When performing the initial dissection, it was easiest to orient the larval SGs with the head facing the non-dominant hand and having the dominant hand gently pull outwards (because the larvae are very actively moving). The improved tissue penetration achieved by washing the glands with a MeOH: glacial acetic acid solution suggests that the lipid content in the larval SG basement membrane and/or cellular plasma membranes is distinct from that of the adult SG. The 90 s acetone wash improved the clarity of glands for imaging. Although the fixation period was recommended to be at least 19 h (overnight), longer periods worked just as effectively.
The morphology of the SGs can vary widely depending on when the larvae are dissected in the 2-day long L4 stage. In early L4 dissections (day 1), the lumen appears much smaller12. In late L4 dissections (second half of day 2), the lumen is large, and the cells are elongated. We found that the optimal period for working with larvae was the L4; L1-L3 SGs are simply too small for manual dissections. In imaging results, glands dissected at mid-to-late-L4 showed smaller cells mixed with larger, laterally elongated cells, particularly in the distal sac.
Previous reports on Anopheles SG development have provided much guidance for the current studies. These reports have included illustrations of dissected embryonic and larval SGs17,18, detailed microscopic analysis of dissected glands19,20, and transcriptomic studies20,21,22. The features of the Anopheles stephensi SG were reported during the 1950s by two different groups10,11. Many of the morphological features of the larval glands were described, including the overall organization of the SG, the relative positions of distinct cell types within the organ (duct, adult SG precursors, and the larval proximal and distal secretory sacs)10,11. Both groups also reported additional morphometric data, including the number and distribution of secretory cells within each sac and their nuclear size10,11. Rishikesh showed that, like the Drosophila larval SGs, the larval SG chromosomes from Anopheles stephensi are polytenized10. These important foundational overviews described broad biological aspects of larval Anopheles SGs.
The method described herein should be useful for studying not only larval SGs of An. gambiae but can also apply to those studying other species of mosquitos. Indeed, the same protocol has been successfully utilized (unpublished) with An. stephensi, a species frequently studied in the laboratory. One limitation to the protocol is the limited number of antibodies that have been specifically generated against mosquito proteins. Although using Drosophila antibodies for highly conserved proteins has circumnavigated this limitation12, the field could benefit from more mosquito protein-specific antibodies. Understanding larval SG cellular and molecular biology can contribute to new control or target strategies and allow for the discovery of new candidate target genes for SG disruption.

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#160;KH2PO4</td>
			<td>Millipore Sigma</td>
			<td>P9791</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Na2HPO4 &#8226; 2H2O</td>
			<td>Millipore Sigma</td>
			<td>71643</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;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&#160; 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 stage, the salivary glands also form a lumen during L412. Salivary glands isolated from early L4 stage larvae stained with Hoechst reveal the proximal and distal lobes separated by a narrow constriction (Figure 3B,C). The forming lumen will extend all the way from the salivary duct at the proximal-most end (not shown) through the distal lobe. The forming lumen can be seen in the immunostained distal salivary gland of a mid-to-late L4 larva (Figure 4). The apical domains of the secretory cells surrounding the forming lumen have intense Nile Red staining (Figure 4B,C) suggestive of microvilli-like structures. Also observed close to the apical surface is Rab11 staining (Figure 4C,D, arrows). Rab11 localizes to apical recycling endosomes. The Rab11 staining that also accumulates along the basal surface of the gland is an artefact due to the stickiness of the basal membrane. Similar background staining is common with immunostaining of both larval and adult salivary glands and has been mistaken for bona fide signal.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62989/62989fig01.jpg" /> 
Figure 1:&#160;Cartoon visualization of the immunostaining process from gland dissection through slide preparation. Abbreviations: SGs = salivary glands; PBS = phosphate-buffered saline; MeOH = methanol; DAPI = 4&#8242;,6-diamidino-2-phenylindole; Abs = antibodies; LH = left hand; RH = right hand. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62989/62989fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62989/62989fig02.jpg" /> 
Figure 2: Male and female L4 stage Anopheles gambaie larvae. (A, B) Early L4 larvae. (C, D) Late L4 Larvae. There is considerable growth during the L4 stage. Females (A, C) have been described as having a distinguishing red stripe down the dorsal thorax (C; black arrow) that is not present in males (B, D), but also their antennae are much less elaborate (frilly) than those of male larvae from both early and late L4 (white arrows in enlarged images). Scale bars = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62989/62989fig02large.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/62989/62989fig03.jpg" /> 
Figure 3: Early L4 salivary glands. (A) Early L4 female larval head with salivary glands and other internal organs still attached. Salivary gland is outlined. Scale bar = 50 &#181;m. (B, C) Isolated larval glands stained with Hoechst. (B) Merge of fluorescent and Nomarski images showing the shape of the proximal and distal lobes and the blue Hoechst staining in nuclei. (C) Hoechst fluorescent image highlights nuclei only. Scale bar in bottom panels = 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62989/62989fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62989/62989fig04.jpg" /> 
Figure 4: Immunostained distal L4 salivary gland. (A) Gland has been stained with Hoechst (nuclear DNA, blue) and (B) Nile Red (membrane marker, red); immunostained for Rab11 (apical recycling vesicles, green) (C). (D) The merged image. Note that a lumen is present at the stage shown (B-D). The arrows in C and D point to the expected Rab11 staining of vesicles near the apical surface that overlaps the Nile Red-positive vesicular staining in the merge (white arrows). Non-specific background staining around the basal surface (asterisks) is also observed, a common type of background observed in larval and adult salivary glands. Scale bars = 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62989/62989fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

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 Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

Dissection z larw CNS na Drosophila melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

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 ...

Protocol for RNAi Assays in Adult Mosquitoes (A. gambiae)

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video protocol illustrates a method used by the Dimopoulos lab to inject dsRNA into Anopheles gambiae mosquitoes, which harbor the malaria parasite.  The technique manipulating the injection setup and injecting dsRNA into the thorax is illustrated.

Protocol for RNAi Assays in Adult Mosquitoes A. gambiae

Reverse genetic approaches have proven extremely useful for determining which genes underly resistance to vector pathogens in mosquitoes. This video protocol illustrates a method used by the Dimopoulos lab to inject dsRNA into Anopheles gambiae mosquitoes, which harbor the malaria parasite. The technique manipulating the injection setup and injecting dsRNA into the thorax is illustrated.

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video ...

Building a Better Mosquito: Identifying the Genes Enabling Malaria and Dengue Fever Resistance in A. gambiae and A. aegypti Mosquitoes

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections.   Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections. Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus ...

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work examining post-embryonic through adult cardiac development has been limited7-10. Examining the changing morphology of the maturing and aging heart are restricted by the lack of techniques available for staging and isolating juvenile and adult hearts. In order to analyze heart development over the fish's lifespan, we dissect zebrafish hearts at numerous stages and photograph them for further analysis11. The morphological features of the heart can easily be quantified and individual hearts can be further analyzed by a host of standard methods. Zebrafish grow at variable rates and maturation correlates better with fish size than age, thus, post-fixation, we photograph and measure fish length as a gauge of fish maturation. This protocol explains two distinct, size dependent dissection techniques for zebrafish, ranging from larvae 3.5mm standard length (SL) with hearts of 100&mu;m ventricle length (VL), to adults, with SL of 30mm and VL 1mm or larger. Larval and adult fish have quite distinct body and organ morphology. Larvae are not only significantly smaller, they have less pigment and each organ is visually very difficult to identify. For this reason, we use distinct dissection techniques.
We used pre-dissection fixation procedures, as we discovered that hearts dissected directly after euthanization have a more variable morphology, with very loose and balloon like atria compared with hearts removed following fixation. The fish fixed prior to dissection, retain in vivo morphology and chamber position (data not shown). In addition, for demonstration purposes, we take advantage of the heart (myocardial) specific GFP transgenic Tg(myl7:GFP)twu34 (12), which allows us to visualize the entire heart and is particularly useful at early stages in development when the cardiac morphology is less distinct from surrounding tissues. Dissection of the heart makes further analysis of the cell and molecular biology underlying heart development and maturation using in situ hybridization, immunohistochemistry, RNA extraction or other analytical methods easier in post-embryonic zebrafish. This protocol will provide a valuable technique for the study of cardiac development maturation and aging.

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

A clear, standardized method for dissection and isolation of the zebrafish heart at multiple developmental stages are described. Annotation and quantification techniques are also discussed.

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work ...

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called imaginal discs. These discs give rise to a high percentage of adult structures that are found within the adult fly. Here we describe a protocol that has been optimized to recover these discs and prepare them for analysis with antibodies, transcriptional reporters and protein traps. This procedure is best suited for thin tissues like imaginal discs, but can be easily modified for use with thicker tissues such as the larval brain and adult ovary. The written protocol and accompanying video will guide the reader/viewer through the dissection of third instar larvae, fixation of tissue, and treatment of imaginal discs with antibodies. The protocol can be used to dissect imaginal discs from younger first and second instar larvae as well. The advantage of this protocol is that it is relatively short and it has been optimized for the high quality preservation of the dissected tissue. Another advantage is that the fixation procedure that is employed works well with the overwhelming number of antibodies that recognize Drosophila proteins. In our experience, there is a very small number of sensitive antibodies that do not work well with this procedure. In these situations, the remedy appears to be to use an alternate fixation cocktail while continuing to follow the guidelines that we have set forth for the dissection steps and antibody incubations.

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

The adult structures of Drosophila are derived from sac-like structures called imaginal discs. Analysis of these discs provides insight into many developmental processes including tissue determination, compartment boundary establishment, cell proliferation, cell fate specification, and planar cell polarity. This protocol is used to prepare imaginal discs for light/fluorescent microscopy.

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called ...

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi), a naturally occurring phenomenon in eukaryotic organisms, is an extremely valuable tool that can be utilized in the laboratory for functional genomic studies. The ability to knockdown individual genes selectively via this reverse genetic technique has allowed many researchers to rapidly uncover the biological roles of numerous genes within many organisms, by evaluation of loss-of-function phenotypes. In the major human malaria vector Anopheles gambiae, the predominant method used to reduce the function of targeted genes involves injection of double-stranded (dsRNA) into the hemocoel of the adult mosquito. While this method has been successful, gene knockdown in adults excludes the functional assessment of genes that are expressed and potentially play roles during pre-adult stages, as well as genes that are expressed in limited numbers of cells in adult mosquitoes. We describe a method for the injection of Serine Protease Inhibitor 2 (SRPN2) dsRNA during the early pupal stage and validate SRPN2 protein knockdown by observing decreased target protein levels and the formation of melanotic pseudo-tumors in SRPN2 knockdown adult mosquitoes. This evident phenotype has been described previously for adult stage knockdown of SRPN2 function, and we have recapitulated this adult phenotype by SRPN2 knockdown initiated during pupal development. When used in conjunction with a dye-labeled dsRNA solution, this technique enables easy visualization by simple light microscopy of injection quality and distribution of dsRNA in the hemocoel.

RNAi Trigger Delivery into Anopheles gambiae Pupae

RNA interference (RNAi) is an extremely valuable tool for uncovering gene function. However, the ability to target genes using RNAi during pre-adult stages is limited in the major human malaria vector Anopheles gambiae. We describe an RNAi protocol to reduce gene function via direct injection during pupal development.

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IR-TEx: An Open Source Data Integration Tool for Big Data Transcriptomics Designed for the Malaria Vector Anopheles gambiae

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IR-TEx: An Open Source Data Integration Tool for Big Data Transcriptomics Designed for the Malaria Vector Anopheles gambiae

IR-TEx explores insecticide resistance-related transcriptional profiles in the species Anopheles gambiae. Provided here are full instructions for using the application, modifications for exploring multiple transcriptomic datasets, and using the framework to build an interactive database for collections of transcriptomic data from any organism, generated in any platform.

IR-TEx is an application written in Shiny (an R package) that allows exploration of the expression of (as well as assigning functions to) transcripts ...

Dissection and Immunostaining of Larval Salivary Glands from Anopheles gambiae Mosquitoes

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