We thank Ella Weinert, Michelle Helms and Marlis Badusche for excellent technical assistance, Jessica B&#246;rstler for analyses of Culex mosquitoes, Norbert Becker and Bj&#246;rn Pluskota for providing Aedes albopictus eggs and Olli Vapalahti for providing the virus stock. ML was supported by the Leibniz Association; grant number SAW-2014-SGN-3.

1. Prepare Female Mosquitoes
<ol>
	<li>Collect mosquitoes from a big cage by using an aspirator (approximately 400 mosquitoes per big cage).</li>
	<li>Anesthetize the mosquitoes with carbon dioxide (CO2) for 7 s.</li>
	<li>Sort 20 females per plastic vial (&#216; 50 mm &#215; 100 mm). 
	Note: If the mosquitoes wake up, use another 3 s of CO2.</li>
	<li>Starve the mosquitoes overnight.</li>
</ol>
2. Artificial Blood Meal
NOTE: All steps are performed under BSL-3 conditions.
<ol>
	<li>Prepare the infectious blood meal.
	<ol>
		<li>Dilute the virus stock to the concentration of 1 x 108 plaque forming units (PFU)/mL. 
		Note: The stock concentration was determined by performing Tissue culture infection dose 50 method (TCID50) and calculated by the Spearman &#38; K&#228;rber algorithm10,11.</li>
		<li>Mix expired human blood (blood preservation) from the blood bank (not suitable for human anymore, but useful for mosquitoes) with fructose (8% solution), filtrated bovine serum (FBS) and working solution of the virus stock (expired human blood:fructose:FBS:working solution = 5:3:1:1). 
		Note: We chose human blood, because this is the known natural mammal host of ZIKV.</li>
	</ol>
	</li>
	<li>Freeze 140 &#181;L of blood meal mix for further analysis via TCID50.</li>
	<li>Perform artificial feeding by using different feeding methods, depending on the species:
	<ol>
		<li>Aedes aegypti: Use a membrane feeding system for 30 min (1 mL per feeder).</li>
		<li>Culex spp.: Provide the infectious blood via a cotton stick overnight and do not warm up the blood (300 &#181;L per vial).</li>
		<li>Aedes albopictus: Put two droplets (50 &#181;L each) of the blood meal mix into the plastic vial and let the mosquitoes feed for 2 h. Do not warm up the blood. 
		Note: Mosquitoes will feed in the first 15 min. The prolonged incubation time is necessary due to safety reasons. The blood droplets are drained after 2 h, thus the contamination risk is reduced during the sorting of fed mosquitoes.</li>
	</ol>
	</li>
	<li>Anesthetize the mosquitoes with CO2 for 7 s. 
	NOTE: If the mosquitoes wake up, use another 3 s of CO2.</li>
	<li>Sort and count fully engorged mosquitoes into a new vial.</li>
	<li>Add a cotton pad, saturated with fructose (8% solution) between the vial and a plug.</li>
	<li>Keep the mosquitos at the designated temperature of 18 &#176;C or 27 &#176;C in the humidity of 80% for 14 or 21 days.</li>
	<li>Feed the mosquitos with fructose-saturated cotton pads. Replenish every 72 h with 1.5 mL of fructose (8% solution) per vial (up to 20 mosquitoes, depending on the feeding rate, see step 2.5).</li>
</ol>
3. Forced Salivation Assay on Day 14 or 21
NOTE: All steps are performed under BSL-3 conditions.
<ol>
	<li>Seed 2 x 104 Vero cells/per well in a 96 well plate with 200 &#181;L of growth medium (Dulbecco&#180;s modified eagle medium (DMEM) supplemented with 3% FBS, 100 U/mL penicillin, 100 mg/mL streptomycin, 5 &#181;g/mL amphotericin B, 2 mM L-glutamine, 1% non-essential amino acids (NEAA), 1% sodium pyruvate).</li>
	<li>Prepare the salivation device:
	<ol>
		<li>Place a squared glass plate (20 &#215; 20 cm) on the bench in an angle of 30&#176;. 
		Note: This is necessary due to the underpressure in the safety lab. If the plate is horizontal or vertical, liquid will leak.</li>
		<li>Place double sided adhesive tape on top of the glass plate.</li>
		<li>Role the modeling clay until it reaches a diameter of 0.5 cm and attach it horizontally 1 cm away from the double-sided adhesive tape.</li>
		<li>Cut off the first 0.3 cm of the tip of a 10 &#181;L filter tip. 
		NOTE: It is advised to prepare this before starting the experiment outside of the biosafety laboratory.</li>
		<li>Fill the filter tips with 10 &#181;L of phosphate buffered saline (PBS, pH 7.4).</li>
		<li>Place the filter tips on the modeling clay with the tip towards the sticky tape.</li>
		<li>Secure the filter tips with gentle pressure.</li>
	</ol>
	</li>
	<li>Prepare the mosquitoes:
	<ol>
		<li>Anesthetize the mosquitoes with CO2.</li>
		<li>Remove the legs and wings to immobilize the mosquitoes.</li>
		<li>Fix the living mosquito bodies on the sticky tape above a filter tip.</li>
		<li>Place the proboscis gently into the filter tip.</li>
		<li>Run the salivation assay for 30 min.</li>
		<li>Remove the filter tips and discard the clay and tape.</li>
	</ol>
	</li>
</ol>
4. Processing of Saliva
NOTE: Steps 4.1&#8211;4.6.2 are performed under BSL-3 conditions.
<ol>
	<li>Expel the contents into the reaction tubes containing 10 &#181;L of PBS.</li>
	<li>Mix gently and transfer the content into the wells of the prepared 96 well plate using one well per sample.</li>
	<li>Incubate the plate for 7 days at 37 &#176;C with 5% CO2.</li>
	<li>Use the binocular to check the cells for the presence of cytopathic effect (CPE).</li>
	<li>If the well is positive for CPE, harvest the supernatant by pipetting 140 &#181;L into a reaction tube for RNA extraction.</li>
	<li>Purify the RNA by using a RNA extraction kit.
	<ol>
		<li>Mix 140 &#181;L of the supernatant with 560 &#181;L of AVL buffer (viral lysis buffer) and incubate for 10 min at room temperature.</li>
		<li>Add 560 &#181;L of ethanol (96%) and vortex the sample. 
		NOTE: The samples are inactivated after this step and can be discharged out of the BSL-3 laboratory.</li>
		<li>Pipet 630 &#181;L of the sample onto a column and centrifuge with 6,000 x g for 1 min.</li>
		<li>Discard the filtrate, pipet the remaining 630 &#181;L on the column and repeat the centrifugation.</li>
		<li>Wash the RNA, bound to the membrane of the column, by adding 500 &#181;L of washing buffer 1 to the column and centrifuge at 6,000 x g for 1 min.</li>
		<li>Discard the collection tube and place the column into a new collection tube. Add 500 &#181;L washing buffer 2 and centrifuge at 12,500 x g for 3 min.</li>
		<li>Discard the collection tube and place the column into a 1.5 mL reaction tube.</li>
		<li>Add 50 &#181;L of elution buffer and incubate for 1 min at room temperature.</li>
		<li>Centrifuge at 6,000 x g for 1 min and then discard the column.</li>
	</ol>
	</li>
	<li>Analyze the samples for ZIKV RNA via RT-qPCR.</li>
</ol>
5. Processing of Mosquito Bodies
NOTE: Steps 5.1&#8211;5.5 are performed under BSL-3 conditions.
<ol>
	<li>Remove the bodies of the mosquito and place each body into a reaction tube. 
	NOTE: It is possible to pause the experiment at this point and store the samples at -80 &#176;C.</li>
	<li>Add 500 &#181;L of homogenization media (DMEM without any supplementals) into the reaction tube.</li>
	<li>Homogenize the body using hand motor mixer and use a new pistil for every sample.</li>
	<li>Add 200 &#181;L of the homogenate into a 96-well-plate for purification. 
	NOTE: It is possible to pause the experiment at this point and store the samples at -80 &#176;C.</li>
	<li>Inactivate the plate by incubation for 60 min at 60 &#176;C.</li>
	<li>Purify the RNA by automated nucleic acid purification. 
	NOTE: It is possible to pause the experiment at this point and store the samples at -80 &#176;C.</li>
</ol>
6. Analysis
<ol>
	<li>Quantify the Viral RNA Copies using RT-qPCR. 
	NOTE: The RNA copies were averaged over all ZIKV-positive mosquito bodies and without including the ZIKV-negative mosquitoes.</li>
	<li>Calculate the IR, defined as the number of ZIKV-positive mosquito bodies per number of fed females.</li>
	<li>Calculate the TR, defined as the number of mosquitoes with ZIKV-positive saliva per number of ZIKV-positive mosquito bodies. The transmission rate could not be calculated for species-temperature combinations with no ZIKV-positive bodies.</li>
</ol>

<ol>
	<li>Musso, D., Gubler, 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=Zika+Virus.">Zika Virus.</a> Clinical Microbiology Reviews. (3), 487-524 (2016).</li><li>Epelboin, Y., Talaga, S., Epelboin, L., Dusfour, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zika+virus:+An+updated+review+of+competent+or+naturally+infected+mosquitoes.">Zika virus: An updated review of competent or naturally infected mosquitoes.</a> PLOS Neglected Tropical Diseases. 11 (11), e0005933 (2017).</li><li>Aliota, M. T., Peinado, S. A., Osorio, J. E., Bartholomay, L. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culex+pipiens+and+Aedes+triseriatus+Mosquito+Susceptibility+to+Zika+Virus.">Culex pipiens and Aedes triseriatus Mosquito Susceptibility to Zika Virus.</a> Emerging Infectious Diseases. (10), 1857-1859 (2016).</li><li>Turell, M. J., Linthicum, K. J., Patrican, L. A., Davies, F. G., Kairo, A., Bailey, C. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vector+competence+of+selected+African+mosquito+(Diptera:+Culicidae)+species+for+Rift+Valley+fever+virus.">Vector competence of selected African mosquito (Diptera: Culicidae) species for Rift Valley fever virus.</a> Journal of Medical Entomology. 45 (1), 102-108 (2008).</li><li>Cornel, A. J., Jupp, P. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+three+methods+for+determining+transmission+rates+in+vector+competence+studies+with+Culex+univittatus+and+West+Nile+and+Sindbis+viruses.">Comparison of three methods for determining transmission rates in vector competence studies with Culex univittatus and West Nile and Sindbis viruses.</a> Journal of the American Mosquito Control Association. 5 (1), 70-72 (1989).</li><li>Hurlbut, H. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosquito+salivation+and+virus+transmission.">Mosquito salivation and virus transmission.</a> The American Journal of Tropical Medicine and Hygiene. 15 (6), 989-993 (1966).</li><li>Anderson, S. L., Richards, S. L., Smartt, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+method+for+determining+arbovirus+transmission+in+mosquitoes.">A simple method for determining arbovirus transmission in mosquitoes.</a> Journal of the American Mosquito Control Association. 26 (1), 108-111 (2010).</li><li>Heitmann, A., Jansen, S., L&#252;hken, R., Leggewie, M., Badusche, M., Pluskota, B., 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=Experimental+transmission+of+Zika+virus+by+mosquitoes+from+central+Europe.">Experimental transmission of Zika virus by mosquitoes from central Europe.</a> Eurovsurveillance. 22 (2), (2017).</li><li>Hierholzer, J. C., Killington, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Virus Isolation and Quantitation - Virology methods manual. , (1996).</li><li>Horzinek, R. W., Marian, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kompendium+der+allgemeinen+Virologie.">Kompendium der allgemeinen Virologie.</a> Zentralblatt f&#252;r Veterin&#228;rmedizin R A. 32 (1-10), 480 (2010).</li><li>Richard, V., Paoaafaite, T., Cao-Lormeau, V. -. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vector+Competence+of+French+Polynesian+Aedes+aegypti+and+Aedes+polynesiensis+for+Zika+Virus.">Vector Competence of French Polynesian Aedes aegypti and Aedes polynesiensis for Zika Virus.</a> PLOS Neglected Tropical Diseases. 10 (9), (2016).</li><li>Fortuna, C., Remoli, M. E., Di Luca, M., Severini, F., Toma, L., Benedetti, E., 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=Experimental+studies+on+comparison+of+the+vector+competence+of+four+Italian+Culex+pipiens+populations+for+West+Nile+virus.">Experimental studies on comparison of the vector competence of four Italian Culex pipiens populations for West Nile virus.</a> Parasites &amp; Vectors. 8, 463 (2015).</li><li>Weger-Lucarelli, J., R&#252;ckert, C., Chotiwan, N., Nguyen, C., Garcia Luna, S. M., Turell, M. 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=Vector+Competence+of+American+Mosquitoes+for+Three+Strains+of+Zika+Virus.">Vector Competence of American Mosquitoes for Three Strains of Zika Virus.</a> PLOS Neglected Tropical Diseases. 10 (10), (2016).</li><li>Dubrulle, M., Mousson, L., Moutailler, S., Vazeille, M., Failloux, A. -. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chikungunya+virus+and+Aedes+mosquitoes:+saliva+is+infectious+as+soon+as+two+days+after+oral+infection.">Chikungunya virus and Aedes mosquitoes: saliva is infectious as soon as two days after oral infection.</a> PLOS ONE. 4 (6), e5895 (2009).</li></ol>

The authors have nothing to disclose.

It has previously been shown that results obtained with forced salivation are in line with classical re-feeding experiments6. However, both in re-feeding experiments and in our presented method of forced salivation, it is impossible to directly proof salivation activity or monitor the release of saliva. To prove salivation activity, samples could be tested for other components present in the saliva (e.g., proteins, carbohydrates, etc.). Furthermore, additional qPCRs would allow detection of viral RNA copies. This, however, requires splitting of saliva samples for various analyses, which may limit sensitivity in the cell culture assay in case of very low particle numbers. In turn, this might lead to an underestimation of the determined transmission rates.
For reproducible results, it is necessary to ensure that all mosquitoes stay alive until the end of the experiment. This is controlled by visually monitoring the body movement activity of the mosquitoes. Furthermore, the principle use of the salivation assay for a given mosquito species has to be tested by feeding control viruses, which are known to be transmitted by this specific mosquito species. Vice versa, every virus introduced into an experiment has to be tested in a susceptible mosquito.
Forced salivation assays have many advantages. High numbers of mosquitoes can be tested simultaneously on a single specimen base under controlled standardized conditions without conflicting with animal welfare regulations9.
The method is used by several laboratories. However, exact setups may differ, which can lead to differences in results between laboratories. One critical component is the capillary to collect the saliva, which can be made of glass14 or plastic15. To comply with the high safety regulations of a biosafety level 3 insectary, we used plastic filter tips instead of glass capillaries, thus reducing the risk of injuries. Furthermore, filter tips have the advantage that the collected liquid can easily be transferred using a pipette.
The setup of the forced salivation assay presented here is based on two investigators working together and sharing duties. Demobilizing of the mosquito is performed by one person, while the other simultaneously prepares the forced salivation setup. This significantly reduces handling time and minimizes the risk of sample switching as changing between the demobilization-workplace and the salivation-plate is not necessary. Moreover, it allows placing the mosquitoes on the forced salivation device immediately after demobilization. This rather short mosquito handling time is essential for successful forced salivation. Lastly, the mosquitoes can be directly monitored for motility throughout the whole experiment.
The salivation assay presented here is used to gain insights into the vector potential of mosquito species. However, to answer some other specific questions (e.g., the numbers of mosquito bites that are needed to infect vertebrates), animal experiments might still be required.

In 2015, Zika virus (ZIKV), a mosquito-borne virus (mobovirus) belonging to the family Flaviviridae, emerged in Columbia and Brazil and spread rapidly across the Americas and the Caribbean, causing an epidemic with notable numbers of associated clinical cases of microcephaly and Guillain&#8211;Barr&#233; syndrome1. Mosquitoes of the species Aedes aegypti and Aedes albopictus are considered the primary and secondary vectors of ZIKV2, but other species of the genus Aedes have also proven to have experimental vector competence3. In contrast to Aedes species, most of the Culex species tested so far are not able to transmit the virus4. Merely, the data for Culex quinquefasciatus provided inconclusive results3. At present, information is lacking on mosquito vector competence for ZIKV under moderate temperature conditions (&#60;20 &#176;C). However, this information is substantial for the risk assessments of possible spreads to regions with temperate climate such as Central Europe.
A competent vector for ZIKV is able to acquire, maintain, amplify and finally transmit the virus. Therefore, testing the mosquito bodies or parts of the bodies, including legs, midgut, head or salivary glands, for ZIKV is not sufficient to determine vector competence. It is mandatory to test for infectious virus particles within released saliva. To assess vector competence, both infection rate (IR, number of ZIKV-positive mosquito bodies per number of engorged mosquitoes) and transmission rate (TR, number of mosquitoes with ZIKV-positive saliva per number of ZIKV-positive mosquito bodies), have to be determined. IRs of mosquitoes can be easily analyzed by simply homogenizing mosquitoes followed by a reverse-transcriptase real-time polymerase chain reaction (RT-qPCR) targeting ZIKV. Moreover, the infection can be further characterized by determining the viral copy numbers per mosquito. In contrast, the analysis of transmission rates relies on the detection of viable virus particles in the saliva of individual mosquitoes. This can be achieved by feeding of infected mosquitoes on susceptible host animals, followed by the analysis of the viremia in the host5. However, this method relies on suitable model organisms, and is costly and restricted by animal-welfare regulations. To avoid the use of laboratory animals for analyzing transmission rates and to reduce costs, artificial re-feeding systems using blood droplets have been described6. However, testing on individual scale combined with a high number of individuals is difficult and time-consuming. The first description of a salivation assay on individual scale was published in 19667 and during recent years, forced salivation experiments have become the method of choice to evaluate the vector competence of mosquitoes3,8.
Based on our vector competence study of Central European mosquitoes published recently9, we describe forced salivation as reported by Anderson et al.8 with some modifications. This method allows high throughput standardized experiments under high biosafety level conditions (BSL-3) and includes the identification of active virus by cell culture based assays. The experiments described here include 856 mosquitoes. They were infected with ZIKV by artificial blood meal and subsequently analyzed for the presence of infectious virus particles in the saliva. The mosquitoes introduced into the experiments comprised two long-established laboratory populations of Ae. aegypti and Culex pipiens pipiens biotype molestus (Cx. p. molestus), as well as field caught Culex pipiens pipiens biotype pipiens (Cx. p. pipiens), Culex torrentium (Cx. torrentium) and two Ae. albopictus populations. With the exception of one of the two Ae. albopictus populations, which was collected in Italy, all mosquitoes were collected in Germany.

<table><tbody><tr><td>Inject+matic Sleeper</td><td>Inject+matic</td><td></td><td></td></tr><tr><td>Bugdorm-1 Insect Rearing Cage</td><td>BugDorm Store</td><td></td><td></td></tr><tr><td>Drosophila cultivation tubes</td><td>carl roth</td><td>PK11.2</td><td>plastic vial for mosquitoes</td></tr><tr><td>Plug for drosophila cultivation tubes</td><td>carl roth</td><td>PK15.1</td><td></td></tr><tr><td>Constant climate chamber</td><td>Binder</td><td>KBF-240</td><td></td></tr><tr><td>Blood</td><td></td><td></td><td>banked human blood, expired, not usable for medical application</td></tr><tr><td>Dumont-Pincette Electronic SS</td><td>Dumont</td><td>B40c</td><td></td></tr><tr><td>Vero cells</td><td>BNITM</td><td></td><td></td></tr><tr><td>Flash light aspirator</td><td>Bioquip</td><td>2809</td><td></td></tr><tr><td>double-sided adhesive tape</td><td></td><td></td><td>no specific provider</td></tr><tr><td>Gammex Powder-free gloves with AMT</td><td>Ansell</td><td></td><td>for safety reasons necessary during work with pricks in the BSL3 laboratory</td></tr><tr><td>RealStar Zika Virus RT-PCR-Kit</td><td>altona diagnostics</td><td>591013</td><td></td></tr><tr><td>MagMax Pathogen RNA/DNA Kit</td><td>Thermo fisher scientific</td><td>4462359</td><td>RNA isolation of mosquito bodies</td></tr><tr><td>Qiamp viral RNA Mini Kit</td><td>Qiagen</td><td>52906</td><td>RNA-Isolation of supernatant</td></tr><tr><td>filter tips (20 &amp;micro;L)</td><td>Sarstedt</td><td>701,116,210</td><td>Cut the first 3 mm of the tip</td></tr><tr><td>hand motor mixer</td><td>carl roth</td><td>sold out</td><td></td></tr><tr><td>micro pestles for hand motor mixer</td><td>carl roth</td><td>CXH8.1</td><td></td></tr><tr><td>DMEM</td><td></td><td>P0403550</td><td></td></tr><tr><td>L Glutamine Penicilin/Streptomycin</td><td>PAN</td><td>P0819100</td><td></td></tr><tr><td>Amphotericin B</td><td>PAN</td><td>P06-0110</td><td></td></tr><tr><td>Fructose</td><td>carl roth</td><td>4981.5</td><td></td></tr><tr><td>Phosphate buffered saline</td><td>PAN</td><td>P04-36500</td><td></td></tr><tr><td>FBS</td><td>PAN</td><td></td><td></td></tr><tr><td>Sodium Pyruvat</td><td>PAN</td><td>P0443100</td><td></td></tr><tr><td>MEM NAA</td><td>PAN</td><td>P0832100</td><td></td></tr><tr><td>Hemotek Feeder</td><td>Hemotek</td><td>PS6</td><td></td></tr><tr><td>Light Cycler</td><td>Roche</td><td>480 II</td><td></td></tr><tr><td>Light Cycler Software</td><td>Roche</td><td>480 SW 1.5.1</td><td></td></tr><tr><td>Modeling clay</td><td></td><td></td><td>no specific provider</td></tr><tr><td>King Fisher Flex Purification System</td><td>ThermoFisher scientific</td><td></td><td></td></tr><tr><td>&lt;em&gt;Aedes albopictus&lt;/em&gt;</td><td></td><td></td><td>eggs were collected in Freiburg, Germany</td></tr><tr><td>Binocular</td><td>MOTIC</td><td>SMZ-168</td><td></td></tr><tr><td>Binocular light</td><td>MOTIC</td><td>MLC-150C</td><td></td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt;-Incubator</td><td>Thermo scientific</td><td>MIDI 40</td><td></td></tr></tbody></table>

forced salivation as a method to analyze vector competence of mosquitoes

Vector competence is defined as the potential of a mosquito species to transmit a mosquito-borne virus (mobovirus) to a vertebrate host. Viable virus particles are transmitted during a blood meal via the saliva of an infected mosquito. Forced salivation assays allow determining the vector potential on the basis of single mosquitoes, avoiding the use of animal experiments. The method is suitable to analyze a large number of mosquitoes in one experiment within a short period of time. Forced salivation assays were used to analyze 856 individual mosquitoes trapped in Germany, including two different Culex pipiens pipiens biotypes, Culex torrentium as well as Aedes albopictus, which were experimentally infected with Zika virus (ZIKV) and incubated at 18 &#176;C or 27 &#176;C for two and three weeks. The results indicated the lack of vector competence of the different Culex taxa for ZIKV. In contrast, Aedes albopictus was susceptible to ZIKV, but only at 27 &#176;C, with transmission rates similar to an Aedes aegypti laboratory colony tested in parallel.

Aedes aegypti is known as a competent vector for ZIKV at least under tropical climatic conditions12. Therefore, we used Ae. aegypti as a positive control to establish the assay and to analyze IRs as well as TRs at an incubation temperature of 27 &#176;C and a humidity of 80%. IR and TR were defined as previously described by Fortuna et al.13. Challenged Ae. aegypti showed IR of 50% and 72% and TR of 42% and 31% at 14 and 21 days post infection (dpi), respectively (Figure 1A and 1B). Subsequently, European mosquitoes were tested for ZIKV infection and transmission using tropical as well as temperate climatic conditions.
At 27 &#176;C incubation temperature, all mosquito species showed ZIKV infection at 14 dpi as well as 21 dpi, except Cx. p. pipiens and Cx. torrentium individuals, which were positive at 14 dpi only. All Aedes species showed higher IRs (between 50% and 72%) as well as higher viral RNA concentration (106 up to 109 RNA copies/mosquito specimens) in comparison to the different Culex taxa, which revealed IRs ranging between 0% and 32% and viral loads of 103 to 104 RNA copies/mosquito specimens. All Aedes species displayed ZIKV-positive saliva (TR 13&#8211;42%), whereas none of the saliva samples from the Culex taxa contained ZIKV (Figure 1A and 1B). Interestingly, transmission rates at 21 dpi were similar in Ae. aegypti and Ae. albopictus from Germany (30%), but were substantially lower in Ae. albopictus from Italy (13%).
Incubation of challenged mosquitoes at a moderate temperature of 18 &#176;C revealed generally lower virus loads in the Aedes species (104&#8211;106 RNA copies/mosquito) as well as in the different Culex taxa (102&#8211;104 RNA copies/mosquito). However, at 18 &#176;C incubation temperature, transmission of ZIKV was not observed for any of the mosquito species investigated.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57980/57980fig1.jpg" /> 
Figure 1: Transmission rates of mosquitoes experimentally infected with ZIKV. Results of vector competence studies with ZIKV in different mosquito species. Six different mosquito taxa were infected with ZIKV and incubated at 27 &#176;C or 18 &#176;C for 14 (A) or 21 (B) days. TR is defined as the number of mosquitoes with ZIKV-positive saliva per number of ZIKV-positive mosquito bodies. The total number of mosquitoes investigated was 856, with a minimum of 30 individuals for each time point/temperature/mosquito taxa. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/57980/57980fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Forced Salivation As a Method to Analyze Vector Competence of Mosquitoes

For efficient control of mosquito borne virus transmission, the knowledge of the vector potential of respective mosquitoes is of particular interest. We describe forced salivation as a method to analyze vector competence of Aedes albopictus and three different Culex taxa for the transmission of Zika virus.

Vector competence is defined as the potential of a mosquito species to transmit a mosquito-borne virus (mobovirus) to a vertebrate host. Viable virus ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

9.5K Views.  Bernhard Nocht Institute for Tropical Medicine. For efficient control of mosquito borne virus transmission, the knowledge of the vector potential of respective mosquitoes is of particular interest. We describe forced salivation as a method to analyze vector competence of Aedes albopictus and three different Culex taxa for the transmission of Zika virus.

Video: Forced Salivation As a Method to Analyze Vector Competence of Mosquitoes

Alphavirus Transducing System:  Tools for Visualizing Infection in Mosquito Vectors

Alphavirus transducing systems (ATSs) are important tools for expressing genes of interest (GOI) during infection. ATSs are derived from cDNA clones of mosquito-borne RNA viruses (genus Alphavirus; family Togaviridae). The Alphavirus genus contains about 30 different mosquito-borne virus species. Alphaviruses are enveloped viruses and contain single-stranded RNA genomes (~11.7 Kb). Alphaviruses transcribe a subgenomic mRNA that encodes the structural proteins of the virus required for encapsidation of the genome and maturation of the virus. Alphaviruses are usually highly lytic in vertebrate cells, but persistently infect susceptible mosquito cells with minimal cytopathology. These attributes make them excellent tools for gene expression in mosquito vectors. The most common ATSs in use are derived from Sindbis virus (SINV). The broad species tropism of SINV allows for infection of insect, avian, and mammalian cells8. However, ATSs have been derived from other alphaviruses as well9,10,20. Foreign gene expression is made possible by the insertion of an additional viral subgenomic RNA initiation site or promoter. ATSs in which an exogenous gene sequence is positioned 5' to the viral structural genes is used for stable protein expression in insects. ATSs, in which a gene sequence is positioned 3' to the structural genes, is used to trigger RNAi and silence expression of that gene in the insect.
ATSs have proven to be valuable tools for understanding vector-pathogen interactions, molecular details of viral replication and maintenance infectious cycles3,4,11,19,21. In particular, the expression of fluorescent and bioluminescent reporters has been instrumental tracking the viral infection in the vector and virus transmission5,14-16,18. Additionally, the vector immune response has been described using two strains of SINV engineered to express GFP2,9.
Here, we present a method for the production of SINV containing a fluorescent reporter (GFP) from the cDNA infectious clone. Infectious, full-length RNA is transcribed from the linearized cDNA clone. Infectious RNA is introduced into permissive target cells by electroporation. Transfected cells generate infectious virus particles expressing the GOI. Harvested virus is used to infect mosquitoes, as described here, or other host species (not shown herein). Vector competence is assessed by detecting fluorescence outside the midgut or by monitoring virus transmission7. Use of a fluorescent reporter as the GOI allows for convenient estimation of virus spread throughout a cell culture, for determination of rate of infection, dissemination in exposed mosquitoes, virus transmission from the mosquito and provides a rapid gauge of vector competence.

Methods for using alphavirus transducing systems to express fluorescent reporters in vitro and in adult mosquitoes are described. This technique may be adapted to express any protein of interest in lieu of or in addition to a reporter.

Alphavirus transducing systems (ATSs) are important tools for expressing genes of interest (GOI) during infection.  ATSs are derived from cDNA clones of ...

Immunology and Infection

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are delivered using a non-quantitative method. Accurate delivery of known quantities of nucleic acids or other chemicals is critical to measure the efficacy of the compound being tested and to allow reliable comparison between compounds.
Here we provide a reproducible, quantitative microinjection protocol that ensures accurate delivery of specific doses of dsRNA, reducing the mortality typically induced by injection injury. These modifications include the addition of Rhodamine B, a graduated injection needle, and an improved recovery method borrowed from Isoe and Collins. This method allows calculation of dose responses and facilitates comparisons between compounds. Versions of this method have been successfully used on three genera of mosquitoes as well as house flies to assess the reduction in fecundity resulting from gene silencing of ribosomal RNA transcripts.
This protocol provides strategies to reduce several challenges of small insect microinjection. Together, mechanical delivery of dsRNAs accompanied by visual verification, identification of effective locations for delivery, and inclusion of a post-injection recovery period ensure accurate dosing and low injury mortality. This protocol also describes an oviposition bioassay for uniform determination of effects on fecundity.

This protocol describes a microinjection methodology that we have standardized and used for several years to deliver specific quantities of nucleic acids directly to the hemolymph of mosquitoes and house flies. This protocol results in minimal injection mortality and allows dose correlated measurements of fecundity.

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are ...

Environment

Use of a Recombinant Mosquito Densovirus As a Gene Delivery Vector for the Functional Analysis of Genes in Mosquito Larvae

In vivo microinjection is the most commonly used gene transfer technique for analyzing the gene functions in individual mosquitoes. However, this method requires a more technically demanding operation and involves complicated procedures, especially when used in larvae due to their small size, relatively thin and fragile cuticle, and high mortality, which limit its application. In contrast, viral vectors for gene delivery have been developed to surmount extracellular and intracellular barriers. These systems have the advantages of easy manipulation, high gene transduction efficiency, long-term maintenance of gene expression, and the ability to produce persistent effects in vivo. Mosquito densoviruses (MDVs) are mosquito-specific, small single-stranded DNA viruses that can effectively deliver foreign nucleic acids into mosquito cells; however, the replacement or insertion of foreign genes to create recombinant viruses typically causes a loss of packaging and/or replication abilities, which is a barrier to the development of these viruses as delivery vectors.
Herein, we report using an artificial intronic small-RNA expression strategy to develop a non-defective recombinant Aedes aegypti densovirus (AaeDV) in vivo delivery system. Detailed procedures for the construction, packaging and quantitative analysis of the rAaeDV vectors, and for larval infection are described.
This study demonstrates, for the first time, the feasibility of developing a non-defective recombinant MDV micro RNA (miRNA) expression system, and thus providing a powerful tool for the functional analysis of genes in mosquito and establishing a basis for the application of viral paratransgenesis for controlling mosquito-borne diseases. We demonstrated that Aedes albopictus 1st instar larvae could be easily and effectively infected by introducing the virus into the water body of the larvae breeding site and that the developed rAaeDVs could be used to overexpress or knock down the expression of a specific target gene in larvae, providing a tool for the functional analysis of mosquito genes.

We report using an artificial intronic small RNA expression strategy to develop a non-defective recombinant Aedes aegypti densovirus (AaeDV) in vivo delivery system. A detailed procedure for the construction, packaging, and quantitative analysis of the rAaeDV vectors as well as for larval infection is described.

In vivo microinjection is the most commonly used gene transfer technique for analyzing the gene functions in individual mosquitoes. However, this method ...

Developmental Biology

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is based on the injection of non-pathogenic particles into insects, as the particles will be detected by the immune system and will induce the production of immune effectors. We focus here on the stimulation of the melanization response in the mosquito Anopheles gambiae. The melanization response results in the encapsulation of foreign particles and parasites with a dark layer of melanin. To stimulate this response, mosquitoes are inoculated with beads in the thoracic cavity using microcapillary glass tubes. Then, after 24 hr, the mosquitoes are dissected to retrieve the beads. The degree of melanization of the bead is measured using image analysis software. Beads do not have the pathogenic effects of parasites, or their capacity to evade or suppress the immune response. These injections are a way to measure immune efficacy and the impact of immune stimulations on other life history traits, such as fecundity or longevity. It is not exactly the same as directly studying host-parasite interactions, but it is an interesting tool to study immunity and its evolutionary ecology.

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

Through inoculation with beads, the described technique enables the stimulation of the mosquito melanization response in the hemolymph circulating system. The amount of melanin covering the beads can be measured after dissection as a measure of the immune response.

The stimulation of immune responses is a common tool in invertebrate studies to examine the efficacy and the mechanisms of immunity. This stimulation is ...

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. ...

Vector Competence Analyses on Aedes aegypti Mosquitoes using Zika Virus

The procedures presented describe a generalized methodology to infect Aedes aegypti mosquitoes with Zika virus under laboratory conditions to determine the rate of infection, disseminated infection, and potential transmission of the virus in the mosquito population in question. These procedures are widely utilized with various modifications in vector competence evaluations globally. They are important in determining the potential role that a given mosquito (i.e., species, population, individual) may play in the transmission of a given agent.

Vector Competence Analyses on Aedes aegypti Mosquitoes using Zika Virus

The presented protocol can determine the vector competence of Aedes aegypti mosquito populations for a given virus, such as Zika, in a containment setting.

The procedures presented describe a generalized methodology to infect Aedes aegypti mosquitoes with Zika virus under laboratory conditions to determine ...

Experimental Viral Infection in Adult Mosquitoes by Oral Feeding and Microinjection

Mosquito-borne viruses (MBVs), which are infectious pathogens to vertebrates, are spread by many mosquito species, posing a severe threat to public health. Once ingested, the viruses must overcome the mosquito midgut barrier to reach the hemolymph, from where they might potentially spread to the salivary glands. When a mosquito bites, these viruses are spread to new vertebrate hosts. Similarly, the mosquito may pick up different viruses. In general, only a tiny portion of viruses may enter the salivary glands via the gut. The transmission efficiency of these viruses to the glands is be affected by the two physical barriers found in different mosquito species: midgut barriers and salivary glands barriers. This protocol presents a method for virus detection in salivary glands of Aedes aegypti&#39;s&#160;following oral feeding and intrathoracic injection infection. Furthermore, determining whether the guts and/or salivary glands hinder viral spread can aid in the risk assessments of MBVs transmitted by Aedes aegypti.

This methodology, which included oral feeding and intrathoracic injection infection, could effectively assess the influence of midgut and/or salivary gland barriers on arbovirus infection.

Mosquito-borne viruses (MBVs), which are infectious pathogens to vertebrates, are spread by many mosquito species, posing a severe threat to public ...

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) ...

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. Insecticide resistance surveillance of mosquito populations is typically done through Centers for Disease Control and Prevention (CDC) bottle bioassays or World Health Organization (WHO) tube tests. However, these methods can result in a high degree of variability in mortality data due to variable insecticide contact with&#160;the insect, the relatively small numbers of organisms tested, extensive variation in mass between populations, and constantly changing environmental conditions, leading to variable outcomes. This paper presents the topical application bioassay, adapted as a high-throughput phenotypic bioassay for both mosquitoes and fruit flies, to test large numbers of insects along&#160;a range of insecticide concentrations.
This assay 1) ensures consistent treatment and insecticide contact with every organism, 2) produces highly specific dose-response curves that account for differences in average mass between strains and sexes (which is particularly important for field-collected organisms), and 3) allows for the calculation of statistically rigorous median lethal doses (LD50), which are necessary for resistance ratio comparisons-an alternative surveillance approach from diagnostic dose mortality, which is also used for larvicide resistance surveillance. This assay will be a complementary tool for accurately phenotyping mosquito populations and, as illustrated using fruit flies, is easily adaptable for use with other insects. We argue that this assay will help fill the gap between genotypic and phenotypic insecticide resistance in multiple insect species.

We describe the methodology and importance of the topical application bioassay to measure insecticide susceptibility in mosquitoes and fruit flies. The presented assay is high-throughput, utilizes insect mass-thus allowing for calculating a mass-relativized lethal dose instead of concentration-and likely has lower variability than other similar methods.

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. ...

Mosquito Feeding Assay: A Method to Evaluate Test Compound Effects on Mortality and Fecundity of Adult Female Mosquitoes

Collect approximately 150 four- to five-day-old adult female mosquitoes with an aspirator, and transfer them to a separate cage. Remove the source of sugar 1 to 24 hours prior to the feeding assay.
Prepare an 80-millimolar stock solution of the testing compound in water. Then, serially dilute it with water to obtain working stock solutions of the desired concentrations.
To obtain desired testing concentrations, add 40 microliters of each dilution to 960 microliters of defibrinated rabbit blood in a 1.5-milliliter tube, and mix by pipetting. Place a membrane filter on a feeding unit, and seal it with a rubber ring.
Next, use a pipette to transfer 1 milliliter of the blood with testing solution through the delivery port located on the reverse side of the feeding unit. Attach the feeding unit to a heating unit. Then, swab the membrane surface gently with freshly prepared 10% lactic acid solution.
Place the feeding unit in the cage. Cover the cage with a dark cloth, and allow the mosquitoes to feed for one hour. After the mosquitoes feed, place the cage in a refrigerator at 4 degrees Celsius for five minutes to anesthetize the mosquitoes. Then, count and record the total number of mosquitoes in the cage.
By examining the abdomen, count and record the total number of fully- and partially-fed female mosquitoes. A minimum of 50 blood-fed mosquitoes should be obtained per dose. Remove any mosquitoes that have not fed. Next, transfer the cage to a growth chamber and use a score sheet to record the number of dead mosquitoes at the appropriate time points.
On day 3 post-blood feeding, place an egg cup in the cage for 72 hours. Use a dissecting microscope to count the total number of eggs produced per treatment.

Forced Salivation As a Method to Analyze Vector Competence of Mosquitoes

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Alphavirus Transducing System: Tools for Visualizing Infection in Mosquito Vectors

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Use of a Recombinant Mosquito Densovirus As a Gene Delivery Vector for the Functional Analysis of Genes in Mosquito Larvae

Inoculating Anopheles gambiae Mosquitoes with Beads to Induce and Measure the Melanization Immune Response

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

Vector Competence Analyses on Aedes aegypti Mosquitoes using Zika Virus

Experimental Viral Infection in Adult Mosquitoes by Oral Feeding and Microinjection

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

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

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