Methods in Anopheles Research - MR4

\Methods inAnopheles ResearchSecond Edition 2010

0-1 Preface.docPage 2 of 2May it serve well those working to reduce malaria and other vector-borne diseases by the study of theirhosts.May, 2007 Mark Q. Benedict CDC, Atlanta USAReferencesClements AN (1992) The Biology of Mosquitoes: Development, Nutrition and Reproduction. Chapman &Hall, LondonClements AN (1999) The Biology of Mosquitoes: Sensory Reception and Behaviour. CABI Publishing,New YorkGerberg EJ, Barnard DR, Ward RA (1994) Manual for mosquito rearing and experimental techniques,revised ed. American Mosquito Control Association, Inc., Lake CharlesTrembley HL (1944) Mosquito culture technique. Mosquito News 4:103-119

Preface to Second EditionWhen Mark Benedict and the MR4 Vector Activity Team launched the first edition of Methods inAnopheles Research in 2007, they never imagined its enormous success. This manual has becomeone of the most accessed items in the MR4 Vector Activity website, providing practical techniques to theintrepid researcher tasked with the “dicey” activity of rearing and investigating anopheline mosquitoes. Ithas become a valuable resource not only for the scientist conducting basic research but also for publichealth entomologist from malaria endemic countries and members of the vector control industry.The first edition of Methods in Anopheles Research was “the latest incarnation” of training materialsthat Mark had developed for his personnel over many years. In the preface of that version, he invitedother researchers to contribute protocols and to make corrections. And they did. Several minor revisionswere made and a few new techniques were added periodically over the following couple of years. As themanual became better known, the number of suggested additions increased. Because the new materialdid not fit well in the original chapters, we realized that we would need to make more than minor changesand could no longer consider it the same edition. This second edition of the Methods in AnophelesResearch reflects those modifications. Many of the previous chapters remain much the same, new oneshave been added to accommodate the new procedures and a few have been rearranged. It still remainsa work in progress and as Mark requested in the first edition, and we continue to urge researchers tocritique and to contribute.We thank all the people in the last several years that have made comments and suggestions and areespecially grateful to those who offered their techniques including the following:Claudia AliagaFrancis AtieliMelissa AveryChris BassJames BecnelNora BesanskyDmitri BoudkoWilliam BrogdonAdeline ChanAnthony CornelMartin DonnellyLin FieldChristen FornadelRalph HarbachClare HolleleyTheresa HowardChristian KaufmannRebekah KentMarc KlowdenLizette KoekemoerGreg LanzaroFrédéric LardeuxPaul LinserYvonne LintonJohn MorganUlrike MunderlohMarco NeiraK.R. Ng’habiDouglas NorrisHilary RansonRosenka TajerinaLeslie VanEkerisJohn VontasBradley WhiteElien WilkinsMartin WilliamsonRobert Wirtz

I would like to give special recognition to the invaluable contributions of Paul Howell and Alice Sutcliffe,whose diligence and persistence made this second edition possible.Finally I would thank all the users who have made this such a popular download and I hope that themanual remains a valuable resource for those working to reduce malaria and other vector-bornediseases.Sincerely,Ellen M. Dotson, Principal Investigator for MR4 Vector Activity, CDC Atlanta GA, USAApril, 2011

Chapter 1 : Insectary Operation1.1 Equipping and Operating an InsectaryPage 1 of 4Chapter 1: Insectary Operation1.1 Equipping and Operating an InsectaryMark BenedictIntroductionMosquito insectaries vary widely in their sophistication and cost. Fortunately, the requirements for goodmosquito culture are easily met and can be achieved by both simple (inexpensive) and complex(expensive) means. In the following section, I will present very personal prejudices and experiences toguide you. My objective is to convey approaches that are sufficiently useful, safe, and - where possible -inexpensive. Additional ideas can be found in Gerberg (1994), and many of those below are also found inBenedict (1997).TemperatureConstant temperature is the most important environmental criterion. Immature mosquitoes are typicallycultured at a water temperature of 27°C. There are several ways in which this can be accomplished.• Standalone incubators. These are a good choice particularly where temperature experiments areplanned or space cannot be dedicated to a mosquito insectary. Care should be given to the shelfspacing and sizes so that the space can be occupied efficiently with the trays you plan to use formosquito culture. The disadvantage is that floor space is not efficiently used relative to someother systems, and the probability of equipment failures multiplies with each additional unit. Thisapproach does provide a good way to divide work spaces for different individuals, stocks orspecies.• Walk-in incubators are often used for larger facilities. These are usually equipped with rustresistantshelving giving greater flexibility for space use. They also allow entry of carts totransport materials and should not have raised thresholds.• Air-conditioned rooms. Dedicated rooms for mosquito culture require a commitment of the spacesfor this activity. Ideally they are designed with water-resistant wall- and floor-coverings such astile or a monolithic material. These provide the greatest flexibility but often mean that all insectaryactivities will be performed in the hot humid environment. They often contain screened enclosuresto separate activities or stocks.• Heater tapes (Dame et al. 1978) and shelves. Means have been devised to heat trays of larvaeby placing heating elements under the trays or shelves. This is a flexible method that providescomfort for personnel. However, the equipment must be devised on an ad hoc basis, and traysmust be covered to prevent water loss and evaporative cooling if the room is not humidified.Relative HumidityIn all of the designs listed under temperature above, humidity is necessary only for adults. 80% relativehumidity to maintain adults is an often-mentioned value that possibly requires more experimental support.None-the-less, one should plan on being able to reach this level. Excessively high humidity is harmful toadult mosquitoes and must be prevented. The only benefit of high humidity to immatures is to preventwater loss and evaporative cooling, both of which can be prevented by covering containers with animpermeable cover. In rough order of descending space capacity, means to generate humidity are:• Steam injection into the central ventilation• Greenhouse-type misting humidifiers• Centrifugal room humidifiers

Chapter 1 : Insectary Operation1.1 Equipping and Operating an InsectaryPage 2 of 4• Evaporative coolers• Home-type steam humidifiers• Boiling water steam generators e.g. in standalone incubators• EvaporationConsistent humidification is a chronically difficult goal due to removal of water from the air the cycling oftemperature control systems and the intrinsic unreliability of humidity creation and control systems withwhich I have had experience. In order to develop a simple and reliable system that satisfies your specificneeds, I recommend you answer the following questions:1. Do you need to humidify the immatures culture space? If you do, a larger capacity active system willbe required. It is preferable for worker comfort and ease to simply humidify only the adult holding areaand cover the immatures’ containers.2. Can I hold the adults I have in a relatively small area? If you are using small cages, cups etc. a verysimple and effective solution is to use a glass or Plexiglas case in which cups of water containingsponges are placed on the shelves. Such a passive system is effective, inexpensive and foolproof.3. Do I need to physically segregate large numbers of adults? In this case, multiple systems, subdividedrooms, or incubators will be required.Two of the above systems are inexpensive and self-regulating: evaporation in a sealed space andevaporative coolers. Both of these methods will attain sufficient humidity and require no controls.Centrifugal room humidifiers and misting systems suffer the problem that droplet sizes are often too largeresulting in puddles and/or reservoirs of water in which microbes can grow. Of the high capacity systems,steam injection into the ventilation system has been most reliable for us. If possible, the ductwork must beconstructed of stainless steel since the high humidity will quickly rust it.LightingMany insectaries use a 12:12 light:dark schedule. This is easily accomplished using a simple light timer.Most laboratories also try to achieve gradual dimming and lightening to stimulate natural behavior. Thisfeature can be purchased with many incubators but is less easily accomplished in dedicated rooms. In thelatter case, a control system, dimming light fixtures and bulbs must be used. These should be capable ofchanging from full light to none in approximately 30 min.SecurityBiosafety issues resulting from escape have been covered adequately in the Arthropod ContainmentGuidelines (Benedict 2003) and will not be addressed here.Physical securityAppropriate means should be in place to prevent casual interference by untrained persons with mosquitoculture. The variety will vary from electronic keypads, locked doors, to no deliberate means at all. Simplelocation of the insectary in the basement or back of a building may be sufficient.Environmental securityEnvironmental alarm systems should be in place to protect valuable stocks and experimental materials.One should ask, “How much would my program suffer if the temperature in this room (or incubator) wereexcessively cold or hot resulting in the death of all the mosquitoes it holds?” This will place a value on analarm system. The MR4 does not have the most reliable insectary environments. Therefore, we havemade it a priority to install a monitoring and alarm system that notifies insectary staff in real time ifconditions are not suitable. We have found it is not sufficient for facilities maintenance staff to receive

Chapter 1 : Insectary Operation1.1 Equipping and Operating an InsectaryPage 3 of 4such alarms. This system has saved stocks numerous times and has used pagers, cell phones, andBlackberries for notification.Comfort of personnelThis should be strongly considered during the design of the insectary. Few enjoy working in a hot humidinsectary, and this can be reduced by subdividing adult and larval holding areas, using lower humidity orrelying on incubators.FurnitureRust-proof metal, fiberglass or plastic furnishings are preferable. Shelving should be easily adjustable andstand-alone units should be equipped with wheels. As is discussed in the chapter on cleanliness andgeneral maintenance, one’s ability to clean equipment – and beneath it – is essential. All furnishings mustbe suitable for being wet frequently.Supplies and culture equipmentFollowing is a list of typical supplies needed to equip a small insectary. The sources for much of thismaterial will be different depending on your location and many substitutions are possible. Though somesources are local, the URLs will provide information to give you an idea of what is described. We indicateFisher Scientific as a source for many items, but these products are widely available. Many practices andinnovative devices are found around the world, so collect good ideas and device solutions in all theinsectaries that you visit.Mosquito Rearing Equipment/Supplies8 oz. white plastic containers (for pupae)-approx 250ml, used in food serviceSource exampleModelexampleURLUS Plastics 81134 www.usplastic.comBugDorm1 adult cage (option 1)MegaViewScienceDP1000www.megaview.com.twwww.bioquip.com12x12x12 Metal cage ( option 2) BioQuip 1450B www.bioquip.comLarval rearing trays BioQuip 1426B www.bioquip.comPlexiglas covers for traysFabricate locally2 ml amber latex pipette bulbs FisherScientific S32324 www.fishersci.comPlastic disposable pipettes (trim end,attach bulb and use as a pupae picker)Stainless steel mesh strainer (to filterlarvae and pupae)FisherScientific 13-711-7 www.fishersci.comLocal source SDM-05 www.atlantafixture.comTubes for mixing yeast e.g. 15 mldisposableFisherScientific05-538-51 www.fishersci.com10 ml pipettes FisherScientific13-678-12Ewww.fishersci.comSucrose (to make 10% sugar solution foradults)Local sourcewww.fishersci.comLarge cotton balls FisherScientific 07-886 www.fishersci.comcolored tape (to label and discriminatestocks-choose 1 color per stock)FisherScientific15-901-15(colorcode)www.fishersci.com

Chapter 1 : Insectary Operation1.1 Equipping and Operating an InsectaryPage 4 of 4Larval diet e.g. Drs. Foster and Smith KoiStaple DietDrs. Foster andSmithKoiStapleDietwww.drsfostersmith.com‘dash, pinch, smidgen’ stainless steelmeasuring spoonsLocal sourceMouth aspirator 1 John Hock Co. 412 www.johnwhockco.comFeather-tip forceps Bioquip 4748 www.bioquip.com2 liter clear plastic pitchers with volumemarkingsLocal sourceFilter paper sheetsFisherScientific09-803-5Ewww.fishersci.comQorpak tubes or similar Qorpak 3891P www.qorpak.com500 ml wash bottles FisherScientific02-897-11 www.fishersci.comWaterproof felt tip markers e.g. ‘Sharpie’ Local source 13-379-1 http://new.fishersci.comTable 1.1.1. Some useful insectary supplies and manufacturers.Common Entomological supply sources:• John Hock Company www.johnwhockco.com• Watkins and Doncaster www.watdon.com• BioQuip www.bioquip.com• Educational Science Co. www.educationalscience.com• MegaView www.megaview.com.twReferencesBenedict MQ (1997) Care and maintenance of anopheline mosquito colonies. In: Crampton JM, BeardCB, Louis C (eds) The Molecular Biology of Insect Disease Vectors. Chapman & Hall, New York, pp 2-12Benedict MQ (2003) Arthropod Containment Guidelines. Vector Borne and Zoonotic Diseases 3:63-98Dame DA, Haile DG, Lofgren CS, Bailey DL, Munroe WL (1978) Improved Rearing Techniques for LarvalAnopheles albimanus : Use of Dried Mosquito Eggs and Electric Heating Tapes. Mosq. News 38:68-74Gerberg EJ, Barnard DR, Ward RA (1994) Manual for mosquito rearing and experimental techniques,revised edn. American Mosquito Control Association, Inc., Lake Charles1 An aspirator can be constructed from a section of rigid transparent plastic tubing 20 cm long with aninside diameter of about 15 mm. One end of the tube is covered with fine cloth netting or metal gauze andthen inserted into a piece of rubber hose/tubing 50-60 cm)

Chapter 1 : Insectary Operation1.2 Cleanliness and General MaintenancePage 1 of 81.2 Cleanliness and General MaintenanceMR4 StaffIntroductionCleaning and general maintenance of insectaries can easily fall to the bottom of the list of things to do.However, daily light cleaning and routine deep cleaning help prevent serious problems such as infectionand predation. Following are some reasons a clean environment has a major impact on mosquito cultureand research results.Promotion of cleanliness and sterilityReduction of infections and pestsMost insectary infections are fungal, protozoan, or bacterial and are routinely transmitted via water or air(see Chapter 2.2). While it is not practical to completely eliminate these pathogens from the environment,it is possible to reduce their prevalence. A primary infection may not be lethal or significantly debilitate themosquitoes, but it may produce conditions that are favorable to the development of secondary infectionsthat are lethal. Often fungal infections may be chronic in nature, diminishing the immune status of larvaeresulting in a secondary, lethal bacterial infection. Microbial control can also lead to a reduction inbiogenic toxins.Minimizing insect pests in the insectary can also be crucial to maintaining healthy stocks. Insect pests,such as predatory roaches and ants, are of greatest concern in an insectary as they can easily consumea colony of adult mosquitoes overnight. Larger pests such as rodents introduce waste products thatharbor pathogens in the rearing environment. The easiest way to minimize pests in the insectary is toreduce or eliminate the conditions that attract them: food, accessible water, and harborages (shelter).Clean conditions alone are usually insufficient to prevent all pest problems. In this case, baits and trapscan be used, but be sure they do not contain insecticides to which your mosquitoes will be exposed (seebelow).Achieving sanitary conditionsAt a given period of log phase growth of a microbial population, the titer of organisms is proportional tothe titer at the beginning. Therefore, minimizing microbial growth by any means can significantly reducethe capacity for growth.Insectaries often use equipment and solutions that cannot be autoclaved or otherwise fully sterilized, norare facilities for gas or irradiation sterilization practical. Nonetheless, measures must be taken thatprovide a reduction in microbial contamination. Heat killing on surfaces and rearing equipment can bedone by boiling, autoclaving, or baking. Exposing fluids, tools and containers to even a sub-sterilizinglevel of heat can allow fewer microbes into your environment. Autoclaving is most effective, but liquidsthat contain components destroyed by autoclaving can be partially decontaminated by an elevated heatprocess such as pasteurization or filtration. Many of these treatments and practices are similar to thosepracticed in restaurants: sanitary, but not sterile conditions are the goal.Worker health and moraleA clean, pleasant-smelling, uncluttered insectary is healthier and more desirable to work in for longperiods of time, and an uncomfortable and smelly insectary is one reason people are not eager to remain.Moreover, an abundance of molds and dust are likely to irritate asthmatics and those with allergies.Techniques for Achieving Clean and Sanitary ConditionsInsectary workers must recognize that sanitation – in addition to sterilization – is an effective way topromote consistent mosquito health. We have listed many options for achieving sanitary conditions, and

Chapter 1 : Insectary Operation1.2 Cleanliness and General MaintenancePage 2 of 8some or all of these can be employed in any laboratory. The consistent, combined use of these isessential.Chemicals including bleach, gases and solventsChlorine bleach (sodium hypochlorite) is commonly used to sterilize plastic containers, countertops, andfloors. Ethanol also has some sterilization effect on bacteria and fungi, but be careful not to exposemosquitoes directly to ethanol as it will kill them instantly. Hydrogen peroxide is another common anduseful chemical that is compatible with many materials for sterilization purposes. Finally, ethylene oxidesterilization can be useful if facilities are available.Cold temperatures including freezing and refrigerationUnless special precautions are taken to protect the organisms, freezing will kill many microbes. Eventhose that will survive cold/freezing to some extent may be reduced in number or their growth-ratediminished. Both larval and adult diets should be stored in a refrigerator or freezer.DesiccationExtremely low humidity, especially in combination with elevated heat, reduces the abundance of manymicrobes. Therefore, plastic rearing containers and other equipment dried and stored in a dry place arelikely to harbor fewer microbes than those dried and stored inside a humid insectary. Drying ovensprovide low humidity and high heat and are useful for sanitizing equipment that cannot withstandautoclaving.DetergentsHand-washing with soap is more effective than just using water since detergents break down cellmembranes and kill microbes in the process. Similarly, detergents will kill microbes and loosen microbialfood sources such as grease and dirt in the insectary better thanwater alone. While excessive detergent residues might also killmosquitoes, surfaces that are cleaned with detergents andrinsed thoroughly will harbor fewer microbes. If you are in doubtabout the toxicity of a detergent, perform a simple doseresponse mortality test with L1s using realistic concentrationsthat might exist as when containers are not completely rinsed.FiltrationUltra-filtration will remove fungi and bacteria from solutions.However, this method is usually only useful for small volumes ofsolutions due to the cost.HeatHeat-treatment via autoclaving is standard for total sterilization.Therefore equipment should be selected with this in mind. Asmentioned above, drying ovens reduce microbes and may becompatible with equipment that cannot be autoclaved. Briefimmersion in hot water is a measure that provides some benefit,and it can be made available in even the most basic insectaries.Irradiation: gamma, X-ray, UV, photonsMany types of irradiation educe the abundance of microbes. AtFigure 1.2.1. Any household water-heatingpot designed for kitchens can work well inthe insectary for sanitizing equipment.Immersing equipment between usesaddresses two problems simultaneously: Itwill clean instruments to protect thetransfer of infection and will also kill anylarvae/pupae that were accidentally leftbehind on the tool.first glance, such methods as listed might not appear to be appropriate for an insectary. However, theymight be used in the ventilation system (UV) or for sanitizing rearing containers. UV rays from sunshinewill even kill some microbes.

Chapter 1 : Insectary Operation1.2 Cleanliness and General MaintenancePage 3 of 8StarvationFew microbes can survive indefinitely without minerals or complex organic compounds. Cleanliness in theinsectary generally reduces such sources.Specific procedures to enhance sanitationAir filtersCentral air-conditioning air filters are effective only if they are changed regularly (Figures 1.2.2-1.2.4).The demand and their performance depend on the cleanliness of the air entering the filter in the firstplace, so routine floor cleaning and dusting have a double benefit.Recirculating filters utilizing activated charcoal, particulate meshes, and HEPA are relatively inexpensiveand readily available (Fig 1.2.4). Consider installation of these in addition to the filtration provided by theair-conditioning system to reduce the number of free-floating particles in the insectary. As with air filters,these filters are only effective if they are changed regularly.Figure 1.2.2. Air filters are great formosquitoes and people if they are changedregularly. Air distribution systems can normallybe adapted easily to include filtration.Figure 1.2.3. Dirty air filters are uselessor even harmful. Filters should bechecked and changed regularly.Figure 1.2.4. (Left)Stand-alone HEPA air filtration units arereadily available and useful, especiallyin confined spaces.

Chapter 1 : Insectary Operation1.2 Cleanliness and General MaintenancePage 4 of 8Humidifier selection and maintenanceMany humidifiers contain a water reservoir that never empties completely. This means that even thoughdeionized or even sterile water may enter the humidifier, airborne particles that fall into the reservoir willintroduce sufficient material to establish microbial growth. These microbes will then conveniently ride onthe water droplets into and onto everything they reach. Steam generators are a better choice for insectarydesign. Routine cleaning of any water system should be done e.g. by flushing with bleach or according tothe manufacturer’s recommendations.Larval dietKeeping the larval food frozen will not sterilize it, but it will prevent microbial growth and decay duringstorage. Process only a small amount of food and aliquot it into smaller portions. Store at -20 o C untilneeded. We recommend keeping any unused food in a refrigerator to reduce contamination since thegrowth rate of microbes is temperature-dependent. When using liquid food, keep it in the refrigerator oncemixed and minimize the amount of time it is out at room temperature. If you pre-mix larval slurry, makeonly as much as you can use in 2-3 days to prevent microbial growth in situ. Refrigerate the foodovernight and discard it if it's left out regardless of how 'good' it smells.Replace the food container between batches. If this is not possible, clean the container with a detergentsoap and thoroughly dry in a warm oven. Likewise, soaking a plastic food container overnight in bleach isgood for reducing pathogens. At a minimum, wash with a brush, detergent, and hot water. Rinsethoroughly in clean water and dry.Never combine batches of old and newly prepared food. Mixing preparations could inadvertentlydisseminate microbes that were growing in the older food to the fresh batch.Adult sugar waterMany laboratories place sugar-water-soaked cotton pads on cages. These require replacement atintervals, in part due to microbial growth. When working with sugar water, keep your hands clean. This isespecially necessary when replacing old with new sugar. For example, if changing cotton sugar pads,think: "Did I just pick up a moldy cotton ball and stick my fingers in the fresh sugar water to get another?"In this example, a solution is to use one hand to remove the old cotton balls, the other for the fresh ones.Wearing a glove on the ‘clean’ hand is a good reminder.Another important measure of mold prevention is to make sure that the feeder you are using is sanitized.Cotton balls can be autoclaved and stored in sealed containers. An open bag of cotton in a humidinsectary is a great settling ground for mold spores, so keep them sealed until use. Feeders of differentsorts, vials or screen covers, can be soaked in bleach and dried prior to re-use. They should also bestored in a closed container prior to use. NOTE: Bleach oxidizes steel very quickly. If you plan to usebleach for sanitizing, choose metals such as aluminum or stainless steel.Finally, autoclave sugar water. Once the container is opened, it begins accumulating microbes. A cup ofsugar water stored in the refrigerator and reused for weeks becomes increasingly contaminated. If youhave a cup of cotton balls in sugar water, discard it weekly and start each week with a clean container,new sugar water, and new cotton balls. Also, you can use a preservative such as methylparaben in thesugar water at low concentrations to reduce microbial growth. See culture section, Chapter 2.4.7, forideas on sugar feeders that lessen mold problems.Mosquito containersIn order to prevent mold growth on mosquito containers, discard dead mosquitoes from used containersas soon as possible. Dead mosquitoes in containers can shed potential primary and secondarypathogens. Even if you autoclave materials, the microbes may have produced toxins before autoclavingor cleaning that will cause problems. For this reason, try to remove dead mosquitoes from active rearingcontainers as much as possible. If a pathogen killed a mosquito that is dead in a rearing pan, when thedead carcass decays it will probably release more pathogens into the water.

Chapter 1 : Insectary Operation1.2 Cleanliness and General MaintenancePage 6 of 8Figure 1.2.7. The mildew and mold growth on the walls and the floor of this insectary canstress mosquitoes causing them to be more susceptible to infections and less able to respondto changing conditions.FloorsWipe up spills and eliminate leaks to keep floors as dry and unfriendly as possible. For these purposes, itmay be helpful to equip the insectary with a wet-dry vacuum cleaner, making sure that the vacuumcleaner is not used elsewhere to vacuum toxins such as under furniture where insecticides have beensprayed. For all of the reasons above, and especially in relationship to desiccation, don't let wateraccumulate on floors, in containers or on counters.Even though detergent may not be necessary to make the floor appear clean, it does have an antimicrobialeffect and should be used for routine mopping.Shelves and countersRemove unused equipment and supplies. Unused materials in the insectary make it more difficult to cleanaround and beneath. Items stored in the insectary will likely begin to accumulate molds: Cardboard isespecially poor in moist environments as it holds water, molds, and provides harborages for cockroachesand other arthropods.Keep shelves uncluttered, dusted, and free of spills, especially sugar water and food sources. Removal ofdust is also important as it is highly organic. It carries mold and bacterial spores and therefore circulationof dust by air in the lab spreads potential sources of infection. When you wipe up a spill, you are not onlyremoving the spill. You are removing the spores of the organisms that grow in the spill, those carried bythe pests attracted to the spill, etc.Use as many sealable storage containers as possible. Tupperware-types are good and withstandbleaching; however, you can't autoclave them. Avoid cardboard, paper and wood. Use instead, plastic,metal, or glass which are easier to sterilize with bleach or heat.Keep items sealed until use. Open one bag of cotton balls or one box of cups at a time. Keep coveredexcept when in use. Consider putting everything into covered containers such as plastic closet boxes orshoe-boxes.

Chapter 1 : Insectary Operation1.2 Cleanliness and General MaintenancePage 7 of 8Figure 1.2.8. (Left) This is an example of wellcleaned working insectary with appropriate storage.Items prone to mold spores from the environment (inthis case sugar-soaked cotton balls) are placed inTupperware containers to shield them. Minimumamounts of daily use items are stored on the shelves.Storage items are placed outside the insectaryenvironment. Using rolling carts with brakes in theinsectary can be a great way to make cleaning easyand efficient. Also, in case of emergency, it is easy tomove items in or out of the insectary.Figure 1.2.9. (Right) Clutter in theinsectary renders the cluttered items dirtyas well as the environment harmful formosquitoes. Any cardboard surfaces suchas the one on the bottom of this shelf canand will hold mold and fungal growth.Pest controlSentinels for cleanlinessInfestations of many insect pests occur due to inadequate cleanliness in the insectary. Periodic cleaningand routine trapping of insects will eliminate food sources thereby reducing their numbers. Cockroaches(Figures 1.2.11 and 1.2.12) typically occur in areas where excess food has been spilled and there arenumerous harborage sites. They are known to passively carry several different pathogenic bacteria ontheir carapaces that they spread while they move about. Cockroaches will also catch and consume livingmosquitoes. Book lice, also known as paper lice, are cosmopolitan insects that live in dark, humidconditions (Figure 1.2.13). They are typically associated with excess food spillage or starchy papergoods on which they feed. Unlike cockroaches, book lice are not known to cause harm to mosquitoes,however their appearance may equate to unclean conditions in the insectary. Once established, they aredifficult to eliminate however turning off the humidity and raising the temperature during a temporary shutdownin the insectary may eliminate large numbers of them. Excess diet should also be disposed of,autoclaved, or kept at 4ºC to kill off lice.

Chapter 1 : Insectary Operation1.2 Cleanliness and General MaintenancePage 8 of 8Figure 1.2.11. Blatellagermanica (German cockroach)Figure 1.2.12. Periplanetaamericana (American cockroach)Figure 1.2.13. Liposcelis corredens(Book louse)Regular preventative trappingEnsure that the insectary is monitored for the presence of rodents, ants, and cockroaches. Antsparticularly can destroy a cage of mosquitoes overnight. Furthermore, both ants and cockroaches canspread microbes and leave feces in the insectary. The MR4 has used both Maxforce ant granules andMaxforce roach killer bait gel without evidence of harm to the colonies. Routine distribution of outdoor antbaits around the perimeter of the insectary building may be a useful preventative measure.Figure 1.2.10. Anticipating the introduction of pests before they areseen can save an insectary from an overnight, unforeseen invasion.Ant baits such as the one shown here are important to place arounddoorways and other entry points. Otherwise, an ant colony canmove in and decimate mosquito stocks overnight.Reduce food sourcesSpilled sugar water and food is difficult to control. One method of prevention is to dispense them onlyover a counter top that gets cleaned daily. Otherwise, make sure any spills are cleaned up as quickly aspossible. Larval food is especially protein and fat-rich so ants and roaches thrive on it. Dead mosquitoescan also be food sources for ants or roaches so clean old cages as soon as possible. Dirty rearing pansare a food rich source for cockroaches so the cleaning as soon as possible applies to the pans as well.Trash cans are also well known food sources for pests. If you dispose of old sugar soaked cotton balls inthe trash, for example, make sure the trash is removed from the insectary daily.Ultimately, the cleaner your insectary, the healthier your mosquitoes will be. Attention to sanitationmethods makes a huge difference in mosquito health management.

Chapter 1 : Insectary Operation1.3 Scheduling and Regulating Your Work LoadPage 1 of 41.3 Scheduling and Regulating Your Work LoadMR4 StaffDevelop and maintain a scheduleRearing multiple stocks and strains of mosquitoes or using large numbers of mosquitoes for experimentsand stock maintenance can be very difficult without thought for scheduling and planning. The first rule isnever to endanger the colony by using too much material for experiments. Once a strain is lost, it is lostforever. You should ensure that your colony is sufficiently large to support current experimental work andthe colony’s future generations. If colonies are reared in a haphazard manner, it is difficult to know whenor if you will have new material available for experiments. However, if the insectary is operated in acontrolled and consistent manner, it will be easy to produce enough material without risking a colony, andfollowing strict standards and schedules makes it effortless to say with assurance when you will havematerial at the needed stage. Some suggestions toward achieving this are outlined below.Decide on discrete or overlapping generationsThere are two general approaches for stock maintenance, each of which has particular advantages:discrete and overlapping generations. The discrete approach produces sufficient material for the nextgeneration which is placed in a fresh cage - there is no mixing between generations. The overlappingapproach produces material which is placed in a cage with adults of the previous generation. so progenyfrom the cage could result from either generation. The MR4 almost exclusively uses discrete generations.Each generation of adults is bloodfed the first and only time for stock and experimental use if there aresufficient numbers of progeny. A second blood-feeding is performed only to produce experimentalmaterial and/or a backup if needed. It is most efficient to label all trays indicating whether they are theprimary stock, experimental material or a backup.If contamination is detected in stocks cultured by the discrete method, previous generations provide abackup generation that may provide pure material if contamination occurs. On the other hand, stocks thatare difficult to bloodfeed or produce few progeny may be best maintained by pooling all the availablematerial in a cage(s) and culturing by overlapping generations.We are aware of no studies of the differences in genetic changes or selection that might occur in eithermode. However, it seems intuitive that maintaining stocks by the discrete method would select individualsthat reproduce early with little effect of greater longevity.Establish a single schedule of activitiesInsectaries are more efficient if there are fixed days for specific tasks such as egging and blood feeding. Ifexperiments require material reared on a different schedule, the individual researcher should beresponsible for keeping their experimental materials separate from the general flow of the insectaryschedule. Having a strict schedule also makes it easier to share chores between technicians as dutiescan be assigned routinely for certain days.Keep the environmental conditions fixed in the insectaryTo ensure predictable development of mosquitoes in the insectary, temperature, and to a lesser extenthumidity, must be controlled. Uncontrolled fluctuations in temperature or humidity will cause colonies todevelop faster or slower, affect fecundity and can cause mortality in extreme cases.Follow culturing density standardsSimilarly, if colonies go underfed or are grown in a more crowded/less crowded density than normal; yourmosquitoes will more than likely not be at the stage you had anticipated for your schedule. There areseveral simple methods for quantifying larvae and eggs though many people can estimate closely enough

Chapter 1 : Insectary Operation1.3 Scheduling and Regulating Your Work LoadPage 2 of 4by eye with experience. Because not all stocks have the same hatching rates, quantitative methods foreggs will require adjustment.Feed larvae appropriately and consistentlyAll trays of larvae should be observed carefully daily and fed and/or the density adjusted because thesepractices affect the success of colony maintenance more than any others. There are several indicators todetermine whether you are feeding too much or too little in Chapter 2.Suggested Schedule 1: a three-week cycle beginning on a FridayBelow is an example schedule based on a typical strain of An. gambiae reared at constant 80% RH, 27 o Cunder the conditions detailed in the culture section of Chapter 2. You will have to make modifications tothis depending on the specific strains you culture and the availability of labor and blood source. Eachmethod referenced is described at length in Chapter 2.Friday: Blood-feed adult females. The mosquitoes should be a minimum of two days post-emergence forthe best results. In many cases, 4-7 days post-emergence is optimal, but do not wait longer for the firstfeeding as mortality will endanger your primary stock and/or opportunity to re-feed.Saturday: No attention required.Sunday: No attention required.Monday: Insert the egging dish into the cage.Tuesday: Remove the egg dish from the cage. Bleach the eggs and store them in a humid sealed cupovernight.Wednesday: Rinse eggs into pans for hatching and feed.Thursday: No attention required.Friday: Split the larvae into pans based on the number you will need but keeping in mind properdensities. Add yeast to a final concentration of 0.02% w/v and a very small amount of the larval diet youwill use.Saturday: No attention is required.Sunday: Feed the larvae a volume of ground diet based on their size and density. If there are too manylarvae in the pan, thin or split into more trays to ensure no crowding occurs.Monday through Wednesday: Continue splitting/thinning and feeding the pans daily as needed. It isbest if the density at this point is the same as the final density; crowding slows development.Wednesday through Friday: Pupae should be collected daily and transferred to a cup with clean waterand placed into a new cage with a sugar source. If you chose to allow adults to emerge in the tray forlater transfer, cover trays at this point. If you are working with a strain that remains in pupal form for 48hours or more, you may want to collect pupae every other day. However, you will need to feed the larvadaily. Most Anophelines have a higher proportion of male pupae developing on the first day so if you arecollecting only 100 for stock you should check to make sure you have a good number of females beforediscarding any remaining larvae.Friday of the following week: Bloodfeed the adults to initiate the cycle again.If you find that the adults are beginning to die before you blood-feed on Friday, alternate the schedulebetween a generation of bloodfeeding on Monday and then Fridays. This way, every other weekend willbe work-free. This makes a 2 1/2 week schedule; better for mosquitoes but not as convenient formosquito culturists.

Chapter 1 : Insectary Operation1.3 Scheduling and Regulating Your Work LoadPage 3 of 4Suggested Schedule 2: a three-week cycle beginning on MondayThis follows the schedule above, but shifted. This schedule will probably result in pupation over theweekend so it may not be as convenient.Monday: Blood-feed adult females.Tuesday: No attention required.Wednesday: No attention required.Thursday: Collect eggs.Friday: Remove the egg dish and bleach the eggs.Saturday: Hatch larvae.Sunday: No attention required.Monday: Feed and split/thin larvae.Tuesday through Thursday: Thin and feed pans as needed.Friday through Sunday: Collect pupae or adults and feed larvae every day.Monday following week: Blood-feed to reinitiate the cycle.Both schedules are laid out in calendar form in Table 1.3.1.Mon Tue Wed Thu Fri Sat SunWeek 1Schedule 1 blood eggSchedule 2 blood eggdish bleach hatchMon Tue Wed Thu Fri Sat SunWeek 2Schedule 1 bleach hatch split/thin split/thinSchedule 2 split/thin split/thin feed feed feed feedMon Tue Wed Thu Fri Sat SunWeek 3Schedule 1 feed feed Feed pupation pupation pupation pupationSchedule 2 pupae pupae Pupae pupaeMon Tue Wed Thu Fri Sat SunWeek 4Schedule 1bloodSchedule 2 blood eggdish bleach hatchTable 1.3.1. Calendar layout of two schedules as described above.

Chapter 1 : Insectary Operation1.3 Scheduling and Regulating Your Work LoadPage 4 of 4Planning experiments: Working backward from the deadlineWhether you are coordinating materials for feeding or simply determining if you can complete anexperiment before a holiday, it is helpful to plan beginning with the deadline date and work backward tothe present using a schedule such as the one presented here. Failure to plan ahead could result in theexperimental material you reared for three weeks being ready on a weekend when you are not at work.You will need to modify the schedule to the actual time periods you experience with your colonies in yourlaboratories. An example of how to plan is given in Table 1.3.2.Sunday Monday Tuesday Wednesday Thursday Friday Saturday1 2 3 4 5 6 7blood8 9 10 11 12 13 14egg dish bleach egg hatch egg feed15 16 17 18 19 20 21check feed check feed feed feed feed22 23 24 25 26 27 28pupae 1 day old 2 days old 3 days old 4 days oldTable 1.3.2. In this example, the researcher needs 4-day old mosquitoes for an infection experiment onThursday the 26 th (yellow highlight). By working backwards on a calendar, one can see that bloodfeedingmust occur on Saturday the 7 th . For convenience, they may wish to bloodfeed on Friday and collect eggson Tuesday.

Chapter 1 : Insectary Operation1.4 Maintaining Stock PurityPage 1 of 21.4 Maintaining Stock PurityMR4 StaffIntroductionAny lab that cultures more than one stock must prevent contamination. Stock identity is determinedultimately by genetic composition; therefore, stocks that are contaminated are of little value, especially iftheir only known distinguishing characteristic was origin location. Physical isolation in different rooms isoften used to prevent contamination but this has limits as the number of stocks increases. Therefore,keeping stocks pure ultimately depends on conscientious methodical attention to detail when makinglabels, transferring pupae and adults, putting egg dishes into cages, etc. Moreover, if your strains are notphenotypically defined, it may be impossible to determine that they are contaminated later.Diligent exercise of precautionary methodology is the only way you will prevent contamination. This canbe augmented by using phenotypically marked stocks when possible. Recessive markers are the bestchoice since contamination is more readily detected. The best advice is to stay conscious, careful andfollow routines designed to avoid contamination.Ways to avoid contamination:There is no substitute to consistent attention to detail, but the following are some ways stocks canbecome contaminated with suggestions for avoiding them.Use carefully decontaminated materialsCause: Pupae and larvae easily get stuck in devices and are very difficult to see at a glance. Whenswitching to another stock, it is easy to not notice the contaminant and transfer from strain to strain(Figures 1.4.1 and 1.4.2).To prevent: visually examine tools and rinse in hot water between handling each stock. If you keep only acouple of stocks, separate, clearly marked tools should be kept for each. Use white and transparentcontainers when possible and white countertops.Figure 1.4.1. Hand held pupa pickers with asingle pupae stuck in the apparatus, shownby arrows.Figure 1.4.2. Larval strainer with a singlelarva stuck in the apparatus, shown witharrow.Cause: Eggs in water can easily spill or splash onto the lid of a pan or cup (Figure 1.4.3). Reusing thesame lid or cup for another stock without decontamination can lead to egg transfer.To prevent: use fresh lids and cups that are decontaminated by desiccation, washing, and/or autoclaving,and consistently return the same lid to each container.

Chapter 1 : Insectary Operation1.4 Maintaining Stock PurityPage 2 of 2Figure 1.4.3. This is a common cause of contamination. Theeggs have splashed onto the lid. Accidental mixing of lids at thispoint can cause transfer of eggs that could go easily unnoticed.Keep all lids exclusive to the cup or pan you are working withassuming contamination has occurred.Cause: Mosquitoes are put into thewrong container e.g. pupae intoadult cages.To prevent: consistently use adifferent color of tape/marker colorfor each stock (Figure 1.4.4). Forsmall numbers of stocks, this allowscolor-coding pans and cageswithout writing labels. Usingdifferent colors makes it difficult tonot notice mixing of strains. Givestocks distinct names.Cause: Free flying adult mosquitoesare a contamination concern. Forexample, a mosquito can be flyingby or biting your hand when you areplacing something inside amosquito cage or blowing inmosquitoes. A gravid female can layeggs in any pans that are leftuncovered. Even covered pans can sometimes have enough of a gap for a mosquito to slip inside and layeggs, thereby contaminating the entire cohort.To prevent: routinely trap free mosquitoes in light traps and make every effort to prevent escapes (Figure1.4.5). Inspect trays daily for pupae. If adults are allowed to emerge from the culture tray before transferto adult cages, covers must be securely fastened.Figure 1.4.4. Different colors of tapeand/or different colors of markers make itobvious to see the difference betweenstocks at a glance. They also make itsimple to locate material rapidly.Figure 1.4.5. Light traps, such as the one picturedhere, are good for trapping any loose adultmosquitoes. Flying mosquitoes are a serious sourceof contamination in an insectary and a risk forescaping into the outside environment.

Chapter 1 : Insectary Operation1.5 Insectary Manager ResponsibilitiesPage 1 of 61.5 Insectary Manager ResponsibilitiesMark Benedict“To provide authenticated, high-quality mosquito reagents, training and information to the researchcommunity of today and the future, in a timely and professional manner.”IntroductionWe begin this section with the mission statement for the MR4 vector activities. In order to accomplish this,the following list of responsibilities was developed as guidelines for the MR4 insectary manager. Whilethe details are specific for the MR4 vector activities at the CDC in Atlanta, it provides a useful guide forsupervisors employing a manager to oversee daily operations and in the development of a jobdescription. With little modification, this has served us well to describe the core activities of the manager.Insectary FacilitiesEnvironmentThe Vector Repository Manager (VRM) shall ensure that...1. Environmental conditions in insectaries are constantly maintained at 80 o F (±1.5 o F). Relative humiditywill be controlled to be in the range of 80% (± 10%) 365 days a year without interruption. Lighting iscontrolled such that a 30 minute sunrise and sunset occur, in between which times, the fluorescentlights will be on continually. The total darkness between the end of sunset and the beginning ofsunrise is 12 hours.2. The environmental conditions, except for lighting, are continually monitored by CDC maintenance staffand changes should be made to settings to achieve the above only after consultation and approval bythe VRM.3. Environmental conditions including lighting are continually and independently monitored by the MR4staff. This is achieved by sensors that are located in all three insectaries and capable of notifying MR4staff of conditions that are outside of the permissible range within 10 minutes regardless of whetherstaff are in the MR4 facility, at home, or traveling as necessary to ensure that at least one staff isaware of the problem.4. Pest insect control is continually performed to ensure essential absence primarily of ants andcockroaches. This is achieved in a way that no harm occurs to the insect colonies either directly or bycontamination with toxicants transported by pests. In the event that other pests are observed (e.g.mice), control is enacted as needed, but again with highest regard for the health of the repositoryinsects. Modifications of the facility are considered that physically reduce entry points, breeding sites,and harborages.5. Insect pest control around the perimeter of the building to reduce external sources is considered andexercised if needed.6. Neither CDC personnel, nor local municipalities conduct insect control in the vicinity of the insectaryfacilities.7. Properly operating mosquito traps or other killing devices operate continuously and are monitored forcatches in all insectaries. These should be capable of trapping primarily Anopheles, but also Aedesand Culex species.8. Cleanliness is maintained in all insectaries and support areas. While hospital cleanliness is neitherattainable nor necessary, a consistent effort should be made to improve the level of cleanliness. Inpart this will require labor, but use of materials and furnishings that do not rust and are easily cleanedwill be helpful. Only cleaning compounds that are non-toxic to the mosquitoes are used, but theseshould be used to reduce cleaning maintenance when possible. Moreover, the CDC maintenance

Chapter 1 : Insectary Operation1.5 Insectary Manager ResponsibilitiesPage 2 of 6staff is instructed to maintain the cleanliness of the floor and other areas within their responsibility. TheVRM is responsible for cleanliness but is not to become the custodian.9. Air filtration is installed and maintained properly to reduce the level of odors, fungi, dust, hair etc.Installing additional equipment or modifying existing equipment is considered to improve the airquality. Mold growing on mosquitoes and the insectary walls can be reduced by consistent attention toeliminating spores. Centralized UV sterilization of the air may be feasible.10. Ensure that documentation of the maintenance of the emergency generator is available and beingmaintained. Notify the PI in the event of any planned power outage.Infrastructure ImprovementsThe VRM shall ensure that...1. Sign-holders are installed that contain information about specific courses of action to take in the eventof various environmental anomalies. These will be located either near the alarms and/or by eachdoorway.2. Signage is current and attractively maintained.3. All infrastructure and environmental changes are consistent with the MR4 objectives. Furthermore,these changes are approved by all insectary users.Infrastructure MaintenanceThe VRM shall ensure that...1. Hallways are kept clear of trash, boxes, unused carts, old equipment etc.2. All lights function. The maintenance personnel should be notified in the event of lights burning out andother electrical problems.3. Timers are properly set and maintained.4. Hallway and insectary walls are kept clean and free of un-necessary notes, tape, scuffs, holes, tacksetc.Insectary SuppliesThe VRM shall ensure that...1. Consumables required for the operation of the insectary are maintained at sufficient levels thatshortages do not occur. The supply should be supplemented long before the need becomes critical.Allowance should be made for shipping delays and incorrect or incomplete orders.2. Establish minimum levels of supplies at which orders will be placed.3. Maintain inventory information sufficiently to ensure above.4. Consumables are safe, and have no characteristics that are an immediate threat to the mosquitostocks.5. Alternative consumables are considered for use. Materials that save time and/or money are soughtand tested.6. Maintain the cleanliness and order of the storage areas.7. Mosquito food and blood sources are safe and of an adequate amount to ensure that shortages do notoccur.

Chapter 1 : Insectary Operation1.5 Insectary Manager ResponsibilitiesPage 3 of 6OtherThe VRM shall insure that...1. Office supplies necessary for the timely shipment and documentation of MR4 reagents is ensured.2. Materials to produce documentation for MR4 reagents are of high quality and of adequate amounts.3. Shipping materials are of good supply, quality, and suitability.4. Computer consumables such as CD/Rs, diskettes, paper etc. is of an adequate supply to producedocumentation, file archives, communication etc.Mosquito AuthenticationThe VRM shall ensure that...1. Only authenticated materials are shipped from and maintained by the MR4.2. Authentication methods are developed that are reproducible with reasonable ease both within therepository and by requesters.3. Materials required for authentication are protected from accidental contamination or loss and can beproduced on demand using independent means.4. Documentation is sufficient to enable requesters to authenticate materials independently.Preservation and ProductionThe VRM shall ensure that...1. Levels of all MR4 stocks are sufficient to ensure a constant supply of material for all MR4 activities.2. Non-MR4 personnel who maintain MR4 stocks are informed about the requirements for theenvironment in the insectary and procedures to follow to ensure that the stocks are maintained withoutcontamination or loss. This must be done without imposing upon them or requiring significantalteration of the existing procedures.3. No MR4 stocks become contaminated or lost. This is very important.4. Sufficient duplication of stocks is implemented to ensure an independent supply that providesinsurance against accidental loss. This may be in the form of on-site maintenance in separatefacilities, or a backup stock in another laboratory from whom the material could be obtained ifnecessary that would notify the VRM in the event of loss. Records of recipients of stocks should bereferred to as a final source of stocks.5. DNAs of stocks are prepared as proposed and distributed to the ATCC and additional backup stocksare maintained at the CDC.6. Sentinel adults are monitored for unusually reduced life span.7. The PI is notified promptly by voice and e-mail in the event of any stock contamination, reduction insupply, or unusual culture conditions.8. Improvements to culture methods are considered if these can save time and/or money.9. A current log is available on the web describing the condition of the stocks at all times including allauthentication.DistributionThe VRM shall ensure that...1. Shipments of mosquitoes are made at first availability of the requested material.2. Contents of shipments are correct, contain appropriate documentation, and are properly packaged.

Chapter 1 : Insectary Operation1.5 Insectary Manager ResponsibilitiesPage 4 of 63. Packaging is of a consistently high quality, is labeled with computer-imprinted labels, andenvironmental conditions of containers are suitable to ensure viability of the product.4. Improved incubation and storage methods are investigated to both prolong the life of laboratorymaterial and longevity in transit.5. The recipient is notified of the anticipated shipment date, actual shipment, and tracking information.This may be done by e-mail, phone, mail, or FAX. A record should be kept for all stages.6. Receipt of a request for materials is promptly acknowledged.7. Shipments are made only to authorized requesters.8. The PI is notified of all intentions to ship mosquitoes before shipment is made.Documentation and RecordsThe VRM shall ensure that...1. Monitoring of all environmental conditions is documented. This means that records of humidity,temperature, and lighting are consistently stored and readily available for the entire 24 hours, 7 daysper week, 365 days of the year.2. Both mosquito culture anomalies and nominal conditions are documented and recorded.3. Records of all requests and shipments are made in a database format. This database should includeat least:a. Date of requestb. Record of confirmationc. Anticipated shipping dated. Actual shipping datee. Carrier and tracking numberf. Record of receipt4. Nominal stock levels and quality should be documented consistently. These records should be publiclyavailable on the web.5. Changes to SOPs should be documented.6. All versions of the handbook should be permanently stored in hard and digital form with date andversion number7. Alterations of the handbook should be coordinated with the requirements of the ATCC.8. An annotated version of the handbook indicating the reasoning behind the changes should beavailable.9. Digital and hardcopy forms of the product information sheets are current and also available on theweb.10. All forms are current.11. All standard operating procedures are detailed sufficiently in hard and digital copy so that a successorknows what to do in every situation. These procedures should be diligently maintained and bound in aclearly divided notebook. Contents should contain SOPs, but also include (for example):a. What to do when the alarms go offb. Nominal environmental parametersc. Where records are stored and how they are backed up

Chapter 1 : Insectary Operation1.5 Insectary Manager ResponsibilitiesPage 5 of 6d. What to do when nobody is here on the weekend and there is water leakinge. What to do when a request for a stock comes and the PI is not available to review the requestf. How to authenticate a DNA sample or mosquito stockg. Who the current contacts are at ATCC with whom to communicate regarding bioinformaticsh. Information required for quarterly and annual reports is consistently recorded and madeavailable to the PI.i. Number of shipmentsj. Most-requested materialsk. Summaries of destinationsl. Summaries of material arriving unusablem. Summaries of replacement requests12. Web information is correct and understandable. This will be accomplished by:a. Coordinating with the ATCC bioinformatics personnelb. Producing all data in database form so that it can easily be sorted, searched, and stored.c. Acquiring new information, photographs, and technologies to make the MR4 web site moreuseful and interesting.d. Informing ATCC of changes needed in catalogues, forms, product information sheets etc. thatare available on the WWW.Budgets and Financial ManagementIn coordination with the Branch Program Specialist, the VRM is expected to ensure that:1. Supplies and equipment budgets for the repository are managed so as to best provide items neededfor the smooth operation of the repository.2. Budgets are not over or under-spent3. Orders are received and billed correctly4. Items are not charged to the VR budget without approval by the VRM or PI.Supervision of PersonnelWhile the ultimate responsibility for the conduct of personnel supervised by the VRM is with the PI, theVRM is expected to:1. Ensure that supervised personnel are aware of their responsibilities2. Be trained to perform all tasks3. Ensure that tasks are performed promptly4. Receive safety and security training

Chapter 1 : Insectary Operation1.5 Insectary Manager ResponsibilitiesPage 6 of 65. Make the PI aware of any problems with managed personnel including:a. Consistently poor technical performanceb. Failure to comply with safety or security requirementsc. Conflicts with other employeesd. Difficulties responding to requests from the VRMe. Time and attendance problems.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 1 of 8Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryAdapted from (Clements 1992)IntroductionBehavior and physiology are important to understand in making decisions in the insectary. The way themosquitoes behave will affect choices of food, blood, egging, insectary supplies, insectary spacedemanded and much more. Additionally, understanding more about the differences between your stockscan be used to give clues of possible contamination along with the morphological and molecularauthentication methods discussed in Chapter 4. These tips can also be practical in understanding whymosquitoes are not thriving or behaving as predicted.EggsCulex, Aedes, and Anopheles eggs are laid in different patterns and observing the patterns on eggcollection can be one way of catching a cross-genus contamination event early. Culex eggs are laid indiscrete rafts of attached eggs by individual females. The eggs are tapered (Figure 2.1.1) and tend todrift to the edges of containers and remain there. Anopheles eggs are unattached and lie in stellatepatterns horizontally on the water surface (Figures 2.1.2 and 2.1.5). Exochorion ‘floats’ on the eggs aid inkeeping them at the surface. Aedes lay their eggs unattached to one another above the water but do nothave floats (Figure 2.1.3 and 2.1.5).Aedes eggs survive drying well (Figure 2.1.4) though the amount of time they can be kept dry prior tohatching varies with the species and conditions of storage. Some strains can be kept dry as much as 6months prior to hatching. Anopheles and Culex eggs do not survive extended drying and should be keptmoist and in a humid atmosphere prior to hatching. The amount of time that can pass before hatchingAnopheles or Culex eggs varies. If an insectary has Aedes and Anopheles or Culex, it is best to alwaysallow the Aedes eggs to dry before hatching to minimize contamination by the more sensitive strains.Figure 2.1.1. Culex eggs less than 24 hours postoviposition. Eggs are cemented together formingan egg-raft that floats on the surface of the water.Figure 2.1.2. Anopheles eggs 30 hours postdeposition. Clear floats are visible on sides of eggs.The non-melanized egg (center) will not hatch.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 2 of 8Larvae hatch from the blunt underside.Figure 2.1.3. Aedes albopictus eggs 48 hours postoviposition on seed germination paper.Figure 2.1.4. Aedes aegypti eggs 2 weeks postoviposition stored under insectary conditions.Figure 2.1.5. Egg cups removed from cages 24 hours after insertion. Aedes eggs (left) were laid on seedgermination paper with only a small amount of water in the bottom to keep the paper wet. Anopheleseggs (right) are laid on the surface of water and will spread across the water surface. In smaller numbers,they accumulate at the edge of the water.When mosquito eggs are laid, they are white. They normally darken and harden within a few hours. Therate at which they change color and harden depends on the strain and temperature. Anopheles eggs thatfail to melanize or sink do not hatch.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 3 of 8Larval FeedingIn the wild, mosquito larvae survive in a large variety of habitats. The food types in these habitats arelargely the same as that in the insectary in that they contain microorganisms, detritus (particulate organicmatter), biofilm, and other organic matter such as dead invertebrates. A major source of nutrients formosquito larvae comes from plant material that has been already degraded by fungi or bacteria.Important in choosing a food is to note the method and location of feeding for the particular strain you areusing. Many Anopheles and Culex use the feeding mode collecting-filtering which is feeding byremoving particles that are suspended in the water column or at the water surface. For Aedes,collecting-gathering is a more common method of feeding which involves first causing materials thathave settled or are attached to surfaces to resuspend and then ingesting them from the resuspensionmixture. Other methods of feeding include scraping (removal and ingestion of the biofilm and protists onthe surface of submerged plants and other surfaces), shredding (biting off small fragments of plants ordead matter), and predation (eating other insects). Much of the differences seen in feeding preferencescan be associated with the differences in mouthparts and head structures (Figures 2.1.6 – 2.1.8). Moredetailed information of the various structures can be found in Clements’ The Biology of Mosquitoes.Figure 2.1.6 Aedes head andmouthparts.Figure 2.1.7 Anopheles head andmouthparts.Figure 2.1.8 Culex head andmouthparts.Even though some Culex and Anopheles share the same method of feeding, the location of the feedingcan be different. Anophelines tend to feed at the air/water interface or on the bottom (Figure 2.1.9) whileCulex and Aedes typically feed throughout the water column (Figure 2.1.10).Figure 2.1.9 Anopheles larvae feed at the watersurface and bottom, but not in the column.Figure 2.1.10 Culex (pictured) and Aedes larvaefeed throughout the water column.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 4 of 8Larval mouthparts are complex and suitable for a form of filter feeding and limited 'chewing.' Themouthparts are well developed but differ among strains. Parts associated with feeding are “teeth” for bothbiting and chewing, curved setae which bring food particles from the water to the mouth, and otherbrushes and combs around the mouth to bring in food. The brush filaments and mandibles are suited tothe type and location of feeding. For example, collector-gatherers are adapted to resuspending settledparticles.Anophelines live at the air/water interface. Typically they are seen lying just below the interface, dorsumup. This is also where they feed as water surfaces are covered with an organic microlayer in the wild. Atthe surface which their head rotated 180 degrees, they beat the mouthpart brushes and create currentswhich bring particles toward the mouth.Collector-filterers such as anophelines have lateral palatal brushes at the mouth that are thought tofunction as paddles rather than as filters as previously thought. However, in the paddling, the movementof the brushes delivers water concentrated with larger particles toward the mouth.The size of the particle that larvae can ingest increases with the size and age of the larva. Factors suchas size and age should be taken into consideration when determining which larval food to use. Also, aslarvae grow, the amount of food they will eat increases by as much as 5 times what they ate in the firstinstar.Growth and DevelopmentIntrinsic EffectsMosquito larvae have four stages. The body size changes continually while the head capsule increases(mainly) only at molts i.e. saltatorially. Thus, the instar is best determined by the head capsule size(Timmermann and Briegel 1993). Performing some measurements on the head capsule of your speciesto determine the range of values that could be observed in any stage is a good idea if working with anexact stage is important for your research project. A series of photographs of stages of larval life mightmake it easier for staging to be apparent by eye until you become familiar with your particular stocks andstrains. Larval stages for An. gambiae are shown in Figure 2.1.11.Generally, males develop faster and are smaller adults than females. Males also typically spend less timein the pupal stage before emerging than females (Haddow et al. 1959; de Meillon et al. 1967). The degreeof sexual size dimorphism varies between stage and species. For example, though the adult size differsquite widely from male to female in Anopheles and Aedes, the pupal size differences are not as apparentin Anopheles as in Aedes (Figures 2.1.12 and 2.1.13).Extrinsic effectsTemperatureTemperature is the most important and easily controlled extrinsic factor affecting growth rates of larvae.The effect of temperature on the growth of mosquito larvae has been studied extensively. Specific foreach species, there is a temperature range in which development can occur. Within this range, growthand development vary dramatically with the temperature fluctuations. For this reason, it is important tocontrol temperature to achieve predictable culture.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 5 of 8Figure 2.1.11 From left to right, Anopheles gambiae larvae 24 hours post hatch (1 st instar or L1), 2 dayspost hatch (2 nd instar or L2), 5 days post hatch (3 rd instar or L3) and 6 days post hatch (4 th instar or L4).All were photographed at the same magnification.Figure 2.1.12. Anopheles gambiae pupae: twofemales (bottom right) and one male (top leftcorner). Size difference is not obvious by eye.Figure 2.1.13. Aedes aegypti pupae: twomales (right) one female (left). Size disparitiesare apparent.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 6 of 8NutritionThe amount of available food significantly affects larval growth. Underfeeding can cause as much delayas overfeeding but will likely be evident later, especially in the adult stage. Much has been written aboutDietary Restriction (DR) in mice and flies and is reviewed in Chapter 3 for its contributions to longevityand fecundity. In short, DR causes the animals to live longer but age faster having negative influence onfecundity and tolerance for environmental fluctuations and infection.Larval DensityAchieving the right density is very important in growth and development. The most common problemsassociated with over crowding are: longer development time, reduced pupation and eclosion, and adecrease in pupal weight. See Chapter 2 Culture section for more on proper density for anophelines.Effect of Larval Health on AdultsAdults from larvae that were crowded are typically smaller and less fecund. The ultimate size of an adultmosquito will be based on genetics in combination with the environmental conditions experienced throughdevelopment. Studies have shown that larvae that are reared in crowded conditions had negative effectson weight at emergence, quantity of the blood meal and overall fertility. Poor larval conditions cannot betotally overcome by good diet or care in later stages, therefore careful attention to larval conditionsdetermines high overall quality of production.Environmental effects on rhythmsStudies show a link between environmental factors and ecdysis. These studies are limited to certainspecies and conditions; however, the evidence supports the shifting of ecdysis under temperaturechanges, light/dark cycles, and larval stress such as salinity. For example, researchers found that incontinual darkness, with variable temperature cycles, the larval-pupal ecdysis was more likely to occurduring the warm phase.The time of day/night the ecdysis will occur is species-dependent. It is thought that the trigger to molt isswitched on and off based on a daily rhythmic activity cycle of 24-hour intervals that is exhibited by manyorganisms, or a circadian rhythm. This is not universal among all mosquitoes. Examples of some foundnot to have such a rhythm are An. quadrimaculatus (Nayar and Sauerman 1970) and Ae. aegypti(Haddow et al. 1959). The best rule of thumb is if your insectary has problems with ecdysis beingtemporally irregular or extended over several days, experiment with your conditions to correct theproblem. Light and temperature are the key factors.Adult feedingPlant JuicesBoth adult male and female mosquitoes will drink plant juices as an energy source (see review in (Foster1995). Plant sugar is the major food resource for mosquitoes. In the wild, the most common source isfloral nectar, but other sources exist such as damaged fruit (Figure 2.1.14) or vegetative tissue. Withthese different meals, the mosquito would be receiving largely sucrose, fructose, or glucose, dependingon the source. Other sources such as maltose or melibiose are seldom found in mosquitoes. Amino acidsneeded for ovary development can be found in some nectar, but the concentrations are not high enoughto replace a blood meal. Natural sugar sources can have a wide range of amounts of sugar from none to50% w/v, though 20-50% w/v is the normal range. Mosquitoes have been seen ingesting crystallizedsucrose by liquefying it with saliva.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 7 of 8Figure 2.1.14. Anophelesgambiae taking sugar mealfrom a tangerine. Female atbottom, center is drinking juice.Two males have beenattracted.Figure 2.1.15. Sugar-fedAnopheles farauti. Plant juicesare stored in the crop ratherthan the midgut. Differentspecies have been observed toingest 29-62% of their unfedweight in sugar. In many tests,certain types of sugar sourceshave been shown to beimportant in increasing the lifespan of females that arefrequently blood fed over thosewho were given only blood.BloodOnly females take blood meals. The blood is their resource for protein needed in ovary development.There is data that substantiates blood as also providing a source of energy since blood fed females havebeen shown to survive longer than females given only water. It is very common for a mosquito to take asmuch as 2-4 times their weight of blood in a single meal. Females can also excrete clear to reddish fluidwhile bloodfeeding in order to concentrate the protein as much as 2 fold (Figure 2.1.16), a process calleddiuresis.Mouthparts Used in FeedingMosquitoes have mouth parts conducive to taking up liquids. Females have more complex mouthpartsbecause they must probe flowers and pierce skin. The mouthparts vary among the types of mosquitoes,especially among those who have different necessary functions such as those that take blood mealsversus those that do not.

Chapter 2 : Anopheles Laboratory Biology and Culture2.1 Behavior and Physiology of Anophelines in the LaboratoryPage 8 of 8Figure 2.1.16. Anophelesstephensi mosquito taking a bloodmeal. Notice the blood movingthrough the proboscis. Themosquito is undergoing a processcalled diuresis by which sheconcentrates the blood.(Photo contributed by JamesGathany, CDC, used withpermission.)Phases of Blood-feeding• Exploration. After a mosquito lands on its host, it goes through an exploratory phase beforepenetrating the skin. Anopheles and Aedes remain stationary for a period of time after landing ona human. Host movement during this time causes the mosquito to leave. Without movement, themosquito explores the skin either in the area of landing or sometimes moving around.• Probing. Prior to penetration, the mosquito goes through a process of probing in which it touchesits labium (not the proboscis) to the skin surface many times to decide where to penetrate withthe proboscis. The mosquito then stabilizes by straightening the legs just prior to penetration.• Penetration. Proboscis is inserted into the skin.• Imbibing. This process begins when the palps stop vibrating from the penetration process.• Withdrawal. This action takes about 3 seconds. A complete withdrawal can occur or only apartial one with re-insertion. After final withdrawal, the female usually flies away quickly.ReferencesClements AN (1992) The Biology of Mosquitoes: Development, Nutrition and Reproduction. Chapman &Hall, Londonde Meillon B, Sebastian A, Khan ZH (1967) The duration of egg, larval and pupal stages of Culex pipiensfatigans in Rangoon, Burma. Bull World Health Organ 36:7-14Foster WA (1995) Mosquito sugar feeding and reproductive energetics. Annual Review of Entomology40:443-474Haddow AJ, Gillett JD, Corbet PS (1959) Laboratory observations on pupation and emergence in themosquito Aedes (Stegomyia) aegypti (Linnaeus). Ann Trop Med Parasitol 53:123-131Nayar JK, Sauerman DM, Jr. (1970) A comparative study of growth and development in Floridamosquitoes. I. Effects of environmental factors on ontogenetic timings, endogenous diurnal rhythm andsynchrony of pupation and emergence. J Med Entomol 7:163-174Timmermann SE, Briegel H (1993) Water depth and larval density affect development and accumulationof reserves in laboratory populations of mosquitoes. Bull. Soc. Vector Ecol. 18:174-187

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 1 of 102.2 Infections in Mosquito CulturesJames J. Becnel and Paul HowellIntroductionFortunately, there are few naturally occurring pathogens that become established in mosquito colonies.There are, however, numerous microbes living within an insectary which, under normal conditions, arenot considered pathogenic (capable of causing disease) but may have deleterious effects when aninsect colony is stressed. Environmental stressors include larval overcrowding, unstable heat or humidity,poor quality diet and overfeeding.Infections in the insectary are spread through one of the following routes: diet, injury, infestation of theegg, or environment. Dietary routes include feeding insects a contaminated diet or the cannibalism ofexpired, infected larvae. Larval injury creates an opening for many water-borne pathogens. Insect eggscan be either internally (viruses) or externally (fungi or microsporidia) infected.Environmental routes are non-specific but can include the following:Introduction of pathogens by wild strains- Wild or newly acquired insects can carry pathogens which donot affect them in the wild, but when introduced in a newly stressed atmosphere, these pathogens canbecome opportunistic. Also, a newly acquired laboratory colony can introduce a chronic infection to aninsectary.Airborne entry- Several fungi, bacteria, and protists can be introduced on airborne particles.People- Some of the more common bacterial contaminants in insect colonies are considered normalhuman fauna like Escherichia coli. These can be passed to the water by touching the water or food whilefeeding.Surfaces- Contaminated or poorly cleaned surfaces and equipment can harbor large numbers ofopportunistic microbes.Knowing what a healthy colony looks and smells like can be the easiest way to detect an infection. Waterthat is malodorous, cloudy, has persistent bubbles or contains excessive foreign matter may bedetrimental to an insect colony (see Chapter 2.4.5 for more on water quality). In the insects, signs andsymptoms can include changes in normal size or color, deformity, lengthened duration of rearing, reducedlongevity of adults, decreased fecundity, and decreased fertility rates. Excess mortality, short life-spans,low reproductive rates, or the presence of fungi on the cuticle are some of the early signs of a possibleinfection in a colony.Infected larvae typically display one of the following: deformation (Figure 2.2.1 and 2.2.2), ordiscoloration of the larvae as well as abnormal larval behavior. In pupae, typical signs of infectionsinclude elongation of early pupae, failure to emerge (Figure 2.2.3), or selective emergence of one sex.

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 2 of 10Figure 2.2.1. Anophelesstephensi larvae that failedto complete metamorphosisdue to infection.Figure 2.2.2. Anopheles gambiaeshowing deformation and signs ofinfection in the larval state.Figure 2.2.3. InfectedAnopheles stephensi thatdied during ecdysis.Commonly reported infections in mosquito coloniesThere are two forms of infections in a colony: chronic and acute. Chronic infections may persist forseveral generations and never be fully apparent but result in poor quality insects, such as infections withfungi and protozoa. Conversely, acute infections quickly spread through a colony and lead to highmortality rates. Pathogens that have been implicated as agents of acute infections include bacteria,viruses, and some fungi. Although the following is not a complete list, it covers the most reportedpathogens and if possible, includes a description of the microbe or the appearance of the infected insect.VirusesThe two types of viruses routinely found to be pathogenic to mosquitoes are defined as occluded or nonoccluded(Federici 1985). Although not routinely found in insectary populations, there have been viralinfections reported in laboratory colonies of An. stephensi. One of these reports showed high mortalityrates in the colony associated with this infection (Bird et al. 1970). At least two other types of viruses havebeen isolated from mosquitoes with only one being reported as detrimental to development of the larvae(Jenkins 1964).Occluded Viruses (Becnel and White 2007)These viruses form proteinaceous crystal occlusions within the mosquito.BaculoviridaeDeltabaculovirus: This Dipteran-specific NPV is the only member of this family that is commonlyassociated with feral mosquito larvae. These viral particles infect the larval midgut epithelium resulting ina stunted appearance, delayed growth and death. Infected larvae often have white cysts or nodulesthroughout the midgut and gastric cecae. These virions are highly virulent and cause mortality within 72-96 hours after the initial infection. Transmission of the virus is enhanced with a high concentration of Mg 2+cations in the larval habitat (Becnel 2007).ReoviridaeCypovirus: These are also referred to as cytoplasmic polyhedrosis viruses (CPV). Infections with CPVsare typically benign in nature but can cause larval mortality when present in high numbers. Infections areeasily seen in L3 or L4 larvae as opaque sections limited to the gastric caeca and/or posterior stomach.Transmission of these viruses is enhanced in the presence of divalent cations or when larvae arestressed.

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 3 of 10Non-Occluded Viruses (Becnel and White 2007)These viruses form paracrystalline arrays of virions in infected cells.IridioviridaeMosquito Iridescent Virus (MIV): These are cosmopolitan in nature in field collected larvae. Infections canbe detected by placing larvae in a dark pan and scanning them with a fluorescent light. Infected larvae willhave iridescent patches of turquoise, green or orange depending on the specific infecting virus. Infectionsare usually localized in the fat body or epidermis. Larvae infected in early instars are highly susceptible.Infections in later instars are typically passive allowing for transovarial transmission. Sub-lethal infectionsoften result in reduced fecundity and longevity in the stock (Becnel 2007).ParvoviridaeMosquito Densovirus (MDV): Infections with the viruses are typically subtle in nature. Infected larvae willbecome lethargic, loose their body color, have a contorted appearance, or appear whitish in color beforeexpiring (Becnel and White 2007) Infections are widespread except in the midgut epithelium. Many ofthese viruses have been isolated from anopheline mosquitoes and from established cell lines.BacteriaAlthough several bacteria are known to be pathogenic to mosquito larvae, relatively few of them occurnaturally in an insectary setting. Infections in the rearing containers are often caused by the inadvertentintroduction of bacteria from the skin of an insectary employee or through the addition of contaminatedwater or food. In Africa, Enterobacteria infections were seen in the haemolymph of insect larvae (Muspratt1966). In advanced stages of disease, black spores were visible inside the larvae and eventually thelarvae displayed a “milky” coloration and swollen appearance (Figure 2.2.4).Figure 2.2.4 Infected Anopheles stephensi. Note the milky swollen appearance indicative ofan infection.Escherichia coli common human fauna. E. coli bacteria have been found to be pathogenic to earlyinstar Culex mosquitoes (Jenkins 1964).Serratia marcescens commonly found growing in standing water. This bacterium has not been reportedto be lethal, but in poor rearing conditions it grows quickly forming a reddish film on the bottom of therearing pans.Pseudomonas fluorescens ubiquitous flagellated bacterium. P. fluorescens is a commonly isolatedbacterium from soil and water sources, and it has been shown to be lethal to mosquito larvae.Pseudomonads are known to cause extensive larval mortality due to their production of toxic substances(Jenkins 1964).

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 4 of 10Leptothrix buccalis common water bacterium isolated in fresh and polluted water sources and found tobe highly lethal to An. maculipennis. With this infection, the larva maintains the disease through eclosion,but death does not occur until sometime after emergence (Jenkins 1964).Streptococcus spp common human fauna. These bacteria can rapidly grow in the warm insectaryconditions and will attach to larvae in large numbers. The bacteria invade the integument (insect's hardouter coat) and cause internal damage leading to mortality in the L3 or L4 stage (Kramer 1964).Treatment of bacterial infectionsAntibiotics such as Penicillin-Streptomycin-Fumigillin (PSF) (Invitrogen 15240-062) can be used in larvalculturing (de St. Jeor and Nielsen 1964). However, it is recommended that before full scaleimplementation, trials are conducted to determine the dosage required to eliminate the infection withoutkilling too many larvae. Additionally, you can feed antibiotics to adults in the sugar meal (Toure et al.2000).In the insectaryTo ensure that an infection does not spread to another colony, sterilize all rearing trays by eitherautoclaving them or soaking them in a 5-10% household bleach solution for 24 hours. Wipe down allcounters with a 5-10% bleach solution and replace any larval rearing diets in the rearing room.FungiFungi have been considered one of the most pathogenic organisms that can infect mosquitoes. A largenumber of fungi have been isolated from mosquitoes, both wild and laboratory reared, recently reviewedby (Scholte et al. 2004). Most fungal infections are transmitted by free-floating spores.Coelomomyces spp This is the one of the most widely studied fungi that infects mosquitoes. Infectionstypically occur in early larval instars, and infected larvae rarely pupate or emerge. The infection can bedetected by locating “buds” or lumped structures in the anal gills of the larva (Figures 2.2.5 and 2.2.6). Ifinfection is suspected, dissecting the larvae should reveal hyphae emanating from several tissuesincluding the malpighian tubes and muscles. The hyphae will be branched, multinucleate, and clavate(club) shaped. Sporongia, the reproductive portion, will develop into dark spherical spores. Although thisis highly pathogenic, it is self limiting and usually only 1 or 2 generations will be affected due to the needof an intermediary host to complete its life cycle (Madelin 1965), (Scholte et al. 2004). In Africa, thesefungi have been often reported to be isolated from the ovaries and fat bodies of females only (Hazard1973). Authors have reported that larvae infected with Coelomomyces fungi often have a yellow, orange,or brownish color due to a number of sporangia that have developed internally (Kramer 1964).Leptolegnia spp. These are typically encountered in wild isolates. They are highly pathogenic, especiallyto L1 and L2 larvae. In An. gambiae it was shown that 100% of larvae were killed within 72 hours postinfection (Scholte et al. 2004).

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 5 of 10Figure 2.2.5. Normal anal gill of larva ofAnopheles gambiae.Figure 2.2.6. Infected anal gill of larva of Anophelesgambiae.Other locations of notable fungal development are in the spermathecae and terminalia (Figures 2.2.7 and2.2.8).Figure 2.2.7 Fungal hyphae growing within aspermathecae of Anopheles quadriannulatus.Figure 2.2.8 Fungal mass on the terminalia ofa male Anopheles dirus.Entomophthora spp. Whereas Coelomomyces infects larvae, Entomophthora typically infects only adultmosquitoes. Spores are transmitted after the sexually mature conidiophore ruptures through the adultcuticle. The easiest method for detecting Entomophthora infections is to remove a recently deceased

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 6 of 10adult, place them on a black piece of paper and cover with a Petri dish. In 24-48 hours inspect the paper;if a white ring is visible around the mosquito, then the colony is infected. The white ring is formed by theforcible release of conidia into the atmosphere (Scholte et al. 2004).Aspergillus spp. As a cosmopolitan fungus, these can be readily isolated from almost any surface.Aspergillus is better known as the causative agent behind bread mold. Like Entomophthora, Aspergillusinfects by penetrating the adult cuticle. Although the exact reason behind its lethality is unknown, it hasbeen shown to cause mortality and reduced egg production in adult mosquitoes. Experimental researchhas also shown that larvae infected by Aspergillus will mature and emerge; however, adults will haveshorter lives and produce fewer offspring (Nnakumusana 1985). Aspergillus colonies typically have a darkgreen to black color and will quickly cover any surface where they are growing.Smittium spp. A recently reported fungus that can cause larval paralysis (personal communication). It isnot commonly encountered but has been reported to be highly lethal to Anopheles larvae compared toaedines.TreatmentAntifungal agents like PSF and fumigillin can be used in either adult diets or in the larval rearing pan. Aswith antibiotics, before any full scale implementation, trials should be conducted to determine the dosagerequired to eliminate the infection without killing large numbers of the larvae or adults.In the insectaryIsolate the infected colony and employ one-time use only cages until the condition improves. If youroutinely maintain colonies in permanent cages, you will want to purchase some temporary cages untilyou can completely clean the old cages. Surface sterilization of cages is typically ineffective since mostfungi produce spores that are resistant to desiccation or chemical treatment. However, soaking cages in astrong bleach solution overnight may help. Additionally, dispose of all diets, even unopened ones, thatmay have been the source of the infection. Clean the insectary with an antifungal agent.ProtistsAlthough these are rarely associated with mortality within a laboratory colony, their presence can meanimpending problems. Often parasitism with protozoans results in reduced mobility in mosquito larvae.Larvae often take on a whitish color and their abdomens swell due an increase in spores internally,eventually leading to deformation. Usually, infections are not evident and are transmitted throughembryos.Protists can be introduced from many sources including rearing water, spores in the air, a previouslyinfected colony, or from wild material. Protozoan infections can occur as internal or external infections.Microsporidians are typically responsible for internal infections while Pertrichidians are responsible forexternal infestations.Internal infectionsLankesteria culicis (Gregarina culicis) This protozoan is often isolated from the gut and malpighiantubules in adult mosquitoes. Aedine colonies are a popular source, but this protozoan has also beenisolated from a few other genera. Larvae can become infected after ingesting spores in the rearing water.Mature forms are then released back into the larval water during the adult emergence stage. L. culicis isnot considered highly pathogenic unless found in high numbers (Jenkins 1964).Amblyosporidae (Amblyospora and Parathelohania). These protists are often isolated from wildmaterial. Parathelohania microsporidians are almost exclusively parasites of anopheline mosquitoes.They are transovarially transmitted and require a copepod intermediate host therefore infections will belimited in an insectary (Becnel and Andreadis 1999). Infections are principally seen within the fat bodies,and mortality occurs before pupation. In Africa, infections were associated with L4 male larvae resulting insex specific mortality (Hazard 1973).

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 7 of 10Brachiola algerae (Nosema algerae) This is considered one of the most important infections in aninsectary. As with other protists, these are often found infecting the tissues associated with larval fatbodies, ovaries, midguts, and gastric systems in mosquitoes. Infections with this parasite are rarelyreported from insectaries. Intra-insectary transmission is reported often from colony to colony (Savageand Lowe 1970). In An. gambiae, infections with Brachiola have caused a reduction in egg production(Jenkins 1964). Infections are transmitted via contaminated surfaces, especially eggs, therefore surfacesterilization of eggs can be used to break the transmission cycle. It is a chronic infection that will sweepthrough a colony until all individuals are infected (J. Becnel personal communication).Vavraia spp (Plistophora). These were formally recognized as Brachiola but have since been movedinto a new genus. As in Brachiola, Vavraia parasitize the malpighian tubules and ovaries and will continueto infect other organs over time. However, mortality is usually negligible. Transmission of this parasite isthought to occur through the embryos. The main effect of parasitism is Vavraia’s ability to interfere withookinete development of Plasmodium parasites within the midgut (Jenkins 1964), (Kramer 1964)).Infections within laboratory colonies have been reported from minimal to critical, and some infectionshave been said to spontaneously disappear.External infectionsEpistylis spp. These protists are recognized as epibionts (An organism that lives on the body surface ofanother) of mosquito larvae. Although they attach themselves to larvae, they do not cause any diseasewithin the animal. These generally will appear as a fuzzy coating around the larvae. Mortality frominfestation with Epistylis is caused by either physically blocking the larva from feeding or by overwhelmingthe larva and reducing its normal movements. Heavily infested larvae may have trouble reaching thesurface to breathe or their siphons may be blocked by attached protists (Larson 1967).Vorticella spp. As in Epistylis, these are considered epibionts of mosquito larvae and appear as a fuzzycoating around the larvae (Figures 2.2.9 and 2.2.10) (Schober 1967). Although they are frequentlyencountered in nature, they have been isolated in laboratory colonies (Armstrong and Bransby-Williams1961; Hati and Ghosh 1961; Larson 1967).Figure 2.2.9 Vorticella-infested Anopheles stephensi L4.

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 8 of 10Figure 2.2.10 Vorticella-infested anal gills of Anopheles stephensi L4.TreatmentThere are no formal treatments for most protozoan infections. Some researchers have had some successin treating Vorticella infestations with mepacrine hydrochloride (Jupp and Smith 1986). Surfacesterilization of eggs can also help to eliminate microsporidian infections (see Chapter 2 Culture section fortechnique).In the insectaryInfected colonies may need to be replaced or isolated to ensure the infection does not spread to otherinsect colonies. Some microsporidian infections are self-limiting and will resolve themselves within 1-2generations. However, others are persistent due to transmission through the embryos. Routinedisinfection of pans and surfaces should limit the number of resistant spores. Epibionts are frequentlyassociated with poor rearing conditions. Therefore, efforts should be made to keep rearing water as cleanas possible and any infected larvae should be removed promptly. For Vorticella treatment, it is possible toreduce the number of protists affecting a colony by adding a 1% bleach solution to the rearing water in a1:20-1:30 ratio.NematodesAlthough these rarely occur within insectaries, they can be introduced into an insectary from fieldcollectedmaterial. Mermithids, the main pathogen from this group, are worms that infect mosquito larvaeand typically cause high mortality before pupation (Kalucy 1972).AcaridsAcarids are mites (see Figure 2.2.11) from several genera that often attach themselves to the abdomenor thorax of an adult mosquito. Light infestations are not fatal; large infestations, however, can lead tomortality (Jenkins 1964).

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 9 of 10Figure 2.2.11. Shown is a mite that wasfound persistently in the insectary,typically in egg dishes and in the bottomof cages. Harm from this mite has notbeen observed.The MR4 would like to acknowledge Dr. Amanda Lawrence from Mississippi State University, Departmentof Entomology for her assistance in providing information used to develop this section.ReferencesArmstrong JA, Bransby-Williams WR (1961) The maintenance of a colony of Anopheles gambiae, withobservations on the effects of changes in temperature. Bull World Health Organ 24:427-435Becnel JJ (2007) Current Status of Deltabaculoviruses, Cypoviruses and Chloriridoviruses Pathogenic forMosquitoes. Virologica Sinica 22:117-127Becnel JJ, Andreadis TG (1999) Microsporidia in Insects. In: Murray Wittner; Louis M. Weiss ce (ed) TheMicrosporidia and Microsporidiosis. American Society for Microbiology, Washington D.C.Becnel JJ, White SE (2007) Mosquito Pathogenic Viruses - The Last 20 Years. AMCA Bulletin No. 723:36-49Bird RG, Draper CC, Garnham PC, Pudney M, Shute PG, Varma MG (1970) Evidence of insect viruses incolonies of Anopheles stephensi. Trans R Soc Trop Med Hyg 64:28-29de St. Jeor SC, Nielsen LT (1964) The use of antibiotics as an aid in rearing larvae of Culex tarsalisCoquillet. Mosq. News 24:133-137Federici BA (1985) Viral Pathogens. In: Biological Control of Mosquitoes American Mosquito ControlAssociation, Fresno BullHati AK, Ghosh SM (1961) Vorticella infestation of mosquito larvae and its effects on their growth andlongevity. Bull. Cal. S.T.M. 9:135Hazard EI (1973) Investigation of pathogens of anopheline mosquitos in the vicinity of Kaduna, Nigeria.In. World Health Organization, Geneva, p WHO/VBC/72.384Jenkins DW (1964) Pathogens, Parasites and Predators of Medically Important Arthropods. AnnotatedList and Bibliography. Bull World Health Organ 30:SUPPL:1-150

Chapter 2 : Anopheles Laboratory Biology and Culture2.2 Infections in Mosquito CulturesPage 10 of 10Jupp PG, Smith AN (1986) The use of mepacrine hydrochloride to control Vorticella on mosquito larvae. JAm Mosq Control Assoc 2:375-376Kalucy EC (1972) Parasitism of Anopheles annulipes Walker by a mermithid nematode. Mosq. News32:582-585Kramer JP (1964) Parasites in Laboratory Colonies of Mosquitoes. Bull World Health Organ 31:- 475-478Larson LV (1967) Association of Vorticella and Epistylis (Protozoa: Ciliata) with mosquito larvae. In. WorldHealth Organization, Geneva, p WHO/VBC/67.20Madelin MF (1965) Further laboratory studies on a species of Coelomomyces which infects Anophelesgambiae Giles. In. World Health Organization, Geneva, p WHO/EBL/52.65Muspratt J (1966) Parasitology of larval mosquitoes, especially Culex pipiens fatigans Wiedemann atRangoon, Burma. In. World Health Organization, Geneva, p WHO/EBL/18Nnakumusana ES (1985) Laboratory infection of mosquito larvae by Entomopathogenic fungi withparticular reference to Aspergillus parasiticus and its effects on fecundity and longevity of mosquitoesexposed to conidial infections in larval stages. Curr Sci. 54:1221-1228Savage KE, Lowe RE (1970) Studies of Anopheles quadrimaculatus infected with a Nosema sp. In: Proc.4th Int. Colloq. Insect Pathol., pp 272-278Schober H (1967) Observations on Culex pipiens larvae infested with Vorticella sp. Mosq. News 27:523-524Scholte EJ, Knols BG, Samson RA, Takken W (2004) Entomopathogenic fungi for mosquito control: areview. J Insect Sci 4:19Toure AM, Mackey AJ, Wang ZX, Beier JC (2000) Bactericidal effects of sugar-fed antibiotics on residentmidgut bacteria of newly emerged anopheline mosquitoes (Diptera: Culicidae). J Med Entomol 37:246-249

Chapter 2 : Anopheles Laboratory Biology and Culture2.3 Modifying Fecundity, Longevity and Size v2Page 1 of 62.3 Modifying Fecundity, Longevity and SizePaul Howell and Liz WilkinsIntroductionMaintaining proper nutrition throughout larval development will have positive effects for the entirety of amosquito’s life. Optimization of nutrition, photoperiod, competition, and temperature will result in healthierlarvae and a more productive colony. If any of these is suboptimal, smaller and generally less vigorousmosquitoes will result. Large size has been often associated with increased fecundity and longevity inmany different species (Blackmore and Lord 2000, Briegel 1990, and Takken et al. 1998). Here wepresent some general information to assist in establishing rearing protocols for anopheline mosquitoes.Larval Diet and NutritionIncreased fecundity and longevity result partly from reserves accumulated during immature stages.Several factors can reduce these reserves and result in poor quality adults. Some studies indicate thatdiets high in protein are superior in producing larger, more fecund mosquitoes. Increases in egg clutchsize have been positively associated with high protein diets (Akoh et al. 1992, Lang 1978). Conversely,sub-optimal diets resulted in the production of smaller adults which were less likely to seek out a bloodmeal (Klowden et al. 1988). A high protein diet also reduced immature development times inToxorhynchites splendens (Amalraj et al. 2005). Therefore the quality of the diet can have significanteffects on colony production and growth.Not only is the quality of diet fed to larvae important, but the quantity is also significant. Reisen et al.(1984) illustrated the relationship between the concentration of food and larval development to pupation.Overfeeding larvae will often lead to high larval mortality (Reisen 1975). However, the surviving larvae willdevelop rapidly and result in large adults (Lillie and Nakasone 1982; Reisen et al. 1984). Conversely,underfed larvae will result in the production of smaller adults. In Cx. pipiens fatigans, underfeeding leadsto a reduction in the number of ovarian follicles (Ikeshoji 1965; Arrivillaga and Barrera 2004). Therefore,measured amounts of a larval diet should be supplied to ensure optimal growth and fecundity.Dietary Restriction (DR) has been studied for its effect on longevity extensively in flies and mice. Thoughthe results are not completely in agreement, the general conclusions are compatible. By and large, inanimals and insects, one of the most successful ways to extend longevity is through DR (Burger andPromislow 2004). For many species of animals, restricting diets yields animals that are leaner and haveincreased longevity (Hopkin 2003). These same animals, however, are typically less fertile and becomeinfected or sick more easily. DR animals are much slower than their better-fed counterparts (Hopkin2003). DR-affected animals live longer but show signs of age much more quickly (Hopkin 2003; Burgerand Promislow 2004). Longevity benefits are easily reversed with the introduction of any stressors to theenvironment as the DR animals are extraordinarily susceptible, especially to infection and sickness. Inthose cases, the better nourished animals will have the greater longevity (Hopkin 2003). A balanced dietat the beginning of life has the greatest benefit on overall lifetime longevity (Rasnitsyn and Yasyukevich1988; McCay et al. 1989).In flies, DR increased the life span of all females including fertile or sterile flies. Dietary manipulationgenerally has a greater and longer effect on females than males, though in a few studies, the opposite istrue (Burger and Promislow 2004).Alternately, it has also been reported that in some animals, the quality of the diet is the more importantfactor in increased growth and longevity than the amount (McCay et al. 1989). Most generally, in the wild,reproduction and speed are the most important elements for survival, and both are severely hindered byDR (Hopkin 2003). So, for a wild animal or insect, the best bet for reproduction and evasion of predatorsis to be big and fat (Hopkin 2003)!

Chapter 2 : Anopheles Laboratory Biology and Culture2.3 Modifying Fecundity, Longevity and Size v2Page 2 of 6Some dietary supplementation can be beneficial in a few insect genera. Folic acid supplementationincreased egg production in the screwworm Cochliomyia hominivorax, even in the presence of a reducingagent (Crystal 1964). The addition of nordihydroguariaretic acid (NDGA) led to increased longevity inadult Ae. aegypti mosquitoes (Richie et al. 1986). The MR4 staff has observed significant increases inlongevity by addition of methylparaben to the sugar meal, though the effect is probably one of reducingmicrobial growth.Larval DensityIn addition to larval diet, the density at which larvae are cultured will affect the size and quality of adultmosquitoes. Overcrowded larvae often become smaller, short lived adults. Ae. sierrensis larvae rearedunder high densities resulted in the production of small adults that had reduced life spans whencompared to a field population (Hawley 1985). Similar results have been reported in Cx. tarsalis (Reisenet al. 1984), Wyeomyia smithii (Bradshaw and Holzapfel 1992) Ae. aegypti (Bedhomme et al. 2003), andAn. stephensi (Reisen 1975). An. gambiae larvae reared in crowded conditions resulted in increasedlarval development times and small adult body size; however, no appreciable effect on longevity wasnoted (Takken et al. 1998; Gimnig et al. 2002). This effect was also seen in An. arabiensis, but the sizelimitingeffects of crowding could be overcome by adding maize pollen to the rearing water (Ye-Ebiyo etal. 2003). When An. gambiae was reared in the same container with An. arabiensis at high densities, theeffects of overcrowding were not seen in An. gambiae. An associated reduction in lifespan of An.arabiensis was noted (Schneider et al. 2000).Increased larval density has also been linked to sex-specific larval mortality. In Cx. quinquefasciatus andAn. stephensi, larvae reared at high densities suffered excess male mortality (Reisen 1975; Suleman1982). It has been hypothesized by one of the authors that this skewed sex ratio may be due to greatermale susceptibility to stressors than females due to their smaller size and reserves (Reisen 1975).Overcrowding also results in increased larval development times. In Cx. tarsalis, overcrowding increaseddevelopment times from 10 days to 14 days (Reisen et al. 1984); in An. gambiae, overcrowding resultedin an average of an additional 1-2 days (Gimnig et al. 2002).Finally, overcrowding affects the fecundity of females. In Wy. smithii, lifetime fecundity can be linked topupal mass whereby larger females produced more eggs over a lifetime than smaller ones (Bradshawand Holzapfel 1992). Similarly, in An. stephensi, it was found that females originating from crowded pansproduced fewer eggs than those from less crowded environments (Reisen 1975). Therefore, rearinganimals at suitable densities e.g. 1 L4 per 1-2 ml of water, will result in long lived and more fecundmosquitoes than if reared at sub-optimal conditions.TemperatureAlthough not as obvious as nutrition and larval density, ambient temperature can dramatically effectmosquito production. Most mosquito larvae are reared around 25-27°C which is similar to theenvironment from which they are isolated. High temperatures can be lethal. In An. albitarsis and An.aquasalis, colder temperatures delayed embryo eclosion by 2 days or more (de Carvalho et al. 2002).Additionally, there was a reported reduction in the hatch rate of An. albitarsis when reared at 21°C. In Ae.albopictus, larvae reared at 26°C pupated faster than those reared at 22°C; however, fewer larvaecompleted ecdysis at the higher temperature (Alto and Juliano 2001). This trend has also been reportedin Cx. tarsalis (Reisen et al. 1984) and An. sergentii (Beier et al. 1987). In addition to reduced ecdysisrates, adults from temperature stressed environments can have reduced longevity. In An. gambiae(Afrane et al. 2006), An. superpictus (Ayetkin et al. 2009) and Cx. tarsalis (Reisen et al. 1984), adultsderived from heat stressed larval regimens were found to have life spans reduced by several days.Interestingly, in Ae. dorsalis, mosquitoes from a high temperature regimen had a reduced number ofovarian follicles (Parker 1982) while An. gambiae mosquitoes derived from the same environment hadincreased fecundity when compared to adults from a lower temperature region (Afrane et al. 2006).Finally, as temperatures increased, it was found that both the larval head capsule size and adult wing sizedecreased in An. merus (le Sueur and Sharp 1991). Wing size and shape were also affected byincreased temperatures in An. superpictus (Ayetkin et al. 2009). Therefore careful maintenance and

Chapter 2 : Anopheles Laboratory Biology and Culture2.3 Modifying Fecundity, Longevity and Size v2Page 3 of 6monitoring of the correct temperature in the rearing environment is necessary in order to produceconsistent-sized mosquitoes.PhotoperiodPhotoperiod is defined as the amount and schedule of light versus darkness. Even less studied thantemperature, the effect of photoperiod can dramatically affect longevity in adult mosquitoes. Manyinsectaries operate on a 12 hour day/night cycle since it is convenient for workers and similar to averageconditions. Cx. pipiens reared under a short photoperiod had smaller ovarian follicles than those rearedunder a long photoperiod (Oda and Nuorteva 1987). This was attributed by the author to the mosquitoespreparing for diapause, which can be induced by shortening the photoperiod.Reducing the photoperiod has also been found to be positively associated with an increased lifespan inAnopheles. In both An. crucians and An. quadrimaculatus, larvae reared under a short photoperiodregimen produced adults that lived longer than those reared under a long photoperiod (defined as greaterthan 15 hours of daylight) (Lanciani 1993; Lanciani and Anderson 1993).The length of photoperiod chosen, therefore, should reflect a balance between what is convenient for theinsectary staff and what is necessary to maintain a healthy colony.Adult dietThough not as critical as other effectors mentioned above, adult diet can also have an affect on thelongevity and fecundity of colonized vectors (reviewed in Foster 1995). Ae. aegypti fed sucrosesupplemented with nodihydroguariaretic acid lived longer than those reared on sucrose alone (Richie etal. 1986). An. gambiae mosquitoes fed sucrose plus blood lived longer than those fed just sucrose orblood alone. However, mosquitoes given only blood were more fecund than those fed sucrose and blood(Gary and Foster 2001). An. gambiae fed sucrose supplemented with 0.2% w/v methylparaben livedlonger than controls fed on sucrose alone (Benedict et al. 2009). Supplementation with uric and ascorbicacids lead to an overall increase in fecundity over the lifetime of a female in An. gambiae (DeJong et al2007). In general, however, sucrose solutions are suitable for regular maintenance of laboratory coloniesand care should be taken before adding supplements to the adult diet.The rearing environment, therefore, can have dramatic effects on laboratory colonies. Subtle changes inany of the above mentioned areas could result in declining fecundity or longevity and lead to colonycollapse. Optimization of some of these areas could result in a healthier colony however. It is suggestedthat testing should be done on a sample population before implementation of any changes as results mayvary between species, colonies and laboratories.ReferencesAfrane YA, Zhou G, Lawson BW, Githeko AK, Yan G (2006) Effects of microclimatic changes caused bydeforestation on the survivorship and reproductive fitness of Anopheles gambiae in western Kenyahighlands. Am J Trop Med Hyg 74:772-778Akoh JI, Aigbodion FI, Kumbak D (1992) Studies on the effect of larval diet, adult body weight, size ofblood-meal and age on the fecundity of Culex quinquefasciatus (Diptera: Culicidae). Insect Sci. Applic.13:177-181Alto BW, Juliano SA (2001) Temperature effects on the dynamics of Aedes albopictus (Diptera: Culicidae)populations in the laboratory. J Med Entomol 38:548-556Amalraj DD, Sivagnaname N, Das PK (2005) Effect of food on immature development, consumption rate,and relative growth rate of Toxorhynchites splendens (Diptera: Culicidae), a predator of containerbreeding mosquitoes. Mem Inst Oswaldo Cruz 100:893-902

Chapter 2 : Anopheles Laboratory Biology and Culture2.3 Modifying Fecundity, Longevity and Size v2Page 4 of 6Arrivillaga J, Barrera R (2004) Food as a limiting factor for Aedes aegypti in water-storage containers. JVector Ecol 29:11-20Ayetkin S, Ayetkin AM, Alten B (2009) Effect of different larval rearing temperatures on the productivity(Ro) and morphology of the malaria vector Anopheles superpictus Grassi (Diptera: Culicidae) usinggeometric morphometrics. J Vector Ecol 34:32-42Bedhomme S, Agnew P, Sidobre C, Michalakis Y (2003) Sex-specific reaction norms to intraspecificlarval competition in the mosquito Aedes aegypti. J Evol Biol 16: 721-730.Beier MS, Beier JC, Merdan AA, el Sawaf BM, Kadder MA (1987) Laboratory rearing techniques andadult life table parameters for Anopheles sergentii from Egypt. J Am Mosq Control Assoc 3:266-270Benedict MQ, Hood-Nowotny RC, Howell PI, Wilkins EE (2009) Methylparaben in Anopheles gambiae s.l.sugar meals increases longevity and malaria oocyst abundance but is not a preferred diet. J InsectPhysiol 55:197-204Blackmore MS, Lord CC (2000) The relationship between size and fecundity in Aedes albopictus. JVector Ecol 25:212-217Bradshaw WE, Holzapfel CM (1992) Reproductive consequences of density-dependent size variation inthe pitcherplant mosquito, Wyeomyia smithii (Diptera: Culicidae). Ann Entomol Soc Am 85:274-281Briegel H (1990) Fecundity, metabolism, and body size in Anopheles (Diptera: Culicidae), vectors ofmalaria. J Med Entomol 27:839-850Burger JM, Promislow DE (2004) Sex-specific effects of interventions that extend fly life span. Sci AgingKnowledge Environ 2004:pe30Crystal MM (1964) Insect Fertility: Inhibition by Folic Acid Derivatives. Science 144:308-309de Carvalho SC, Martins Junior Ade J, Lima JB, Valle D (2002) Temperature influence on embryonicdevelopment of Anopheles albitarsis and Anopheles aquasalis. Mem Inst Oswaldo Cruz 97:1117-1120DeJong RJ, Miller LM, Molina-Cruz A, Gupta L, Kumar S, Barillas-Mury C (2007) Reactive oxygenspecies detoxification by catalase is a major determinant of fecundity in the mosquito Anophelesgambiae. Proc Natl Acad Sci USA 104:2121-2126Farkas MJ, Brust RA (1985) The effect of a larval diet supplement on development in the mosquitoWyeomyia smithii (Coq.) under field conditions. Can J. Zool. 63:2110-2113Foster WA (1995) Mosquito sugar feeding and reproductive energetics. Annual Review of Entomology40:443-474Gary RE, Jr., Foster WA (2001) Effects of available sugar on the reproductive fitness and vectorialcapacity of the malaria vector Anopheles gambiae (Diptera: Culicidae). J Med Entomol 38:22-28Gimnig JE, Ombok M, Otieno S, Kaufman MG, Vulule JM, Walker ED (2002) Density-dependentdevelopment of Anopheles gambiae (Diptera: Culicidae) larvae in artificial habitats. J Med Entomol39:162-172Hawley WA (1985) The effect of larval density on adult longevity of a mosquito; Aedes sierrensis:epidemiological consequences. J Anim Ecol 54:955-964

Chapter 2 : Anopheles Laboratory Biology and Culture2.3 Modifying Fecundity, Longevity and Size v2Page 5 of 6Hopkin K (2003) Dietary drawbacks. Sci Aging Knowledge Environ 2003:NS4Ikeshoji T (1965) The influence of larval breeding conditions on fecundity of Culex pipiens fatigans Wied.In. WHO, Geneva, p WHO/Vector Control/135.165: Report no.111Klowden MJ, Blackmer JL, Chambers GM (1988) Effects of larval nutrition on the host-seeking behaviorof adult Aedes aegypti mosquitoes. J Am Mosq Control Assoc 4:73-75Lanciani CA (1993) Photoperiod and longevity in Anopheles crucians. J Am Mosq Control Assoc 9:308-312Lanciani CA, Anderson JF (1993) Effect of photoperiod on longevity and metabolic rate in Anophelesquadrimaculatus. J Am Mosq Control Assoc 9:158-163Lang JT (1978) Relationship of fecundity to the nutritional quality of larval and adult diets of Wyeomyiasmithii. Mosq. News. 38:396-403le Sueur D, Sharp BL (1991) Temperature-dependent variation in Anopheles merus larval head capsulewidth and adult wing length: implications for anopheline taxonomy. Med Vet Entomol 5:55-62Lillie TH, Nakasone RI (1982) An evaluation of commercial diets for rearing Wyeomyia smithii. MosqNews 42:225-231McCay CM, Crowell MF, Maynard LA (1989) The effect of retarded growth upon the length of life spanand upon the ultimate body size. 1935. Nutrition 5:155-171; discussion 172Oda T, Nuorteva P (1987) Autumnal photoperiod and the development of follicles in Culex pipiens pipiensL. (Diptera, Culicidae) in Finland. Ann. Entomol. Fennici. 53:33-35Parker BM (1982) Temperature and salinity as factors influencing the size and reproductive potentials ofAedes dorsalis (Diptera: Culicidae). Ann. Entomol. Soc. Am. 75:99-102Rasnitsyn SP, Yasyukevich VV (1988) On the ability of mosquito larvae (Diptera, Culicidae) to endurestarvation. Entomologicheskoye Obozreniye 4:708-715Reisen WK (1975) Intraspecific competition in Anopheles stephensi Liston. Mosq. News. 35:473-482Reisen WK, Milby MM, Bock ME (1984) The effects of immature stress on selected events in the lifehistory of Culex tarsalis. Mosq. News. 44:385-395Richie JP, Jr., Mills BJ, Lang CA (1986) Dietary nordihydroguaiaretic acid increases the life span of themosquito. Proc Soc Exp Biol Med 183:81-85Schneider P, Takken W, McCall PJ (2000) Interspecific competition between sibling species larvae ofAnopheles arabiensis and An. gambiae. Med Vet Entomol 14:165-170Suleman M (1982) The effects of intraspecific competition for food and space on the larval developmentof Culex quinquefasciatus. Mosq. News. 42:347-356Takken W, Klowden MJ, Chambers GM (1998) Effect of body size on host seeking and blood mealutilization in Anopheles gambiae sensu stricto (Diptera: Culicidae): the disadvantage of being small. JMed Entomol 35:639-645

Chapter 2 : Anopheles Laboratory Biology and Culture2.3 Modifying Fecundity, Longevity and Size v2Page 6 of 6Ye-Ebiyo Y, Pollack RJ, Kiszewski A, Spielman A (2003) Enhancement of development of larvalAnopheles arabiensis by proximity to flowering maize (Zea mays) in turbid water and when crowded. AmJ Trop Med Hyg 68:748-752

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.1 Collecting Anopheles EggsPage 1 of 22.4 Anopheles Culture2.4.1 Collecting Anopheles EggsMR4 StaffIntroductionFor colony maintenance, we blood feed females only once in their lifetime, between 3 and 7 days postemergence(see Chapter 1, Rearing Schedule). After subsequent feedings, anophelines will lay eggsagain, which is helpful for maintaining a colony and having plenty of experimental material. Afterbloodfeeding, allow 2-3 days for embryo development prior to egging. Certain species may take moretime (An. dirus - 4 days), so you may have to modify your schedule according to the needs of yourparticular colony.Among the water types we have used, we have observed no effect on oviposition. While there are almostcertainly measurable differences that would be important in mass rearing facilities, an excess of eggsbeyond what is required for stock keeping is almost always obtained, and efficiency is not an issue.Most eggs are laid at night, and egg dishes are typically removed the day following their insertion. Thereare several ways to collect eggs. One method is to fill a cup with clean water to about 1 cm depth and linethe edges with filter paper such that half of the paper is submerged (Figure 2.4.1.1). The filter paperprevents the eggs from sticking to the dry plastic sides of the cup when it is sloshed.Other methods include allowing mosquitoes to lay eggs on wet filter paper supported by wet cotton orsponges which are then floated in water the next day, filter paper funnels partially submerged in water,and dark bowls lined with filter paper (Figure 2.4.1.2-Figure 2.4.1.4).Regardless of the method used, after collecting eggs, remove all dead adults to help prevent carryinginfections into the new larva pans. Adults can be removed individually with forceps or filtered out bywashing the egg/adult mix through a screen and collecting the eggs beneath.Figure 2.4.1.1. Egg collection cupcontaining standing water to a depth that isapproximately half way up the filter paperliner.Figure 2.4.1.2. Filter paper on a damp sponge in aplastic dish. The sides are lined with either white orblack paper, and it is apparent that An. gambiae andarabiensis prefer resting on the latter for oviposition.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.1 Collecting Anopheles EggsPage 2 of 2Figure 2.4.1.3. Filter paper funnel partiallysubmerged in water.Figure 2.4.1.4. Dark bowl lined with filter paper.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.2 Bleaching Anopheles EggsPage 1 of 22.4.2 Bleaching Anopheles EggsMR4 StaffIntroductionMany laboratories find that surface-cleansing of eggs with bleach is a helpful routine procedure tominimize the growth of microsporidians (Alger and Undeen 1970; Robert et al. 1993). Mortality due tothese pathogens is sometimes observed only after several generations of culture without surfacecleansing. The following method describes a method to ‘surface sterilize’ eggs and also allows forconcentrating eggs on a filter paper disk. This only works well if eggs are collected in water or on asurface from which they are easily removed.Materials• Filtration device (A Buchner funnel is an option, but take care to clean carefully if used for severalstocks since eggs will leak around the edge of the paper). Figure 2.4.2.1 shows an example of aNalgene 500 ml disposable filter modified by carefully breaking off the upper chamber andremoving the membrane. It can be reused indefinitely.• 1% v/v household bleach in wash bottle• Sterile water in wash bottle• A vacuum source equipped with trapProtocol1. Remove adults from surface by filtering through screen or with forceps.2. Center filter paper on platform with vacuum applied.3. Lightly wet the filter paper with sterile water and ensure that the filter paper is held securely by thevacuum.4. Slowly pour water/eggs onto the center of the disk so that eggs do not spill outside of the depression(Figure 2.4.2.2).5. Remove vacuum.6. Add the bleach solution to the entire depression for up to 1 minute. Remove bleach by applyingvacuum.7. Add water until depression is full. Apply vacuum.8. Wash with water two times. Apply vacuum until enough surface water remains to keep eggs moistwithout allowing them to run off the sides of the paper. Eggs can typically be stored in a humidcontainer on filter paper for up to 24 hours before hatching.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.2 Bleaching Anopheles EggsPage 2 of 2Figure 2.4.2.1 Modified chamberattached to vacuum.Figure 2.4.2.2 Pour eggs gently into the center of thefiltration device. Use bleach solution to remove finaleggs from the sides of the container.ReferencesAlger NE, Undeen AH (1970) The control of a microsporidian, Nosema sp., in an anopheline colony by anegg-rinsing technique. J Invertebr Pathol 15:321-327Robert V, Tchuinkam T, Carnevale P (1993) Successful eradication of a microsporidian, Nosema sp., in amosquito colony. Ann Soc Belg Med Trop 73:71-72

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.3 Hatching Anopheles EggsPage 1 of 22.4.3 Hatching Anopheles EggsMR4 StaffIntroductionAnopheles eggs are usually hatched as soon as they are mature – normally about two days after they arelaid. Methods to dry and store Anopheles eggs for a couple of weeks have been developed (Bailey et al.1979), but these are useful primarily in larger production settings. Eggs are usually hatched at a highdensity and given a diet consisting of small particles such as live baker’s or brewer’s yeast. Whileexcessive crowding can stunt their growth irreversibly, culture at too low a density often results inexcessive microbial growth and larval death.This protocol starts with eggs that have been collected and possibly bleached and are sufficiently matureto hatch immediately. See the sections on Collecting (2.4.1) and Bleaching Anopheles Eggs (2.4.2) formore information.Materials• 2 % w/v active (live) baker’s or brewer’s yeast in purified water. Mix and use only that day.• water wash bottle• mosquito tray• 5 ml pipet• 50-100 ml bottle• mosquito culture waterProcedure1. For each egg batch, add deionized water to a rearing pan so that water just completely covers thebottom of the pan. Add more if your trays will be uncovered (not recommended). Avoid adding toomuch water as this will increase larval mortality (Timmermann and Briegel 1993).2. Add a yeast suspension to each pan to a final concentration of 0.02% (Figure 2.4.3.1). Forexample, in the pan shown in Figure 2.4.3.1, we add 300 mL of water and 2 mL of a 2% w/v yeastsolution. The pan shown will rear approximately 300 L4s at a nice density. Swirl until the yeast hasdispersed throughout the pan. Place the pan on the shelf or rack where it will be kept.3. Hold the egging paper by the edge to avoid touching the eggs (Figure 2.4.3.2) and gently rinsethe eggs into the pan (Figure 2.4.3.3). Ensure that no eggs stick to your fingers as they can easilybe accidentally transferred to the next pan. Be careful that eggs do not splash into an adjacentpan or onto the water bottle. After rinsing the eggs into the pan and before rinsing the next strain,clean your fingers using a paper towel. Be careful not to disturb or move the pan at all after addingthe eggs. Disturbing the pan can cause the eggs to stick to the sides of the pan above the waterwhere they will dry and not hatch. If you accidentally jar the pan, gently rinse the eggs down fromthe side of the pan into the rearing water.4. Make sure the pans are clearly labeled with strain name and date of hatching.5. Cover the pan to prevent contamination that could occur from accidental splashing when rinsingeggs into an adjacent pan and oviposition by loose females that can contaminate the stock.6. Anopheles eggs will generally hatch immediately or within 24-48 hours after placement in water.On the day after placing the eggs in water, without disturbing the pans, gently uncover and scan tosee if there are L1s present. They are very small and hard to see in many genera (Figure 2.4.3.4).If you do not see any L1 larvae, carry the pan into a well lit area and check again. For more onscreening hatching of individual eggs microscopically, see Chapter 3.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.3 Hatching Anopheles EggsPage 2 of 27. Allow one day between placing the eggs in water and splitting or thinning.Figure 2.4.3.1. Add yeast slurry to pan with justenough water to cover the bottom.Figure 2.4.3.2. Hold egg paper in such a way thatyour fingers do not touch the eggs.Figure 2.4.3.3. Gently rinse eggs into pan. Coverand do not disturb for 24 hours.Figure 2.4.3.4. 24 hours after rinsing eggs into tray,check for hatch (stock G3 An. gambiae sensu strictoshown).NotesMany labs line the trays with a strip of filter paper. This prevents the eggs from becoming stranded anddrying out if the water is sloshed. This technique is useful if you are adding the eggs to the tray and thenmoving it to the shelf where the eggs will hatch. When using this method, discard the paper whenhatching is complete.An efficient method for food preparation is to pre-package 1 g of yeast in 50 ml disposable Falcon tubesand store them at -20°C until needed. Simply remove as many tubes as are needed that day, mix withwater, and use immediately.ReferencesBailey DL, Thomas JA, Munroe WL, Dame DA (1979) Viability of eggs of Anopheles albimanus andAnopheles quadrimaculatus when dried and stored at various temperatures. Mosquito News 39:113-116Timmermann SE, Briegel H (1993) Water depth and larval density affect development and accumulationof reserves in laboratory populations of mosquitoes. Bull. Soc. Vector Ecol. 18:174-187

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.4 Determining Egg Hatch RatesPage 1 of 22.4.4 Determining Egg Hatch RatesMR4 StaffIntroductionEgg hatching rates vary between stocks depending on intrinsic fertilization rates, semisterility due tocrossing type or presence of chromosomal aberrations and the methods used to handle eggs afteroviposition. Egg hatch rates >80% are typical. Counting hatched larvae is not a proxy for determining thehatch rate as mortality may occur in the L1 stage and these larvae are very difficult to detect. Typically,anopheline eggs that sink to the bottom of the pan do not hatch; however, they should be inspected andincluded in hatch rate data.Unhatched eggs fall into several classes which may be of interest to record: (1) Unmelanised eggs areoften observed but will inevitably fail to hatch; (2) Unhatched, melanised eggs in which no indication of anembryo can be seen; (3) Unhatched, melanised eggs in which a developing embryo is seen but neverhatches; (4) Unhatched, melanised eggs in which an embryo is alive but has not hatched.The last category is problematic. In some species, hatch is very synchronous, but in others –An. gambiae – it occurs over several days even when eggs are moist (Lehmann et al. 2006). We haveoften observed egg batches in which most eggs have hatched a day earlier, but the activity of countingthe eggs stimulates further hatch. These various types and timings of hatching should be taken intoaccount when collecting and interpreting hatching data.Materials• Filter paper• Wash bottle containing water• Fine probesEquipment• Stereoscope• 2-place denominator (counter)Determining hatch rates for eggs collected fromindividual females1. Assuming eggs have been collected in tubes, theseshould have been lined previously with filter paper(See Chapter 3.9). If the eggs are not all at the edges,carefully touch the center of the water. The oil on yourfingers will usually cause the eggs to move to the side.If not, tease them to the edge with a probe.2. Very slowly and smoothly slide the papers up the sideof the tube and transfer to a rigid, movable, flatsurface. A small piece of Plexiglass (10 X 20 cM) isideal for this.Figure 2.4.4.1. An unhatched (left) andhatched egg (right) from the same female,laid on the same day. It is apparent thelower one has hatched because of thedislocated operculum. Un-melanised eggsdo not hatch.3. Count the eggs in situ under a stereoscope recording hatched vs. unhatched on the denominator.You may need to prod or burst the egg with a probe to determine whether it is hatched. Typically theoperculum of hatched eggs will be dislodged somewhat (Figure 2.4.4.1). If only a sample of eggs isneeded for rate information, counting 50 eggs is sufficient. Otherwise, count all the eggs.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.4 Determining Egg Hatch RatesPage 2 of 2Determining hatch rates for en masse egg collectionsIn this case, the container from which eggs are taken may or may not be lined with filter paper. If it is, skipto step 2.1. Slide a small piece of filter paper (approx 2 X 6 cm) down the side of the tray above the eggs to becollected. When the paper begins to wet, slowly slide it behind the eggs until the paper is wellsubmerged.2. Slowly slide the paper on which the eggs are resting up the side of the tray until it is above the water.The eggs should adhere to the paper as it is raised.3. Transfer the paper to the counting board as described above and count.ReferencesLehmann T et al. (2006) Genetic contribution to variation in larval development time, adult size, andlongevity of starved adults of Anopheles gambiae. Infect Genet Evol 6:410-416

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.5 Estimating the Number of Eggs and LarvaePage 1 of 42.4.5 Estimating the Number of Eggs and LarvaeMR4 StaffIntroductionIn order to maintain mosquito colonies, a sufficient number of individuals most be produced within aspecific time. The most efficient way to accomplish this is to consistently rear a fixed number of larvae pertray and volume of water and diet. This prevents waste of resources, creates healthier and more fecundcolonies, and you can reliably predict when you will have material available for experiments. Crowding theearly stages in anopheline larvae is detrimental and should be avoided in most species (exceptionsinclude An. quadrimaculatus and An. freeborni which seem to require higher densities in the earlyinstars). If you cannot consistently estimate the appropriate number of larvae, to prevent over-crowding inthe early instars you can either (1) measure the number of eggs or (2) the number of larvae per pan(Dame et al. 1978).In the following sections, we suggest two general approaches by which repeatable numbers of larvae canbe placed in trays.Estimating the number of eggs• If one is culturing large amounts of a single stock and the egg hatching rate is consistent,volumetric estimation of the number of eggs to place in each tray is a feasible approach. Sincelarvae are more susceptible to damage when handled individually in the early stages, Dame et al.(1978) suggested that this is the best method. They collected An. albimanus eggs, bleachedthem, (see Bleaching Anopheles Eggs, Chapter 2.4.2) and immediately floated them on 28°Cwater for 30 hours. Afterward, the eggs were strained through a screened bottom cup (100micron pore size) and allowed to dry by drawing air over the eggs at a rate of 600-700m/sec for30 minutes (Dame et al. 1978). Eggs were then measured into a severed 1ml disposable pipettefor dispensing. It was reported in An. albimanus that .085ml of eggs was equal to 5000-6000eggs.• For implementation, one would choose some small volume container into which semi-dry eggswould be placed using a fine paint brush. After filling a predetermined volume, the eggs would beremoved and counted. An average and standard deviation could be determined. Use of glassineweighing papers and methods to reduce static electricity will improve this method.• Depending on the sophistication and needs of the system, one could also devise photographicand image analysis methods to count eggs in a monolayer on paper. However, this would notresult in specific amounts for dispensing as the above method does.Estimating the number of larvaeIf one is working with numerous stocks, it is difficult to dry and measure eggs. Hence it is more commonto dispense a known number of larvae into rearing pans for culturing. Use L2s as they are less delicatethan L1s.• The simplest yet most time-consuming and accurate method is to individually count larvae intothe rearing pan. Using a pipette, aspirate and count several larvae and place the drop in thebottom of the pan until the desired number has been reached (e.g. 300 larvae per tray).• A faster but less accurate method is to use a photographic scale (Rutledge et al. 1976). Place anunknown number of larvae into a Petri dish. Place the dish near the photographs of countedlarvae at the same stage (Figure 2.4.5.1) and determine which one contains a similar number oflarvae. Adjust the number of larvae until the desired density is achieved.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.5 Estimating the Number of Eggs and LarvaePage 2 of 4100 200300 400Figure 2.4.5.1. Photographic guide to thevolume of larvae in a fixed volume. Allpictures are of a 6 cm diameter Petri dishcontaining 5 ml of water. These images are ofAn. quadrimaculatus L2s. From top left tobottom: 100, 200, 300, 400, and 500 larvae.500• More recently, a graphical display was developed for culicid larvae, which is similar in nature tothe photographic method above. A dish of larvae is compared to the pictograph from which thedensity is estimated (Carron et al. 2003).

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.5 Estimating the Number of Eggs and LarvaePage 3 of 4• Finally, one could pour the concentrated larvae into a known volume of water and mix them untilthey were uniformly dispersed (Gerberg et al. 1968, 1994). From the known volume 20-25 1mlaliquots are removed so as to create an estimate of larvae per ml from which aliquots can bedispensed into new rearing pans.ReferencesCarron A, Duchet C, Gaven B, Lagneau C (2003) An easy field method for estimating the abundance ofculicid larval instars. J Am Mosq Control Assoc 19:353-360Dame DA, Haile DG, Lofgren CS, Bailey DL, Munroe WL (1978) Improved Rearing Techniques for LarvalAnopheles albimanus : Use of Dried Mosquito Eggs and Electric Heating Tapes. Mosq. News 38:68-74Gerberg EJ, Barnard DR, Ward RA (1994) Manual for mosquito rearing and experimental techniques,revised ed. American Mosquito Control Association, Inc., Lake CharlesGerberg EJ, Gentry JW, Diven LH (1968) Mass rearing of Anopheles stephensi Liston. Mosq. News 28342-346Rutledge LC, Sofield RK, Piper GN (1976) Rapid counting methods for mosquito larvae. Mosq. News 36537-539

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Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.6 Anopheles Larval CulturePage 1 of 62.4.6 Anopheles Larval CultureMR4 StaffIntroductionExcept in rare cases, mosquito larval culture is septic, and the diet consists of both added food and themicrobial growth that results. Reliable diets probably provide good nutrition directly and also promote amicrobial environment that nourishes larvae. The relationship between microbial growth, larvalconsumption, water volume and larval density is complex, but the following recommendations are typicalfor Anopheles culture.Studies have shown that above a weight threshold, the number of eggs laid per female is in fairly directproportion to their adult weight. Adult size is ultimately determined by larval size, and that on larvalculture. So, it is logical that larger larvae yield higher numbers of eggs per female – a desirable outcomein most laboratory cultures.Consistent use of successful practices developed and proven in each lab should be adhered to faithfully.Culture water and the environmentWhen larval culture fails, laboratorians often suspect something is wrong with the water. In spite of thisconcern, many different sources of water have been used successfully - in combination with good rearingtechniques: chlorinated/fluoridated municipal supply, deionized, untreated deep well, and distilled water.Water source is probably not a critical factor for most species unless it contains high levels of toxicchemicals. If in doubt, try changing to a more purified form to see if conditions improve. The consistencyof the source should be considered in making the choice as this will determine long-term success.Mosquito rearing water is a rich environment in which to grow microbes. Therefore, 'more microbes in,even more microbes out' is a good way to think of larval culture. Use of almost any treated water willreduce microorganisms relative to natural sources. However, some chemical treatment methods used inmunicipal supplies may not be compatible with larval mosquitoes. Chloramine is reportedly of particularconcern for invertebrate culture. The rule of thumb is to test the water source with a small number oflarvae before using it on a larger scale. If you are concerned about chlorine residues, water from a hotwater source that is allowed to cool, or chlorinated water that is allowed to sit overnight should beadequate. For critical applications where physiologically uniform water is required, we usereverse/osmosis deionized UV sterilized water to which we add 0.3 g / liter artificial sea salts such asInstant Ocean.A water temperature of 27°C is suitable for rearing most anophelines. The room or incubator temperatureshould be adjusted to achieve this depending on whether the trays are covered or not. Uncovered trayswill have a water temperature significantly below the air temperature due to evaporative cooling. Traysplaced higher in a room will often be at temperatures significantly above that of lower ones.Diets and preparationDiet can be provided as a water slurry or dry powder. The former offers ease of measuring while the latteris simple and places the diet – at least initially - at the surface where much feeding occurs. Any powderedfood can be provided either way, but if a slurry is fed, ensure that it is well suspended before eachfeeding.0.02% w/v baker’s or brewer’s yeast (final concentration) is perfect for the day of hatching and 48 hourspost-hatching (see Hatching Anopheles Eggs, Chapter 2). These are often inexpensive and readilyavailable. Their high nutritional content and size make them a good choice.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.6 Anopheles Larval CulturePage 2 of 6For later stages, switch to either a powdered larval food or a slurry consisting of larger particles. Differenttypes of food may superficially appear suitable for larval feeding, but to ensure high quality, measuresurvival from egg hatch to eclosion at least once with any new food source before routine use. Larvalculture affects adult longevity and fecundity in the long term (see Modifying Fecundity, Longevity, andSize), so using the best larval food available will save you time in the future.One well-tested larval food is finely ground Koi Staple Diet from Drs. Foster and Smith, though TetraMinflake food is widely used alternative. For large scale production where cost may be a greaterconsideration, inexpensive and readily available diets such as Farex baby food, hog chow, dog chow, andliver powder have also been used. Since Anopheles feed primarily at the water surface (see Behavior andPhysiology of Anophelines in the Laboratory), food that floats on the water surface is ideal. Grinding foodand sifting for a small size can ensure the food will be of a small enough particle-size for the larvae toingest.Koi pellets and similar pellet and flake diets can be prepared in a grinding mill or blender and siftedthrough a 250 micron sieve (Figure 2.4.6.1). When fed as a powder, this will remain temporarily on thesurface. Such finely ground food is suitable for feeding from L2 to pupation. If you sift food, the largerparticles of food that did not go through the sieve can be saved for feeding L4s or more tolerant strainssuch as Aedes. Larval food should be stored at -20 o C until ready for use if possible to prevent microbialgrowth.When feeding, disperse dry food evenly across the top of the water (Figure 2.4.6.2). It may be dispensedfrom a salt shaker, some other simple improvised device or shaken from a tiny weighing spoon.Figure 2.4.6.1. Preparing food. Use a generallaboratory grinder (pictured here), a grinding mill ora household coffee grinder. To ensure food is smallenough for the earliest stages, use a sieve withholes no larger than 250 microns in size.Figure 2.4.6.2. Check pans daily and assess forfeeding and density. Adjust the density by splittingor thinning to about 200-300 larvae in a typical tray.DensityIt has been shown experimentally that eclosion rates diminish as density in the pan increases(Timmermann and Briegel 1993). High larval density has also found to distort sex ratios by favoring malesover females in An. stephensi (Reisen and Emory 1977). Larval vigor irreversibly restricts adult health soattention to this factor cannot be over-emphasized. Larval crowding stresses larvae and is implicated insome larval infections.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.6 Anopheles Larval CulturePage 3 of 6A reasonable density for most L3-4 anophelines is 1 larva per ml with the water level 0.5-1 cm in depth.When it is impractical to estimate the exact density in the early (L1-2) stages, larvae are usually culturedat a high density. For this reason, the rearing schedule (Chapter 1) is designed for thinning progressivelyin stages. Some species such as An. quadrimaculatus and An. freeborni may culture more successfully at ahigher density in the early instars than is typical for An. gambiae.FeedingAt a constant temperature and given an appropriate amount of diet, the time from hatch to pupationshould be predictable within one day from generation to generation. If pupation is delayed more than aday or two, any of the following could be responsible: temperature is too low, inadequate food was givenat some stage, the density was too high, or excessive food was given in the early stages. Poor cultureresults in disparate developmental rates leading to pupation over the course of several days. This resultsin a great deal of extra work for the technicians. Ideally, almost all pupae form over 2 to 3 days.Examine the pans daily (even when you do not feed) to ensure that the larvae are developing asexpected and the density is appropriate. If you notice great differences in sizes of larvae between orwithin one tray of a cohort at any time, you likely have them too crowded and/or they are underfed. Notsurprisingly, the amount of food provided daily must increase as the larvae develop and as their densityincreases. However, there is a limit to the amount of food and larvae you can place in one tray, so werecommend adhering initially to the density guidelines above and modifying only the amount of diet.Underfed trays will contain larvae that die, are slow-growing or are variable in their development rate. Inextreme cases, unusually long fecal pellets will be observed due, presumably, to re-ingestion of feces.L4s that are overcrowded and/or underfed will be small and have little fat body accumulation.Overfeeding is common and is indicated by numerous observations that precede larval death.1. Foul smell. If you smell a foul odor when you remove the cover, you’re feeding too much. A healthyorganic odor is normal. However, what is considered healthy is admittedly dependent on personalaesthetics!.2. Excessive turbidity. Yellowish to greenish-colored water is fine and often appears in later stages ofrearing (referred to as gelbstoff). However, if the water is turbid, feed less or not at all until the waterclarifies. If turbidity persists, filtering the larvae out from the old culture water and at least a partialwater change may be necessary. Greater turbidity is tolerable during the L3 and L4 stages whereasL1s and L2s are more sensitive. You will develop judgment regarding how much turbidity isappropriate.3. Excessive surfactants. When the water in the pan is agitated, bubbles that form should burst rapidly.If they persist, bacterioneuston has formed an excessive surface microlayer which is not healthy forAnopheles larvae. Check for these by sloshing the water gently. Larvae exposed to water with highlevels of surfactants often do not survive and re-feeding of the adult stock may be necessary. Ifbubbles persist, filtering the larvae out from the old culture water or dragging a tissue over the surfaceand at least a partial water change are recommended. If this is observed routinely, the cultureconditions must be changed.Overfeeding and microbes in the larval culture can cause irregular scum to form on the bottom of thetrays. These pans should be scrubbed with a detergent, rinsed thoroughly and autoclaved or bleached. Itis important to remember that in almost all cases, a thin layer will form on the bottom of the pan fromwaste materials and settled diet. It is a problem only when the layer becomes gelatinous or jelly-like inappearance, irregular in its reflectance, or contains bubbles. Change the container and reduce thefeeding rate.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.6 Anopheles Larval CulturePage 4 of 6More signs of poor larval health related to density and feeding ratesThe slowest -growing larvae and those cultured in turbid water often develop melanic nodules in theabdomen and thorax or black patches on the cuticle. These are both bad signs, and the individuals thathave these should be discarded - they seldom survive. See the section on minimizing infections forphotos of infected larvae. Sub-optimal larval rearing conditions can also result in missing setae or thosethat are covered with black film (probably fungus). This can often be observed in the slowest-growinglarvae even under good conditions, but if the condition is prevalent, a change in your methods iswarranted.When food is overly abundant, Vorticella reproduce to such an extent that they cover the larvae and givethem a fuzzy appearance. This is sometimes evident even without microscopic examination.Pupae that are not curled into the typical ‘comma’ shape but have a horizontally extended abdomen willnot emerge. If you observe this among the first-forming pupae, it may not be too late to rescue theremaining larvae by changing the culture conditions.The metamorphic transitions are the most sensitive stages to the effects of poor larval health. This can beobserved as failure of larva to pupate or pupae to emerge as adults. One should observe >95% of adultshave emerged from pupa cups under good conditions. The effects of poor larval/pupal conditions areoften evident in a short adult life span, and males are especially sensitive to this effect.Day Stage Amount of diet1 Hatching L1 60 mg yeast2 L1/L2 No additional diet needed3 L2 60 mg L3/4 diet4 L2/3 No additional diet needed5 L3 120 mg L3/4 diet6 L3/4 120-300 mg L3/4 diet7 L4 120-300 mg L3/4 diet8 L4/pupae 120-300 mg L3/4 dietTable 2.4.6.1. Approximate food amounts to be fed to the L1-L4 larvae per day assuming L3 and L4densities of 1 larva/ml. The table assumes younger larvae are at a higher density – at least 2 X.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.6 Anopheles Larval CulturePage 5 of 6Figure 2.4.6.3. An example of a density reference photo of 50 (left) and 100 (right) L4 An. albimanus.Creating a photograph of the correct density in the pan size that you use may be helpful for technicians inroutinely estimating a good density by eye.ReferencesReisen WK, Emory RW (1977) Intraspecific competition in Anopheles stephensi (Diptera Culicidae) II.The effects of more crowded densities and the addition of antibiotics. Can. Ent. 109:1475-1480Timmermann SE, Briegel H (1993) Water depth and larval density affect development and accumulationof reserves in laboratory populations of mosquitoes. Bull. Soc. Vector Ecol. 18:174-187

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Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.7 Separating Larvae and PupaePage 1 of 42.4.7 Separating Larvae and PupaeMR4 StaffIntroductionFour features used alone or in combination to separate pupa from larva are: 1) buoyancy 2) size 3)activity and 4) appearance. Pupa of most species should be separated from larva daily; otherwise, adultswill emerge. Some species require a longer pupal development and can be separated every other day.There are several larva/pupa separation methods that use equipment varying in sophistication. All will notwork equally well for all species and must be tested. While pipetting will select pupae of all species basedon appearance, the most uniformly effective en masse method for all species is size separation using aglass plate pupa separator (Fay and Morlan 1959). Another device based on the size differences is that ofMcCray (1961) but this does not allow on-the-fly adjustment as readily as does the Fay and Morlandevice.Separate by appearanceThe simplest, but most time consuming method involves using a “pupa picker” to individually manuallyremove pupae from the rearing pan and place them into an emergence cup. This method is appropriateonly when small numbers of pupae are present. Examples of some utensils are shown in Figure 2.4.7.1.Figure 2.4.7.1. Examples of utensilsuseful for manual pupae separation or“hand picking”. On the left is a piece ofwire mesh glued to a metal spatula and isused to scoop pupae. The middleimplement is made of a 1,000 µldisposable pipette tip, the tip of which hasbeen trimmed off and fitted with a 2 mlpipette bulb. The pupa picker on the rightis a disposable pipette especially good foravoiding contamination as it istransparent. Rinsing these in hot wateroften is necessary to preventcontamination.Separation by differential buoyancy of pupae and larvaeLarvae are negatively buoyant whereas pupae are positively buoyant. Pupal and larval activitycounteracts this useful difference, but chilled water can be used to reduce this.SwirlingOne quick method to try is swirling larvae and pupae a cup: larvae will accumulate in the middle on thebottom and the pupae at the sides of the cup. Using a pipette such as one shown in Figure 2.4.7.1,remove the larvae from the middle or the pupae from the sides.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.7 Separating Larvae and PupaePage 2 of 4Swirling with chilled waterA variation is to use ice water to stun the mosquitoes allowing them to be separated in water without theirbehavior interfering (Weathersby 1963).1. Collect the pupae and larvae together in a strainer and wash them into a small bowl (Figure 2.4.7.2).2. Add ice-cold water free of ice (Figure 2.4.7.3).3. Quickly swirl and wait until separation occurs (Figure 2.4.7.4-5).4. Gently pour the pupae into a sieve and transfer to emergence cups containing culture temperaturewater.NotesA separatory funnel or Imhoff cone can also be used with the ice water method (Figure 2.4.7.6) (Hazard1967). This is a good method for large numbers of a single strain.Ice water increases larval mortality in some strains/species so the time of ice water exposure should beminimized. Larvae typically only survive this treatment 2 or 3 times, so this is most useful when largenumbers are cultured and picking is limited to 2 to 3 days.Carbon dioxide anesthesia has also been used for the purpose of immobilizing pupae and larvae forseparation, though we have not tried it (Lin and Georghiou 1976). This method could be similar to icewater in its usefulness.Figure 2.4.7.2. Place pupae andlarvae in a small volume of water.Figure 2.4.7.3. Pour in ice waterleaving enough room for swirlingand pouring.Figure 2.4.7.4. Swirl gently andthen allow separating. Larva willaccumulate in the bottom middle ofthe cup and pupae will float at thetop and around the sides.Figure 2.4.7.5. Side view oflarval/pupal separation in ice water.Figure 2.4.7.6.Chilled watermethod as shownto left but usingan Imhoff cone.Larvae are on thebottom andpupae at the topof the funnel.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.7 Separating Larvae and PupaePage 3 of 4Separation by larger size of pupaeFay-Morlan SeparatorA custom-built glass-plate pupa separator (Figure2.4.7.7) is good for large numbers of one strain and hasminimal harmful effects. Two roughly parallel glassplates are adjusted with knobs to control the spacing ofthe upper and lower portions of the glass. By controllingthe space you can trap the pupae (larger diameter) andallow the larvae (smaller) to flow through into a tray.Then the glass plates are loosened and the pupaeflushed into a separate pan. This method can also beused to separate the sexes of pupae when male andfemale size differs considerably though it is not usuallypossible with Anopheles mosquitoes since both sexesare roughly the same size in the pupal stage (seeChapter 2, Physiology section). Care must be taken notto overload the device or it will become congested withpupae and the flow of larvae will be prevented. The riskof contamination is the main drawback to this methodas larvae and pupae may get caught on the edges andbe inadvertently transferred to another stock. Rinsingthoroughly between strains can prevent such transfer.Figure 2.4.7.7. Custom-built pupa separatorrelying on the size difference between pupaand larva for separation (produced by theJohn Hock Company).Figure 2.4.7.8. The McCray gateseparator. The closest gate tothe viewer has a narrower gapthan the farthest gate; this is setup to separate dimorphic pupaeand larvae.Gate SeparatorA smaller machine is the McCray Separator (1961). The separatorshown utilizes a series of pre-cut aluminum plates that can beadjusted to catch pupae of varying sizes; however these cannot beadjusted during use - only between (Figure 2.4.7.8). It is especiallygood for separating pupae that have distinct size dimorphism suchas Ae. aegypti. The contents of the larval rearing pans are pouredinto the upper chamber of the sluice where the plate gap is set tocatch the largest pupae first. By applying a gentle stream of water,the smaller size pupae and larvae are washed down to the lowerchamber which is set at a slightly narrower gap to catch any smallerpupae. From here, the larvae are flushed down into a new rearingpan where they can be fed and left to pupate. After all the larvaehave been rinsed through, remove the aluminum dividers and rinsepupae into a new pan and transfer to a cage for emergence. As withthe Fay-Morlan Separator, care must be made to ensure that all thepupae have been washed from the apparatus to avoid contaminationwhen handling several different stocks.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.7 Separating Larvae and PupaePage 4 of 4Gentle vortex: Separate by both activity and buoyancyThe vortex method not using chilled water ranges from greatto poor depending on the mosquito strain. A variation is tocollect pupae and larvae in a strainer and washing them intoa Florence flask with fresh water (Figure 2.4.7.9). Swirl asyou fill to the top. Most of the larvae will dive and most of thepupae will rise to the neck. When separation is maximal,pour the pupae off. This method is seldom 100% effective,so some hand-picking is required. This can also work withan Imhoff funnel (as shown in Figure 2.4.7.6) in which casethe larvae are drained from the bottom. Cool water willimprove the separation, but take care that repeated chillingis avoided as much as possible to prevent mortality.Figure 2.4.7.9. An. gambiae larvae andpupae in the process of separation basedon activity and buoyancy. Pupae arefloating to the surface.ReferencesFay RW, Morlan HB (1959) A mechanical device for separating the developmental stages, sexes, andspecies of mosquitoes. Mosq News 19 144-147Hazard EI (1967) Modification of the ice water method for harvesting Anopheles and Culex pupae. Mosq.News 27 115-116Lin CS, Georghiou GP (1976) Tolerance of mosquito larvae and pupae to carbon dioxide anesthesia.Mosq News 36:460-461McCray EM (1961) A mechanical device for the rapid sexing of Aedes aegypti pupae. J Econ Entomol 54819Weathersby AB (1963) Harvesting mosquito pupae with cold water. Mosqss News 23:249 -251

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.8 Anopheles Adult CagingPage 1 of 42.4.8 Anopheles Adult CagingMR4 StaffIntroductionCages can be quite simple and are often improvised from easily obtained materials. The type of cage youwill use for adults will depend on amount of mosquitoes, availability of cage material, security requiredand behavioral constraints of the stock. The size of your cage must be large enough to promote or allowmating. Following are some caging options.Before purchasing or building cages, consider the number of adults that will be housed, ease of cleaning,and whether you prefer to dispose of them or reuse them. Purpose-built improvised cages are often asuseful as commercial options.Paper cansCylindrical paper containers are popular in insectary usage as they come in different sizes, areinexpensive and are disposable (Figure 2.4.8.1). A relatively small number (e.g. about 10-50 females andmales) of An. gambiae, An. stephensi, and An. albimanus adults will mate well in pint (~1/2 liter) papercups but can hold up to 100 adults without crowding, so this is a good container for many genetic crossesand small stocks. For a colony of 500-1,000 adults, a 3.8 liter (1 gallon) size container is moreappropriate.You can create a hole in the screen plugged with cotton for introducing mosquitoes or use tube-gauzestapled into a hole cut in the side to permit the introduction of cups for pupae or egg collection (Figure2.4.8.1). Using mesh netting on the top allows blood- and sugar feeding. Because of dripping blood andsugar water, paper containers will grow fungi in the bottom and must be disposed after use.Figure 2.4.8.1. Paper cartons of two sizes, pint(left) gallon (right), showing top entry (left) andside entry (right) solutions.Figure 2.4.8.2. Left: Small vial perfect forsingle pair matings or single female oviposition.Right: Single female placed in small vial withwater and egging paper for oviposition.Single-pair matings work well in small (approx 120 ml) plastic vials (Qorpak No. 3891, 6.75 cm deep, 4.5cm diameter) such as the one shown in Figure 2.4.8.2. It has been modified by cutting a hole in the topand covering with mesh for easy feeding and aspirator entry. Using small containers for single pair matingcan delay mating from the typical 1 or 2 days after emergence to 6-7 days. These small vials are alsouseful for collecting eggs from individual females. To egg with this container, simply fill to about 1 cmdepth with water and line with two filter paper strips that extend half way under water (Figure 2.4.8.2).

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.8 Anopheles Adult CagingPage 2 of 4Metal-frame cagesMetal cages are good alternatives since they are autoclavable and therefore easily sterilized andreusable. One metal cage developed by Savage and Lowe (1971) consists of a single-piece bent-toshapealuminum sheet covered with tube-gauze (Medical Action Industries White T-1 Tubegauze ®#58205) (Figure 2.4.8.3). This cage is appropriate for small colonies of up to 500 adults if the colonymates well in a cage of this size. Removing the mesh netting between uses and autoclaving the metalportions make this a great choice for avoiding infection in colonies. The netting can be cleaned andreused or discarded.Another example of a larger design is shown in Figure 2.4.8.4 (BioQuip, www.bioquip.com) and worksbetter for colonies of the same number but that need more space for mating. It does require cleaningbetween uses. With either of these choices, introducing cups and bloodfeeding is easy.Figure 2.4.8.3. Savage cage, perfect for smallcolonies that mate well in small spaces.Figure 2.4.8.4. Autoclavable cage especiallyuseful for colonies that mate poorly andlarger numbers.The best cage solutions have plenty of resting space for the mosquito. Though having such space doesnot seem to be important for mating, it is preferred by some species and therefore assumed important for‘comfort’. With An. arabiensis mosquitoes, if a resting space is offered, usually more than 90% of themosquitoes will be inside that space (assuming it is large enough) at daylight times, see Figure 2.4.8.6.The use of red boxes has been published as preferred resting sites over some alternative trappingdevices in anophelines (Goodwin 1942). Thus, if mosquitoes choose red resting boxes in the wild, theywill most likely work well inside your cages. Red, most notably, and some shades of blue have beenshown to be preferred to black or other colors for Anopheles resting (Nuttall and Shipley 1902) and aretherefore good color choices.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.8 Anopheles Adult CagingPage 3 of 4Figure 2.4.8.6. An. arabiensis adults crowding into a single resting tube in daylight hours.ReferencesGoodwin MN (1942) Studies on artificial resting places of Anopheles quadrimaculatus Say. In: EmoryUniv. Field Sta., Newton, GANuttall GHF, Shipley AM (1902) Studies in relation to malaria. The structure and biology of Anopheles.Amer. Journal Hyg. 2:58-84Savage KE, Lowe RE (1971) A one-piece aluminum cage designed for adult mosquitoes. Mosq. News.31:111-112

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Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.9 Anopheles Adult DietPage 1 of 22.4.9 Anopheles Adult DietMR4 StaffIntroduction6% glucose, 10% v/v corn syrup, 10% w/v sucrose, and dilute honey are common sugar sources for adultmale and female mosquitoes. These can be provided using soaked cotton balls lying on top of a cage ifthe mesh is non-absorbent (e.g. nylon rather than cotton). This requires ensuring that the cotton stays wetenough for the mosquitoes to drink the sugar. Sugar pads normally need to be changed no less thanevery week because mold spores and fungi grow well on exposed sugar pads. 0.2% methylparabenadded to a sugar source can extend the time before the sugar source begins to collect mold spores thatare harmful to mosquitoes without causing early mortality or reduced longevity (Benedict 2009). In thiscase, sugar sources can be left unchanged for 30 days as long as they stay wet.Cleanliness of sugar water and cotton balls is of the utmost importance.After a bag of cotton balls is opened, mold spores can settle on themimmediately. Cotton balls can be sterilized by autoclaving and should bestored in sealed containers.One way to avoid having to moisten sugar pads is to use a hangingfeeder that holds soaked cotton (Figure 2.4.9.1). This example consistsof an inverted sample vial (Fisherbrand Polystyrene Sample Vial (20ml)Cat. No. 03-341-13) hung in the cage. A hole has been punched in thecap (at the bottom) to allow mosquitoes access to the soaked sugar pad.The vial hangs from a bent wire inserted into the vial base by heating thewire and forcing it through the plastic. Two large cotton balls are soakedwith sterilized sugar solution and then placed in the vial. This orientationhas the advantage that Anopheles will seldom lay eggs in it, and it willremain moist for 1 week without attention. If using such a source, thesugar vials and covers should be bleached between use and stored in aclosed container to prevent mold spores from accumulating on them.Figure 2.4.9.1. Invertedhanging sugar vial.Another alternative is to fit theopening in the cap (as shown inFigure 2.4.9.2) of the invertedfeeder with a piece of porousplastic large enough for the mosquitoes to feed through but smallenough that water does not drip. The porous plastic center in thephotograph to the left was obtained from Small Parts Inc., catalogno. SPE-040-20. This modification allows for pouring the sugarwater directly into the feeder with no need for cotton. Using thishanger in combination with sugar water containing methylparabensolution should require no attention for about 30 days.Figure 2.4.9.2. Cap of feeder witha porous plastic insert.ReferencesBenedict MQ, Hood-Nowotny RC, Howell PI, Wilkins EE (2009) Methylparaben in Anopheles gambiae s.l.sugar meals increases longevity and malaria oocyst abundance but is not a preferred diet. J. Insect Phys.55: 197-204.

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Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.10 Bloodfeeding : Membrane Apparatuses and AnimalsPage 1 of 62.4.10 Bloodfeeding : Membrane Apparatuses and AnimalsMR4 StaffBlood Feeding Strategies:Blood-feeding is one of the most problematic activities of anopheline culture. Even when ethical rulespermit the use of a live animal, their care and housing are expensive and time-consuming. In spite of this,many scientists have concluded that there is no equal to a live animal regarding attraction, satiation, andegg production. There are many reasons why animals are employed as a blood source: animals provide aconstant source of blood which is at the proper temperature, the animal provides necessary stimuli forfeeding, and it is not necessary to handle blood directly. However, these must be weighed against thecons which include the potential for animal biting, accidental disease transmission, and the procurement,maintenance, and care of animals.There has been one reported success of using an artificial blood meal to produce eggs in Aedes;however, it has never been reported successful in anophelines (Kogan 1990).One full blood meal is enough to mature oocysts in most well-cultured anophelines. The length of feedingtime needed to ensure females are fed to repletion or the type of blood necessary will depend on themosquito you are using. Usually 3-5 day old females feed most readily. However, some colonies of An.atroparvus and An. minimus feed more avidly at 6-9 days.Keeping in mind the amount of blood that a single animal can safely provide, it is best to maximize thenumber of cages you are feeding at one time to reduce the amount of total time the animal is restrainedand/or anesthetized. Also, feed no more frequently nor more mosquitoes than are necessary to generatethe number of progeny you need.In some instances a mosquito colony may not feed on the animal provided. Sugar-starving females for 5-12 hours prior to blood feeding can increase success rates in some hesitant strains. However, removingsugar jeopardizes males and should be used only when necessary or on separated females. If starvationdoes not help bloodfeeding rates and the females are of optimum age, then there may be other factorsaffecting the feeding rate:A new species of animal. Mosquitoes can be selected to feed on one animal source by propagating thecolony only with those mosquitoes that fed a particular species. It may take a few generations before themosquitoes readily accept a new host. In the effort to increase feeding, increase the feeding time and thefrequency. Supplement with another blood source if critical to maintenance of the colony.Infection in the colony. Mosquitoes infected with fungi (Scholte et al. 2006) may not respond to a host orbe unable to imbibe properly. Some infections will stall the digestion process resulting in a reduction inegg production. See Chapter 2 for information about preventing infections.Improper feeding time: This is important in both newly acquired strains and material brought from thefield. The new mosquito colony may be accustomed to feeding at a specific time or under certainconditions. Wild mosquitoes feed in darkness, so lighting is a factor that should be considered. With anew or wild colony, try to determine when the mosquitoes are actively searching for a blood meal andschedule feeds for that time if possible. If they prefer to feed at night, cover the cages with a dark shroudfor several hours before feeding and turn out the lights while offering the bloodmeal. Also, the length oftime you bloodfeed may be too short. Some colonies require time near the blood source before they willtake a meal.Use of Live AnimalsColonizing mosquitoes involves locating a source of blood for the propagation of the colony. Live animalswere utilized more often than not in the past; however, their use has diminished due to theimplementation of strict guidelines governing the use of animals in a research setting. Likewise,

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.10 Bloodfeeding : Membrane Apparatuses and AnimalsPage 2 of 6Institutional Review Boards (IRBs) disallow any use of live animals when deemed unnecessary or if analternate, less intrusive method for feeding is known. It is best to check with your local IRB beforeconsidering blood sources to determine what blood sources and conditions are allowed. If you decide touse live animals, obtaining an approved protocol from a colleague in your country is a good start. Thefollowing are a good list of sources for rules and regulations for the U.S. to consider when writing aprotocol.The Public Health Services Policy on humane care and use of laboratory animals is provided by theNIH as a resource in caring for animals in a research setting.http://grants.nih.gov/grants/olaw/references/phspol.htmThe Animal Welfare Act sets rules for how animals are to be used.http://www.nalusdAn.gov/awic/legislat/awAn.htmThe Guide for the Care and Use of Laboratory Animals gives standard criteria used by the PublicHealth Services (PHS) in the production of the PHS Policy.http://www.nap.edu/readingroom/books/labrats/chaps.htmlThe Association for Assessment and Accreditation of Laboratory Animal Care International(AAALAC) is a body that accredits local institutions if the facilities meet the standardized requirementsabove. http://www.aaalac.org/The World Organization for Animal Health (OIE) is an international agency that also sets standards forthe use of animals in research. http://www.oie.int/eng/en_index.htmInstitutional Animal Care and Use Committee (IACUC) is another resource for the use of any animal ina laboratory setting. This group also has some links to other government resources as well as anextensive online library. http://www.iacuc.org/Institute for Laboratory Animal Research (ILAR) provides information and publications on the use andcare of several animals common to laboratory settings. http://dels.nas.edu/ilar_n/ilarhome/There are a few types of animals which are more commonly used in a research setting. They are chosenfor many reasons and are all considered small animals which can be easily procured.Lagomorphs-rabbits (Figure 2.4.10.1)Rodents-mice, rats, and guinea pigsChickensBirds are a special case and there are additional resources for their use located at the websites below:http://www.nmnh.si.edu/BIRDNET/GuideToUse/index.htmlhttp://www.mbc.edu/osp/docs/IACUC_Proposal_Sheet.dochttp://www.fiu.edu/~dsrt/animal/birds_lab.htmMedical schools may have research animals that they plan to destroy after their experiments have ended.This can be a good source for single use feeding (i.e. the animal will be euthanized after feeding). Avoidusing any animals that were infected with any pathogen if you use animals from a medical school.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.10 Bloodfeeding : Membrane Apparatuses and AnimalsPage 3 of 6Figure 2.4.10.1. Use of arabbit for mosquito feedingutilizing a combination ofdrugs for anesthesia andpain under an IRBapprovedprotocol.Restraining the animalensures uninterruptedfeeding for the mosquitoesand safety for the rabbitshould it awakenunexpectedly. Shaving thebelly prior to feedingmakes the skin easilyaccessible.Membrane feedingDue in part to the restrictions on the use and difficulty of live animals in a research setting, artificialmembrane methods for feeding have been developed. Designs all have two basic features: a heatingelement and a membrane to hold the blood. The heating element is necessary to maintain the blood at areasonable temperature at which mosquitoes will imbibe, typically between 35- 40°C. The membraneshould be relatively thin so mosquitoes can pierce it easily but sturdy so that spilling does not occur.Membrane typesThere are several materials that have been used in membrane feeding. Natural membranes are claimedto work best (e.g. animal skin membranes), but they are can be difficult to procure and can usually onlybe used once (Novak et al. 1991). Other membranes used with some success include: Parafilm M®,collagen sheets, latex membranes e.g. gloves or dental dam, sausage casings (natural or synthetic),Baudruche membranes (Joseph Long Inc, N.J.), and condoms (lambskin or latex).Membrane FeedersSeveral different methods have been developed for feeding mosquitoes, some simple, some are morecomplex. A short list of some reported examples from the literature are:Latex or lambskin condoms: Fill the condom with heparinized blood. Tie it closed and soak it in a warmwater bath (~40°C) for 30 minutes to 1 hour to fully heat the blood. Condoms have been reported to havevarying levels of success.Tseng feeder (Tseng 2003): This employs a Parafilm packet filled with blood then wrapped in a wire meshcasing. It is best to heat the blood before pouring it into the Parafilm packet.Mishra feeder (Mishra et al. 2005): A Petri dish is wrapped with Parafilm on the outside leaving a smallpocket in the center of the bottom. Warm water is poured into the Petri dish and a small amount of bloodis injected into the Parafilm. This unit can be placed directly on the cage.Mourya feeder (Mourya et al. 2000): A 10 mm hole is drilled into a sheet of acrylic. Parafilm is thenstretched over the sheet and warmed blood added into the 10 mm hole. A beaker of warm water placedon the top of the hole maintains the temperature.More Complex Designs for Membrane FeedersGlass feeders: Although there are several models of these, most are built in the same manner: An outerarea contains circulating warm water and an inner chamber where the blood is poured, see Figure2.4.10.2. A nice reference to several styles is given by Kasap et al. (2003).

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.10 Bloodfeeding : Membrane Apparatuses and AnimalsPage 4 of 6Figure 2.4.10.2. Example of a membrane feeder using dental dam and an electric pump andheater to circulate warm water.Electric units: These models e.g. Hemotek (Figures 2.4.10.3-2.4.10.5) employ an electric heatingelement to maintain the temperature of the blood meal. These are very reliable and easy to maintain(Hagen and Grunewald 1990; Cosgrove et al. 1994).Figure 2.4.10.3. The Hemotek systemoffered by Discovery Workshops(hemotek@discoveryworkshops.co.uk)Figure 2.4.10.4. The electric heating unit with amembrane feeder attached to the bottom.

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.10 Bloodfeeding : Membrane Apparatuses and AnimalsPage 5 of 6Figure 2.4.10.5. The components that make up themembrane feeder. Clockwise from top, a membrane(in this case Parafilm; the manufacturer’smembranes are recommended), a completelyassembled membrane unit ready to be attached to theheating unit, the metal membrane unit without amembrane, and the “O” ring used to attach themembrane.Switching from a live animal to a membrane system can be difficult if a colony is accustomed to feedingon a live animal. There are some methods commonly employed to increase success rates when using amembrane feeder.Starvation: sugar-starving females 5-12 hours can be helpful in increasing desire to take a blood meal;sugar-starving can cause mortality in males and should be limited if the males are not first removed.Lighting: most successful membrane feeds are conducted in the dark; you can either cover the cagesbeing fed or turn off the lights.Stimuli: gently rubbing the membrane on your arm will transfer human volatiles needed for phagostimulationin mosquitoes.ATP: adding this chemical to the blood has been shown to improve membrane feeding with some strainsof Aedes but not Anopheles.Blood sourcesA reliable source of blood for membrane feeding must be found. Although blood can be drawn from ahuman volunteer or from a laboratory animal, the use of either type of blood may require specialpermission through an IRB or an IACUC (Institutional Animal Care and Use Committee). This typicallyinvolves a review as well as a trained specialist to perform the venipuncture procedure.Blood can sometimes be obtained from a slaughter house (an abattoir). If properly treated (eithercollected in a heparinized vial or provided some anti-coagulant agent and stored at 4°C), these sources ofblood have been used successfully for up to 2 months. Heparinized or citrinated animal blood can also beprocured from specialized vendors such as Lampire, HemoStat, and Bioreclamation.Personal safety while using a membrane feeding apparatus is critical since all blood should be handledas if it were infected. Proper training in the use of Personal Protective Equipment (PPE) such as latexgloves, lab coat, and face-shield should be provided to all personnel handling the blood or performing thefeedings. Personnel should receive appropriate instructions according to their institutions rules.ReferencesCosgrove JB, Wood RJ, Petric D, Evans DT, Abbott RH (1994) A convenient mosquito membrane feedingsystem. J Am Mosq Control Assoc 10:434-436

Chapter 2 : Anopheles Laboratory Biology and Culture2.4 Anopheles Culture2.4.10 Bloodfeeding : Membrane Apparatuses and AnimalsPage 6 of 6Hagen HE, Grunewald J (1990) Routine blood-feeding of Aedes aegypti via a new membrane. J AmMosq Control Assoc 6:535-536Kasap H, Alptekin D, Kasap M, Guzel AI, Luleyap U (2003) Artificial bloodfeeding of Anopheles sacharovion a membrane apparatus. J Am Mosq Control Assoc 19:367-370Kogan PH (1990) Substitute blood meal for investigating and maintaining Aedes aegypti (Diptera:Culicidae). J Med Entomol 27:709-712Mishra K, Kumar Raj D, Hazra RK, Dash AP (2005) A simple, artificial-membrane feeding method for theradio-isotope labelling of Aedes aegypti polypeptides in vivo. Ann Trop Med Parasitol 99:803-806Mourya DT, Gokhale MD, Barde PV, Padbidri VS (2000) A simple artificial membrane-feeding method formosquitoes. Trans R Soc Trop Med Hyg 94:460Novak MG, Berry WJ, Rowley WA (1991) Comparison of four membranes for artificially bloodfeedingmosquitoes. J Am Mosq Control Assoc 7:327-329Scholte EJ, Knols BG, Takken W (2006) Infection of the malaria mosquito Anopheles gambiae with theentomopathogenic fungus Metarhizium anisopliae reduces blood feeding and fecundity. J Invertebr Pathol91:43-49Tseng M (2003) A simple parafilm M-based method for blood-feeding Aedes aegypti and Aedesalbopictus (Diptera: Culicidae). J Med Entomol 40:588-589

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 1 of 82.5 Basic Anopheles Mendelian GeneticsMark BenedictIntroductionIt is difficult to culture stocks on a long-term basis without some knowledge of their genetics. After all,what ultimately distinguishes stocks and species is their genetic constitution, not the name that we assignto them. The following are aspects of Anopheles genetics that are very relevant to understanding the dayto-daystability and integrity of stocks. Exceptions can be applied to every generalization made here.However, the following will ground you in Anopheles genetics.As a basis for the modern discussions of anopheline phylogenetics and genomics, the existing reviewsare still very useful (Kitzmiller and Mason 1967; Kitzmiller 1976).In this chapter, we provide the minimum background to understand anopheline genetics and exercisesusing anopheline characters to help make the information relevant. Answers are provided after thequestions. If you have no understanding of Mendelian genetics, study the basic genetic principles ofdiploid organisms in a basic text before proceeding. You should particularly understand Punnett squaresand basic probability.Genetic GlossaryAllele: Variant forms of genesCoupling: Also called ‘cis.’ Two alleles being referred to are on the same homolog. In contrast, see‘repulsion..Diploid: Having two sets of chromosomes.Expressivity: The degree or Extent of expression of the phenotype. This addresses the issue e.g. not ofwhat proportion of individuals have freckles, but how many or how large the freckles are. Or foranophelines not, do they have a red stripe, but how intense and clear is it? In contrast, see ‘penetrance.’Gene: The basic unit determining heritable expression. (There are other kinds of heritable expression, butgene covers 99.9% of them.)Genotype: The allelic constitution of an individual, but sometimes applied to tissues or cells.Haploid: Having one set of chromosomes.Hemizygote: Usually refers to the state of having only one copy of a gene located on the X chromosomein males (which have only one X).Heterozygote: The condition in which a diploid individual has two different alleles of a gene.Homolog: One member of a pair of chromosomes.Homozygote: The condition in which a diploid individual has two apparently identical alleles of a gene.Linkage group: May temporarily be the genes or loci that are experimentally identified as not segregatingindependently from one another, but ultimately refers to the collection of everything located on onechromosome. The number of linkage groups identified experimentally may be greater than the actualnumber of chromosomes. (Think of contigs in a DNA sequence. One usually starts with several, but asthe sequence in the gaps is obtained, multiple contigs coalesce into one.Locus: A general term for a place on the chromosomes. It may be a region, a base-pair, or functionallydefined.

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 2 of 8Paracentric inversion: A rearrangement of the chromosome in which a portion of a homolog is flipped andthe centromere is NOT included in the flipped region. Important because they often occur naturally andareuseful as phylogenetic and population genetic tools.Penetrance: A qualitative (low vs. high) or numerical value that refers to the Proportion of individuals in apopulation that express the phenotype that definitively identifies a particular genotype when observed.For example, in An. gambiae, c+ / c females have a red stripe on the larval dorsum generally. Somethinglike 5% of c+ / c females do not appear to have a red stripe. So one could say that red stripe is about95% penetrant. In contrast, see ‘expressivity.’Pericentric inversion: A rearrangement of the chromosome in which a portion of a homolog is flipped andthe centromere is included in the flipped region. Virtually all inversions of this sort in anophelines areinduced by irradiation.Phenotype: The expressed manifestation of a genotype.Quantitative vs. discrete traits: Traits whose expression varies primarily in degree. For example: plantheight, intensity of flower color, malaria parasite encapsulation. Discrete traits can be classified intoclasses e.g. white eye vs. wild eye, collarless vs. collarless+, ebony vs. ebony+.Repulsion: Also called ‘trans’. Two alleles being referred to are on the opposite homologs. In contrast,see ‘coupling.’Trait: Rough term meaning the same as a ‘character’. (Sounds more scientific than ‘a thingie!’) It is afunctional description of some distinct behavior, form, color etc. Used the same as the way we would useit in common speech.Allelic relationshipsComplete dominant: In a heterozygote, only the dominant allele is expressed, and the recessive allele isnot. For example: c+ is completely dominant over c. Stripe+ is dominant over st in An. albimanus.Codominant: In a heterozygote, the phenotypes associated with both of two alleles present are observed.For example: microsatellite alleles are usually referred to as codominant markers because the repeat sizeof both alleles in a heterozygote can be observed. Another example would be enzyme electromorphs.Epistasis: The phenotype of the expression of one gene eliminates the expression of another gene i.e.the phenotype of gene A prevents expression of gene B. white anophelines have no pigment associatedwith the stripe and collarless genes.Gene interaction: The phenotype that is observed is different from that associated with gene A or gene B,but is a result of their combined effect. For example: An. gambiae that have pink-eye mutations generallyhave white or pink eyes. An. gambiae that have red-eye mutations have red eyes. When they have bothpink-eye and red-eye mutations, the have ‘pumpkin’ colored eyes. Neither gene alone can produce thiseffect.Partially dominant or semi-dominant: In a heterozygote, the dominant allele is expressed to a lesserdegree than in a homozyous dominant individual. For example: A. albimanus ebony heterozygotes areintermediate in darkness to either homozygoteThe Facts of LifeFemale Monogamy & Male PolygamyTo generalize the observations of numerous studies, Anopheles females mate only once. The sperm withwhich the male inseminates the female are stored in the spermatheca and are sufficient to fertilize severalbatches of eggs. Under typical insectary conditions, the sperm in the spermathecae are never depleted.Males, on the other hand, will mate several females if given the opportunity. One male can fertilizeapprox. 6-10 females. But again, males are seldom depleted of sperm in typical insectaries due to thelimited number of virgin females and the presence of competing males. The genetic significance of these

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 3 of 8facts is that progeny from one female can almost always be considered as resulting from the mating ofone male. The significance of single-male mating is that among the progeny from a single female (i.e. a‘family’), at most four alleles will be observed.Chromosome NumberAll anophelines have a haploid number of 3: there are two autosomes (chromosomes not involved in sexdetermination) named chromosome 2 and 3, and sex chromosomes named X or Y. These can beobserved in mitotic spreads of brains and testes (Figures 2.5.1 and 2.5.2). However, salivary glandsand/or ovaries may contain polytenized chromosomes in various species.Sex determinationSex determination appears superficially similar to humansand Drosophila melanogaster: Females are XX and males areXY. The X chromosomes contain a region or regions ofeuchromatin, in which most expressed genes are located,and highly condensed heterochromatin which contains highlyrepetitive DNA and presumably few expressed genes. Theseregions are fairly distinguishable in chromosomepreparations.RecombinationRecombination frequency is roughly proportional to distanceon the chromosomes over most of the genome. Its frequencyis similar in males and females in some species (e.g. An.gambiae, An. albimanus) but is higher in females in others(An. quadrimaculatus). Though observations have been madeof differences between the sexes, there is insufficientknowledge to predict in which species a difference will beobserved.Figure 2.5.1. Female brain metaphasechromosomes of A. gambiae s.s.Effects of sex on expressionSex-limited traitsTraits that are expressed in only one of the sexes. Mostobvious are sexual morphologic characters like ovariandevelopment. To use the same example again, why do onlyfemales express the red stripe since both males and femalescan have the c+ / c genotype? This observation is expressedby saying that red stripe is a sex-limited trait. This should notbe confused with sex-linked traits. It has nothing to do withthe location of the gene.Sex-influenced traitsTraits that can be observed in both sexes, but the kind ordegree of expression is influenced by the sex. For example: Ilied for heuristic purposes. Sometimes red stripe can beobserved in c+ / c males. However it is so faint relative tofemales that depending on where one draws the line betweenFigure 2.5.1. Male testes metaphasechromosomes of A. gambiae s.s. Whilethe Y is of variable size in this species,this stock has a very smallheterochromatic Y.individuals that have a ‘red stripe’ and those that do not, you could say they don’t show red stripe.Regardless, red stripe is so much fainter in males so that even if we do not consider it sex-limited, it isstrongly sex-influenced. Traits of this class may also be determined by autosomal genes.

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 4 of 8Sex-linked traitsExcept for maleness, which is associated with the presence of a Y chromosome, sex-linked traits aredetermined by genes on the X chromosome.Linkage RelationshipsAutosomal linkage or inheritanceProblem 1: Use the Punnett Square to describe the frequency of gametes, genotypes, and phenotypeswhen two collarless (Mason 1967) heterozygotes mate: The “c+” allele of this autosomal gene is fullydominant over “c.” Fill in the genotypes and phenotypes in the blanks. Answers can be found at the endof this section.Female Gametes1a.1b.c+ proportion = …………….c proportion = …………….Male Gametes1c.c+ proportion =…………….1d.c proportion =…………….1e. proportion =………………..1g. proportion =………………..1f. proportion =…………………1h. proportion =……………….Note that among the three genotypic classes of progeny, females and males will occur at equalfrequencies in each.Determine the genotypic frequencies1i. c+ / c ____________1j. c+ / c+ ____________1k. c / c ____________Determine the phenotypic frequencies1l. c+ ____________1m. c ____________

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 5 of 8Sex linkageProblem 2: Again, use a Punnett Square to illustrate the pattern. This example uses the sex-linkedmarker white (w) which is located on the X chromosome (Benedict et al. 1996). The w+ allele is fullydominant over the w allele. The cross is between a heterozygous female and a hemizygous male whocarries the w+ allele. Fill in the pheno- and genotypes including sex.Female Gametes2a. w+ proportion = ……………. 2b. w proportion = …………….2c. w+ proportion =2e. w+ / w+ proportion =2e. w+ / w proportion =Male Gametes…………….2d. Y proportion =…………….……………………Sex = ………2e. w+ Y proportion =…………………Sex = …………………………..Sex = ………2e. w Y proportion =………………..Sex = ………Note that among the progeny, females and males occur at different frequencies in each eye-colorphenotypic class.Determine the genotypic frequencies2i. w+ / w female ____________2j. w / w female ____________2k. w+ / w+ female ____________2l. w / Y (male) ____________2m. w+ / Y (male) ____________Determine the phenotypic frequencies2n. w+ female ____________2o. w female ____________2p. w+ male ____________2q. w male ____________

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 6 of 8Inheritance of two linked autosomal genesProblem 3: In this case, we consider two loci that are linked. We will use the Punnett Square to describethe pattern using collarless and Dieldrin resistance (Rdl) (Davidson 1956; Davidson and Hamon 1962)assuming 15% recombination. Rdl R is fully dominant. The recombinant gametes of either complementaryclass will be formed at equal rates. This example is set up as a typical testcross in which a doubleheterozygousfemale is crossed to a homozygous male that has the recessive alleles for both markers.Female GametesNon-recombinantRecombinant3a. Rdl R c+proportion =3b. Rdl S cproportion =3c. Rdl R cproportion =3d. Rdl S c+proportion =………………………………………………………………Male Gametes3e. Rdl S c proportion =………………3f. Rdl S c proportion =………………3g = …………. 3h = …………. 3i = …………. 3j = ………….3k = …………. 3l = …………. 3m = ………. 3n = ………….Determine the genotypic frequencies.3o. Rdl R c+ / Rdl S c __________ This is one 'parental' class3p. Rdl S c / Rdl S c __________ This is the other 'parental' class3q. Rdl R c / Rdl S c __________ This is one 'recombinant' class3r. Rdl S c+ / Rdl S c __________ This is the other 'recombinant' classDetermine the phenotypic frequencies3s. Rdl R c+ ____________ parental3t. Rdl S c ____________ parental3u. Rdl R c ____________ recombinant3v. Rdl S c+ ____________ recombinantNotice that in a testcross, the genotypic and phenotypic frequencies are identical. The frequency of allgenotypes can be determined conclusively. In contrast, a backcross to one of the parents may or may notbe a testcross.

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 7 of 8ReferencesBenedict MQ, Besansky NJ, Chang H, Mukabayire O, Collins FH (1996) Mutations in the Anophelesgambiae pink-eye and white genes define distinct, tightly linked eye-color loci. J Heredity 87:48-53Davidson G (1956) Insecticide resistance in Anopheles gambiae Giles: a case of simple Mendelianinheritance. Nature 178:861-863Davidson G, Hamon J (1962) A case of dominant dieldrin resistance in Anopheles gambiae Giles. Nature196:1012Kitzmiller JB (1976) Genetics, Cytogenetics, and Evolution of Mosquitoes. In: Caspari E (ed) Advances inGenetics. Academic Press, New York, pp 315-433Kitzmiller JB, Mason GF (1967) Formal Genetics of Anophelines. In: Wright JW, Pal R (eds) Genetics ofInsect Vectors of Disease. Elsevier, Amsterdam, pp 3-15Mason GF (1967) Genetic studies on mutations in species A and B of the Anopheles gambiae complex.Genetical Research, Cambridge 10:205-217

Chapter 2 : Anopheles Biology Laboratory and Culture2.5 Basic Anopheles Mendelian GeneticsPage 8 of 8Problem answers1a-d: all 0.51e-h: all 0.25. This is calculated as the product of the gamete frequencies of that row and column.1i: 0.5 (0.25 + 0.25 since there are two sources of these genotypes)1j: 0.251k: 0.251l: 0.75 (since c+ is fully dominant, this is 0.25 + 0.5)1m: 0.252a-d: all 0.52e and f: both 0.25 and females2g and h: both 0.25 and males2i: 0.252j: 02k: 0.252l: 0.252m: 0.252n: 0.52o:02p:0.252q:0.253a and b: 0.4253c and d: 0.0753e and f: both 0.53g and h: 0.21253i and j: 0.03753k and l: 0.21253m and n: 0.03753o and p: 0.4253q and r: 0.0753s and t: 0.4253u and v: 0.075

Chapter 2 : Anopheles Biology Laboratory and Culture2.6 Basic Anopheles Population GeneticsPage 1 of 62.6 Basic Anopheles Population GeneticsMark BenedictIntroductionMost stocks kept in insectaries are of value because they contain one or more interesting alleles that areeither fixed (pure-breeding) or polymorphic in the populations. On the other hand, some laboratories keepa single wild-type stock, and no other stocks are present in the laboratory. In the latter case, the mainconcern is rarely contamination with a different species, but changes in allele frequency, excessiveinbreeding etc. In that case, the relative fitness of various alleles will determine whether one or another islost or becomes fixed over time.The following information is primarily directed toward the issues of maintaining alleles in polymorphicpopulations, and of the fate of alleles in contaminated stocks. As in the other genetics chapter, wepresent some simple problems that are similar to those encountered in the insectary. These will assistyou in planning crosses, isolating mutations etc. Make plenty of drawings and your own Punnett squaresto figure these out.Before you read this section, you should be very comfortable with the contents of the chapter on the basicMendelian genetics of Anopheles and the background to that chapter.Hardy-Weinberg Equilibrium and the Binomial EquationYou’re probably familiar with the assumptions of Hardy-Weinberg Equilibrium: a hypothetical model thatpredicts the frequency and stability of allele frequencies in populations given certain assumptions.According to its principles, the fundamental dynamics of the fate of alleles introduced into populations canbe predicted by some fairly simple equations and applications of probability. While these predictionsassume a few facts to be true that may not be, for Mendelian traits in laboratory stocks of mosquitoes,these assumptions approximate reality sufficiently that you can predict probable mating, allele, and genoandphenotypic frequencies for planning experiments.Two conclusions flow from their principle (quoted from (Strickberger 1968). As we are using theterminology, it would be more proper to substitute “allele” for “gene” in the following description.):1. Under conditions of random mating in a large population where all genotypes are equally viable, genefrequencies of a particular generation depend upon the gene frequencies of the previous generationand not upon the genotype frequencies.2. The frequencies of different genotypes produced through random mating depend only upon the genefrequencies.The best-known method for determining allele and genotype frequencies in a randomly mating populationis a simple binomial equation (considering only two alleles per gene or locus):p 2 + 2 p q + q 2 which is the same as ( p + q ) 2 and these must equal 1The variables ‘p’ and ‘q’ are the frequencies of either of two alleles where p + q = 1. The frequency of the“ p p “ (homozygous) genotype is simply “ p 2 “ and the frequency of “ q q “ genotype is “ q 2 .” Thefrequency of the heterozygous genotype is “ 2 p q .”

Chapter 2 : Anopheles Biology Laboratory and Culture2.6 Basic Anopheles Population GeneticsPage 2 of 6Problem 1: red eye (r) is recessive to wild type (r+, (Beard et al. 1994). In a cage of randomly matingmosquitoes, you observe 4 out of 100 individuals that have the red eye ( r r ) phenotype.If p = the allelic frequency of r and q = the allelic frequency of r+, what are the allelic frequencies in thispopulation? Substitute the value you know into the binomial equation and solve for the other variable.1a. frequency p = ………………..1b. frequency q = ………………..Using the binomial equation, what are the expected frequencies of r+ r+ and r+ r genotype individuals?1c. Proportion r+ r+ ( i.e. q 2 ) = ………………..1d. Proportion r+ r ( i.e. 2pq ) = ………………..You have now estimated the frequencies of both the alleles and genotypes based on a quickdetermination of the number of red eye individuals. The simple binomial equation allows you to makeuseful conclusions about the population!Since we are assuming that individuals of all genotypes mate with one another randomly with nopreference, we can estimate the probability of various matings by simply multiplying the genotypicfrequencies.Problem 2: From the above cage of adults, you selected only wild eye-color males and virgin females andmated them to one another. You have collected eggs from 21 of these females and obtained 20 familiesof larvae.Among what proportion of families do you expect to observe red-eye progeny? In the table, first circle thematings that will produce red-eye progeny. (Use the Punnett square as an aid to calculate the expectedmating frequencies. There is a little trick here. You have to think!)Proportion Females (hint: these proportions must equal 1)2a. proportion heterozygotes( r+ r )= ………………..2b. proportion homozygotes( r+ r+ )= …………………..Proportion Males2c. proportionheterozygotes ( r+ r )= …………………….2d. proportionhomozygotes ( r+ r+ )=………………………2e. proportion = ……………….. 2f. proportion = ………………..2g. proportion = ……………….. 2h. proportion = ………………..

Chapter 2 : Anopheles Biology Laboratory and Culture2.6 Basic Anopheles Population GeneticsPage 3 of 6Approximately how many families will contain red eye progeny?2i. number of families = …………………You may be wondering, “What is the probability that if I have 20 families I won’t get any that have red-eyeindividuals just by chance?” Simple probability can be used to estimate the probability of getting variousnumbers of families. (Usually, one is interested in knowing how many families you need to isolate toensure you’ll get at least one that you desire; in this case a r+r X r+r mating.)As a heuristic device to determine the value, let’s think of the recurrence of the worst case: What is theprobability of isolating one family and it will NOT have any red-eye individuals? It is simply…1 minus the probability that one DOES contain red eye which = 2j. …………………The probability of isolating 2 families, neither of which contain any red eye progeny is simply theprobability of isolating the one above times the probability of isolating a second whose probability isEQUAL TO isolating the first family. (NOTE that these are independent events.)p X p = p 22k. …………………..So if we take p to the 2 nd power for two families, we take it to the ‘nth’ power for ‘n’ families. So for 20families, the probability of NOT seeing any families containing red eye is simply2l. ……………………Problem 3: collarless is a very common polymorphism in Anopheles gambiae s.l.(Mason 1967). The callele is recessive to c+ and therefore both “ c+ c ” and “ c+ c+ “ individuals have identical phenotypes.Suppose you wanted to purify a pure-breeding “ c+ c+ “ stock. You go to the G3 (an old wild stock fromThe Gambia) stock tray and out of 100 larvae, you find that 90 are phenotypically c+. You inbreed the c+phenotype individuals.In the following exercise, we will determine the probability that you will get at least one family that was theresult of a mating between two “ c+ c+ “ individuals if you obtain 30 families from the c+ individuals matedto one another.First, calculate the allele frequencies in this population using the binomial equation.If p = the allelic frequency of c and q = the allelic frequency of c+3a. p 2 = ………………...3b. p = …………………3c. q = …………………

Chapter 2 : Anopheles Biology Laboratory and Culture2.6 Basic Anopheles Population GeneticsPage 4 of 6Using the binomial equation, what are the expected frequencies of c+ c+ and c+ c genotype individuals?3d. proportion c+ c+ ( i.e. q 2 ) = ………………..3e. proportion c+ c ( i.e. 2pq ) = ……………….Remember, as in the previous example where individuals have been removed from a population, we’reinbreeding ONLY the c+ phenotype individuals so you must adjust the frequencies of the above c+phenotypes before proceeding to reflect the fact that they now make up all ( = 1 ) of the individuals beingconsidered.Proportion c+ c+ ( i.e. q 2 ) + Proportion c+ c ( i.e. 2pq ) = 1Proportion FemalesHeterozygotes ( c+ c )3f. proportion 2pq= ………………..Homozygotes ( c+ c+ )3g. proportion q 2= ………………..Proportion MalesHeterozygotes ( c+ c )3h. proportion 2pq =………………..Homozygotes ( c+ c+ )3i. proportion q 2 =………………..3j. proportion =……………….. 3k. proportion = ………………..3l. proportion = ……………….. 3m. proportion = ………………..We asked, “If you isolate 30 families, what is the probability that you will get at least one family that wasthe result of a mating between two ‘ c+ c+ ‘ individuals?” This is the converse of asking, “If you isolate 30families, what is the probability that ALL will be from matings between something other than c+ c+ types?”3m: p 30 = ………………..3n: and the likelihood of getting at least 1 family is 1 − p 30 or ………………..Fitness Effects in Polymorphic PopulationsFinally, we consider polymorphic alleles in populations where the fitness of the various alleles is notequal. Fitness in the genetic sense refers simply to the relative reproductive success of an allele or agenotype. Mutant alleles are usually assumed to have reduced fitness relative to wild-types, but this is notalways so. Perhaps the collarless phenotype is less fit than wild-type. If so, what happens to thefrequency of that phenotype and the collarless allele frequencies over time? Other alleles whose fitnesswe might want to consider in the insectary are recessive lethals, insecticide resistance alleles, andparasite susceptibility alleles.

Chapter 2 : Anopheles Biology Laboratory and Culture2.6 Basic Anopheles Population GeneticsPage 5 of 6Alleles can have a positive or negative fitness depending on the environment.Fitness can be expressed as an adjustment variable ‘s’ to the allele frequency. For example, a neutralallele with no effect on fitness has a relative fitness of ‘ s = 0 ’ such that the allelic frequency adjustmentwould be q ( 1- s ) or q (1 - 0 ) or simply q. On the other hand, a lethal has a fitness of 1 so that q ( 1- s )or q (1 - 1 ) becomes simply 0 (zero).Alleles with fitness of

Chapter 2 : Anopheles Biology Laboratory and Culture2.6 Basic Anopheles Population GeneticsPage 6 of 6ReferencesBeard CB, Benedict MQ, Primus JP, Finnerty V, Collins FH (1994) Eye pigments in wild-type and eyecolormutant strains of the African malaria vector Anopheles gambiae. J Heredity 86:375-380Mason GF (1967) Genetic studies on mutations in species A and B of the Anopheles gambiae complex.Genetical Res, Cambridge 10:205-217Strickberger MW (1968) Genetics. Macmillan, New YorkAnswers1a: 0.2 (this is the square root of the frequency of “ r r ” individuals)1b: 0.81c: 0.641d: 0.322a: 0.33 3k: 0.252b: 0.67 3l: 0.252c: 0.33 3j: 0.232d: 0.67 3n: 7.9 X 10 -52e: 0.1089 3o: 0.9992f: 0.2112g: 0.2112h: 0.44892i: about 22k: 0.792l: 0.0997 or odds of about 1 in 103a: 0.103b: 0.3163c: 0.6843d: 0.473e: 0.433f: 0.48 (2pq X 1.111)3g: 0.523h: 0.483i: 0.523m: 0.27

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosPage 1 of 8Chapter 3: Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosMark BenedictIntroductionWe present two methods that have been successful (and a variation of Method 1 using oil). The first wasdeveloped by John R. (Randy) Clayton for injection of An. gambiae embryos, and it has been used toobtain high frequency egg hatching and EGFP transient expression rates as described by Grossman etal., 2001 (Grossman et al. 2001). This method is similar to that used by Dave O’brochta’s group at theUniv. Maryland. They successfully transformed An. gambiae using a similar method by covering theembryos with halocarbon oil prior to injection (Kim et al. 2004). Using oil should provide better visibility ofthe needle flow rate.The second method is fast and requires less judgment than those above. It was developed by HervéBossin and Mark Benedict for An. arabiensis and An. gambiae. However, it should be useful for manymosquito species. Anecdotally, mosquito species vary in the ease with which they can be microinjected.Both of the methods above have been used successfully with gambiae s.l. which is supposed to be one ofthe more difficult to inject.We recommend mounting the injection needle in a fixed position and moving the slide holding the alignedembryos using the stage controls to the appropriate place for injection. This allows one to use a rathersimple needle positioner mounted on or by the microscope.Anopheles embryos cannot be dechorionated, so use of rigid needles and firm positioning of the embryosis necessary. Quartz glass injection needles are by far preferable to aluminosilicate or borosilicateneedles. These require higher pulling temperatures than the other glasses and therefore a laser needlepullermust be used.Materials:• Mated adult females bloodfed 3-5 days post-eclosion.• Clean water in a wash bottle• Pipettor e.g. P20• Fine paint brushes 1 and forceps• Filter paper• 2X Na phosphate Injection buffer (see below for preparation; requires KCl and di- and monobasicsodium phosphate)• Minimum fiber filter paper e.g. (Whatman 1450-090, ‘Hardened circles’)• Eppendorf Microloader tips (no. 5242-956-003)• Ultrafree-MC filters (no. UFC30HV00)• Quartz glass capillaries, 1 mm OD, 0.7 mm ID X 10 cm length (e.g. Sutter no. QF100-70-10 )1 Select the brushes carefully from among the finest at an art or craft store. Sable brushes are excellentand more expensive, but regardless, a very fine pointed tip is essential.

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosPage 2 of 8• Double-sided adhesive tape. 3M type 415 has been tested for vertebrate toxicity and is a goodchoice (Method 1 only)• 22 x 22 glass or plastic coverglasses with a strip of pre-cut double-sided tape attached (Method 1only)• 25 mM NaCl (or 10-50 mM range for testing, Method 1 only)• Millipore (or other) mixed cellulose ester membranes (e.g. HAWP02500- US or HAWP03700,Method 2 only) 2Equipment• Either a compound or high-quality dissecting microscope can be used for injections. It ispreferable if it can be dedicated to this purpose.• Sutter P-2000 Micropipette puller or similar device 3• Needle positioner and holder• pH meter• Eppendorf Femtojet or similar device equipped with a foot pedal• Dissecting scope and illuminator for embryo alignmentSolutionsThis recommendation is for Drosophila melanogaster from Bill Engels lab, but it seems suitable formosquitoes. Prepare two 0.1 M solutions of monobasic and dibasic sodium phosphate. Mix the two andadjust pH with one or the other to pH 6.8-7.8. Prepare a solution of 0.5 M KCl in purified water.2X injection buffer is:0.2 mM Na phosphate10 mM KClFilter sterilize and store at room temperature or lower.Starting procedures common to both methods1. Prepare the capillaries by flushing them several times with purified water followed by ethanol toremove lint and glass chips. Blot the remaining ethanol and flame the capillaries briefly to remove allliquid. It is convenient to store them in a covered glass culture tube.2. Immediately before use, thaw the DNA and filter through a 0.2 micron Millipore Ultrafree-MC filterfilter to remove particulates. This latter measure (suggested by D. O’brochta) is simple and effective.Store on ice until use.2 Any non-fibrous membrane that is very thin, hydrophilic and does not contain detergent would probablywork for this e.g. Southern blotting membrane. If in doubt regarding the presence of wetting agents etc.rinse the membrane well before use. These types of membranes are perfect because when wet, theyadhere closely to a microscope slide so that the embryos don't slide beneath the membrane, and they areabout the same thickness as an embryo so visualization is easy.3 The program we use for the P-2000 is: HEAT: 650, FIL: 4, VEL: 40, DEL: 150, PUL: 157. However,conditions necessary to produce suitable needles may differ on your device and may require slightchange, even from day to day.

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosPage 3 of 83. Harvest eggs 60-96 hours after females are bloodfed by placing 10 to 15 females in a transparentcylindrical container (~40 ml) open at one end and covered with rubber dental dam at the other(Figure 3.1.1.1). Alternatively, eggs may be collected on damp filter paper in a small Petri dish 4 .Figure 3.1.1.1. Cylindrical container (~40ml) containing 10-15 previously bloodfedfemales open at one end and covered withrubber dental dam at the otherFigure 3.1.1.2. Container with females resting overa single water-filled well on a cobalt blue ceramicdepression plate. Embryos deposited in singledepression shown (inset).4 Cooling the eggs will extend their useful time, but MQB’s experience has been that this also reducessurvival.

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosPage 4 of 8Figure 3.1.1.3. Egg darkening at 20 min intervals (top left to bottom right) beginningapproximately 15 m after oviposition and incubated at room temperature (~22°C). Most of thedarker eggs in the lower two left panels are suitable for injection. Note that one egg did notdarken. It will not hatch. When eggs have fully darkened (as in the lowest right panel) but areoften still at the blastoderm stage, they are more difficult to inject (photographs courtesy of G.Labbe, Oxitec, used with permission). Non-melanized eggs cannot be handled without sufferingmortality.4. Slide the container over a single water-filled well on a depression plate such that the females canaccess the water (see Figure 3.1.1.2). Cover to darken for 30 minutes and then remove the females.5. Age the new embryos for at least 30 minutes at insectary conditions (28°C; 80% humidity) or at roomtemperature. After this time, the eggs should be medium gray (Figure 3.1.1.3).Method 1:The distinctive feature is that embryos are injected under saline, the concentration of which is determinedempirically to balance internal pressure. The balance pressure must be such that the turgidity is sufficientfor needle penetration yet low enough that oozing and needle backflow are minimized. The saltconcentration can be adjusted as needed to achieve this for each species.1. At room temperature (~ 24 o C), transfer eggs from the depressions to a glass slide with a finepaintbrush and align with the dorsal (flattened, concave) surface facing up.2. Align the anterior ends of 25-35 eggs in 25 mM NaCl against a strip of reduced-fiber filter paper.3. Remove the filter paper by tugging it away sharply, so as not to disturb the alignment of the embryos.4. Allow eggs to desiccate slightly.5. Press a taped coverslip gently against the eggs’ dorsal surface and immediately invert. Getting theembryos to stick to the tape is the most difficult part of this procedure. They cannot be too wet - theywill not stick - or too dry - in which case they die.6. Cover with a solution of 25 mM NaCl (or Halocarbon oil) to prevent drying and place eggs in a humidbox at room temperature until injection. Eggs are appropriate for injecting around 2 hours afterdeposition when they have are medium dark. Choosing eggs for injection is discussed in Figure3.1.1.3.7. Immediately prior to injection, add more 25 mM NaCl to the coverslip. A large volume surrounding theembryos is desirable as it reduces distortion of the image. Attach the coverslip to a glass slide with abit of double-stick tape along the edge and place on the stage.8. Inject embryos on the ventral surface, near the posterior end, with the embryo turned at an angle ofabout 15-25 degrees. The horizontal angle of the needle can vary, but should be roughly within 30degrees vertical from the plane of the stage. Take care to avoid injection into the periplasmic space.Instead, inject immediately anterior to the periplasmic space and posterior to the egg floats. Injectionsshould be carried out at 100X magnification.9. Immediately after all of the embryos on a slide are injected, remove the slide from the scope and andplace the coverslip carrying the eggs into a cup of reverse-osmosis/deionized sterilized (RO/DI) H 2 0at room temperature to recover.10. When all injected coverslips from a cohort have been aligned and injected, placed all coverslips in acup of 50 ml RO/DI H 2 0 under insectary conditions to hatch. It is not necessary that the eggs float forhatching.11. Hatching will begin in approximately 48 hours after which the larvae can be handled as described inthe chapter on family culture.

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosPage 5 of 8Method 2:This method differs in that embryos are aligned against a thin membrane and injected semi-dry. Noadhesive tape is necessary.1. Cut pieces of membrane with a scalpel or razor blade at an approximately 45° angle so that theposterior edge for injection will be perpendicular to the needle. A cleanly cut edge is desirable.2. Cut a piece of filter or blotter paper smaller than the height of the slide. You may wish to stack acouple of pieces to provide a larger water reservoir.3. Assemble the membrane and filter paper as shown (Figure 3.1.1.4) and wet with water so that allpaper is wet and the membrane is moist but not dripping.4. Using a brush, transfer 30-50 embryos to the edge of the membrane.5. Distribute them as shown (Figure 3.1.1.5) with the narrower posterior end toward the bottom. Whenaligning the embryos, roll them over so that the ventral side (convex) is upward, and they will nestnicely in the 90º niche between the membrane and slide.6. Orient all in the same direction. As you work, keep the papers moist by adding small volumes (10 μl)of water to the blotter paper. You should maintain a meniscus of water around the eggs, but do notwet excessively causing the eggs to become dislocated.7. When you have filled the edge of the membrane with eggs (~50), transfer to the scope for injection.Keep in mind that when using this technique, the needle will not be submerged in liquid, so keepsufficient back-pressure on the needle to keep it cleared. Frequently check the needle flow bywithdrawing the needle and ‘inject’ into air. You should see a small droplet appear or run back up theneedles into a larger droplet that often hangs on the needle shaft. Add small volumes of water to theblotter paper as the eggs dry during injection.8. After injection rinse the eggs off into a Petri dish using water and incubate as in Method 1.Figure 3.1.1.4. Diagram of microinjection methodusing membranes as opposed to the adhesivedouble-stick tape.Figure 3.1.1.5. Eggs aligned for injection with acloseup of the needle approaching perpendicularto the injection area (photograph courtesy of G.Labbe, Oxitec).

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosPage 6 of 8Other things you might want to knowQ: Do you remove the chorion before injection as has been described in Drosophila?A: No. Endochorion removal in anophelines has not been accomplished. This is why the quality of theneedles and turgor of the eggs is crucial.Q. What does a good An. gambiae injection look like?A: Larval hatch rates vary between 10% and 50% using either method. The most probable cause of thisvariation is physical wounding of the embryo during injection. If the needle does not slide easily throughthe chorion of the egg during injection then something is wrong. It is the ease of penetration that allowscontinuous injection without needle clogging or breakage.Under good conditions, the needle will slide in and out of the egg with little effort. Slight resistance topenetration is apparent when entering the egg and a small volume of yolk can sometimes be seen flowinginto the tip of the capillary, only to be expelled immediately during injection. Although visibility is worseinjecting under aqueous solution rather than halocarbon or mineral oil, a slight clearing of the yolk is oftenseen, even through the dark chorion. Injected eggs sometimes recoil and bulge briefly and slightly when asufficient volume has been released into them and this is also a good sign as long as a minimal amountof yolk escapes from the wound site.Q: I don’t have a laser needle puller. Will this method work with boro- or aluminosilicate needles?A: In principle, there is no reason why this method would not work with a softer glass but with frequentneedle replacement; however, an attempt at this has not been published. Quartz needles may simplyallow a larger degree of error on the part of the person injecting. Aluminosilicate glass needles arepreferable to borosilicate because of their greater hardness.Q: Have you used a chorion hardening inhibitor?A: No. Many inhibitors have been tested of the prophenoloxidase activation cascade (pNpGB,benserazide, PTU), but we have found nothing that clearly resulted in an increase in embryo injectability.Q: Do you bevel your needles?A: No. Non-beveled quartz is hard and sharp enough so that needles can be pulled and usedimmediately.Q: How do you prepare your DNA for injection?A: DNA was prepared with a Qiagen Endo-Free kit and resuspended in injection buffer. It was then storedat -80 o C until use.Q: What do you feed your hatching larvae?A: We feed L1 larvae two drops of 2% w/v baker's yeast on day two post-injection and another two dropson day four. Beyond day four, we feed as appropriate with our standard food mixture of finely ground KoiFloating Blend.Q: I thought anopheline eggs floated when they hatched. Aren’t your injected embryos submergedwhen they hatch in Method 1?A: Yes. While Anopheles eggs typically do float, submerging eggs post-injection does not seem to have astrong effect on mortality relative to floating controls. In addition, this method avoids the large degree ofmortality which was inflicted when attempting to remove the eggs from the adhesive surface of the tape.Q: How hard do you press the coverslip down on the embryos when you’re picking them up inMethod 1?A: Delicately but firmly (!?). Just hard enough to see that the eggs have come into contact with the tapeand bulge slightly.Q: What happens if I inject the embryos earlier than you describe?

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.1 Microinjection Methods for Anopheles EmbryosPage 7 of 8A: Younger embryos are difficult to inject due to sensitivity to handling. Simply moving them early indevelopment kills them.ReferencesGrossman GL, Rafferty CS, Clayton JR, Stevens TK, Mukabayire O, Benedict MQ (2001) Germlinetransformation of the malaria vector, Anopheles gambiae, with the piggyBac transposable element. InsectMol Biol 10:597-604Kim W et al. (2004) Ectopic expression of a cecropin transgene in the human malaria vector mosquitoAnopheles gambiae (Diptera: Culicidae): effects on susceptibility to Plasmodium. J Med Entomol 41:447-455

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Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.2 Anopheles Embryo FixationPage 1 of 43.1.2 Anopheles Embryo FixationMR4 StaffIntroductionThe following embryo fixation method is suitable for preparing An. gambiae embryos for immunostainingand may be suitable for other anophelines or even genera of mosquitoes. It was developed and used byYury Goltsev (2004) and further tested by John Yoder (2006). After fixation, embryos of the properdevelopmental stage must be selected from the pool for analysis. Removal of the relatively impermeablechorion of An. gambiae requires a method different from that used for Drosophila melanogaster.Solutions• purified water e.g. distilled or reverse-osmosis/deionized• 25% household bleach diluted in purified water• heptane• 9% formaldehyde in purified water, adjust to pH 7 with NaOH.• methanolMaterials• glass vials e.g. scintillation vials• 100 micron nylon mesh or similar (for device in Figure 3.1.2.1)• Pasteur pipettesProcedure:Figure 3.1.2.1. Eggs must be contained in amanageable mesh container in which solutionscan be added and removed by draining andrinsing. Shown is an example of a possiblecontainer improvised from a 50 ml disposabletube that was cut. The lid clamps the meshand has a hole to allow solutions to pass.1. Remove egging cup containing newly laidembryos from mosquito cage and hold at20 o C. Embryos can be collected for about 3hours and then held until the desireddevelopmental stage.2. Rinse eggs into a fine mesh basket (e.g. 100micron nylon mesh) with deionized water.Place mesh with eggs into an empty Petri dishunder a stereo microscope. An example of apossible container for eggs is shown in Figure3.1.2.1.3. While watching the embryos through themicroscope, gently add bleach solution to theegg container until the eggs are floating. Thisstep washes away the exochorion. Swirlgently 1-2 times; remove the mesh containerwhen approx 50% of the eggs sink.4. Rinse eggs and mesh container thoroughlywith deionized water.

5. Place eggs in a new Petri dish containingpurified water while you “test crack” a sampleof the embryos. This step is necessary toensure that the exochorion was removedduring the bleaching step. If this layer is notremoved, the fixatives will not permeate theembryo and fixation will fail.a. Aliquot approx. 25 test embryos into ascintillation vial. Remove the waterwith a pipette and add 5 ml heptane.b. Incubate at room temperature for 5minutes while occasionally swirlinggently.c. Add 5 ml of methanol and vigorouslyswirl once to mix. Place scintillationvial on its side under the stereoscopeand watch for cracking (Figure3.1.2.2). It is normal for the embryosto seep out of their chorion. If theembryos crack, discard these testChapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.2 Anopheles Embryo FixationPage 2 of 4eggs and proceed with the protocol. If they do not, longer bleaching is needed.6. Rinse embryos into a new scintillation vial with deionized water. Remove as much water aspossible using a Pasteur pipette.7. Add 5 ml heptane and shake 3-4 times gently by hand to mix. Remove as much remaining wateras possible. Add 5 ml formaldehyde. Shake on rotary platform for 25 minutes on a medium speedsetting. Eggs will accumulate between layers as shown inFigure 3.1.2.3.8. Remove formaldehyde phase only (leaving heptanephase) using a fresh pipette. Replace with a large volumeof deionized water. Briefly shake 3-4 times gently by handand remove only the water phase (leaving heptanephase). Add 10ml of fresh deionized water.9. Shake on platform an additional 30 minutes on a mediumspeed setting.10. Remove only water phase (leaving heptane phase). Fillvial to the top with boiling deionized water. Incubate for 30seconds.11. Remove hot water phase (leaving heptane phase) andreplace with ice-cold deionized water.12. Place vial on ice for 10 minutes.13. Remove water phase using a glass pipette.14. Remove as much of the heptane phase as possible.Figure 3.1.2.3. Nondamagedeggs willaccumulate at theinterface between twolayers.Figure 3.1.2.2. Examples of eggs crackingproperly. Cracking is an indication that theprocess is proceeding correctly, and you cancontinue. Discard these test eggs.15. Add 5ml of fresh heptane. Remove as much water aspossible.16. Add 5ml methanol and swirl vigorously once, placescintillation vial on its side under stereo scope to watch forcracking, a sign the fixation has been successful to this

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.2 Anopheles Embryo FixationPage 3 of 4point.17. Let stand 15-20 minutes.18. Remove as much of both phases as possible. Rinse with 5 ml methanol twice removing anyexcess liquid, then add 5 ml fresh methanol.19. At this point, the embryos can be stored at -20 o C in methanol for several months.20. The endochorion must be manually peeled away using fine needles and double stick tape beforestaining.ReferencesGoltsev Y, Hsiong W, Lanzaro G, Levine M (2004) Different combinations of gap repressors for commonstripes in Anopheles and Drosophila embryos. Dev Biol 275:435-446Yoder JH, Carroll SB (2006) The evolution of abdominal reduction and the recent origin of distinctAbdominal-B transcript classes in Diptera. Evol Dev 8:241-251

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Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.3 Establishing Cell Lines from Anopheles spp. Embryonic TissuesPage 1 of 23.1.3 Establishing Cell Lines from Anopheles spp. Embryonic TissuesUlrike MunderlohMaterialsAnopheles eggsMosquitoes are commonly reared at ~28°C; other temperatures are suitable, but will influence the timingof egg production and embryonic development. Female mosquitoes are provided a blood meal from asuitable host in the afternoon of day 0. The afternoon/evening of day 2, a dish with clean water is placedin the cage, to allow females to deposit eggs over night.Eggs aged 24-36 hrs are collected using a transfer pipette, strip of screen, or filter paper, and added to a35 mm diameter Petri dish containing 70% ethanol with a drop of Tween 80 (e.g., Sigma-Aldrich catalogNr. P4780). The eggs will sink, and should be agitated by swirling the dish. The ethanol is replaced with0.5% benzalkonium chloride (e.g., Sigma-Aldrich catalog Nr. 09621) with a drop of Tween 80, and thedish again agitated for 5 min. The benzalkonium chloride is removed, and the eggs are rinsed 2-3 times insterile, distilled water. 50~100 eggs are transferred to a new dish containing 0.2 ml of culture mediumsupplemented with 10-20% fetal bovine serum (FBS, heat-inactivated), 5-10% tryptose phosphate broth(TPB), and a mixture of penicillin (50-100 units/ml) and streptomycin (50-100 µg/ml; e.g., Invitrogencatalog Nr. 15140-122) and fungizone (0.25 – 0.5 µg/ml; e.g., Invitrogen catalog Nr. 15290-018).MediaWe have used Leibovitz’s L-15 medium successfully, as well as a modification thereof, L-15B, diluted to~300 mOsm/L using sterile cell culture grade water (Munderloh and Kurtti 1989; Munderloh et al. 1999).Other media may be substituted, such as RPMI1640, Medium 199, Eagles’ MEM, or Ham’s F12 (e.g.,from Invitrogen, http://www.invitrogen.com/site/us/en/home/Applications/Cell-Culture/Mammalian-Cell-Culture.reg.us.html) with 10% - 20% FBS (Invitrogen or Sigma) and 5-10% TPB (Becton Dickinson,catalog Nr. 260300), but should be tested for their ability to sustain primary and established cell lines. ThepH of the medium should be adjusted to 7.0 to 7.2 using either sterile 1-N NaOH or 1-N HCl, as needed.If the medium pH drifts up beyond 7.8, it may be useful to add a buffer such as HEPES (e.g., Invitrogencatalog Nr. 15630) or MOPS (e.g., Sigma-Aldrich catalog Nr. M1442) at ~25 mM concentrationMethodsEggs are crushed by applying gentle downward pressure using a sterile glass or plastic plunger from a 3or 5-ml syringe, the flattened end of a sterile glass rod, or similar device. Crushed eggs and tissues arecollected with a 2-ml pipette, and transferred to a 5.5 cm 2 flat-bottom tube (Nunc, catalog Nr. 156758) in1-2 ml of complete medium containing antibiotics and antifungal solution as above, and the tubes aretightly capped. Cultures are incubated flat side down at 28-31°C. Use of a CO 2 incubator is not necessaryand not recommended.Cultures are fed approximately once a week by removing as much of the medium as possible withoutaspirating any tissue fragments or cells and replacing it with 2 ml of fresh medium. Antibiotics/antifungalsshould be included in the medium for the first few weeks, and can be omitted subsequently. If it is desiredto continue using antibiotics, the antifungal component should be omitted. A mixture of penicillin andstreptomycin is preferable over gentamycin as the latter may adversely affect mosquito cell lines in thelong term.The progress of the cultures is best monitored using an inverted phase contrast microscope. During thefirst days after adding the embryonic fragments, most tissue clumps will remain non-adherent, and organssuch as guts and Malpighian tubules should show active peristaltic movements. Within days to weeks,cells should be migrating out from the torn tissue ends, and will often anchor to the bottom of the tube.Commonly, hollow balls consisting of a “monolayer” of cells surrounding a fluid-filled interior, will be seen

Chapter 3 : Specific Anopheles Techniques3.1 Embryonic Techniques3.1.3 Establishing Cell Lines from Anopheles spp. Embryonic TissuesPage 2 of 2sprouting from embryonic fragments. Once cells in a primary culture have replicated sufficiently, a portion(~1/2 to 1/3) of the tissues can be removed by pipetting, and transferred to a new tube to set up asubculture. It is advisable to keep the other portion of cells in the parent flask or tube, as it is common thatthey will remain vigorous even if the subculture should fail.This process is repeated many times until cultures can be subcultured or split on a regular basis, and theculture is considered established. During this process increasingly larger culture vessels will be used,e.g., 12.5-cm 2 flasks, then 25-cm 2 flasks, etc. Although the first several subcultures are usually made bytransferring 30 – 50% of the cells to a new culture vessel, with time it is advisable to “push” a cell line byusing higher dilutions of 1:10 or more. Some mosquito cell lines can be diluted up to 100-fold. Seedingsubcultures at relatively high densities (dilutions of 1:2 or 1:3) will depress cell replication and slowgrowth, often resulting in cultures of poor condition.It is not uncommon that a single primary culture will give rise to sublines displaying differingmorphologies. Some sublines may continue growth as adherent cells, and others may becomeestablished as suspension cultures. In particular, the “hollow ball” or “vesicle” phenotype frequentlydevelops, and may become fixed. Sublines with particular, desired characteristics can also be selected byculture manipulation (e.g., adherent lines can be developed by continuously discarding non-adherent cellsduring medium changes). Once cells are growing reliably, it is a good idea to try to reduce the amount ofFBS, and TPB. Established mosquito cell lines commonly grow quite well with only 5% of FBS. Adding alipoprotein supplement (such as the one from Rocky Mountain Biologicals, Missoula, MT, or the CellPro-LPS from Fisher Scientific), if available, can further reduce the requirement for FBS, and reduce cost.These methods can equally be applied to other mosquito species, keeping in mind the length of timerequired for embryonic development. Eggs should be at least at their half point before hatching, all theway up to just before hatching. Although open culture vessels can be used, such as small Petri dishes ormulti-well plates, they require a humidified atmosphere as well as a CO 2 incubator when media containingbicarbonate are employed. Also, open culture vessels are far more susceptible to contamination thanclosed ones.Additional references: (Munderloh et al. 1982) (Mazzacano et al. 1991)ReferencesMazzacano CA, Munderloh UG, Kurtti TJ (1991) Characterization of a New Continuous Cell Line from theFlood Water Mosquito, Aedes vexans. Cytotechnology 5:147-154Munderloh UG et al. (1999) Invasion and intracellular development of the human granulocytic ehrlichiosisagent in tick cell culture. J Clin Microbiol 37:2518-2524Munderloh UG, Kurtti TJ (1989) Formulation of medium for tick cell culture. Exp Appl Acarol 7:219-229Munderloh UG, Kurtti TJ, Maramorosch K (1982) Anopheles stephensi and Toxorhynchites amboinensis:aseptic rearing of mosquito larvae on cultured cells. Journal of Parasitology 68:1085-1091

Chapter 3 : Specific Anopheles Techniques3.2 Eye Color Mutant ScreeningPage 1 of 23.2 Eye Color Mutant ScreeningMark BenedictIntroductionAnopheles mosquitoes undergo induced color change (called homochromy) based on perception of thebackground against which they are cultured. 1 When larvae are reared on either a dark black or whitebackground, they become pigmented dark or pale respectively as shown by the pair of An. albimanuslarvae in Figure 3.2.1. The degree of darkening depends in part on the length of time the larvae havebeen cultured in a black container and the degree of fat body development. Therefore, larvae culturedduring their entire development in a dark container at a low density show this change most dramatically.This color change depends on the normal eye pigmentation and, presumably, on the proper function ofany pigmentation and signaling pathways involved in the response.The method simply requires culturing the larvae from at least the second stage in black or dark-coloredcontainers that are illuminated. (Larvae cultured in darkness will develop typical pale pigmentation.) Thesource of illumination does not appear to be critical. Occasional transfers of a few minutes to whitecontainers for feeding or correcting the density does not interfere with the effect.At the L3 or L4 stage, larvae are scanned en masse in the dark tray in a well-illuminated location for thosethat appear lighter in color. They are usually quite apparent as demonstrated by the two larvae in Figure3.2.2, but purposely seeding a sample of dark larvae with a few that are pale will demonstrate the degreeof effect that can be expected. These individuals are transferred to a dish for microscopic examination.Leaving the larvae undisturbed during examination provides better visualization since the dorsal sidecoloration is a more consistent indicator of general color. After this initial selection, it is also helpful totransfer the larvae to a white tray and scanning for pale individuals. Usually, no more than approximately25 larvae per thousand cultured in this way require individual examination.Figure 3.2.1. Anopheles albimanus larvae rearedon a black background (top) and a whitebackground (bottom).Figure 3.2.2. Eye color mutants are easier to detecton a dark background as they will not changepigmentation and will appear much lighter in color.1 Before beginning a large screen, it is advisable to culture a thousand or so larvae of the species ofchoice in dark containers. While all individuals of most species change color, some laboratory stockshave a low frequency of individuals that do not.

Chapter 3 : Specific Anopheles Techniques3.2 Eye Color Mutant ScreeningPage 2 of 2ReferencesBenedict MQ, Besansky NJ, Chang H, Mukabayire O, Collins FH (1996) Mutations in the Anophelesgambiae pink-eye and white genes define distinct, tightly linked eye-color loci. Journal of Heredity 87:48-53Benedict MQ, Chang H (1996) Rapid isolation of anopheline mosquito eye-colour mutants based on larvalcolour change. Medical and Veterinary Entomology 10:93-96

Chapter 3 : Specific Anopheles Techniques3.3 Determining the Sex of Anopheles Larvae and PupaePage 1 of 43.3 Determining the sex of Anopheles larvae and pupaeMR4 StaffIntroductionThere is often an experimental need to separate the sexes before they emerge e.g. in order to preserveunmated status of females, to obtain material for molecular analysis, or to determine male/female larvalratios. Anopheles spp. differ from many other mosquitoes in that there is often no easily discernabledifference in the female/male larval or pupal size or pigmentation. Some, but not all, anophelines L4females can be identified based on the generally darker color and larger size. Here we present threemethods for determining the sexes based on larval and pupal characteristics.Larval sex determination: Option 1Figure 3.3.1. Cartoon of a ‘sandwich’ slide whichworks well to position and immobilize a larva forviewing without causing injury.An early method for sexing Anopheles larvaebased on the form of the imaginal antennal lobeshas been reported (Jones 1956), but the graphicsin the manuscript can be difficult to interpret –particularly in copies. Here we offer a refinementof the method and new images developed for An.gambiae. (Note: This method is not very usefulwith An. stephensi because the imaginal disks aredifficult to see.) The best results are usuallyobtained with 2 nd day L4s as the pre-antennal lobeis almost fully formed. All observations andphotographs were made on a stereoscope, and itis important to use the dark-field setting.Constructing the viewing slide:Materials• Standard glass microscope slide• 0.3 - 0.5 mm thick plastic spacer e.g. a thin plastic laboratory ruler cut into 1 X 1.5 cm pieces. Thethickness must be selected to support a coverslip over the gap so that a larva is held firmly butnot crushed. A stack of plastic coverglasses may be stacked and glued together to obtain theappropriate thickness.• Epoxy glueConstruction and use1. Clean the slide with ethanol and dry.2. Apply a small drop of epoxy glue to the plastic spacers.3. Glue the spacers onto the slide 0.8 - 1cm apart from each other and allow to cure.

Chapter 3 : Specific Anopheles Techniques3.3 Determining the Sex of Anopheles Larvae and PupaePage 2 of 44. Place a larva, dorsum upward, between the spacers. Add sufficient water to fill the gap, and place acoverglass on top such that it bridges the spacers. The pre-antennal lobes can be seen between theimaginal eyes while viewing the dorsal side of the head. In males, the lobe is large, circular andeasier to see. The female’s lobe is smaller and it is only easily seen in the second and third days ofthe fourth instar. Males are typically a bit easier to identify than females (see Figures 3.3.2 and3.3.3).Figure 3.3.2. L4 Anopheles gambiaemale. Region of interest for sexdetermination is circled.Figure 3.3.3. L4 Anopheles gambiaefemale. Region of interest for sexdetermination is circled.Larval sex determination: Option 2In An. gambiae and arabiensis, the "Red stripe" character can be visualized in L3s through mid pupae(Benedict et al. 2003) and provides a character that can be used to positively identify females with highcertainty. When the collarless alleles (see Morphological Characteristics, Chapter 4) are heterozygous (c+/ c), a red stripe is evident on the female dorsum. The collarlesstrait is polymorphic in most colonies and wild populations andappears to have little if any effect on vigor. In Figure 3.3.3, thedorsum of this L4 larva has both white and red pigmentcharacteristic of a c+ / c heterozygous female. While thepresence of the red stripe can be used to select females withhigh certainty, the absence of the red stripe does not necessarilyindicate a male in a polymorphic population. Though this methoddoes not allow one to distinguish males, a cross between ahomozygous c+ / c+ and c / c individuals would create F1heterozygotes in which both sexes could be distinguished withhigh certainty. X-chromosome markers could also be used in agenetic scheme to produce progeny whose sex could bedetermined as early as the L1 stage. (Strains suitable for suchcrosses are available from the MR4.)Figure 3.3.3. Red stripe characterindicates a female larva.

Chapter 3 : Specific Anopheles Techniques3.3 Determining the Sex of Anopheles Larvae and PupaePage 3 of 4Pupal sex identificationAnopheles pupae are much simpler to sex thanlarvae. The pupa should be lying on its side semidryin order to see the genitalia easily, and it maybe necessary to use a small brush or forceps togently lift the paddles.1. Using a pipette, gently transfer 1 pupae toeither a depression well plate or a piece ofdamp filter paper. If using a depression wellplate, remove as much water as possible sothat the pupa is lying on its side.2. Under a stereoscope, observe the prominentgenitalia for comparison with Figure 3.3.4.ReferencesBenedict MQ, McNitt LM, Collins FH (2003)Genetic traits of the mosquito Anopheles gambiae:Red stripe, frizzled, and homochromy1 Journal ofHeredity 94 227-235Jones JC (1956) A Simple Method for sexing livingAnopheles larvae (Diptera, Culicidae). Annals ofthe Entomological Society of America 50:104-106Figure 3.3.4. The terminalia of the pupae are verydistinctive. However, the paddles can easily get inthe way making it difficult to distinguish. Gentlypoking the pupa will usually make them changeposition to reveal the terminalia.

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Chapter 3 : Specific Anopheles Techniques3.4 Mosquito AnesthesiaPage 1 of 43.4 Mosquito AnesthesiaMark Benedict and Paul HowellIntroductionAdult mosquitoes can be immobilized by chemical anesthesia or by chilling. In the following, we describesimple methods and apparatuses to accomplish this. Be aware that in excess, all of these methods resultin mortality and must be tested before routine use. Generally, highest survival is obtained with theminimum exposure sufficient to immobilize the mosquitoes. A common indicator of stress due toanesthesia short of lack of recovery is tarsi falling off.CO 2 and N 2Carbon dioxide and nitrogen gas are both useful for anesthesia and are very safe for human exposure.They can be supplied as compressed gas, as vapor from sublimation of dry ice or evaporation of liquidnitrogen. CO 2 has the distinct advantage over nitrogen that, its density being greater than air, it pools intrays where mosquitoes are placed. Furthermore, its smell is distinct and unpleasant making detectionsimple. Nitrogen has no odor and regulation of the amount and its presence is more difficult to determine.However, given the apparent disadvantages of N 2 , I have heard reports that nitrogen anesthesia ispreferable to maintain some behaviors e.g. for forced copulation (for which see McCuiston and White1976).Shown is a small chamber made for anesthesia ofmosquitoes (Figure 3.4.1). It is constructed ofPlexiglas and has a white plastic platform made ofporous polyethylene (e.g. Small Parts no. SPE-040-20) through which gases can pass. The gas is fedinto the lower portion from which it oozes up ontothe mosquitoes lying on the tray. The raised walls ofthe upper portion help retain a reservoir of CO 2 .This same container can be used for nitrogenanesthesia. Similar simple improvised anesthesiacontainers can be constructed using fine nylonmesh and plastic containers at hand.Figure 3.4.1. This improvised chamber providessufficient area for as many mosquitoes as is safeto keep anesthetized at one time.A simple modification is to bubble the gasesthrough water to humidify them. A device that willperform this can be made from a flask and rubberstoppers. We have no information indicatingwhether this measure increases longevity and/orrecovery, but it is a prudent measure.Ethyl Ether and ChloroformVapors of both of these are useful for anesthesia when used safely (see safety measures at the end ofthis section). Their volatility and combustibility combined with their effects on humans make them asecond choice to CO 2 and N 2 . However, they are very portable and require little equipment in use.Moreover, their effects are generally longer-lasting than those of CO 2 and N 2 meaning that more workingtime with an immobilized mosquito can be achieved. Chloroform kills mosquitoes more readily.Both can be administered by pouring the minimum effective amount (e.g. 1 ml) onto an absorbentmaterial such a sponge or cotton wool. The container holding this should be air tight and sufficiently largeto introduce a mosquito holding tube. Mosquitoes are blown into the tube which is then placed in thechamber until the adults are knocked down. The tube is then removed and the adults poured out.

Chapter 3 : Specific Anopheles Techniques3.4 Mosquito AnesthesiaPage 2 of 4TriethylamineThis chemical is used for Drosophila quite successfully because it is extremely safe. It is available for thispurpose from Carolina Biological Supply as “FlyNap.” Our limited experience with this is that it‘anesthetizes‘ Anopheles mosquitoes irreversibly. In this regard, it is similar to chloroform but would be agood choice when extended immobilization but not recovery is acceptable. Normal biological activities ofseveral types during anesthesia have been confirmed in Culex (Kramer et al. 1990), but we are not awareof similar observations of Anopheles.ChillingFigure 3.4.2. A small Peltier cold table on whichdamp filter paper is placed to conduct cold andprevent mosquitoes from sticking to thecondensation.All mosquitoes with which I have had experience willwithstand some degree of chilling on ice followed byresting on a near-freezing surface. Shown is a smallchilling table that can be used for this purpose. Formosquitoes it is usually necessary to cover the coldsurface with a thin piece of damp paper to prevent themosquitoes from sticking to the condensate on thebare platform. Being slightly damp makes the paperadhere uniformly to the plate and increases heattransfer. Even this must be changed frequently as itbecomes sticky. Entire cages or cups of mosquitoescan be placed briefly in a freezer in order to knockthem down and then transferred to a chilled surface,but make the time as short as possible since mostanophelines will not survive total freezing.Ether and Chloroform SafetyExposure to ether and chloroform should beminimized by keeping containers sealed, dispensingminimum amounts, and using them for the shortestpossible durations of time.Diethyl ether should be stored in a flammable storagecabinet or an explosion proof refrigerator not longerthan 3 - 6 months. This cabinet should not be used tostore oxidizing agents. Explosive peroxides can formwith long term storage, so purchase and store onlyenough for immediate needs. The occupationalexposure limit is 400 ppm as a time-weighted average(TWA, 8 hour exposure) and it has a short term limitexposure (15 minutes) of 500 ppm. Keep theanesthesia chamber closed as much as possible. Review the specific MSDS of the manufacturer youpurchase ether from for any additional handling and storage recommendations, as well other relevanthealth and safety information. Some manufacturers recommend that you do not open unless contentsare at room temperature or below, and that after opening the container, any unused ether be discarded ordisposed of after 2-3 days. Only dispense ether in a chemical fume hood. Avoid agitation and sparksduring all phases of use.Chloroform is considered a known animal carcinogen with unknown relevance to humans. Itsoccupational exposure limit for an 8-hour TWA exposure is 10 parts per million (ppm). However,exposures to chloroform should never exceed 50 ppm at any time. This is referred to as OSHA’s ceilingoccupational exposure limit. Chloroform has a low odor threshold of 85 ppm, so by the time you smellchloroform the ceiling concentration would have already been exceeded. Anesthetizing procedures

Chapter 3 : Specific Anopheles Techniques3.4 Mosquito AnesthesiaPage 3 of 4should be performed in an area with good ventilation, preferably a chemical fume hood or other form oflocal exhaust enclosure.Employees using this chemical should be trained to recognize the acute and chronic health effectsassociated with an over-exposure that can occur by inhalation, absorption through the skin and byingestion. Selection of gloves is of particular importance since permeation of some nitrile gloves canoccur within as little as 3 minutes. Contact the glove manufacturer for specific selectionrecommendations. Employees should always be informed of glove limitations and trained accordinglyeven for incidental use. Chloroform should not be stored with caustics. Review the MSDS for additionalsafe handling, storage and disposal information.AcknowledgmentsThanks to Paul Vinson and Cheryl Connell of the CDC Office of Health and Safety for safety advice.ReferencesKramer LD, Presser SB, Houk EJ, Hardy JL (1990) Effect of the anesthetizing agent triethylamine onwestern equine encephalomyelitis and St. Louis encephalitis viral titers in mosquitoes (Diptera:Culicidae). J Med Entomol 27:1008-1010McCuiston LJ, White DJ (1976) Laboratory colonization of Aedes sollicitans (Walker) with a review of thetechnique of induced copulation. Proc N.J. Mosq Control Assoc.:164-175

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Chapter 3 : Specific Anopheles Techniques3.5 Protocol for 96 Well DNA ExtractionPage 1 of 43.5 Protocol for 96 Well DNA ExtractionClare Holleley and Alice SutcliffeIntroductionThis protocol describes the expansion of a common genomic DNA extraction method (salting-out method)to a 96-well platform adapted for anopheline mosquitoes. DNA extractions of individual insects havetraditionally been conducted by grinding individuals separately in 1.5ml tubes which can be a very timeconsumingprocess, especially when studies require large numbers. Expansion of the salting-out methodto 96-well PCR plates dramatically reduces the amount of time it takes to perform a large number ofextractions. Simultaneous 96-well insect maceration is achieved using a commercially available bacterialcolony replicator tool (Figure 3.5.1). The novel application of this tool as a maceration device coupledwith the salting-out procedure described below radically improves the efficiency of individual genomicDNA extraction without having a large impact on yield or failure. This protocol has been adapted formosquitoes from the original protocol used in Drosophila (Holelley 2007). Using this protocol, we obtainedaverage yields of 6.15µg and 5.65µg per mosquito by processing live and desiccated mosquito samplesrespectively. This compares to yields of 5.90µg and 3.50µg we obtained in our laboratory using the DNAextraction protocol described by Collins et al (1987).MaterialsForcepsIncubator or thermocycler48 or 96-pin bacterial replicator (See Figure 3.5.1)microplates and plastic capsUltra-Centrifuge (capable of holding 96-well microplates)Bunsen burner, striker, and gas sourceReagents1M Tris-HCl (pH 9.0 at 25°C)1M KClTriton® X-1005M Potassium AcetateProteinase K (Roche, 03 115 887)100% Isopropanol70% ethanolTE buffer 0.01 M, pH 7.4Reagent PreparationThermophilic DNA polymerase 1X buffer1. Add 46.95ml deionized water, 2.5 ml 1M KCl, 500µl 1M Tris-HCl (pH 9.0 at 25°C) and 50µl Triton X-100.2. Prepare 5ml aliquots of 1X buffer for long term storage at -20°C otherwise store at 4°C for short termuse.

Chapter 3 : Specific Anopheles Techniques3.5 Protocol for 96 Well DNA ExtractionPage 2 of 4Bacterial replicator sterilization:1. Place bacterial replicator in ethanol for 5 minutes ensuring the entire length of the pins is immersed.2. Heat replicator over Bunsen burner or similar until ethanol has evaporated for sterilization (5-10seconds).3. Set bacterial replicator aside to cool to room temperature before use in next steps.OPTIONAL: Other methods for sterilization include autoclaving or 2 hours of UV light.Mosquito preparation:1. Prepare a master mix of buffer as shown in Table 3.5.1.2. Aliquot 50µl of master mix into each well of a 48 or 96-well microplate.3. Using forceps, place one mosquito in each well cleaning forceps between samples.4. Carefully place sterilized, room temperature bacterial replicator into microplate (containing samples).5. Carefully grind mosquito samples for 10 minutes by moving replicator up and down and/or rockingreplicator from side to side. It is important that this is not done too vigorously as this can cause thebottom of the well to break or samples to be splashed, contaminating adjacent wells. When extractingDNA from mosquitoes and other insects, it is particularly important to thoroughly macerate the insectto allow cell lysis.6. Place plastic caps onto microplate and incubate 12-15 hours at 55°C.Number of samples96 48 11X DNA polymerase buffer 5000µl 2500µl 50µlProteinase K 20µl 10µl 0.2µlTable 3.5.1. Master mix volumes for 96, 48 or one 25μl PCR reactions. Amounts for larger master mixeshave been adjusted upwards to be for 50 and 100 reactions compensate for imprecise measurementsGenomic DNA extraction1. Remove microplate from thermocycler and cool to room temperature before proceeding.2. Add 25µl of 5M potassium acetate to each well containing sample.3. Reseal with plastic caps and briefly vortex.4. Centrifuge at 4100rpm and room temperature for 15 minutes to pellet cell debris.5. In a new microplate, add 60µl of 100% isopropanol to each well.6. Carefully remove the supernatant (approximately 60µl) from each well and add to the new microplatecontaining isopropanol. Avoid dislodging cell pellet.7. Seal with new plastic caps and invert 30 times to mix. Microplate containing cell pellets can bediscarded.8. Centrifuge at 4100rpm for 15 minutes at room temperature.9. Carefully remove and discard supernatant. The DNA is should be visible at the bottom of each welland should not be dislodged or removed. The pellet may appear purple in color.10. Add 50µl of 70% ethanol to each well. Replace caps and invert 20 times to wash DNA.

Chapter 3 : Specific Anopheles Techniques3.5 Protocol for 96 Well DNA ExtractionPage 3 of 411. Centrifuge at 4100rpm for 15 minutes at room temperature.12. Carefully remove and discard supernatant as before.13. Allow remaining ethanol to evaporate from microplate by leaving uncovered at room temperature for30 minutes or until no ethanol remains.14. Resuspend DNA in 50µl of TE buffer to each well.15. Seal microplate with new plastic caps and incubate at 65°C for 1 hour tapping periodically.16. DNA samples can be stored at -20°C or -80°C.Figure 3.5.2 shows the result of 23 An. gambiae s.s., 23 An. arabiensis and 23 desiccated mosquitosamples analyzed with the An. gambiae authentication PCR (Wilkins et al. 2006) and 23 An.quadrimaculatus samples analyzed with the Anopheles ITS2 Amplification (Beebe and Saul 1995), using1µl of template genomic DNA extracted with this protocol.Figure 3.5.1. Example of a 96-bacterial replicator tool usedfor maceration of mosquitosamples and reservoir.Figure 3.5.2. Lanes 1, 50, 51, 100 1kb ladder, lanes 25, 49, 75and 99 control wells. Lanes 2-24 An. gambiae s.s., lanes 26-48 An.quadrimaculatus, lanes 51-74 An. arabiensis and lanes 76-79desiccated mosquito samples. Bands are species specific. 5μl ofsample loaded and run on a 2% agarose EtBr gel.ReferencesBeebe NW, Saul A (1995) Discrimination of all members of the Anopheles punctaulatus complex bypolymerase chain reaction-restriction fragment length polymorphism analysis. The American Journal ofTropical Medicine and Hygiene 53:478-481Collins FH, Mendez MA, Rasmussen MO, Mehaffey PC, Besansky NJ, Finnerty V (1987) A ribosomalRNA gene probe differentiates member species of the Anopheles gambiae complex. American Journal ofTropical Medicine and Hygiene 37:37-41Holelley CE (2007) Economical high-throughput DNA extraction procedure in a 96-well format forDrosophila tissue. Dros Inf Serv 90:137-138Wilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect single nucleotide polymorphisms forAnopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin inAnopheles arabiensis. Malar J 5:125

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Chapter 3 : Specific Anopheles Techniques3.6 Determination of Lipid, Glycogen, and Sugars in MosquitoesPage 1 of 43.6 Determination of Lipid, Glycogen and Sugars in MosquitoesAdapted from (Van Handel 1985a; Van Handel 1985b; Van Handel and Day 1988; Kaufmann andBrown 2008)Introduction by C. KaufmannThis method determines the lipid, glycogen, and sugar content of a single insect. It has the advantagethat reasonable results can be achieved with only a few numbers insects, which may be of greatsignificance in the field.The separation of lipid, glycogen and sugar is rather important because vast amounts of ingested sugar,as given via a feeding solution in the laboratory or sugar sources like nectar and honeydew that areingested in nature, can interfere with the lipid analysis (blue/violet coloring of the vanillin test). Also, thehot anthrone test does not differentiate between the ingested sugar within the crop and the carbohydratethat is already transferred to its storage form glycogen.The homogenization with sodium sulfate solution will help to co-precipitate the glycogen after the additionof the chloroform-methanol solution, in which the lipids and sugar dissolve; the addition of water allowsthe separation of the sugar and lipid (sugar in the upper phase and lipid in the lower phase). Be carefulwhen handling the chloroform and acid.Note that for larger insects, you should use different values of chloroform/methanol (2.8 ml instead of 1.6ml) and water (2 ml instead of 0.6 ml); the rest is the same.Materials1. Glass centrifuge tube2. Glass grinding pestle or stir rod3. Heating block at 90-110 o C4. Erlenmeyer flask5. CentrifugeReagents1. Sulfuric acid (95-98%)2. Anthrone3. Anhydrous glucose4. Sodium sulfate5. Methanol6. Commercially available vegetable oil (e.g. soybean oil)7. Vanillin8. Phosphoric acid (85%)9. ChloroformSolutions1. 2% sodium sulfate (NaSO 4 ) solution2. chloroform/methanol mixed 1:1 (v/v)3. Vanillin-phosphoric acid reagenta. Dissolve 600 mg vanillin in 100 mL DI hot water.b. Add 400 mL 85% phosphoric acid.c. Store in the dark. Stable for several months but discard if it darkens.4. Anthrone reagenta. Add 385 mL sulfuric acid (95-98%) to 150 mL DIb. Dissolve 750 mg anthrone

Chapter 3 : Specific Anopheles Techniques3.6 Determination of Lipid, Glycogen, and Sugars in MosquitoesPage 2 of 4c. Store at 4°C. Stable for several weeks.Standards1. Lipid: 100 mg per 100 mL of a commercial vegetable oil (e.g. soy bean oil) in chloroforma) In triplicate, add 50, 100, 200 and 400 μl of solution to glass tube.b) Place in heating block at 90-110 o C to evaporate the solvent.c) Add 0.2 mL of sulfuric acid and heat for 10 min at 90-110 o Cd) Add vanillin reagent to 5 mL level and mix.e) Remove from heating block and allow to cool.f) Allow reddish color to develop; this will take approximately 5 min and will be stable up to30 min.g) Determine OD at 625 nm and plot μg lipid vs. OD for calibration line.2. Sugar and glycogen: 100 mg per 100 mL of anhydrous glucose in deionized water.a) In triplicate, add 25, 50, 100, 150 and 200 μl of glucose solution to glass tubeb) Add anthrone reagent to 5 mL level and mix.c) Heat for 17 minutes at 90-110 o C.d) Remove from heating block and allow to cool.e) Determine OD at 625 nm and plot μg glucose vs. OD for calibration standard.Extraction of Lipid, Glycogen and Sugar Fractions from Mosquito1. Add mosquito to glass centrifuge tube.2. Add 0.2 mL sodium sulfate solution.3. Homogenize mosquito in solution until no identifiable parts remain (glass rod or other utensil).4. Wash glass rod into centrifuge tube with two x 0.8 mL volumes of chloroform/methanol solution.5. Centrifuge (3000 rpm, 1 min).6. Transfer supernatant to clean centrifuge tube. Retain pellet for glycogen analysis.7. Add 0.6 mL DI water to supernatant. Mix.8. Centrifuge (3000 rpm, 1 min).9. Separate top fraction (water/methanol) for sugar analysis.10. Bottom portion (chloroform) holds the portion for lipid analysis.Lipid Analysis1. Place portion for lipid analysis in a tube with a marking at the 5 mL level.2. Place in heating block at 90-110 o C to evaporate the solvent.3. Add 0.2 mL of sulfuric acid and heat for 10 min at 90-110 o C4. Add vanillin reagent to 5 mL level and mix.5. Remove from heating block and allow to cool.6. Allow reddish color to develop; this will take approximately 5 min and will be stable up to 30 min.7. Determine OD at 625 nmSugar Analysis1. Place portion for sugar analysis in a tube with a marking at the 5 mL level.2. Place in heating block at 90-110 o C to evaporate the solvent down to 0.1-0.2 mL.3. Add anthrone reagent to 5 mL level and mix.4. Heat for 17 minutes at 90-110 o C.5. Remove from heating block and allow to cool.6. Determine OD at 625 nmGlycogen Analysis1. Add anthrone reagent to 5 mL level and mix.2. Heat for 17 minutes at 90-110 o C.3. Remove from heating block and allow to cool.4. Determine OD at 625 nm.

Chapter 3 : Specific Anopheles Techniques3.6 Determination of Lipid, Glycogen, and Sugars in MosquitoesPage 3 of 4ReferencesKaufmann C, Brown MR (2008) Regulation of carbohydrate metabolism and flight performance by ahypertrehalosaemic hormone in the mosquito Anopheles gambiae. J Insect Physiol 54:367-377Van Handel E (1985a) Rapid determination of glycogen and sugar in mosquitoes. Journal of theAmerican Mosquito Control Association 1:299-304Van Handel E (1985b) Rapid determination of total lipids in mosquitoes. J Am Mosq Control Assoc 1:302-304Van Handel E, Day JF (1988) Assay of lipids, glycogen and sugars in individual mosquitoes: correlationswith wing length in field-collected Aedes vexans. J Am Mosq Control Assoc 4:549-550

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Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.1 Mating : General ConsiderationsPage 1 of 23.7 Anopheles Mating3.7.1 Mating : General considerationsPaul HowellIntroductionMating in most anopheline mosquitoes occurs during the early evening and is believed to occur primarilyin swarms. Anopheles male mosquitoes aggregate just before dusk and commence swarming at theonset of sunset. Swarming males use their erect antennal fibrillae to detect a nearby female mosquito’swing beat frequencies (Nijhout and Sheffield 1979; Ikeshoji et al. 1985; Leemingsawat 1989; Clements1992). In Toxorhynchites it was found that males will actually harmonize their wing beat with females asthey near, possibly as a form of species recognition, before mating commences (Gibson and Russell2006). In many species, copulation is initiated in flight with males and females meeting within the swarms(Clements 1992). Once a male anopheline has grasped a receptive female, it reorients itself so it is in theventer-to-venter position allowing the reproductive organs to meet. After coitus commences, the malemoves into an end-to-end position with the female as the pair falls (Charlwood and Jones 1979).Copulation may continue for a short period of time after alighting, but in most genera it is a very quickprocess which ceases before the pair reaches the ground.Newly emerged anophelines are not sexually mature. Male mosquitoes require about 24 hours beforetheir terminalia have rotated and their fibrillae are mature enough to become erect and detect females(Clements 1992). Female mosquitoes, however, typically need 48-72 hours before they become receptiveto males - usually prior to blood feeding in the wild. Anopheles males can mate several times, but femalesbecome refractory to re-insemination and re-mating is rare (Villarreal et al. 1994; Yuval and Fritz 1994). InAn. culicifacies, it was found that a proportion of colonized females had multiple inseminations, but thiswas attributed to the laboratory setting and not a natural behavior (Mahmood and Reisen 1980).In the laboratory, it is often not feasible to maintain a colony in a large enough cage to promote naturalswarming behavior. Instead, selection of a stenogamous colony - one that breeds in a small cage - isperformed. During colonization, only a proportion of individuals will respond to the novel environment, anda genetic ‘bottleneck’ occurs - loss of heterozygosity. Norris et al. showed that even a newly colonizedstrain has an extreme loss of heterozygosity compared to field samples from the same area (Norris et al.2001). Often, fixation of particular alleles is a quick inevitable process and cannot be remedied withoutthe introduction of new field material.Although little is known about cues that are needed to stimulate mating within the laboratory, someexperiments have been done to develop methods to improve colony mating. The addition of a simulatedsunrise and sunset has been shown to have a positive effect on colonization efforts – presumably due toimproved mating (Charlwood and Jones 1980; Panicker and Bai 1980). There are also reports ofresearchers utilizing a low watt colored light prior to the dark period to stimulate mating (Pan et al. 1982;Villarreal et al. 1998). Cage size also has an impact on the success of a colony. Some species will notmate within a small cage due to some unknown parameter, such as eurygamy, and require larger cagesto complete mating (Pan et al. 1982; Marchand 1985). Marchand (1985) and Peloquin (1988) utilized anartificial sky and horizon to promote mating.ReferencesCharlwood JD, Jones MDR (1979) Mating behaviour in the mosquito, Anopheles gambiae s.l. I. closerange and contact behaviour. Phys Entomol 4:111-120Charlwood JD, Jones MDR (1980) Mating in the mosquito, Anopheles gambiae s.l. II. Swarmingbehaviour. Phys Entomol 5:315-320

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.1 Mating : General ConsiderationsPage 2 of 2Clements AN (1992) The Biology of Mosquitoes: Development, Nutrition and Reproduction. Chapman &Hall, LondonGibson G, Russell I (2006) Flying in tune: sexual recognition in mosquitoes. Curr Biol 16:1311-1316Ikeshoji R, Sakakihara M, Reisen WK (1985) Removal sampling of male mosquitoes from fieldpopulations by sound-trapping. Jpn J Sanit Zool 36:197-203Leemingsawat S (1989) Field trials of different traps for malaria vectors and epidemiologicalinvestigations at a foot-hill basin in Kanchanaburi, Thailand. Jpn J Sanit Zool 40:171-179Mahmood F, Reisen WK (1980) Anopheles culicifacies: the occurrence of multiple inseminations underlaboratory conditions. Entomol Exp Applic 27:69-76Marchand RP (1985) A new cage for observing mating behavior of wild Anopheles gambiae in thelaboratory. J Am Mosq Control Assoc 1:234-236Nijhout HF, Sheffield HG (1979) Antennal hair erection in male mosquitoes: a new mechanical effector ininsects. Science 206:595-596Norris DE, Shurtleff AC, Toure YT, Lanzaro GC (2001) Microsatellite DNA polymorphism andheterozygosity among field and laboratory populations of Anopheles gambiae ss (Diptera: Culicidae). JMed Entomol 38:336-340Pan JF, Yu TR, Zhu HK (1982) Study on laboratory breeding of Anopheles balabacensis balabacensisBaisas. WHO/MAL/82.989Panicker KN, Bai MG (1980) A note on laboratory colonization of Anopheles subpictus Grassi. Indian JMed Res 72:53-54Peloquin JJ, Asman SM (1988) Use of a modified Marchand cage to study mating and swarm behavior inCulex tarsalis, with reference to colonization. J Am Mosq Control Assoc 4:516-519Villarreal C, Arredondo-Jimenez JI, Rodriguez MH, Ulloa A (1998) Colonization of Anophelespseudopunctipennis from Mexico. J Am Mosq Control Assoc 14:369-372Villarreal C, Fuentes-Maldonado G, Rodriguez MH, Yuval B (1994) Low rates of multiple fertilization inparous Anopheles albimanus. J Am Mosq Control Assoc 10:67-69Yuval B, Fritz GN (1994) Multiple mating in female mosquitoes-evidence from a field population ofAnopheles freeborni (Diptera: Culicidae). Bull Entomol Res 84:137-140

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.2 Forced CopulationPage 1 of 63.7.2 Forced CopulationMR4 StaffIntroductionAdults from several Anopheles - An. dirus, An. funestus, and An. darlingi - have proved difficult speciesfrom which to establish a stenogamous laboratory colony. The main obstacle has been to stimulatemating. In the late 1950’s, McDaniel and Horsfall (1957) developed an induced copulation technique toproduce Aedes mosquitoes that are intractable in the insectary. Their method was based on reportedobservations on the mating behavior of European mantids which decapitated their mates prior tocopulation. When the male mantis was decapitated, the suboesophageal ganglion was severed therebyovercoming the innate inhibition of copulatory muscles in the male (Baker 1964).It is sometimes necessary to use the induced, or ‘forced,’ copulation technique in order to initiate andeven to maintain a colony or to obtain matings with rare or specific individuals. Often, colonies initiated byforced copulation will eventually become stenogamous. Forced mating has been applied to severalmosquito genera. Specific adaptations increase the success rate for mating of anophelines (Baker et al.1962), (Ow Yang et al. 1963). Caravaglios found that males did not need to be decapitated and could besimply held with a small suction pipette, mated, and then returned to the colony without harm(Caravaglios 1961). Conditioning the males by placing them in a 15°C room for 12-24 hours was found toincrease copulation rates (Baker et al. 1962).Materials• Minutien pins mounted on small wooden sticks (10-20)• Ethyl ether• 50 ml Falcon tubes modified to hold mosquitoes (or similar)• Vacuum source with modified tip to prevent mosquitoes from being damaged• Glass container with tight fitting lid (staining jar)• Cotton ballsMethod - modified from the An. maculatus technique (Ow Yang et al. 1963)Success depends on the preparation of high-quality males and females. Both sexes should be of anappropriate age that reflects when the species mates. Males should be at least 72 hours old and femalesat least 48-72 hours old to ensure the reproductive organs have fully matured.1. Two to four hours prior to mating, bloodfeed 3-4 day old females to repletion. Using bloodfed femalesis not absolutely necessary, but it has been reported that the engorged abdomen of the female makesforced copulation more successful (McCuiston and White 1976). Moreover, this ensures that at leastone of the requisites for obtaining progeny has been accomplished!2. Separate males from females and place them in separate containers. 13. Gently aspirate approximately 10 blood fed females into an anesthetizing container (Figure 3.7.2.1).Before proceeding, see Mosquito Anesthesia section (Chapter 3.4). The anesthetizing container canbe made from a 50 ml Falcon tube with the tip removed and both ends covered in mesh held in place1 In some Anopheles species, it is has been found that “seasoning” the males by placing them atapproximately 15-20° Fahrenheit at least 12 hours prior to copulation increases the mating success rate(Ow Yang CK, Sta Maria FL, Wharton RH (1963) Maintenance of a laboratory colony of Anophelesmaculatus Theobald by artificial mating. Mosq News 23:34-35). This is not true for all species.

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.2 Forced CopulationPage 2 of 6with rubber bands or any container that can withstand ether. In a glass container with a lid, place 10-20 cotton balls in the bottom and pour in 5-10 ml of ether. Nitrogen or carbon dioxide gas can be usedas well to anesthetize the females with little to no affect on the rate of successful copulation (Fowler1972).Figure 3.7.2.1. An example of an anesthetizingcontainer.Figure 3.7.2.2. Proper vacuum pressure aspiration ofa male prior to pinning.4. Prepare un-anesthetized males by attaching them to a fine pipette attached to a mild vacuum, the tipof which is just large enough to hold the male by the thorax without damaging it (Figure 3.7.2.2). Theoptimal place to capture a male is on the mesonotum. However, if the vacuum is weak you cancapture the male by slipping the pipette over the male’s abdomen. Caution should be used whencapturing males in this manner as the vacuum may damage the male’s claspers.5. Once the male is captured, gently pierce the side of the thorax with a minutien pin mounted on asmall wooden stick (Figures 3.7.2.3 and 3.7.2.4) e.g. the stick from an oral cotton swab could bemodified with a pin for this purpose. It will be necessary to support the male against a firm surface toenable the pin to penetrate. Prepare 5-10 males at a time in this manner. Use only males that are stillmoving for matings.

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.2 Forced CopulationPage 3 of 6Figure 3.7.2.3. Holding male with vacuumpressure, gently stab the male through thethorax taking care to not crush the head.Figure 3.7.2.4. Properly pinned male.6. Place one container of 10 females into the anesthetizing chamber and leave it for 6-10 seconds(depending on the species and strength of ether).7. Watch the females closely, and once they have all fallen from the sides, remove them from thechamber. Do not leave females in the ether too long or they will not recover from the treatment.8. Gently disperse the females onto a piece of filter paper and position ventral side up. Take a mountedmale, pinch off the head and hind-tarsi (Figure 3.7.2.5), and then gently stroke the abdomen of themale over the female’s abdomen to stimulate the claspers to open (Figure 3.7.2.6).9. Place the male at a 45-90° angle venter-to-venter with the female until the male clasps the female(Figure 3.7.2.7). Leave clasped for 1-2 seconds then pick up both using the male on the pin. If matedsuccessfully, they will remain attached for several seconds. 2 Successful mating has usually occurredif they remain attached for 3-5 seconds (Figure 3.7.2.8).10. Place male and female together into a new cage to allow female to recover from the anesthesia.11. Ensure that the females are recovering from the anesthesia by gently blowing into the recovery cage.If females are not waking up within 10 minutes, anesthetize more lightly.2 It is possible to reuse males to mate more than one female.

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.2 Forced CopulationPage 4 of 6Figure 3.7.2.5 Male with head and legs removed.Figure 3.7.2.6 Positioning of the male prior tostroking the females abdomenFigure 3.7.2.7 Pairing of male and female, note themale is near a 90°angle to the femaleFigure 3.7.2.8 If pairing is successful, you areable to lift the female using the male.

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.2 Forced CopulationPage 5 of 6ReferencesBaker RH (1964) Mating problems as related to the establishment and maintenance of laboratorycolonies of mosquitoes. Bull World Health Org 31:467-468Baker RH, French WL, Kitzmiller JB (1962) Induced copulation in Anopheles mosquitoes. Mosq News22:16-17Caravaglios N (1961) Riv Parassit 22:149Fowler HW (1972) Rates of insemination by induced copulation of Aedes vexans (Diptera: Culicidae)treated with three anesthetics. Ann Entomol Soc Am. 65:293-296Kreutzer RD, Kitzmiller JB (1969) Colonization of Anopheles earlei Vargas. Mosq News 29:589-590McDaniel IN, Horsfall WR (1957) Induced copulation of aedine mosquitoes. Science 125:745Ow Yang CK, Sta Maria FL, Wharton RH (1963) Maintenance of a laboratory colony of Anophelesmaculatus Theobald by artificial mating. Mosq News 23:34-35

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Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.3 Pair MatingsPage 1 of 23.7.3 Pair MatingsMR4 StaffBackgroundPair mating (or single-pair mating) is most commonly used when knowledge of the genetic makeup ofboth parents is important. The isolation of specific phenotypes within Anopheles mosquitoes is useful indetermining vector population species/genetic composition (Rabbani et al. 1976). Phenotypes such asinsecticide resistance as well as other novel genetic mutations can be quickly isolated and purified usingpair mating (Collins et al. 1986).Beside forced copulation (Chapter 3.7.2), mating between a particular male and female can be obtainedby free-mating in small numbers in small containers. Allowing a male and a female to mate freely is lessinvasive and time consuming than the method of forced copulation. Neither method is efficient, however.Benedict and Rafferty (2002) reported a method for obtaining reasonable frequencies of free matings.They observed that mating did not occur until later than is typical, so we recommend keeping the pair inthe mating tube for 7 or 8 days.Materials• Qorpack tubes with modified lids, or similar• Cotton balls• 10% sucrose solution• Rack for holding the single pair mating tubes• AspiratorProcedure1. Ensure the lighting in the space in which the mosquitoes will beheld is adjusted to obtain sunset and sunrise periods to entrain thelarvae before adulthood.2. After emergence, place a male and female in an individual rearingcontainer (Figure 3.7.3.1) and leave for 5-8 days. Additionalfemales may be added if desired – 5 would be a reasonablemaximum.3. Maintain the adults by placing a sugar pad on top of the tube andkeeping it wet.4. Obtain eggs as described in Chapter 3.9 - Family Culture.Figure 3.7.3.1. This particularchoice of tube that we use forfamily oviposition is modifiedfrom a chamber purchasedthrough Qorpack (Bridgeville,PA. No. 3891). Many similartubes are suitable. The bottommay be covered with a filterpaper disk to absorb fluids.ReferencesBenedict MQ, Rafferty CS (2002) Unassisted isolated-pair mating of Anopheles gambiae (Diptera:Culicidae) mosquitoes. Journal of Medical Entomology 39:942-944Collins FH et al. (1986) Genetic selection of a Plasmodium-refractory strain of the malaria vectorAnopheles gambiae. Science 234:607-610Rabbani MG, Seawright JA, Leatherwood LB (1976) A method for culturing single families of Anophelesalbimanus. Mosq. News 36:100-102

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Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.4 An. gambiae and arabiensis Mating Status DeterminationPage 1 of 43.7.4 An. gambiae and arabiensis Mating Status Determination in PreservedFemalesK.R. Ng’habi and Greg LanzaroIntroductionThe An. gambiae mosquito has a karyotype consisting of two pairs of autosomes (Chromosome 2 and 3)and one pair of sex chromosomes (Chromosome X and Y). The Y chromosome constitutes ~10% of thewhole genome and contains a male determining factor which, when present in a XX/XY system, inducesmale development (Clements 1992). Y-chromosome linked DNA fragments have been characterized andY-chromosome specific PCR markers have been developed (Krzywinski et al. 2004; Krzywinski et al.2005). Previously, detection of mating success among females relied on microscopic dissection of femaleovaries or examination of sperm in the female spermatheca. This method is reliable and robust but withthe limitations that it is time consuming, labor intensive, and requires fresh specimens. A simple andrapid method to determine the mating status of dried female An. gambiae is therefore useful in order toanalyze large sample sizes within a short period of time.Prepare PCR mixture for 96, 48 or 1 50µl PCR reactions 1 . Add reagents in the order presented.96 48 1 Reagent3.73 ml 1.87 ml 38.8 µl Sterile water18.63 ml 9.31 ml 5 µl Taq 10X PCR Buffer with MgCl 2 (1.5 mmol/L)3.73 ml 1.87 ml 0.2 µl dNTP (2.5 mmol/L)931.2 µl 465.6 µl 0.25 µl S23 (F, 25pmol) [CAAAACGACAGCAGTTCC]232.8 µl 116.4 µl 0.25 µl S23 (R, 25pmol ) [TAAACCAAGTCCGTCGCT]116.4 µl 58.2 µl 0.5 µl Taq polymerase27.354 ml 13.677 ml 45 µl Total (To each 45µl add 5µl of DNA template)Table 3.7.4.1. F and R indicate forward and reverse primers, respectively. DNA extractions of males mayalso be used as positive controls.96 48 1 Reagent3.73 ml 1.87 ml 38.8 µl Sterile water18.63 ml 9.31 ul 5 µl Taq 10X PCR Buffer with MgCl 2 (1.5 mmol/L)3.73 ml 1.87 ml 0.2 µl dNTP (2.5 mmol/L)931.2 µl 465.6 µl 0.25 µl 128125I (F, 25pmol) [GGCCTTAACTAGTCGGGTAT]232.8 µl 116.4 µl 0.25 µl 128125I (R, 25pmol) [TGCTTTCCATGGTAGTTTTT]116.4 µl 58.2 µl 0.5 µl Taq polymerase27.354 ml 13.677 ml 45 µl Total (To each 45µl add 5µl of DNA template)Table 3.7.4.2. F and R indicate forward and reverse primers, respectively. DNA extractions of males mayalso be used as positive controls.PCR cycle conditions94 o C/3min x I cycle(94 o C/20sec, 55 o C-64 o C/30sec and 72 o C/1 min)* x 35 cycles72 o C/10min x 1 cycle4°C hold1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.4 An. gambiae and arabiensis Mating Status DeterminationPage 2 of 4Run samples on a 3.0 % agarose EtBr gel for visualization. *The primers have different amplificationtemperatures but within this range should work.Figure 3.7.4.1: Agarose gel electrophoresis showing amplification of Y chromosome sequences in An.arabiensis (primer 128125I) males and mated females. There was no amplification in virgin females.Lanes 1 and 26, ladder, lanes 2-13 mated, lanes 14-19 unmated, lanes 20-25 males. (Ng'habi et al.2007), used with permission.Figure 3.7.4.2: Agarose gel electrophoresis showing amplification of Y chromosome sequences in An.gambiae s.s. (Primer S23) males and mated females. There was no amplification in virgin females. Lanes1 and 26, ladder, lanes 2-13 mated, lanes 14-19 unmated, lanes 20-25 males. (Ng'habi et al. 2007), usedwith permission.

Chapter 3 : Specific Anopheles Techniques3.7 Anopheles Mating3.7.4 An. gambiae and arabiensis Mating Status DeterminationPage 3 of 4References:Clements AN (1992) The Biology of Mosquitoes: Development, Nutrition and Reproduction. Chapman &Hall, LondonKrzywinski J, Nusskern DR, Kern MK, Besansky NJ (2004) Isolation and characterization of Ychromosome sequences from the African malaria mosquito Anopheles gambiae. Genetics 166:1291-1302Krzywinski J, Sangare D, Besansky NJ (2005) Satellite DNA from the Y chromosome of the malariavector Anopheles gambiae. Genetics 169:185-196Ng'habi KR, Horton A, Knols BG, Lanzaro GC (2007) A new robust diagnostic polymerase chain reactionfor determining the mating status of female Anopheles gambiae mosquitoes. Am J Trop Med Hyg 77:485-48796 well sample preparation template

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Chapter 3 : Specific Anopheles Techniques3.8 Protocol for Forcing Female Anophelines to OvipositPage 1 of 43.8 Protocol for Forcing Female Anophelines to OvipositJohn MorganIntroductionThis forced oviposition technique has proved extremely useful when collecting Anopheles species fromfield material where large numbers of families are required, e.g. for association mapping and microarraygene expression analyses. However, the method will be of use for any work benefitting from improvedsynchronicity of egg-laying. If this method is used with mosquitoes collected from the field, it is essentialthat all mosquitoes be removed from the tube and preserved before dispatching. On no account shouldlive mosquitoes be transferred using this method because of the potential health hazard they present. Todate, this protocol has been used to obtain eggs from wild-caught resting Anopheles gambiae s.s., A.arabiensis and A. funestus.The forced laying takes place within 1.5 ml Eppendorf tubes which should be prepared as follows.Materials• 1.5 ml Eppendorf tubes• Whatman no.1 filter paper• Dissecting needles(seeker)• ForcepsIn the Field1. Collect resting blood-fed females using an aspirator and transfer to a paper storage cups with aflexible gauze cover. Provide mosquitoes with 10% sugar solution by placing moistened cotton woolon top of the gauze.2. Maintain mosquitoes for 3-5 days to allow them to become gravid. During this period they should bemaintained at ≈25°C and RH >70%.Preparation of Oviposition Containers3. Carefully pierce the Eppendorf tube with 2 holes in the cap and also 2 holes near the base; a seekerproduces holes of the appropriate size (Figure 3.8.1).4. Cut isosceles triangles of Whatman no 1 filter paper approximately 1cm wide at base and 2cm long.5. Insert the paper onto the side of the Eppendorf tube with forceps (Figure 3.8.2).Technique6. Moisten the filter paper with water and tap out any excess water. The amount of water on the paper isintegral to this protocol. (Figure 3.8.3).7. Position the filter paper so that the condition of the mosquito and the presence of eggs can bedetermined without the need to open the tube.8. Using an aspirator, gently introduce a gravid mosquito into each Eppendorf tube (Figure 3.8.4).9. Label the tube with a unique identifier.10. Enclose the tubes in a sealable plastic bag if necessary to maintain humidity and check the tubesdaily to ensure the filter paper remains moist.

Chapter 3 : Specific Anopheles Techniques3.8 Protocol for Forcing Female Anophelines to OvipositPage 2 of 411. When the eggs are seen (Figure 3.8.5), carefully remove the female parent with forceps andpreserve individually over silica gel. A convenient method for storing dry individuals on silica gel is touse pierced numbered 0.2ml color-coded PCR tubes.12. Cohorts of sample tubes are enclosed with silica gel in labeled snap closure plastic bags. Groups(e.g. daily collections) are then enclosed in larger snap bags and transported in sealed plastic boxes.13. Tubes containing eggs should be kept cool in a sealed plastic bag, monitoring the moisture levelbefore dispatch, transportation or synchronized emerging.14. For most purposes it will be preferable to place egg papers into individual containers for hatching inorder to maintain iso-female families and to assess quality and size of families (Figure 3.8.6) see also(Chapter 3.9 Family Culture).Figure 3.8.1 Preparing thetubeFigure 3.8.2 Inserting the filterpaper triangleFigure 3.8.3 Moistening the filterpaperFigure 3.8.4 Aspirating thefemale into the tubeFigure 3.8.5 Checking forovipositionFigure 3.8.6 Single family rearing

Chapter 3 : Specific Anopheles Techniques3.8 Protocol for Forcing Female Anophelines to OvipositPage 3 of 4ReferenceMorgan JC, Irving H, Okedi LM, Steven A, Wondji CS (2010) Pyrethroid resistance in an Anophelesfunestus population from Uganda. PLoS ONE 5(7):e11872. doi:10.1371/journal.pone.0011872

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Chapter 3 : Specific Anopheles Techniques3.9 Family CulturePage 1 of 23.9 Family CultureMR4 StaffIntroductionFamily – or single family - culture is useful for many experimental plans or for the development of newstrains. It is particularly important when the mother’s genetics must be known, in genetic crossing, andwhen genotypic frequencies are of interest. If specific knowledge of the father’s genetics is important tostudies, pair matings must be used (Chapter 3.7.3) or inferred from progeny analysis. Otherwise, enmasse -mated females can be isolated for individual egging. It some cases, the genetics of the motherwill be determined based on the genotypes/phenotypes of her progeny.Another application of family culture is to establish wild colonies. In species complexes such asAn. gambiae, An. funestus and An. dirus where sympatric forms co-exist, it is essential to isolate thespecies of interest (Mpofu et al. 1993). By utilizing family rearing techniques, individual females areseparated so that eggs from a single pair are segregated, thus allowing the establishment of purebreedinglines that can be transformed into laboratory colonies.It is best to culture individual families in the dish in which the eggs were laid for the first couple of days toavoid moving fragile eggs/larvae. The Qorpak vial shown in Figure 3.9.1 are suitable for 100 or fewerprogeny, but larger numbers should be collected in larger cups to prevent early larval mortality. Thelarvae should be progressively transferred to larger containers and water volumes as they develop. It isnot uncommon to culture a family in three different containers before pupation to ensure good survival.A common error is to conclude that one’s family culture method is suitable even in the absence ofhatching data based on total egg hatch. Because L1s may die and decay rapidly, their presence can onlybe known based on a count of the total number of eggs that hatched. Particularly for frequency andsurvival data this is essential. Do not rely on counts of larvae even one day after hatching for quantitativedata.Obtaining eggs from reluctant femalesLow rates of oviposition often hinder successful family culture. Several methods have been developedthat have been shown to increase oviposition within the laboratory. The use of a dark oviposition dish ismore attractive than a clear or white dish in An. quadrimaculatus (Lund 1942), An. gambiae (Huang et al.2005), and An. arabiensis (MQB pers. comm.). In An. albimanus, wild caught blood fed females that wereallowed to oviposit in a 5 dram vial laid more eggs than those allowed to oviposit in a large cage (Baileyand Seawright 1984). The complete removal of one wing of a gravid female that is lightly anesthetized willpromote oviposition soon thereafter, though this is time consuming for the technician. This method is alast resort because mortality results.Rearing schedule for individual familiesIf you are starting with bloodfed material, begin with schedule at day 4.Day 1- Blood feed females.Day 2- No attention is required.Day 3- No attention is required.Day 4- Transfer gravid females to vials lined with filter paper (for example: Qorpack Bridgeville, PA. No.3891 containing strips of filter paper cut to size, Figure 3.9.2) and containing 1-2 cm of water.Day 5- Remove the females from their vials.Day 6- add 2 drops of a 2% w/v yeast slurry to each vial.

Chapter 3 : Specific Anopheles Techniques3.9 Family CulturePage 2 of 2Day 7-count hatch rate and transfer larvae to a larger container (Figure 3.9.2) containing 0.02% w/v finalconcentration of yeast (see Determining Egg Hatch Rates, Chapter 2).Day 8-observe, but feeding is usually not neededDay 9 through pupation-feed ground fish food or other larval diet. Monitor water quality carefully in thesesmaller pans as it is easier to accidentally over feed such a small number of larvae. Transfer to largercontainers as needed.Figure 3.9.1. This particular individual familyoviposition tube is modified from a chamberpurchased through Qorpack Bridgeville, PA.No. 3891.Figure 3.9.2. Examples of family rearing trays.ReferencesBailey DL, Seawright JA (1984) Improved techniques for mass rearing Anopheles albimanus. In: King EG(ed) Advances and challenges in insect rearing. USDA, New OrleansHuang J, Walker ED, Giroux PY, Vulule J, Miller JR (2005) Ovipositional site selection by Anophelesgambiae: influences of substrate moisture and texture. Med Vet Entomol 19:442-450Lund HO (1942) Studies on the choice of a medium for oviposition by Anopheles quadrimaculatus Say. JNatl Mal Soc 1:101-111Mpofu SM, Masendu HT, Kanyimo KH, Mtetwa C (1993) Laboratory colonization of Anophelesquadriannulatus from sympatry with other sibling species of the Anopheles gambiae complex inZimbabwe. Med Vet Entomol 7:122-126

Chapter 4 : Stock Authentication4.1 Authentication by Morphological CharactersPage 1 of 6Chapter 4 : Stock Authentication4.1 Stock Authentication by Morphological CharacteristicsMR4 StaffIntroductionIt is common for laboratories to rear several different species and/or stocks of the same species in oneinsectary. Often it is difficult to determine whether a colony has been contaminated, especially when thestocks appear identical. A PCR method to distinguish four anopheline species based on their 28Sribosomal subunit was developed as a quality control method to ensure contamination had not occurredbetween colonies (Kent et al. 2004). Although this method is highly specific, it can be very costlyperforming several PCR assays to detect a rare contaminant, so it is desirable to develop simple directmethods to verify colony purity.Morphological discrimination of adultsIt is not necessary to have extensive knowledge of mosquito identification to develop methods to keepstocks in order. A simple method to confirm identity is to develop ‘local authentication standards’ basedon morphological characteristics of adults. These standards are not meant to distinguish your mosquitofrom all of those in the world but rather to distinguish the ones you maintain from one another. Thereforethe standards are ‘local’. The features can be described in very general language e.g. large adults, whiteknees, grey. Although these methods are not useful for members of cryptic species complexes likeAn. gambiae, it does work well when several different species of different appearance are maintained.The local authentication standard consists simply of a chart that lists useful morphological characters thatindividually, or in some combination, distinguish all the species you keep. Its creation is simple. Removeseveral male and female members of each species and stun them in the freezer for a few minutes oranesthetize them by some other method. Place them side-by-side under a dissecting scope and scanprominent morphological landmarks (see below) to see if any differ. After selecting some candidatefeature(s), scan larger numbers to ensure all individuals have the characteristic and it can be seen evenin older individuals in which e.g. the scales may have rubbed off. An example of such a local standard isshown in Table 1.Common Morphological Characteristics• Protarsi: In An. gambiae and An. farauti you will usually find three white bands on the distal endof the protarsus. This characteristic is not seen in An. dirus, An. freeborni, or An.quadrimaculatus. Figures 4.1.1-3.• Metatarsi: Unique, species specific, white banding patterns are often seen in An. dirus (whitebanding on the femur-tibia joint) and An. albimanus (prominent broad white bands on metatarsi).Figures 4.1.4-6.• All tarsi: In some species the legs will appear spotted or speckled under magnification. This canbe seen in An. stephensi, An. dirus, and An. farauti.• Abdominal banding patterns: The ventral side of the abdomen can look very similar betweenspecies e.g. An. stephensi and An. gambiae. However, among others, there are various sizes ofbands seen (e.g. An. freeborni have narrow transverse banding while An. quadrimaculatus, An.atroparvus, and An. minimus have wide transverse abdominal banding). Figures 4.1.7-9.• Halteres: We have found that coloration of these structures is a good separation technique for afew strains. An. dirus and An. farauti both have halteres that are black ventrally and whitedorsally. Figures 4.1.10-12.

Chapter 4 : Stock Authentication4.1 Authentication by Morphological CharactersPage 2 of 6• Anterior wing margin: Although the specific banding pattern is highly distinct, often the presenceor absence of dark scaling on the wing margin is enough to distinguish between 2 species.Figures 4.1.13-15.• Palps: Most anophelines have some banding on their palps. The number or width of bands or thelack of bands can be very diagnostic. Figures 4.1.16-18.ProtarsiFigure 4.1.1. An. gambiae. Figure 4.1.2. An. farauti. Figure 4.1.3. An.quadrimaculatus.MetatarsiFigure 4.1.4. An. gambiae. Figure 4.1.5. An. albimanus. Figure 4.1.6. An. dirus.Abdominal pigmentFigure 4.1.7. An. gambiae. Figure 4.1.8. An. dirus. Figure 4.1.9. An. albimanus.HalteresFigure 4.1.10. An. gambiae.Figure 4.1.11. An. farauti(dorsal side).Figure 4.1.12. An. farauti(ventral side).

Chapter 4 : Stock Authentication4.1 Authentication by Morphological CharactersPage 3 of 6WingsFigure 4.1.13. An. gambiae. Figure 4.1.14. An. albimanus. Figure 4.1.15. An.quadrimaculatus.PalpsFigure 4.1.16. An. gambiae. Figure 4.1.17. An.Figure 4.1.18. An. dirus.quadrimaculatus.Once you have made a checklist of which traits each species has, simply tabulate the results to see howthey differ (Table 4.1.1). For example, if you have An. stephensi and An. gambiae, the two can be difficultto separate with most features but can be separated by the heavily spotted tarsi on An. stephensi.Likewise, An. farauti and An. stephensi can be separated from one another based on the presence of aventrally black haltere as seen in An. farauti.STECLA ORLANDO F1 GA/AR/ME/QD FAR1 STE-2Character albimanus quadrimaculatus freeborni gambiae complex farauti stephensiliberally spotted tarsi N N N N Y Yhaltere black ventrally N N N N Y Nprominent broad white band distalon metatarsi Y N N N N Nsooty dark wings N Y N N N Nprominent dark anterior marginwing pigment Y N N Y Y Ymottled abdominal pattern similarto camouflage Y N N N N NTable 4.1.1: A simple table of a few adult morphological characteristics useful to distinguish differentspecies in laboratory settings.Useful traits of immaturesAlthough not as useful as adult characters, there are some unique phenotypes that vary and are easilyobserved. The use of these phenotypes in conjunction with adult characteristics ensures strain purity.Larval / pupal stripe: Anopheles larvae and pupae often have a unique white (Figure 4.1.19) or red(Figure 4.1.20) stripe on their dorsum (French and Kitzmiller 1964; Mason 1967), and its pattern andintensity varies. The An. freeborni F1 colony uniformly carries a white stripe phenotype which is especiallynoticeable in the pupal stage. The red stripe characteristic in An. gambiae is typically sex-limited tofemales. Not every gambiae stock will have this phenotype (e.g. An. arabiensis from Sudan do notexpress this whereas our An. arabiensis from Tanzania do) and the variation itself is a useful observation.

Chapter 4 : Stock Authentication4.1 Authentication by Morphological CharactersPage 4 of 6Figure 4.1.19. An. freeborni white stripe larvae(bottom) shown for comparison with An.quadrimaculatus (top).Figure 4.1.20. An. gambiae larvae shown with(bottom) and without (top) red stripecharacteristic.Larval color: Mutants with differing larval color have been widely reported from An. stephensi in India aswell as An. quadrimaculatus and An. albimanus (Seawright et al. 1985). Often these are genetic, but theyalso may be linked to diet. Culturing a pure-breeding variant-color colony can make separation of thatcolony from others quite easy (Figure 4.1.21).Eye color mutations: Eye color mutants can be separated from wild-eye larvae based on their inability tomelanize when reared in a dark or black pan (See Eye-Color Mutant Screening). Most larvae detect theirenvironment and darken. Eye color mutants cannot discern their backgrounds so they will not melanize(Figure 4.1.22).Collarless trait: Some larva will have a “collar” on the dorsum of the abdomen and some will not (Figure4.1.23) (Mason 1967). Many wild strains are polymorphic for this trait. However, choosing those thateither have the trait or do not to continue a colony can make it easy to quickly note a contamination event.Larval postures when resting on the bottom: These are not definitive by any means. However, somespecies rest differently when compared side by side. An. farauti has a “U” shape, An. gambiae rest in an“L” shape, and some An. atroparvus larvae will appear to rest in a “?” manner (Figure 4.1.24). Otherbehaviors are distinct: An. minimus larvae cluster around the edge of the pan while very few will ventureinto the center. Once you learn what is customary for your strains, watch for changes.

Chapter 4 : Stock Authentication4.1 Authentication by Morphological CharactersPage 5 of 6Figure 4.1.21. Examples from two An. stephensistrains carried simultaneously in an insectary,GREEN1 (left) and STE2 (right). GREEN1 wasselected for a green mutation from the wild typestrain. The green mutant is likely that of Suguna(1981).Figure 4.1.22. An. gambiae (ASEMBO1) larvareared in a white pan (bottom) and a blackpan (top) demonstrating the melanizationcapabilities of this wild-type strain.Figure 4.1.23. Collarless trait shown on dorsumof An. gambiae. Compare the white pigment tocollarless-minus individuals shown in Figure4.1.22.Figure 4.1.24. Curled larval resting postureseen commonly in a disturbed pan of A.farauti.ReferencesFrench WL, Kitzmiller JB (1964) Linkage groups in Anopheles quadrimaculatus. Mosq News 24:32-39Kent RJ, West AJ, Norris DE (2004) Molecular differentiation of colonized human malaria vectors by 28Sribosomal DNA polymorphisms. Am J Trop Med Hyg 71:514-517Mason GF (1967) Genetic studies on mutations in species A and B of the Anopheles gambiae complex.Genet Res, Camb 10:205-217Seawright JA, Benedict MQ, Narang S (1985) Color mutants in Anopheles albimanus (Diptera: Culicidae).Ann Entomol Soc Am 78:177-181Suguna SG (1981) The genetics of three larval mutants in Anopheles stephensi. Indian J Med Res73:120-123

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Chapter 4 : Stock Authentication4.2 Authentication by PCR4.2.1 Anopheles gambiae Thioester-Containing Protein (TEP1) PCR AssayPage 1 of 24.2 Authentication by PCR4.2.1 Anopheles gambiae Thioester-Containing Protein (TEP1) PCR AssayMR4 StaffIntroductionRecent work has shown a relationship between the presence of a TEP1 mutation with an increase inP. berghei parasite mortality in An. gambiae mosquitoes (Blandin et al. 2004). An SNP-based PCR assaywas developed to detect the mutation in An. gambiae mosquitoes. Several MR4 stocks are distinguishedby being pure-breeding for either allele.PCR assay for the TEP1/16 mutation in An. gambiaePrepare PCR Master Mix for 96, 48 or 1 25 μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent785 μl 517.5 μl 7.85 μl sterile H 2 O500 μl 250 μl 5.0 μl 5X Taq PCR Buffer250 μl 125 μl 2.5 μl dNTP (2.5 mM mix)250 μl 125 μl 2.5 μl TEP1 (F, 10pmol/µl) [AAAGCTACGAATTTGTTGCGTCA]250 μl 125 μl 2.5 μl TEP1R (R, 10pmol/µl) [ATAGTTCATTCCGTTTTGGATTACCA]250 μl 125 μl 2.5 μl TEP16 (R, 10pmol/µl) [CCTCTGCGTGCTTTGCTT]100 μl 50 μl 1.0 µl MgCl 2 (25 mM)15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase (5U/μl)2.4 ml 1.2 ml 24 μl Total (To each 24 ul reaction add 1 μl template DNA)Table 4.2.1.1. F and R indicate forward and reverse orientation. Use 1 μl DNA template.PCR cycle conditions94°C/5min x 1 cycle(94°C/30sec, 52°C/30sec, 72°C/45sec) x 40 cycles72°C/5min x 1 cycle4°C holdRun samples on a 2% agarose EtBr gel; load 15 μl sample on gel.This PCR will yield fragments of 372 bp for homozygous susceptible and 349 bp for homozygousrefractory, respectively. Heterozygous samples will have both bands.Figure 4.2.1.1. The TEP1 assay performed on a homozygous andheterozygous samples. Lane 1, 1kb ladder.1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 4 : Stock Authentication4.2 Authentication by PCR4.2.1 Anopheles gambiae Thioester-Containing Protein (TEP1) PCR AssayPage 2 of 2ReferencesBlandin S, Shiao SH, Moita LF, Janse CJ, Waters AP, Kafatos FC, Levashina EA (2004) Complement-likeprotein TEP1 is a determinant of vectorial capacity in the malaria vector Anopheles gambiae. Cell116:661-67096 well sample preparation template

Chapter 4 : Stock Authentication4.2 Authentication by PCR4.2.2 Anopheles arabiensis ND5 Molecular Authentication v 1Page 1 of 24.2.2 Anopheles arabiensis ND5 Molecular AuthenticationLiz WilkinsIntroductionAnopheles arabiensis is a member of the An. gambiae s.l. complex. An. arabiensis strains from differentregions have been found to be variable in the region of the mtDNA dehydrogenase gene subunit 5 (ND5)(Temu and Yan 2005). This region yielded the single nucleotide polymorphisms (SNPs) necessary tocreate a PCR to distinguish two laboratory colonies of An. arabiensis: KGB which originated fromZimbabwe about 1975, and A. arabiensis Dongola which originated from Sudan in 2004. The two aremorphologically indistinguishable in the larval or adult stages and both are susceptible to insecticidesmaking molecular distinction necessary. Part of the ND5 region was amplified and sequenced from eachstrain, and SNP sites were used to create primers using the intentional mismatch primer (IMP) (Wilkins etal. 2006) method of design at the SNP sites. This protocol is useful in distinguishing these two strains onefrom the other.PCR authentication for the members of the Anopheles arabiensis complexPrepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1185 μl 592.5 μl 11.85 μl sterile H 2 O500 μl 250 μl 5.0 μl 5X PCR Buffer200 μl 100 μl 2.0 μl dNTP (2.5 mM mix)100 μl 50 μl 1.0 μl DF (F, 25pmol/μl) GATAAAGCAATAATTTTCTTTAAAGCG100 μl 50 μl 1.0 μl KR (R, 25pmol/μl) GGTGCAAATTTTGAATTTGATTTACAA100 μl 50 μl 1.0 μl IPCF (R, 25pmol/μl) GCATGAGTTAATAAATGAAAAAAAGC100 μl 50 μl 1.0 μl IPCR (R, 25pmol/μl) CTATAACTAAAAGTGCCCAAATTC200 μl 100 μl 2.0 μl MgCl 2 (25mM)15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase (5U/μl)2.5 ml 1.25 ml 25 μl TotalTable 4.2.2.1. F and R indicate forward and reverse orientation.PCR Cycle conditions95°C/5min x 1 cycle(95°C/30sec , 58°C/30sec , 72°C/30sec) x 30 cycles72°C/5min x 1 cycle4°C holdUse a 2% Agarose ethidium bromide gel for visualization.Primers create an internal positive control of 350 bp and a species specific fragment of 260 bp forDongola and 130 bp for KGB (Figure 4.2.2.1).

Chapter 4 : Stock Authentication4.2 Authentication by PCR4.2.2 Anopheles arabiensis ND5 Molecular Authentication v 1Page 2 of 2Figure 4.2.2.1. Lane 1 1Kb marker, lane 2 An. arabiensis KGB,lane 3 An. arabiensis Dongola, Lane 4 1Kb marker.ReferencesTemu EA, Yan G (2005) Microsatellite and mitochondrial genetic differentiation of Anopheles arabiensis(Diptera: Culicidae) from western Kenya, the Great Rift Valley, and coastal Kenya. Am J Trop Med Hyg73:726-733Wilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect single nucleotide polymorphisms forAnopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin inAnopheles arabiensis. Malar J 5:12596 well PCR sample preparation template

Chapter 4 : Stock Authentication4.2 Authentication by PCR4.2.3 Anopheles gambiae white gene v 1Page 1 of 24.2.3 Anopheles gambiae white geneLiz Wilkins, Alice Sutcliffe, Paul HowellIntroductionThe white gene of Anopheles gambiae is located on the X chromosome and known for intron variation,making it a good marker for differentiating closely related species (Mukabayire et al. 2001). In laboratorieswhere several wild stocks from the same sub-species and origin are reared concurrently, it can be difficultto ensure that contamination events between the stocks has not occurred. The MR4 cultures two pairs ofsuch closely related stocks, MOPTI and MALI, ZAN/U and PIMPERENA. An 800 bp segment of intron 3of the white gene ((Besansky et al. 1995); GenBank accession number U29485) was amplified usingprimers WG2 GAGCATCATTTTTTGCTGCG and WG5 CGTGGTTATCGTATCAAAAG as published by(Mukabayire et al. 2001) and sequenced to discover single nucleotide polymorphisms (SNPs) betweenthe stocks (Figure 4.2.3.2.). Primers were designed using the intentional mismatch primer (IMP) method(Wilkins et al. 2006) at these sites for a stock specific authentication method.PCR authentication for stock specific SNP sites in the white genePrepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 196 48 1 Reagent1085 μl 542.5 μl 10.85μl Distilled H2O500 μl 250 μl 5.0 μl 5X GoTaq PCR Buffer200 μl 100 μl 2.0 μl dNTP (2.5 mM concentration)100 μl 50 μl 1.0 μl UFOR (25 pmol/ μl) ATT ATC TGA TGA AGC TTG GAG TCT TTT100 μl 50 μl 1.0 μl UREV (25 pmol/ μl) ATG AAA ATA AGG AGC TTC CTG GCA T100 μl 50 μl 1.0 μl PIMPR (25pmol/ μl) TCA ATG ACA TGA CGT TAT AAT CTG TCT TT100 μl 50 μl 1.0 μl MOPR (25pmol/ μl) CTG TTG TCT TAC AGT AGG GTT ATG T100 μl 50 μl 1.0 μl MOP2R (25pmol/ μl) AAC GTA CGA CGT ATG ATC TAA CTG A100 μl 50 μl 1.0 μl MALR (25pmol/ μl) CTC ATA TTC AAG GAT GAA CAC AAT AC100 μl 50 μl 1.0 μl MgCl2 (25 mM)15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase2.5 ml 1.25 ml 25 μl Total (To each 25 μl reaction add 1 μl template DNA)Table 4.2.3.1. Add reagents in order presented.PCR cycle conditions94°C/5min x 1 cycle(94°C/30sec , 56°C/30sec , 72°C/30sec) x 35 cycles72°C/10min x 1 cycle4°C hold1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 reactionsto compensate for imprecise measurements.

Figure 4.2.3.1 Lane 1 1kb ladder, lane 2Chapter 4 : Stock Authentication4.2 Authentication by PCR4.2.3 Anopheles gambiae white gene v 1Page 2 of 2Run samples on a 2% agarose EtBr gel. (Figure 4.2.3.1).Primers create fragments of 478 universal, 413 mopti, 350 mopti & zan/u,292 mali, 116 pimperenaFigure 4.2.3.2. Specific SNP sites in the white gene used to create this assay. Numbering based onaccession number U29485.SNP sites11591 11767 11826 11885U29485 TTACTATGAC GATTTCTTGT GCTTGTTAGA GACGCTTTACMopti TTACTATGAC GATTTCTTGT GTCTGTTAGA GACACTTAACMaliTTACTATGAC GATGTCTTGT GCTTGTTAGA GACGCTTTACPimperena TTACAATGAC GATTTCTTGT GCTTGTTAGA GACGCTTTACZan/u TTACTATGAC GATTTCTTGT GTCTGTTAGA GACGCTTTACReferencesBesansky NJ, Bedell JA, Mukabayire O, Hilfiker D, Collins FH (1995) The white gene of Anophelesgambiae: structure and sequence of wild type alleles. submittedMukabayire O, Caridi J, Wang X, Toure YT, Coluzzi M, Besansky NJ (2001) Patterns of DNA sequencevariation in chromosomally recognized taxa of Anopheles gambiae: evidence from rDNA and single-copyloci. Insect Mol Biol 10:33-46Wilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect single nucleotide polymorphisms forAnopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin inAnopheles arabiensis. Malar J 5:12596 well PCR sample preparation template

Chapter 5 : Insecticide Resistance Monitoring5.1 Insecticide Resistance Bioassays5.1.1 Larval Insecticide Resistance AssaysPage 1 of 4Chapter 5 : Insecticide Resistance Monitoring5.1 Insecticide Resistance Bioassays5.1.1 Larval Insecticide Resistance AssaysMR4 StaffIntroductionInsecticide-susceptibility defined stocks require routine quality control measures to determine whether acontamination event has occurred. Exposure is also conducted to ensure that the level of resistance hasnot changed. Larval exposures are simple to conduct since they are a non-flying stage and exposurelevels are consistent in the aquatic environment. There are numerous variations for larval exposures, butthe MR4 uses fixed time and dose treatments of L4s routinely because of their ease. Exposure of otherstages and even embryos is also possible using similar methods.In each of these assays, L4s are subjected to a known concentration of an insecticide for a fixed timeperiod (French and Kitzmiller 1963). Because some mortality may occur even among resistant individuals,if the purpose is selection for a resistant colony, a large cohort should be tested to perpetuate the colony.Experience with a particular insecticide will indicate how long the exposure should be and what timeperiods are appropriate for your application. The exact time and concentration necessary for causingmortality or boosting insecticide resistance is best determined utilizing dose-response curves.Some typical discriminating concentrations and treatment times for L4s (solvent).• DDT: 0.4ppm DDT (ethanol) for 24 hours.• Permethrin: 1ppm permethrin (ethanol) for 24 hours.• Propoxur: 20ppm propoxur (acetone) for 1 hour.• Dieldrin: 1ppm dieldrin (ethanol) for 1 hour.• Malathion: 1ppm malathion (ethanol) for 24 hours.Materials 1• Concentrated insecticide from which stock dilution will be prepared (1000 X the final treatmentconcentration is recommended)• Glass graduated cylinders• Volumetric flask1Many of the insecticides will be formulated in solvents that dissolve plastics - especially acetone - soglass bottles and pipettes are recommended.

Chapter 5 : Insecticide Resistance Monitoring5.1 Insecticide Resistance Bioassays5.1.1 Larval Insecticide Resistance AssaysPage 2 of 4• Glass pipettes• Pipetting device• 100 ml glass bottles with screw tops for insecticide stock storage• Dedicated treatment containers. These can be assigned to a particular insecticide, or if they aremade of glass or metal, they can be cleaned thoroughly with solvent for reuse. Polypropylenefood cups and other disposable containers are good for treatments. Treatment containers shouldnot be mixed with colony maintenance trays.• Solvents for dilution of insecticide (see above)• Stainless steel or nylon strainer fine enough to collect L4s.ProtocolExposure:1. Prepare the stock solution of insecticide. 22. Prepare the insecticide dilution in water at the desired concentration.3. Collect early stage L4s ensuring that sufficient numbers for a backup are retained in the event of anunexpected result or accident. For example, MQB once accidentally washed resistant larvae withacetone! Resistance was not observed. Do not expose pharate pupae. They are often more resistantthan L4s. 34. Drain water off of larvae using a fine mesh. Stainless steel kitchen handheld strainers or improviseddevices are suitable.5. Transfer the larvae to the empty treatment pans by tapping the screen containing larvae sharply onthe side of the treatment tray or by washing them in. Attempt to add as little water with the larvae tothe pans as possible (Figure 5.1.1.1) to avoid insecticide dilution. Aspirate out excess water.6. Add insecticide to the pans using a volume similar to that used for larval culture (~ 1 ml / L4) (Figure5.1.1.2). After a few minutes, susceptible larvae can often be recognized by unusual twitchingmovement. Typically, these larvae will not recover and will be classified as moribund (French andKitzmiller 1963).7. For all assays that are longer than 1 hour, feed larvae to ensure mortality is due to the insecticide andnot starvation.8. At the end of the exposure, strain off the larvae and wash thoroughly with water before returning to aculture tray.2 For routine use of insecticides at the same concentration, we have found it convenient to formulateinsecticides at 1000 X the concentration desired. This makes formulation of the final dilution simple (1 mlper liter) and keeps the solvent concentration sufficiently low that it alone does not cause mortality.3 A susceptible reference stock is often necessary to ensure that a lethal dose has been delivered. Forroutine authentication purposes, this may be a stock of a different species.

Chapter 5 : Insecticide Resistance Monitoring5.1 Insecticide Resistance Bioassays5.1.1 Larval Insecticide Resistance AssaysPage 3 of 49. Determine mortality at fixed time intervals (Figure 5.1.1.3). It is often helpful to sink dead larvae bystirring and poking. Affected larvae are usually unable to return to the surface. 4Figure 5.1.1.1. L4 larvae in a dedicated treatmentcup. Most of the water has been removed using aplastic pipette.Figure 5.1.1.2. 200ml of a 1ppm dieldrin solutionwith approximately 200 L4 larvae. Since this is a1 hour treatment, no food has been added.4 There is often excessive concern that insecticides used for treating larvae will kill susceptible stocks inthe insectary due to carryover e.g. on trays, gloves, larvae. While this is possible and should beconsidered, calculation of realistic amounts of insecticide contamination can show that carryover amountsare usually too low to cause mortality even among susceptible individuals.

Chapter 5 : Insecticide Resistance Monitoring5.1 Insecticide Resistance Bioassays5.1.1 Larval Insecticide Resistance AssaysPage 4 of 4Figure 5.1.1.3. 2 hours post treatment, the pan on the left contains the resistant population which isevenly dispersed throughout the container while the pan on the right contains the susceptible control.Note the clumped L4 larvae on the bottom of the pan.ReferencesFrench WL, Kitzmiller JB (1963) Time in concentration: a simple technique for the accurate detection ofresistance to insecticides in mosquito larvae. In. World Health Organization, Geneva, WHO/MAL401

Chapter 5 : Insecticide Resistance Monitoring5.1 Insecticide Resistance Bioassays5.1.2 WHO Paper AssayPage 1 of 25.1.2 WHO Paper TestingMR4 StaffIntroductionThis test involves the use of a specially designed plastic container lined with insecticide impregnatedpapers. Adult mosquitoes are aspirated into the containers forcing exposure to the insecticide for a fixedtime period. The limitations reported for this type of assay include expensive test kits and the inability todetect low frequency resistance in a population (Brogdon 1989). At the WHO website you can find theInsecticide Resistance Monitoring PDF file in which they list how the assays should be run and how todetermine the proper discriminating dose for the species you are handling. The necessary supplies canbe obtained from the WHO using the link: http://www.who.int/whopes/resistance/en/. Examples of theWHO tube are shown in Figure 5.1.2.1.Figure 5.1.2.1. WHO paper insecticide resistance testing tube for adult mosquitoes.ReferencesBrogdon WG (1989) Biochemical resistance detection: an alternative to bioassay. Parasitol Today 5:56-60

Chapter 5 : Insecticide Resistance Monitoring5.1 Insecticide Resistance Bioassays5.1.2 WHO Paper AssayPage 2 of 2

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 1 of 245.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using theCDC Bottle BioassayWilliam Brogdon and Adeline ChanPREFACEInsecticide resistance in a vector population is initially detected and characterized by using some sort ofbioassay to determine whether a particular insecticide is able to control a vector at a given time. Ideally,this fundamental question should be answered before a particular insecticide is chosen and procured forvector control.The Centers for Disease Control and Prevention (CDC) bottle bioassay is a surveillance tool for detectingresistance to insecticides in vector populations. It is designed to help determine if a particular formulationof an insecticide is able to control a vector at a specific location at a given time. This information,combined with results of bioassays using synergists and those of biochemical and molecular assays, canassist in determining which insecticide should be used if resistance is detected.The aim of this document is to provide a practical laboratory guideline that describes how to perform andinterpret the CDC bottle bioassay. Information for resistance testing can also be obtained from the CDCwebsite at http://www.cdc.gov/malaria.We hope you find this tool useful in the support of vector control programs.Sincerely,William G. Brogdon, PhDResearch EntomologistEntomology BranchDivision of Parasitic DiseasesCenters for Disease Control and Prevention4770 Buford Hwy, MS F-42Atlanta, GA 30341 USATel: +1 770.488.4523Fax: +1 770.488.4258Email: WBrogdon@cdc.govAdeline Chan, PhDResearch EntomologistEntomology BranchDivision of Parasitic DiseasesCenters for Disease Control and Prevention4770 Buford Hwy, MS F-42Atlanta, GA 30341 USATel: +1 770.488.4987Fax: +1 770.488.4258Email: AChan@cdc.govACKNOWLEDGEMENTWe wish to thank the enormous number of people who have contributed both to the development andimplementation of this method. Particular thanks go to the United States Agency for InternationalDevelopment (USAID) and the Pan American Health Organization (PAHO) through the Amazon MalariaInitiative (AMI) for adopting this method as part of this initiative. Scientists and control program personnelfrom many countries have expended great energy and resources in the collection of field data thatallowed both refinement and realistic evaluation of the method alongside other methods, such as theWorld Health Organization (WHO) paper-based bioassay. While too many to mention individually, thesepeople have our grateful thanks. Finally, we would like to thank Alexandre Macedo de Oliveira, MD, PhDand Beatie Divine, MA, MBA for the careful review of this document and their input in the editorialprocess.

INTRODUCTIONChapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 2 of 24Bioassays allow for the detection and characterization of insecticide resistance in a vector population.This guideline will describe the Centers for Disease Control and Prevention (CDC) bottle bioassay, a toolfor detecting resistance to insecticides. The information provided by this bioassay, combined with resultsof bioassays using synergists and those of biochemical and molecular assays, can also assist indetermining mechanisms associated with resistance.The CDC bottle bioassay relies on time mortality data, which are measures of the time it takes aninsecticide to penetrate a vector, traverse its intervening tissues, get to the target site, and act on thatsite. Anything that prevents or delays the compound from achieving its objective — killing insects —contributes to resistance. Information derived from the CDC bottle bioassay may provide initial evidencethat an insecticide is losing its effectiveness. This methodology should be considered for routine useeven before an insecticide is considered, and procured, for vector control.The CDC bottle bioassay can be performed on vector populations collected from the field or on thosereared in an insectary from larval field collections. It is not recommended to use mosquitoes that haveemerged from eggs laid in the insectary.A major advantage of this bioassay is that different concentrations of an insecticide may be evaluated.Furthermore, the technique is simple, rapid, and economical compared to other alternatives. The CDCbottle bioassay can be used as part of a broader insecticide resistance monitoring program, which mayinclude the World Health Organization (WHO) paper-based bioassay, and biochemical and molecularmethods.The CDC bottle bioassay can be used for any insect species. For the purposes of this guideline,mosquitoes will be used as an example.Material and Reagents250 ml Wheaton bottles with screw lids (Figure 5.1.3.1). Each bioassay typically requires fivebottles: four for replicates and one for control;Graduated disposable plastic pipettes that can measure 1ml; or micropipetters and tips;Aspirator apparatus for collecting mosquitoes;Containers for transferring/holding mosquitoes;Bottles for stock solutions. These can be amber-colored or foil-wrapped if clear bottles are used(100–1,000ml depending on the user’s choice of stock solution volume);Timer capable of counting seconds;Permanent markers for labeling bottles, caps, and pipettes;Masking tape for labeling bottles, caps, and pipettes;Disposable gloves;Sheets, pens, and pencils for data recording.ReagentsInsecticide(s) to be tested (technical grade or formulations);Acetone or technical grade absolute ethanol.NB: Use safety procedures as recommended by your institution when handling insecticides (e.g.,procedure gloves, laboratory coat).

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 3 of 24Figure 5.1.3.1: Material and reagents for the CDC Bottle bioassayInitial considerations: Diagnostic dose and diagnostic timeThe first step in standardizing the CDC bottle bioassay is to determine the diagnostic dose and thediagnostic time. The diagnostic dose is a dose of insecticide that kills 100% of susceptible mosquitoeswithin a given time. The expected time for the insecticide to achieve this objective is called the diagnostictime. Those are the reference points against which all other results are compared. Resistance isassumed to be present if a significant portion of the test population survives the diagnostic dose at thediagnostic time.The diagnostic dose and the diagnostic time should be defined for each insecticide, each region, andeach vector species that is monitored. The diagnostic dose and the diagnostic time are validated using asusceptible population of vectors collected from the field. Once the diagnostic dose and the diagnostictime for a species from a given location have been determined, these parameters should be used fortesting that particular vector population from that location from that time on. Use of the same parametersis required to detect changes in the response of the population over time (e.g., number of test mosquitoessurviving after an exposure time that originally killed 100% of the test population). Detailed informationabout diagnostic doses, diagnostic times, and calibration of the CDC bottle bioassay is given in Appendix2.Diagnostic doses and diagnostic times have been determined for mosquitoes from many geographicalregions. Table 5.1.3.1 shows diagnostic doses and diagnostic times applicable to Anopheles and Aedesmosquito populations. The diagnostic doses and the diagnostic times for anophelines shown below wereagreed upon for use on anophelines collected in South America as part of the Amazon Malaria Initiative(AMI). These doses and times, as well as those listed for Aedes, are well within the range of diagnosticdoses and diagnostic times for use worldwide. Therefore, the diagnostic doses and the diagnostic timesin Table 5.1.3.1 serve as sample reference points for the main insecticides used globally. Diagnosticdoses and diagnostic times for other insect species may still need to be determined.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 4 of 24Table 5.1.3.1: Sample diagnostic doses and diagnostic times for Anopheles and Aedesmosquitoes.InsecticideInsecticide concentrationper species (µg/bottle)AnophelesAedesDiagnostic time(minutes)Bendiocarb 12.5 12.5 30Cyfluthrin 12.5 10 30Cypermethrin 12.5 10 30DDT 100 75 45Deltamethrin 12.5 10 30Fenitrothion 50 50 30Lambdcyhalothrin 12.5 10 30Malathion 50 50 30Permethrin 21.5 15 30Pirimiphos-methyl 20 -- 30In summary, determining the diagnostic dose and the diagnostic time is the first step to standardize theCDC bottle bioassay. This step should be done at the national or regional level in a given country orregion to allow for comparability among different laboratories over time. Once the diagnostic dose anddiagnostic time are agreed upon for a particular insecticide and mosquito species, there is no need toredo this exercise until evidence of high levels of resistance in this species is documented.Preparation of stock solutionsThe bottles used for the bioassay need to be coated inside with the diagnostic dose of the insecticideunder evaluation. As can be seen from Table 5.1.3.1, the diagnostic dose is a determined amount ofinsecticide per bottle. Therefore, if 12.5µg of deltamethrin is to be added to a testing bottle, it would beadvisable to have a stock solution with 12.5µg/ml, which means that 1 ml of the solution would contain thedesired amount of insecticide to be added to the bottle. This is equivalent to saying that it is practical tomake stock solutions with concentrations that can be easily correlated to the dose needed to coat thebottles.To make insecticide stock solutions, dilute the appropriate amount of insecticide (technical grade orformulation) in acetone or technical grade ethanol. Examples of quantities of technical grade insecticideneeded to prepare 100 ml, 500 ml, and 1,000 ml of stock solutions are shown in Table 5.1.3.2. Technicalgrade insecticide may be solid or liquid and need to be of good quality and not be expired. It is importantto label the stock solution bottle with the name of the insecticide, concentration, and date of preparation.Examples of preparation of stock solutions from technical grade and formulations are shown in Box 1.Once the stock solution is made, it can be stored in the refrigerator (4°C) in light-proof bottles (ambercoloredbottles or foil-wrapped if clear) for future use. At the CDC, refrigerated stock solutions of many

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 5 of 24insecticides have been used for 2–3 years without degradation of activity. It is recommended to take thestock solutions out of the refrigerator at least 1 hour before running the bioassay to allow them to come toroom temperature before use. The stock solution should be gently swirled before use to mix it.Table 5.1.3.2: Quantities of technical grade insecticide required for preparation of differentvolumes of stock solution.InsecticideWeight (mg) of technical gradeinsecticide needed per volume ofstock solution (Anopheles)Weight (mg) of technical gradeinsecticide needed per volume ofstock solution (Aedes)100 ml 500 ml 1,000 ml 100 ml 500 ml 1,000 mlBendiocarb 1.25 6.25 12.5 1.25 6.25 12.5Cyfluthrin 1.25 6.25 12.5 1 5 10Cypermethrin 1.25 6.25 12.5 1 5 10DDT 10 50 100 7.5 37.5 75Deltamethrin 1.25 6.25 12.5 1 5 10Fenitrothion 5 25 50 5 25 50Lambdcyhalothrin 1.25 6.25 12.5 1 5 10Malathion 5 25 50 5 25 50Permethrin 2.15 10.75 21.5 1.5 7.5 15Pirimiphos-methyl 2 10 20

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 6 of 24Box 1: Examples of stock solution preparation.1. Preparing stock solutions from technical grade insecticideAssume a 100% deltamethrin technical grade insecticide. To obtain a concentration of 12.5µg/bottle, dissolve 12.5 mg the insecticide in enough acetone or absolute ethanol to make 1 liter oftotal solution. Each 1 ml of this solution will contain 12.5 µg of the insecticide. Stock solutions ofvarying volumes (for example 100 ml) can be made for the convenience of the user as long as theproportion of insecticide and solvent remains constant.2. Preparing stock solutions from concentrations other than technical gradeTo calculate the volume of a formulation to be added to the solvent to reach the desiredconcentration in the stock solution, it is necessary to consider the concentration of the activeingredient in the formulation. To do that, simply divide the desired amount of milligrams needed for1 liter of the stock solution by the concentration in the formulation available. This will give thevolume of the formulation needed to make 1 liter of stock solution. The formula:Milligrams of technical grade% of active ingredient in formulationExample:Using a formulation of 10% deltamethrin, calculate how much is needed to obtain 12.5 mg.Volume needed = 12.5 mg = 125 mg of the 10% formulation0.10So, 125 mg of the 10% concentration of deltamethrin will need to be added to enough acetone orabsolute ethanol to make 1 liter of total solution. By doing so, each 1 ml of the stock solution willcontain 12.5 µg of deltamethrin.Mosquito handlingFemale mosquitoes to be used in the bioassay can be collected as adults from the field (of mixed age andphysiological status) or as adults of a known age reared from field larval collections. Use of mosquitoesthat have emerged from eggs laid in the insectary is not recommended. If field-collected adults are used,their physiological status (i.e., unfed, blood fed semi-gravid, gravid) should be recorded on the resultsheet. Female mosquitoes should be fed only with 10% sugared solution the day before testing. It isrecommended that a minimum of 100 mosquitoes, divided among four replicate bottles, should be testedfor an insecticide at a given concentration. When it is not possible to collect this number of mosquitoeson a single occasion, results of multiple bioassays over a few days may be pooled to achieve therecommended sample size, 100 mosquitoes. In either case, each bioassay must include a control bottlewith 10–25 mosquitoes.Some field collections may contain different species. Therefore, species must be identified either beforeor after the bioassay is conducted to validate its results (Box 2). To determine the species composition ofmosquito collections before the bioassay, it is possible to “knock down” (anesthetize) mosquitoes withethyl acetate.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 7 of 24Box 2: Guidelines for situations where different mosquito species exist in samplecollections.In those situations where different mosquito species exist, it is recommended that species beidentified, either before or after the CDC bottle bioassay. If a predominant species is detected(i.e., more than 95% belong to one single species), consider this the species tested, and theresults of the CDC bottle bioassay can be considered adequate for the predominant species.If no particular species represents at least 95% of the mosquitoes being tested, account forthis heterogeneous population. To achieve this:1. Identify the species and sort before the bioassay using ethyl acetate. Conduct separatebioassays for each, or predominant, species; or2. Start the bioassay without pre-identification if this is not possible (lack of expertise withmosquito “knock down,” or presence of closely related or cryptic species). If there aresurviving mosquitoes at the diagnostic time, stop the bioassay and separate live fromdead mosquitoes. Identify the species for both live and dead mosquitoes, and considerthem separately for analysis.Procedures for cleaning and drying bottles before coating1. Wash the bottles with warm soapy water and rinse thoroughly with water at least three times.Tap water can be used for this step;2. Place bottles in an oven (50°C) for 15–20 min or until they are thoroughly dry before using them;3. If there is no oven, leave bottles to dry completely at room temperature or in the sun, with thecaps off. In humid situations, bottles can be left to dry with caps off overnight or longer;4. To assure that the cleaning procedure is adequate, introduce some susceptible mosquitoes into asample of recently washed and dried bottles. Mosquitoes should not die right away. If they do,repeat the washing and drying procedure.Marking of bottles1. Since the bottles will be reused, consider using a piece of masking tape on the bottles and capsfor marking them instead of writing directly on the bottles and caps (Figure 5.1.3.2). This mayfacilitate the cleaning of the bottles after the bioassay is completed;2. Mark one bottle and its cap as control;3. Mark the other four bottles and caps with the replicate number (1–4) and the bioassay date;4. If more than one type of insecticide or more than one concentration of the insecticide is beingtested at the same time, also label the bottles and their caps with the insecticide name andconcentration;5. Mark both the cap and the bottle so that bottles are associated with their respective caps. This isvitally important because the inside of the entire bottle will be coated, including the inside of thecap.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 8 of 24Figure 5.1.3.2: Labeling bottles and capsBottle coating1. Make sure that bottles and caps are completely dry;2. Remove caps from the bottles;3. If using disposable pipettes, label one pipette as ‘solvent only’ for the control bottle, and anotherpipette as ‘insecticide solution’ for the test bottles;4. Add 1 ml of acetone/ethanol to the control bottle and put the cap back on tightly;5. Add 1 ml of the stock insecticide solution to the first test bottle and put the cap back on tightly.This step is to be done if the stock solution has the same concentration per ml as is desired forthe bottle (Table 5.1.3.1);6. Repeat Step 5 with the other three test bottles;7. Swirl the contents inside the bottle so that the bottom is coated (Figure 5.1.3.3);8. Invert the bottle and swirl to coat the inside of the cap (Figure 5.1.3.4);9. Place the bottle on its side for a moment to let the contents pool. Gently rotate while rocking thebottle gently so that the sides all the way around are coated;10. Repeat this for all the test bottles (Figure 5.1.3.5);11. Remove the caps and continue rolling bottles on their side until all visible signs of the liquid aregone from inside and the bottles are completely dry (Figure 5.1.3.6);12. Leave bottles on their sides and cover with something that will keep them protected from light;13. If bottles are not used right away, store bottles in a dark place (such as a drawer) with the capsoff to avoid moisture build-up. If shipping pre-coated bottles, ship the bottles with the caps on.More information on the storage of coated bottles is given on page 11 of this chapter.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 9 of 24Figure 5.1.3.3: Coating the bottom of the bottleFigure 5.1.3.4: Coating the top of the bottleFigure 5.1.3.5: Coating the sides of the bottleFigure 5.1.3.6: Removing caps and rolling thebottles on their sidesCDC bottle bioassay methodGeneral considerations1. Use a filter in the aspirator to avoid inhaling mosquitoes or insect fragments;2. Blow gently to expel the mosquitoes into the bottles. If you blow too hard, the mosquitoes can bedamaged by hitting the sides of the bottle and killed before the insecticide has a chance to do so;3. Be careful not to touch the inside of the bottle with the aspirator, as this may contaminate theaspirator;4. Remember that the number of mosquitoes in the each of the test bottles does not need to beequal;5. Determine species composition of mosquitoes either before or after the bioassay is conducted(Box 2).

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 10 of 24Bioassay procedureThe bioassay can be performed with the bottles in an upright position or with the bottles lying on theirsides. The important thing is to be consistent and follow the same procedure each time.The steps:1. Using an aspirator, introduce 10–25 mosquitoes into the control bottle. It is not necessary tocount the mosquitoes; the exact number does not matter;2. Introduce 10–25 mosquitoes into each test bottle; again, the exact number does not matter(Figure 5.1.3.7);3. Start a timer. Be sure to examine the bottles at Time 0 and count the number of dead and/or livemosquitoes;4. If you find dead mosquitoes at Time 0, make a note of them on the form (Appendix 3);5. Record how many mosquitoes are dead or alive, whichever is easier to count, every 15 minutesuntil all are dead, or up to 2 hours (Figure 5.1.3.8). It is not necessary to continue the bioassaybeyond 2 hours;6. Record these data on the reporting form (Appendix 3);7. Graph the total percent mortality (Y axis) against time (X axis) for all replicates consideredtogether using a linear scale;8. Remember that mortality at diagnostic time is the most critical value because it represents thethreshold between susceptibility and resistance. Refer to Table 5.1.3.1 for diagnostic doses andtimes for commonly used insecticides;9. Take into consideration mortality in the control bottle at 2 hours (end of the bioassay) whenreporting the results of the bioassay (See validity of the data section below). Use Abbot’s formulato correct results if the mortality at 2 hours in the control bottle is between 3% and 10%. You mayneed to discard the bioassay results if mortality in the control bottle at the end of the test was>10%.Mosquitoes are considered dead if they can no longer stand. See Box 3 for more information.A timer could be started for each bottle, but it is sufficient to start one timer when the first or last bottlereceives its mosquitoes. It is, however, important to be consistent and follow the same timer startprocedure each time. Mosquitoes alive at the diagnostic time (Table 5.1.3.1) represent mosquitoesresistant to the insecticide being tested. These mosquitoes may be transferred to a sleeved carton forfurther analysis (e.g., molecular or biochemical assays). Mosquitoes flying at the end of the bioassay inthe control bottle may need to be killed to get an accurate count. Mosquitoes can be killed by freezing orstunning them.Box 3: Notes about mortality criteria. “Dead” mosquitoes are mosquitoes that cannot stand. It helps to gently rotate the bottle while taking the count. Immobile mosquitoes that slide along the curvature of the bottle can be easily categorizedas dead. It is easier to count the number of dead mosquitoes in the first readings of the bioassay, andit is easier to count the number of live mosquitoes when few remain alive. In the end, the percentage of dead mosquitoes at the diagnostic time (dead mosquitoes/total of mosquitoes in the assay) is the most important value in the graph.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 11 of 24Figure 5.1.3.7: Transferring mosquitoes intoinsecticide-coated bottlesFigure 5.1.3.8: CDC bottle bioassay in progressHandling of coated bottlesMore than one batch of mosquitoes can be run in a single bottle in one day. However, the main limitingfactor for reusing previously coated bottles is moisture build-up with successive introductions ofmosquitoes, especially in humid conditions. If the bottles are to be reused on the same day, it isnecessary to leave some time (2–4 hours, longer if in a humid climate) between the bioassays for thebottles to dry out (with caps off) before introducing more mosquitoes. If the bottles are to be reused thefollowing day, bottles with caps off can be left to dry overnight protected from direct light. It is prohibitedto dry bottles in the oven after they have been coated with insecticide.If the bottles are not to be used soon after coating them with insecticide, it is recommended to let themdry with their caps off. When the bottles are dry, they should be stored in a dark place (such as a drawer)with their caps off. Depending on the insecticide used, bottles can be stored from 12 hours to 5 days inthis manner. The length of time bottles can be stored depends on the insecticide. Resmethrin- andNaled-coated bottles do not store well, so they should be used immediately after being prepared.Organophosphate-coated bottles should be used within 24 hours. To check if a stored bottle is stilladequate, it is possible to put some mosquitoes known to be susceptible in the bottle. If they die in theexpected time frame (within the diagnostic time), the bottle can still be used. Bottles can be coated in acentral laboratory and shipped for use in the field. During transport, bottles should have their caps on.Mosquito preparation for testing to determine mechanisms of insecticide resistanceResistance is assumed to be present if a portion of the test population survives the diagnostic dose at thediagnostic time. Mosquitoes that survive the bioassay can be used for testing to identify mechanisms ofresistance using enzymatic assays or molecular methods. Surviving mosquitoes may be easily releasedfrom bottles into a sleeved holding carton to separate them from those killed during the CDC bottlebioassay. Mosquitoes that will be further tested using enzymatic assays should be stored frozen.Mosquitoes to be used for molecular studies can be frozen, dried, or stored in 70% (or higher) ethanol. Inaddition, it may be necessary to use products like RNALater® (Applied Biosystems [Ambion], Foster City,California) to preserve samples for measurement of RNA levels associated with up-regulated enzymemechanisms.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 12 of 24Validity of the dataWith practice, the mortality of mosquitoes in the control bottle at 2 hours (end of the bioassay) should bezero. In most cases, mortality of up to 3% in the control bottle may be ignored. In cases where mortalityis 3%–10% in the control bottle at 2 hours, it is possible to either use Abbot’s formula to correct thefindings (see Box 4), or discard results and repeat the bioassay. If mortality in the control bottle is greaterthan 10% at the end of the bioassay, the results of this particular run should be discarded, and the CDCbottle bioassay should be repeated. If a particular mosquito collection is essentially irreplaceable and thebioassay cannot be repeated, Abbot’s formula can be considered even when control mortality is >10%.Box 4: Abbott's formula.Corrected mortality = (mortality in test bottles [%] - mortality in control bottle [%]) x 100(100% - mortality in control bottle [%])For example: If mortality in test bottles is 50% at diagnostic time and control mortality is 10% at 2hours, the corrected mortality is [(50%-10%) / (100%-10%)] x 100 = 44.4%Note: In cases of 100% mortality in test bottles, Abbott’s formula has no effect. For example:[(100% - 10%) / (100% - 10%)] x 100 = 100% corrected mortalityInterpretation of resultsAs with other resistance bioassays, data from the CDC bottle bioassay using test mosquitoes need to becompared with data from susceptible mosquitoes or from a population that will serve as baseline.Resistance thresholds for each insecticide can be determined by calibrating the CDC bottle bioassay(Appendix 2). Calibration entails determining the diagnostic dose and the diagnostic time for a particularspecies in a given region, which correspond to the dose and time at which all of susceptible mosquitoesdie (Figure 5.1.3.9). If test mosquitoes survive beyond this threshold, these survivors represent aproportion of the population that has something allowing them to delay the insecticide from reaching thetarget site and acting. In other words, they have some degree of resistance. In the example shown inFigure 5.1.3.9, all mosquitoes that died before the diagnostic time when exposed to insecticide-coatedbottles were susceptible. Test mosquitoes surviving beyond the diagnostic time threshold were assumedto have some degree of resistance. In the example, only 23% of the test population was susceptible.Recommendations for interpretation of bioassay data are shown in Box 5. The most importantinformation is the mortality at the diagnostic time, but the bioassay is carried out beyond the diagnostictime to evaluate the intensity of resistance.Box 5: Interpretation of data for management purposes.WHO recommendations for assessing the significance of detected resistance: 98%–100% mortality at the recommended diagnostic time indicates susceptibility; 80%–97% mortality at the recommended diagnostic time suggests the possibility of resistancethat needs to be confirmed;

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 13 of 24Figure 5.1.3.9: Determination of resistance threshold.Mortality (%)1009080706050403020100SusceptibleResistant0 15 30 45 60 75 90 105 120Time (min)Resistance surveillanceBackgroundAlthough resistance data are often collected as part of vector control programs, this is often not done asregularly as it should be in a true resistance surveillance effort. Surveillance requires the regularcollection and interpretation of epidemiological data to support changes in public health programs. It isimportant to consider the CDC bottle bioassay an instrument to collect information to support aninsecticide resistance surveillance system. Resistance data are most valuable when collected over timeto allow for comparisons and for monitoring of trends.It is important to consider how information collected as part of an insecticide resistance surveillancesystem will be used. Most malaria control programs carefully assess the efficacy of their vector controlprogram by, for example, plotting incidence of malaria cases or by counting adult mosquitoes or larvalcollections at sentinel sites. The integration of insecticide resistance data and other kinds of malariarelateddata needs to be taken into consideration before proposing and implementing remediationstrategies for insecticide resistance.Features of resistance emergenceSeveral genetic, biologic, and operational factors influence the development of insecticide resistance. Inmany respects, resistance is a complex problem, with different outcomes possible in a particular area,depending on the influence of diverse factors on initial conditions. Even so, certain factors affectresistance development throughout the world. Major resistance characteristics are discussed below,showing why each manifestation of resistance is potentially unique and therefore must be evaluated oncase-by-case basis.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 14 of 24Focal nature of resistanceVector control personnel frequently assume that resistance in a particular species occurs throughout theircontrol area, but insecticide resistance can be focal. In Guatemala, sampling sites for Anophelesalbimanus only a few kilometers apart varied not only by presence or absence of resistance, but also bylevel of resistance and by dominant mechanism responsible for resistance. Generally speaking, areas ofongoing vector control activities tend to have higher levels of resistance; when resistance levels inadjacent areas are compared, levels may be higher in areas of more intensive mosquito control.Resistance and disease controlIn some cases, vector control strategies in a given area may not be affected by the level of insecticideresistance. For example, a control program may be able to control only 75% of the vector population. Inthese cases, an insecticide resistance level lower than 10% will likely not affect disease control efforts. Insuch a situation, it would be sufficient to increase surveillance and monitor the level and frequency ofresistance but no change in control strategies would be needed.Guiding principlesIn general terms, resistance surveillance should be conducted in areas where disease transmission is aconcern and where insecticide-based control measures are contemplated, ideally before purchase ofinsecticide. In addition to constraints imposed by economic resources, the number of sites that can besampled is highly dependent on the size of the area contemplated for insecticide use. Due to thepotential focal nature of resistance, efforts must be made to choose spatially distributed sites in the areaof interest, if possible. Areas 20 km or more apart should not be assumed to have similar resistancepatterns. Another means of deciding on surveillance sites is to focus on those areas of active diseasetransmission. Even if only one or a few sites can be monitored, this is far preferable to having nosurveillance sites. In addition, efforts should be made to operate sites for at least a few years, sincecomparative data are the most meaningful information.Ideally each site should be monitored once a year. Where control efforts are seasonal, it may be usefulto monitor at the beginning and at the end of the control season. This does not apply to situations suchas the use of insecticide-treated bednets, where the insecticide exposure is year round. If several vectorsin the area are seasonal, the resistance testing schedule should be adjusted to the species of interest.It is also important to consider that it will be necessary to try to identify resistance mechanisms onceresistance is detected with the CDC bottle bioassay, whether using the CDC bottle bioassay withsynergists, or biochemical and/or molecular methods. Decisions on which insecticide to change to willdepend upon the specific mechanism(s) of resistance.Finally, some countries have found it useful to centralize preparation of bottles and administrativeorganization of surveillance. A central reference laboratory can provide support for technical assistanceand quality assurance. It can also serve as a reference laboratory for training, provision of supplies,species identification, and enzymatic assays and molecular methods for determination of resistancemechanisms.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 15 of 24CDC bottle bioassay and synergistsBackgroundThe CDC bottle bioassay using bottles that were coated with a single insecticide provides information oninsecticide resistance to that particular insecticide in adult vectors. These data may provide earlyevidence that an insecticide is losing its effectiveness.Once resistance is detected, or at least suspected, one must decide what to do next and which othercompounds are likely to still be effective and not compromised by cross resistance. This requiresknowledge of the resistance mechanism(s) in place; information usually acquired using either biochemical(microplate) assays or molecular methods. A rapid and inexpensive alternative to assess resistancemechanisms is to use the CDC bottle bioassay with synergists. Synergists are enzyme inhibitors ofinsecticide detoxification enzymes. Synergists are available for the metabolic detoxification enzymes:esterases, oxidases, and glutathione s-transferases.Synergists act by abolishing the apparent resistance observed in the CDC bottle bioassay if adetoxification enzyme plays a role in that particular resistance mechanism (Figures 5.1.3.10a and5.1.3.10b). Data for resistant and susceptible populations are shown (Figure 5.1.3.10a). Once asynergist is used on the resistant population, one of three things might happen (Figure 5.1.3.10b):a. Resistance to the insecticide is abolished (time-mortality line A), which suggests that themechanism related to that synergist is playing a role in the insecticide resistance observed;b. Resistance to the insecticide is partially abolished (time-mortality line B). This suggests that themechanism related to that synergist is involved in the resistance, but it is not the only mechanisminvolved in this particular case;c. Resistance to the insecticide is unaffected (time-mortality line C). This indicates that themechanism related to that synergist is not involved in the resistance.It is also possible to determine if a target site mechanism, such as the presence of the kdr gene (sodiumchannel mutation) or insensitive acetylcholinesterase, is involved. This is done by using the synergists incombination. Their combined use will not abolish the resistance in the bioassays when a target sitemechanism is present. It is crucial in areas where pyrethroids and/or DDT are used to evaluate therelative role of detoxification and target site mechanisms involved in a particular incidence of resistance.A target site mechanism confers DDT–pyrethroid cross-resistance, while a detoxification mechanism mayor may not. Knowledge of the resistance mechanism involved is required to select a replacementinsecticide.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 16 of 24Figures 5.1.3.10a and 5.1.3.10b. Effects of synergists on resistant vector populations. Figure 5.1.3.10ashows data for a population of resistant vectors compared to a susceptible population. Figure 5.1.3.10bshows the three possible outcomes of synergist exposure (Line A — Resistance to the insecticide isabolished; Line B — Resistance to the insecticide is partially abolished; and Line C — Resistance to theinsecticide is unaffected).Fig. 5.1.3.10a.Fig. 5.1.3.10b.10 010 080SusceptibleResistant80ABMortality (%)6040Mortality (%)6040C202000 30 60 90 120Ti m e (m i n )00 30 60 90 120Ti m e (m i n )

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Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 18 of 24Use of synergistsCommonly used synergists in conjunction with the CDC bottle bioassay: Piperonyl butoxide (PBO), which inhibits oxidase activity; S.S.S-tributlyphosphorotrithioate (DEF), which inhibits esterase activity; Ethacrynic acid (EA), diethyl maleate (DM), and chlorfenethol (CF), which inhibitglutathione transferase activity.It is also possible to use a combination of the above synergists. Testing mosquitoes with synergists is atwo-step procedure. Mosquitoes are first exposed to the synergist(s) for 1 hour and then tested forinsecticide resistance using the CDC bottle bioassay. A schematic representation of performing abioassay with synergist(s) is shown in Figure 5.1.3.11.Preparation of bottles for synergist bioassaysTo use the bioassay with synergists:1. Prepare the synergist stock solution by diluting the appropriate amount of synergist in acetone ortechnical grade ethanol to be able to coat the bottles with the concentrations shown in Table 3.To make these stock solutions, use the same procedure used for making insecticide stocksolutions. In brief, dilute the appropriate amount of synergist in acetone or technical gradeethanol. To get a concentration of 400 µg/bottle of piperonyl butoxide, dissolve 400 mg inenough acetone or absolute ethanol to make 1 liter of solution. Each 1 ml of this solution willcontain 400 µg of piperonyl butoxide.2. Mark one bottle and its cap as the synergist-control bottle (without synergist);3. Mark a second bottle and its cap to be the synergist-exposure bottle;4. Add 1 ml of acetone or ethanol to the synergist-control bottle and put the cap back on tightly;5. Add 1 ml of the synergist stock solution to the synergist-exposure bottle and put the cap on backtightly;6. Coat the bottles, remove the caps, and let the bottles dry (Figures 5.1.3.3 – 5.1.3.6)7. Prepare two test sets of bottles to run the CDC bottle bioassay.Table 5.1.3.3: Synergist concentrations used in the CDC bottle bioassay.SynergistSynergist concentration(µg/bottle)Chlorfenethol 80Diethyl maleate 80Ethacrynic acid 80Piperonyl butoxide 400S.S.S-tributlyphosphorotrithioate 125

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 19 of 24CDC bottle bioassay with synergistTo run the CDC bottle bioassay with synergists:1. Introduce equal numbers of mosquitoes into the synergist-control bottle and into thesynergist-exposure bottle (about 125 mosquitoes in each bottle);2. Keep the mosquitoes in the bottles for 1 hour to allow the synergist to act;3. After the 1-hour exposure is completed, transfer the mosquitoes to two holding cartons, onefor the synergist-control mosquitoes and another for the synergist-exposed mosquitoes. Thismakes it easier to transfer mosquitoes into the insecticide-treated bottles;4. Perform the CDC bottle bioassay using one set of insecticide-coated bottles (one control andfour test bottles) for the synergist-control mosquitoes and another set (one control and fourtest bottles) for the synergist-exposed mosquitoes;5. Compare the data for the two populations of test mosquitoes.Interpretation of bioassays with synergistsPages 15-16 provide information on how to interpret the results of the CDC bottle bioassay usingsynergists. Resistance that cannot be attributed to one of the detoxification mechanisms after allsynergists have been used is likely to be due to a target site mechanism, such as kdr (sodium channelmutation) or insensitive acetylcholinesterase.ReferencesBrogdon, WG and McAllister JC, 1998. Simplification of adult mosquito bioassays through use oftime-mortality determinations in glass bottles. J. Am. Mosq. Control Assoc. 14(2):159–64.National Center for Infectious Diseases, Centers for Disease Control and Prevention (CDC).Evaluating mosquitoes for insecticide resistance: Web-based instruction. Available from:http://www.cdc.gov/malaria.Zamora Perea E, Balta Leon R, Palomino Salcedo M, Brogdon W, Devine G, 2009. Adaptation andevaluation of the bottle assay for monitoring insecticide resistance in disease vector mosquitoes inthe Peruvian Amazon. Malar. J. 8: 208.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 20 of 24Appendix 1. Frequently asked questions (FAQs)1. What happens if there are not enough mosquitoes for a complete bioassay?When the number of mosquitoes captured in the field is insufficient for a full bioassay (four coated andone control bottles), you can reduce the number of bottles to be tested, but each bioassay must ALWAYSbe run with a control until the required number is completed. If the testing takes place over a long periodof time, use recently coated bottles if necessary. See expected lifetime of coated bottles in the guideline.Except in the case of organophosphate-coated bottles, coated bottles can be use multiple times overseveral days until the bioassay is completed, as long as moisture build-up from aspiration does notbecome excessive.2. Should some bottles be designated solely as control bottles?No, some bottles should not be designated as control bottles. Bottles should randomly be assigned astest or control bottles. This will provide an additional quality control to the adequacy of the washingprocedure.3. What if there are no susceptible mosquitoes available for CDC bottle bioassay calibration?The diagnostic dose and diagnostic time for a particular species in a given area are similar. Use thediagnostic dose and the diagnostic time published in this guideline or consult the authors of this guidelineor other users with experience in the method for that particular vector. Note that the value of the CDCbottle bioassay lies in showing changes over time in the characteristics of vector populations. Therefore,a baseline is useful even if some individual mosquitoes show resistance when the initial baseline isestablished.4. Can male mosquitoes be used for the control bottle?No, males should not be used for the control bottles. Some resistance mechanisms are sex- linked, andone can be misled by using males in the control. In addition, most mosquitoes collected will be females.5. How can mosquitoes be introduced into the bottle without letting other mosquitoes escape?Some people have found it useful to employ a piece of cotton wool held against the aspirating tube at thetop of the bottle as the mosquitoes are being introduced into the bottles. As the aspirator is withdrawnafter the mosquitoes are introduced, the cotton wool can be used to close the bottle top, until the bottlecap is put in place. In our experience, a swift decisive puff of air will introduce mosquitoes without loss.Attempting to introduce mosquitoes into a bottle more than once may allow some to escape. Thissometimes happens if the user attempts to put exactly the same number of mosquitoes into each bottle,which is not necessary.6. What happens if there are fed and unfed mosquitoes among the field-collected mosquitoes tobe used in the bioassay?A collection of mosquitoes from the field may contain female mosquitoes in various physiological states,e.g., fed and unfed mosquitoes. There are two ways that this can be dealt with. First, mosquitoes can berandomly selected. Alternatively, mosquitoes can be held for one or two days for the blood meal to bedigested and then used for the bioassay.

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 21 of 24Appendix 2. Diagnostic doses and CDC bottle bioassay calibrationIt is assumed that resistance is present if a diagnostic dose, proven and validated against a susceptibleinsect population, is survived by members of a test population at a predetermined diagnostic time. Thediagnostic dose and diagnostic time are optimal parameters for detecting insecticide resistance. Adiagnostic dose that is too low will not kill susceptible mosquitoes during the bioassay, providing a falsepositiveresult for resistance. On the other hand, a diagnostic dose that is too high will kill resistantmosquitoes during the bioassay, masking resistance.For some insect vectors from some geographic regions, diagnostic doses and diagnostic times for severalinsecticides have already been determined. It is recommended that countries in these regions use thealready established parameters to allow them to compare data across countries or within regions.However, if this information is not available, the diagnostic dose and the diagnostic time will need to bedefined for each insecticide, in each region, and for each main vector species that is to be monitored. Todetermine the diagnostic dose and the diagnostic time for use in the CDC bottle bioassay, the assay willhave to be calibrated.Calibration assayThe assay is calibrated by first selecting the testing population and possible lengths of test time, and thenby determining possible diagnostic doses, given preferred diagnostic times.Population: The first step is to select a susceptible vector population to use as a baseline. If such apopulation is not available, it is possible to use the vector population from the area where the chemicalvector control measures are to be applied. This will be the reference point against which all futurepopulations can be compared.Diagnostic time: For practical reasons, the diagnostic time should be between 30 and 60 minutes.Diagnostic dose: The diagnostic dose will be a dose of insecticide that can kill 100% of mosquitoessometime between 30 and 60 minutes and that is below the saturation point. To determine possiblediagnostic doses, first prepare bottles with a range of different concentrations of insecticides per bottle, asoutlined in the guideline. Using each of these different bottles, run separate CDC bottle bioassays on 25mosquitoes of the susceptible population to determine upper limit of the diagnostic dose, which is thesaturation point. The saturation point can be defined as a concentration above which no additionaldecrease in the time required to kill 100% of the mosquitoes with an increase in insecticide concentration.See a more detailed explanation on how to determine the saturation point below.It may be necessary to run additional sets of concentrations around that range until the optimal diagnosticdose is determined. For example, starting with 10 µg/bottle, increase concentration with increments of 5µg/bottle, and continue to a final concentration of 200 µg/bottle. If no clear saturation point can bedetermined, run more assays using bottles with 200 µg/bottle, with increments of 5µg. If the saturation point still cannot be determined, more assays may be run with bottles using smallerincrements of insecticide.Interpretation of calibration dataGraphing the results of the calibration assay will show that the time-mortality line becomes straighter,steeper, and moves toward the Y-axis as the insecticide concentration increases (Figure 5.1.3.4). Thismeans that by increasing insecticide concentration the time-mortality line will reach a point whereincreasing the concentration of insecticide will not kill all mosquitoes any faster. In the example below, 15

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 22 of 24µg/bottle is the toxicological saturation point for insecticide entering the mosquito and reaching its target.Increasing the concentration to 25 µg/bottle does not cause the insecticide to penetrate the mosquito,reach the target site, and kill the mosquito any faster. Therefore, 15 µg/bottle is the saturation point andthe maximum concentration to use as the diagnostic dose. Otherwise, there is a risk that resistantmosquitoes will be killed by doses higher than the saturation point and then be recorded as susceptible,i.e., false negatives for resistance.A slightly smaller concentration compared to the toxic saturation point will kill mosquitoes in an amount oftime perhaps more convenient for the user (e.g., 30 to 60 minutes). So, it is possible to choose a lowerdiagnostic dose that kills 100% of mosquitoes within 30 to 60 min. It must be understood that severaldifferent pairs of diagnostic doses and diagnostic times will give interpretable results, but it is necessaryto consistently use the same diagnostic dose and diagnostic time for that particular insecticide on thatvector in future assays over long periods of time to allow for comparability. Otherwise, the method will notallow assessment of changes in resistance over time for that species.Figure 5.1.3.4: Determining diagnostic doses and diagnostic times.In the example shown below, 15 µg/bottle is the saturation point because higher doses did not decreasethe time for 100% of susceptible mosquitoes to be killed. A concentration of

Chapter 5 : Insecticide Resistance5.1 Insecticide Resistance Bioassays5.1.3 Guidelines for Evaluating Insecticide Resistance in Vectors using the CDC Bottle BioassayPage 23 of 24Appendix 3. CDC bottle bioassay data recording formDate: _________________ Mosquito species: _________________________________Insecticide: ________________________________Diagnostic dose: _____________________ Diagnostic time: ______________________Location of mosquito collection: ___________________________________________Time(min)Bottle 1 Bottle 2 Bottle 3 Bottle 4 All test bottles ControlAliveDeadAliveDeadAliveDeadAliveDeadTotaldeadTotal%deadTotaldeadTotal%dead01530354045607590105120TotalinbottleComments: _____________________________________________________________________________________________________________________________________

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Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity AssaysPage 1 of 45.2 Introduction to Microplate Enzyme Activity AssaysWilliam G. BrogdonIntroductionIn this section, preparation of the mosquito homogenates for analysis and data analysis are described.Specific assays are presented in the following sections.The presence of enzyme activities relevant to insecticide-resistance is often performed using microplateassays. These can be conduced with either pupae or adults. Since mosquitoes must be destroyed, itshould only be used to sample the population to determine if a specific mechanism is present and at whatfrequency it occurs. This method does, however, allow you to detect underlying resistance mechanismsthat may not be detected using bioassays. A disadvantage of these assays is that there is no way toreliably detect heterozygotes for many enzymes. These assays should always be performed with asusceptible control so baseline values can be used for comparison. Suitable samples can be obtainedfrom among the MR4-held stocks.Specimens for biochemical analysis must befreshly killed and immediately used or frozenfor later analysis. No chemicals of any sortshould be used for killing since they caninterfere with the assays. Samples may befrozen a stored a -20 o C for up to 7 days.Thereafter, -80 o C or colder storage isadvised. Samples may be stored only brieflyin the refrigerator.Alcohol-persevered specimens cannot beused for biochemical analysis. The alcoholwill reduce or eliminate the enzyme activitiesthat these procedures are measuring.Figure 5.2.1. An example of an oxidase enzymebioassay with elevated enzyme levels (possibleresistance) indicated by the darker colors.You may wish to perform total solubleprotein analysis on your samples. Thisenables size-correction when comparingdifferent species or different "broods" of thesame species which allows you to correctfor higher enzyme levels due only to theirsize. The standard curve is usuallydetermined using bovine serum albumin.Procedures for how to do this are provided by the manufacturer of the protein detection system used orare widely available and will not be discussed further.Collecting and interpreting dataA plate-reading spectrophotometer will be used to collect data at the appropriate absorbing wavelength(described in the protocol for each assay). The software for these instruments will probably include anumber of data handling features that may be especially useful.Following is a brief introduction to data analysis. A susceptible population shows an upper absorbancerange limit for susceptibility in terms of activity. Individuals with levels above that threshold are lesssusceptible (Figure 5.2.2).The susceptible population (white bars) is normally distributed with regard to enzyme activity. The upperrange limit at e.g. 570 nm (in this esterase assay) is 0.9. This becomes the resistance threshold. Thepopulation shown in black is a hypothetical field population containing individuals with elevated activity

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity AssaysPage 2 of 4(possibly corresponding to resistance) causing the distribution to be skewed to the right. Most fieldpopulations will contain both resistant and susceptible individuals. When this method is performed inconjunction with bioassays, the levels of resistance observed in the bioassay should correspond to theenzyme activity. If a higher frequency of resistance is observed in the bioassay than what is suggested bythe enzyme assay, a different resistance mechanism may be responsible.10090susceptibleresistantnumber mosqs80706050403020100susceptiblefield0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4 1.5 1.6absorbance 570 nmFigure 5.2.2. The absorbance of individuals of two different populations. Note that the referencesusceptible population contains no individuals having an absorbance > 0.9. In contrast, the fieldpopulation contains not only individuals with absorbance levels similar to those of the susceptibles butalso many with much higher absorbance levels. One may conclude that the latter individuals representinsecticide-resistant mosquitoes if similar proportions are confirmed by bioassays.It is sometimes useful to directly compare the absorbances obtained from two different enzyme assays oftwo populations when different mechanisms may be responsible for resistance observed in a bioassay.This also allows one to identify the presence of resistant individuals using a smaller sample. Such arepresentation is shown in Figure 5.2.3.Materials:• 1.5 ml tubes (e.g. Kontes)• Disposable or reusable pestles for above tubes• Pipettors and tips• Multichannel pipettors• Forceps• Freezer (used to kill or anesthetize samples)• Timer (digital counter capable of counting seconds)• Microplates• Microplate-reading spectrophotometer• Analytical balance• Dibasic potassium phosphate• Monobasic potassium phosphate

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity AssaysPage 3 of 4• Bottles of various sizes to store chemical solutions• Graduated cylinders• pH meter• Laboratory stirrerabsorbance oxidase21.81.61.41.210.80.60.40.20susceptiblepermethrin-selected0 0.5 1 1.5absorbance esteraseFigure 5.2.3. Oxidase and esterase absorbances of individual mosquitoes have beenplotted for two populations in an XY graph. In this example, permethrin exposure has beenapplied and has increased the level of resistance observed in bioassays. The upper limit ofoxidase activity in the unselected population is indicated by the horizontal line. Manyindividuals in the permethrin-selected stock now have elevated levels of oxidase thoughthe level of esterase has not been affected.Reagent Preparation:0.25 M Potassium Phosphate Buffer [KPO 4 ]1. Place 800 ml purified water in a glass beaker on a stirrer2. Add 6.6 g dibasic potassium phosphate3. Add 1.7 g monobasic potassium phosphate4. Adjust to pH 7.2 using one of the above.5. Store at room temperature6. Adjust to 1000 ml final volume

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity AssaysPage 4 of 4Sodium Acetate Buffer [NaOAc]1. Place 900 ml purified water in a glass beaker on a stirrer.2. Add 83 ml 3M sodium acetate.3. Adjust to pH 5 with glacial acetic acid.4. Adjust to 1000 ml final volume.5. Store at room temperature.Protocol for mosquito preparation:1. Kill the mosquitoes by placing them in a freezer for at least 10 minutes. Mosquitoes revive afterbrief exposures to freezing temperatures and may escape. 12. Homogenize 1 adult or pupa in 100 µl of KPO 4 buffer in a grinding tube.3. Dilute to 1000 µl final concentration with KPO 4 .4. OPTIONAL: To increase the number of assays that can be performed from each mosquito, aliquot500 µl of the homogenate into separate tubes and dilute each to 1000 ml.Loading homogenates into microplates1. At room temperature (or on ice if desired), load 100 µl aliquots of homogenate into the microplatewells in triplicate on the same plate for each enzyme assay. Use a new pipette tip for each sample.Load the first mosquito sample three wells across (A 1-3) and the next mosquito in the wells directlybelow the first (B 1-3). Continue down the plate until you reach the bottom, then shift right to the nextthree vacant set of columns and continue at the top working downward. Wells A 4-6 should containyour 9th mosquito if you have followed this pattern.2. Load the positive and negative controls into the last 6 wells on the plate.3. See specific assays for detection of various activities in the following sections. These should beperformed immediately.Notes on using multi-channel pipettors• Tips must be firmly attached by strong quick pressure of the pipettor.• Use your gloved fingers to individually check the tightness of the tips before attempting to load thepipettor with reagent.• Particular care must be taken to directly observe that each tip has loaded with the same volume asothers.• Tight-fitting tips and slow deliberate loading are essential to obtaining accurate and precise data.References:http://www.cdc.gov/ncidod/wbt/resistance/assay/microplate/index.htm1 See alternatives in chapter on Mosquito Anesthesia. Killing mosquitoes with insecticide may drasticallyaffect resistance enzymes levels measured in the assays and is not recommended.

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity Assays5.2.1 Microplate Insensitive Acetylcholinesterase AssayPage 1 of 25.2.1 Microplate Insensitive Acetylcholinesterase AssayWilliam G. BrogdonIntroductionInsensitive acetylcholinesterase (AChE) has been associated with resistance to carbamates andorganophosphates. Propoxur is used in this assay to inhibit the activity of the sensitive (i.e. susceptible)AChE, allowing the detection of the altered enzyme when it is present: The number of alleles ofinsensitive AChE is greater as the yellow color darkens. It may be possible to clearly distinguishhomozygous resistant, heterozygous and homozygous susceptible individuals by their discreteabsorbance classes.Materials• Acetone• Acetylthiocholine iodide (ATCH)• Dithiobis (2-nitrobenzoic acid), (DTNB)• 0.25 M KPO 4 buffer prepared as described in Microplate Enzyme Assays Introduction• Propoxur, technical gradeReagent preparationATCH1. Dissolve 75 mg acetylthiocholine iodide (ATCH) and 21 mg propoxur in 10 ml acetone.2. Add 90 ml 0.25 M KPO 4 buffer.3. Store at 4° C for up to 3-4 days.DTNB (Ellman’s reagent)1. Dissolve 13 mg Dithiobis (2-nitrobenzoic acid), (DTNB), in 100 ml 0.25 M KPO 4 buffer.2. Store at 4° C for up to 3-4 days.Protocol1. To the plate containing the mosquito homogenates (see Microplate Enzyme Assays Introduction),add 100 µl of KPO 4 to the negative control wells.2. Add 100 µl ATCH to each well.3. Add 100 µl DTNB to each well.4. Read plate immediately (T 0 ) with microplate reader at 414 nm.5. Read plate at 10 minutes (T 10 ).6. Subtract the T 0 reading from the T 10 reading and use this for your statistical analyses.

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Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity Assays5.2.2 Microplate Glutathione S-Transferese AssayPage 1 of 25.2.2 Microplate Glutathione S-Transferase AssayWilliam G. BrogdonIntroductionElevated glutathione s-transferase (GST) activity has been associated with resistance to DDT. Usersshould be familiar with the contents of Microplate Enzyme Assays Introduction before proceeding.Materials• Acetone• Purified water• Reduced glutathione (e.g. Sigma G4251)• 0.25 M KPO 4 buffer (prepared as described in Microplate Enzyme Assays Introduction)• Brown glass storage bottlesReagent preparationGST solution1. Dissolve 61 mg reduced glutathione in 100 ml KPO4 buffer.2. Store at 4° C for up to 3-4 dayscDNB solution1. Dissolve 20 mg 1-chloro-2,4'-dinitrobenzene (cDNB) in 10 ml acetone.2. Add 90 ml 0.25 M KPO 4 buffer.Protocol1. To the plate on which you have previously added the 100 µl of mosquito homogenates (seeMicroplate Enzyme Assays), add 100 µl of 0.25 M KPO 4 buffer in three negative control wells on thelower right corner of the plate.2. To each well, add 100 µl reduced glutathione solution.3. To each well, add 100 µl cDNB solution.4. Read plate immediately (T 0 ) with microplate reader using 340 nm filter.5. Read plate at 5 minutes (T 5 ).6. Subtract the T 0 reading from the T 5 reading and use this for your statistical analysis.

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity Assays5.2.2 Microplate Glutathione S-Transferese AssayPage 2 of 2

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity Assays5.2.3 Microplate Nonspecific Esterase AssayPage 1 of 25.2.3 Microplate Nonspecific Esterase AssayWilliam G. BrogdonIntroductionMeasures levels of non-specific - and ß-esterases present. These enzymes have been implicated inresistance organophosphates and pyrethroids. Users should be familiar with the contents of MicroplateEnzyme Assays Introduction before proceeding.Materials• Acetone• - or ß-naphthyl acetate (e.g. Sigma, N6875)• 0-dianisidine tetrazotized (e.g. Sigma, D9805)• 0.25 M KPO 4 buffer prepared as described in Microplate Enzyme Assays Introduction• Cytochrome-C from bovine heart• - or ß- napthol (e.g. Sigma, S477753)Reagent PreparationOxidase positive control1. Add 10 mg cytochrome-C to 100 ml 0.25 M Na Acetate buffer, pH 5.2. Use fresh.Esterase activity stock solution1. Dissolve 50 mg - or ß-naphthyl in 10 ml acetone depending on the assay you have chosen.2. Add 90 ml 0.25 M KPO 4 .3. Place 1 to 1.5 ml aliquots of solution in microfuge tubes and freeze. Use a lightproof storagecontainer. 1Standard1. For positive controls, dilute the esterase stock 1:35 (i.e 35 µl ß-naphthyl stock, 1.2 ml KPO 4 buffer)and 1:70 (i.e. 17.5 µl ß-naphthyl stock, 1.2 ml KPO 4 buffer).or ß-naphthyl acetate1. Dissolve 56 mg or ß-naphthyl acetate in 20 ml acetone2. Add 80 ml 0.25 M KPO 43. Store at 4° C in a light-proof bottle. Check color of dianisidine before use as it will degrade. Colorshould be pale yellow. If color is amber, discard and make fresh.Dianisidine solution1. Dissolve 100 mg 0-dianisidine tetrazotized in 100 ml purified water immediately before use.1 You can re-freeze the extra stocks if you keep them protected from light e.g. by wrapping the containerin foil or placing in a dark box.

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity Assays5.2.3 Microplate Nonspecific Esterase AssayPage 2 of 2Protocol1. If previously frozen, remove esterase activity stock solution from the freezer and thaw.2. If previously refrigerated, bring the diasnisidine solution to room temperature. Solid may comepartially out of solution when cold. If this has happened, swirl until dissolved.3. To the plate containing the mosquito homogenates (see Microplate Enzyme Assays Introduction),add 100 µl of KPO 4 to the negative and positive sample wells.4. Positive Control= or ß-naphthol. Add 100 µl of the appropriate control to each of three wells oneach plate you run as a positive control.5. Add 100 µl or ß-naphthyl acetate to each well.6. Incubate at room temperature for 10 minutes.7. Add 100 µl dianisidine to each well8. Incubate 2 minutes9. Read at 620 nm if using -naphthyl or at 540 nm if using ß-naphthyl.

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity Assays5.2.4 Microplate Oxidase AssayPage 1 of 25.2.4 Microplate Oxidase AssayWilliam G. BrogdonIntroductionElevated oxidase levels are associated with resistance to many classes of insecticide. Users should befamiliar with the contents of Microplate Enzyme Assays Introduction before proceeding.Materials• Pure methanol• Sodium acetate• Glacial acetic acid• Purified water• TMBZ or TMBZ[2HCl]• > 3% hydrogen peroxide (H 2 O 2 )• 0.25 M KPO 4 buffer prepared as described in Microplate Enzyme Assays Introduction• Cytochrome-C from bovine heart• Brown glass storage bottlesSolution Preparation0.25 M Sodium Acetate Buffer pH 5.01. To 800 ml of purified water in a beaker, add 0.25 moles of NaAc and dissolve by stirring.2. Adjust pH to 5.0 with acetic acid. 13. Store at room temperature.TMBZ solution1. Dissolve 20 mg 3,3',5,5'-Tetramethyl-Benzidine Dihydronchloride* (TMBZ [2HCL] or TMBZ) in 25 mlmethanol. 22. Add 75 ml 0.25 M Na Acetate, pH 5.0 buffer (prepared above).3. Store for up to a few days at 4°C. If this reagent turns light blue, discard and make a fresh batch.3% hydrogen peroxideHydrogen peroxide is available in many concentrations. Prepare a 3% solution in purified water usingwhat is available.Oxidase positive control stock1. Add 10 mg Cytochrome-C to 100 ml 0.25 M Na Acetate buffer, pH 5.1 It is very important the pH is exactly 5.0.2 TMBZ [2HCI] will dissolve if left to sit for a few minutes. TMBZ will dissolve if swirled under hot waterfrom the tap. Do not heat with an open flame or on a hot pad. Do not shake vigorously to aid in dissolving.

Chapter 5 : Insecticide Resistance Monitoring5.2 Microplate Enzyme Activity Assays5.2.4 Microplate Oxidase AssayPage 2 of 22. Prepare two dilutions of the above for positive controls: 1:55 (i.e. 22 µl stock, 1.2 ml KPO 4 buffer) and1:110 (i.e. 11µl cytochrome-stock, 1.2 KPO 4 buffer).3. Use fresh.Protocol1. To the plate containing the mosquito homogenates (see Microplate Enzyme Assays Introduction),add 100 µl of KPO 4 to the negative and positive sample wells.2. Add 100 µl of the cytochrome-C positive control to three wells at the lower right side of the plate –undiluted and the two dilutions prepared above. 33. Add 200 µl TMBZ solution to each well.4. Add 1 drop (or 25 µl) 3% hydrogen peroxide (H 2 O 2 ) to each well.5. Incubate for 5 minutes.6. Read with microplate reader at 620 nm.3 Cytochrome-C is very photo labile. Make sure it is the last chemical you add to your oxidase platesbefore adding the TMBZ and H 2 O 2 .

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.1 Knockdown Resistance – Anopheles gambiaePage 1 of 25.3 Insecticide Resistance Allele Assay by PCR5.3.1 Knockdown Resistance - Anopheles gambiaeIntroductionKnockdown Resistance, or KDR, is a commonly occurring permethrin resistance mutation foundthroughout Africa. Currently there are two main PCR assays to detect the West and East African forms(Martinez-Torres et al. 1998; Ranson et al. 2000). An RT-PCR based assay can be found in Chapter8.5.1.3.PCR authentication for KDR resistance in An. gambiaePrepare PCR Master Mix for 96, 48 or 1 25 μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1710 μl 855 μl 17.1 μl sterile H 2 O250 μl 125 μl 2.5 μl 10X PCR Buffer100 μl 50 μl 1.0 μl dNTP (2.5 mM mix)100 μl 50 μl 1.0 μl AgD1 (2.5 pmol/μl) [ATAGATTCCCCGACCATG]100 μl 50 μl 1.0 μl AgD2 (2.5 pmol/μl) [AGACAAGGATGATGAACC]100 μl 50 μl 1.0 μl AgD3 (2.5 pmol/μl) [AATTTGCATTACTTACGACA]100 μl 50 μl 1.0 μl AgD4 (2.5 pmol/μl) [CTGTAGTGATAGGAAATTTA]30 μl 15 μl 0.3 μl MgCl 2 (25 mM)12.5 μl 6.25 μl 0.125 μl Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 25 μl TotalTable 5.3.1.1. PCR for West African KDR resistance mechanism.Prepare PCR Master Mix for 96, 48 or 1 25 μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1710 μl 855 μl 17.1 μl sterile H 2 O250 μl 125 μl 2.5 μl 10X PCR Buffer100 μl 50 μl 1.0 μl dNTP (2 mM mix)100 μl 50 μl 1.0 μl AgD1 (2.5 pmol/μl) [ATAGATTCCCCGACCATG]100 μl 50 μl 1.0 μl AgD2 (2.5 pmol/μl) [AGACAAGGATGATGAACC]100 μl 50 μl 1.0 μl AgD4 (2.5 pmol/μl) [CTGTAGTGATAGGAAATTTA]100 μl 50 μl 1.0 μl AgD5 (2.5 pmol/μl) [TTTGCATTACTTACGACTG]30 μl 15 μl 0.3 μl MgCl 2 (25 mM)12.5 μl 6.25 μl 0.125 μl Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 25 μl TotalTable 5.3.1.2. PCR for East African KDR resistance mechanism.PCR Cycle sequence (for both East and West versions):94°C/5min x 1 cycle(94°C/1min, 48°C/2min, 72°C/2min) x 40 cycles72°C/10min x 1 cycle4°C holdRun samples on a 2% agarose EtBr gel; load 5 μl sample.Primers create fragments of 293 internal control, 195 resistant, 137 susceptible. (Figure 5.3.1.1).

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.1 Knockdown Resistance – Anopheles gambiaePage 2 of 2Figure 5.3.1.1. East AfricanPCR. Lane 1, 1 kb marker, 2,resistant, 3, susceptible, and4, heterozygous.ReferencesMartinez-Torres D et al. (1998) Molecular characterization of pyrethroid knockdown resistance (kdr) in themajor malaria vector Anopheles gambiae s.s. Insect Mol Biol 7:179-184Ranson H, Jensen B, Vulule JM, Wang X, Hemingway J, Collins FH (2000) Identification of a pointmutation in the voltage-gated sodium channel gene of Kenyan Anopheles gambiae associated withresistance to DDT and pyrethroids. Insect Mol Biol 9:491-49796 well sample preparation template

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.2 Kdr – Knockdown Resistance in Anopheles gambiae – Hyunh et al.Page 1 of 45.3.2 Kdr – Knockdown Resistance in Anopheles gambiae - Huynh et al.MR4 StaffIntroductionResistance to pyrethroids in An. gambiae is commonly associated with a single base-pair mutation in thevoltage-gated sodium channel referred to as knockdown resistance (KDR). In Africa there are two formsof this mutation, the west African form which results from a leucine to phenylalanine substitution(TTA/TTT) (Martinez-Torres et al. 1998) and the east African form which results from a leucine to serinesubstitution (TTA/TCA) (Ranson et al. 2000). The method of Huynh et al. uses the intentional mismatchprimer method described by Wilkins et al. (2006), and results using this assay were reported (Huynh et al.2007). An RT-PCR based assay can be found in Chapter 8.5.1.3.Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 rxn 48 rxn 1 rxn Reagent680 μl 340 μl 6.8 μl sterile H 2 O500 μl 250 μl 5.0 μl GoTaq 5X PCR Buffer (containing 15 mM MgCl 2 )250 μl 125 μl 2.5 μl dNTP (2-2.5 mM mix)200 μl 25 μl 0.5 μl MgCl 2 (25mM)200 μl 100 μl 2.0 μl IPCF: (F, 2.5 pmol/μl) [CTAACGCGAATTAAATGCTTTGTGACAG]200 μl 100 μl 2.0 μl IPCR: (R, 2.5 pmol/μl) [CAAAAGCAAGGCTAAGAAAAGGTTAAGC]200 μl 100 μl 2.0 μl WT: (5.0 pmol/μl) [GGTCCATGTTAATTTGCATTACTTACGAaTA]200 μl 100 μl 2.0 μl East: (2.5 pmol/μl) [CTTGGCCACTGTAGTGATAGGAAAaTC]20 μl 10 μl 0.2 μl Go-Taq DNA polymerase (5 U/μl)2.3 ml 1.15 ml 23 μl Total (To each 23 ul reaction add 2 μl template DNA)Table 5.3.2.1. East African knock-down resistance. Lower case nucleotides indicate the intentionalmismatch in the primer sequences. Nucleotides in bold are located at site of SNP (where applicable). Fand R indicate forward and reverse orientation. Add 2 μl DNA template per reaction.96 rxn 48 rxn 1 rxn Reagent880 μl 440 μl 8.8 μl sterile H 2 O500 μl 250 μl 5.0 μl GoTaq 5X PCR Buffer (containing 15 mM MgCl 2 )250 μl 125 μl 2.5 μl dNTP (2-2.5 mM mix)100 μl 25 μl 0.5 μl MgCl 2 (25mM)100 μl 50 μl 1.0 μl IPCF: (F, 2.5 pmol/μl) [CTAACGCGAATTAAATGCTTTGTGACAG]100 μl 50 μl 1.0 μl IPCR: (R, 2.5 pmol/μl) [CAAAAGCAAGGCTAAGAAAAGGTTAAGC]100 μl 50 μl 1.0 μl WT: (25.0 pmol/μl) [GGTCCATGTTAATTTGCATTACTTACGAaTA]100 μl 50 μl 1.0 μl West: (8.8 pmol/μl) [CTTGGCCACTGTAGTGATAGGAAATgTT]20 μl 10 μl 0.2 μl Go-Taq DNA polymerase (5 U/μl)2.3 ml 1.15 ml 23 μl Total (To each 23 ul reaction add 2 μl template DNA)Table 5.3.2.2. West African knock-down resistance. Add 2 μl DNA template per reaction.1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxns tocompensate for imprecise measurements.

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.2 Kdr – Knockdown Resistance in Anopheles gambiae – Hyunh et al.Page 2 of 4PCR cycle conditions (East African kdr)95°C/5min x 1 cycle(95°C/30sec , 57°C/30sec , 72°C/30sec) x 35 cycles72°C/5min x 1 cycle4°C holdPCR cycle conditions (West African kdr)95°C/5min x 1 cycle(95°C/30sec , 59°C/30sec , 72°C/30sec) x 35 cycles72°C/5min x 1 cycle4°C holdRun samples on a 2% agarose EtBr gel; load 5 μlsample.All successful reactions should contain a band of 285bp. (Figure 5.3.2.1). In addition, a band of 210 bpindicates the susceptible (wild type) allele and one of188 bp the resistant allele.Figure 5.3.2.1. Gel electrophoresis ofknockdown resistance assay, east and westsamples from separate PCRs. The first lanecontains a 1kb ladder marker, Lane 2, westAfrican homozygous resistant (VK), Lane 3,west African homozygous susceptible (MOPTI),Lane 4, west African heterozygous (VK XMOPTI), Lane 5, east African homozygousresistant (RSP-ST), Lane 6, east Africanhomozygous susceptible (KISUMU1), Lane 7,east African heterozygous (RSP-ST xKISUMU1), Lane 8, 1kb ladder marker.96 well sample preparation template

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.2 Kdr – Knockdown Resistance in Anopheles gambiae – Hyunh et al.Page 3 of 4ReferencesHuynh LY, Sandve SR, Hannan LM, Van Ert M, Gimnig JE (2007) Fitness costs of pyrethroid insecticideresistance in Anopheles gambiae. In: Annual Meeting of the Society for the Study of Evolution,Christchurch, New ZealandMartinez-Torres D, Chandre F, Williamson MS, Darriet F, Berge JB, Devonshire AL, Guillet P, Pasteur N,Pauron D (1998) Molecular characterization of pyrethroid knockdown resistance (kdr) in the major malariavector Anopheles gambiae s.s. Insect Mol Biol 7:179-184Ranson H, Jensen B, Vulule JM, Wang X, Hemingway J, Collins FH (2000) Identification of a pointmutation in the voltage-gated sodium channel gene of Kenyan Anopheles gambiae associated withresistance to DDT and pyrethroids. Insect Mol Biol 9:491-497Wilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect single nucleotide polymorphisms forAnopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin inAnopheles arabiensis. Malar J 5:125

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.2 Kdr – Knockdown Resistance in Anopheles gambiae – Hyunh et al.Page 4 of 4

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.3 Dieldrin Resistance – Anopheles gambiae and An. arabiensisPage 1 of 45.3.3 Dieldrin Resistance - An. gambiae and An. arabiensisIntroductionDue to the ban on the use of dieldrin as an insecticide, resistance to dieldrin (Rdl) is not prevalent in thewild; however, several laboratory colonies are maintained that have this mutation. The Rdl mutation hasbeen found to be associated with cross-resistance to newer insecticides that are currently beingemployed (Brooke et al. 2000; Kolaczinski and Curtis 2001). Two PCR assays have been developed todetect the Rdl mutation; one in An. gambiae (Du et al. 2005) and one in An. arabiensis (Wilkins et al.2006). An RT-PCR based assay can be found in Chapter 8.5.1.5.An. gambiae (Du et al. 2005)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1555 μl 777.5 μl 15.55 μl sterile H 2 O500 μl 250 μl 5.0 μl 5X GoTaq PCR Buffer100 μl 50 μl 1.0 μl dNTP (2.5 mM mix)100 μl 50 μl 1.0 μl RDLF (F, 25 pmol/µl) [AGTTGGTACGTTCGATGGGTTA]100 μl 50 μl 1.0 μl RDLR (R, 25 pmol/μl) [CCAGCAGACTGGCAAATACC]100 μl 50 μl 1.0 μl DF1RDL (F, 25 pmol/μl) [AATGCTACACCAGCACGTGTTGG]30 μl 15 μl 0.3 μl MgCl 2 (25 mM)15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 25 μl TotalTable 5.3.3.1. F and R indicate forward and reverse orientation. Use 1 μl DNA template.An. arabiensis (Wilkins et al. 2006)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1455 μl 727.5 μl 14.55 μl sterile H 2 O500 μl 250 μl 5.0μl 5X PCR Buffer100 μl 50 μl 1.0 μl dNTP (2 mM mix)100 μl 50 μl 1.0 μl RDLF (F, 25 pmol/µl) [AGTTGGTACGTTCGATGGGTTA]100 μl 50 μl 1.0 μl RDLR (R, 25 pmol/μl) [CCAGCAGACTGGCAAATACC]100 μl 50 μl 1.0 μl AARDL (F, 25 pmol/μl) [GCTACACCAGCACGTGaTT]100 μl 50 μl 1.0 μl RDLSS (R, 25 pmol/µl) [CAAGACAGTAGTTACACCTAAaGC]30 μl 15 μl 0.3 μl MgCl 2 (25 mM)15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 25 μl TotalTable 5.3.3.2. In primer sequence, lower case nucleotides indicates the intentional mismatch, nucleotidesin bold are located at site of SNP (where applicable); F and R indicate forward and reverse orientation.Use 1 μl DNA template.

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.3 Dieldrin Resistance – Anopheles gambiae and An. arabiensisPage 2 of 4PCR Cycle conditions for both assays94 o C/3min x 1 cycle(94 o C/1min -0- 53 o C/2min -0- 72 o C/2min) x 30 cycles72 o C/10min x 1 cycle4 o C holdRun samples on a 2% agarose EtBr gel; load 5 μl sample.An. gambiaePrimers create fragments of: control band 390 bp, An. gambiae resistant 160 bpAn. arabiensisPrimers create fragments of: control band 255 bp, An. arabiensis resistant 157 bp, and An. arabiensissusceptible 120 bp (Figure 5.3.3.1)Figure 5.3.3.1 An. arabiensis RDl assay. Lane 1 1kb ladder, lanes 2-9 resistant An. arabiensis, lanes 10-24 susceptible An. arabiensis.ReferencesBrooke BD, Hunt RH, Coetzee M (2000) Resistance to dieldrin + fipronil assorts with chromosomeinversion 2La in the malaria vector Anopheles gambiae. Medical and Veterinary Entomology 14:190-194Du W et al. (2005) Independent mutations in the Rdl locus confer dieldrin resistance to Anophelesgambiae and An. arabiensis. Insect Molecular Biology 14:179-183Kolaczinski J, Curtis C (2001) Laboratory evaluation of fipronil, a phenylpyrazole insecticide, against adultAnopheles (Diptera: Culicidae) and investigation of its possible cross-resistance with dieldrin inAnopheles stephensi. Pest Management Science 57:41-45Wilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect single nucleotide polymorphisms forAnopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin inAnopheles arabiensis. Malar J 5:125

96 well sample preparation templateChapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.3 Dieldrin Resistance – Anopheles gambiae and An. arabiensisPage 3 of 4

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.3 Dieldrin Resistance – Anopheles gambiae and An. arabiensisPage 4 of 4

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.4 ACE-1 Resistance in An. gambiaePage 1 of 25.3.4 ACE-1 Resistance in An. gambiaeMR4 StaffIntroductionInsensitive acetylcholinesterase (AChE) is a resistance mechanism associated with tolerance tocarbamate and organophosphate insecticides. Mutations within AChE genes in Dipterans are widespreadand have varying effects on the tolerance levels to insecticides. This is of great importance due to theincreased interest in utilizing bendiocarb as a potential treatment for bed-nets and increased resistancelevels to common insecticides across Africa. (Weill et al. 2004) isolated a unique mutation found in bothold and new world vectors in the ace-1 allele. From this a PCR-RFLP was designed based on the G119Smutation isolated from An. gambiae. An RT-PCR based assay can be found in Chapter 8.5.1.4.Anopheles spp. (Weill et al. 2004)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1635 μl 817.5 μl 16.35 μl sterile H 2 O500 μl 250 μl 5.0 μl 5X GoTaq PCR Buffer100 μl 50 μl 1.0 μl dNTP (2.5 mM mix)125 μl 62.5 μl 1.25 μl MOUSTDIR1 (25 pmol/µl) [CCGGGNGCSACYATGTGGAA]125 μl 62.5 μl 1.25 μl MOUSTREV1 (25 pmol/µl) [ACGATMACGTTCTCYTCCGA]15 μl 7.5 μl 0.15 μl Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 25 μl TotalTable 5.3.4.1. Use 1 μl DNA template.PCR Cycle conditions93 o C/5min x 1 cycle(93 o C/1min -0- 53 o C/1min -0- 72 o C/1.5min) x 35 cycles72 o C/10min x 1 cycle4 o C holdRestriction enzyme digestAdd 1μl AluI restriction enzyme, 2 µl of H20, and 2µl of buffer to 15 μl PCR product from above reaction.Allow to incubate at 37°C for 8-24 hr. For shorter times, incomplete digests could be a problem.Visualize on a 2% agarose ethidium bromide gel.Anopheles spp.Primers create a 194 bp amplicon, after restriction enzyme digest homozygous resistant individuals willhave 120bp and 74bp fragments (Figure 5.3.4.1).

Chapter 5 : Insecticide Resistance Monitoring5.3 Insecticide Resistance Allele Assay by PCR5.3.4 ACE-1 Resistance in An. gambiaePage 2 of 2Figure 5.3.4.1 Ace-1 resistance PCR-RFLP. Lane 1,1kb ladder, lanes 2-4 An. gambiae homozyogous forace-1 resistance mutation, lane 5 homozygous An.arabiensis negative for the ace-1 mutation.ReferencesWeill M et al. (2004) The unique mutation in ace-1 giving high insecticide resistance is easily detectable inmosquito vectors. Insect Mol Biol 13:1-796 well sample preparation template

Chapter 6 : Dissection Techniques6.1 General Dissection BuffersPage 1 of 2Chapter 6 : Dissection Techniques6.1 General Dissection BuffersMarc KlowdenBackgroundWhen dissecting mosquitoes for examination or for studying the functions of their isolated tissues, aphysiological saline should be used to prevent them from drying out and to maintain these tissues in areasonably normal state. These solutions will ideally mimic the composition of hemolymph. However,because insect hemolymph can often vary considerably during the life cycle in response to feeding andreproduction, insect tissues generally show a wide tolerance to variations in composition, and the salinesolutions that have been formulated are sufficiently generic to enable them to be used for most species.Saline solutions suitable for dissection(Hayes 1953)Component g/lNaCl 9.0CaCl 2 0.2KCl 0.2NaHCO 3 0.1(Ephrussi and Beadle 1936)Component g/lNaCl 7.5KCl 0.35CaCl 2 0.21Saline solutions suitable for long-term tissue survival:(Beyenbach and Masia 2002). Used for adult Malpighian tubules.Component g/lNaCl 8.76KCl 0.25CaCl 2 0.19MgCl 2 0.09Hepes 5.96Glucose 0.90NaHCO 3 0.15Adjust to pH 7 with 1M NaOH

Chapter 6 : Dissection Techniques6.1 General Dissection BuffersPage 2 of 2(Onken et al. 2006). Used for larval midgut tissue.Component g/lNaCl 2.48KCl 0.22MgCl 2 0.06CaCl 2 0.55NaHCO 3 0.42Succinic acid 0.59Malic acid 0.67L-proline 0.58L-glutamine 1.33L-histidine 1.35L-arginine 0.57Glucose 1.80Hepes 5.96Adjust to pH 7 with 1M NaOHReferencesBeyenbach KW, Masia R (2002) Membrane conductances of principal cells in Malpighian tubules ofAedes aegypti. J Insect Physiol 48:375-386Ephrussi B, Beadle GW (1936) A technique of transplantation for Drosophila. Am. Nat. 70:218-225Hayes RO (1953) Determination of a physiological saline solution for Aedes aegypti (L.). J. Econ.Entomol. 46:624-627Onken H, Moffett SB, Moffett DF (2006) The isolated anterior stomach of larval mosquitoes (Aedesaegypti): voltage-clamp measurements with a tubular epithelium. Comp Biochem Physiol A Mol IntegrPhysiol 143:24-34

Chapter 6 : Dissection Techniques6.2 Rapid Larval Midgut ExtractionPage 1 of 26.2 Rapid Larval Midgut ExtractionMarco Neira, Dmitri Boudko, Leslie VanEkeris, Paul LinserIntroductionOnce mastered, this technique allows the researcher to quickly separate the midgut from the rest of thebody, and is therefore suitable for procedures which require the pooling of large amounts of tissue, suchas protein and/or RNA extraction.Materials• Dissection dish (a Petri dish that has its bottom coated with a fine layer of Sylgard ® (silicone,Dow-Corning Corporation, Midland, Michigan)• Stereoscope• Fine forceps• Microdissection scissors• Dissecting needles or minutien pin mounted on a long wooden stick• Clean microscope slides and cover slipsProtocol1. Place larvae on ice for 10-15 minutes in order to immobilize them.2. Transfer each larva to a dissection dish containing 70% ethanol.3. Pin the head capsule to the bottom of the dish using a minutien pin (sometimes placing the larvaventral-side up can facilitate this task).4. Using sharp microdissection scissors, clip-off abdominal segments 8-10.5. Use the sides of two sets of forceps to carefully ‘squeeze’ the gut out of the larval body. Start byapplying pressure at the head/thorax junction (Figure 6.2.1 A), and work your way across the rest ofthe thorax and the abdomen using a stepwise motion. The gut should progressively protrude out ofthe distal end of the abdomen (Figure 6.2.1 B).6. After the gut has been extracted, it might need to be carefully cleaned of attached fat-body and/orlarge tracheal trunks. Clean guts should immediately be transferred to the appropriate reagents forprotein or RNA isolation.

Figure 6.2.1. Larval mosquito midgut extraction.Chapter 6 : Dissection Techniques6.2 Rapid Larval Midgut ExtractionPage 2 of 2

Chapter 6 : Dissection Techniques6.3 Larval Midgut VivisectionPage 1 of 26.3 Larval Midgut VivisectionMarco Neira, Dmitri Boudko, Leslie VanEkeris, Paul LinserIntroductionThis technique is used for studies that require direct access to the internal organs of a live larva, such aselectrophysiological measurements. It can also be used to perform whole-mount immunostaining ofinternal organs, in which case the specimen should be dissected in fixative solution instead of hemolymphsubstitute.Materials• Dissection dish (a Petri dish that has its bottom coated with a fine layer of Sylgard ® (silicone,Dow-Corning Corporation, Midland, Michigan)• Stereoscope• Fine forceps• Dissecting needles or minutien pin mounted on a wooden applicator stick• Clean microscope slides and cover slipsHemolymph substitute42.5mM NaCl, 3mM KCl, 0.6mM MgSO 4 , 5mM CaCl 2 , 5mM NaHCO 3 , 5mM succinic acid, 5mM malicacid, 5mM L-proline, 9.1mM L-glutamine, 8.7mM L-histidine, 3.3mM L-arginine, 10mM dextrose, 25mMHepes, pH 7.0Fixative SolutionTo make fixative solution for whole-mounts, mix equal volumes of 4% paraformaldehyde in 0.1M sodiumcacodylate bufferVariant 1:1. Place larvae on ice for 10-15 minutes to immobilize them.2. Transfer each larva to a dissection dish containing Ringer’s solution, hemolymph substitute, oranother isotonic medium.3. Place the larva dorsal-side up, and pin the head capsule to the bottom of the dish using a minutienpin.4. Using sharp microdissection scissors, clip-off abdominal segments 8-10.5. Using fine forceps, carefully grab the integument of seventh abdominal segment and pull it justenough to straighten the larva’s body (Figure 6.3.1 A).6. While keeping the body straight, use microdissection scissors to cut across the lateral side of theabdomen and thorax, and then across the dorsal side of the thorax, as shown by the dashed arrowsin Figure 6.3.1 A.7. Move the dorsal integument aside to expose the internal organs. Use minutien pins to secure theexoskeleton to the bottom of the dish.Variant 2:1. Place larvae on ice for 10-15 minutes in order to immobilize them.2. Transfer each larva to a dissection dish containing Ringer’s solution, hemolymph substitute, oranother isotonic medium.

Chapter 6 : Dissection Techniques6.3 Larval Midgut VivisectionPage 2 of 23. Place the larva dorsal-side up, and pin the head capsule and the last abdominal segment to thebottom of the dish using minutien pins.4. Insert microdissection scissors at the head/thorax junction and cut the integument following the middorsalline of the body (between the two main dorsal tracheal trunks). Then, cut the integumentacross the dorsal side of the thorax as shown by the dashed arrows in Figure 6.3.1 B.5. Using fine forceps, grab the dorsal integument and pull it gently towards the sides (Figure 6.3.1 B),exposing the internal organs. Use minutien pins to secure the exoskeleton to the bottom of the dish.Figure 6.3.2 shows the final result of this kind of dissection.Figure 6.3.1. Larval mosquito vivisection. A) variant 1; B) variant 2Figure 6.3.2. Final result of larval vivisection. AMG: anterior midgut; CMG: central midgut; GC:gastric caeca; MT: Malpighian tubes; PMG: posterior midgut; Py: Pylorus. Numbers indicateabdominal segments. Figure modified from (Boudko et al. 2001). Used with permission.ReferencesBoudko DY, Moroz LL, Linser PJ, Trimarchi JR, Smith PJ, Harvey WR (2001) In situ analysis of pHgradients in mosquito larvae using non-invasive, self-referencing, pH-sensitive microelectrodes. J ExpBiol 204:691-699

Chapter 6 : Dissection Techniques6.4 Adult Male Testes DissectionPage 1 of 26.4 Adult Male Testes DissectionMR4 StaffIntroductionMale reproductive organs can provide valuable data about population age and mating history (Mahmoodand Reisen 1982; Huho et al. 2006). Testes size has been associated with male age in An. stephensi inwhich it was found that longer testes were seen in sexually mature individuals while shorter ones werefound in sexually immature or older mosquitoes (Mahmood and Reisen 1982). Additional studies in An.gambiae have found that differences in the appearance of the male accessory glands (MAG) were relatedto the age of the mosquito (Huho et al. 2006). Mosquitoes that were greater than 4 days old typically hadno visible clear area in their MAG and large sperm reservoirs while mosquitoes that were younger than 4had a small, transparent area on the edge of their MAG and small sperm reservoirs. Adult male testes arealso a good source of metaphase chromosomes (French et al. 1962). The dissection technique describedbelow should successfully remove all the reproductive organs.Materials• Stereoscope• Compound microscope• Fine forceps• Dissecting needles or minutien pin mounted on a long wooden stick• Clean microscope slides and cover slips• PBS solution (0.01M, pH 7.2)Protocol1. Aspirate males into a container and anesthetize by gently chilling them at -20ºC for 5-7 minutes.2. Place a drop of PBS on a clean slide3. Under the stereoscope, gently grasp the male by the thorax with a pair of forceps and place ventralside up with the abdomen resting in the PBS.4. Take a fine tip needle or forceps and gently remove the claspers of the male by piercing them andgently pulling them away while holding the thorax with forceps (Figure 6.4.1).5. Using the dissecting needles, gently remove the extraneous tissues isolating the male reproductiveorgans then cover with a clean cover slip.6. Examine both the testes and MAG to determine their dimensions and presence or absence of a clearzone (Figures 6.4.2-9).ReferencesFrench WL, Baker RH, Kitzmiller JB (1962) Preparation of mosquito chromosomes. Mosq News L1 -2927pdf 22:337-383Huho BJ, Ng'habi KR, Killeen GF, Nkwengulila G, Knols BG, Ferguson HM (2006) A reliablemorphological method to assess the age of male Anopheles gambiae. Malar J 5:62

Chapter 6 : Dissection Techniques6.4 Adult Male Testes DissectionPage 2 of 2Mahmood F, Reisen WK (1982) Anopheles stephensi (Diptera: Culicidae): changes in male matingcompetence and reproductive system morphology associated with aging and mating. J Med Entomol19:573-588Figure 6.4.1. Dissected maleAnopheles gambiae mosquito.The MAGs are visible as theyellow sacs near the terminalia.Figure 6.4.2. MAG of a mosquitothat is 4 days old.Note the absence of a clearzone.Figure 6.4.4. MAG and testes ofa 1 day old An. gambiaemosquito.Figure 6.4.5. MAG and testes of a 5day old An. gambiae mosquito.Figure 6.4.6. MAG and testesof a 7 day old An. gambiaemosquito.Figure 6.4.7. Testis of a 1 dayold An. gambiae mosquito.Figure 6.4.8. Testis of a 5 dayold An. gambiae mosquito.Figure 6.4.9. Testis of a 7 dayold An. gambiae mosquito.

Chapter 6 : Dissection Techniques6.5 Dissecting Plasmodium-Infected Mosquitoes6.5.1 MidgutPage 1 of 26.5 Dissecting Plasmodium-Infected Mosquitoes6.5.1 MidgutMR4 StaffIntroductionExperimental Anopheles infections may be performed todetermine the length of sporogony for various Plasmodiumspecies or to establish susceptibility or refractoriness toPlasmodium infections. Dissection of the midgut to observeoocysts should be performed 5-7 days after infection occurs.The precise number of days that is best will vary with parasitespecies. Some examples of variability are shown in Table6.5.1.1 (WHO 1975). An alternate method by Looker andTaylor-Robinson can be found in Methods in Malaria Research(Looker and Taylor-Robinson 2004).SpeciesP. vivax 4-7P. ovale 3-4P. falciparum 3-7P. malariae 3-11Days to oocystsTable 6.5.1.1. Adapted from WHOManual on Practical Entomology, 1975.MaterialsStereoscopeDissecting needlesMicroscopeForcepsPBS or saline solutionMicroscope slidesMercurochromeCover slipsChloroformDetecting the presence of Plasmodium oocysts in the midgut:1. Anesthetize the females (see section on Mosquito Anesthesia).2. Place a drop of PBS on a clean microscope slide.3. Place a female in the drop of PBS ventral side up with the terminalia in the center of the drop.4. Grasp the female by the thorax using forceps.5. Lay a dissecting needle across the 7 th abdominal segment (without slicing the abdomen) and gentlydetach the terminalia.6. While still grasping the thorax with forceps, gently pull the terminalia away from the abdomen using thedissecting needle. Pull very slowly to ensure that the midgut does not detach from the terminal end(Figure 6.5.1.1).7. If the midgut detaches, gently cut the edges of the 1st abdominal segment and gently pull the cuticleover the contents.8. Remove the Malpighian tubes as well as any other accessory tissue and debris leaving only themidgut.

Chapter 6 : Dissection Techniques6.5 Dissecting Plasmodium-Infected Mosquitoes6.5.1 MidgutPage 2 of 29. On a new microscope slide, place a small drop of mercurochrome.10. Place the dissected midgut in the stain and cover with a clean cover slip.11. Visualize under a compound microscope. Depending on the model, you may have to use phasecontrast in order to identify the oocysts.What you will see:In most mosquitoes you will find varying numbers of encapsulated and unencapsulated (normal) oocysts.Encapsulated oocysts appear as disk-shaped structures in the midgut (small melanized particles) (Figure6.5.1.2, right panel). Normal oocysts appear as large spherical objects, usually slightly lighter incoloration and attached to the apical surface (Figure 6.5.1.2, left panel). There are certain strains ofmosquitoes that have been selected to highly express an immune response to Plasmodium infection (L3-5, An. gambiae) or to be completely susceptible to infection (4ARR, An. gambiae).Figure 6.5.1.1. Dissection of midgut (showninside circle). Cut away other portions to avoidconfusion under the microscope.Figure 6.5.1.2. Left panel shows wild-type or normalunencapsulated oocysts. Right panel shows allencapsulated oocycsts with the exception of onenormal oocyst toward the center (Collins et al. 1986).ReferencesCollins FH et al. (1986) Genetic selection of a Plasmodium-refractory strain of the malaria vectorAnopheles gambiae. Science 234:607-610Looker M, Taylor-Robinson AW (2004) Methods in Malaria Research, 4th edn. MR4/ATCC, Manassas,VAWHO (1975) Manual on practical entomology in malaria. Part II. In. World Health Organization, Geneva, p191pp.

Chapter 6 : Dissection Techniques6.5 Dissecting Plasmodium-Infected Mosquitoes6.5.2 Salivary GlandsPage 1 of 26.5.2 Salivary GlandsMR4 StaffIntroductionDissection of the salivary glands should be done 10-18 dayspost-infection. The precise number of days that is best willvary with species. Some examples of variability are shown inTable 6.5.2.1 (WHO 1975). An alternate method by Lookerand Taylor-Robinson can be found in Methods in MalariaResearch (Looker and Taylor-Robinson 2004).Materials• Stereoscope• Dissecting needles• Microscope• Forceps• PBS or saline solution• Microscope slides• Giemsa stain• Cover slips• Chloroform, ethyl ether or ethylacetateSpeciesDays to sporozoitesP. vivax 8-10P. ovale 10-15P. falciparum 10-18P. malariae 15-21Table 6.5.2.1. Adapted from WHOManual on Practical Entomology, 1975.ProtocolDetecting the presence of sporozoites in the salivary glands.1. Anesthetize adult females (see 3.8 Mosquito Anesthesia).2. Place a drop of PBS on a clean microscope slide.3. Place a female on her side with her thorax in the PBS solution.4. While firmly holding the female with forceps or a needle, gently lay another needle across the necknear the head and slowly pull the head away (Figure 6.5.2.1).5. Gently detach the salivary glands from the head.6. If the salivary glands remain within the thorax, using the needle, apply gentle pressure to the thoraxtowards the mesonatum end and squeeze the glands out.7. At this stage you can either process the slides for staining or perform a brief inspection to determine ifsporozoites are present.Brief inspectionCover the dissected salivary glands with a cover slip and inspect the glands under a microscope for thepresence of the threadlike sporozoites. Gentle pressure on the glands will help rupture the tissues freeingthe sporozoites into solution.

Chapter 6 : Dissection Techniques6.5 Dissecting Plasmodium-Infected Mosquitoes6.5.2 Salivary GlandsPage 2 of 2StainingLet glands fully dry on the slide. Once dried, fix with methanol for 1 minute, rinse with water, then stainwith Giemsa stain for 40 minutes. Rinse again and air dry before inspecting under a microscope.What you will seeIn uninfected salivary glands, you will see a tissue layer with no visible threadlike sporozoites (Figure6.5.2.2). In infected mosquitoes, you will see the threadlike parasites both in the tissues or, if the gland isruptured, in the field surrounding the tissues (Figure 6.5.2.3). Giemsa staining helps estimate lowparasite loads by increasing resolution of the sporozoites within the glands.Figure 6.5.2.1. Dissection of salivary glands.Figure 6.5.2.2. Stained, uninfected salivaryglands.Figure 6.5.2.3. Stained, infected salivaryglands. Note the thin, threadlike sporozoites(circled) of P. vivax spilling from the salivarygland.ReferencesLooker M, Taylor-Robinson AW (2004) Methods in Malaria Research, 4th edn. MR4/ATCC, Manassas,VAWHO (1975) Manual on practical entomology in malaria. Part II. In. World Health Organization, Geneva, p191pp.

Chapter 6 : Dissection Techniques6.6 Examination of Ovaries by Tracheal Distension to Determine ParityPage 1 of 46.6 Examination of Ovaries by Tracheal Distension to DetermineParityFrancis AtieliIntroductionParity is used to determine the age structure of a feral population, and it can also be used to ascertain thenet reproductivity of a colony (Githeko et al. 1993). Parous mosquitoes are those that have taken a bloodmeal and oviposited at least once. Nulliparous mosquitoes have never oviposited. Net reproduction rateof a colony can be determined by dissecting several females to determine the number of parousindividuals within the population.A more advanced technique was developed by Detinova (1962) that assesses how many egg batcheshave been developed by an individual female (reviewed by Hoc and Wilkes 1995). In this method, asembryos develop within the ovaries they stretch the ovariole sheath. After oviposition, these sacs whichcontain remnants from oogenesis shrink and develop into permanent dilatations. After each subsequentfeeding, a new embryo will form anterior to the previous dilatation. A mosquito with three dilatations wouldbe labeled “3 parous”. Caution should be taken when examining for dilatations as resorbed embryos willalso form dilatations. Likewise in some species such as An. atroparvus, dilatations may not be formed atall (Service 1993).The simplest technique for determining parity is to examine the tracheoles within the ovaries (Kardos andBellamy 1961). As the ovaries expand after the primary blood meal, the tracheoles that are associatedwith them are permanently distended (Hoc and Charlwood 1990). Therefore, in nulliparous females thetracheoles are tightly wound coils called ‘skeins.’ Parous females will have tracheoles that havedistended. Note that in younger females there may be a mixture of both skeins and ‘extended’ tracheoles;if any distended tracheoles are present, you can assume that the female is parous and has fed and laideggs at least one time.Materials• Microscope slides• PBS or another physiological dissecting solution• Forceps• Dissecting needles• Stereoscope• Compound Microscope (200 X magnification)Technique1. Gently anesthetize adult females.2. Place a drop of PBS on a clean microscope slide.3. Under the stereoscope, gently grasp female by the thorax with forceps, and place ventral side up withher abdomen in the PBS.4. While viewing the specimen under the stereoscope, take a fine tip needle or forceps and gentlyremove the 7th and 8th abdominal segments of the female by grasping them and pulling away slowly.5. Locate the ovaries; they will appear as a pair of white oval objects attached to the removed segments(Figure 6.6.1). Dissect away the accessory tissues and isolate the ovaries.6. Transfer ovaries to a new slide and allow to air dry.

Chapter 6 : Dissection Techniques6.6 Examination of Ovaries by Tracheal Distension to Determine ParityPage 2 of 47. Multiple pairs of ovaries can be mounted on a single microscope slide, but ensure that all the sampleshave dried before viewing them under a microscope.8. View under a compound scope at 200 X magnification. Locate the tracheoles and determine if thespecimen is nulliparous (Figure 6.6.2) or parous (Figures 6.6.3 and 6.6.4).Figure 6.6.1. Dissection of female terminaliaunder stereoscope. The ovaries are circled.Terminalia and segments VII and VIII are visible inthe upper right hand corner.Figure 6.6.2. An example of a tightly coiledtracheole called a “skein” is shown inside the circle.This is a nulliparous female.Figure 6.6.3. Inside the circle is the loosestructure of the tracheoles after oogenesis. This isreferred to as the extended state. This female isparous.Figure 6.6.4. Both types of tracheoles are seen inthis photograph. The circle on the left side shows atightly coiled skein while the circle on the rightshows an extended tracheole. Even though both arepresent in this example, this female would beconsidered parous.ReferencesDetinova TS (1962) Age grading methods in Diptera of medical importance with special reference tosome vectors of malaria. Monograph Series, World Health Organization 47:1-216

Chapter 6 : Dissection Techniques6.6 Examination of Ovaries by Tracheal Distension to Determine ParityPage 3 of 4Githeko AK, Service MW, Atieli FK, Hill SM, Crampton JM (1993) Field Testing an Enzyme-LinkedSynthetic Oligonucleotide Probe for Identification of Anopheles gambiae ss and An. arabiensis. Annals ofTropical Medicine and Parasitology 87:595-601Hoc TQ, Charlwood JD (1990) Age determination of Aedes cantans using the ovarian oil injectiontechnique. Med Vet Entomol 4:227-233Hoc TQ, Wilkes TJ (1995) The ovarioles structure of Anopheles gambiae (Diptera: Culicidae) and its usein determining physiological age. Bull Entomol Res. 85:56-69Kardos EH, Bellamy RE (1961) Distinguishing nulliparous from parous female Culex tarsalis byexamination of the ovarian tracheoles. Ann Entomol Soc Am 54:448-451Service MM (1993) Mosquito Ecology: Field Sampling Methods, 1st edn. Chapman and Hall, New York,New York

Chapter 6 : Dissection Techniques6.6 Examination of Ovaries by Tracheal Distension to Determine ParityPage 4 of 4

Chapter 6 : Dissection Techniques6.7 Dissecting Spermathecae to Determine Insemination StatusPage 1 of 26.7 Dissecting Spermathecae to Determine Insemination StatusMR4 StaffIntroductionIn order to monitor colony mating activity and to assess wild-mosquito mating status, it is often necessaryto determine whether females are inseminated. The least complicated method to determine this is todissect spermathecae and determine whether spermatozoa are present. The spermatheca is located nearthe terminalia inside segment VIII and can often be seen through the cuticle while viewing the ventralside. It is a spherical, fenestrated dark-brown organ which has a golf-ball-like appearance. After gentlyremoving the terminalia and segment IX, the spermathecae will be exposed along with the malpighiantubules, ovaries, and digestive tract. Then the spermathecae can be easily removed and viewed undermagnification, and insemination status is usually readily apparent.Materials:• Fine forceps• Dissecting needles or minutien pin mounted on a long wooden stick• Clean microscope slides and cover slips• PBS solution• DropperEquipment:• Stereoscope• Compound microscope with dark field capacityProcedure:1. Anesthetize or kill the adult females.2. Place a drop of PBS on a clean slide.3. Gently grasp the female by the thorax with a pair of forceps and place ventral side up with theabdomen resting in the PBS under the stereoscope.4. Gently remove the terminalia of the female by grasping them and pulling away slowly using a fine tipneedle or forceps.5. Locate the spermatheca within the 8th segment and terminalia section removed previously. It shouldappear as a dark sphere that may or may not be surrounded by accessory tissues (Figure 6.7.1).Dispose of the mosquito carcass and remaining tissues.6. Gently lower a cover slip onto the spermatheca using a needle (to avoid rupturing the spermathecae).On the underside of the slide, circle the area surrounding the spermathecae using a permanentmarker. The marking will make it much simpler to locate the tiny organ. With experience, you willdetermine how much PBS to use to prevent rupturing the spermatheca.7. Under 100X magnification on a compound microscope, look for movement of the long thread-likespermatozoa within the spermathecae (Figure 6.7.2). They will appear as fine concentric threadswithin the spermathecae and are often seen rotating as a cluster. If the spermatheca ruptured afterplacing the cover slip, scan the surrounding field for the spermatozoa. Uninseminated females willhave a fairly transparent spermatheca (Figure 6.7.3).

Chapter 6 : Dissection Techniques6.7 Dissecting Spermathecae to Determine Insemination StatusPage 2 of 2Figure 6.7.1. Golf ball shapedspermatheca.Figure 6.7.2. 100X magnificationof a full spermatheca. The brightring is the bundle of sperm.Figure 6.7.3. Uninseminatedfemale spermatheca.

Chapter 6 : Dissection Techniques6.8 A. gambiae s.l. Ovarian Polytene Chromosome PreparationPage 1 of 66.8 A. gambiae s.l. Ovarian Polytene Chromosome PreparationAnthony Cornel 1IntroductionThe highest quality Anopheles gambiae s.l. polytene chromosomes are prepared from nurse cells in eggsat Christophers’ III stage (Clements 1992) of ovarian development. Chromosomes of adequatepolytenization also occur in salivary gland cells of late fourth stage larvae which are useful in larvalecological studies. Anopheles gambiae s.l. has a diploid chromosome complement of 2n = 6 with one pairof heteromorphic X and Y sex chromosomes and two pairs of autosomes numbered as 2 and 3 (seesection on Anopheles Mendelian Genetics).A typical polytene chromosome squash will produce a spread of arm configurations known as X, 2R, 2L,3R and 3L (Figure 6.8.1). The entire Y and portions of the X cannot be visualized because they areheterochromatic and under-replicated. Drawings depicting the banding patterns of An. gambiae s.l.polytene chromosomes are available to identify the divisions and subdivisions and locations ofparacentric inversions (Coluzzi et al. 2002; Holt et al. 2002).Wherever possible, multiple methods have been described from which you can choose depending onlaboratory equipment and the needs and quality of chromosome preparation required. The most crucialstep to obtain excellent chromosomes is to recognize the appropriate half gravid state of the female andlate 4 th stage larvae from which to dissect. Thereafter, the most crucial steps are the tapping proceduresto spread the chromosomes into recognizable arms. It will take a little while and practice to achieve theappropriate skills, and I suggest you spend a few days learning these skills with someone who is bothfamiliar with chromosome preparation techniques and An. gambiae s.l. chromosome banding patterns.Solutions• Anesthetic (ethyl ether, triethylamine, CO 2 , -20C freezer space)• Modified Carnoy’s fixative (three parts pure (100%) ethanol and one part glacial acetic acid).Ethanol absorbs water from the atmosphere so ensure that the ethanol is pure, as water infixative compromises quality of spreads.• Sigmacote® - (Sigma-Aldrich – St. Louis MO, USA)• Propionic acid - prepare 5% and 50% in water.• 2% lacto-aceto orcein - prepare this solution by adding slowly 2% by weight of synthetic orceinpowder to a solution of 1 part glacial acetic acid and 1 part pure lactic acid under constant stirring(use a magnetic stirrer). Remove un-dissolved orcein particulates by filtering the solution throughWhatman 3MM paper. The solution can be stored indefinitely at room temperature.• Ethanol – 70%, 90% and 100% in water.Materials• Anesthetizing mosquito chamber: 45 ml Falcon® tubes, razor blade, 925 mµ mesh or smallerscreening, glue, rubber bands, dental rubber latex sheeting (Super Dam- Patterson Brand),regular household sponge, desiccator.• 4C refrigerator• -20C freezer1 Mosquito Control Research Laboratory, Department of Entomology and Center for VectorborneDiseases, University of California at Davis, Parlier, CA 93648, E-mail: cornel@uckac.edu

Chapter 6 : Dissection Techniques6.8 A. gambiae s.l. Ovarian Polytene Chromosome PreparationPage 2 of 6• 0.13- 0.17 mm thick cover glasses of dimensions 22L X 22W mm• Frosted 1mm thick microscope slides 75 x 25 mm in size.• Beaker• Aluminum foil• Forceps• Pasteur pipettes• Absorbent paper (Whatman 3MM paper)• Dissecting needles and minutien pins• Dissecting Microscope• 1.5 ml screw cap polypropylene vials• Phase contrast compound microscope with 10X, 40 or 60 X and 100X objective lenses.• Digital imaging systemFigure 6.8.1. Chromosome squash showing the five reasonably well spread chromosome arms X, R,L, 3R and 3L of An. gambiae sensu stricto. Magnification X 600 under phase contrast. T = Telomere;C = Centromere. Specimen from Eau et Foret, Cameroon 2006.

Chapter 6 : Dissection Techniques6.8 A. gambiae s.l. Ovarian Polytene Chromosome PreparationPage 3 of 6CAFigure 6.8.2. Selection of items useful for preparingAn.gambaie s.l. polytene chromosomes. A -Anesthetizing chamber; B - mattress needle with endcurved downwards for tapping; C – rolled up absorbentpaper for drawing up small volumes of fluids onmicroscope slides.BProcedure:1. Siliconized cover-glasses are not crucialfor temporary squashes but they are ifcover-glasses need to be removed forpreparation of permanent mounts and insitu hybridization. Prepare siliconizedcover-glasses by fully immersing themindiviually in Sigmacote® in a beaker orother container made of glass (repelsilane solutions from other companiescan be used as well). Place the beaker ina fume hood, and cover tightly with foil.Use gloves when handling Sigmacote®.Leave overnight. The next day, lift eachcoverslip by its corner with forceps anddip into a beaker of water and then 100%ethanol to clean. Place each cover-glasson a lint free surface to dry, andthereafter store them in a box untilrequired.2. Blood feed adult females between thehours of 8 to 10:00 am. Remove the fullyblood fed mosquitoes, and hold them in a separate cup or cage in the insectary and supply with 10%sucrose solution. Quarter pint ice cream carton cups with mosquito-proof netting on the top are ideal.When an insectary is not available or if you are in the field collecting blood fed females resting insidedwellings, holds the mosquitoes in cups inside or in the shade of a tree and keep humid by coveringwith a moist cloth.3. Place 1 ml of modified Carnoy’s solution into each 1.5 ml polypropylene vial. Ovaries from onemosquito will be placed into each vial. Anticipate the number of mosquitoes to be dissected to ensuresufficient numbers of vials are prepared in advance. Place the vials at 4C.4. Select half gravid females 18-33 hours after blood feeding. Ovaries will develop to the half gravidstage at different rates depending on the mosquito strain and the temperature and humidity at whichthey are held. Generally at 26C ovaries take about 20 hours and at 30C ovaries take 16- 18 hoursto reach the half gravid stage. Half gravid females (ovaries at Christopher III stage) will appear similarto the image in Figure 6.8.3. When the ovaries take up less than 3/5 of the abdomen they aregenerally too young. When the ovaries extend beyond 3/5 of the abdomen and the developing eggshave a greenish tinge under sunlight they are too old for cropping. Identifying half gravid females withovaries at the appropriate age for harvesting chromosomes will take some practice, and you shoulderr on the side of harvesting a little young at Christopher’s II stage if you are unsure. Polytenizationbegins in Christopher’s II stage and the chromosomes at this stage will be thin, brittle and have lesswell defined bands than at Christopher’s III stage but can be scored by an experienced person afterlight tapping.5. Anesthetize the mosquitoes. This can be done in various ways depending on availability of equipmentand chemicals (see section on Mosquito Anesthesia). The least invasive and easiest is to aspiratehalf gravid females into a container and hold at -20C for five minutes to knock the mosquitoes down.Do not hold them for longer to kill them. Removal of ovaries is best done when the mosquitoes arestill alive as chromosomes degenerate very quickly after death. Place the mosquitoes back into thefreezer for a few minutes if they recover before dissecting. If no freezer is available then anesthetizethe mosquitoes with ethyl ether, triethylamine or C0 2 using an anesthetizing chamber (Figure 6.8.2).Make the chamber by cutting the end of a 45 ml Falcon® tube and covering the cut end with twosheets of rubber latex sheeting with slits just large enough to insert the end of an aspirator blowingmosquitoes into the chamber. Drill holes into the cap of the Falcon® tube and cover the holes with

Chapter 6 : Dissection Techniques6.8 A. gambiae s.l. Ovarian Polytene Chromosome PreparationPage 4 of 6mosquito proof mesh so the mosquitoes do not escape but vapor can pass through. Place theFalcon® tube chamber, with mosquitoes in, into a dessicator saturated with C0 2 gas or a spongesoaked with the anesthetizing chemical (ethyl ether or triethylamine). Wait until the mosquitoes areknocked down before dissection.Figure 6.8.3. Examples of adult females atvarious stages of ovarian development. A –too young, B at correct age and C too old.OV indicates the developing ovaries whileBL indicates undigested blood meal.Photographs taken by Marshall Johnson.6. In conditions where no anesthetizing facilities are available, such as in the field, knock out themosquitoes by shaking the cup or container they are in from side to side for about 30 seconds. Themosquitoes will be stunned by hitting against the sides, but it takes some experience to judge howvigorously to shake the container without damaging the mosquitoes.7. Removing ovaries can be done with or without a dissecting microscope depending on your eyesightand dexterity. Without a microscope - which is by far the quickest way - gently hold the head, thoraxand base of abdomen of the mosquito between the tips of your thumb and index finger. Grab the lasttwo segments of the abdomen with forceps. Gently squeeze the base of the abdomen andsimultaneously pull on the last two abdominal segments with the forceps to extract the ovaries. In thisway, only the ovaries will pull out. Immediately place the ovaries into the vial containing modifiedCarnoy’s solution. Do not let the ovaries dry out after dissecting. After each dissection place a label(write label with a pencil, not a pen) with ovaries into the vial. The remaining head and thorax can bediscarded or preserved in a separate vial for other purposes such as for DNA extraction. If preserved,label the remaining carcass with an accession number corresponding to that of the ovary.8. To remove ovaries under a microscope, pierce the side of the mosquito with a minutien pin and on aclean microscope slide pull the last two apical abdominal segments with a forceps or needle until theovaries have extruded. Immediately place the ovaries into modified Carnoy’s.9. Dissected ovaries fixed and preserved in modified Carnoy’s for more than 24 hours typically providethe best preserved and clean chromosome spreads, especially if required for in situ hybridization.Ovaries preserved in modified Carnoy’s can be stored for many months at 4ºC or for years at -20ºC.10. Placing the whole abdomen in modified Carnoy’s also produces readable spreads but thechromosomes can have a washed out appearance after longer storage due to presence of morewater in the whole abdomen versus ovaries alone. It is not recommended to fix the entire adult inModified Carnoy’s solution as then too much water enters the preservation solution.

Chapter 6 : Dissection Techniques6.8 A. gambiae s.l. Ovarian Polytene Chromosome PreparationPage 5 of 611. To make the chromosome squashes, remove the ovaries from the vials with a pair of forceps andplace them into a drop of modified Carnoy’s (approximately 25 µl) on a dust- and grease-freemicroscope slide. Quickly separate half the ovules or follicles from one ovary, and return the rest ofthe ovaries to the vial to use as back-up for later preparations. You have to do this quickly before themodified Carnoy’s solution dries out on the slide as no spreads can be recovered from dry ovules.12. Drop about 50 µl of 50% propionic acidonto the ovules and leave them for about 3n memABminutes until they have cleared andswollen to about twice their original size.Consult Figure 6.8.4 for further guidanceovulento visualize appropriately aged ovules.Once again do not let the ovules dry out.ovule13. Under a dissecting microscope carefullyovuleseparate the ovules from each other andCDgently squeeze each ovule (follicle) out ofmemits surrounding membrane. Removeovulenconnecting tissues, trachea andmembranes by sliding them away from theovules and draw up the waste tissues withthe tip of tightly rolled up absorbent paper.Figure 6.8.4. Examples of ovules after 5 minutes in 5% 14. From my experience, staining thepropionic acid that were dissected at various stages of chromosomes is unnecessary particularlydevelopment. A – ovule at early Christopher’s II stage in these days of improved microscopewhich is too young for harvesting, Note for comparison optics. Staining with 2% lacto-aceto orceinthe edge of an ovule of appropriate age and size in the is optional and this step should be donetop right hand corner; B and C – ovules at Christopher’s after the ovules have been separated andIII stage of development which are at the correct age cleaned (after completion of step 13). Doand size for producing good chromosome squashes. In this by adding a drop of the stain to theimage B the ovule to the right is too old. In both B and C ovules in the 50% propionic acid. Createthe ovules have popped out of their surroundingan even-staining mixture around the ovulesmembranes (mem) and the nuclei of nurse cells are by stirring gently with a needle. Leave thisenlarged (n); C- ovule too old. All images taken at 100 for about 4 minutes for the ovule cells toX mag. under phase contrast.absorb the stain. Gently absorb the excessstain with a piece of tightly rolled upabsorbent paper and add a drop of 50% propionic acid to the ovules which by now should have apale pink hue to them. Proceed to step 15.15. Place a further drop of 25µl of 50% propionic acid onto the clean ovules.16. Wipe a cover-glass with a clean sheet of lint free paper – lens cleaning tissue or KimWipe®(Kimberley-Clark- Roswell GA) and place on top of the ovules. Ensure the cover-glass is dust andgrease free.17. Carefully set the microscope slide on a piece of absorbent paper on a flat surface.18. Various people use different tapping techniques to make chromosome squashes, but essentially allbegin by tapping quite firmly to break open the nuclear membranes and spreading out thechromosomes followed by harder tapping or pressing to flatten the chromosomes. I prefer to firstgently tap the cover-glass with a mattress needle about 8 to 10 times (end bent downwards asdepicted in Figure 6.8.2). While tapping with the needle the cover-glass will shift around a little whichhelps to shear the nuclear membranes and spread the chromosome arms out. View the slide at 100Xmagnification under phase contrast in a compound microscope to determine if more tapping to spreadchromosomes is required.

Chapter 6 : Dissection Techniques6.8 A. gambiae s.l. Ovarian Polytene Chromosome PreparationPage 6 of 619. Once the first round of tapping has been completed, remove excess propionic acid by gently pressingabsorbent paper over the cover-glass edges. Place a new dry piece of absorbent paper over thecover-glass, and press firmly downwards with a thumb directly over the cover-glass. Do not move thecover-glass laterally while thumb pressing as the chromosomes will roll and be unrecognizable. Scanthe slide at 100X magnification to get an overall perspective if the chromosomes are flat enough. Ifthey are not sufficiently flat then give them a further thumb press. Chromosome spreads can also beflattened by holding the slide at about 70C for 5 to 10 seconds on a slide warmer. Don’t overheat thechromosomes as the banding will fade. View again at 100X magnification and select the bestchromosome compliment to examine at higher power.20. On occasions you may wish to view the spreads under oil at 600 to 1,000X magnification and thendecide to go back to a lower magnification that does not require oil. The oil can be absorbed relativelyeasily from the surface of the cover-glass with tightly rolled up absorbent paper. Oil cannot beremoved effectively from an unsiliconized cover-glass which is another reason why I recommend youroutinely coat cover-glasses (refer to step 1).21. On occasions the slide will begin to dry up (air creeps in between the cover-glass and microscopeslide) while you are examining the spreads under the microscope. Simply add a small drop of 50%propionic acid to the edge of the cover-glass and the acid will, by capillary action, draw under thecover-glass.22. If results are required immediately, chromosomes can be prepared from freshly dissected ovaries byfirst dissecting them out of the female mosquito in 5% propionic and then transferring them into adrop of 50% propionic acid on a clean microscope slide. Continuing with the process as describedfrom steps 11 to 20.23. Interpretable chromosome spreads, but of lesser quality, can be extracted from ovaries preserved forless than 1 hour in modified Carnoy’s solution. This comes in useful in the field when on the spotdecisions are required to know which An. gambiae s.l. member or chromosomal forms of An.gambiae occur in an area. Use the same procedure as described from steps 11 to 20.24. Permanent records of chromosome preparations can be kept either by taking photographs or bymaking permanent mounts. Digital cameras attached to the microscope and computer makecapturing images much easier than the past when emulsion films were used. If permanentpreparations are still required then proceed to the next step.25. Remove the cover-glass by holding the microscope slide in one corner with a forceps and holding inliquid nitrogen until the bubbling stops. Take it out of the liquid nitrogen and immediately place themicroscope slide on a flat surface and briskly pry the cover-glass off with a razor blade form onecorner. Immediately begin dehydrating the preparations placing the slides into an ethanol series of70%, 90% and 100% for 5 minutes each at 4C. Air dry the slide and add a drop of mounting mediumused for histology onto the chromosomes and cover with a cover-glass (does not have to besiliconized). Leave to dry and ring the cover-glasses with sealant according to recommendedmounting medium instructions.ReferencesClements AN (1992) The Biology of Mosquitoes: Development, Nutrition and Reproduction. Chapman &Hall, LondonColuzzi M, Sabatini A, della Torre A, Di Deco MA, Petrarca V (2002) A polytene chromosome analysis ofthe Anopheles gambiae species complex. Science 298:1415-1418Holt RA et al. (2002) The genome sequence of the malaria mosquito Anopheles gambiae. Science298:129-149

Chapter 6 : Dissection Techniques6.9 A. gambiae s.l. Salivary Gland Chromosome PreparationPage 1 of 46.9 An. gambiae s.l. Salivary Gland Chromosome PreparationAnthony Cornel 1IntroductionChromosome preparations from larval salivary glands first require a clean dissection of the salivaryglands from L4 larvae. Fourth stage larvae of appropriate age are at about two to three hours beforepupal trumpets can be seen developing in the thorax under the cuticle. Thereafter, histolysis of thesalivary glands renders them useless for this purpose.Solutions• Modified Carnoy’s fixative (three parts pure (100%) ethanol and one part glacial acetic acid).Ethanol absorbs water from the atmosphere so ensure that the ethanol is pure, as water infixative compromises quality of spreads.• Propionic acid - prepare 5% and 50% in water.• 2% lacto-aceto orcein - prepare this solution by adding slowly 2% by weight of synthetic orceinpowder to a solution of 1 part glacial acetic acid and 1 part pure lactic acid under constant stirring(use a magnetic stirrer). Remove un-dissolved orcein particulates by filtering the solution throughWhatman 3MM paper. The solution can be stored indefinitely at room temperature.• Ethanol – 70%, 90% and 100% in water.Materials• 0.13- 0.17 mm thick cover glasses of dimensions 22L X 22W mm• Frosted 1mm thick microscope slides 75 x 25 mm in size.• Beaker• Aluminum foil• Forceps• Pasteur pipettes• Absorbent paper (Whatman 3MM paper))• Dissecting needles and minutien pins• Dissecting Microscope• 1.5 ml screw cap polypropylene vials• Phase contrast compound microscope with 10X, 40 or 60 X and 100X objective lenses.• Digital imaging systemProcedure1. Dissect salivary glands by placing a larva in a drop of 5% propionic acid on a microscope slide. Severthe abdomen from the thorax and insert a needle from the rear of the thorax along the mid dorsalsurface just underneath the cuticle up to just inside the head. Rub another needle over the inserted1 Mosquito Control Research Laboratory, Department of Entomology and Center for Vector borneDiseases, University of California at Davis, Parlier, CA 93648, E-mail: cornel@uckac.edu

Chapter 6 : Dissection Techniques6.9 A. gambiae s.l. Salivary Gland Chromosome PreparationPage 2 of 4needle to abrade and break the larval cuticle along the mid dorsal line. Carefully open up the thoraxand separate the glands from the connecting tissue but not from their ducts that lead to the head.Gently pull the head away from the thorax and the glands should come out still attached via the ducts.2. Place the head and attached glands in a drop of modified Carnoy’s on a microscope slide for freshpreparations or in a vial of modified Carnoy’s for later squashing.3. For fresh preparations, under the dissecting microscope separate the salivary glands from the headand discard the head.4. Place another drop of modified Carnoy’s onto the salivary glands to fix for a minute.5. Drop about 50 µl of 50% propionic acid onto the salivary glands and leave them for about 3 minutesuntil they have cleared and swollen to about twice their original size.6. Under a dissecting microscope carefully separate the glands from each other. Remove connectingand fatty tissue and trachea if any are still attached to the glands by sliding them away and draw upthe waste tissues with the tip of tightly rolled up absorbent paper.7. Staining with 2% lacto-aceto orcein is optional and this step should only be done after the glandshave been separated and cleaned. Do this by adding a drop of the stain to the glands in the 50%propionic acid. Create an even staining mixture around the glands by stirring gently with a needle.Leave this for about 4 minutes for the gland cells to absorb the stain. Gently absorb the excess stainwith a piece of tightly rolled up absorbent paper and add a drop of 50% propionic acid to the glandswhich by now should have a pale pink hue to them. Proceed to step 8.8. Place a further drop of 25µl of 50% propionic acid onto the clean glands.9. Wipe a cover-glass with a clean sheet of lint free paper – lens cleaning tissue or KimWipe®(Kimberley-Clark- Roswell GA) and place over the glands. Ensure the cover-glass is dust and greasefree.10. Carefully set the microscope slide on a piece of absorbent paper on a flat surface.11. Various people use different tapping techniques to make chromosome squashes but essentially allbegin by tapping quite firmly to break open the nuclear membranes and spreading out thechromosomes followed by harder tapping or pressing to flatten the chromosomes. I prefer to firstgently tap the cover-glass with a mattress needle about 8 to 10 times (end bent downwards). Whiletapping with the needle the cover-glass will shift around a little which helps to shear the nuclearmembranes and spread the chromosome arms out. View the slide at 100X magnification under phasecontrast in a compound microscope to determine if more tapping to spread chromosomes is required.Once the first round of tapping has been completed, remove excess propionic acid by gently pressingabsorbent paper over the cover-glass edges. Place a new dry piece of absorbent paper over thecover-glass and press firmly downwards with a thumb directly over the cover-glass. Do not move thecover-glass laterally while thumb pressing as the chromosomes will roll and be unrecognizable. Scanthe slide at 100X magnification to get an overall perspective if the chromosomes are flat enough. Ifthey are not sufficiently flat then give them a further thumb press. Chromosome spreads can also beflattened by holding the slide at about 70C for 5 to 10 seconds on a slide warmer. Don’t overheat thechromosomes as the banding will fade. View again at 100X magnification and select the bestchromosome compliment to examine at higher power.12. On occasions you may wish to view the spreads under oil at 600 to 1,000X magnification and thendecide to go back to a lower magnification that does not require oil. The oil can be absorbed relativelyeasily from the surface of the cover-glass with tightly rolled up absorbent paper. Oil cannot beremoved effectively from an unsalinized cover-glass which is another reason why I recommend youroutinely salinize cover-glasses (refer to step 1).13. On occasions the slide will begin to dry up (air creeps in between the cover-glass and microscopeslide) while you are examining the spreads under the microscope. Simply add a small drop of 50%

Chapter 6 : Dissection Techniques6.9 A. gambiae s.l. Salivary Gland Chromosome PreparationPage 3 of 4propionic acid to the edge of the cover-glass and the acid will, by capillary action, draw under thecover-glass.14. Salivary glands attached to the head can be preserved up to several weeks and longer for laterchromosome preparation. I recommended dissecting and preserving the salivary glands still attachedto the head via the salivary gland ducts for handling purposes. It is much easier to grab the head withforceps than the smaller salivary gland. Furthermore, there is a much higher likelihood of gettingchromosomes out of intact salivary glands rather than broken ones because only few salivary glandcells produce polytene chromosomes.15. Permanent records of chromosome preparations can be kept either by taking photographs or bymaking permanent mounts. Digital cameras attached to the microscope and computer makecapturing images much easier than the past when emulsion films were used. If permanentpreparations are still required then proceed to the next step.16. Remove the cover-glass by holding the microscope slide in one corner with a forceps and holding inliquid nitrogen until the bubbling stops. Take it out of the liquid nitrogen and immediately place themicroscope slide on a flat surface and briskly pry the cover-glass off with a razor blade form onecorner. Immediately begin dehydrating the preparations placing the slides into an ethanol series of70%, 90% and 100% for 5 minutes each at 4C. Air dry the slide and add a drop of mounting mediumused for histology onto the chromosomes and cover with a cover-glass (does not have to besiliconized). Leave to dry and ring the cover-glasses with sealant according to recommendedmounting medium instructions.

Chapter 6 : Dissection Techniques6.9 A. gambiae s.l. Salivary Gland Chromosome PreparationPage 4 of 4

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 1 of 12Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesTheresa Howard, Ralph Harbach, and Yvonne LintonINTRODUCTIONThe primary purpose of this section is to provide uniform methods for the collection, preservation andrearing of material for the project. The emphasis here is on methods suited for obtaining Anophelesmosquitoes for taxonomic studies. Essential for this type of study is a large amount of uniformly preparedmaterial with all the stages individually associated, general information on bionomics and conspicuousenvironmental factors, and a sample from as many habitats as possible in the study areas. Importantconsiderations in selecting the methods and techniques adopted here have been simplicity and suitabilityfor use under field and laboratory conditions and standardisation and simplification of records andlabelling to minimise errors and to save time.COLLECTION RECORDSCOLLECTION FORM. A standard form (see attached) for recording all the data pertaining to a collectionhas been developed for the project. The form must be filled out in the field as completely as possible andthe remainder added in the laboratory. A pencil should be used for all entries. All measurements shouldbe indicated in the metric system. There is a minimum of writing to be done (only in lined open spaces) onthe form, the rest is to be done by circling or underscoring appropriate words or statements or by placingcheck marks or other signs in appropriate columns.COLLECTION OF ADULTSEQUIPMENT. The basic equipment and supplies needed for the collection of adults are (1) aspirators,(2) plastic vials, (3) covered cups (4) cages and (5) flashlight (torch).CAPTURING. Resting, landing or biting mosquitoes are readily collected with an aspirator one or a fewat a time and then transferred to individual collection vials or covered cups. It is always preferable tocapture mosquitoes with an aspirator but it may not be practicable when very large numbers areencountered. It is also possible to place a tube directly over a resting or biting specimen but to do thisnumerous tubes may be neededKILLING AND STORING. The most satisfactory killing tubes are charged with ethyl acetate but otherkilling agents may be used (e.g. chloroform). The killing tubes should be used exclusively for mosquitoes,and should ideally contain Plaster of Paris to absorb the killing fluid (strips of dry absorbent tissue paperor paper towelling may be used, but when these strips get damp they must be replaced with fresh dryones). The specimens must be removed from the killing tubes within a few minutes after beingintroduced.ISOLATING FOR OVIPOSITION. A number of species of mosquitoes which are very common as adultsare very seldom encountered as larvae and pupae and the immature stages and breeding sites of someof these are poorly known. To obtain the immature stages of these it is necessary to isolate live individualfemales collected in the field, induce them to oviposit and to rear all the stages from the eggs (see thesection on PROGENY REARINGS). It is also frequently desirable to identify cryptic species bychromosomes, enzymes or DNA using this technique for obtaining material. The progeny broods can bedivided for study using different techniques.

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 2 of 12COLLECTING AND RECORDING. Collect mosquitoes in houses and outdoor sites selected for study.Record all the data for every collection on the standard record form (see Appendix 7.1.1 & 7.1.2). Firstenter the locality and general information in the top section of the form. Usually several collections will bemade in one locality. A separate collection number is assigned to every collection in a specific site, of aspecific type on a specific host or bait, and at a specific time of capture. A collection of adults for progenyrearings must be assigned a number distinct from any general adult collection made at the same time andplace. Record all the appropriate items on the collection form. If hourly collections are made to studybiting activity, separate cups must be used for each hour of collection, appropriately labelled, and thespecies and specimens collected should be recorded on an hourly biting record (see Appendix 7.1.3 &7.1.4).COLLECTION OF IMMATURE STAGESEQUIPMENT. The basic equipment and supplies needed for the collection of immature stages are: (1)dippers or pans, (2) aquatic and dip nets, (3) collection bags or vials, (4) plastic pipettes and (5) plastic,enamel or porcelain sorting bowls.COLLECTING AND RECORDING. Because of the much greater percentage of species that can becollected as immature stages as compared to adults the emphasis in taxonomic surveys should be placedon the collection of immature stages, which can then be reared with relative ease in the laboratory toprovide definite association of both sexes and all stages.The immature stages should be collected with great care to prevent injury and should be provided with asufficient volume of water and fine debris from the original breeding site to insure adequate food supplyfor rearing. Larvae and pupae are removed from the breeding site with the appropriate tool such as adipper, aquatic net, dip net, or pan. All immature stages of Anopheles are placed with ample water fromthe breeding site into a plastic cup, sorting bowl or pan until the desired number is obtained. Large debrisis removed from the container and the sediment (if present) is allowed to settle. The larvae and pupae arethen transferred to a plastic bag (twirlpak ® ) or vials for transport to the laboratory. These bags or vialsmust be carefully marked on the outside with the collection number.TRANSPORTING COLLECTIONS. The various containers with immature stages collected in the fieldshould be handled very carefully. Usually no difficulty is encountered and little or no mortality occurs ifsome care is taken in transporting live mosquitoes in field vehicles. First, the containers should be placedaway from direct sunlight and excessive heat; wet towels may be placed over and around the containerswhen high temperatures are encountered. Second, drive carefully avoiding high speeds, sudden stopsand excessive bouncing. Third, transport larvae and pupae over long trips or rough terrain by floatingcollection bags (these are preferred over vials for this reason) in water contained in a cool box or largeplastic bag. The water absorbs the shock of rough travel.INDIVIDUAL REARINGS. Individual rearings are made from either pupae (pupal rearings) or larvae(larval rearings), which have been isolated in individual plastic vials and marked with the country identifierand collection number. Each individually reared specimen must be provided with an individual rearingnumber, which will identify the stages of each individual.FACILITIES AND EQUIPMENT. A large cool room with electricity, running water and several tables orlaboratory benches serves very well as a simple laboratory for mosquito rearing. The room should beant-proof but if it is not, the legs of the tables or benches should be placed in cans of oil. Air-conditioningis a convenience for the workers but is not necessary or even desirable for mosquito rearing.A laboratory or a collapsible stereoscopic field microscope is extremely useful. The basic equipment andsupplies required for rearing and processing are: (1) pipettes and medicine droppers, (2) aspirators, (4)plastic, enamel or porcelain collecting bowls, (5) plastic rearing vials, (6) applicator sticks, (7) glass vials

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 3 of 12and lids, (8) 80% and 95% ethanol, (9) labels and (10) pencils, grease pencils. All the equipment shouldbe for use exclusively in the laboratory and should not be taken into the field.CARE OF COLLECTIONS AND SORTING. Immediately after a field trip all the collections must bechecked carefully. Labels on all containers should be made plainly visible and legible.Set up immature collections by emptying contents of collection bags and vials into separate bowls. Labeleach bowl with the same number marked on the collection bag or vial. The sorting of the immature stagesand their separation for individual rearings and preservation can now be carried out. The sooner this isdone the better, and this should always be completed within 24 hours of capture. Work with onecollection at a time going through the whole process before turning to the next collection. If more thanone collection bag or vial contains immature stages from the same collection, assemble them all together.Generally, more than one species is found in a single collection of immature stages even whencollections are made from individual breeding sites, but as a rule one species is dominant in a particularcollection and the others are frequently represented by few specimens or by younger larval instars.First, all the pupae are transferred one to a plastic vial in about 2 cm (somewhat less than one inch) offresh clean water (always use rainwater or distilled water). The plastic vials are marked with thecollection number (grease or wax pencil is preferred but paper labels can be used). These will be laterprocessed as indicated below under EMERGENCE VIALS. As a rule, all the pupae present in a collectionshould be isolated individually unless the collection consists primarily of pupae and the total numberexceeds 10. If it is obvious that several species are represented among the pupae, a larger numbershould be isolated individually (see EMERGENCE VIALS below).Second, the fourth-instar larvae are picked up one by one and placed into separate plastic vials. Isolateall mature larvae in the collection, or a maximum of 10 per species if the collection is very large. Theindividual vials are filled with distilled water or rainwater to a height of about 2 cm (somewhat less thanone inch), marked with the collection number in grease pencil and set aside to be processed as indicatedbelow under PUPATION VIALS. The remaining larvae in each collection, normally up to a maximum of20, are set aside for killing and preservation (see WHOLE LARVAE in the section on KILLING ANDPRESERVATION), and any larvae left over are retained in the bowl and removed for individual rearing asthey become fourth instars. In case it is not practical or desirable to take care of a large numbers ofcollections or rearings, all the larvae remaining after isolation of individuals should be killed and preserved(NEVER DISCARD ANY MATERIAL ONCE COLLECTED). Great care should be taken to thoroughlyrinse sorting pans and pipettes between the processing of different collections to eliminate contamination.WASHING CONTAINERS. All containers used for collecting, sorting and rearing must be washedthoroughly before they are used again. First, wipe off the grease pencil markings (if used) from the drybowls and vials with a piece of cotton or paper. Containers that are reasonably clean may be merelyrinsed several times in clean fresh water. Aspirators should also be cleaned periodically and the nettingreplaced on the plug.PUPATION VIALS. Vials containing isolated larvae, marked with collection numbers, should beexamined twice a day for pupation, preferably in early morning and late afternoon. Check through all thevials and set aside all those containing larval skins before further processing. Remove the larval skin withan applicator stick (without dragging it along the side of the rearing vial) and transfer it into a storage vialwith 80% alcohol and attach it with an elastic band to the rearing vial containing the pupa. Place a lid onthe rearing vial to prevent escape of the adult mosquito after it emerges from the pupa. This marked vialis now set aside to be processed as indicated under REARED ADULTS in the section on KILLING ANDPRESERVATION below.EMERGENCE VIALS. Vials containing isolated pupae should be examined twice a day, preferably inearly morning and late afternoon. Check all the vials and set aside all those with emerged or drowned

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 4 of 12adults for processing. To remove the viable emerged adult loosen the lid and carefully slip in and replaceit by the mouth of an inverted plastic holding vial. Tap on the side of the pupal container to induce themosquito to enter the holding vial and quickly stopper it with the original lid. Label this holding vial with thesame number that appears on the rearing vial containing the pupal skin. Remove the pupal skin with anapplicator stick (without dragging it along the side of the vial) and place it in the vial of 80% ethanol withthe associated larval skin. Completely fill the vial with ethanol to exclude air and attach it to the plasticholding vial containing the adult mosquito. This vial will now be processed as indicated below underREARED ADULTS. The entire process is repeated for all the pupal vials.All vials containing dead pupae and drowned, partially emerged adults stuck to the water should bediscarded. These specimens are not useful for taxonomic, but if they are alive they may be preserved forenzyme or DNA analysis.HOLDING VIALS. Reared adults for taxonomic study should be kept for at least 24 hours before beingkilled and processed as indicated in the section on REARED ADULTS. Many adults may die before theyare processed, in which case they will more difficult to mount on points on pins for study.The individual holding vials require relatively little attention. Check at least twice a day and process deadspecimens immediately or as soon as possible, following the directions in the section on REAREDADULTS. After the required 24-hour holding period, process the live specimens following the samedirections.PROGENY REARINGSGravid females collected in the field will generally oviposit within 3 to 4 days if they will lay eggs at all.Females should be retained in a cool damp environment and provided with 5-10% sucrose solution insaturated wads of cotton. The cotton wads should be moistened or replaced once or twice daily. On thethird day following capture, each female is lightly anaesthetised with ethyl acetate or knocked down in afreezer (about 1 minute) for the purposes of removing a wing. This is accomplished by grasping eachwing with a forceps and gently pulling until one of the wings is torn from the thorax. The anaesthetisedfemale is then placed on the surface of water (containing a hatching stimulus) in a small cup. Femalesstressed in this manner usually oviposit soon after being placed on the water. Some gravid females willrefuse to lay eggs and these can be preserved for enzyme or DNA studies. Once larvae become late firstor early second instars, they should be transferred to a larger bowl or pan and fed at least once per dayby sprinkling powdered food onto the surface. A portion of fourth-instar larvae (pre-pupae) should beremoved and reared individually for taxonomic study as described in the section on INDIVIDUALREARINGS. Remaining larvae can be preserved for enzyme or DNA analysis.KILLING AND PRESERVATIONEQUIPMENT AND SUPPLIES. The following equipment and supplies are needed for killing andpreserving mosquitoes: (1) killing tubes (preferably with ethyl acetate), (2) aspirators, (3) forceps, (5)labels, (6) 80% and 95% ethanol, (7) applicator sticks or scoop, (8) glass vials with lids, (9) beaker or panfor hot water.FIELD-COLLECTED ADULTS. Adult mosquitoes collected in the field are usually killed, identified andpreserved immediately on return to the laboratory. The method of preservation will depend on the type ofstudy they will be used for. Some blood-fed adults will be retained and set-up to obtain progeny broods(see PROGENY REARINGS).REARED ADULTS. All reared adults after being held for 24 hours in plastic vials (see HOLDING VIALS)should be killed and processed as indicated below. Parent females in PROGENY REARINGS should beprocessed immediately after oviposition following the procedure required for the type of study to beundertaken (i.e. morphology, enzymes or DNA).

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 5 of 12Process all the adults (dead or alive) from individuals that have been held for the length of time (24 hours)together at some convenient time in the afternoon or evening. Numbers are assigned to individualspecimens at this stage, and the necessary information is recorded on the appropriate collection andrearing form (see Figures 7.1.1 and 7.1.2). For example, an adult female with larval and pupal skins isgiven the next available number on the form, perhaps 12-2, indicating that this is the second specimenreared from collection 12. The first specimen from this collection would have been labelled 12-1. Thenumber needs to be written on 2 labels, one for the adult and one for the vial containing the associatedskins. Pre-printed labels may be used to avoid errors and increase efficiency.To speed the process, have at hand 2-5 killing tubes to be used serially. Loosen the lid of a holding vialand holding the vial upside down slip its mouth over that of a killing tube. The adult will be knocked downby the ethyl acetate within seconds and will fall into the killing tube. To hasten knockdown, tap on thevial. Remove the vial and hold a thumb over the mouth of the killing tube until the specimen ceasesstruggling. Place the label inside the killing tube, stopper the tube and set it aside. Proceed in the samemanner with 1-4 other adults and place the killing tubes in sequence. After all the tubes are used, returnto the first one and process it as indicated in the next paragraph. After this step is finished kill anotheradult with the emptied tube and place it in sequence after the others. Continue in this manner until all theadults are processed. Time the processing to allow the adult to remain in a tube about 5 minutes, but notlonger than 10 minutes by using the appropriate number of killing tubes.Transfer the adult and its label from the killing tube to a clean white card. Using fine forceps pick up theadult by one leg and place it on an elevated surface (e.g. empty slide box) with a white background.Orient the specimen with its left side facing down. This should be accomplished without grasping thespecimen with the forceps. Move the specimen to the edge of the surface with its legs projecting beyondthe edge. Insert a heavy paper point an appropriate distance from the head of an insect pin and place atiny droplet of ambroid ® cement or other suitable glue on the upper apical angle of the point. Holding thepin so that the point is upside down, gently touch the droplet of the glue to the thorax (right side) of themosquito. Final orientation of the mosquito on the upper surface of the point is with the left side up, headfacing left and the legs extended toward the pin. This orientation protects the specimen from damage andcorresponds to the preferred orientation of illustrations in taxonomic publications. Place the label with thecollection and rearing number on the pin and store the specimen in an appropriate insect storage box.WHOLE LARVAE. It is essential to preserve an adequate sample of whole larvae of every species fromevery field collection and from all progeny rearings (see CARE OF COLLECTIONS AND SORTING in thesection on INDIVIDUAL REARINGS). To be useful for taxonomic purposes, whole larvae must be killedand preserved carefully so that the body shape and all structures, particularly setae, are retained. Thelarvae set aside for killing and preservation (identified by the collection number) are first transferred with adropper to a small cup or bowl with fresh clean water as a washing procedure. If much debris or sedimentis still present, additional serial transfers should be made until it is eliminated. Remove as much water aspossible from the cup or bowl using a fine pipette. Next heat a beaker or pan of water is about 60ºC(140ºF) and pour the hot water into the cup or bowl. As soon as the larvae float up to the surface, thewater is removed with a fine pipette and replaced with a quantity of 80% ethanol. After 5 minutes thelarvae are transferred with a lifter (do not use forceps) to a vial with 80% ethanol. Completely fill the vialwith 80% ethanol to remove all air and cap it tightly. No more than 20 larvae should be placed into asingle vial as the water contained in the bodies of the larvae will significantly dilute the concentration of asmall quantity of ethanol and jeopardise preservation. Prepare a paper label in pencil and tape it to a vial.The label may be placed inside the vial if the larvae are separated from the label by a small wad of cotton.Larvae to be used for DNA studies should ONLY be killed by placing them into a vial of 95% ethanol.SKINS. The most valuable material for taxonomic purposes is the associated larval and pupal skins fromindividual rearings and the corresponding adults. The greatest care must be taken in processing these.Labels should not be enclosed in vials with skins unless these are separated by a small wad of cotton. It

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 6 of 12is far better to tape a label to the outside of the vial, or to write the specimen number on a piece of tapeand wrap it around the vial in a lightly overlapping manner to enhance adhesion.PRESERVATION AND MOUNTING TECHNIQUESWHOLE LARVAE1. Kill larvae in hot water (not boiling), remove promptly with a lifter. Store in small vial containing 80%ethanol (ethyl alcohol). For material to be used for DNA studies, larvae should be killed by placingdirectly into 95% ethanol – DO NOT kill in hot water.2. Transfer specimens from alcohol to cellosolve for 15 minutes or more (dark specimens can be storedin cellosolve for 8 hours or overnight).3. Lift the specimen from cellosolve and place on the centre of glass microscope slide with the dorsalside up.4. Drop a small amount of Euparal on the specimen. Mount specimen dorsal side up with the headpointing down; arrange head, thorax and abdomen in natural position, then cut the abdomen betweensegment VI and VII (with a fine scalpel blade). Place the terminal segments with siphon to the left inculicine larvae or segment X to the right in anopheline larvae (see FIGURE 7.1.3).5. Place more Euparal on the specimen and check the arrangement of setae and larval position, thencarefully cover the specimen with a 22 mm rectangular cover glass.6. Dry in an oven at 45º-55ºC for 4 weeks or more.LARVAL AND PUPAL EXUVIAEThe fourth-instar larval and pupal exuviae from an individually reared adult should be mounted on thesame slide.1. Store in 80% alcohol.2. Transfer the specimens into cellosolve for 15 minutes.3. Lift the specimen from cellosolve placing it on a glass microscope slide, the larval exuviae on the leftand pupal exuviae on the right (pointing head down and dorsal side up).4. Drop a small amount of Euparal on the specimens. Arrange and spread the body and setae of larvalexuviae into better position, then separate the pupal cephalothorax just cephalad of the wing, leavingthe metanotum attached to the abdomen. Open the cephalothorax, mount it ventral side up andplace it below the metanotum (see FIGURE 7.1.3).5. Add more Euparal, check the position of the exuviae then cover the specimens with a 15 mm roundcover glass.6. Dry in an oven at 45º-55ºC for 4 weeks or more.ADULTS1. After emergence adults should be held for at least 24 hours before killing.2. Kill in a killing tube charged with ethyl acetate or chloroform 1 (ethyl acetate is preferred because itkeeps specimens relaxed longer).3. Apply a small amount of Ambroid ® cement 2 or other glue to the tip of a paper point and affix thespecimen on the right side of the thorax with the legs toward the pin (see FIGURE 7.1.3).1 CAUTION! Chloroform and Ethyl Acetate are toxic and dangerous to breathe. These chemicals are stored in liver tissues andmay cause health problems if used frequently. Always use in well ventilated areas.2 Ambroid ® cement should be thinned down with amyl acetate.

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 7 of 12Pinned specimens should be kept in boxes. Seal each box in a self-seal or taped closed plastic bag. Ifpossible include a sachet of Silica Gel in the bag to keep specimens dry. The bag should prevent thespecimens from being eaten by beetles, cockroaches, mites and other insects.LABELING (see FIGURE 7.1.3)Label the slide with 2 labels:Left label contains:Country, collection numberProvince, date collectedLocationCollector's nameHabitatSlide number3 4Right label contains:GenusSubgenusSpeciesPerson making the determinationLabel the adult with 2 labels. Each label should be ¼” x ½” in size or smaller.Upper label contains:Country, provinceLocationCollection NumberDate/YearLower label contains:Genus, subgenus and speciesSexPerson making the determinationSTORING, PACKING AND SHIPPINGSTORING. All preserved adults and all supplies used for the preservation of adults must be stored in adry pest-proof box to protect them from mold and insects. A simple cardboard shipping box sealed in aplastic bag or beem capsules in individual sealed plastic bags with a drying agent placed inside theplastic bags or air-tight wood, pest-proof box can be used to store the material. The vials with ethanolshould be kept in a cool place. The vials should be packed neatly in cardboard containers of a convenientsize or wrapped together in paper towelling in bundles of 20 to 50. Vials containing alcohol should havethe tops tightly sealed to prevent evaporation and be packed in small cardboard boxes. Microscopeslides should be stored in slide boxes with the cover-slip parallel to the surface of the storage shelf toreduce the risk of slippage in cases where the mounting media is not 100% dry.PACKING AND SHIPPING. Material should be shipped as soon as possible for final processing andmounting. Do not let large quantities accumulate but send it in small parcels. Be sure to include thecollection forms with the preserved specimens.

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 8 of 12It is very important to pack the material very carefully or it may be completely ruined. Use a sturdycorrugated cardboard box lined with appropriate packing material. All the material to be shipped mustfirst be carefully packed in small cardboard boxes as indicated in the section above. Check every box forloose containers and fill in the unused space with cotton or crumpled paper so that nothing will movewhen the box is shaken but do not pack too tightly. Next seal the individual boxes with tape. Place all theboxes in the shipping container and fill all the spaces between them tightly with crumpled paper so thatnothing moves and the container is completely filled. Enough packing material should be placed on top ofthe boxes so that when the lid is put on a slight pressure will be needed to keep it down. Now close theshipping container and seal the top with tape. The container can then be covered with wrapping paper ifrequired.If pinned specimens in Schmitt or other boxes are shipped, a corrugated cardboard container lined with 2inches of packing material should be used. Make certain that all the pins are inserted tightly in the box.Remove all the foreign material such as drying agents or naphthalene flakes from the box. If large labelshave been used, insert pins on each side of each label to prevent it from swinging in transit. Microscopeslides should be secured in slide boxes with strips of paper or foam rubber to prevent movement duringshipment. All packages should be shipped by air, preferably by AIR PARCEL POST. All packages shouldbe plainly marked "Preserved material for scientific study, no commercial value."

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 9 of 12Figure 7.1.1.Comprises 2-letter country identifier andindividual collection locality numbere.g. IN12 (12th collection site in Indonesia)Any additional information that may be usefuli.e. how many whole larvae preserved in ethanol

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 10 of 12Figure 7.1.2.The same country identifier and numberas on the main collection sheet - these formsare usually printed back-to-backIndividual specimen number -each specimen will have a uniquenumber comprising the CollectionNumber and this numbere.g. IN12-1Which life stage rearedor obtained - may onlybe an adult collectionSex of individualspecimen

Figure 7.1.3.Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 11 of 12

Chapter 7 : Taxonomy and Systematics7.1 Methods for Collecting and Preserving MosquitoesPage 12 of 12APPENDECIES 7.1.1 – 7.1.4Appendix 7.1.1 & 7.1.2: Collection Forms - pages 13-14Appendix 7.1.3 & 7.1.4: Biting Collection Record - pages 15-16(When printing - print page 14 on the reverse of page 13 (additional rearing forms can be printedseparately) and 16 on the reverse of 15)

Collection No. Nearest Town DateCountry Specific Locality TimeProvince Latitude/Longitude Collector(s)Second Administrative DivisionElevationOrganisationmCOLLECTION TYPE ENVIRONMENT LARVAL HABITAT WATER:Immature Rain Forest Pond - Lake PermanentResting - Evergreen Forest Ground Pool TemporaryHouse Deciduous Forest SwampAnimal Shelter Cloud Forest Marshy Depression WATER MOVEMENTCave Coniferous Forest Stream Margin StagnantTree Hole Scrub/Bush Stream Pool SlowVegetation Savanna Rock Pool ModerateOther: ____________ Prairie Seepage - Spring FastBiting/ Landing - Island DitchHuman Swamp Well SALINITYAnimal: ___________ Salt Marsh Artificial Container FreshNet Beach Hoof Print BrackishLight Trap: ____________ Mangrove RutBait Trap Orchard - Plantation Rice Field TURBIDITYSwarming Cultivated Field: ____________ Mangrove ClearAt Light Bamboo Grove Other: ____________ ColouredOther: ____________ Urban TurbidVillage ALGAE PollutedTERRAIN Other: ____________ FilamentousMountain Green PHYSICAL FACTORSHill ENVIRONMENTAL MODIFIERS Blue-Green pHValley Primary Brown ConductivityPlateau Secondary Other: ____________ Temperature (ºC)PlainTDSAgricultureALGAL DENSITYDISTANCE FROM HOMES Pasture None AQUATIC VEGETATIONm Grove/Plantation: ____________ Scarce SubmergedOther: ____________ Moderate FloatingSKY Abundant EmergentClear WIND Submerged and FloatingPartly Cloudy None DIMENSIONS OF SITE Submerged and EmergentOvercast Light m X m Floating and EmergentFog Gusts All TypesMist Strong Depth mLight RainQUANTITY OF AQUATIC VEG.Heavy Rain HEIGHT ABOVE GROUND NonemScarceSHADEModerateNoneAbundantPartialHeavyREMARKSHOSTHumanHorseCowOther: ____________

Collection NumberCountryNumber Le Pe Sex Identification / Notes

MOSQUITO BITING COLLECTION RECORDColl. No. ___________________________________Date ___________________________Host _____________________________________ Collector ________________________Location _________________________________________________________________________Weather ___________________________________Temp. (W/D) _____________________Habitat __________________________________________________________________________Collection: Inside Outside OtherSPECIES 2400-0059 0100-0200-0259 0300-0359 0400-0459 0500-0559 TOTAL0159TOTAL

MOSQUITO BITING COLLECTION RECORDColl. No. ___________________________________Date ___________________________Host _____________________________________ Collector ________________________Location _________________________________________________________________________Weather ___________________________________Temp. (W/D) _____________________Habitat __________________________________________________________________________Collection: Inside Outside OtherSPECIES 1800-1859 1900-2000-2059 2100-2159 2200-2259 2300-2359 TOTAL1959TOTAL

Chapter 7 : Taxonomy and Systematics7.2 Ribosomal DNA PCR Assays7.2.1 Amplification of the Second Internal Transcribed Spacer Region (ITS2) in AnophelinesPage 1 of 27.2 Ribosomal DNA PCR Assays7.2.1 Amplification of the Second Internal Transcribed Spacer Region (ITS2)in AnophelinesMR4 StaffIntroductionIn many insect genera, often the amplification and sequencing of the ITS2 region is useful in order todetect intraspecific differences. The ribosomal DNA region is made up of 3 functional rDNA genesseparated by two spacer regions: the ITS1 and ITS2 regions. In many anopheline species it has beenfound that the ITS2 region is more informative, especially in designing PCR assays to distinguishmembers of cryptic complexes. Universal primers designed by Beebe and Saul (1995) and have beenshown to be suitable in a number of anopheline species.PCR assay for amplifying the ITS2 region in anopheline mosquitoes (Beebe and Saul 1995)Prepare PCR Master Mix for 96, 48 or 1 25 μl PCR reactions 1 . Add reagents in the order presented.96 48 1 Reagent1555 μl 777.5 μl 15.55 μl sterile H 2 O500 μl 250 μl 5.0 μl GoTaq 5X PCR Buffer with MgCl 2100 μl 50 μl 1.0 μl dNTP (2.5 mM mix)150 μl 75 μl 1.5 µl ITS2 A primer (1 pmol/μl) [TGT GAA CTG CAG GAC ACA T]150 μl 75 μl 1.5 µl ITS2 B primer (1 pmol/μl) [TAT GCT TAA ATT CAG GGG GT]30 μl 15 μl 0.3 µl MgCl 2 (25 mM)15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 24 μl Total (To each 24 ul reaction add 1 μl template DNA)Cycle conditions94°C/4min x 1 cycle(94°C/30sec, 53°C/40sec, 72°C/30sec) x 35 cycles72°C/10min x 1 cycle4°C holdPCRRun samples on a 2.0% agarose EtBr gel for visualization. Band sizes will vary by species (Figure7.2.1.1).ReferencesBeebe NW, Saul A (1995) Discrimination of all members of the Anopheles punctaulatus complex bypolymerase chain reaction-restriction fragment length polymorphism analysis. Am J Trop Med Hyg53:478-4811 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 7 : Taxonomy and Systematics7.2 Ribosomal DNA PCR Assays7.2.1 Amplification of the Second Internal Transcribed Spacer Region (ITS2) in AnophelinesPage 2 of 2Figure 7.2.1.1. Lane 1, 1kb ladder, lane 2, An.gambiae, lane 3, An. arabiensis, lane 4, An.freeborni, lane 5, An. sawadwongporni, lane 6,An. albimanus, lane 7, An. quadrimaculatus, lane8, An. stephensi, lane 9, An. minimus96 well sample preparation template

Chapter 7 : Taxonomy and Systematics7.2 Ribosomal DNA PCR assays7.2.2 PCR amplification of expansion segments within the 28S subunitPage 1 of 27.2.2 PCR amplification of expansion segments within the 28S ribosomal DNA inAnopheles mosquitoesIntroductionThe most common molecular systematic and phylogenetic assays for anophelines target the ribosomalDNA (Krzywinski and Besansky 2003). Ribosomal DNA (rDNA) is a tandem-repeated array comprised ofthree subunits (18S, 5.8S, and 28S) and several intergenic spacer regions (reviewed in Hwang and Kim1999). Although the spacer regions evolve faster than the transcribed regions, the 18S and 28S subunitsare comprised of several expansion segments which have varying levels of interspecific polymorphism(Hancock et al. 1988). The large subunit (28S) is the largest of the three units and has been found toshow more interspecific polymorphism within its domains compared to the small subunit (18S) (Hwangand Kim 1999). Here we present PCR assays for three of the domains located within the 28S subunit.PCR primers for specific expansion segments and their associated product sizesApprox.Domain Primer sequence Tm Product size ReferenceD2 forward GTG GAT CCA GTC GTG TTG CTT GAT AGT GCA G 60 560 bp Porterreverse GTG AAT TCT TGG TCC GTG TTT CAA GAC GGG 60D3 forward GAC CCG TCT TGA AAC ACG GA 50 415 bp Baldwinreverse TCG GAA GGA ACC AGC TAC TA 50D7 forward CTG AAG TGG AGA AGG GT 42 480 bp Friedrichreverse GAC TTC CCT TAC CTA CAT 42Table 7.2.2.1. Primers are presented 5’ to 3’. Product sizes may vary between species due to expansionof the domains.PCR amplification of the expansion segments of the 28S subunit in anophelinesPrepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1.49 ml 745 µl 14.9 µl sterile H 2 O500 µl 250 µl 5.0 µl 5X PCR Buffer100 µl 50 µl 1.0 µl dNTP (2 mM mix)100 µl 50 µl 1.0 µl Forward primer (25 pmol/µl)100 µl 50 µl 1.0 µl Reverse primer (25 pmol/µl)100 µl 50 µl 1.0 µl MgCl 2 (25mM)10 µl 5.0 µl 0.1 µl Taq DNA polymerase (5U/μl)2.4 ml 1.2 ml 24.0 µl TotalTable 7.2.2.2. Use 1 µl of DNA template per reaction.PCR Cycle conditions95°C/5min x 1 cycle(95°C/30sec , **Tm 1 °C/30sec , 72°C/45sec) x 30 cycles72°C/5min x 1 cycle4°C hold1**Tm, Information for specific annealing temperatures is shown in Table 7.2.2.1 for each of the expansion segments.

Chapter 7 : Taxonomy and Systematics7.2 Ribosomal DNA PCR assays7.2.2 PCR amplification of expansion segments within the 28S subunitPage 2 of 296 well PCR sample preparation templateRun samples on a 1.5% Agarose EtBr gel; load 10 μl of the sample.Primers create a single band that is specific for each domain as listed in table 7.2.2.1 (Figure 7.2.2.1).Figure 7.2.2.1. D7 expansion segment: lane 1 1Kb marker,lanes 2-9 An. subpictus samples.ReferencesBaldwing JG, Frisse LM, Vida JT, Eddleman CD, Thomas WK (1997) An evolutionary framework for thestudy and developmental evolution in a set of nematodes related to Caenorhabditis elegans. Mol PhylogEvol 8: 249-259.Friedrich M, Tautz D (1997) An episodic change of rDNA nucleotide substitution rate has occurred duringthe emergence of the insect order Diptera. Mol Biol Evol 14: 644-653.Hancock JM, Tautz D, Dover GA (1988) Evolution of the secondary structures and compensatorymutations of the ribosomal RNAs of Drosophila melanogaster. Mol Biol Evol 5: 393-414.Hwang UW, Kim W (1999) General properties and phylogenetic utilities of nuclear ribosomal DNA andmitochondrial DNA commonly used in molecular systematics. Korean J Parasit 37: 215-228.Krzywinski J, Besansky NJ (2003) Molecular systematics of Anopheles: from subgenera tosubpopulations. Ann Rev Entomol 48: 111-139.Porter CH, Collins FH (1996) Phylogeny of Nearctic members of the Anopheles maculipennis speciesgroup derived from the D2 variable region of 28S ribosomal RNA. Mol Phylog Evol 6: 178-188.

Chapter 7 : Taxonomy and Systematics7.2 Ribosomal DNA PCR assays7.2.3 General PCR to amplify a portion of the 18S subunit in anophlinesPage 1 of 27.2.3 General PCR to amplify a portion of the 18S rDNA subunit in anophelinesMR4 Vector ActivityIntroductionThe rDNA is a multicopy (>100) locus useful for phylogenetic studies due to its rapid, concerted evolution.It is employed to distinguish between cryptic species since nucleotide differences will quickly fix aftergene flow between two populations ceases.PCR assay for the amplification of the 18S subunit regionPrepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1.54 ml 770 μl 15.4 μl Distilled H2O500 μl 250 μl 5.0 μl 5X PCR buffer150 μl 75 μl 1.5 μl dNTP (2mM concentration)100 μl 50 μl 1.0 μl 18SForDros (25pmol/ul) [GAG GGA GCC TGA GAA ACG GCT AC]100 μl 50 μl 1.0 μl 18SRevDros (25pmol/ul) [CCT TCC GTC AAT TCC TTT AAG TTT C]100 μl 50 μl 1.0 μl MgCl2 (25 mM)10 μl 5.0 μl 0.1 μl Taq DNA polymerase (5U/ μl)2.5 ml 1.25 ml 25 μl Total (To each 25 μl reaction add 1.0 μl template DNA)Table 7.2.3.1. F and R indicate forward and reverse orientation. DNA extraction negative control to beincluded in addition to PCR reaction mix negative control.PCR Cycle conditions94°C/5min x 1 cycle(94°C/1min , 54°C/1min , 72°C/1min) x 35 cycles72°C/7min x 1 cycle4°C holdUse a 1.5% Agarose ethidium bromide gel for visualization.Primers create an approximately 900 bp band (Figure 7.2.3.1).Figure 7.2.3.1. Lane 1 1Kb marker, Lane 1, 1kb ladder, lanes 2-9 An.subpictus, lanes 10-11 An. gambiae.

Chapter 7 : Taxonomy and Systematics7.2 Ribosomal DNA PCR assays7.2.3 General PCR to amplify a portion of the 18S subunit in anophlinesPage 2 of 296 well PCR sample preparation templateReferencesBeebe NW, Foley DH, Cooper RD, Bryan JH, and Saul A. 1996. DNA probes for the Anophelespunctulatus complex. Am J Trop Med Hyg. 54: 395-398.

Chapter 7 : Taxonomy and Systematics7.3 Mitochondrial DNA PCR assays7.3.1 General PCR for amplification of Cytochrome c Oxidase I and II in anophelinesPage 1 of 27.3 Mitochondrial DNA PCR Assays7.3.1 General PCR for amplification of Cytochrome c Oxidase I and II inanophelinesIntroductionMitochondrial DNA (mtDNA) is one of the most commonly studied regions in insect systematics due to itshigh rate of homoplasmy (Caterino et al 2000). Within the mtDNA there are several segments of whichonly a few are routinely examined: cytochrome c oxidase I (COI), cytochrome c oxidase II (COII), andcytochrome b (cytb – see Chapter 7.3.2).Primer sequenceReferenceCOI forward GGA GGA TTT GGA AAT TGA TTA GTT CC Beard 1993COI reverse GCT AAT CAT CTA AAA ATT TTA ATT CCCOII forward TCT AAT ATG GGA GAT TAG TGC Simon 1994COII reverse ACT TGC TTT CAG TCA TCT AAT GTable 7.3.1.1. Primer sequences and references used in these assays.Prepare PCR Master Mix for 96*, 48* or 1 25μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent1.24 ml 620 μl 12.40 μl sterile H 2 O500 μl 250 μl 5.0 μl 5X PCR Buffer (go taq Promega)200 μl 100 μl 2.0 μl dNTP (25 mM mix)150 μl 75 μl 1.5 μl Forward primer (600 nM/ μl)150 μl 75 μl 1.5 μl Reverse primer (600 nM /μl)250 μl 125 μl 2.5 μl MgCl 2 (25mM)10.0 μl 5.0 μl 0.1 μl Taq DNA polymerase (5U/μl)2.5 ml 1.25 ml 25 μl TotalTable 7.3.1.2. Add 1 µl of template gDNA to each reaction. DNA extraction negative control to beincluded in addition to PCR reaction mix negative control.PCR Cycle conditions95°C/3min x 1 cycle(95°C/30sec , 55°C/30sec , 72°C/45sec) x 35 cycles72°C/10min x 1 cycle4°C holdRun samples on a 1.5% agarose gel. You will expect the following fragment sizes: COI – 700bp, COII –730 bp (Figure 7.3.1.1.).

Chapter 7 : Taxonomy and Systematics7.3 Mitochondrial DNA PCR assays7.3.1 General PCR for amplification of Cytochrome c Oxidase I and II in anophelinesPage 2 of 2Figure 7.3.1.1. Lane 1, 1kb ladder, lanes 2-6 COI ampliconfor An. subpictus, lane 7 negative control, lane 8 1kb ladder,lanes 9-13 COII amplicon for An. subpictus, lane 14negative control.96 well PCR sample preparation templateReferencesb) c)Beard CB, Hamm DM, and Collins FH. 1993. The mitochondrial genome of the mosquito Anophelesgambiae: DNA sequence, genome organization, and comparisons with mitochondrial sequences of otherinsects. Insect Mol Biol 2: 103-124.Caterino MS, Cho S, and Sperling FAH. 2000. The current state of insect molecular systematic: a thrivingtower of Babel. Ann Rev Entomol. 45: 1-54.Simon C, Frati F, Beckenbach A, Crespi B, Liu H, and Flook P. 1994. Evolution, weighting, andphylogenetic utility of mitochondrial gene sequences and a compilation of conserved polymerase chainreaction primers. Ann Entomol Soc Am. 87: 651-701.

Chapter 7 : Taxonomy and Systematics7.3 Mitochondrial DNA PCR assays7.3.2 General PCR to amplify the cytochrome B region in anophlinesPage 1 of 27.3.2 General PCR to amplify the cytochrome B region in anophelinesMR4 Vector ActivityIntroductionAlthough most phylogenetic studies involve one of the cytochrome oxidase genes (COI and COII) or oneof the D domains in the 28S subunit of the rDNA, the cytochrome B (cytb) gene has been usedextensively by vertebrate scientists (Simmons 2001). The utility of mitochondrial DNA (mtDNA) forphylogenetic studies is based on its lack of recombination, maternal inheritance, rapid evolution, andintraspecific polymorphisms (Avise et al. 1987).PCR assay for the amplification of the cytochrome B regionPrepare PCR Master Mix for 96, 48 or 1 50μl PCR reactions. Add reagents in the order presented.96 48 1 Reagent3.32 ml 1.66 ml 33.2 μl sterile H 2 O1.0 ml 500 μl 10.0 μl 5X PCR Buffer200 μl 100 μl 2.0 μl dNTP (2 mM mix)200 μl 100 μl 2.0 μl cytbF (25 pmol/ μl) GGA CAA ATA TCA TTT TGA GGA GCA ACA G200 μl 100 μl 2.0 μl cytbR (25pmol/ μl) ATT ACT CCT CCT AGC TTA TTA GGA ATT G60 μl 30 μl 0.6 μl MgCl 2 (25mM)20 μl 10 μl 0.2 μl GoTaq DNA polymerase (5U/μl)5.0 ml 2.5 ml 50 μl Total (To each reaction add 1.0 µl of DNA)Table 7.3.2.1. F and R indicate forward and reverse orientation.PCR Cycle conditions94°C/5min x 1 cycle(94°C/30s , 50°C/30s , 72°C/1min) x 35 cycles72°C/10min x 1 cycle4°C holdUse a 1.5% Agarose ethidium bromide gel for visualization.Primers create an approximately 459 bp band (Figure 7.3.2.1).Figure 7.3.2.1. Lane 1 1Kb marker, lane 2 An. atroparvus, lane 3 An.gambiae, lane 4 An. arabiensis, lane 5 An. funestus, lane 6 An.pharoensis.

Chapter 7 : Taxonomy and Systematics7.3 Mitochondrial DNA PCR assays7.3.2 General PCR to amplify the cytochrome B region in anophlinesPage 2 of 296 well PCR sample preparation templateReferencesAvise JC, J Arnold, RM Ball, E Bermingham, T Lamb, JE Neigel, CA Reeb, and NC Saunders. 1987.Intraspecific phylogeography: the mitochondrial DNA bridge between population genetics andsystematics. Ann Rev Ecol Syst. 18: 489-522.Krzywinski, J, RC Wilkerson, and NJ Besansky. 2001. Evolution of mitochondrial and ribosomal genesequences in Anophelinae (Diptera: Culicidae): Implications for phylogeny reconstruction. Mol Phyl Evol.18 (3): 479-487.Simmons RB and SJ Weller. 2001. Utility and evolution of cytochrome b in insects. Mol Phyl Evol. 20 (2):196-210.

Chapter 8 : Field Techniques8.1 Molecular Identification of Plasmodium spp. in AnophelinesPage 1 of 4Chapter 8 : Field Techniques8.1 Molecular Identification of Plasmodium spp. in AnophelinesFrédéric Lardeux, Rosenka Tejerina, Claudia AliagaIntroductionThe identification of Plasmodium species in whole mosquitoes by PCR is difficult because of thepresence of reaction inhibitors from the insects (Schriefer et al. 1991). The present protocol is based on achelex extraction which overcomes the PCR inhibition phenomenon. The semi-nested multiplex PCRtechnique detects and distinguishes among the four human Plasmodium species in single mosquitoesand in pools of up to 100 mosquitoes. The extraction and PCR technique presented here can be usefulto: (1) estimate by mosquito pooling Plasmodium prevalence in Anopheles populations in low prevalenceareas where large numbers of individual mosquitoes would need to be processed to obtain a reliableestimate; (2) incriminate Anopheles species as malaria vectors; (3) identify at one time all the circulatingPlasmodium species in vectors from an area; (4) detect mixed infections in mosquitoes; and (5) detectmosquitoes with low-level parasite infections. An alternate, RT-PCR method can be found in Chapter8.5.1.2.DNA extraction (Lardeux et al. 2008)1. Homogenize mosquito heads + thoraces in saline solution (NaCl 0.9%).2. Add Chelex 100X (for volumes see Table 8.1.1) and vortex.3. Incubate at 100°C for 10 min.4. Centrifuge at 13 000 rpm for 5 min.5. Mix the supernatant with 1:1 vol. phenol-chloroform and centrifuge 3 times at 10,000 rpm for 5min.6. Mix the supernatant with 1:1 vol. 70% ethanol and centrifuge at 14,000 rpm for 20 min.7. Dry the pellet at 37°C8. Suspend the pellet in 100 µl sterile H 2 O (nuclease-free H 2 O).9. Keep at 4°C until PCR processed.Mosquito pool size Volume NaCl (µl)Concentration Chelex 100X Volume Chelex 100X(w/v)°(µl)1-10 50 5 % 24020 100 5 % 48030 150 5 % 75040 200 10 % 80050 250 10 % 80060 300 10 % 90070 350 10 % 90080 400 10 % 100090 450 10 % 1000100 500 10 % 1000Table 8.1.1. Concentration of Chelex 100X (%), volumes (µl) of chelex 100X and of saline solutionused in the preparation of the DNA template in accordance with the size of the pool of mosquitoes(number of mosquitoes processed)

Chapter 8 : Field Techniques8.1 Molecular Identification of Plasmodium spp. in AnophelinesPage 2 of 4Semi-nested multiplex PCR for Human Plasmodium species identification (Lardeux et al. 2008)Prepare PCR Master Mix for 96, 48 or 1 20 μl PCR reactions 1 . Add reagents in the order presented.96 48 1 Reagent259.2 µl 129.6 µl 2.7 µl Nuclease free H 2 O384.0 µl 192.0 µl 4.0 µl Taq 1X PCR Buffer with MgCl 296.0 µl 48.0 µl 1.0 µl dNTP’s (0.5 mM mix)96.0 µl 48.0 µl 1.0 µl Universal reverse. UNR (R, 0.05 µM) GAC GGT ATC TGA TCG TCT TC96.0 µl 48.0 µl 1.0 µl Plasmodium. PLF (F, 0.05 µM) AGT GTG TAT CAA TCG AGT TTC28.8 µl 14.4 µl 0.3 µl Go Taq DNA Polymerase ( 5U/µl)960 µl 480 µl 10 µl Total (to each 10µl reaction add 10 µl template DNA)Table 8.1.2. F and R indicate forward and reverse orientation.Prepare PCR 2 Master Mix for 96, 48 or 1, 20 µl PCR reactions; Add reagents in the order presented96 48 1 Reagent912.0 µl 456.0 µl 9.5 µl Nuclease free H 2 O384.0 µl 192.0 µl 4.0 µl Taq 1X PCR Buffer with MgCl 296.0 µl 48.0 µl 1.0 µl dNTP’s (0.5 mM mix)153.6 µl 76.8 µl 1.6 µl PLF (F, 0.08 µM) AGT GTG TAT CAA TCG AGT TTC38.4 µl 19.2 µl 0.4 µl P. falciparum. FAR (R, 0.04 µM) AGT TCC CCT AGA ATA GTT ACA38.4 µl 19.2 µl 0.4 µl P. vivax. VIR (R, 0.04 µM) AGG ACT TCC AAG CCG AAG C38.4 µl 19.2 µl 0.4 µl P. malariae. MAR (R, 0.04 µM) GCC CTC CAA TTG CCT TCT G38.4 µl 19.2 µl 0.4 µl P. ovale. OVR (R, 0.04 µM) GCA TAA GGA ATG CAA AGA ACA G28.8 µl 14.4 µl 0.3 µl Go Taq DNA Polymerase ( 5U/µl)1.73 ml 864 µl 18 µl Total (to each 18 µl reaction add 2 µl of PCR1 amplicon)Table 8.1.3. F and R indicate forward and reverse orientation.PCR cycle conditionsPCR 1 : UNR-PLF94°C/ 5 min x 1 cycle(94°C/ 1 min, 60°C/ 1 min, 72°C/ 90 sec) x 40 cycles4°C holdPCR 2: PLF-MAR, FAR, VIR, OVR94°C/ 5 min x 1 cycle(94°C/ 30 sec, 62°C/ 30 sec, 72°C/ 60 sec) x 35 cycles72°C/ 10 min x 1 cycle4°C holdRun samples on a 1.5 % agarose EtBr gel for visualization.If needed, positive samples from agarose can also be run on an 8% polyacrylamide gel, staining with0.2% silver nitrate and revealed with a 2:1 volume of 30 g / l sodium carbonate: 0.02% formaldehyde.Primers create fragments of 269 bp (P. malariae) , 395 bp (P. falciparum), 436 bp (P. ovale), 499 bp (P.vivax)1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.1 Molecular Identification of Plasmodium spp. in AnophelinesPage 3 of 4ReferencesLardeux F, Tejerina R, Aliaga C, Ursic-Bedoya R, Lowenberger C, Chavez T (2008) Optimization of asemi-nested multiplex PCR to identify Plasmodium parasites in wild-caught Anopheles in Bolivia, and itsapplication to field epidemiological studies. Trans R Soc Trop Med Hyg 102:485-492Schriefer ME, Sacci JB, Wirtz RA, Azad AF (1991) Detection of Polymerase Chain Reaction-AmplifiedMalarial DNA in Infected Blood and Individual Mosquitoes. Experimental Parasitology 73:311-31696 well sample preparation template

Chapter 8 : Field Techniques8.1 Molecular Identification of Plasmodium spp. in AnophelinesPage 4 of 4

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 1 of 128.2 Plasmodium Sporozoite ELISARobert Wirtz, Melissa Avery, Mark Benedict, Alice SutcliffeIntroductionEnzyme-linked immunosorbent assays (ELISAs) were developed to detect Plasmodium falciparum, P.vivax-210, and P. vivax-247 circumsporozoite (CS) proteins in malaria-infected mosquitoes. Thesensitivity and specificity of the ELISAs are based on the monoclonal antibodies (Mabs) used. TheELISAs detect CS proteins, which can be present in the developing oocysts, dissolved in haemolymph,and on sporozoites present in the haemocoel or in the salivary glands. A positive ELISA on a mosquitodoes not establish that species as a vector, and ELISA results may not be synonymous with salivarygland sporozoite dissections.ELISAs can be carried out on fresh, frozen, or dried mosquitoes. If specimens are to be dried, they mustbe processed quickly and kept dry (stored with desiccant) to prevent microbial growth that can result inhigh background values. Before collection of the mosquitoes is initiated, consideration should be given tothe possibility of conducting other tests (e.g., molecular, host blood meal, etc.) that may require differentstorage conditions or extraction buffers. Voucher specimens should also be collected and saved.The "sandwich" ELISA is begun by adsorption of the capture Mab to the wells of a microtiter plate (Figure3.3.1). After the capture Mab has bound to the plate, the well contents are aspirated and the remainingbinding sites are blocked with blocking buffer. Mosquitoes to be tested are ground in blocking buffercontaining IGEPAL CA-630, and an aliquot is tested. Positive and negative controls are also added tospecific plate wells at this time. If CS antigen is present (depicted as diamond in Fig. 1.B) it will form anantigen-antibody complex with the capture mAb. After a 2-hour incubation at room temperature, themosquito homogenate is aspirated and the wells are washed. Peroxidase-linked Mab is then added to thewells, completing the formation of the "sandwich" (Fig. 1.C). After 1 hour, the well contents are aspirated,the plate is washed again and the clear peroxidase substrate solution is added (Fig 1.D). As theperoxidase enzyme reacts with the substrate, a dark green product is formed (Fig 1.D), the intensity ofthe color is proportional to the amount of CS antigen present in the test sample.Results are read visually or at 405-414 nm using an ELISA plate reader 30 and/or 60 minutes after thesubstrate has been added.The assay involves 2 steps:1. Screening phase where the ELISA is used to identify positive samples (Worksheet 1).2. Quantification ELISA - where the ELISA positive samples from the initial screening are retestedto: a) confirm positives and b) quantify the amount of CS protein per sample (Worksheet 2).If it is necessary to test for all three Plasmodium species (Pf, Pv210 and Pv247) these tests can be runconcurrently on 3 plates or if there is not enough material, the 3 assays may be run consecutively.For technical advice or recommendations regarding the use of this protocol, please contact:Robert A. Wirtz, Ph.D., Chief, Entomology Branch, MS F-42, Phone: 770-488-4240, Fax: 770-488-4258Centers for Disease Control and Prevention, 4770 Buford Highway, NE, Atlanta, GA 30341-3724 USAE-mail: rwirtz@cdc.govAcknowledgmentsDevelopment of these assays was a cooperative effort among the National Academy of Sciences,National Institutes of Health, Centers for Disease Control and Prevention, Naval Medical ResearchInstitute, New York University, Walter Reed Army Institute of Research, Protein Potential and the WorldHealth Organization. These instructions were developed by Robert Wirtz, Melissa Avery and MarkBenedict.

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 2 of 12Figure 8.2.1. The “sandwich” ELISA for detection of Plasmodium falciparum and P. vivaxcircumsporozoite proteins.A. Anti-sporozoite monoclonal antibodyadsorbed to the plateB. Blocking buffer added to prevent nonspecificbindingC. Mosquito triturate added to the well. D. Peroxidase-linked anti-sporozoitemonoclonal antibody added.E. ABTS substrate added to the wells..

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 3 of 12Sporozoite ELISA Solutions, Capture and ConjugatePHOSPHATE BUFFERED SALINE (PBS), pH 7.2: Use stock laboratory PBS OR Dulbecco's PBS(Sigma #D5773); adjust pH if necessary. Add 0.01 gm phenol red or 100l of phenol red stock solution(1gm/10ml water)/L PBS. Store 4°C; shelf life is 2 weeks.BLOCKING BUFFER (BB): Shelf life is 1 week at 4°C; BB may be frozen. Use only ELISA grade Sigmacasein.BB – blocking buffer: Use only ELISA grade casein (Sigma C7078). Shelf life is 1 week at 4°C;BB may be frozen.Boiled casein (BB): ½ liter 1 literPBS, pH 7.4 450 ml 900 mlcasein 2.5 g 5.0 g0.1 N NaOH 50 ml 100 mlphenol red* 100 μl 200 μl1. Bring to a boil the 0.1 N NaOH and slowly add the casein (Sigma C7078).2. After casein is dissolved, allow to cool and slowly add the PBS, adjust the pH to ~7.4 with HCl, andadd the phenol red.*A stock solution of phenol red (1g/10ml water) eliminates the need to weigh out small amounts of thismaterial. Phenol red added to BB is only used as a dye to aid in visualization and is optional.Grinding Buffer and Mosquito Sample Preparation: Shelf life at 4°C is 1 week.Grinding solution for approximately one plate:1. Combine 25ml of BB and 125µl of Igepal CA-6302. Mix well to dissolve the Igepal CA-630 in the BB.(Note: IGEPAL CA-630 (Sigma I3021) replaces NONIDET P-40, no longer available from SIGMA-ALDRICH. If available, NONIDET P-40 can be used.)To Grind:1. Place the mosquito, head-thorax only, of no more than 10 pooled mosquitoes in a labeled1.5ml micro centrifuge grinding tube2. Add 50µl of grinding buffer.3. Grind well4. Rinse the pestle with two 100µl volumes of Grinding Solution , catching the rinses in the tubecontaining the mosquito triturate. Final volume will be 250µl. Test or freeze for later use.5. Rinse the pestle in PBS-Tween twice; dry with tissue to prevent contamination betweenmosquitoes.Wash Solution (PBS-Tween): PBS plus 0.05% Tween 20. Add 0.5 ml Tween 20 to 1 liter of PBS. MIXWELL. Store at 4°C. Shelf life is 2 weeks.Monocolonal Antibodies (mAb) Capture and Conjugate: Monoclonal antibodies (mAb) capture andconjugate received from KPL will be lyophilized. Label will list the amount of glycerol water to be added.Glycerol water is a 1:1 mixture of distilled water and glycerol. Glycerol water allows for storage at -20°Cwithout freeze-thawing. This step only needs to be performed when a new vial of capture or conjugateneeds to be reconstituted.

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 4 of 12Notes and Troubleshooting for Sporozoite ELISAs1. Do not add sodium azide to solutions as it is a peroxidase inhibitor. We no longer add thimerosal tothe solutions, as this is mercury-based and presents problems with proper disposal.2. To fill each of the 96 wells on a plate with 50μl requires 4.8ml. It is convenient to make up 5.0ml ofeach mAb solution and 10.0ml of substrate (100µl/well) per plate.Species mAb g/50μl/WELL g/5ml l STOCK/5ml PBSPf capture 0.200 20.0 40Pv - 210 capture 0.025 2.5 5Pv - 247 capture 0.025 2.5 5Pf peroxidase 0.050 5.0 10Pv - 210 peroxidase 0.050 5.0 10Pv - 247 peroxidase 0.050 5.0 10l STOCK/5ml BB3. Cover plate during incubations to prevent evaporation. Incubate plate in the dark (especially forincubation of peroxidase conjugate and substrate solution). An efficient way of doing this is to use asmall cardboard box lid or to line the lid of a pipette tip box with aluminum foil.4. ELISAs can be carried out on fresh, frozen or dried mosquitoes. If specimens are to be dried, theymust be processed quickly and kept dry (store with desiccant) to prevent microbial growth that canresult in high background values in the ELISAs. Once identified to species, mosquitoes should bepooled in groups of maximum 10 and stored at -20°C or dried over silica gel until they can beprocessed. Store the triturate at -20°C until tests are to be performed.5. Negative controls: Triturate laboratory reared, known uninfected female mosquitoes (same as testspecies if possible) in 50μl BB:IG-630, dilute with 150μl BB (total volume 200μl) and place 50μl fromeach into negative control wells 1B-1H for initial testing or wells 1A-1H for conformational testing. Todetermine the negative cut off value for initial testing calculate mean on these 7 negative controls andretest any mosquito with a value two times the mean.6. Phenol red added to BB is only used as a dye to aid in visualization and is optional.7. Only high, lab grade paper towels should be used while performing CS-ELISA testing, particularly to“bang” the ELISA plate on, to remove excess solutions. Use of brown paper towels and kitchenpaper towels should be avoided as the result is high background values. It is thought that fibers fromthese towels adhere to the plates and interfere with detection. If high, lab grade paper towels are notavailable, perform this step over the sink or ensure careful removal of all solutions with a vacuumsystem for each required step.8. Do not vortex samples as this can lead to high background and inaccurate absorbance values. Shortcentrifugation can but used to settle body parts and allow pipetting of a clean sample. Thiscentrifugation will not pellet sporozoites and they will remain suspended in the supernantant.9. Wipe the bottom of the ELISA plate with alcohol to remove debris and oils from ELISA plate beforereading. Fingerprints, oils and debris on the plate can lead to inaccurate absorbance values.10. BB should be placed in wells without negative mosquito controls, positive controls, or samples as“filler”.11. The Corning® 96 well clear round bottom PVS (soft plate) cannot be reliably used with certain,especially newer, models of plate readers. This is due to distortion of the plate upon entering thereader carriage or a poor fit of plate in the reader carriage. This can be solved by using Costar 96-Well EIA/RIA plates (Round well; high binding), available from www.fishersci.com see supplies list).

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 5 of 12Dilutions for Positive Control Antigens for CS-ELISASerial dilutions are easily prepared by one of the two following methods:1. All working concentrations required can be prepared and labeled ahead of time. When addition ofpositive controls to wells of ELISA plate is required, 50l of each solution is added. This method issuited for inexperienced pipetters.Label microfuge tubes with solution number and prepare sequentially according to chart. Remainingvolumes can be stored at -20ºC and used again when necessary.Solution Volume of Positive Control Volume of BlockingNumber AntigenBuffer (l)PlasmodiumspeciesAntigenConcentration(pg/50l)P. falciparum Stock Lyophilized Pf 1000 500,000I 20l of stock 1000 5,000*II 10l of I 500 100*III 500l of II 500 50*IV 500l of III 500 25*V 500l of IV 500 12*VI 500l of V 500 6*VII 500l of VI 500 3*VIII 500l of VII 500 1.5*IX 0 (Blank) 500 0 (Blank)P. vivax 210 Stock Lyophilized Pv210 500 500,000A 10l of stock 1000 5,000B 20l of A 500 200*C 200l of B 800 40*D 500l of C 500 20*E 500l of D 500 10*F 500l of E 500 5*G 500l of F 500 2.5*H 500l of G 500 1.25* I 500l of H 500 0.6*J 0 (Blank) 500 0P. vivax 247 Stock Lyophilized Pf 1000 227,500*A 20l of stock 1000 4,450*B 500l of I 500 2,275*C 500l of II 500 1,140*D 500l of III 500 570*E 500l of IV 500 285*F 500l of V 500 140*G 500l of VI 500 70*H 0 (Blank) 500 0 (Blank)* Amounts used in ELISA plate wells for the quantitative assays

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 6 of 122. The highest concentration to be used on ELISA plate is prepared ahead of time. When addition ofpositive controls to wells of ELISA plate is required, serial dilutions are made directly on the plate.This requires very accurate and advanced pipetting skills.Positive controls (be sure to label all vials):P. falciparum:STOCK: Add 1000l BB to lyophilized positive control (5g) (do not use glycerol:water or wateralone) to give 10,000pg/l BB.Vial I = Transfer 20l (200,000pg) of STOCK to 1000l BB to for 100pg/l BB.Vial II = Transfer 10l (1000pg) of Vial I to 500l BB for 2pg/l or 100 pg/50l BBBegin in well 2A as wells 1A-1H should be reserved for 8 negative control mosquitoes whenretesting. Freeze the stock solution and Vials I and II for continued use.Use 100µl Vial II in well 2A and add 50l BB to wells 2B through 2H. Pipette 50l from well 2Aand add to well 2B. Continue this serial dilution through well 2H and discard 50l from well 2H soeach well contains 50l of BB + diluted positive control.Concentrations of positive controls are 100, 50, 25, 12, 6, 3, 1.5 and 0pg/50l BB (starting with2A and finishing with 2H)Run standard curve in triplicate (wells 2A-4A to 2H-4H) as well as 8 negative control mosquitoes(wells 1A-1H) when retesting.P. vivax-210:STOCK = Add 500l BB to lyophilized positive control (5g) (do not use glycerol:water or wateralone) to give 10,000pg/l BB.Vial A = Transfer 10l (100,000pg) of STOCK to 1,000l BB for 100pg/l BB.Vial B = Transfer 20l (2,000pg) of Vial A to 500l BB for 4pg/l BB.Vial C = Transfer 200l (800pg) of Vial B to 800l BB for 0.4pg/l or 40 pg/50l BB.Begin in well 2A as wells 1A-1H should be reserved for 8 negative control mosquitoes whenretesting. Freeze the stock solution and Vials I and II for continued use.Use 100µl Vial C in well 2A and add 50l BB to wells 2B through 2H. Pipette 50l from well 2Aand add to well 2B. Continue this serial dilution through well 2H and discard 50l from well 2H soeach well contains 50l of BB + diluted positive control.Concentrations of positive controls are 40, 20, 10, 5, 2.5, 1.25, 0.6 and 0pg/50l BB (starting with2A and finishing with 2H)Run standard curve in triplicate (wells 2A-4A to 2H-4H) as well as 8 negative control mosquitoes(wells 1A-1H) when retesting.P. vivax-247:STOCK = Add 1000 µl BB to lyophilized positive control (4.55ug) (do not use glycerol:water orwater alone) to give 4,550pg/l BB .Vial 1 = Transfer 20l (91,000pg) of STOCK to 1,000l BB to give 4,550pg/l or BB.Begin in well 2A as wells 1A-1H should be reserved for 8 negative control mosquitoes whenretesting. Freeze the stock solution and Vials I and II for continued use.Use 100µl Vial 2 in well 2A and add 50l BB to wells 2B through 2H. Pipette 50l from well 2Aand add to well 2B. Continue this serial dilution through well 2H and discard 50l from well 2H soeach well contains 50l of BB + diluted positive control.Concentrations of positive controls are 4450, 2275, 1140, 570, 285, 140, 70 and 0pg/50l BB(starting with 2A and finishing with 2H)Run standard curve in triplicate (wells 2A-4A to 2H-4H) as well as 8 negative control mosquitoes(wells 1A-1H) when retesting.

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 7 of 12Sporozoite ELISA DirectionsPlease be sure to read the notes preceding and following these directions before beginning.1. Fill out the top portion of the appropriate Sporozoite ELISA worksheet. Mark the ELISA plate in orderto maintain correct plate orientation.2. Prepare a working solution of mAb capture by adding PBS to the reconstituted capture mAb based onthe volumes by species listed below. Vortex gently.Species mAb g/50l/WELL g/5ml l STOCK/5mlPf capture 0.200g/50l 20.0g 40l stock+ 5ml PBSPv-210 capture 0.025g/50l 2.5g 5l stock+ 5ml PBSPv-247 capture 0.025g/50l 2.5g 5l stock+ 5ml PBS3. Place 50l of mAb solution made in step 2 in each well of the ELISA plate. Use a separate plate foreach sporozoite species.4. Cover plate and incubate for 0.5 HOUR room temperature.0.5 HOUR5. Aspirate well contents and bang plate upside down on paper towel 5 times, holding sides only. Note:If aspiration system is not available, bang plate on sink edge into the sink and then again on papertowels. Do not use brown paper towels.6. Fill wells with 200l BB7. Cover plate, leaving space between well and top of lid. Incubate for 1 hour at room temperature.1 HOUR8. Aspirate well contents and bang plate upside down on paper towel 5 times holding sides only.9. Load samples and controls into the plate (initial and quantitative tests).-Use 9a when first testing the samples to determine if there are positives-Use 9b when retesting, confirming and quantifying any positives from the initial/first test.9a. Initial test see Worksheet 1 (plate #1, typically Day 1):i. Add 50l of positive controls to well A1. See the notes on 6 and 7 for positive controldilutions. (Pf = Vial II, Pv210= Vial C, Pv247= Vial 2)ii. Add 50l of negative control(s) to wells B1-H1.iii. Add 50l of mosquito triturate per well to remaining wells.iv. Cover and incubate 2 hoursb. Quantification test see worksheet 2 (plate #2, typically Day 2):i. Add 50l of negative control(s) to wells A1-H1.ii. Add 50l last vial dilution of positive control to wells A2, A3, and A4 and 50l of eachserial dilution to wells B2-H2, B3-H3 and B4-H4. See pages 7 and 8 for positive controldilutions. (Pf = Vial II, Pv210= Vial C, Pv247= Vial 2)iii. Add 50ul of mosquito triturate per well to remaining wells.iv. Cover and incubate 2 hoursPlease note, steps 10-12 can be performed just before the end of the 2 hour incubation2 HOURS

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 8 of 1210. Prepare substrate by mixing Substrate A and Substrate B at a 1:1 ratio. A full 96-well plate willbe 5ml of Substrate A + 5ml of Substrate B11. Prepare a working solution of mAb conjugate by adding BB to the reconstituted capture mAbbased on the volumes by species listed below. Vortex gently.Species mAb ug/50 l/WELL g/5ml l STOCK/5mlPf peroxidase 0.050g/50l 5.0g 10l stock + 5ml BBPv-210 peroxidase 0.050g/50l 5.0g 10l stock + 5ml BBPv-247 peroxidase 0.050g/50l 5.0g 10l stock + 5ml BB12. Check enzyme activity by mixing 5l of the mAb conjugate made in step 11 with 100l of thesubstrate made in step 10 in a separate tube. Vortex gently. There should be rapid color changeindicating that the peroxidase enzyme and the substrate are functional.13. Aspirate well contents bang plate upside down on paper towel 5 times holding sides only.14. Wash wells two (2) times with 200l of PBS-Tween, aspirating and banging plate 5 times witheach wash.15. Add 50l of peroxidase conjugate solution made in step 11 to each well.16. Cover and incubate one (1) hour1 HOUR17. Aspirate well contents and bang plate upside down on paper towel 5 times holding sides only.18. Wash wells 3 times with 200l of PBS-Tween, aspirating and banging plate 5 times with eachwash.19. Add 100l substrate solution per well.20. Cover plate and incubate thirty (30) minutes. Handle plate carefully to avoid splashing.21. Read visually, or at 405-414nm0.5 HOUR22. Cover plate and incubate an additional thirty (30) minutes if testing Pv-210 or Pv-247 species,for a total incubation time of 1 hour.23. Read visually, or at 405-414nm0.5 HOUR

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 9 of 12WORKSHEET 1: ELISA Template for Screening (Initial test)ELISA Plate No:______________Capture mAb: Lot #____________DATE: _______________Peroxidase-mAb: Lot #:________________Positive control: Lot#______________NEG1 2 3 4 5 6 7 8 9 10 11 12ABCDEFGHPosNegNegNegNegNegNegNeg____ 1) Coat PVC plate with 50l capture mAb. 0.5 hr incubation____ 2) Aspirate wells; fill wells with 200l blocking buffer (BB). 1 hr incubation____ 3) Aspirate wells; Add 50l mosquito triturate and positive control 2 hr incubation____ 4) Aspirate and wash two times with 200l PBS-0.05% Tween 20.____ 5) Add 50l peroxidase-mAb. Mix substrate (1:1). 1 hr incubation (in the dark)____ 6) Aspirate and wash 3 times with 200l PBS-0.05% Tween 20.____ 7) Add 100l substrate: ____ a) Enzyme check; ____ b) 100l/well. 0.5 hr incubation (in the dark)____ 8) Read absorbency 405nm.Analysis:Samples which have OD values above the cut-off (cut-off = 2 X mean OD of negative samples)are considered positive and should be followed up with quantitative testing.

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 10 of 12WORKSHEET 2: ELISA Template for Quantitative Testing (Quantification Test)ELISA Plate No:______________Capture mAb: Lot #____________DATE: _______________Peroxidase-mAb: Lot #:________________Positive control: Lot#______________NEG POS. CONTROL1 2 3 4 5 6 7 8 9 10 11 12ABCDEFGH____ 1) Coat PVC plate with 50l capture mAb. 0.5 hr incubation____ 2) Aspirate wells; fill wells with 200l blocking buffer (BB). 1 hr incubation____ 3) Aspirate wells; Add 50l mosquito triturate and positive control 2 hr incubation____ 4) Aspirate and wash 2 times with 200l PBS-0.05% Tween 20.____ 5) Add 50l peroxidase-mAb. Mix substrate (1:1). 1 hr incubation (in the dark)____ 6) Aspirate and wash 3 times with lPBS - 0.05% Tween 20.____ 7) Add 100l substrate: ____ a) Enzyme check; ____ b) 100 μl/well. 0.5 hr incubation (in the dark)____ 8) Read absorbency 405nm.Analysis: Compare to standard curve from positive control antigen stocks to estimate CS antigen in eachsample. From this estimate, calculate the equivalent number of sporozoites for that sample by chartcomparison.

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 11 of 12ReferencesBurkot TR, Williams JL, Schneider I (1984) Identification of Plasmodium falciparum-infected mosquitoesby a double antibody enzyme-linked immunosorbent assay. Am J Trop Med Hyg 33:783-788Collins FH et al. (1984) First field trial of an immunoradiometric assay for the detection of malariasporozoites in mosquitoes. Am J Trop Med Hyg 33:538-543Rosenberg R et al. (1989) Circumsporozoite protein heterogeneity in the human malaria parasitePlasmodium vivax. Science 245:973-976Wirtz RA, Burkot TR, Andre RG, Rosenberg R, Collins WE, Roberts DR (1985) Identification ofPlasmodium vivax sporozoites in mosquitoes using an enzyme-linked immunosorbent assay. Am J TropMed Hyg 34:1048-1054Wirtz RA, Sattabongkot J, Hall T, Burkot TR, Rosenberg R (1992) Development and evaluation of anenzyme-linked immunosorbent assay for Plasmodium vivax-VK247 sporozoites. J Med Entomol 29:854-857Wirtz RA et al. (1987) Comparative testing of monoclonal antibodies against Plasmodium falciparumsporozoites for ELISA development. Bull World Health Organ 65:39-45

Chapter 8 : Field Techniques8.2 Plasmodium falciparum Sporozoite ELISAPage 12 of 12

Chapter 8 : Field Techniques8.3 Molecular Identification of Mammalian Blood Meals from MosquitoesPage 1 of 48.3 Molecular Identification of Mammalian Blood Meals fromMosquitoesChristen M Fornadel, Rebekah J Kent, Douglas E NorrisIntroductionIdentification of blood meals is an important step in understanding mosquito ecology. The followingprotocol was developed to differentiate between a select group of potential mammal host bloods inengorged anophelines, but may be adapted or expanded to suit particular needs (i.e. most often anexpanded or altered host list). This protocol was designed for use on genomic DNA extractions ofmosquito abdomens.Initial Blood Meal Identification PCR (Kent and Norris 2005; Kent et al. 2007)This multiplexed PCR produces species-specific fragments of varying sizes amplified from thecytochrome b gene, encoded in the mitochondrial genome. Host DNA is detectable up to 24-30 hourspost feeding.Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent Product size (bp)1.83 ml 915 μl 18.3 μl sterile H2O250 μl 125 μl 2.5 μl Taq 10X PCR Buffer with MgCl2100 μl 50 μl 1.0 μl dNTP (final concentration of 100 μM of each dNTP)50 μl 25 μl 0.5 μl UnRev1025 (50 pmol/μl) [ggttg[t/g]cctccaattcatgtta]50 μl 25 μl 0.5 μl Pig573F (50 pmol/μl) [cctcgcagccgtacatctc] 45350 μl 25 μl 0.5 μl Human741F (50 pmol/μl) [ggcttacttctcttcattctctcct] 33450 μl 25 μl 0.5 μl Goat894F (50 pmol/μl) [cctaatcttagtacttgtacccttcctc] 13250 μl 25 μl 0.5 μl Dog368F (50 pmol/μl) [ggaattgtactattattcgcaaccat] 68050 μl 25 μl 0.5 μl Cow121F (50 pmol/μl) [catcggcacaaatttagtcg] 56120 μl 10 μl 0.2 μl Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 25 μl TotalFor DNA template use up to 3 µl DNA sample (from abdomen extraction eluted in 50μl dH 2 0)PCR Cycle conditions95°C/5min x 1 cycle(95°C/60sec , 56°C/60sec , 72°C/60sec) x 40 cycles72°C/7min x 1 cycle4°C hold1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 reactionsto compensate for imprecise measurements.

Chapter 8 : Field Techniques8.3 Molecular Identification of Mammalian Blood Meals from MosquitoesPage 2 of 4Small Blood Meal PCR and Enzyme Digest (Fornadel and Norris 2008)For samples that fail to produce a reaction product with the multiplexed PCR the following small bloodmeal PCR/enzyme digest may be used. This assay allows identification of host source from partiallydigested blood meals (out to 60 hours post feeding), as well as from partially degraded DNA extractions.The PCR was designed to produce a small 98bp amplicon from the mammals tested above, which canthen be incubated with specific restriction enzymes to determine host source.Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions 1 . Add reagents in the order presented.96 48 1 Reagent size (bp)1.83 ml 915 μl 18.3 μl sterile H2O250 μl 125 μl 2.5 μl Taq 10X PCR Buffer with MgCl2100 μl 50 μl 1.0 μl dNTP (final concentration of 100 μM of each dNTP)50 μl 25 μl 0.5 μl UnRev1025 (50 pmol/μl) [ggttgtcctccaattcatgtta]50 μl 25 μl 0.5 μl UniForA (50 pmol/μl) [tccaaacaac[a/g][a/c]agcataatatt] 9820 μl 10 μl 0.2 μl Taq DNA polymerase (5 U/μl)2.3 ml 1.15 ml 23 μl TotalFor DNA template use 2 µl DNA sample (from abdomen extraction eluted in 50μl dH 2 0)Save 6 µl of the PCR products to run as undigested controls on a 3% agarose gel alongside the digestedamplicons.Note: The master mix can be tripled for a total reaction volume of 78 µl. This will allow one to perform upto 4 digests of the amplified product with enough undigested sample leftover as a control.PCR Cycle conditions95°C/5min x 1 cycle(95°C/60sec , 55°C/60sec , 72°C/60sec) x 40 cycles72°C/7min x 1 cycle4°C holdRestriction Enzyme DigestsThe following digests can be performed in whichever order/combination provides the best efficiency forthe samples under study.Host Cut position Enzyme Recognition Seq.Human 59 Fnu4HI GC^N_GCCow 54 BanII G_RGCY^CDog 24 MspI C^CG_GGoat 43 NsiI A_TGCA^TPig 54 SpeI A^CTAG_TRestriction Digest ReactionPer reaction (1)Reagent7.5 µl sterile H2O2.5 µl Taq 10X PCR Buffer with MgCl21-2 U Enzyme15 µl PCR product.025 µl only for SpeI digests BSA 100X~25 µl TotalAll digests are carried out at 37C for at least 3hrs but can be left overnight.

Chapter 8 : Field Techniques8.3 Molecular Identification of Mammalian Blood Meals from MosquitoesPage 3 of 496 well PCR sample preparation templateReferencesFornadel CM, Norris DE (2008) Increased endophily by the malaria vector Anopheles arabiensis insouthern Zambia and identification of digested blood meals. Am J Trop Med Hyg 79:876-880Kent RJ, Norris DE (2005) Identification of mammalian blood meals in mosquitoes by a multiplexedpolymerase chain reaction targeting cytochrome B. Am J Trop Med Hyg 73:336-342Kent RJ, Thuma PE, Mharakurwa S, Norris DE (2007) Seasonality, blood feeding behavior, andtransmission of Plasmodium falciparum by Anopheles arabiensis after an extended drought in southernZambia. Am J Trop Med Hyg 76:267-274

Chapter 8 : Field Techniques8.3 Molecular Identification of Mammalian Blood Meals from MosquitoesPage 4 of 4

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.1 Anopheles gambiae Complex – Scott et al.Page 1 of 28.4 Species Complex Authentication by PCR8.4.1 Anopheles gambiae complex (Scott et al.)MR4 StaffIntroductionThe Anopheles gambiae complex is comprised of 7 cryptic species: An. gambiae s.s, An. arabiensis, An.bwambe, An. melas, An. merus, An. quadriannulatus A and B, some of which are sympatric.Distinguishing the members of the complex was first done based on karyotype and chromosomalinversions (Coluzzi et al. 1979). Recently, however, easier PCR based assays have been developed thatdistinguished several of the members based on species-specific single nucleotide polymorphisms (SNPs)in the intergenic spacer region (IGS) (Scott et al. 1993; Fettene and Temu 2003; Besansky et al. 2006;Wilkins et al. 2006). A good overview and the fine points of this assay has been published (Cornel andCollins 1996). Alternatives to this assay can be found in Chapters 8.4.3 and 8.5.1.1.PCR authentication for the members of the Anopheles gambiae complex (Scott et al. 1993)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent835 μl 417.5 μl 8.35 μl sterile H 2 O500 μl 250 μl 5.0 μl GoTaq 5X PCR Buffer with MgCl 2250 μl 125 μl 2.5 μl dNTP (2.5 mM mix G,A,T,C)100 μl 50 μl 1.0 μl MgCl 2 (25 mM)100 μl 50 μl 1.0 μl UN (F, 25 pmol/μl) [GTGTGCCCCTTCCTCGATGT]100 μl 50 μl 1.0 μl AR (R, 25 pmol/μl) [AAGTGTCCTTCTCCATCCTA]100 μl 50 μl 1.0 μl GA (R, 25 pmol/μl) [CTGGTTTGGTCGGCACGTTT]100 μl 50 μl 1.0 μl ME (R, 25 pmol/μl) [TGACCAACCCACTCCCTTGA]200 μl 100 μl 2.0 μl QD (R, 25 pmol/μl) [CAGACCAAGATGGTTAGTAT] OR200 μl 100 μl 2.0 μl QDA (R, 25 pmol/µl) [CATAATGAGTGCACAGCATA] )15 μl 7.5 μl 0.15 μl Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 24 μl Total (To each 24 ul reaction add 1 μl template DNA)Table 8.4.1.1. F and R indicate forward and reverse orientation. It may improve specificity to leave outprimers for species that do not occur in the area of sample collection. If removing a primer, replace primervolume with an equal volume of sterile water. Use the QDA primer instead of the QD primer in areaswhere An. merus and An. quadriannulatus are sympatric. Use 1 μl DNA template.PCR cycle conditions95°C/5min x 1 cycle(95°C/30sec , 50°C/30sec , 72°C/30sec) x 30 cycles72°C/5min x 1 cycle4°C holdRun samples on a 2% agarose EtBr gel; load 5 μl sample (Figure 8.4.1.1).1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.1 Anopheles gambiae Complex – Scott et al.Page 2 of 2Primers create fragments of 153bp (QD) or 415bp (QDA) An. quadriannulatus, 464bp/466bp An.melas/merus, 390bp An. gambiae, 315bp An. arabiensis.ReferencesBesansky NJ, Collins FH, Townson H (2006) A species-specific PCR for theidentification of the malaria vector Anopheles bwambae. Ann Trop MedParasitol 100:277-280Coluzzi M, Sabatini A, Petrarca V, Di Deco MA (1979) Chromosomaldifferentiation and adaptation to human environments in the Anophelesgambiae complex. Trans Roy Soc Trop Med Hyg 73:483-497Cornel AJ, Collins FH (1996) PCR of the ribosomal DNA intergenic spacerregions as a method for identifying mosquitoes in the Anopheles gambiaecomplex. In: Clapp JP (ed) Methods in Molecular Biology: Species DiagnosticProtocols: PCR and other nucleic acid methods. Humana Press, Totowa NJ, pp321-332Fettene M, Temu EA (2003) Species-specific primer for identification ofAnopheles quadriannulatus sp. B (Diptera: Culicidae) from Ethiopia using amultiplex polymerase chain reaction assay. J. Med. Entomol. 40:112-115Scott JA, Brogdon WG, Collins FH (1993) Identification of single specimens ofthe Anopheles gambiae complex by the polymerase chain reaction. Am J TropMed Hyg 49:520-529Wilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect singlenucleotide polymorphisms for Anopheles gambiae species identification, Moptiand Savanna rDNA types, and resistance to dieldrin in Anopheles arabiensis.Malar J 5:125Figure 8.4.1.1. An.gambiae species ID.Lane 1, An. gambiae,Lane 2, An.arabiensis, Lane 3, 1kb ladder.96 well PCR sample preparation template

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.2 An. gambiae Ribosomal DNA Type – Fanello et al.Page 1 of 28.4.2 An. gambiae Ribosomal DNA Type (Fanello et al.)MR4 StaffIntroductionThe Fanello et al. (2002) method for differentiating between the rDNA types of An. gambiae uses arestriction enzyme, Hha I, to digest the PCR product from the Scott et al. assay (1993) discussed inAnopheles gambiae complex authentication section. The HhaI enzyme will specifically digest theSavanna form leaving 2 discernable bands while the Mopti form will only have 1 discernable band. Asmall 23 bp band is also created, but it is not seen on standard agarose gels. An alternate PCR basedmethod is presented in Chapter 8.4.3.PCR discrimination of Mopti & Savanna members of the Anopheles gambiae complex (Fanello etal. 2002)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent1235 μl 617.5 μl 12.35 μl sterile H 2 O500 μl 250 μl 5.0 μl GoTaq 5X PCR Buffer250 μl 125 μl 2.5 μl dNTP (2.5 mM mix)100 μl 50 μl 1.0 μl UN (F, 25 pmol/μl) [GTGTGCCCCTTCCTCGATGT]100 μl 50 μl 1.0 μl GA (R, 25 pmol/μl) [CTGGTTTGGTCGGCACGTTT]100 μl 50 μl 1.0 μl AR (R, 25 pmol/μl) [AAGTGTCCTTCTCCATCCTA] 2100 μl 50 μl 1.0 μl MgCl 2 (25 mM)15 μl 7.5 μl 0.15 μl Taq DNA polymerase (5 U/μl)2.4 ml 1.2 ml 24 μl Total (To each 24 ul reaction add 1 μl template DNA)Table 8.4.2.1. F and R indicate forward and reverse orientation. Use 1 μl DNA template.PCR cycle conditions94°C/5min x 1 cycle(94°C/30sec -o- 50°C/30sec -o- 72°C/30sec) x 30 cycles72°C/5min x 1 cycle4°C holdRestriction enzyme digestAdd 0.5 μl HhaI restriction enzyme to 10 μl PCR product from above reaction. Allow to incubate at 37°Cfor 3-24 hr. For shorter times, incomplete digests could be a problem.Run samples on a 2% agarose EtBr gel. Primers create fragments of 397 bp for Mopti and 225 and110bp for Savanna forms (Figure 8.4.2.1).1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.2 Arabiensis primer not necessary if certain of working with gambiae strain.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.2 An. gambiae Ribosomal DNA Type – Fanello et al.Page 2 of 2Figure 8.4.2.1. Gel electrophoresis of Mopti/SavannarDNA assay. Lanes 2-5 and 6-9 contain Mopti andSavanna PCR products respectively. Lanes 1 and 10contain 1kb ladder marker. Photo from (Wilkins et al.2006); used with permission.ReferencesFanello C, Santolamazza F, della Torre A (2002) Simultaneous identification of species and molecularforms of Anopheles gambiae complex by PCR-RFLP. Med Veterinary Entomol 16:461-464Scott JA, Brogdon WG, Collins FH (1993) Identification of single specimens of the Anopheles gambiaecomplex by the polymerase chain reaction. Am J Trop Med Hyg 49:520-529Wilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect single nucleotide polymorphisms forAnopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin inAnopheles arabiensis. Malar J 5:12596 well PCR sample preparation template

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.3 Combined An. gambiae complex and ribosomal DNA type assay for M/S discriminationPage 1 of 28.4.3 Combined An. gambiae complex and ribosomal DNA type assay forM/S discriminationLiz WilkinsIntroductionThe Wilkins et al. (2006) method of Anopheles gambiae complex discrimination is based on speciesspecificsingle nucleotide polymorphisms (SNPs) in the intergenic spacer region (IGS) (section 8.4.1). Wehave added primers to this method to simultaneously elucidate the Ribosomal DNA type. Theseadditional primers also incorporate the intentional mismatches into the primers (Intentional MismatchPrimers (IMPs)) to increase the specificity (Wilkins et al. 2006). An RT-PCR based method is alsoavailable in Chapter 8.5.1.1.PCR authentication for the members of the Anopheles gambiae complex (Wilkins et al. 2006) withadditional primers for Ribosomal DNA typePrepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent910 μl 455 μl 9.1 μl sterile H 2 O500 μl 250 μl 5.0 μl 5X GoTaq PCR Buffer250 μl 125 μl 2.5 μl dNTP (2.5 mM mix)30 μl 15 μl 0.3 μl MgCl 2 (25mM)100 μl 50 μl 1.0 μl IMP-UN (F, 25pmol/μl) [GCTGCGAGTTGTAGAGATGCG]100 μl 50 μl 1.0 μl AR-3T (R, 25pmol/μl) [GTGTTAAGTGTCCTTCTCCgTC]100 μl 50 μl 1.0 μl GA-3T (R, 25pmol/μl) [GCTTACTGGTTTGGTCGGCAtGT]100 μl 50 μl 1.0 μl ME-3T (R, 25pmol/μl) [CAACCCACTCCCTTGACGaTG]200 μl 100 μl 2.0 μl QD-3T (R, 25pmol/μl) [GCATGTCCACCAACGTAAAtCC]100 μl 50 μl 1.0 μl IMP-S1 (R, 25pmol/μl) [CCAGACCAAGATGGTTCGcTG]100 μl 50 μl 1.0 μl IMP-M1 (R, 25pmol/μl) [TAGCCAGCTCTTGTCCACTAGTtTT]15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase2.5 ml 1.25 ml 25 μl Total (To each 25 μl reaction add 1 μl template DNA)Table 8.4.2.1. Lower case nucleotides indicates the intentional mismatch in the primer sequences.Nucleotides in bold are located at site of SNP (where applicable), F and R indicate forward and reverseorientation. Use 1 μl DNA template. Improved specificity can be achieved by eliminating primers forspecies not found in the sampling area. Add water to compensate for this volume if all are not included.PCR cycle conditions95°C/5min x 1 cycle(95°C/30sec , 58°C/30sec , 72°C/30sec) x 30 cycles72°C/5min x 1 cycle4°C holdRun samples on a 2% agarose EtBr gel; load 5 μl sample.1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.3 Combined An. gambiae complex and ribosomal DNA type assay for M/S discriminationPage 2 of 2Primers create fragments of 636 An. quadriannulatus, 528 An. merus, 463 An. gambiae, 387 An.arabiensis, 333 An. gambiae M, 221 An. gambiae S. (Figure 8.4.2.1).Figure 8.4.2.1. Lane 1 An.quadriannulatus, lane 2 An.merus/melas, lane 3 An. gambiae,Lane 4 An. arabiensis, Lane 5 1kbladder.Figure 8.4.2.2. Top portion of gel, An. gambiae and M type;bottom portion of gel, An. gambiae and S type. 1kb ladder,first and last lanes.ReferencesWilkins EE, Howell PI, Benedict MQ (2006) IMP PCR primers detect single nucleotide polymorphisms forAnopheles gambiae species identification, Mopti and Savanna rDNA types, and resistance to dieldrin inAnopheles arabiensis. Malar J 5:125

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.4 Anopheles funestus Complex - KoekemoerPage 1 of 28.4.4 Anopheles funestus ComplexLizette KoekemoerIntroductionThe Anopheles funestus group consists of at least eleven species: An. funestus Giles, An. vaneedeniGillies and Coetzee, An. rivulorum Leeson, An. rivulorum-like, An. leesoni Evans, An. confusus Evansand Leeson, An. parensis Gillies, An. brucei Service, An. aruni Sobti, An. fuscivenosus Leeson and anAsian member An. fluviatilis James. These species are not all sympatric. Originally, distinguishing themembers of the group was mainly based on karyotyping (Green and Hunt 1980; Green 1982). Recently,however, easier PCR based assays have been developed that distinguished the most common membersof the group. The PCR assay presented is based on species-specific single nucleotide polymorphisms(SNPs) in the internal transcribed spacer region 2 (ITS2) (Koekemoer et al. 2002; Cohuet et al. 2003).PCR authentication for the members of the Anopheles funestus group (Koekemoer et al. 2002;Cohuet et al. 2003)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent300 μl 150 μl 3.0 μl sterile H 2 O250 μl 125 μl 2.5 μl Taq 10X PCR Buffer with MgCl 2250 μl 125 μl 2.5 μl dNTP (2 mM mix)150 μl 75 μl 1.5 μl MgCl 2 (25mM)200 μl 100 μl 2.0 μl UV (F, 33 pmol/μl) [TGT GAA CTG CAG GAC ACA T]200 μl 100 μl 2.0 μl FUN (R, 33 pmol/μl) [GCA TCG ATG GGT TAA TCA TG]200 μl 100 μl 2.0 μl VAN (R, 33 pmol/μl) [TGT CGA CTT GGT AGC CGA AC]200 μl 100 μl 2.0 μl RIV (R, 33 pmol/μl) [CAA GCC GTT CGA CCC TGA TT]200 μl 100 μl 2.0 μl PAR (R, 33 pmol/μl) [TGC GGT CCC AAG CTA GGT TC]200 μl 100 μl 2.0 μl RIVLIKE (R, 33 pmol/μl)[CCG CCT CCC GTG GAG TGG GGG]200 μl 100 μl 2.0 μl LEES (R, 33 pmol/μl) [TAC ACG GGC GCC ATG TAG TT]50 μl 25 μl 0.5 μl Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 24 μl Total (To each 24 ul reaction add 1 μl template DNA)Table 8.4.4.1. F and R indicate forward and reverse orientation. Volume can be reduced to 12.5 ul foreconomy. DNA extraction negative control to be included in addition to PCR reaction mix negativecontrol. If all primers are not needed, complete the total volume with water.PCR cycle conditions94°C/2min x 1 cycle(94°C/30sec , 45°C/30sec , 72°C/40sec) x 36 cycles72°C/5min x 1 cycle4°C holdRun samples on a 2.5% agarose EtBr gel; load 10 μl sample1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.4 Anopheles funestus Complex - KoekemoerPage 2 of 2Primers create fragments of 587bp An. vaneedeni, 505bp An. funestus, 411bp, An. rivulorum, 313bp An.rivulorum-like (West Africa), 252bp An. parensis and 46bp An. leesoni 636 (Figure 8.4.4.1).Figure 8.4.4.1. Amplified fragments using thespecies-specific polymerase chain reaction for theidentification of members of the Anophelesfunestus group. Lanes 1 and 9, 100-basepair DNAsize marker ladder; lane 2, An. vaneedeni; lane 3,An. funestus; lane 4, An.rivulorum; lane 5, An.rivulorum-like; lane 6, An. parensis; lane 7,An.leesoni; lane 8, negative control. The sizes ofthe fragments of the ladder are 1,000, 800, 700,600, 500, 400, 300, 200, and 100 bp.ReferencesCohuet A, Simard F, Toto JC, Kengne P, Coetzee M, Fontenille D (2003) Species identification within theAnopheles funestus group of malaria vectors in Cameroon and evidence for a new species. Am J TropMed Hyg 69:200-205Green CA (1982) Cladistic analysis of mosquito chromosome data (Anopheles (Cellia) Myzomyia. JHeredity 73:2-11Green CA, Hunt RH (1980) Interpretations of variation in ovarian polytene chromosomes of Anophelesfunestus Giles, An. parensis Gillies, and An. aruni. Genetica 51:187-195Koekemoer LL, Kamau L, Hunt RH, Coetzee M (2002) A cocktail polymerase chain reaction assay toidentify members of the Anopheles funestus (Diptera: Culicidae) group. Am J Trop Med Hyg 66:804-81196 well PCR sample preparation template

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.5 Anopheles funestus Complex – MR4Page 1 of 28.4.5 PCR Discrimination of the Anopheles funestus ComplexLiz Wilkins, MR4 StaffLike the Koekemoer et al.(2002) assay, the MR4 method of Anopheles funestus complex discrimination isbased on species-specific single nucleotide polymorphisms (SNPs) in the second internal transcribedspacer region (ITS2). However, it also incorporates intentional mismatches into the primers (IntentionalMismatch Primers (IMPs)) to increase the specificity (Wilkins et al. 2006).PCR authentication for the members of the Anopheles funestus groupPrepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent1000 μl 500 μl 10.0 μl sterile H 2 O500 μl 250 μl 5.0 μl Taq 5X PCR Buffer with MgCl 2100 μl 50 μl 1.0 μl dNTP (2 mM mix)200 μl 100 μl 2.0 μl MgCl 2 (25mM)100 μl 50 μl 1.0 μl UV (F, 5 pmol/μl) [CCG ATG CAC ACA TTC TTG AGT GCC TA]100 μl 50 μl 1.0 μl FUN (R, 5 pmol/μl) [CTC GGG CAT CGA TGG GTT AAT CAT G]100 μl 50 μl 1.0 μl VAN (R, 5 pmol/μl) [AAC TCT GTC GAC TTG GTA GCC GAA C]100 μl 50 μl 1.0 μl RIV (R, 5 pmol/μl) [AAT CAG GGT CGA ACG GCT TGC CG]100 μl 50 μl 1.0 μl PAR (R, 5 pmol/μl) [GCC CTG CGG TCC CAA GCT AGA TT]100 μl 50 μl 1.0 μl RIVLIKE (R, 5 pmol/μl) [CTC CCG TGG AGT GGG GGA TC]100 μl 50 μl 1.0 μl LEES (R, 5 pmol/μl) [GAC GGC ATC ATG GCG AGC AGC]10 μl 5 μl 0.1 μl Taq DNA polymerase (5 U/μl) –MR4 uses GoTaq, Promega2.5 ml 1.25 ml 25 μl Total (To each 25 μl reaction add 1 μl template DNA)Table 8.4.5.1. F and R indicate forward and reverse orientation. DNA extraction negative control to beincluded in addition to PCR reaction mix negative control. If all primers are not needed, complete the totalvolume with water.PCR cycle conditions94°C/4min x 1 cycle(94°C/30sec , 58°C/30sec , 72°C/45sec) x 30 cycles72°C/7min x 1 cycle4°C holdRun samples on a 1.5% agarose EtBr gel; load 10 μl sample1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.5 Anopheles funestus Complex – MR4Page 2 of 2Primers create fragments of 496bp An.vaneedeni, 424bp An. funestus, 346bp, An.rivulorum, 241bp An. rivulorum-like (WestAfrica), 176bp An. parensis and 93bp An.leesoni (Figure 8.4.5.1)References:Koekemoer LL, Kamau L, Hunt RH, Coetzee M(2002) A cocktail polymerase chain reactionassay to identify members of the Anophelesfunestus (Diptera: Culicidae) group. Am J TropMed Hyg 66:804-811Wilkins EE, Howell PI, Benedict MQ (2006) IMPPCR primers detect single nucleotidepolymorphisms for Anopheles gambiae speciesidentification, Mopti and Savanna rDNA types,and resistance to dieldrin in Anophelesarabiensis. Malar J 5:125Figure 8.4.5.1 Lane 1 1kb ladder,lane 2 An. funestus, lane 3 An.rivulorum, lane 4 An. parensis, lane5 An. leesoni.96 well PCR sample preparation template

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.6 Anopheles minimus ComplexPage 1 of 28.4.6 Anopheles minimus Species ComplexMR4 StaffIntroductionAnopheles minimus is a complex comprised of three members: species A (minimus sensu stricto), C(harrisoni), and E. Anopheles minimus and An. harrisoni occur sympatrically throughout Southeast Asiaand are implicated in malaria transmission while species E is limited to the Ryuku Islands of Japan(Garros et al. 2006). An original method for distinguishing the various members was determining thepresence of a pale humeral spot; however this method has proven unreliable in use (Sungvornyothin etal. 2006). A PCR-RFLP (restriction fragment length polymorphism) designed utilizing SNP differences inthe ITS2 regions has been developed to distinguish complex members of An. minimus (Van Bortel et al.2000).PCR authentication for the members of the Anopheles minimus complex (Van Bortel et al. 2000)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent1555 μl 777.5 μl 15.55 μl sterile H 2 O500 μl 250 μl 5.0 μl GoTaq 5X PCR Buffer with MgCl 2100 μl 50 μl 1.0 μl dNTP (2.5 mM mix)30 μl 15 μl 0.3 µl MgCl 2 (25 mM)150 μl 75 μl 1.5 µl ITS2 A primer (1 pmol/μl) [TGT GAA CTG CAG GAC ACA T]150 μl 75 μl 1.5 µl ITS2 B primer (1 pmol/μl) [TAT GCT TAA ATT CAG GGG GT]15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase (5 U/μl)2.5 ml 1.25 ml 25 μl Total (To each 24 ul reaction add 1 μl template DNA)Table 8.4.6.1. Prepare PCR Master Mix for 96, 48 or 1 25 μl PCR reactions. Add reagents in the orderpresented. Use 1 μl DNA template.PCR cycle conditions94°C/4min x 1 cycle(94°C/30sec, 53°C/40sec, 72°C/30sec) x 35 cycles72°C/10min x 1 cycle4°C holdRFLP ProcedurePer reaction well:2 μl sterile H 2 O2 μl Restriction Enzyme buffer1 μl Sau 96I restriction enzyme (GGNCC cutting site)15 μl PCR product20 μl Total1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.6 Anopheles minimus ComplexPage 2 of 2Incubate at 60 o C for 2 hours.Run samples on a 2.5% agarose EtBr gel for visualization.The PCR-RFLP procedure will yield the following products: An. minimus 220+200bp and An. harrisoni300+220bp products.ReferencesGarros C, Van Bortel W, Trung HD, Coosemans M, Manguin S (2006) Review of the Minimus Complex ofAnopheles, main malaria vector in Southeast Asia: from taxonomic issues to vector control strategies.Trop Med Int Health 11:102-114Sungvornyothin S, Garros C, Chareonviriyaphap T, Manguin S (2006) How reliable is the humeral palespot for identification of cryptic species of the Minimus Complex? J Am Mosq Control Assoc 22:185-191Van Bortel W, Trung HD, Roelants P, Harbach RE, Backeljau T, Coosemans M (2000) Molecularidentification of Anopheles minimus s.l. beyond distinguishing the members of the species complex.Insect Mol Biol 9:335-34096 well sample preparation template

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.7 Anopheles quadrimaculatus Complex – Cornel et al.Page 1 of 28.4.7 Anopheles quadrimaculatus Species Complex (Cornel et al.)MR4 StaffIntroductionThough malaria is no longer endemic in the United States, the competent vector An. quadrimaculatus isstill present. It has been determined that the An. quadrimaculatus complex is made of at least fivemembers: An. inundatus, An. diluvialis, An. maverlius, An. smaragdinus, and An. quadrimaculatus(Narang et al. 1990). A PCR based on SNPs in the internal transcribed spacer region has beendeveloped for An. quadrimaculatus that can distinguish the various members of the complex (Cornel et al.1996).PCR authentication for the members of the Anopheles quadrimaculatus complex (Cornel et al.1996)Prepare PCR Master Mix for 96, 48 or 1 25 μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent450 μl 225 μl 4.5 μl sterile H 2 O250 μl 125 μl 2.5 μl 10X Perkins Elmer PCR Buffer100 μl 50 μl 1.0 μl dNTP (2 mM mix)200 μl 100 μl 2.0 µl MgCl 2 (25 mM)160 μl 80 μl 1.6 μl AQU (F, 1 pmol/μl) [CGACACAGCTCGATGTACAC]100 μl 50 μl 1.0 μl AQA (R, 1 pmol/μl) [TCCGTAGGAGGCTGCATTTT]460 μl 230 μl 4.6 μl AQB (R, 1 pmol/μl) [CACACTACACAACACGCTTT]230 μl 115 μl 2.3 μl AQC (R, 1 pmol/μl) [TACCCCGGCCTTGTAGCAAA]540 μl 270 μl 5.4 μl AQD (R, 1 pmol/μl) [ATGCAAAAGGTGGTGTTGTG]12.5 μl 6.25 μl 0.125 μl Taq DNA polymerase (5U/μl)2.5 ml 1.25 ml 25 μl Total (To each 24 ul reaction add 1 μl template DNA)Table 8.4.7.1. F and R indicate forward and reverse orientation. Use 1 μl DNA template. If all primers arenot used, complete the volume with water.PCR Cycle conditions94°C/2min x 1 cycle(94°C/1min, 50°C/2min, 72°C/2min) x 25 cycles72°C/10min x 1 cycle4°C holdRun samples on a 2% agarose EtBr gel; load 5 μl samplePrimers create fragments of 319bp An. quadrimaculatus, 227bp An. smaragdinus, 293bp An. inundatusor An. diluvialis, and 141bp An. maverlius.1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.7 Anopheles quadrimaculatus Complex – Cornel et al.Page 2 of 2ReferencesCornel AJ, Porter CH, Collins FH (1996) Polymerase chain reaction species diagnostic assay forAnopheles quadrimaculatus cryptic species (Diptera: Culicidae) based on ribosomal DNA ITS2sequences. J Med Entomol 33:109-116Narang SK, Seawright JA, Kaiser PE (1990) Evidence for microgeographic genetic subdivisions ofAnopheles quadrimaculatus species C. J Am Mosq Control Assoc 6:179-18796 well sample preparation template

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.8 Anopheles quadrimaculatus Complex – Levine et al.Page 1 of 28.4.8 Anopheles quadrimaculatus Species Complex (Levine et al.)Mark BenedictIntroductionThough malaria is no longer endemic in the United States, the competent vector An. quadrimaculatus isstill present. It has been determined that the An. quadrimaculatus complex is made up of at least fivemembers: An. inundatus, An. diluvialis, An. maverlius, An. smaragdinus, and An. quadrimaculatus(Narang et al. 1990). A PCR based on SNPs in the internal transcribed spacer region was developed forAn. quadrimaculatus that can distinguish the various members of the complex (Cornel et al. 1996),however a later modification was developed that uses primers that have similar 60°C annealingtemperatures and the concentrations recommended are the same as one another (Levine et al. 2004).Fragment sizes are also slightly more distinct (Figure 4.2.6.1).PCR authentication for the members of the Anopheles quadrimaculatus complex (Cornel et al.1996)Prepare PCR Master Mix for 96, 48 or 1 25 μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent380 μl 190 μl 3.5 μl sterile H 2 O250 μl 125 μl 2.5 μl 10X Promega Taq PCR Buffer w/o MgCl 2250 μl 125 μl 2.5 μl dNTP (2 mM mix each G,A,T,C)250 μl 125 μl 2.5 µl MgCl 2 (32 mM)250 μl 125 μl 2.5 μl AquadU-2 (F, 10 pmol/μl) [GTGCGACACAGCTCGATG]250 μl 125 μl 2.5 μl AquadQ-2 (R, 10 pmol/μl) [CCGTAGGAGGCTGCATTTTA]250 μl 125 μl 2.5 μl AquadS-2 (R, 10 pmol/μl) [GAACACACTACACAACACGCTTT]250 μl 125 μl 2.5 μl AquadD1-2 (R, 10 pmol/μl) [AGGCCCATGTACTCCGTAGG]250 μl 125 μl 2.5 μl AQD (R, 10 pmol/μl) [ATGCAAAAGGTGGTGTTGTG]20 μl 10 μl 0.5 μl Taq DNA polymerase (5U/μl)2.4 ml 1.2 ml 24 μl Total (To each 24 ul reaction add 1 μl template DNA)Table 8.4.8.1. F and R indicate forward and reverse orientation. Use 1 μl DNAtemplate. If all primers are not used, complete the volume with water.PCR cycle conditions95°C / 2min x 1 cycle(95°C / 30s, 57°C / 30s, 72°C / 30s) x 30 cycles4°C holdRun samples on a 2% agarose EtBr gel; load 5 μl sample.Primers create fragments of 321 bp for An. quadrimaculatus, 233 bp forAn. smaragdinus, 353 bp for An. diluvialis, 365 for An. inundatus and 141bp forFigure 8.4.8.1. PCRproducts obtained fromamplification of A.quadrimaculatus s.s.(Q), A. smaragdinus(S), A. diluvialis (D)and A. maverlius (M)1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 rxnscompensate for imprecise measurements.

Chapter 8 : Field Techniques8.4 Species Complex Authentication by PCR8.4.8 Anopheles quadrimaculatus Complex – Levine et al.Page 2 of 2An. maverlius.ReferencesCornel AJ, Porter CH, Collins FH (1996) Polymerase chain reaction species diagnostic assay forAnopheles quadrimaculatus cryptic species (Diptera: Culicidae) based on ribosomal DNA ITS2sequences. J Med Entomol 33:109-116Levine RS, Peterson AT, Benedict MQ (2004) Distribution of members of Anopheles quadrimaculatusSay s.l. (Diptera: Culicidae) and implications for their roles in malaria transmission in the United States. JMed Entomol 41:607-613Narang SK, Seawright JA, Kaiser PE (1990) Evidence for microgeographic genetic subdivisions ofAnopheles quadrimaculatus species C. J Am Mosq Control Assoc 6:179-18796 well sample preparation template

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCRPage 1 of 48.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool Using Real-Time PCRChris Bass, Martin Williamson, John Vontas, Hilary Ranson, Martin Donnelly and Lin FieldIntroductionThe ‘Vector Population Monitoring Tool’ (VPMT) consists of a suite of high-throughput assays that can beused to screen mosquito disease vector populations for a number of traits. These include: Anopheles gambiae s.l. species identification (Section 8.5.1.1) Detection of infection with Plasmodium parasites (Section 8.5.1.2) Detection of insecticide resistance mechanisms. To date this includes:1) Detection of knock-down resistance (kdr) mutations (Section 8.5.1.3)2) Detection of insensitive acetylcholinesterase (iAChE) (Section 8.5.1.4)3) Detection of resistance to dieldrin (rdl) (Section 8.5.1.5)The assays that constitute the VPMT have been designed to be high-throughput and require very smallamounts of starting material (DNA) which can be extracted from mosquitoes that have been stored in awide variety of ways (e.g. in ethanol, isopropanol, dried, frozen). This is to ensure that they can be usedfor assaying field collected mosquitoes without the need for a cold chain to preserve the specimens. Inthe case of the species identification assays, a single mosquito leg can be used as template for PCRwithout the need to first extract DNA.The assays presented in this section are based on TaqMan SNP genotyping. The TaqMan assay is aPCR method employing oligonucleotide probes that are dual-labeled with a fluorescent reporter dye and aquencher molecule. Amplification of the probe-specific product causes cleavage of the probe, generatingan increase in reporter fluorescence as the reporter dye is released away from the quencher. By usingdifferent reporter dyes, cleavage of allele-specific probes can be detected in a single PCR. The ‘closedtube’nature of the TaqMan platform means there is no requirement for post-PCR processing andconsequently assays are simple to perform and rapid to run.Ordering ReagentsPrimers: These are standard oligonucleotides and are available from a range of suppliers.Probes: Protocols described here use 2 types of fluorescently labeled probes:TaqMan® MGB Probes: available from Applied Biosystems. Each probe is labeled with a 5′ reporterdye (e.g. Vic or Fam) and also carries a 3′ non-fluorescent quencher and a minor groove binder (MGB) atthe 3′ end. The minor groove binder provides more accurate allelic discrimination by increasing the T Mbetween matched and mismatched probes. Store aliquots in aluminum foil.LNA Probes: These probes employ locked nucleic acid (LNA) modified nucleotides which have a similareffect to the MGB moiety. They can be purchased from Sigma or from Thermo Scientific. These areusually delivered lyophilized, with the yield given in nanomoles. Prepare aliquots and store each aliquotwrapped in aluminum foil/out of light. The LNA probe used in the species identification assay described inthis guide is labeled with Cy5. (note: Sigma class the Cy5 label as non-standard so the probe is moreexpensive than standard). When ordering: Type: mix DNA/LNA, backbone: PO, 5-Modification: Cy5, 3-Modification: BHQ2; [Cy5] AC+A+T+AG+GATGGA+G+A+AGG [BHQ2]. Thermo scientific uses LA, LC,LG, LT for LNA bases, so the sequence of the probe above should be copied in asacLALTLAgLGatggaLGLALAgg. Fill out the modifications part of the table as follows: at 5'-end: CY5, at 3'-end: Black Hole Quencher 2.

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCRPage 2 of 4PCR Mastermix: All the assays have been optimized using SensiMix NoRef DNA kit (Cat # QT505) fromQuantace.Assay CostAssay cost will vary from country to country so the following should be used as a general guide and is anapproximate price, correct as of 2008. In each case costs were calculated in US $ by using retail pricesfrom US websites. These prices do not include promotional or negotiated discounts or the cost ofdelivery. All primer/probe/PCR master-mix costs are from ordering at the largest available scale(substantial cost savings are made by ordering at the largest available scales). Cost calculations includeall the plastics required to set up the reactions (PCR tubes/filter tips etc.).Species ID (2 plex assay) US$ 0.75/specimenSpecies ID (3 plex assay) US$ 1/specimenPlasmodium detection assay US$ 0.75/specimenKdr assay US$ 1.5 (this is the combined cost of running two assays (kdr-w+kdr-e)iAChE assay US$ 0.75/specimenRdl assay US$ 1.5 (this is the combined cost of running two assays (A296G+A296S) /specimenReal-time PCR machinesThe assays described in this guide were originally developed using a Rotor-Gene 6000 (CorbettResearch). We have also trialed several of the assays using a Chromo4 and Mini-Opticon (Bio-Rad). Inthe case of the latter machines, both clear and white PCR tubes/plates were used successfully withslightly improved results using the white tubes/plates. Users of alternative real-time machines may find asmall amount of optimization is required to achieve optimal results.General advice and TroubleshootingMake aliquots of all reagents to avoid repeated freeze-thawing. This is especially important for thefluorescently labeled probes.Examine both the fluorescent traces and scatter plots when scoring samples.Use the auto-scale function of the real-time PCR machine software with caution as this can artificiallyelevate the fluorescent trace resulting in mis-scoring.If further assay optimization is required, the annealing temperature, the concentration of probes in thereaction, and number of temperature cycles are good parameters to alter in attempts to enhancesensitivity or reduce non-specific signals.If mosquito individuals are to be tested with the Plasmodium detection assay, DNA should be extractedfrom the head/thorax of mosquito specimens to avoid detecting Plasmodium stages in the blood meal/gutin addition to sporozoites in the salivary glands.Always include controls in assay runs. We recommend that these include one or more no-templatecontrols, and in the case of the An. gambiae s.l. species ID and Plasmodium assays, a positive control foreach probe (e.g An. gambiae s.s, An. arabiensis and An. quadriannulatus for the 3 plex species ID assayand P. falciparum and P. vivax for the Plasmodium assay). In the case of the kdr, iAChE and rdl assays, acontrol template for each genotype should be included (e.g., when running the kdr-w assay, include a kdrwhomozygous sample, a wild-type homozygous sample, and a kdr-w heterozygous sample). We havefound that including these controls greatly facilitates the interpretation of results and generally speeds upthe scoring of unknown samples.

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCRPage 3 of 4Assay ControlsThe following table details the control genotypes currently available from the Malaria Research andReference Reagent Resource Center (MR4) www.mr4.org. We have also created additional plasmidand whole genome amplified controls for these assays available from the authors on request.Anopheles gambiae s.l. species identification (Section 8.5.1.1)MRA-495 Anopheles arabiensis Genomic DNA from An. arabiensis KGBMRA-142 Anopheles gambiae Genomic DNA from An. gambiae, G3 strainMRA-761G Anopheles quadriannulatus Genomic DNA from An. quadriannulatus, SKUQUA strainDetection of infection with Plasmodium parasites (Section 8.5.1.2)MRA-102G Plasmodium falciparum Genomic DNA from P. falciparum 3D7MRA-341G Plasmodium vivax Genomic DNA from P. vivax ONGDetection of knock-down resistance (kdr) mutations (Section 8.5.1.3)MRA-762 Anopheles gambiae KISUMU Wild-type susceptible homozygous controlMRA-334 Anopheles gambiae RSP Kdr L1014S (Kdr-e) homozygousDetection of insensitive acetylcholinesterase (iAChE) (Section 8.5.1.4)MRA-762 Anopheles gambiae KISUMU Wild-type susceptible homozygous controlMRA-913 Anopheles gambiae AKRON Wild-type resistant controlDetection of resistance to dieldrin (rdl) (Section 8.5.1.5)MRA-115 Anopheles gambiae IN22C+ Wild-type resistant homozygous control (containedwithin IN22C+ strain, only c+ individuals)MRA-762 Anopheles gambiae KISUMU Wild-type susceptible homozygous controlMRA-495 Anopheles arabiensis KGB Wild-type susceptible homozygous controlMRA-764 Anopheles arabiensis SENN Wild-type resistant homozygous controlAuthors also have the following plasmid controls available:Wild-type (susceptible homozygous control), Kdr L1014S (kdr-e) homozygous and Kdr L1014F (kdr-w)homozygousWild-type (susceptible homozygous control), and the G119S homozygous

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCRPage 4 of 4Authors AffiliationsChris Bass, Martin Williamson, and Lin FieldAddress: Insect Molecular Biology Group, Department of Biological Chemistry, Rothamsted Research,Harpenden, AL5 2JQ, UKEmail:Chris Bass - chris.bass@bbsrc.ac.uk;Martin S Williamson - martin.williamson@bbsrc.ac.uk;Linda M Field - lin.field@bbsrc.ac.ukHilary Ranson, Martin DonnellyAddress: Vector Group, Liverpool School of TropicalMedicine, Pembroke Place, Liverpool L35QA, UKEmail:Martin J Donnelly - m.j.donnelly@liverpool.ac.uk;Hilary Ranson - Hranson@liverpool.ac.ukJohn VontasAddress: Laboratory of Pesticide Science, Agricultural University of Athens, Iera Odos 75, 118 55,Votanikos, Athens, GreeceEmail:John Vontas - vontas@aua.gr

8.5.1.1 An. gambiae s.l. species complex ID assayChapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.1 An. gambiae s.l. species complex ID assayPage 1 of 6Chris Bass, Martin Williamson, John Vontas, Hilary Ranson, Martin Donnelly and Lin FieldIntroductionTwo alternative assays have been developed. The first (Bass et al. 2007) (2-Plex Assay) uses two probesto distinguish between the main malaria vectors An. gambiae s.s. and An. arabiensis as one group andAn. quadriannulatus, An. melas or An. merus as a second group. This assay is suitable for use on realtimePCR machines that have two detection channels (such as Biorad’s Mini Opticon). This assay can beused alone to discriminate vector from non-vector (in regions where An. merus/melas/bwambae are notpresent) or in combination with an existing TaqMan assay (Walker et al. 2007) to further distinguish An.arabiensis from An. gambiae s.s..The second assay (Bass et al. 2008) (3-Plex Assay) is an enhancement of the first assay and uses threeprobes to distinguish between An. arabiensis, An. gambiae s.s. and An. quadriannulatus/merus/melas/bwambae as a group. This assay requires a real-time PCR machine that has a least threedetection channels. Traditional PCR assays can be found in Chapters 8.4.1 and 8.4.3.2-Plex Assay96 48 1 Reagent500 μl 250 μl 5.0μl sterile H 2 O1 ml 500 μl 10.0 μl SensiMix DNA kit (Quantace)160 μl 80 μl 1.6 μl primer ComF (800 nM) GCTTGGTGGTTTGTCCG160 μl 80 μl 1.6 μl primer ComR (800 nM) CTGTGTCGACGTGGTCCC40 μl 20 μl 0.4 µl probe AG/AA (200 nM) 6FAM- GACCAAGACGAGC40 μl 20 μl 0.4 µl probe AQ/AM (200 nM) VIC- GACCAAGACGCGC1.9 ml 950 μl 19 μl Total (To each 19 μl reaction add 1-2 μl genomic DNA)2-Plex Assay PCR cycle conditions95°C/10 min x 1 cycle; (95°C/15sec, 50°C/20sec, 72°C/20sec) x 45 cyclesMeasure fluorescence at the end of each cycle3-Plex Assay96 48 1 Reagent630 μl 315 μl 6.3μl sterile H 2 O1.25 ml 625 μl 12.5 μl SensiMix DNA kit (Quantace)200 μl 100 μl 2.0 μl primer Uni F (800nM) GTGAAGCTTGGTGCGTGCT200 μl 100 μl 2.0 μl primer Uni R (800nM) GCACGCCGACAAGCTCA50 μl 25 μl 0.5 µl LNA probe Aa+(200nM)[Cy5]AC+A+T+AG+GATGGA+G+A+AGG [BHQ2]20 μl 10 μl 0.2 µl TaqMan MGB probe (80 nM) Ag + VIC-TGGAGCGGaACAC50 μl 25 μl 0.5 µl TaqMan MGB probe (200nM) Aq + 6FAM-TGGAGCGGgACAC2.4 ml 1.2 ml 24 μl Total (To each 24 μl reaction add 1-2 μl genomic DNA)3-Plex Assay PCR cycle conditions95°C/10 min x 1 cycle(95°C/25sec, 66°C/60sec) x 40 cyclesMeasure fluorescence at the end of each cycle

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.1 An. gambiae s.l. species complex ID assayPage 2 of 6Interpreting Results and Examples: 2-Plex AssayInterpreting results is carried out by examining the levels of FAM and VIC fluorescence during PCR. Asubstantial increase in FAM fluorescence during PCR indicates an An. gambiae s.s. or An. arabiensisspecimen while a substantial increase in VIC fluorescence indicates an An. quadriannulatus, An. melas orAn. merus specimen (Figure 8.5.1.1.1.). An increase in both dyes would indicate a hybrid or acontaminated sample.To help identify species the software that accompanies the real-time PCR machine may allow endpointfluorescence values for the two dyes to be automatically corrected for background and plotted againsteach other in bi-directional scatter plots (Figure 8.5.1.1.2.).3025Cycling of FAM probe (An.arabiensis/An. gambiae)Fluorescence20151055 10 15 20 25 30 35 40 45Cycle6050Cycling of VIC probe (An.quadriannulatus/An. melas/An. merus)Fluorescence403020105 10 15 20 25 30 35 40 45CycleFigure 8.5.1.1.1 Species identification using the 2-plex TaqMan assay designed to distinguish theprincipal vector species An. gambiae s.s. and An. arabiensis from the other members of the complex. Inthis example two or more specimens of An. gambiae s.s., (red trace) An. arabiensis, (blue trace) An.melas (green trace), An. merus (purple trace) and An. quadriannulatus (orange trace).

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.1 An. gambiae s.l. species complex ID assayPage 3 of 6An. gambiae s.s. /An. arabiensisAn. quadriannulatus/An.merus/An. melasFailedreactionsFigure 8.5.1.1.2. Scatter plot analysis of TaqMan fluorescence data. In this example real time PCR wascarried out using the newly developed TaqMan assay designed to distinguish the principal vector speciesAn. gambiae s.s. and An. arabiensis from the other members of the complex on 96 samples.Fluorescence values of the FAM labeled probe specific for An. gambiae s.s. and An. arabiensis were thenplotted against the VIC labeled probe specific for An. quadriannulatus, An. melas and An. merus.Interpreting Results and Examples: 3-Plex AssayA substantial increase in Cy5 fluorescence (probe Aa) during PCR identifies an An. arabiensis specimen,an increase in VIC fluorescence (probe Ag) identifies an An. gambiae s.s. sample, and an increase in6FAM fluorescence (probe Aq) identifies An. quadriannulatus, An. melas or An. merus specimen (Figure8.5.1.1.3.). An increase in two or more of the dyes would indicate a hybrid or a contaminated sample.

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.1 An. gambiae s.l. species complex ID assayPage 4 of 6Fluorescence7060504030Cycling of VIC probe (An.gambiae)20105 10 15 20 25 30 35 40CycleFluorescence4035302520Cycling of Cy5 probe (An.arabiensis)1510Fluorescence54540353025205 10 15 20 25 30 35 40CycleCycling of FAM probe (An.quadriannulatus/An. melas/An.merus)151055 10 15 20 25 30 35 40CycleFigure 8.5.1.1.3. Species identification using the multiplex real-time PCR assay. 20 or more specimensof An. gambiae s.s. (red trace), An. arabiensis (blue trace) and An. quadriannulatus/An. melas/An. merus.(green trace).Assay Notes and TroubleshootingThe 3-plex species ID was initially run on a Corbett rotorgene PCR machine with an annealing/extensiontime of 60°C. At this temperature the probes Aa and Aq (Cy5- and 6FAM-labelled) showed specificamplification of An. arabiensis and An. quadriannulatus/An. melas/An. merus respectively. However whilethe probe Ag (VIC-labelled) also gave sensitive detection of An. gambiae s.s., when An. quadriannulatus,An. melas or An. merus DNAs were tested a low level ‘background’ fluorescence signal was observed,presumably from non-specific binding of this probe (see figure 4 below). This could be eliminated byincreasing the annealing/extension temperature to 66°C and lowering the final probe concentration in thePCR from 200 to 80 nM (Figure 8.5.1.1.3.).However, when this assay was run recently on an alternative PCR machine (Chromo4, Bio-Rad) usingthe modified conditions and white PCR tubes the same low level ‘background’ was seen once more.Increasing the annealing/extension temperature to 67°C and lowering the final probe concentration in the

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.1 An. gambiae s.l. species complex ID assayPage 5 of 6PCR from 200 to 50 nM showed a degree of improvement but did not completely eliminate thebackground signal. In practice this background does not inhibit scoring as an An. quadriannulatus, An.melas or An. merus specimen shows a strong signal in the Cy5 channel while a true Anopheles gambiaes.s. individual only shows signal in the VIC channel.10080Cycling of VIC probe (An.gambiae s.s.)Fluorescence604020605005 10 15 20 25 30 35 40CycleCycling of Cy5 probe (An.arabiensis)Fluorescence40302010155 10 15 20 25 30 35 40CycleCycling of FAM probe (An.quadriannulatus/merus/melas)Fluorescence1055 10 15 20 25 30 35 40CycleFigure 8.5.1.1.4. Species identification using the multiplex real-time PCR assay. When the assay is run at60°C a background signal from the VIC probe is observed when using An. quadriannulatus, An. melas orAn. merus DNAs. An. gambiae s.s. (red trace), An. arabiensis (blue trace) and An. quadriannulatus/An.melas/An. merus (green, pink and light blue traces).ReferencesBass C, Williamson MS, Field LM (2008) Development of a multiplex real-time PCR assay foridentification of members of the Anopheles gambiae species complex. Acta Trop 107:50-53Bass C, Williamson MS, Wilding CS, Donnelly MJ, Field LM (2007) Identification of the main malariavectors in the Anopheles gambiae species complex using a TaqMan real-time PCR assay. Malar J 6:155Walker ED et al. (2007) Identification of field caught Anopheles gambiae s.s. and Anopheles arabiensisby TaqMan single nucleotide polymorphism genotyping. Malar J 6:23

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.1 An. gambiae s.l. species complex ID assayPage 6 of 6

8.5.1.2 Plasmodium detection assayChapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.2 Plasmodium detection assayPage 1 of 4Chris Bass, Martin Williamson, John Vontas, Hilary Ranson, Martin Donnelly and Lin FieldIntroductionThis assay detects all four malaria-causing Plasmodium species and discriminates P. falciparum from P.vivax, P. ovale and P. malariae. This method is not inhibited by the storage of mosquito specimens bydrying or in ethanol or isopropanol (Bass et al. 2008). The assay is suitable for use on real-time PCRmachines that have two or more detection channels. This assay will detect stages of Plasmodium in theblood meal/gut of the mosquito; therefore, to ensure only sporozoites within the salivary glands aredetected, DNA should be extracted from head-thorax only and the abdomen removed prior to extraction.An alternate technique can be found in Chapter 8.1.Assay conditions96 48 1 Reagent480 μl 240 μl 4.8 μl sterile H 2 O1.25 ml 625 μl 10.0 μl SensiMix DNA kit (Quantace)200 μl 100 μl 1.6 μl primer PlasF (800nM) GCTTAGTTACGATTAATAGGAGTAGCTTG200 μl 100 μl 1.6 μl primer PlasR (800nM) GAAAATCTAAGAATTTCACCTCTGACA20 μl 10 μl 0.6 µl TaqMan MGB probe (200 nM) Falcip+ 6FAM-TCTGAATACGAATGTC50 μl 25 μl 0.4 µl TaqMan MGB probe (200 nM) OVM+ VIC-CTGAATACAAATGCC2.2 ml 1.1 ml 19 μl Total (To each 19 μl reaction add 1-2 μl genomic DNA)NOTE: In tests of this assay using the Rotor-Gene 6000 (Corbett Research) no loss of sensitivity wasobserved for half volumes of reagents (10μl total volume).Assay PCR cycle conditions95°C/10 min x 1 cycle; (95°C/10sec, 60°C/45sec) x 40 cyclesMeasure fluorescence at the end of each cycleInterpreting Results and ExamplesThe Plasmodium assay uses two probes, the first labeled with 6FAM detects P. falciparum and thesecond, labeled with VIC, detects P. vivax/P. ovale/P. malariae. A substantial increase in FAMfluorescence during PCR indicates the presence of P. falciparum whilst a substantial increase in VICfluorescence indicates the presence of P. vivax, P. ovale or P. malariae (Figure 8.5.1.2.1). An increase inboth dyes would indicate a mixed infection.To help with species assignment, the Rotor-Gene software allows endpoint fluorescence values for thetwo dyes to be automatically corrected for background and plotted against each other in bi-directionalscatter plots (Figure 8.5.1.2.2).

20Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.2 Plasmodium detection assayPage 2 of 415Cycling of FAM probe (P.falciparum)PfFluorescence105 10 15 20 25 30 35 40 45Cycle403530Cycling of VIC probe (P.malariae/P. ovale/P. vivax)PoPmFluorescence2520Pv151055 10 15 20 25 30 35 40 45CycleFigure 8.5.1.2.1. Detection of Plasmodium species by TaqMan assay. In this example two or morespecimens of P. falciparum (blue trace), P. vivax (green trace), P. ovale (yellow trace) and P. malariae(red trace) were tested.

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.2 Plasmodium detection assayPage 3 of 4P. falciparumP. vivax/P.ovale/P. malariaenegativeFigure 8.5.1.2.2. Scatter plot analysis of TaqMan fluorescence data. In this example the TaqMan assaywas carried out on ~30 Plasmodium genomic DNA samples and five no template negative controls.Fluorescence values of the FAM labeled probe specific for P. falciparum were plotted against those of theVIC labeled probe specific for P. vivax, P. ovale and P. malariae.Notes and TroubleshootingThe Plasmodium detection assay was originally optimized using purified Plasmodium genomic DNA astemplate and carrying out 45 cycles of PCR. When tests were carried out on blood fed mosquitoes usinga standard ‘quick and dirty’ DNA extraction, a small amount of non-specific fluorescence was sometimesseen after 40 cycles (see figure 7 below). For this reason we recommend restricting the number of cyclesin PCR to 40.70Fluorescence60504030Non-specific productsappearing after 40cycles of PCR20105 10 15 20 25 30 35 40 45CycleFigure 8.5.1.2.3. Cycling of VIC labelled probe specific for P. ovale, P. vivax, and P. malariae.

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.2 Plasmodium detection assayPage 4 of 4ReferencesBass C, Nikou D, Blagborough AM, Williamson MS, Vontas J, Sinden RE, Field LM (2008) PCR-baseddetection of Plasmodium in Anopheles mosquitoes: a comparison of a new high-throughput assay withexisting methods. Malaria J 7:177

8.5.1.3 Knock down resistance (kdr) assaysChapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.3 Knockdown resistance (kdr) assaysPage 1 of 6Chris Bass, Martin Williamson, John Vontas, Hilary Ranson, Martin Donnelly and Lin FieldIntroductionTwo separate assays have been developed for the detection of kdr-w (L1014F) or kdr-e (L1014S) (Basset al. 2007). This assay is suitable for use on real-time PCR machines that have two or more detectionchannels. Alternate PCR assays can be found in Chapters 5.3.1 and 5.3.2.Assay conditions for kdr-w (L1014F)96 48 1 Reagent500 μl 250 μl 5.0 μl sterile H 2 O1.0 ml 500 μl 10.0 μl SensiMix DNA kit (Quantace)160 μl 80 μl 1.6 μl primer kdr-forward (800nM) CATTTTTCTTGGCCACTGTAGTGAT160 μl 80 μl 1.6 μl primer kdr-reverse (800nM) CGATCTTGGTCCATGTTAATTTGCA40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) WT VIC-CTTACGACTAAATTTC40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) KdrW 6FAM-ACGACAAAATTTC1.9 ml 950 μl 19 μl Total (To each 19 μl reaction add 1-2 μl genomic DNA)Assay conditions for kdr-e (L1014S)96 48 1 Reagent500 μl 250 μl 5.0 μl sterile H 2 O1.0 ml 500 μl 10.0 μl SensiMix DNA kit (Quantace)160 μl 80 μl 1.6 μl primer kdr-forward (800nM) CATTTTTCTTGGCCACTGTAGTGAT160 μl 80 μl 1.6 μl primer kdr-reverse (800nM) CGATCTTGGTCCATGTTAATTTGCA40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) WT VIC-CTTACGACTAAATTTC40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) KdrE 6FAM-ACGACTGAATTTC1.9 ml 950 μl 19 μl Total (To each 19 μl reaction add 1-2 μl genomic DNA)Assay PCR cycle conditions95°C/10 min x 1 cycle; (95°C/10sec, 60°C/45sec) x 40 cyclesMeasure fluorescence at the end of each cycleInterpreting Results and ExamplesBoth kdr assays use two probes, the first specific for the wildtype allele is labeled with VIC, and thesecond, specific for the mutant allele (kdr-w or kdr-e), is labeled with FAM. In either assay, a substantialincrease in VIC fluorescence indicates a homozygous wildtype, a substantial increase in FAMfluorescence indicates a homozygous mutant, and a, usually intermediate, increase in both signalsindicates a heterozygote (Figure 8.5.1.3.1-2). Individuals homozygous for the kdr-e mutation display noincrease in VIC or FAM fluorescence in the kdr-w assay and vice versa. To help score the genotypes, theRotor-Gene software allows endpoint fluorescence values for the two dyes to be automatically correctedfor background and plotted against each other in bi-directional scatter plots (Figure 8.5.1.3.3.).

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.3 Knockdown resistance (kdr) assaysPage 2 of 6Fluorescence100806040Cycling of FAM probe(phenylalanine)Rw/RwS/Rw20S/S5 10 15 20 25 30 35 40Cycle3530Cycling of VIC probe (leucine)S/SFluorescence2520S/Rw1510Rw/Rw5 10 15 20 25 30 35 40CycleFigure 8.5.1.3.1. Real-time TaqMan detection of kdr-w (L1014F)S: Wild type allele (L1014), Rw:Resistant allele, West African mutation (L1014F).

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.3 Knockdown resistance (kdr) assaysPage 3 of 66050Cycling of FAM probe (serine)Re/ReFluorescence4030S/Re2010 S/S405 10 15 20 25 30 35 40Cycle3530Cycling of VIC probe (leucine)S/SFluorescence2520S/Re1510Re/Re5 10 15 20 25 30 35 40CycleFigure 8.5.1.3.2. Real-time TaqMan detection of kdr-e (L1014S) S: Wild type allele (L1014), Re:Resistant allele, East African mutation (L1014S)

8070Mutant (S1014)Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.3 Knockdown resistance (kdr) assaysPage 4 of 6Heterozygous60FAM fluorescence (kdr-e allele)50403020100-10-20No template-10 0Wild Type (L1014F)10 20 30 40 50 60 70 80VIC fluorescence (wild type allele)Figure 8.5.1.3.3. Scatter plot analysis of TaqMan fluorescence data In this example real time PCR wascarried out using the east kdr assay on ~70 samples then fluorescence values of the FAM labeled probespecific for the kdr-e mutation were plotted against the VIC labeled probe specific for the wild type allele.Notes and TroubleshootingIf separate runs have been performed on the same samples for kdr-e and kdr-w, when interpreting theresults it is useful to have two copies of the real-time PCR machine software open on the computer atonce. In one copy open the kdr-w run and in the other open the kdr-e run. The same sample can thenviewed in each assay by moving between the two runs/copies of the software. This helps to rapidly assigneach sample a genotype. For example, sample one is examined in the kdr-W run and shows a signal inthe FAM channel and not in the VIC channel and when examined in the kdr-E run shows a signal inneither channel. Sample one is therefore scored kdr-W homozygous. Sample two is examined in the kdr-W run and shows an intermediate signal in the VIC channel but no signal in the FAM channel, whenexamined in the kdr-E run it shows an intermediate signal in both the FAM and VIC channels. Sample twois therefore scored kdr-E heterozygous. Alternatively if the east and west assays were run together on thesame samples the same approach can be followed using a single copy of the software.In most of our experiments using wild-caught mosquito specimens, we have found that interpreting theresults of the kdr TaqMan assays is relatively straightforward. However in some instances when using‘quick and dirty’ DNA extraction protocols, we have sometimes seen background signals which have

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.3 Knockdown resistance (kdr) assaysPage 5 of 6made assigning genotypes more difficult. Difficulty in scoring genotypes is often associated with very lowsignals due to the poor or low yield of DNA obtained.In our experience, distinguishing between Rw/Rw and S/Rw genotypes can seem less obvious thanscoring other genotypes because a Rw/Rw genotype shows a very low level signal in the VIC channelrather than a completely flat line (as with a no-template control). In addition the signal strength from theFAM and VIC probe can be different. Running controls of the two genotypes is very helpful and allows fora direct comparison with unknown samples. Further examples of RwRw and S/Rw genotypes (without theautoscaling) illustrating these points are included below in Figure 8.5.1.3.4.10080Cycling of FAM probe(phenylalanine)Fluorescence6040Rw/RwS/Rw2005 10 15 20 25 30 35 40Cycle10080Cycling of VIC probe (leucine)Fluorescence6040S/Rw20Rw/Rw05 10 15 20 25 30 35 40CycleFigure 8.5.1.3.4. Real-time TaqMan detection of kdr-w (L1014F). Blue trace, R/Rw: Resistant allele,Green trace S/Rw, Black trace, blank. Note the autoscale function was not used in this example and thesignal from the VIC probe is lower than the FAM probe. In addition note that the Rw/Rw signal in the VICchannel is above that of the blank shown in black.Use of Assay on single legsIn our experiments to date when we have run the assay on DNA extracted from a single leg we haveobtained mixed results (possibly due to the low quality/concentration of DNA extracted) and therefore wecannot recommend the TaqMan kdr assays for this approach.ReferencesBass C et al. (2007) Detection of knockdown resistance (kdr) mutations in Anopheles gambiae: acomparison of two new high-throughput assays with existing methods. Malar J 6:111

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.3 Knockdown resistance (kdr) assaysPage 6 of 6

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.4 Insensitive acetylcholinesterase (iAChe) assayPage 1 of 28.5.1.4 Insensitive acetylcholinesterase (iAChe) assayChris Bass, Martin Williamson, John Vontas, Hilary Ranson, Martin Donnelly and Lin FieldIntroductionThis assay detects the G119S mutation in the gene ace-1 which encodes the acetylchoinesteraseenzyme. This assay is suitable for use on real-time PCR machines that have two or more detectionchannels. A PCR based assay can be found in Chapter 5.3.4.Assay conditions96 48 1 Reagent500 μl 250 μl 5.0 μl sterile H 2 O1.0 ml 500 μl 10.0 μl SensiMix DNA kit (Quantace)160 μl 80 μl 1.6 μl primer ACE1-F (800nM) GGCCGTCATGCTGTGGAT160 μl 80 μl 1.6 μl primer ACE1-R (800nM) GCGGTGCCGGAGTAGA40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) Ace1G119 VIC-TTCGGCGGCGGCT40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) Ace1S119 6FAM-TTCGGCGGCAGCT1.9 ml 950 μl 19 μl Total (To each 19 μl reaction add 1-2 μl genomic DNA)Assay PCR cycle conditions95°C/10 min x 1 cycle; (95°C/10sec, 60°C/35sec) x 40 cyclesMeasure fluorescence at the end of each cycleInterpreting Results and ExamplesThe iAChE assay uses two probes, the first specific for the wild-type allele is labeled with VIC and thesecond, specific for the mutant allele (S119), is labeled with FAM. A substantial increase in VICfluorescence indicates a homozygous wild-type, a substantial increase in FAM fluorescence indicates ahomozygous mutant and a, usually intermediate, increase in both signals indicates a heterozygote(Figure 8.5.1.4.1).

AChapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.4 Insensitive acetylcholinesterase (iAChe) assayPage 2 of 2110105100959085R/R8075Fluorescence7065605550S/R45403530252015S/S1050110105B5 1 0 1 5 2 0 2 5 3 0 3 5 4 0Cycle100959085S/S80757065Fluorescence605550S/R45403530252015R/R10505 1 0 1 5 2 0 2 5 3 0 3 5 4 0CycleFigure 8.5.1.4.1. Real-Time TaqMan detection of G119S. A) Cycling FAM probe (Serine) B) Cycling VICprobe (Glycine).

8.5.1.5 Resistance to dieldrin (rdl) assayChapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.5 Resistance to dieldrin (rdl) assayPage 1 of 2Chris Bass, Martin Williamson, John Vontas, Hilary Ranson, Martin Donnelly and Lin FieldIntroductionTwo TaqMan assays have been developed, one for each of the two alternative mutations found in An.gambiae s.s. (A296G) and An. arabiensis (A296S). This assay is suitable for use on real-time PCRmachines that have two or more detection channels. An alternate PCR based assay can be found inChapter 5.3.3.A296G Assay conditions96 48 1 Reagent500 μl 250 μl 5.0 μl sterile H 2 O1.0 ml 500 μl 10.0 μl SensiMix DNA kit (Quantace)160 μl 80 μl 1.6 μl primer GlyRdlF (800nM) TCATATCGTGGGTATCATTTTGGCTAAAT160 μl 80 μl 1.6 μl primer GlyRdlR (800nM) CGACATCAGTGTTGTCATTGTCAAG40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) WT1 VIC-ACGTGTTGCATTAGG40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) Gly 6FAM-ACGTGTTGGATTAGG1.9 ml 950 μl 19 μl Total (To each 19 μl reaction add 1-2 μl genomic DNA)A296G Assay PCR cycle conditions95°C/10 min x 1 cycle; (95°C/10sec, 62°C/45sec) x 40 cycles. Measure fluorescence at the end of eachcycle.A296S Assay conditions96 48 1 Reagent500 μl 250 μl 5.0 μl sterile H 2 O1.0 ml 500 μl 10.0 μl SensiMix DNA kit (Quantace)160 μl 80 μl 1.6 μl primer SerRdlF (800nM) TCATATCGTGGGTATCATTTTGGCTAAAT160 μl 80 μl 1.6 μl primer SerRdlR (800nM) TCGTTGACGACATCAGTGTTGT40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) WT2 VIC-TTACACCTAATGCAACACG40 μl 20 μl 0.4 µl TaqMan MGB probe (200 nM) Ser 6FAM-CACCTAATGAAACACG1.9 ml 950 μl 19 μl Total (To each 19 μl reaction add 1-2 μl genomic DNA)A296S Assay PCR cycle conditions95°C/10 min x 1 cycle; (95°C/10sec, 60°C/45sec) x 40 cycles. Measure fluorescence at the end of eachcycle.Interpreting Results and ExamplesBoth rdl assays use two probes, the first specific for the wild type allele is labeled with VIC and thesecond, specific for the mutant allele (S296 or G296), is labeled with FAM. In either assay, a substantialincrease in VIC fluorescence indicates a homozygous wild type, a substantial increase in FAMfluorescence indicates a homozygous mutant and a, usually intermediate, increase in both signalsindicates a heterozygote (Figures 8.5.1.5.1 and 8.5.1.5.2).

Chapter 8 : Field Techniques8.5 Real-Time PCR Assays8.5.1 Vector Population Monitoring Tool using Real-Time PCR8.5.1.5 Resistance to dieldrin (rdl) assayPage 2 of 225A) Cycling FAM probe (Glycine)R/RFluorescence2015S/R105S/S5 10 15 20 25 30 35 40Cycle1510B) Cycling VIC probe (Alanine)S/SFluorescence5S/RR/R5 10 15 20 25 30 35 40CycleFigure 8.5.1.5.1. Real-time TaqMan detection of A296G6050A) Cycling FAM probe (Serine)R/RFluorescence40302010S/RS/S5 10 15 20 25 30 35 40CycleFluorescence4540353025B) Cycling VIC probe (Alanine)S/SS/R201510R/R5 10 15 20 25 30 35 40CycleFigure 8.5.1.5.2. Real-time TaqMan detection of A296S

Chapter 8 : Field Techniques8.6 Molecular Karyotyping PCR Assays8.6.1 Anopheles gambiae 2La inversion – White et al.Page 1 of 28.6 Molecular Karyotyping PCR Assays8.6.1 Anopheles gambiae 2La inversion (White et al.)Bradley White and Nora BesanskyIntroductionThe 2La inversion in An. gambiae is associated with tolerance to aridity and indoor resting behavior. Thisinversion may also be involved in the incipient speciation between the Mopti and Savanna chromosomalforms (White et al. 2007). Traditionally, polytene chromosomal banding patterns were used to differentiatebetween the 2La and 2L+ a forms. However, due to the complex procedures needed to produce these,few laboratories have trained personnel. In order to better understand the effects of karyotype onphenotype, a PCR-based method was developed based on the published sequence of An. gambiae. Thismethod has been shown to be robust in identifying the inversion within all members of the An. gambiaecomplex.PCR authentication for the 2La inversion in the Anopheles gambiae complex (White et al. 2007)Prepare PCR Master Mix for 96, 48 or 1 25μl PCR reactions. 1 Add reagents in the order presented.96 48 1 Reagent1185 μl 592.5μl 11.85μl Distilled H2O500 μl 250 μl 5.0 μl 5X PCR buffer200 μl 100 μl 2.0 μl dNTP (2.5 mM concentration)100 μl 50 μl 1.0 μl 23A2 (R, 25 pmol/ μl) CTC GAA GGG ACA GCG AAT TA100 μl 50 μl 1.0 μl 27A2 (F, 25 pmol/ μl) ACA CAT GCT CCT TGT GAA CG100 μl 50 μl 1.0 μl DPCross 52L+ (F, 25pmol/ μl) GGT ATT TCT GGT CAC TCT GTT GG200 μl 100 μl 2.0 μl MgCl2 (25 mM)15 μl 7.5 μl 0.15 μl Go-Taq DNA polymerase (5U/ μl)2.4 ml 1.20 ml 24 μl Total (To each 24 μl reaction add 1 μl template DNA)Table 8.6.1.1. F and R indicate forward and reverse orientationPCR cycle conditions94°C/2min x 1 cycle(94°C/30sec , 60°C/30sec , 72°C/45sec) x 35 cycles72°C/10min x 1 cycle4°C holdRun samples on a 2% agarose EtBr gel. (Figure 8.6.1.1).Primers create fragments of 492 and 207bp for 2La and 2L+ a arrangements, respectively.1 Amounts for larger master mixes have been adjusted upwards to be sufficient for 50 and 100 reactionsto compensate for imprecise measurements.

Chapter 8 : Field Techniques8.6 Molecular Karyotyping PCR Assays8.6.1 Anopheles gambiae 2La inversion – White et al.Page 2 of 2Figure 8.6.1.1 Lane 1 1kb ladder, lane 2 2La/a homozygous An. gambiae,lane 3 2La/+ a hybrid, lane 4 2L+ a /+ a homozygous.Troubleshooting:Lowering the annealing temperature (to 58°C or 55°C) may effectively improve PCR amplification undersome conditions.96 well PCR sample preparation templateReferencesWhite BJ et al. (2007) Molecular karyotyping of the 2La inversion in Anopheles gambiae. Am J Trop MedHyg 76:334-339

Chapter 9 : Obtaining Live Material9.1 Permits and RegulationsPage 1 of 4Chapter 9 : Obtaining Live Material9.1 Permits & RegulationsPaul HowellIntroductionObtaining insect vectors for research purposes is often a lengthy process involving the procurement ofpermits and the development of an adequate space to contain them. In the past 5 years, several rulesand regulations have changed concerning the importation of insect vectors into the United States as wellas transfer of vectors within the country. Permits are necessary before any species can be imported intothe United States; however, in most states permits are not needed in interstate transfers of vectors.California, Florida, Texas, and Hawaii often have more strict rules for importing exotic vectors that couldpotentially become established due to the hospitable climate in these states. Although there are noreports of a vector escaping from a laboratory into the wild in North America, the potential exists for anescapee becoming established as evidenced by the inadvertent introduction of An. gambiae into Brazil inthe early 20th century (Soper and Wilson 1943).Import and Export Permits (updated May 2010)USPHS 42 CFR Part 71.54 states (a) A person may not import into the United States, nor distribute afterimportation, any etiologic agent or any arthropod or other animal host or vector of human disease, or anyexotic living arthropod or other animal capable of being a host or vector of human disease unlessaccompanied by a permit issued by the Director. As of 2010, the CDC Etiologic Agent Import PermitProgram (EAIPP) states that they will only issue a permit for “living vectors containing, or suspected ofcontaining an etiologic agent”. Importations of live eggs from long-established laboratory colonies or deador preserved materials no longer require a permit from the CDC.Per the CDC EAIPP: To facilitate clearance of materials that do not require a U.S. PHS Import Permit, it isrecommended that each shipment of this material be accompanied by a signed statement, on officialletterhead, from the person responsible for the shipment of this material with the following information:(1) A description of the material;(2) A statement that this material meets one of the above criteria (e.g., this material is not known orsuspected to contain an etiological agent, host, or vector of human disease); and,(3) Verification that it has been packaged, labeled, and transported in accordance with all applicableregulations. Note that other permits may be required (e.g., USDA).In addition, as of 2007 a valid USDA-APHIS permit is needed in order to complete any internationalimportation. As of 2010 the USDA will no longer allow hand-carrying of any samples into the UnitedStates. All materials, either living or dead, will require a permit and must be shipped using a commercialcarrier service such as Federal Express. This is in response to a GAO mandate that all material enteringthe United States be sent using a commercial carrier so that there are physical records showing that thematerials have been received by the appropriate party.Information on these permits is given below. Shipments without a USDA-APHIS permit are subject tobeing held at customs for an extended period of time or returned to the sender depending on the carrierbeing used.

Chapter 9 : Obtaining Live Material9.1 Permits and RegulationsPage 2 of 4United States Public Health Services form CDC 0728Information on how to apply as well as PDF forms are available at:http://www.cdc.gov/od/eaipp/Unites States Department of Agriculture APHIS Permit VS 16-3A site visit by a USDA official may be required before issuance of a VS 16-3 permit. This site visit is notfree, fees are listed at the website as well as information on how to apply, fee schedule, and PDF formsare available at:http://www.aphis.usda.gov/import_export/index.shtmlRegulations on the transfer of insectsWhen importing or transferring materials, it is recommended that non-motile forms are shipped sincethere is little possibility of these forms escaping into the wild if the package is accidentally opened(Benedict 2003). Packaging information can be obtained from the International Air TransportationAssociation (IATA) under CR 61-66. Requirement 62 directly deals with insects used for researchpurposes (see addendum) and outlines packaging and labeling requirements necessary for shipments. Ifadult forms are to be sent, it is recommended that animals be triple packaged to ensure that if onecontainer is compromised, the others will contain the material inside. All packages should have thefollowing information prominently displayed on them: Shippers information, Recipients information, genusand species of all materials being sent and the quantity of animals being sent. Alternately, you can shipmaterials using the IATA Biological Material Packaging Instructions 650 (PI 650).To determine if materials can be transferred to your location, contact your local Department of Agriculture,Health, or Natural Resources. Some states may not allow the importation of exotic vectors that couldbecome established if they escaped such as Aedes albopictus or Ae. aegypti. International recipientsshould check with their respective Secretariats or Departments of Agriculture, Natural Resources, orwhichever body governs the importation of live animals. The European Union as well as Japan currentlydo not require import permits for insect vectors however some nations do. Other nations, such as Austria,may have increased local regulations at their respective quarantine stations which may cause delays inprocessing shipments. Certain documents should always accompany an international shipment, theseinclude: recipients import permit (if applicable), Shippers Export Declaration (SED), a commercial invoicestating what is contained within the package, and in some cases a letter stating that the material is free ofany known etiologic agent (Sanitary Certificate).Information at a glance:International Air Transportation Association (www.iata.org)This association develops guidelines used by most major airlines and international shippers for thetransportation of various materials including live animals. These guidelines are often used by the USDAand US Fish and Wildlife Services for enforcement issues.Information on shipping hazardous materials (research insects are included in this heading)https://iataonline.com/Store/Products/Product+Detail.htm?cs_id=9515%2D48&cs_catalog=PublicationsLive animal regulations (Requirement 62)https://www.iataonline.com/Store/Products/Product+Detail.htm?cs_id=9105%2D33&cs_catalog=PublicationsShipping RulesDomestic shippers have implemented several varying rules as to which types of insects can betransferred and to where. Below is a list of the 4 most common shippers in the United States and theirregulations as they apply to the transfer of insect vectors.

Chapter 9 : Obtaining Live Material9.1 Permits and RegulationsPage 3 of 4The United States Department of Transportation, as well as other agencies have created regulations thatpertain to the shipment of insect vectors, both infected and uninfected. The following regulations shouldbe viewed in order to help decide if an insect can be shipped. All of these regulations can be viewed athttp://www.gpoaccess.gov/cfr/index.htmlPublic Health Service 42 CFR Part 72 and 42 CFR Part 71-Foreign quarantine, importation of vectors.The USPHS CFR specifically deals with the direct importation and transfer of vectors of human disease.Department of Transportation 49 CFR Parts 171-178. These regulations pertain to the packaging ofHazardous Materials including specifications on types of containers and their proper use.Department of Agriculture 9 CFR Parts 102 and 120. These regulations are used by the USDA in order todetermine if the vector that is being imported can be a vector of animal disease. Although there areseveral other parts to this CFR, these seem to be the most applicable.Federal Express (www.fedex.com)FedEx allows the transfer of insect eggs, including mosquitoes, from institute to institute only without anyclearance. Clearance to ship adult insects, however, must be gained by contacting the PackagingDepartment (800-633-7019, option 5) and then your local Sales Representative (call Live Animal Desk at800-405-9052 to determine who is your local representative). The Sales Representative will then contactthe Legal Department to obtain a Waiver of Liability, once this is granted your institution is on file as beingpermitted to ship live adult insects.Information at a glance:Live Animal Desk 800-405-9052http://www.fedex.com/us/government/intl/termsandconditions.html#liveanimalsPackaging Department 800-633-7019 option 5United Parcel Service (www.ups.com)UPS does not currently allow the transfer of mosquitoes, defined as obnoxious insects on their website,even if they are classified as research insects.Information at a glance:http://www.ups.com/content/us/en/resources/prepare/guidelines/animals.htmlDHL Worldwide Express (www.dhl.com)DHL does permit the transfer of live insects (non-venomous) throughout the U.S. Importation into the U.S.must be accompanied by a USPHS Import Permit and 24 hour notice must be given to DHL Customs atthe airport stating where the package originated, what the contents are, is the material infected, andwhere the material originated from (wild or laboratory collected material). DHL can then expedite theshipment through customs once it arrives in the U.S.Information at a glance:Quarantine and Customs 718-995-0001http://www.dhl-usa.com/resources/Prohibited_Restricted_Commodities.pdfUnited States Postal Service (www.usps.com)The USPS will accept shipments of live animals only if cleared by their Expedited Services Office. Contactyour local District Representative to determine if they will accept your package. You can locate thenumbers for your local representative at the website address listed below.Information at a glance:

Chapter 9 : Obtaining Live Material9.1 Permits and RegulationsPage 4 of 4http://www.usps.com/send/waystosendmail/extraservices/specialhandlingservice.htmArthropod Containment Level (ACL) guidelinesThis document provides a wealth of information on insectary design, containment level designation, riskassessments, transportation of vectors, as well as several other issues dealing with operating aninsectary.http://www.liebertonline.com/doi/pdf/10.1089/153036603322163475ReferencesBenedict MQ (2003) Arthropod Containment Guidelines. Vector Borne and Zoonotic Diseases 3:63-98Soper FL, Wilson DB (1943) Anopheles gambiae in Brazil: 1930-1940. In. Rockefeller Foundation, NewYork

Methods in Anopheles Research - MR4

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