DOCUMENT RESUME ED 045 449 SE 010 47U AUTHOR Sudit, W. Daniel; Chamberlain, Roy W. TITLE Collection F, Processing of Medically Important Arthropods for Arbovirus Isolation. INSTITUTION National Communicable Disease Canter (DHEW), Atlanta, Ga. PUB DATE Jan 67 NOTE 36p. AVAILABLE FPCM Center for Disease Control, Arbovirus-Ecology Laboratory, 1600 Clifton Rd., Atlanta, Ga. 30313 (free) EDRS PRICE DESCRIPTORS EDRS Price MF-40.25 HC-$1.85 Biology, *Entomology, *Laboratory mechniques, *Medical Research, *Public Health, *Resource Materials, Science Equipment ABSTRACT The methods given for collecting, preserving, and processing mosquitoes and other archropods for isolation of arboviruses are those used by the National Communicable Disease Center. Techniques of collecting mosquitoes as they bite, using light or bait traps, and from their daytime resting sites are described and illustrated. Details of subsequent storage and processing for virus extraction and identification are included. A short section describes differences in procedure when ticks or mites are collected. (AL)
Transcript
DOCUMENT RESUME
ED 045 449 SE 010 47U
AUTHOR Sudit, W. Daniel; Chamberlain, Roy W.TITLE Collection F, Processing of Medically Important
Arthropods for Arbovirus Isolation.INSTITUTION National Communicable Disease Canter (DHEW),
Atlanta, Ga.PUB DATE Jan 67NOTE 36p.AVAILABLE FPCM Center for Disease Control, Arbovirus-Ecology
ABSTRACTThe methods given for collecting, preserving, and
processing mosquitoes and other archropods for isolation ofarboviruses are those used by the National Communicable DiseaseCenter. Techniques of collecting mosquitoes as they bite, using lightor bait traps, and from their daytime resting sites are described andillustrated. Details of subsequent storage and processing for virusextraction and identification are included. A short section describesdifferences in procedure when ticks or mites are collected. (AL)
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COLLECTION & PROCESSINGOF MEDICALLY IMPORTANT
ARTHROPODSFOR ARBOVIRUS ISOLATION
ay W. Imolai. iaDIA and ROI" W. ONAMS1RLAIN
U.S. DEPARTMENT OP HEALTH, EDUCATION, AND WELFAREPUBLIC HEALTH SERVICE
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PREFACEThis bu:letin was prepared in response to numerous requests for detailed information on
methods for catching, preserving and processing mosquitoes and other arthropods for isolationof orbeviruses. The methods given have evolved and been tested over the past kw years inthe Arbovirms Vector Laboratory, Arbovirus Unit, CDC. Failure to consider special methodsused by many eminent arbovirus workers from other areas does not reflect discredit upon thembut rather points ot.i the limitations of the present work.
Crateful acknowledgement is made to the many resourceful people, within and withoutCDC, who have directly or indirectly contributed to the development and testing of the methodsherein deter ibed.
MOSQUITOES
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COLLECTION AND PRESERVATION
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Collection with CDC light traps 1
Collection from daytime resting sites
Biting collections 10
Collecting with bait traps 10
PROCESSING MOSQUITOESIN THE LABORATORY 12
Facilities for processing field-collected materials 12
Identification and pooling itGrinding, centrifugation, and storage of suspensions 10
Animal inoculation and observation 1$
Harvesting mouse brains for passage 22
Grinding, centrifuging, and passing mouse brain material - 27
ARTHROPODS OTHER THANMOSQUITOESVERTEBRATE BLOOD AND TISSUE
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MOSQUITOES
COLLECTION ANDPRESERVATION
For arbovirus survey purposes, a greatmany species of mosquitoes must be takenalive in large enough numbers for significantvirus tests. A variety of collecting methodsmay be required to produce adequate samples,since no one method will yield all species.Once collected, the mosquitoes must behandled and processed carefully to preventthe loss of the virus they may contain.
Cillection with C013 light traps
Experience has shown light traps to bea productive means of collecting mosquitoes,both in consideration of numbers of indi-viduals captured and diversity of speciesrepresented. A miniature battery-operated light
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trap has been developed at CDC1/ (Figure I),adaptable for either a 4 or 6 volt battery.Because of its light weight and small electri-cal current demand, it is highly portable andcan be used advantageously in even the mostprimitive areas. It is used extensively inmany CDC field programs, supplemented whennecessary by other collecting methods.
Depending upon the size and ecologicaldiversity of the study area, and the number ofmosquitoes desired from a particular site asa sample for virus testing, the traps may beclustered only 50 to 100 feet apart (Figure 2)or may be widely separated over many miles.A total of 20 to 30 traps are operated effec-tively by a two-man crew if placement is inreasonably accessible locations. All equip-ment needed may be transported in a standardstation wagon (Figure 3 ). A man can easilyemu 6 small motorcycle batteries or 12 dis-mantled traps at a time into the field in light-weight cases supplied with shoulder straps(Figure 4).
Site selection for light traps is importantand improves with the experience of the col-lector. In generals the beat catches are madewhere the cover is good and the humidityrelatively high. Locations a short distanceinto the margins of woods or swamps are par-ticularly favorable; traps hung over openwater or in open pastures are, on the otherhand, usually leas productive. If a site failsto produce the expected number of mos-quitoes, judging from collections in othertraps in the area, the trap is relocated. Some-times a shift of only a few yards makes aconsiderable difference in the number ofmosquitoes attracted.
Care is taken to place the traps wherethey will be shaded from the morning sun, toIncrease the chance of mosquito survival.
The beat light trap catches are madeduring the dark of the moon or on overcastnights; the weak light of the CDC trap com-petes poorly with bright moonlight. For thisreason light - trapping is usually restricted to
Sudta, W. D. and Chombatloin, R. W., 1962. Battey*owed light imp, an Proeved Pale]. Mosq.Neal, 22:126-129.
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'LIGHT TRAPS
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the darkest half of the month unless supple-mented by other means of mosquito collection.
Rainfall during the night generally doesnot reduce the light trap catch; to the con-trary, intermittent showers appear to enhancethe catch. Recent studies have shown that acarbon dioxide supplement to the CDC lighttrap greatly increases the trap's efficiency 21A 1 or 2 pound piece of dry ice is wrapped innewspaper or aluminum foil (Figure 5) andsuspended immediately above or beside thetrap. Greater numbers of mosquitoes arecaught, the ,pecies spectrum increased, andthe use of the trap less restricted as to place-me*t and moonlight conditions. Also, if thetrap is set in late afternoon, diurnal speciescan be captured. fEWEISIr
The traps are visited as early as prac-ticable the next morning. The mosquito col-lecting bag containing the captured mosqui-toes is removed from the trap and the sleeveloosely lc.otted (Figure 5). The bag is thengently collapsed, labeled as to rite and date(Figure 1), and placed on edge in a styrofoampicnic chill chest containing Zwo frozen quart-sized refreetant cans (Figure Mosquitoescooled in this manlier r -n'ain inactive andalive for several hours, even In summer heat,permitting the pick-up of bags at widely scat-tered points prior to killing and freezing themosquitoo storage. It is important not tocrowd the .oliecting bags oy placing too manyin the el at, as the mosquitoes may be killed
2/ Newhouse, V. F,, Chomberloin, R. W., Johnston,.1. G., Jr., and Svdio, W. D., 1966. Use of dry leeto increase mosquito conches of the CDC miniatureTight trap. Moog. News, A (1): 30-35.
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by crushing and the inside bags are not suf-ficiently cooled. Fifteen bags is a maximumnumber to be cooled by the two refreezantcans in a 11/2 cubic foot chest.
The bags of mosquitoes aia taken to aconvenient work site (field lab, tourist court,or merely a shady place and station wagontailgate) for rough sorting and storage. Thebags are transferred to a portable (5 cubicfoot) styrofoam dry ice chest and allowed toremain there about 15 minutes to kill themosquitoes by freezing (Figure 9). The bagsare then carefully removed two or three at atime and allowed to warm up a few minutes tothaw the mosquitoes. When a bag is warmedsufficiently to open it without mosquito break-age, a gentle swinging in the air will hastenthawing of the mosquitoes (Figure 10). Ifthey are handled while frozen, the mosquitoeswill shatter. The sleeve of the bag is untiedand the thawed mosquitoes gently shaken outonto a white light-trap lid for sorting (Fig-ures 11, 12). They are quickly separated(within 10-15 minutes, maximum) from the"trash" insects by aspirator and forceps,placed in flat-bottomed shell vials (17 by55 mm) and tightly plugged with rubber stop-pers (Figure 13). The tubes receive adhesivetape labels giving the date and location, andare Maher sealed by wrapping the junctureof glass lip and stopper with several roundsof 1/2-inch waterproof adhesive tape (Figure14). If more than one tube is required for asingle trap collection, each is numbered (e.g.,1 of 3, 2 of 3, etc.) to simplify associatingthem later when being processed in the lab-oratory. A field work-sheet is made up at thetime of mosquito tubing, giving trap numbers,dates, and approximate number of mosquitoesin each catch. These data are helpful laterin making decisions as to pooling and identi-fication schedules.
The mosquitoes may be anesthetized orkilled by chloroform instead of freezing, pro-vided the chloroform is used conservativelyand the mosquitoes handled as quickly aswith the other method. After tubing, the mos-quitoes are immediately stored on dry ice inlabeled cylindrical ice-cream cartons (Figure15) and are later transported to the mainlaboratory. There they are stored in a mechan-ical freezer at -00° C until they are identi-
fied, suitably pooled, and tested for virus. Noevidence of virus lads has been noted in mos-quito specimens stored at this temperaturefor a year in the field tubes.
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Collection from daytime resting sites
Many species of mosquitoes can be col-lected from various daytime resting sites,including back-yard chicken houses, out-buildings, garages, crawl spaces beneathporches, in culverts, beneath bridges, and inhollow trees and stumpholes. Under some cir-cumstances, the inside of houses may be in-spected. Specially constructed structures,such as privy-like sheds or 1-cubic-foot blackor dark-red boxes, may be provided in plannedstudies. The resting collections are particu-larly useful for obtaining species which areonly weakly attracted to light traps, such asCu lex pipiens Linn. and C. quinquefasciatusSay, both of which are important urban vectorsof St. Louis encephalitis.
In making the collections, the insidewalls of chicken houses, garages or otheroutbuildings, close to the ceiling as well asnear the floor, should be examined for restingmosquitoes since varying temperature, humidi.ty and light conditions influence the restingposition of the mosquitoes. Old spider websare usually favorite sites. A flashlight isneeded for examining the darker portion ofthe site, and collections are most easily madeusing a mouth or battery-powered aspirator(Figure 16).
A battery-powered aspirator can be adapt-ed from a vacuum flashlight (vacuum lintbrush) by removing the brush from the portand attaching a mosquito-collecting unit.2/Such an aspirator greatly accelerates the col-lection of mosquitoes without the danger anddiscomfort of inhaling the contaminated dustusually encountered.
If the mosquito conditions in the areaunder study are not already known to the col-lectors, a general survey is first made tolocate productive collecting sites. Using alocal map, the area is divided into sectionsfor systematic inspection and collection fromfavorable resting places (Figure 17). In anurban epidemic area, two men are assigned toa section as a team, one to serve as driver,the other as map reader and guide. If thecity is large, several teams may be requiredfor coverage of all sections within a reason-able time. Some portions of an average city,such as well-kept residential blocks, canusually be dismiseed after cursory inspectionas being unfavorable for mosquito productionand harborage; the older sections and out-skirts, on the other hand, are generally morefavorable for mosquitoes and require moretime.
2/ Husbands, R. C., 1958. An improved mechanicalaspirator. Calif. Vector Views, 5: 72-73.
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Culverts and bridges (Figure 18) can belocated by examining the map for roads overstreams or ditches. Chickens can usually beseen or heard while driving slowly through asection. If none are seen, stopping in themiddle of a block and looking in back yardsor questioning residents may yield loca-tions of chicken houses (Figures 18 , 20). Per-mission must always be obtained from theresidents before entering a premise to col-lect mosquitoes.
During this general search, small numbersof mosquitoes may be caught in many differentcollecting sites. To test these most econom-ically in the laboratory, all mosquitoes cap-tured in a section may be pooled and labeledwith the names of two streets which intersectcentrally in the section. Larger collectionsof mosquitoes may be labeled with a specificaddress (Figure 21 ). Well distributed exam-ples of these better sites can serve as routinecollecting stations to be revisited periodical-ly to reveal changes in virus activity andfluctuations in the mosquito population. The
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number of such permanent stations and thefrequency of visits to them are determined bythe over-all plan of operation. Usually it isbest to delay collecting around human habita-tions until 8:00 a.m. to avoid disturbing theoccupants. Collecting becomes progressivelyless profitable beyond 3:00 or 4:00 p.m. sincethe mosquitoes begin to disperse by that time.
The mosquitoes collected are transferredto small cages made from one-pint cylindricalice cream cartons by covering one or bothends with fine meshed netting (Figure 22). Acork-stoppered hole in the side permits ac-cess. The cages 07 covered with a dampenedtowel and transpo to a temporary labora-tory. There the mosquitoes may be held alivefor 24 hours for partial digestion of bloodmeals to permit inactivation of antibodywhich might be contained, or they may bekilled immediately by freezing on dy ice.After killing, they are sealed in labeled flat-bottomed vials as for light trapped specimens,at approximately 60° C until they are identi-fied and tested.
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Biting Collections
Some day-biting mosquito species arenot caught in light traps in sufficient numbersto make up adequate samples for virus tests,nor are they found readily in daytime restingsites. The same is true of some crepuscularand night-active mosquitoes. In these caseshuman biting collections may be profitable.Most conveniently, two collectors work to-gether. Sleeves and trouser legs may be rolledup, or shirts removed if necessary. The mos-quitoes attracted should be captured quicklyby aspirator before they have an opportunityto feed (Figure 23 ). The individual found tobe the more attractive to the mosquitoesserves as the principal bait while the otherdoes the bulk of the collecting. A large ani-mal such as a cow or horse may serve as aneffective attractant.
Collecting periods of 15 to 30 minutesper station are generally adequate for routinesurveillance during the day or after dark.During the dusk period, however, a singlesite is usually worked from about a half-hourbefore sunset until 30 to 45 minutes aftersunset since some species bite avidly foronly a short time during this period and mightbe missed if the collectors are traveling be-tween stations.
The mosquitoes caught at each collec-tion site or cluster of associated sites areplaced in labeled pint-carton cages untilthey can be killed by freezing or chloroform,sealed in tubes, and stored on dry ice fortransport to the processing laboratory.
Collecting with bait traps
A wide variety of mosquito bait trapshas been devised and used in many parts ofthe world. The basic principle of most ofthese traps is the same, i.e., the mosquitoesenter an ingress baffle to get to the attractant(usually a live animal) but are unable to findtheir way back out again. Bait traps areusually more selective than light traps, at-tracting some species of mosquitoes but notothers. This is partly due to the host prefer-ence of the mosquitoes and partly to a re-luctance of some species to enter the baffles.Bait traps are commonly used in host prefer-ence studies but are also valuable for supple-
menting other methods of collecting or forselectively capturing large numbers of certainspecies.
Some bait traps, called stable traps, areshed-like and large enough to house an ani-mal the size of a horse or cow. Figure 24shows a stable trap with a special catchingchamber mounted above the entry slot on eachside to prevent feeding upon the horse. It
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is a more common practice, however, to per-mit the mosquitoes to enter into the mainbody of the trap; they are then collected eachmorning from the walls with an aspirator(Figure 25).
An effective trap for use in conjunctionwith a sentinel chicken pen is described byRainey et al 4/ A slotted, rectangular capturechamber is affixed to the rear side of thechicken roosting area and captures the mos-quitoes upon entry (Figure 26).
Many small types of traps have been de-vised which are easily portable and can bebaited with various small animals rangingfrom birds and chicks to reptiles, rodents,and rabbits. Sometimes dry ice as a source ofcarbon dioxide, serving as an animal simu-lant, is used. An effective and convenientsmall type is represented by the lard-can trapof Bellamy and Reeves 5/, Which has beenused successfully in many field operations.It is a large lard can with a screen cone lead-ing inward from each end. A side door isprovided for insertion of a bait animal (Fig-ure 21). It is hung in a horizontal positionProm the limb of a tree (Figure 28). The de-sign may be variously modified to expose the
qt to mosquito bites or to protect it byscreening.
Ai Rainey, M. B., Warren, G. V., Hess, A. D. and Black-more, J. S., 1962. A sentinel chicken shed and mos-quito trap for use in encephalitis field studies.Mosq. News, 22: 337-342.
Bellamy, R. E. and Reeves, W. C., 1952. A portablemosquito bait trap. Mosq. News, 12: 256-258.
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Removal of captured mosquitoes from alard-can trap may be accomplished in severalways, but one especially applicable to fieldconditions is as follows: First remove thebait animal through the side door; then standthe trap on end, lid uppermost. With a house-hold hand sprayer of the insecticide type,spray chloroform into the top of the can insufficient quantity to anesthetize the mos-quitoes. Next, flip the can over endwise andbump it against the ground, lid end down.This jars the mosquitoes loose from thescreen and onto the peripheral area of thelid. The can is then lifted off the lid, andthe mosquitoes aspirated (Figure 29) andplaced into pint carton cages for recoveryand later processing. If a mouth aspirator isused, be certain that the chloroform fumeshave dispersed before drawing up the mos-quitoes.
PROCESSING MOSQUITOESIN THE LABORATORY
Facilities for processing fieldcollectedmaterials
One of the problems of greatest concernin the processing of field-collected mosqui-toes in the laboratory is to eliminate faultytechniques or conditions which might causecross-contamination. To solve this problemat CDC a separate laboratory is used ex-clusively for the sorting, identification, grind-ing, centrifugation, and storage of the fieldspecimens. No materials known to be virus-infected are permitted in this room. The basicequipment provided, used for no other pur-pose, includes a refrigerated centrifuge, re-frigerator, 60° C mechanical freezer, dis-secting microscopes and a chill table forsorting mosquitoes.
All virus work on virus strains isolated,such as passages and virus identificationprocedures, is done in separate facilities.Even then, field isolates are not handled con-currently with known infected experimentalmaterial.
This same caution applies to animalrooms in which the suckling mice are in-oculated and kept. Preferably "primary"inoculations (those of field suspensions) areplaced in rooms separate from "passage"inoculations, which of course are much morelikely to be infected with virus. Mice inocu-lated experimentally with known viruses arenever placed in the same room with fieldmaterial but are relegated to "hot" areas.
Identification and pooling
Mosquitoes must be accurately identifiedby species before they are tested. Since it isneither economically feasible nor necessaryto test each mosquito separately, they arepooled- by species. But first, before identifi-cation is commenced, a decision is made asto which collections are closely enoughassociated in space and time to be groupedtogether for sorting and pooling purposes.This decision is based mainly upon the dis-tances between collecting sites, the ecologi-cal differences or similarities in the varioussites, and the degree of precision desired in
determining the time of mosquito infection,should viruses be found present. Generally,collections from closely associated sitesover 1- to 2-week collection periods can begrouped without serious loss of specific in-formation. This grouping is done in the inter-ests of economical pooling of the individualspecies for testing. When several collectionsare lumped together, more of the mosquitopools, particularly of the minor species, canbe made up tc maximum size. Since each poolis inoculated into a litter of suckling micevalued at approximately $1.50, conscientiouspooling is not only advisable but usuallyan absolute necessity.
The number of mosquitoes making up in-dividual test pools may range from a singlemosquito, if that happens to be the onlyspecimen of a particular species taken, to amaximum of approximately 100. The maximumpool size at the Communicable Disease Cen-ter is arbitrarily set at 100 for average sizedmosquitoes and fewer for the large mosqui-toes such as Psorophora confinnis (L.A.)and P. ciliate (Fab.). Very small mosquitoessuch as the Uranotaenia species are proc-essed in pools of up to 200 each. The maxi-mum figure is flexible and can be loweredif a high field infection rate is expected. Forexample, if as many as 1 of each 5 pools of100 of a particular species are found infected,the pool size is reduced to 50 in subsequenttests to decrease the possibility of doubleinfections.
Males are generally discarded unlesssaved for terminalia examination to confirmidentification of as so -Rated females.
The mosquitoes to be sorted are removedfrom the freezer one or two tubes at a timeand thawed in the hand or by laying on atable top for a few minutes. They are thenspilled out onto the inverted lid of a glass14.5 cm Petri dish lined with a single thick-ness of slightly moistened filter paper. Thedesired degree of moistness is obtained bywetting the paper with water, pouring off theexcess, then blotting as dry as possible withfacial tissue or a snft towel. At CDC thedish of mosquitoes is kept cold during sort-ing by a specially designed chill table, thesurface of which is provided with a refrig-eration unit s/ In the absence of such re-frigerating equipment, however, a suitable
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cold surface can be prepared by placing a1-in. thick slab of dry ice in a shallow trayabout 16-in. square. The surface of the dryice should be insulated with a cloth toweland enough thicknesses of paper toweling toprevent refreezing of the specimens; the sur-face temperature of the mosquito dish shouldbe approximately 3° to 5° C. Flatsided quart-sized frozen refreezant cans or pliofilm bagsof cracked water-ice can also be used assorting surfaces. When using the cracked ice,it is advisable to provide each bag with avent tube to prevent ballooning as the icemelts.
The mosquitoes are sorted according tospecies with fine-tipped forceps into smallerPetri dish halves (55 to 60 mm diameter),similarly lined with damp filter paper andconveniently arranged semicircularly on thecold surface around the large sorting dish(Figure 30 ). To speed up the mosquito han-dling, the initial sorting is done mostly bynaked eye by skilled technicians, trained todistinguish the different species by mac-roscopic characters. Periodically, whenever
Sudlo, W. D., Chamberlain, R. W. and Collier, M.,1965. The CDC entomological chill table, a re-frigeroted unit for us, in processing mosquitoes forvirus Isolation studies. Mosq. News,25(4):385-389.
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a maximum pool size is reached, mosquitoesare listed on an Arthropod Pooling Recordsheet (Figure 31), transferred to a clean, un-lined, 10 cm Petri dish, and the lid given acode number with glass-marking ink or a softwax pencil (Figure 32). These dishes areaccumulated in a refrigerator at approximately4°C for up to 4 hours until their contents arechecked for accuracy against the poolingrecord sheet by an entomologist (Figures 33,34)."This checking is done with a dissectingmicroscope, the stage of which is kept coldby resting it upon a chilled surface.
The pooling record sheet provides forrecording the unfed (non-engorged), gravid,and blood-engorged (red or black) specimensseparately, although they may be ground upin the same pool. However, the abdomens ofthe more freshly engorged (red) specimensare removed with a clean, sharp scalpel,placed in labeled tubes and saved for laterprecipitin testing to determine the host fedupon. The remaining portions of these mos-quitoes are returned to the pool. If the num-bers of freshly engorged mosquitoes aregreat, or if their abdomens are not removed,they are tested in separate pools which aresmaller than those for the other feeding cate-gories. This is done to minimize the possibil-ity of neutralizing virus of an infected mos-quito pool by antibody which might be presentin the host blood.
The importance of keeping the mosqui-toes chilled while sorting cannot be toostrongly emphasized. Best knowledge indi-cates that most arboviruses are highly un-stable in dead mosquitoes at room tempera-ture. In view of the considerable effort andexpense of obtaining .and testing the speci-mens, one should not take chances of losingeverything by careless handling. Not only willvirus be lost, but an erroneous picture of theinfection rate in the survey area will bo ob-tained which may lead to epidemiologic mis-interpretation.
The importance of double-checking theidentifications must also be stressed. Re-gardless of the skill of the initial sorter,mechanical errors invariably occur. Even ifa single person is responsible for all mos-quito identifications, he should check his ownpoolings prior to their being ground for test-ing. The seriousness of misidentification may
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be extreme, as a virus isolation from a mis-identified species may send yotr own orother research groups after faulty leads con-cerning vector-virus-host relations.
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Grinding, centrifugation, and storage ofsuspensions
A supply of sterile diluent is made upeach day; however, any left over can beplaced in the refrigerator and used again thenext day but is not kept beyond that time. Itconsists of 25 percent normal rabbit serumin 0.05 M phosphate-buffered distilled water,pH 7.6 to 7.8, and contains 1.6 mg of strep-tomycin sulfate and 1,000 units of sodiumpenicillin 0 per ml. The high proportion ofserum gives this diluent excellent virusstabilizing properties. The formula for itspreparation is as follows:
Penicillin working stock: To 1,000,000unit bottle, add 10 ml 0.05 M phos-phate buffered water (pH 7.8) to pro-duce working stock containing 100,000units ml.
Streptomycin working stock: To 5.0 gmbottle add 12.6 ml buffered water (pH7.8) to yield a 400 mg/m1 concentratedstock. To prepare working stock con-taining 40 mg /ml, add 54 ml bufferedwater to 6.0 ml of the concentratedstock.
For 26 percent normal rabbit serum dil-uent:
26.0 ml normal rabbit serum1.0 ml penicillin (100,000 units /ml)4.0 ml streptomycin (40 mg /ml)
70.0 ml buffered water (pH 7.8)
100.00 ml yield
A double set of labels are prepared foreach pool using Si-in. adhesive tape and aballpoint pen with waterproof Ink. Time canbe saved by using a tubber stamp for theparts of the labels which are repeated. Thelabels are placed on sterile, corked Kahntubes, one pair per tube, and the tubes linedup in order in a rack. The corks are removedand placed temporarily into a clean Petridish. With an automatic pipette, 2 al of the25 percent rabbit serum diluent are dispensedinto each of the tubes, and the corks replaced(Figure Si ). The rack of tubes is then placedin a shallow pan of ice.
Five Petri dishes of the identified mos-quitoes are removed from the refrigerator at atime. These are lined up in order on a towel-covered work table, about 6 inches apart; infront of each a chilled, sterile 3-in. G. D.mortar is placed..Then, starting from left toright, the mosquitoes are dumped into theirrespective mortars and the labeled lids againrealined with the mortars to prevent confusionas to identity (Figure 38). Any mosquitoes
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which adhere to the dish are dislceged with asterile applicator stick, which is then dis-carded. Two or three drops of diluent aredispensed by dropper bottle into each mortar(Figure 31), followed by a small amount ofpowdered Alundum* as abrasive (Figure 38).In order, each pool is ground well in thesmall amount of diluent to make a smoothpaste. Do not short-cut the grinding. Thenthe 2 ml of diluent are dumped from the firstKahn tube into the first mortar, the labelchecked against that on the Petri dish lid toavoid error, and the grinding continued (Fig-ure 39). When a smooth suspension is ob-tained, it is poured back into the same Kahntube (Figure 40), which is replaced in theiced rack. This procedure is repeated untilall of the suspensions are ground. The sameamount of diluent (2 ml) is used for eachpool, regardless of whether it contains aminimal or maximal number of mosquitoes.The 2 ml volume is convenient to work with,neither restricting reinoculations nor undulydiluting the virus in an infected pool.'Use of trode nomes is for identificotion only and doesnot constitute endorsement by the Public HeolthService or the U.S. Deportment of Health, Educotion,and Welfore.
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18
The above method of predispensing thediluent into the Kahn tubes eliminates thedanger of contaminating the entire diluentsupply, as can happen when the diluent ispipetted directly to each mortar from a com-mon flask at time of grinding.
The suspensions are spun for 30 minutesat 3000 RPM (1700 X g) in a 60-place anglehead in a refrigerated centrifuge (Figure 41).At completion of centrifugation each super-nate is poured into a clean corked Kahn tube(Figure 42) and the two labels transferred.Use caution in handling the stoppers or thefingers may become contaminated; never touchthe lip of a tube. The tubes are then placedin pint-sized cylindrical ice cream cartons,the lids of which are labeled as to con-tents, and stored in a mechanical freezer at60° C until the suspensions are inoculatedintracerebrally into suckling mice (Figure43). If preferred, the inoculations can beperformed immediately after the centrifuga-tion, before storage. However, there is noevidence that the extra freeze-thaw decreasesin appreciable degree the virus titer of an in-fected suspension, provided the 26 percentserum diluent is used. Also, suspensionshave been stored fot a year with little or noindication of virus loss.
elN.
Animal Inoculation and obi erwation
Suckling mice are used for the initialisolation of arboviruses from the mosquitopools, in the present absence of a singletissue culture system approaching their broadsusceptibility. Wean ling mice, although satis-factory for some srboviruses, fail to demon-strate the broad susceptibility required forgeneral arbovirus survey purposes, Pregnantfemale mice are placed in separate mousecans when they are nearly ready to litter andare checked each morning. The new littersand their mothers are dated and set aside as"zero days old," to be used on day 1 or day 2.They can be inoculated on the day of birth,but the chance of their being destroyed bytheir mothers is somewhat increased. Theyare also sometimes used when three days ofage; however, the resistance of mice to someof the arboviruses increases with age, so thispractice is ordinarily reserved for Mondayinoculations, to use the litters remaining fromthe venous Friday. Mice older than threedays are never used to attempt field isola-tions where the type of virus to be encoun-tered is unknown.
Each litter is culled to six baby mice(Figure 44). One of the tube labels is placed
19
on the mouse can (Figure 45). The -nice areeach inoculated intracerebrally with 0.02 mlof the mosquito suspension, using a 'A -mltuberculin syringe and 'A-in. 27-gauge needle. Aseparate syringe and needle is used for eachmosquito pool. Syringe filling (Figure 46)
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20
and the mouse inoculations (Figure 41) areperformed over the mouse can to minimizecontamination of the table area. Special cau-tion is observed to avoid touching the tubelip, plunger shaft, or needle hub since cross-infections from one litter to the next mayoccur during inoculation if the hands be-come contaminated. The hands are bracedagainst the edge of the can for steadiness.A disposable pliofilm glove is worn on thehand which holds the mice during inoculation,The tubes of mosquito suspensions are keptin a rack in a shallow pan of ice during theanimal room operations.
The inoculated mice are generally placedback into the can with their own mother; how-ever, some technicians pool all the young andplace them with mothers at random after in-oculation with good success. In fact, themothers of litters up to a week old may beused by discarding their own young and re-placing them with 1- or 2-day-old mice thinnedfrom other litters.
The teat mice are checked each morningfor 12 to 14 days, using 10-inch forceps (Fig-ure 411). The forceps are alternated frequent-ly with another pair in a jar of disinfectant,especially when an apparently infected mouseis touched. The disinfectant is blotted offwith a paper towel before use. The mousecount and symptoms are recorded on an Ani-mal Inoculation Record sheet, using stand-ardized abbreviations: 14, missing; E, eatenby mother; D, dead; Pa, paralyzed; Pr, pros-trate; and Co, convulsing. Additional designa-tions may also be helphil, such as Tw (twitchy),Wo (wobbly), and Pa? (questionably paralyzedor with indefinite symptoms, to be observedclosely on future chectinga). When symptomsbegin to occur, it is wise to check the miceagain in the late afternoon. A dash () in thechecking column indicates normality.
A sample Animal Inoculation Recordsheet is shown in Figure 41. These are keptIn loose leaf binders in page order. For con-venience, inoculation sheets of field suspen-siona we kept in a "Primary" book; those ofpassages are kept in a separate "Passage"book.
The dead mice and those showing defi-nite signs of infection are saved for passageor later reference. llsaally the mice withsevere symptoms, such as general paralysis
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or prostration, and those very recently dead,contain more virus in their brains than micewith lesser symptoms. The sick mice arekilled by brief exposure to chloroform fumesin a killing jar, then are placed in suitablylabeled end-flap 3- x 6-in. manila envelopes.The label indicates the mouse involved,symptom, date, and page in the record book(Figure 50). The mice saved are stored indate order in a mechanical freezer at 60° Cuntil processed further.
Strict maintenance and sanitation prac-tices are obnerved in the animal rooms at alltimes. Ample food and water is constantlyavailable to discourage the mothers from eat-ing their own young. At the close of the ob-servation period, all surviving mice are killedby chloroform. Then they, their bedding ma-terial, and any left -over mouse food are in-cinerated, and the mouse cans and waterbottles are sterilized by autoclaving.
Novelties am. Niles he passage
Periodically, usually at weekly intervals,the Animal Inoculation Record sheets areinspected, and mice are selected from theReeser for brain passage into other sucklingmice. A serial code dumber (passage number)is given each mouse selected, and is record-ed both on the Animal Inoculation Recordsheet aid the mouse envelope. At the same
time a passage listing is made out on aPassage List sheet (Figure 51), associatingthe passage number with mosquito suspen-sion, mouse source, and record book pages.Columns are niso provided for recordingresults of sterility teats on brain suspen-sions. This list is valuable in trackingdown the laboratory history of material testedmince it shows in one place the source, dis-position, dates of handling and associationwith other materials.
Generally all single, scattered "deeds"are passed unless the mice are obviouslydecomposed or have died of some cause easi-ly recognized as foreign to arboviruses. Ifmore than one mouse in a litter dies, in apattern consistent with virus infection, usual-ly only a single mouse of the group is passedand the remainder held until the outcome isknown. If the first mouse passed is negative,a second mouse is passed, provided thesymptom or day of death was different fromthat of the first mouse.
The goal is a clear-cut positive or nega-tive result from each mosquito pool inocu-lated, but frequently a fully qualified negativeteat is difficult to obtain. Ideally, a pool isconsidered negative if all mice inoculatedsurvive for the full checking period, or whendead mice are involved, they are passed withnegative results. In many instances, however,non-specific deaths way have occurred soearly in the majority of mice in an inoculatedlitter as to cause the test to be consideredunlitlefactorY, in which case the originalsuspension must be reinoculated.
Also, the mother mice may eat some oftheir young. This is usually not critical ifonly one or two are eaten on the first day ofinoculation; four survivors past the first dayare generally considered a legitimate test&Inc*. few arboviruses kill suckling mice with-in one day. When missing mice occur at laterincubation periods, however, or when an arbo-virus is suspected which kills within one day,a strong possibility exists that they wereeaten because they were sick. These suspen-sions must be reinoculated.
Some suspensions appear to be toxic,killing the mice on the first day. If this re-peats on second inoculation, and passagefails to indicate presence of a virus, thematerial should either be diluted 1:10 fat
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reinoculation or placed in hamster kidney orduck embryo tissue cultures. It may also beinoculated into suckling mice by the sub-cutaneous or intraperitoneal routes, sincemost arboviruses which kill by the intra-cerebral route are also infectious when inocu-lated peripherally. Mice are less subject totoxicity of materials inoculated peripherally.
The positive pools are more easily rec-ognized, since one or two passages generallyresult in all mice dying within two or threedays of each other. The brains of some ofthese mice are used for identification bycomplement fixation, hemagglutination, andneutralization tests. Some are also saved asstock for future reference. When a mosquitopool has been found to contain virus, theoriginal suspension is generally reinoculatedto confirm the validity of the isolation.
It is during the passaging procedure thatlaboratory contamination and serious errorare most likely to occur. Infected sucklingmouse brain tissue often contains very highconcentrations of virus; titers of 109.0 withsome viruses are not unusual. This meansthat the material could be diluted a billionfold end still contain enough virus to killsuckling mice. In handling viruses in suchconcentration, considerable caution must betaken to prevent risk to the technician andcross-contaminations between different brainspecimens. Carelessness during the prepara-tions can result in apparent "isolations"from associated non-infected suspensions,leading to considerable contusion and seriousmisinterpretations.
Manipulations should be curled out ina properly e;hausted hood. Dissecting instru-ments (curved iris scissors and 8-In. thumbforceps) are sterilized by boiling for 20minutes in an Instrument sterilizer. Two setsate used for each mouse, one to remove theskin from the cranium and the other to takeout the brain. The method described belowcan be used for either suckling or weanlingmice.
The frozen mice are partially thawed,dipped in 2 percent Lysol solution, blottedbriefly on paper toweling, and pinned to asoftwood dissecting board covered with apaper towel and a piece of butcher parchment.Map pins ate used to fasten the mouse down,one placed through the bridge of the nose and
another through the base of the tail. Thescalp is removed with the .first set of instru-ments (Figure 52). These instruments are thenwiped on a gauze pad and stacked in a jarof 2 percent Lysol disinfectant, to awaitlater sterilization by boiling. The skull capand adjacent skin are swabbed with 1:1000Merthiolate and the swab-stick discarded(Figure 53 ). A drop or two of acetone arethen delivered from a height of about 2 inchesby dropper to wash away the excess Merthio-late and to cause rapid drying of the skullsurface (Figure 54). It is dropped from thisheljht to minimize chance contamination ofthe dropper tip, since the same one is usedfor each mouse. A second set of instrumentsis then used to cut off the skull cap and takeout the brain (Figure 55). It is advisable tohave the brain only partially thawed or itwill be too viscous to handle. Freshly killedmice of course pose no problem in this to-sped,. The brain is transferred to a labeled,sterile, corked, flat-bottomed shell vial (17X 55 mm) (Figure 56); then a culture is madeby immersing the tip of the scissors into atube of thioglycollate broth and wiping againstthe side of the tube. These scissors andforceps are then also set aside to be steri-lized. The label on the tube is checkedagainst that on the envelope from which themous,' was taken, since confuaion of labelsis a serious source of error.
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The pins are removed and placed in acontainer of Lysol solution. The mouse car-cass is wrapped in the butcher parchmentand paper towel and placed in a plastic bagfor disposal. The mouse board receives a newset of covering papers, and sterile instru-ments are used for the next mouse.
The vials containing the mouse braintissue are accumulated in a refrigerator dur-ing the dissection period. These tissues arethen either ground immediately into sus-pensions or are frozen at 60°C to be groundat a later date.
Another method can be used for sucklingmouse brain harvesting which is considerablyspeedier but which requires special cautionto preclude dangerous aerosols. Antibiotic-treated 25 percent rabbit serum diluent isdistributed in 2m1 amounts into sterile corkedKahn tubes. These tubes are appropriatelylabeled, placed in a rack, and the rack is setin an ice bath in a safety hood (Figure 51).
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26
Frozen suckling mice are allowed tothaw completely. With forceps, each mouse tobe harvested is placed belly down, head awayfrom the operator, upon a dissecting boardcovered with a sheet of butcher parchment.A 3-in. length of 4- or 1-in. adhesive tapeis placed transversely over the mouse tohold it firmly down onto the paper; the lead-ing margin of the tape should be across theshoulders, leaving the head free. The mouse'srecord numbe is double checked against thenumber of the previously labeled tube. Thenthe head and neck are swabbed thoroughlywith 1:1000 Merthiolate and the swab dis-carded. Using a disposable 1.0 ml syringeand a 19 or 20 gauge 1-in. needle, the cra-nium is pierced from its posterior aspect and0.1 to 0.2 ml of the viscous brain materialdrawn up (Figure 58). Disposable syringesare used for seater vacuum. The syringe isinserted into the labeled tube but the con-tents are not yet discharged. Instead about0.6 ml of the antibiotic-treated diluent isslowly drawn into the syringe along with thebrain material (Figure 59). Then, holding theneedle below the leeel of the remainingdiluent, all the material in the syringe isslowly ejected a single time. No additionaldrawing-up and discharging should be donesince du:knolls aerosols will be produced.The syringe and needle are discarded. Alwaysuse a well ventilated hoed. A gentle shakingafter replacement of the cork will assure
adequate mixing (Figure SO ); if virus causedthe mouse's death, it wit, be present in suchhigh concentration that better mixing is notrequired. The suspension is now ready forcentrifugation, described blow. Where cul-tures are desired, these are made by insertingthe needle tip into a tube of thioglycollatemedium immediately after the brain harvest-ing, before dilution.
Grinding, centrifuging and passing mousebrain material
Each dissected mouse brain is trans-ferred to a cold, sterile, 3-in. 0. D. mortarwith a sterile applicator stick (Figure 61) andground in 2 ml (for suckling mouse brain4 or3 ml (for weanling mouse brain) of the 26percent rabbit serum diluent to make an ap-proximate 10 percent brain suspension.(Grinding is not required for suckling mousebrains harvested by the syringe technique).Each suspension is poured into a sterile.corked, double-labeled Kahn tube and centri-fuged in the cold for 20 minutes at 2500 RPM(1100 X g) in an angle-head centrifuge. A sep-arate centrifuge from the one used to spindown the mosquito suspensions is employedto minimize the chance of accidental con-tamination of primary material. The super-nates may be poured off into clean corkedKahn tubes and the labels transferred; how-ever, for routine passages, this step may beskipped. The brain tissue forms such a tightpellet at the bottom of the tube that the clearfluid above may be drawn up into a syringefor inoculation without unduly disturbing it.The suspensions are either inoculated im-mediately by the intracerebral route intolitters of suckling mice or are frozen at 600C to be inoculated later. If a high proportionof the passages are expected to be viruspositive, it is a good practice to intersperseseveral normal brain suspensions in machinoculation series as a control against care-less handling and cross-contamination..
27
Daily mouse checks are mad as before.In the event of apparent arbovirus isolations,freshly dead, prostrate, or paralyzed miceare saved and brains harvested for virus iden-tific ation procedure 3.
In a busy laboratory a considerable num-ber of arbovirus isolations may be made fromseveral different field areas concurrently,and following the results of the variousinoculations may become cumbersome. Thevirus isolation progress form shown in Fig-ure 62 provides for convenient recording ofdata as it develops.
ARTHROPODS OTHER THANMOSQUITOES
The methods of collection of variousother medically important arthropods are manyand varied and will not be discussed here.However, the same principles expounded forhandling the mosquitoes apply: keep thespecimens alive or keep them frozen at dryice temperature.
Some arthropods, particularly ticks, willoften be engorged with vertebrate blood whencollected. If the blood is partially digested,there is no laboratory problem. lf, however,the blood is fresh, it may still contain someactive arbovirus antibody of the host andcould conceivably inactivate virus in the poolduring the grinding process. It is thereforerecommended that freshly engorged arthro-pods be held alive until partial digestion oc-curs, to denature any arbovirus antibodypresent. If this is not possible, they shouldbe tested in smaller pools than ordinarilywould be the case, separate from those notcontaining red blood.
With some arthropods, particularly mites,bird lice and immature ticks, accurate identi-fication is impossible without specially pre-pared mounts. In these instances, separationas to different kinds is made to the best ofone's ability under a dissecting microscope,and a 10 percent sample of each of these kindsis preserved for identification purposes.
The pool site may vary according to theexpected infection tate and the site of thearthropod. If the infection tate is high, thepool site must be reduced to preveat doubleinfections is single pools. If the arthropod is
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large, only a small number may be ground in2.0 ml of diluent. It may be advisable in someinstances when the arthropod is large to in-crease the amount of diluent per pool to 5.0ml. Although an occasional weak virus in-fection may go urdetected because of theadditional dilution, the number lost wouldnot, ordinarily be great enough to seriouslyaffect the significance of survey data.
The methods already described for lab-oratory processing of mosquitoes generallyapply, except that a greater amount of abra-sive powder (Alundum) may be required forthe grinding of ticks since they are leatheryand tough. Sometimes they are ground whilestill frozen.
VERTEBRATE BLOODAND TISSUE
The methods of trapping and bleedingvarious vertebrates in the field are beyondthe scope of this manual and will not be dis-cussed here. However, in view of the factthat the samples obtained are generally pro-cessed in association with the arthropodmaterial, the handling of vertebrate sampleswill be considered briefly.
Whole blood specimens are allowed toclot and the serum is removed. Serum removalis facilitated by use of a small, portableangle-head centrifuge carried with the fieldequipment. Both the clot and the serum are
29
then sealed and frozen on dry ice for trans-port to the laboratory, or they may be heldunfrozen on wet ice if rapid transport to thelaboratory is possible. The clot can be groundlater in two to four times its volume of dilu-ent, centrifuged, and used for virus isolationtests, thus conserving the serum for serologictests if it is in short supply.
Blood specimens diluted in part withantibiotic-fortified 25 percent normal rabbitserum diluent may be kept on wet ice forseveral days. If tests are for virus only, or ifthe only aerological tests to be run areneutralization tests, the whole mixture maybe frozen. If, however, other serologic testsare contemplated, the specimens should becentrifuged and the diluted serum removedbefore freezing. Generally the small clotis discarded.
The carcasses of small rodents areplaced in plastic bags, labeled, and frozen toreturn to the laboratory. There they are storedat 60° C in a mechanical freezer untilready to dissect. A small piece of braintissue weighing approximately 0.2 gm is re-moved from the thawed animal with sterileprecautions, ground in 2.0 ml of the 25 per-cent rabbit serum diluent, centrifuged, andinoculated into suckling mice for testing. Asimilar pool is made of combined smallpieces of heart, liver, and spleen.
The other laboratory isolation proceduresare similar to those already described in de-tail in the mosquito section.
