DARWIN INITIAVE “Capacity building for biodiversity studies of freshwater insects in Argentina”
Recommendations by the Natural History Museum (NHM) on METHODS FOR COLLECTING, PRESERVING,
REARING AND MAILING OF SIMULIID SPECIMENS
Luis M. Hernández
INTRODUCTION
The primary purpose of theses instructions is to provide uniform method for the collection, preservation and rearing of Simuliidae
commonly used by the research team at the NHM. These instructions are intended mainly for the field worker who needs to collect and
handle specimens of simuliids as r efer ence material fo r b io d iversity o r vecto rs stud ies. The collecting techniques
and rearing methods in Simuliidae in this protocol stems up from those outlined in Anonymous (1994) mainly by A.J.Shelley & C.
Lowry. Similar techniques also follows that of Crosskey (1990) and Lane & Crosskey (1993). The collection forms for recording
data has been developed from colleagues at the Mosquitoes Research Programme at the Natural History Museum (NHM) and
adapted for studies in Simuliidae.
Essential for this type of study is a uniformly prepared material, where specimens are collected and preserved in different
methods for integrated taxonomic studies. It is paramount in Simuliidae taxonomy that adults are individually reared, dried-pinned,
and kept together with their respective pupa for identification. The coloration pattern of the thorax is an important taxonomic
character and it will not visible when specimens are in ethanol. Therefore, the majority of the adults should be micropinned and
the pupa kept in a plastic microvial with glycerine attached to the pinned-adult; some other material should be stored in ethanol for
dissection. General information on bionomics and environmental factors, and a sample from as many habitats as possible in the
study area is also essential for species distribution. Important considerations in selecting the methods and techniques adopted here
have been simplicity and suitability for use under field and laboratory conditions and standardisation and simplification of records
and labelling to minimise errors and to save time.
The Simuliidae are specialized insects as the female require a blood meal for egg maturation. Being specialized this
family require some special collecting and handling techniques, as well as variety of preservation methods for identification.
COLLECTION RECORDS
Collection form. A standard form (see Appendix 1) for recording all the data pertaining to a collection has been developed from
colleagues at the mosquitoes research group and this has been adapted for studies in Simuliidae. The form must be filled out in the
field as completely as possible and the remainder added in the laboratory. A pencil should be used for all entries.
All
measurements should be indicated in the metric system. There is a minimum of writing to be done (only in lined open spaces) on
the form, the rest is to be done by circling or underscoring appropriate words or statements or by placing check marks or other
signs in appropriate columns.
COLLECTING SPECIMENS
The equipment and chemicals required to collect blackflies are given in Appendix 2.
Immature stages (Larvae and pupae): These are found by searching likely submerged substrates, from which they are easily removed with fine forceps. It
is desired to take larvae and pupae back alive and placed on a self-sealed polythene bag to take to a laboratory base from the field
collecting sites. The bags should be placed in a cooler that contains ice or cold water to keep a cool temperature inside it. The substrate will later be
examined in the laboratory base for selecting the mature pupae to rear.
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As pupae are firmly fixed to their holds they need careful side to-side pressure to loosen them and prevent pinching as they are
removed; once loose they suffer least damage if picked up by the hind end of the cocoon. Diversion of water flow may be needed to allow
collection of specimens attached to smooth rock surfaces that cannot be lifted.
Early stages can be conveniently collected straight into small vials containing a preservative fluid. The recommended vial is a small
glass bottle with a non-contracted neck and a screw cap. Such bottles measuring about 4.5x2 cm are ideal, tough for field use and capable
of being stood down safely whilst substrate and specimens are handled; narrow tubes and vials with press-over or press-in plastic caps are
unsatisfactory.
The recommended collecting preservative for all general purposes is ethyl alcohol (ethanol). For field collecting this should be at a
minimum of 80% strength but a 95% strength should be used if larvae are wanted with opened head-fans and extruded rectal o r g a n s .
A l c o h o l a t 7 0 % s t r e n g t h i s w i d e l y u s e d b u t i s unsatisfactory because water on and in the larvae when they are placed in the
collecting bottles dilutes the alcohol to an unsafe level for preservation (if its use is unavoidable the specimens should be transferred to fresh
alcohol as soon as possible after collection).
Larvae intended for cytological study must be collected into Carnoy's fixative (1 part of glacial acetic acid mixed with
two or three parts of absolute ethyl alcohol) and the fixative should be made up fresh at the time of collection. Larvae need not be cut open.
Adult flies. Wild female flies are usually collected in vector/biodiversity studies by exposing a person as human bait and catching
flies when they are settled on the skin to take blood. A suitable collecting tube can be placed over the fly as it probes; if the
specimen does not immediately enter the tube it can be induced to do so by touching it carefully with the rim of the tube. The best
specimens are obtained if flies are collected singly into individual tubes, but if biting numbers are high a sucking-tube (pooter) is
useful for rapid collection of many flies into a single container. Glass tubes with plastic lids or corks and measuring 3"xl"
(7.5x2.5 cm) are recommended for individual fly catching.
Flies are prone to die quickly under the tropical conditions where vectors occur and should be held in a cool and shaded place if
they are needed alive, e.g. whilst being taken to a laboratory base. A cold-box is useful as a container for live flies because darkness inhibits fly activity
and minimises possible damage to specimens.
T r a p s o f v a r i o u s k i n d s , i n c l u d i n g a n i ma l - b a i t t r a p s , C O attractant traps, sticky traps, light-traps and suction traps,
can be used to collect flies, but trapping techniques are more for specialised research purposes than for routine field collecting
(for which catches from human bait remain the most effective procedure).
Sweep-netting of likely resting sites, such as river -side vegetation, is seldom useful as a collecting
method. Hand netting of ovipositing or swarm-mating flies is occasionally possible.
PRESERVING SPECIMENS
Larvae and pupae. These should never be stored for long in the original tubes or bottles in which they were collected; however good the corks or
caps the risk of drying-out is high in the t r o p i c s a n d l a r v a e a n d p u p a e c a n n o t b e s a t i s f a c t o r i l y reconstituted
once partial or complete drying has occurred. Specimens should be transferred from their original vials into tubes intended for
permanent storage, and these tubes plugged with cotton-wool and inverted in container jars filled with the s a me p r e s e r v a t i v e fl u i d
a s t h e s p e c i me n t u b e s . Inversion of tubes ensures that specimens cannot become dried out even if the fluid level becomes very low in the container
jar ( F i g . 1 ) .
The ideal preserving fluid is 80% ethyl alcohol. Alcohol at 70% is not strong enough to guarantee the safety of specimens from
decomposition and strengths above 80% are unnecessary and too expensive. Decolorization of specimens occurs through long-term
alcohol storage, but happens also with other conventional preserving fluids (e.g. formalin) that are otherwise effective. The
practice of adding glycerine to alcohol storage vials is not reco mmend ed ; it can produce an undesirable clarifying of
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specimens and is unnecessary if care is taken to see that vials and jars have good seals (corks and screw-caps are poor seals and do not
prevent alcohol evaporation for long).
Recommended storage tubes are flat-bottomed glass tubes measuring 2"x0.5" (50x12 mm); this size permits storage of many
specimens, allows space for data labelling, and ensures that specimens are easily removed when needed. Small narrow-bore tubes are
useful for storage of important single specimens within a larger storage t u b e , b ut lo ng narro w t u b e s ( u n d e r 1 0 mm d i a me t e r )
a r e unsatisfactory as standard storage tubes because specimens are hard to remove. Whatever tube is used, care is needed to ensure that air
is completely excluded; soaking of cotton wool plugs in alcohol before they are inserted in the tubes helps to preclude air bubbles.
Recommended container jars are simple straight-sided jam jars provided with plastic-coated metal twist-grip caps. These
are easily obtainable and in every way better than expensive laboratory jars from specialist suppliers. The best size of
standard jam jar as container for the 2-inch tubes is height 3.5" (9 cm) and diameter 3" (7.5 cm).
Larvae that have been collected into Carnoy's fixative for chromosome study should be transferred to fresh 'Carnoys'
within a few hours of collection and stored in 'Carnoys' until they are studied. They should be kept refrigerated as far as possible. Larvae and
pupae should be removed from their substrates before storage. Alcohol makes pupal cocoons rather brittle and liable to damage if
pupae need to be removed from substrates after immersion in preservative.
Adult flies. Flies can be killed with ethyl acetate, ether, chloroform, or other standard killing agent, or with tobacco smoke
or exposure to heat. Once dead they shrivel and harden rapidly and become liable to fragment unless quickly preserved in a suitable
manner. They can either be kept as dry flies, which is the best method for reference collection purposes, or as alcoholpreserved specimens. Alcohol material is useful for some purposes, especially as an adjunct to dry material when flies are plentiful, but preference
should be given to dry preservation if specimens are scarce. Freeze-drying produces the most perfect specimens and should be used if
the facility is available. Alcohol material should be stored as described above for larvae and pupae.
Dry flies must be individually preserved as they cannot safely be kept en masse in tubes or other containers. Each fly needs
mounting, for example micro-pinning or glueing to a support (Fig. 1)
Micro-pinning: this is the strongly recommended method for preserving adult flies. Fine-calibre stainless steel micro-pins
(Minuten), as supplied by entomological equipment firms, are used and each fly mounted on its own micro-pin. Each micro-pinned fly is
'staged ' ( Fig . 1 ), i.e. sup p o r ted o n a sho r t mo unt o f Polyporus (the best material), plastazote, cork or soft cellular plastic strip
(polystyrene does not grip pins properly and should not be used), and the whole carried on a long strong pin that bears the data labels. Micro-pins are
best handled with curved insect forceps ('pinning' forceps). Flies are most easily pinned through the side, and if possible pinning should be
done under the stereo microscope to ensure that the specimen is pierced in the best position, i.e. through the thorax behind the pleural
membrane and below the wing base (Fig. 1). The impaled fly should be centred on the micro-pin for safety; 12.5 mm
length micro-pins should be used to allow sufficient length for staging the handling. (The micro-pinning method recommended in
some textbooks by which the micro-pin is passed through the staging material first and the insect positioned on the tlp of the pin should
never be used. The specimen is too vulnerable to loss and cannot conveniently be restaged if this becomes necessary.)
Glue-mounting: The fly is glued to the tip of a tapered mount of card or other material or is attached (e. g. by a dab of shellac) to the mid-shaft of a
long carrier pin. Glueing methods are not recommended as specimens are liable to break free from their adhesive and unless glueing is
expertly done it obscures much of the specimen.
Collections of dry adult flies must be kept free from mould and insect pests, to which they are specially liable in the tropics. Small air-tight
wooden store boxes that have been treated with me r t h i o l a t e o r s i mi l a r f u n g i c i d e a n d h a v e p r o v i s i o n fo r naphthalene
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preservative are the best storage containers for pinned flies under field conditions. Each box should ideally be wrapped tightly in a
polythene bag, preferably containing some silica gel to keep the specimens dry.
Slide preparations: Specimens intended for a specialist or for deposit as voucher material in a reference collection should not be mounted
on slides. Slide preparations of whole larvae, pupae or adult flies are of little use. Specimens should never, as a substitute for alcohol
preservation, be routinely mounted on slides in media such as chloral gum or polyvinyl lactophenol; t h i s d i s t o r t s t h e i r
s h a p e a n d o v e r - c l e a r s t h e m, t h e r e b y destroying the natural appearance (see SLIDE PREPARATIONS IN SIMULIIDAE)
Data labelling: All preserved specimens must be accompanied by collection data, otherwise they have little value (Fig. 1, 2). Data
labels should be placed inside each vial of alcohol material with the wording outwards so that it can be read without removing them;
labels on pinned specimens should be spaced when possible so that all can be read. Printed labels are desirable to avoid problems with
indecipherable handwriting. Code letters and numbers for temporary field use are inadequate for voucher specimens and data of reference material
should include (i) collection locality, (ii) date of collection, (iii) name of collector. The locality data should include the following whenever practicable:name of country, name of major subdivision ( usually province o r district), nearby town or village, name of river or
stream, altitude, latitude and longitude, grid reference (if any). Useful supplementary data include the host for flies
caught biting and notes on atypical capture sites ('in vehicle', 'in light-trap' etc.). When specimens have been identified they should be
labelled with their scientific names, each dry fly being given its own identification label if individually mounted.
REARING ADULT FLIES FROM PUPAE
For experimental purposes or reliable identification it is often necessary to obtain reared adult flies from pupae. Procedures f o r d o i n g
s o a r e s i m p l e , b u t c a r e i s n e e d e d f o r t h e m o s t efficient results and the most perfect specimens; potentially important vector
material is often spoiled through poor handling. According to the purpose, flies may be individually reared or mass reared (see Fig. 1)
Individual rearing: This method should be used to obtain voucher specimens as it allows each fly to be kept correctly associated with the
pupal skin from which it emerged. This association is often essential for correct identification.
Choosing pupae for adult emergence: Flies emerge best from mature or almost mature pupae, inside which they are already 'pharate
adults', and these should be used for quick and depend able results. Suitable pupae are recognised by being very dark
to almost black or by having the eyes very conspicuously darkened.
Setting up emergence containers: Each pupa is placed separately in a small covered container that is kept slightly damp
(not wet). The most useful containers are either individual vials (corked glass tubes measuring about 7.5x2.5 cm are excellent)
(Fig. 1), or the individual cells of a plastic grid-tray. The pupae can usually be removed without damage from their stream
substrates, and such removed pupae should be laid on dampened filter paper, in the emergence tube or on damp filter paper or
blotting paper placed as a floor to the cells of a grid-tray. Alternatively, pupae can be left attached to small pieces of
vegetation which are laid on strips of damp filter paper and propped against the side of the emergence containers, but in this case the substrate
should be trimmed to a minimum around the pupa to reduce condensation. Whether removed from the substrate or not the pupae should be
touched against absorbent tissues before being placed in emergence vessels to remove excess moisture from them. Emergence containers
must not be too wet or allowed to 'sweat' - excess water is the main cause of emergence failures or o f s p o i l e d e me r g e d fl i e s . O n c e
s e t u p fo r e me r g e n c e t h e containers must be kept cool, and it is useful to keep them in darkness to reduce activity by emerged flies.
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Containers should be examined frequently so that pupae can be removed to drier conditions if necessary and any adult flies removed
as soon as
possible.
Handling of emerged flies: Flies will emerge at irregular intervals for several days after the pupae are set up but if not
emerged in about 4-5 days it is seldom worth retaining pupae any longer. Flies are easily spoilt bybeing trapped on moist parts of the emergence
containers and should be transferred to clean dry tubes as soon as possible after emergence, and kept alive for a t least few hours
(best 24 hours) in a cool and dark place t o harden (for reference and identification the most perfect specimens possible are
needed) ; unhardened ('teneral' ) flies make bad reference material because their final colouring is undeveloped and they shrivel
excessively when dead. Flies emerging in, for example, 3"xl" tubes can be easily transferred to dry tubes of the same size by holding
one tube-mouth against the other and extending the dry tube towards the light (flies tending to go upwards and towards light sources).
Flies emerging in grid-trays need to be removed by cutting the covering material (e.g. cling-film) of the cell and removing the fly by suckingtube. Killing and pinning can be done as described above under preserving methods.
Handling of pupal skins: Empty pupal skins and cocoons should be remo ved from emergence tub es and placed in
alcohol in an individual microvial, they must always rema in w ith their associated adult. When the adult is micropinned its correlated pupal skin and cocoon is attached to the carrier pin below the adult mount. Pupal skins should be kept in a fluid
such as glycerine in a micro-vial (Fig. 1).
Mass rearing: If large numbers of adults are wanted without need for the associated skins from the pupae they can be obtained by
mass rearing in large containers. Stones or vegetation bearing quantities of pupae at all stages of their development can be placed in a
damp vessel such as large jar layered on the bottom with wetted filter-paper and any emerging flies extracted as required. A convenient
method with material on grasses or other trailing vegetation is to suspend the vegetation in large polythene bags, removing the
flies by sucking-tube alive or (if not needed alive) byfirst killing them with ethyl acetate or a similar killing reagent introduced into the bag.
SLIDE PREPARATION IN SIMULIIDAE
One method commonly used to preserve dissected specimens of simuliids using Berlese as mounting medium is given in
Appendix 3 (after Anonymous, 1994). However, there are several problems that may occur with Berlese mountant, e.g., it sometimes
becomes very dark (almost black), the choral hydrate crystallizes or the mountant may dehydrate if not properly ringed. It is therefore
suggested that for permanent mounts Euparal is used.
Dissections of Simuliidae are now carried out following a modified technique routinely used by colleagues in the
Mosquitoes Research Programme at the Natural History Museum (BMNH) which have been outlined in Hernández & Shelley
(2005). This technique follow closely the one detailed in Appendix 3 in terms of dissecting steps and equipments, but differ in
some of the chemicals used for final positioning of main structure. The main morphological character used in Simuliidae
identification and how they are positioned on the slide are detailed in Figs. 3-7.
When specimens are to be dissected from dried pinned adults, these are left in a relaxing box overnight to soften and then
dissected the following day. Specimens preserved in ethanol they are placed in single petri dishes or in a 10 or 12 hole porcelain
plates for dissection, and following the same detailed procedure: All specimens are individually dissected in 80% alcohol. The
adult’s head, thorax and abdomen were placed in hot 10% KOH for 10-15 minutes. These body parts were then rinsed in water for
5-10 minutes and after two changes of 80% alcohol, all unwanted body tissues and membranes were removed. The head, thorax
and abdomen were placed together with the remaining parts of the adult’s body and pupa, transferred to Cellosolve (2-
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Ethoxyethanol) for 10-15 minutes and mounted in a cavity slide using Euparal® as the mounting medium after the genitalia and
cibarium had been dissected in a small drop of phenolic gum Copal (D.J. & D. Henshaw supplier). The slide was left to dry
overnight and then a coverslip with a drop of Euparal® was placed on top the following morning. All slide should be kept
horizontally for storage. The slide should be put in an oven at 45ºC-55ºC to dry for about four weeks, if necessary.
IDENTIFICATION OF ARGENTINEAN SIMULIIDAE
The most complete references for identification of Simuliidae in Argentina are that of Coscarón (1987, 1991), which include keys and diagnosis of
genera, subgenera and species in this country.
MAILING SPECIMENS
The following points should be observed when mailing specimens to specialists as voucher material or for some other purpose.
Avoiding damage: Many specimens reach their destination in broken or irretrievably damaged condition. Nearly always
this is because of poor packing of specimens before dispatch. Damage can be avoided by taking the following simple precautions.
Specimens in fluid: These should be mailed in screw-cap vials, which are usually leak-proof even if not taped. Vials
with press-in or press-over caps are not satisfactory and corks, if used, should be waxed for leak-proofing. Glass vials are as safe as plastic
vials when properly packed. Each vial should be individually wrapped in tissue or cotton-wool packing. After individual
wrapping vials need to be placed in a strong container such as a wooden box or tin, and this in turn packed in a large carton containing shock-absorbent
material such as cotton-wool, polystyrene chips or wood-wool; for individual vials specially hollowed wooden containers are available
and are ideal. Each vial should be lightly plugged within by tissue or similar material to prevent specimens from
slopping back and forth in
transit, and air-bubbles should be rigorously excluded (many fluid-preserved specimens reach specialists in a damaged or partdried
condition because steps have not been taken to prevent swilling and to eliminate air).
Pinned adult flies: These should be securely pinned into a rigid box of wood or stout cardboard or plastic. This container should then
be packed into a much larger carton so that there is a minimum of three-inch thickness of soft packing material (e.g.
polystyrene beans or wood-wool) around each surface of even a small specimen-box - and more than this for large store-boxes. If the specimens
are staged the mounts should be cross-pinned to prevent them swinging, or if not cross-pinned then the specimens positioned in the box so that they
could not touch one another if the mounts rotated. The specimen-box should contain a cornerpiece of pinned cotton-wool to catch any
specimen or structure that breaks loose, and its opening should be covered with cellophane or cling-film.
Marking packages: Mail office requirements vary, but packages should carry "fragile" stickers and labels
indicating the contents as "Specimens for scientific study: no commercial value" or equivalent statement.
Advising specialists: Specialists should be advised in advance if specimens are being mailed to them, and prior arrangements should be
made with a specialist if a sending is likely to involve a major identification or research task. Specimens should be sent in a condition that
minimises expenditure of specialist time: e.g. larvae and pupae should have been removed from substrates. If the return of specimens is
expected it should be kept in mind that, by convention, a specialist is considered free to retain up to one-third of the material sent for
identification without special agreement.
SUMMARY OF RECOMMENDED METHODS WITH VOUCHER SPECIMENS
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1. Collect and preserve larvae and pupae in ethyl alcohol ( e t h a n o l ) a t m i n i m u m 8 0 % s t r e n g t h . D o n o t a d d
glycerine.
2. Store preserved larvae and pupae and alcohol-preserved adults in tubes in larger alcohol jars for safety.
3. Preserve adult flies as dry micro-pinned specimens for p r e f e r e n c e , u s i n g a l c o h o l - p r e s e r v e d f l i e s a s a secondary
adjunct to pinned material.
4. Keep flies reared from pupae alive for at least 24 hours to harden.
5. Keep pupal skins that correlate with reared flies in microvials containing glycerine mounted with the adults.
6. Ensure that specimens are accompanied by full collecting data and by name labels if they have been identified.
7. Inform specialists in advance if voucher specimens are being mailed to them.
8. State the whereabouts of voucher specimens in publications to which they relate.
REFERENCES
Coscarón, S. (1987) El género Simulium Latreille en la Región Neotropical: Análisis de los Grupos Supraspecíficos,
Especies que los Integran y Distribución Geográfica (Simuliidae, Diptera). Museu Paraense Emílio Goeldi, 112 pp.
Coscarón, S. (1991) Fauna de Agua Dulce de la República Argentina. 38. Insecta, Diptera. 2. Simuliidae. vi+7-304 [+ 67
figures on unnumbered pages], Fundación para la Educación, la Ciencia y la Cultura, Buenos Aires.
Anonymous (1994) Manual of Annual International Course on Identification of insects a\ns Arthropods of Medical and
Veterinary Importance, 18 April-13 May, 1994 [Dissection and mounting techniques in Simuliidae by AJ.Shelley & C.
Lowry.]
Crosskey, R.W. (1990) The Natural History of Blackflies. Ix+ 711 pp. John Riley, Chester.
Hernández, L.M. & Shelley, A.J. (2005) New specific synonymies and taxonomic notes on Neotropical blackflies (Diptera:
Simuliidae). Zootaxa, 853: 1-46.
Lane, R. & Crosskey, R.W. (1993) Medical Insects and Arachnids, 688 pp, Chapman & Hall, Andover.
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FIGURE 1. Recommended methods for preserving and rearing adults in Simuliidae (after Anonymous, 1994; Lane & Crosskey,
1993). 1. Specimens in ethanol. 2. Single reared micropinned adults showing data labels. 3. Recommended method to single
reared simuliids species. 4. Recommended method for micropinned adults, black circles and arrows indicates the different position
of the pin
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FIGURE 2. Recommended recording of data labels in Simuliidae (after Anonymous, 1994).
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FIGURE 3. Mouth parts and genitalia of Simuliidae (modified after Anonymous, 1994).
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FIGURE 4. Pupal and larval terminology (modified after Anonymous, 1994).
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FIGURE 5. Example of a slide dissection of a female simuliid (modified after Anonymous, 1994).
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FIGURE 6. Example of a slide dissection of a female simuliid (modified after Anonymous, 1994).
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FIGURE 7. Example of a slide dissection of a female simuliid (modified after Anonymous, 1994).
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Collection No. ARG-2006-
COUNTRY: ARGENTINA
Country
Locality
ARGENTINA
Nahuel Hupai National Park
Province
Specific locality (e.g., name of stream
Rio Negro province
San Carlos de Barriloche , Rio…..
Latitude/Longitude
Elevatio (m)
Date:
Time:
Collector(s)
Julieta Masaferro
Number
L
P
Sex
Identification / Notes
Appendix 1. Collection form of Simuliidae.
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COLLECTION TYPE
Immature: Larvae
Immatutre: Pupae
Adults:
Man-Biting (MB)
Landing
No biting
Larvae/pupae in Carnoys:
Larvae/pupae/adults in Alcohol:
Females MB to be pinned:
Specimens to rear:
ENVIRONMENTAL MODIFIERS
Primary
Secondary
Agriculture
Pasture
Grove/Plantation: ____________
Other: ____________
TERRAIN
Mountain
Hill
Valley
ENVIRONMENT
Rain Forest
Evergreen Forest
Deciduous Forest
Cloud Forest
Coniferous Forest
Scrub/Bush
Savanna
Prairie
WATER:
Permanent
Temporary
LARVAL./PUPAL HABITAT
On submerged branches
On submerged leaves
Attached to trailing vegetation
Attached to submerged plants
On rocks
On waterfalls
On rapids
Other:
WATER MOVEMENT
Slow
Moderate
Fast
WIND
None
Light
Gusts
Strong
SKY
Clear
Partly Cloudy
Overcast
Fog
Mist
Light Rain
Heavy Rain
Green
Blue-Green
Brown
Other: ____________
TURBIDITY
Clear
Coloured
Turbid
Polluted
DIMENSIONS OF SITE
Width m
Depth m
HEIGHT ABOVE GROUND
m
RIVER BED
AQUATIC VEGETATION
Submerged
Floating
Emergent
Submerged and Floating
Submerged and Emergent
Floating and Emergent
All Types
QUANTITY OF AQUATIC
VEG.
None
Scarce
Moderate
Abundant
rocky
sandy
calcareous
muddy
with algae
PHYSICAL FACTORS
PH
Conductivity
Water Temperature (ºC)
TDS
Water speed
Appendix 1. Collection form of Simuliidae. Continue…
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ALGAL DENSITY
None
Scarce
Moderate
Abundant
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FIELD
In the Cool Box
Cold Box (cooler) [to put inside in the cold box]
Plastic bags, self sealed- [more than 300] (approximately to take more than 20 per collecting day)
Pinning box
Standard insect pins, approximately 30 or 38 mm (more than 300)
Polyporous- cut in rectangles- (more 300)
Micropins A1 (more than 300)
Small plastic vials (about 12 mm) and plastic bungs for pinned link-reared material
Small glass vials (Durham tubes) for individually link-reared material or man-biting in spirit [more 300]
Glass vials of 12 mm (more than 300) for specimens in alcohol
Honey size plastic or glass jar to put vials of 12 mm with material in alcohol
Cotton wool (more than 4 packs) [one pack per collecting trip is enough]
Fine forceps (normally 4 pairs)
Pipettes- [more than 4, preferably plastic- Luis might take sample for NHM]
Small plastic petri dish or watch glass
Chemicals
Glycerol [To place pupa in small plastic vials for individually link-reared material]
Ethyl acetate or chloroform [killer agent]
Alcohol 100% (2-3 bottles per collecting day)
Alcohol 80%
Glacial Acetic Acid (100 %)
For Rearing of adults and collecting of larvae. Material in Carnoy’s
Small rearing tubes with plastic top- [more than 300-they can be re-use; size approximately 25 cm long]
McCarthy tubes for Carnoy’s [e.g., more 20 tubes per day; 10 tubes with 1 part of Acetic Acid and 20 with two parts
of 100%, to give a tube 2:1 parts of Carnoy’s] [More than 500] [Change the fluid after returning back to the lab]
[Tubes preferably with silver screw top to avoid spillage of fluid]- material should be put in fridge at cold
temperature, normally at 4ºC [Luis to take a sample from NHM]
Appendix 2. Equipment used for collecting and rearing Simuliidae in the field and in the laboratory.
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General
Scissors; Notebook; Whatman filter paper; Pencils; Pencil sharpener; Blades; Tissues; Paper for labelling; Rubbers;
Large piece of plastozote [Insect postal box size]; Maps of collection area; Thermometer; PH reader; Small petri
dish; GPS + Batteries; Digital Camera [For survey the different collecting sites; habitats]
LABORATORY
Dissections
Cavity slides- 16 mm (more than 1000)
Cover slips- 6-10-13-16 mm (more than 1000)
Hot plate [Square surface] for dissection of genitalia
Chemicals
Phenol gum copal [Luis to take some from NHM]
Cellosolve (2-Ethoxyethanol) (2 bottles)
KOH at 10-20%
Porcelain cavity plates (with 10 cavities each) for separation of material (around than 10) [cost £10 in UK]
Dissecting microscope
--------------------------------------------------------------------------------------------------------------------Companies
*
BioQuip
17803 LaSalle Avenue; GARDENA, CA 90248-3602, USA
Tel: 001 310 3240620
Fax: 001 310 3247931
E-Mail: bioquip@aol.com; http://www.bioquip.com/
STARLAB (UK); LTD.
4 Tanners Drive
Blakelands, Milton Keynes MK14 5NA, Great Britain, Tel: 01908 283800 , Fax: 01908 283802
E-mail: info@starlab.co.uk
VWR International Limited
Merck House, Poole, BH15 1TD, UK
ASCOT LABORAOTRIOES
52 Levenshulme Road, Gorton, Manchester, M18 7NN, UK, Tel: 0161 224 5184
Appendix 2. Equipment used for collecting and rearing Simuliidae in the field and in the laboratory. Continue…
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Dissection Techniques for Simuliidae (using Berlese mountant medium)
Equipment required. micro-pin dissecting needles with straight and hooked ends, Berlese mounting and storage medium,
10% KOH, glacial acetic acid, heating block, cavity blocks, cavity slides, fine forceps, cover slips.
A modified technique is given in the section “SLIDE PREPARATION IN SIMULIIDAE”. For positioning of body
parts see Figs. 3-7.
ADULTS
[Males and females are dissected in basically the same way.]
1. Place selected specimen in cavity block containing 70-80% alcohol. 2. Remove legs and wings at bases.
3. Transfer legs and wings to a second cavity block containing Berlese Storage Medium.
4. Separate head, thorax and abdomen and place in 10% KOH on a heating block set at about 70°C. Leave for 45 mins - 1
hr (depending on specimen size and length of storage), until the abdomen appears clear and red/brown 'fluid' is released from
the occipital foramen when the surrounding region of the head is gently pressed.
5. Transfer head, thorax and abdomen into glacial acetic acid for at least. 15 mins (timing is here is not vital; it is preferable
for material to be left for longer). [The effect of glacial acetic acid is to neutralise the KOH.]
6. Transfer from acetic acid to the cavity block containing Berlese Storage Medium with the legs and wings.
[Dissections are performed in this medium rather than in the mountant itself because it is less viscous than the
mounting medium and will not begin to solidify making manipulation impossible].
7. Remove all remaining soft tissue etc. from head, thorax and abdomen so that taxonomic characters are clearly visible.
For the head this is done by gently pressing the occiput region so that degenerated tissues are released through occipital
foramen. For slide mounting, only scutum, scutellum and postnotum are required from thorax. Superfluous regions and
remaining muscle tissue are picked (or cut) away. The abdomen should already be relatively clear. Residual tissues are
'hooked' out using a hooked needle, taking care not to damage the genital fork and spermatheca.
8. FEMALE ONLY: Remove cibarium by placing micro-needle (with slightly angled tip) inside the head (via occipital foramen)
onto frontal region thus holding the head firm, and gently pulling forward with another needle (with slightly angled tip)
on hypopharynx.
9. Dissect genitalia by placing whole abdomen on flat slide in drop of Berlese storage medium.
FEMALES: remove tenth segment with genital fork and spermatheca attached. Gonapophyses (ovipositor valve) should
remain intact on ninth segment. Separate spermatheca, then genital fork from remaining terminalia (including the cercus and
paraproct).
MALES: remove tip of abdomen from the ninth segment onwards. Dissect out gonostyles (styles, distimeres or
claspers), ventral plate, median sclerite and paramere (endoparameral organ).
[Note: sometimes median sclerite is so small that it is best left attached to paramere rather than risk losing it].
10. Mount with rest of dissection on cavity slide as in diagram.
Appendix 3. Dissecting method of Simuliidae using Berlese medium (after Anonymous, 1994).
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PUPAL EXUVIAE
1. Place cocoon and pupal exuviae in cavity block containing Berlese Storage Medium.
2. Gently remove pupal exuviae from cocoon by slitting underside edge of cocoon and teasing hooks on pupal abdomen
out to cocoon mesh.
3. Remove pupal abdomen.
4. Remove frontoclypeus and divide thorax into two by tearing along median line on dorsal suface.
5. Mount with corresponding adult as shown in diagram.
[Whole pupae are usually only dissected when there are no adults available.
In this situation mature pupae are necessary. Dissection is then as for exuviae and adults.]
LARVAE
[Only mature larvae with dark gill histoblasts are usually dissected]. 1. Place specimen in cavity block containing 70-80%
alcohol.
2. Remove histoblasts by peeling away the cuticle covering them and gently prizing them out. Transfer to a second
cavity block containing Berlese Storage Medium. (If the material has been stored in Carnoys fluid, then placing them in 50%
acetic acid for about 15 minutes will usually cause them to uncurl.) The histoblasts will usually need to be teased opened
whilst mounting to show length, number and configuration of individual filaments.
3. Remove anal papilla (if everted). Mount immediately in indicated position o n slid e (see d iagrams) as B erlese
tend s to clear this str uctur e and it may be lost if stored in the storage medium.
4. Transfer larva to cavity block containing 10% KOH on a heating block set at about 70°C for about 45 mins, or
until body has cleared. (The digestive tract contents and salivary glands should still be visible.)
5. Transfer to glacial acetic acid for at least 15 minutes (timing is not vital; it is preferable for material to be left for
longer). [The effect of glacial acetic acid is to neutralise the KOH.]
6. Transfer to cavity block containing Berlese Storage Medium with gill histoblasts.
7. Remove residual debris from larval body by forcing it along the body and then through the openings made when
removing histoblasts (and anal papillae, if removed).
8. Place larva ventral side up and remove head fans (taking care not to damage antennae), mandibles, maxilla, labrum and
hypopharynx.
9. Mount dissected larva on a cavity slide as in diagrams.
All preparations are left overnight so that the Berlese can set. A small fresh drop of Berlese is then added and the cover slip
immediately put in place. This prevents too much movement of the dissected specimen, ensuring the required parts
can be clearly observed.
Appendix 3. Dissecting method of Simuliidae using Berlese medium (after Anonymous, 1994). Continue…
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