VOLUME VIII
                              MOSQUITOES, BLACK FLIES, MIDGES AND SAND FLIES

                          REPORT To  THF

                          VOLUME VIII
    The work upon which this publication is based was performed in whole
or in part under Contract No. 68-01-2457 with the Office  of Pesticide Programs,
Environmental Protection Agency.

                           Report To The
                  Environmental Protection Agency

                              By The

             American Institute of Biological Sciences
                     Arlington, Virginia 22209
                         EPA REVIEW NOTICE

This Report has been reviewed by the Office of Pesticide Programs,
Criteria and Evaluation Division,  and approved for publication.
Approval does not signify that the contents necessarily reflect
the views and policies of the Environmental Protection Agency, nor
does mention of trade names or commercial products constitute
endorsement of recommendation for use.


                   PR.  CALVIN  G.  ALVARE2
                  American Cyanamid Company
New Orleans Mosquito Abatement
 District Director
University of California,
USDA-ARS, Insect Attractants and
 Biological Research Lab
State Education Building,
 Albany, New York
EPA Observer;
Criteria and Evaluation Division
AIBS Coordinators:


                         Table of Contents


Introduction 	    1

General Considerations 	    2

  Biological Parameters  	    2
  Experimental Parameters  	    2
  Data Analysis	    4
  References 	    6

CULICIDAE (Mosquitoes) 	    7

  Larvae 	    7

    Laboratory Evaluation  	    8
      Technical and Emulsifiable Formulations  	    8
      Granules 	    9
      Briquettes	   10
      Petroleum Oils	   11
      Insect Growth Regulators and Antimetabolites 	   12

    Field Evaluation	   12
      Liquid Formulations  	   12
      Granules	   14
      Briquettes	   14
      Petroleum Oils	   14
      Insect Growth Regulators and Antimetabolites 	   14

  Adults	   16

    Laboratory Evaluation  	   16
      Filter Paper Residue Test   	   17
      Wind Tunnel Aerosol Test	   17
      Topical Application Test 	   17

    Field Evaluation	   18
      Ground Applications  	   18
      Aerial Applications	   22

  References	   24

                    Table of Contents Continued


SIMULIIDAE (Black Flies) 	   29

  Larvae	   29

    Laboratory Evaluations 	   29
      Jar Technique	   30
      Cloth Bags Technique	   31
      Flushing and Draining Technique  	   31
      Trough Technique 	   32

    Field Evaluation	   33
      Single Stream Evaluations  	 . 	   33

  Adults	   34

    Laboratory Evaluation  	   34

  References	   35


Chironomid Midges  	   37

  Larvae	   37

    Laboratory Evaluation  	   37
      Conventional Larvicides  	 . 	   37
      Insect Growth Regulators and Antimetabolites 	   39

    Field Evaluation	   40
      Conventional Larvicides  	 . 	   40
      Insect Growth Regulators and Antimetabolites 	   41

  Adults	   42

    Laboratory Evaluation  	   43

    Field Evaluation	   43

Chaoborid Midges 	   44

  Larvae	   44

    Laboratory Evaluation  	   44

    Field Evaluation	   45

  Adults	   46

  References	   46

                    Table  of  Contents  Continued


CEBATOPOGONIDAE (Biting Midges, Punkies,  Sand Flies)  	    49

  Larvae	    49

    Laboratory Evaluation   	    49

    Field Evaluation	    49

  Adults	'	    50

    Laboratory Evaluation   	    50

    Field Evaluation	    51

  References	    52

     This document presents test procedures for the assessment of the
biological efficacy of pesticides against several groups of important
insect pests which pose an economic and public health threat to man in
urban, rural, agricultural and wilderness habitats.  While these protocols
do not cover all procedures and techniques employed to date, they, however,
include the most widely used and accepted procedures by researchers in
this field.  Deviation from and modification of these procedures may be
deemed desirable when required by specific conditions.

     The insects for which evaluation procedures are outlined here, be-
long to four families:  Culicidae (mosquitoes), Simuliidae (black flies),
Ceratopogonidae (biting midges) and Chironomidae (nonbiting midges, includ-
ing chaoborid midges).  These insects inhabit a variety of ecological
niches and show diverse biological, physiological and behavioral patterns.
In the evaluation of any bioactive compounds, the unique features of
each species must be taken into consideration.

     The procedures and techniques outlined here are mere guidelines for
securing meaningful information on the efficacy of pesticides for the
control of pest and vector insects.

                          GENERAL CONSIDERATIONS
     Members of the four families for which these protocols are developed,
are mostly aquatic or semi-aquatic in their immature stages.  The mature
stage is terrestrial, dispersing beyond the immediate vicinity of its
breeding source.  Pesticides are evaluated against the immature Cegg, larva,
pupa), as well as the adult stage.  In assessing the activity and efficacy
of pesticides against these insects, the following factors should be con-
sidered and standardized.  Deviations necessitated by test conditions are
given in the separate sections of this document.
                          Biological Parameters

     Species:—The insect should be identified to species whenever possible.
If not, it should be determined at least to the generic level.  Additional
information as to subspecific level, strains, etc., should be noted.

     Stage;—The susceptibility and response of living organisms to pesti-
cides varies a great deal depending on the size, stage and instar of the
insect.  Methods and procedures utilized for assessment are dictated by
the stage and habitat of the insect.  Pesticides are usually evaluated
against the larval and adult stages, while in certain situations, it will
be necessary to gather data on the egg and pupal stages.  In larval evalua-
tions, the instar should be standardized and specified.  Most laboratory
tests in this referenced group employ fourth instars.  In some situations,
younger or older (if existing) instars are employed.

     Age;—This factor plays an important role in influencing the perform-
ance of pesticides.  The age of the test insects should be reported or an
age range, if precise age is not known.  In dealing with larvae, it is not
essential to report the calendar age, if the larval instars are specified.
Approximate age of adult test insects should be specified.

     Sex;—Sex of the test insects should be specified.  This is an im-
portant consideration in evaluation procedures against adults.  It is
difficult to determine the sex of the immature stages and generally this
information is not needed.
                         Experimental Parameters

     Experimental parameters should be developed to meet the needs of a
specific situation.   The procedures followed under laboratory and field
conditions may be similar or quite different from each other.  Some general
guidelines for experimental design and data analysis are as follows:


     Test procedures and conditions should be standardized in the laboratory,
so that the results will be reproducible.  In addition to specifying the
biological parameters as discussed above, the following experimental con-
ditions and techniques should be given due consideration and recorded:

     Test Container:—The volume and size of the test container will vary
according to the requirements of the species and the nature of the test.
Guidelines are given under each group.

     Contamination;—This is an important problem in the evaluation of
pesticidal chemicals.  Laboratories used for testing should be kept free
of insecticidal contamination.  Contamination is minimal where disposable
equipment and containers are employed.  If glass or other nondisposable
items are used, they should be thoroughly washed, cleaned and decontaminated
prior to reuse.

     Test Solutions:—Solutions should be made in acetone, ethanol, or
other appropriate organic solvents.  Stock solutions should be sealed
tightly and refrigerated to preclude degradation.  If water is used as a
solvent, diluent or carrier, fresh preparations should be made prior to
each use.

     Number of Organisms;—This will vary according to the species and
availability of material for testing.  In general, no less than 10 individ-
uals should be employed per replicate.  The most desirable number of test
animals is 20-50 per unit.

     Replications, Number of Tests and Dosages:—Each treatment (test con-
centration) preferably should he repeated on a minimum of three separate
occasions, yielding a total of six replicates per treatment.  Each compound
should be evaluated at three to four discriminating dosages producing mortal-
ity in the range of 10-90 percent.
     Physical Factors;—Temperature, relative humidity and photoperiod
conditions should be standardized.  These parameters will vary according
to the species and test requirements.  In general, insects during exposure
and holding are subjected to normal daylight periods, 20-30°C temperature
and 40-80% RH.

     Duration of Exposure:—Varies according to species, age, stage and
the mode of action of test compounds.  For quick acting compounds, mortality
is assessed 24 to 48 hours after exposure.  In some tests, the insects are
exposed for 15-60 minutes, then transferred to or held in untreated contain-
ers for 24-48 hours prior to reading of mortality.  In the case of delayed
acting substances, such as insect growth regulators, the test duration may
involve the entire life stage or beyond.

     Food Provision:—For short exposure periods, food is generally not
required.  However, for longer exposure or holding periods, food is provided

the test insects.  In some situations, such as in the case of black fly
larvae, food particles may be employed as a vehicle for the ingestion of
stomach poisons.


     It is essential that bioactive compounds be evaluated against various
species in a variety of biotopes.  The following factors should be considered
in an evaluation of pesticides under field conditions:

     Sampling Techniques:—Various species and the different stages of the
same species require specific sampling techniques.   Applicable techniques
for assessment of populations of test species or groups  should be employed.

     Plot Size:—This can vary from a few square meters  to hundreds of
hectares,  depending on the objectives of the test,  species and the type of
control desired.

     Experimental Design:—Each treatment and untreated  check should be
replicated at least twice.  A minimum of three samples of the organisms
should be taken per plot at each interval.  Larger plots may require a
greater number of samples.  The treatments should be assigned to the plots
at random, or in other manners as dictated by the test conditions.
                              Data Analysis
     Data gathered in laboratory bioassay and field conditions should be
anlayzed for significant differences.   Standard statistical procedures and
computer programs are available for expediting such an analysis (Busvine,
1957; WHO, 1970).  For analysis and interpretation of the data, the follow-
ing procedures should be followed:
     In laboratory bioassay experiments,  the extent of mortality in the
checks will provide information on natural mortality.  If check mortality
is more than 20%, the tests should be discarded.   If mortality in the
checks is between 5-20%,  mortality induced by the test compound should be
corrected by Abbott's Formula:

% corrected mortality = % test  mortality  - % control mortality
                                100 - % control mortality       X  10°

     Corrected mean mortalities may be plotted against log concentration
on probit paper.   The points for three or four discriminating concentrations
of dosages showing activity are fitted with a straight line from which I^Q
and LCgo values are read  off.   The data may also  be treated and analyzed by
computer and the  confidence limits determined.

     In the field, the data may be analyzed and interpreted in various
ways.  If no check plots are employed, then the extent of control is
determined by comparing the mean values of posttreatment intervals with
those obtained prior to treatment using Formula I:
% Control of Inhibition of Emergence (El) = 100
                                               /T2 \
100 where:
     T  = mean number per sample prior to treatment

     T  = mean number per sample after treatment
     Where values for both treated and check plots for both pretreatment
and posttreatment periods are available, the calculations for extend of
control or inhibition of emergence are made by Mulla's Formula II  (Mulla
et al. 1971):
                                                 /Ci     T9 \
% Control or Inhibition of Emergence  (El) = 100-I ^  x ~-j  100 where:
     C-j^ = mean number per sample pretreatment in check

     T-, = mean number per sample pretreatment in treated

     G£ = mean number per sample posttreatment in check

     T2 = mean number per sample posttreatment in treated

     If no counts are available for pretreatment periods, and both check
and treated plot designs are utilized, then the extent of control and in-
hibition of emergence are calculated by Formula III  (Mulla et al. 1975):

% Control or Inhibition of Emergence (El) = 100[I.]100 where:

     T = mean number per sample in treated

     C = mean number per sample in check
     Additional techniques and computer methodologies may be employed in
the analysis of data as dictated by the condition of a given experiment.
Detailed methodologies are outside the scope of this document.

Busvine, J.R.  1957.  A Critical Review of the Techniques for Testing
  Insecticides.   Commonwealth Institute of Entomology.  London.  208 pp.

Mulla, M.S., G.  Majori, and H.A. Darwazeh.  1975.  Effects of the
  insect growth regulator Dimilin or TH-6040 on mosquitoes and some non-
  target organisms.  Mosq.  News  35:211-16.

Mulla, M.S., R.L. Norland,  D.M. Fanara, H.A. Darwazeh, and D.W. McKean.
  1971.  Control of chironomid midges in recreational lakes.  J. Econ.
  Entomol.   64:300-07.

World Health Organization.   1970.  Insecticide resistance and vector
  control.   WHO Tech.  Kept.  Ser. 443:279.

                         CULICIDAE (Mosquitoes)
     Mosquitoes rank as the most important insect pests and vectors of
human diseases.  They are prevalent throughout the United Stated causing
discomfort not only to humans but also to domestic animals.  Mosquitoes,
if not controlled, may cause substantial economic losses to real estate,
tourism and agricultural industries.  Mosquitoes are well known as the
vectors of human and equine encephalides.  In 1975, over 2,000 cases of
human encephalitis occurred in over 30 states and 30 deaths resulted.
Mosquitoes are vectors of dog heartworm which occurs throughout the
United States.  Furthermore, the mosquito species which transmit malaria,
dengue and yellow fever are present in many of our states.  The reintro-
duction of these diseases remains a possibility because of the extensive
world wide travel of our citizenry.

     There are at least 260 organized mosquito control districts in the
United States.  New districts are being formed continually as the need
for improved mosquito control is recognized by state and local govern-
ments.  In addition to the organized mosquito control efforts, many cities,
towns, communities and private citizens exercise chemical control of
mosquitoes when mosquito annoyance becomes severe or when the threat of
disease transmission exists.  Most of these agencies utilize three basic
methods of mosquito control.  These are physical control (breeding source
reduction or elimination), biological control and chemical control.  At
the present time, emphasis is placed on chemical control strategies be-
cause physical control measures are not always environmentally or econom-
ically feasible and biological control measures are not technically feasi-
ble for most species of mosquitoes.

     Mosquito control operations rely heavily on larviciding.  Since
immature mosquito populations are confined to breeding areas, less
chemical is required than in adulticiding programs.  However, adulticiding
forms an important part of most mosquito control programs since it is not
always feasible to achieve satisfactory control with larvicides and the
other available methods.

     Development of resistance to chemicals in use has brought about a
need to develop new classes of chemicals that are effective mosquito
control tools.  Furthermore, there is a need for the development of new
chemicals that are more specific, have greater efficacy and pose minimal
or no risk to humans and the environment when applied as mosquito control
     Vector and pestiferous mosquito larvae in the United States breed  in  a
variety of habitats, ranging from stagnant ponds to vast expanses

of coastal salt marshes and flood plains of rivers to rainwater caught in
tin cans, tree holes and the leaf axils of plants.  The control of these
mosquitoes is generally best accomplished by measures directed during this
somewhat confining and limiting period of larval development as opposed to
measures directed at the free-flying adult stage.

Laboratory Evaluation

     Laboratory testing of conventional mosquito larvicides, i.e. compounds
causing mortality within 24-48 hours as opposed to insect growth regulators
(IGR's) which may act more slowly, frequently begins with the exposure of a
given number of larvae to serial dilutions of candidate materials in a
suitable container.  Larvae should be of uniform instar and may be obtained
from laboratory-reared colonies.  If a laboratory population is not avail-
able, field collected specimens may be utilized.  The larval source in the
field must be sufficient to provide 20-25 larvae per each of two replicates
per treatment, including an untreated check.  Larvae for testing should be
third or early fouth instars, but earlier instars and eggs may also be
collected in the field and used in laboratory tests.  Field collected material
should be carefully handled and transported to the laboratory to minimize
injury.  Packaging into secured containers in a cool ice chest is desirable.
The larvae should be allowed an adjustment period of about one hour upon
arrival to the laboratory after transport from the field.  Any larvae
showing abnormalities, such as a fuzzy or discolored appearance due to the
presence of parasites, should be discarded.  Washing of larvae, if necessary,
may be accomplished by transferring the larvae to clean water or by strain-
ing larvae and then rinsing under slowly running water.  Lots of 20-25
larvae should be selected by pipet, strainer, or screen loop and distri-
buted into small beakers or containers, each containing 25 ml of water,
which are transferred to larger containers for bioassay.

     Technical and Emulsifiable Formulations:—Place 225 ml water into
appropriate containers 7.5-10 cm in diameter such that the depth of water
is between 2.5 and 7.5 cm.  Glass containers (jars, bowls or 500 ml beakers)
may be used, but they must be cleaned and decontaminated after each use.
Waxed paper cups are acceptable but must be discarded after each use.  Dis-
tilled water, rainwater or tap water may be used.  Tap water may cause pro-
blems in some areas because of the addition of disinfectants that may have
deleterious effects on the larvae.  Distilled water, obtained commercially,
may also contain traces of poisonous heavy metals.  Certain species, such
as salt-marsh or tree-hole mosquitoes, may suffer on transfer to relatively
pure water, an effect that will be reflected in high control mortalities.
In some cases, water from the breeding site should be used, provided that
it is free from insecticides and care is exercised to exclude detritus.
For salt-marsh mosquitoes, add sufficient salt to the solution.  The average
temperature of the water should be recorded and should be approximately
25°C; it must not be below 20°C or above 30°C.

     Stock solutions are prepared from technical or formulated supplies.
A 1% stock solution should be prepared using acetone, ethanol, or other

appropriate  solvent, with  serial  solutions  prepared  from this  stock
solution which is stored under refrigeration in a sealed glass vial.
Test concentrations are prepared by pipetting 1 ml or less of the
appropriate solution under the surface of the water into each of the
containers.  In preparing a series of concentrations, the most dilute
should be pipetted first.  Each concentration should be duplicated
with duplicate controls included for a solvent check (1 ml of acetone
or ethanol) and a standard insecticide, if possible.   Within 15-30
minutes of preparation of the test concentrations, add the mosquito
larvae by tipping the contents of the small beakers into the vessels
(WHO 1975, Mount et al. 1971, Clancy et al. 1969).  A variation of this
techinque can be accomplished by adding 20-25 larvae directly into 100
ml of water in waxed paper cups.  To obtain the desired concentrations,
1 ml or less of test solutions are added to each cup (Mulla et al. 1966).

     Mortality counts are made after 24-48 hours exposure.  The number
of moribund and dead larvae should be combined.  Dead larvae are those
that cannot be induced to move when probed with a needle in the siphon
or cervical region.  Moribund larvae are those incapable of rising to
the surface within a reasonable period of time or of showing the
characteristic diving reaction when the water is disturbed.  If more
than 20 percent of the control larvae pupate or die,  then the test should
be discarded and repeated.

     Each material should be tested on three separate occasions.  Percent
mortality should be corrected with Abbott's formula for each treatment
dosage.  A minimum of three discriminating dosages must fall between
10 and 90 percent mortality.  The mortality for each concentration is
plotted on probit log paper and a dosage response line fitted through
the points.  From this line or from a computer program, the LCrr, and
LCqQ for each material can be determined as described under Data Analysis.

     Granules:—Insecticides incorporated onto granular carriers are
highly successful as larvicides against mosquitoes, particularly when
the vegetative cover is too dense for penetration by sprays and dusts.
However, granulated formulations of insecticides are complex in nature
and require a special knowledge of the basic physico-chemical character-
istics of each of the ingredients.  The size and composition of the
granule, the solvent used for impregnation and various extrinsic factors
have been found to influence the release rate and effectiveness of candi-
date insecticides (LaBrecque et al. 1956, Mulla and Axelrod 1960a, 1960b,
1962) .  Different techniques of evaluation are required for granules than
for emulsions and evaluation of the toxicant is also generally accompanied
by evaluation of the various factors of the particular carrier.

     The effectiveness of granular insecticides (less than 0.5% in
concentration) can be determined by adding various quantities of the
formulation to water, adding larvae and assessing mortality after 24-48
hours.  Granules are dropped into quart jars containing 900 ml water.
High concentrate granules cannot be tested in this manner unless diluted
with the inert carrier.  The concentration of the active ingredient
should be 2X to 3X the LCgQ level.  Do not stir the water containing the

granules and add 20-25 third or early fourth instar larvae, without food
(LaBrecque et al. 1956),   The test should be conducted at a constant
temperature, within the range of 20-30°C, since temperature markedly
influences the rate and extent of release (Mulla and Axelrod 1960a).
This test procedure may be modified by adding the test organism to the
water after addition of granules at intervals of 24 hours or longer and
assessing the mortality.

     Diffusion of the toxicant can be determined by siphoning from the
top, without disturbing the portion remaining in the bottom of the jar,
portions to be jar tested.  Twenty to 25 larvae should be exposed to each
portion and mortality recorded after 24 hours or longer (LaBreque et al.

     Effectiveness and release of toxicants from granules can be deter-
mined by the aliquot testing method.  The rate of release of the toxicant
is measured by withdrawing aliquots of treated water at various time
intervals after addition of the granules.  One gallon jars containing
3,500 ml of water are used in evaluating the rate of release of granular
formulations (Mulla and Axelrod 1960a).  Aspirate 10 ml of water each
from the top and bottom portions (one inch above granules), combine and
add  varying volumes to 100 ml water in cups containing larvae to obtain
theoretical concentrations equaling 1-5X the LC^Q concentration of the
active ingredient.  Record larval mortality after 24 hours (Mulla and
Axelrod 1960a, 1962).  The % release for each formulation is calculated
from the observed mortality of larvae at a given concentration in the
cups.  The amount of toxicant corresponding to the observed mortality
is determined from standard dosage mortality lines established for the
toxicant against larvae of that species.  Knowing this concentration,
the extent of release can be determined in the jar.

     Larvae requiring a plant substrate for attachment, such as
Mansonia perturbans  (Walker), may be used in bioassay tests with
granular insecticides.  They may be collected from field sources and
shaken from the roots into water.  The clean plants should then be placed
in battery jars, or other appropriate vessels, containing 2 liters of
water.  Add 20-25 fourth instar larvae and allow them to reattach.  Add
various amounts of granules to jars and record mortality at 24 hours or
longer (Chapman 1955).

     Briquettes:—Insecticidal briquettes are used as mosquito larvi-
cides to obtain a longer lasting'residual action than afforded by other
formulations.  Such residual action is important in specialized uses
involving standing water in small containers such as cemetery urns,
street catch basins and stagnant ponds and, in some cases, to dispense
low levels of toxicant to flowing streams.  Briquettes are made of
plaster and sawdust, sand and cement, charcoal, polymers and various
combinations of plaster of paris (Raley and Davis 1949, Elliott 1955,
Barnes et al. 1967, Nelson et al. 1973).

     The length of effectiveness of briquettes can be measured by
periodically exposing briquettes to water and then bioassaying with
mosquito larvae.  Removing aliquots of the treated water after various
exposure intervals to the briquettes and bioassay as mentioned above.

     Petroleum Oils:—Petroleum hydrocarbon formulations have been
used for a long time to control mosquito larvae.  Although these oils
are primarily larvicidal, most formulations also induce mortality in
the eggs and pupae.  The oils are applied to the surface of water where
they form a thin film.  Larvae and pupae coming to the surface for
breathing receive toxic action.

     The biological activity of petroleum hydrocarbons is influenced by
the characteristics of the films.  Uniform film, having long duration,
generally results in greater mortality of immature mosquitoes coming
in contact with the film.  Films consisting of patches, lenses and globs
do not result in good contact with the larvae, and therefore, show low
biological activity (Hagstrum and Mulla 1968).  Due to the unique feature
of petroleum hydrocarbons, their biological assay is somewhat different
from that of the other insecticides.

     Techniques developed for evaluation of petroleum oils mostly use
glass or enamel containers.  Coated or plastic containers absorb oils
and should not be used.  Oil formulations have to be applied without
any dilution which will affect film characteristics.  Glass beakers
holding 250 ml water with 20-25 third or early fourth instar mosquito
larvae are treated by releasing droplets of oil from a micro-syringe
over the water.  The test containers are held for 24-48 hours at 20-30°C
and mortality recorded (Micks et al. 1967, 1968).

     Another variation of the above method is the use of shallow pie
glass dishes.  These dishes have a capacity of 1-liter, when filled
to the lip, and thus provide greater surface area than  beakers and
simulate field conditions in terms of the evaporation pattern of the oils.
Since the volume of water used here is greater than that in the beakers,
50 larvae may be employed per unit.

     Petroleum oil formulations are evaluated against the pupae in exact-
ly the same manner as against the larvae, while against eggs, a slight
variation may be used.  Assessment of ovicidal activity is determined by
counting larvae (Micks et al. 1967), or assessing the number of hatched
ova (with the operculum popped out) where larval counts are not possible
(Mulla 1964, Mulla and Chaudhury 1968).  The eggs are normally exposed
for 48-96 hours before mortality is assessed.  In order to assess the
extent of hatch of eggs laid in an egg raft (laid by Culex, Culiseta,
Manson-ia species and others), the egg rafts, after desired exposure,
are placed in 2 ml of 1% KOH in a screw cap vial for 12-24 hours.  The
volume of KOH is then brought to 10 ml with water, the vial capped and
shaken vigorously for one minute.  The KOH disintegrates the raft into
individual eggs.  For counting the hatchability, the egg suspension is
streaked out on strips of filter paper, observed and counted under a
dissecting microscope.  Hatched eggs are easily distinguished from unhatch-
ed eggs by the operculum opened out.  Eggs laid individually need not be
treated in this manner (Mulla 1964).

     Insect Growth Regulators and Antimetabolites;—The development of
compounds which inhibit mosquito development has occurred_ in the recent
past which have been called the "third generation pesticides'  (Williams
1967).  These synthetic compounds display juvenile hormone-like activity
which is characterized by inhibition or prevention of emergence of adults
by mimicking the biological activity of the natural insect hormones.
The use of the term "juvenile hormone" originally was used to  describe
this group of compunds, but it is no longer appropriate because it may
connote potentially undesirable interpretations, and because other com-
pounds not so closely related to the natural juvenile hormone  have shown
antimetabolic effects on growth development.  For the purpose  of this
discussion, both insect growth regulators (IGR's) and antimetabolites will
be collectively termed IGR's.

     The criteria for effectiveness of IGR's are different from those
used for measuring larval and pupal response to conventional larvicides.
In general, IGR's induce a response in stages beyond that treated.  In
evaluating the effects if IGR's, developmental events are monitored to
the stage of the adult emergence.  In some cases, it might be  necessary
to assess the longevity and fecundity of the resulting adult population.
Criteria for "normal" adults, or completely separated pupal exuviae, must
be established.  Normal adults are those displaying no abnormalities in
physical appearance or behavior.

     Laboratory assays are conducted primarily in glass containers, but
other types may be employed.  Paper cups are inadequate because they
deteriorate due to the length of the duration of the test.  Place 200-
250 ml of water into each container.  Add 25-50 fourth instar  larvae
from standardized larval source.  Add food in quantity and selection as
determined for each test species.  Pipet from stock solution the desired
amount of test compound, not exceeding 1 ml of solution.  Place test
units at constant temperature 20-30°C, and relative humidity,  ca. 40-80%.
Mortality of each stage is assessed every two or three days and dead or
moribund organisms are removed.  Normal adult emergence is assessed by
counting and removing completely separated pupal exuviae and collecting
adults in emergence units.  The overall effectiveness is determined by
calculating the inhibition of adult emergence in the treatments by formula
III as given under Data Analysis (Hsieh and Steelman 1974, Jakob 1972,
Jakob and Schoof 1971, 1972, Mulla et al. 1974, Schaefer and Wilder, 1972).
A variation of this technique involves the use of larger containers and
the removal of pupae to cages for assessment of eclosion (Dame et al. 1976).

Field Evaluation

     Liquid Formulations:—Compounds with proven laboratory effectiveness
are tested out-of-doors in small replicated man-made or naturally occurring
ponds.  An important feature of such experimental ponds is knowing the
volume of water for administering exact application rates.

     Laboratory outdoor ponds have been constructed by digging holes 3 m
long by 1,5 m wide and  ,75 m deep with sloping sides.  The ponds are lined
with a double layer of  8-mil polyethylene plastic film which laps over the
sides.  About 15 cm of  soil and sod are placed over the film and 1,100
liters of water added»  Manure and other organic materials are added.
Water hyacinths are placed in the ponds to shade the water and keep it
cool.  The ponds are allowed several days to stabilize.  Measurements are
made of final length and width at predetermined levels to calculate surface
area and pool volume.   The ponds are filled to the desired depth the day
before testing, and the water level is kept constant throughout the test
(Bailey et al. 1970).

     Other ponds are constructed in sizes of 5X5X0.3m and 3.5X7X0.3m
with an automatic water replenishment device triggered by a float valve.
These ponds have natural bottom, vegetation, and fauna, and support wild
populations of three to four species of mosquitoes.  For sampling sites,
each pond is provided with five bundles of straw, one in each corner and
one in the center, as a source of organic matter (Mulla and Darwazeh
1971).  Organisms may also be sampled along the vegetated margins or from
other portions of the ponds.  Fifty-five gallon drums are also used for
outdoor bioassay (Brooks and Schoof 1965).

     The pond systems are employed to evaluate compounds against indigenous
mosquito populations.   Pre-treatment counts of larvae are made by dips in
areas of highest larval concentration.  This sampling procedure is biased
and not random by design because larvae tend to clump and a true random
sample would be less definitive.  Samples from the margins of the pools
or next to artificial sampling sites yield the least variable results
(Mulla et al. 1975)-  Post-treatment samples should be collected at
regular intervals and mortality calculated according to the appropriate
formula given in the Data Analysis section.

     At times, it may be necessary to evaluate the longevity of a larvicide
by treating water in outdoor ponds and bioassaying it against laboratory
populations.  In this scheme, collect water samples prior to and at 2, 24 and
48 hours, or other intervals after treatment, return to the laboratory
and strain through a fine mesh screen to remove detritus, naturally
occurring mosquito larvae and other organisms if present.  Place one
sample into a container along with 25-50 early fourth instar larvae.
Evaluate mortality as determined in laboratory tests (Dame et al. 1976).

     Evaluation of mosquito larvicides at field scale level requires
special considerations.  Care should be exercised in calculating dosage
levels.  Sampling should be conducted by the same individual throughout
the trial.  The larger plot, the greater the number of samples required.
Sample at the same time of day in the same general area.  Sample those
areas most likely to have the largest concentration of larvae, i.e. ,
along margins of grassy shore, near organic accumulations or in the shade.
Count only third and fourth instar larvae and pupae to avoid recently
hatched eggs not exposed to toxicant.

     Tests in irrigated pastures with naturally occurring larvae can be
conducted in 1/32 acre, or longer, plots.  Small dikes may be employed
to separate the plots.  Mix the required amount of toxicant with 1,000 ml
of water and apply evenly with a hand sprayer.  Ten to 15 dips per plot
are taken prior to and 24 hours after treatment (Mulla et al, 1975,
Mulla and Darwazeh 1975a).  Effectiveness is determined by comparing
mean number of larvae in posttreatment with those in pretreatment of the
same plots (Formula I).

     Plots may also be established in river bottoms or other breeding
sources by diking and treating as above  (Mulla et al. 1960),  Rice fields
with established levees can be divided into plots for trials.  Treat each
plot with the proper dosage of chemical with a hand sprayer.  Assessment
is made by comparing pre- and post-treatment counts of mosquitoes.  Depend-
ing on the size of the plot, 20 samples per plot may be sufficient (Gahan
et al. 1976) .  Effectiveness is determined by comparing larval counts in
the post-treatment samples with that of the pre-treatment (Formula I).

     Granules:—Evaluation of granular insecticides under small field plot
trials can be accomplished as reported for liquid formulations.  Pre-and
post-treatment larval samples are taken from treated and untreated plots.
Roadside ditches have been used as granular test sites.  Ten to 20 foot
plots are treated and 10 dipper samples taken pre-treatment and 24 hours
post-treatment (LaBreque et al. 1956).  Granular formulations may be
evaluated against Mansonia perturbans (Walker).  Determination of control
is made by pre- and post-treatment collections of larvae.  A standard
quantity of plant material supporting larval populations is collected
and quickly transferred to white pans one-half full of water.  The plants
are shaken into the water to dislodge the larvae for counting (Chapman

     Briquettes:—The field effectiveness of briquettes is measured in the
same manner as for liquid formulations, except that evaluation periods are
extended.  Treated ponds are assayed by placing 25 early fourth instar
larvae into floating cages in the pond.  The cages are made of 32-ounce
waxed paper cups that have the bottoms replaced with a fine mesh screen.
Fishing floats are attached for flotation.  Assays can be made at regular
intervals until the formulation ceases to give control (Bailey et al. 1970).
Field water from the plots can also be bioassayed against larvae of known
susceptibility in the laboratory.

     Petroleum Oils;—The efficacy of oils can be assessed as reported
under liquid formulations.

     Insect Growth Regulators and Antimetabolites:—IGR's are field tested
in man-made ponds or small naturally occurring pools and other breeding
sources similar to those described for field testing conventional
larvicides.   When appropriate modifications as described for laboratory
testing of IGR's are employed, which measure their delayed effects, these
breeding sources can provide useful field trial information.  It is necessary
to follow a representative portion of the population through to emergence,

and this is accomplished by means of a floating cage,  The cage affords
constant exposure of the test organism to the treated water, and yet,
permits removal of the test population for recording of developmental
events.  The cage consists of a 1-quart polystyrene cup fitted inside a
styrofoam ring float.  The container is covered with lid screen to permit
ventilation.  Holes are cut in the lower section of the cup and are
covered with a fine mesh screen to allow water and food particles to pass
through, but not the larvae (Mulla et al, 1974).

     Ponds treated with IGRfs at various rates are assayed at various
intervals by isolating a portion of the natural populations of larvae
in two or more floating cages per pond.  Each floating cage contains
20 fourth or other instar larvae.  Emergence and mortality are monitored
every other day, or at other appropriate intervals and they may be
provided with food.  The percent inhibition of emergence based on the
number of larvae corrected by the formula given in Data Analysis is then
calculated (Mulla et al. 1974).

     Effectiveness of treatments in ponds or other breeding sources can
also be determined by collecting larval and pupal samples with a dipper
and assessing the density of the various instars.  This is especially
needed for the assessment of compounds that induce mortality at the
interstadial period or during molting.  In these situations, disappear-
ance or absence of the pupal stage provides a good criterion of effective-
ness (Mulla et al. 1975).  Water from treated ponds or other breeding
sources can also be returned to the laboratory for bioassay.  After
filtering through organdy or cheesecloth (discard cloth after each use
to avoid contamination), the  filtrate is assayed using the standard
laboratory techniques described for IGR's (Dame et al. 1976, Schaefer
and Wilder, 1972).  Field population in the treated and untreated
water may be brought into the laboratory for assessment.

     Natural field pools are utilized as testing units by dividing them
by dikes into replicated plots.  Emulsifiable concentrates,
microencapsulated concentrates and wettable powder formulations are ap-
plied by mixing into 1,000 ml of water and applying by hand sprayer
over the plot.  Percent reduction of populations is determined by five
dips per plot (1/32 acre plots) prior to and 24 and 48 hours after
treatment.  Percent emergence inhibition is determined by isolating
20 fourth instar larvae or pupae in floating cages as described above.
Emergence of pupae may be assessed by placing 20 per 100 ml water in
paper cups.  The number of dead pupae and adults is recorded after
completion of emergence.  The % El is determined as given in the Data
Analysis section (Mulla and Darwazeh 1975).

     Rice field mosquitoes can be assayed by dividing the field into
small plots prior to flooding.  Several days after flooding, a natural
population is treated at various rates with candidate compounds,
Following treatment, 20 larvae are collected from each plot and placed
in floating cages, as described above, back into the same plots.

Observations for mortality and adult emergence are made 24, 48, and 72
hours post-treatment.  Larvae may also be transported back to the
laboratory for observation and assessment of mortality and emergence
of adults (Steelman et al. 1975, Schaefer and Wilder 1972),  When the
test insects reach the pupal stage in the field, cubic foot cages (with
one side removed) are placed over concentrations of pupae in the various
tests and control plots to obtain adult emergence CSchaefer and Wilder
     Adulticides are important tools used by most mosquito control
districts since this method can be employed when other methods are either
not feasible or have failed to provide satisfactory control.  Adulticides
are primarily applied as ground or aerial aerosols and sprays.  In recent
years, ultralow volume aerosols and sprays have become popular because of
substantial economic and environmental advantages inherent with this
method.  Ground aerosols are useful in controlling adult mosquitoes in
urban and suburban areas where road networks allow adequate coverage.
Aerial applications are useful in large-scale adulticiding and in target
areas inaccessible to ground equipment.  Both aerosols and sprays are
applied at relatively low dosages which do not provide residual activity.
Thus, their effect is of a temporary nature and applications must be
repeated as reinfestation is indicated by mosquito surveys.
Laboratory Evaluation

     An important consideration in testing insecticides against adult
mosquitoes is susceptibility variation among species and strains of the
same species.  New insecticides should be evaluated against a wide range
of species and especially against the more important target species.
Variations in the susceptibility of strains usually depend on their history
of exposure to various classes of insecticides.  Thus,  the most useful
data are those obtained by using either laboratory strains of known
susceptibility (or resistance) or from field collected specimens.  If
field specimens are used, they must be collected from areas which have not
received recent insecticide applications.

     Test specimens may be collected as eggs, larvae or pupae and
reared to adults using standard rearing techniques (AMCA 1970).
Immature mosquitoes may be transported long distances if they are kept in
insulated jugs to protect them from excessive cold or heat.  They should
be kept in their breeding water until transfer to appropriate rearing
media in the laboratory.  In the field, adult mosquitoes can be collected
by aspiration or baited traps and transferred to the laboratory in
insulated chests containing moist cotton and also canned ice if being
transported during hot weather.

     In general, adult female mosquitoes are less susceptible than
males; therefore, the former are usually utilized in laboratory testing
program since they represent the field problem,  If both sexes are used,
data for each sex should be recorded and analyzed separately for maximum
statistical reliability.

     The age of test specimens should be as uniform as practicable within
a range of two to eight days since it does, to some extent, govern
physiological condition.  Other than age, the physiological condition
of test specimens is controlled by using standard rearing techniques which
require consistent handling, environment and nutrition.

     There are two widely used test methods available that permit the
rapid evaluation of large numbers of insecticides against adult mosquitoes
with a standardized equipment, solvents and handling procedures.  These
are the filter paper residue method and the wind tunnel contact aerosol
method.  The topical application method has also been used for many years,
but it has never gained popularity because it is relatively slow and

     Filter Paper Residue Test:—The insecticides to be tested are dissolved
in acetone (w/v) and pipetted  (1 ml) onto Whatman No. 2 9-cm diameter
filter paper placed horizontally on pin points.  After five minutes of
drying, a section ca. 5 cm wide at the widest point is trimmed off one
side of the paper and the latter is rolled and placed inside a shell vial
(2.1 X 8.4 cm) lining the sides almost completely.  Approximately one hour
later, the mosquitoes are anesthetized with cold or CC^, counted into
groups of 25 and transferred to the test vials.  The vials are covered at
the open end with cheesecloth and placed flat on their side for a one hour
exposure period.  After exposure, the mosquitoes are anesthetized and
transferred into holding cages (0.47 liter unwaxed paper cups fitted with
a net cover (Georghiou and Metcalf 1961, Georghiou and Gidden 1965).

     Wind Tunnel Aerosol Test:—The wind tunnel test simulates field
aerosol applications in the laboratory, thus it is a useful method of
testing chemicals intended for use in aerosol programs.

     Test chemicals are diluted in acetone or other suitable solvents
and pipetted as small aliquots (0.25-0.5 ml) into a nozzle designed for
total delivery into the wind tunnel tube.  Tunnel air velocities are
3-4 mph to simulate natural wind currents.  Adult female mosquitoes
are exposed in 14-18 mesh screen wire cages in groups of 25.  After
exposure, the mosquitoes are anesthezied with CC>2 and transferred to
clean holding cages for knockdown and mortality observations  (Mount
and Pierce 1975, Mount et al. 1976, Rathburn 1969, Boike and Rathburn

     Topical Application Test:— Application of diluted  (usually in
acetone) chemicals is made with a calibrated repeatable microsyringe.
Volumes applied are usually 0.3-0.5 yl/mosquito.  Specimens can be
anesthetized with either C0~ or cold during actual application.  For
consistent treatment, the droplet of insecticide should be applied to

the same body area (usually the mesonotum) each time.  One refinement that
is sometimes employed with this method is the coordination of dosage with
mosquito weight.  Adequate results can be obtained without using this
additional step, especially when working with mosquito populations of
uniform size and weight.  After treatment, the mosquitoes are held in
clean cages and provided a sucrose solution on cotton (Ludwick 1953,
Georghiou and Metcalf 1961, Mount and Pierce 1975),

     Since test specimens are treated individually with uniform dosages,
the repeatability of results with this method is somewhat higher than with
the two previously described methods.  Past results indicate that reliable
lethal doses may be obtained with an exposure of a minimum of 160 specimens
per chemical.  This total would be made up of four discriminating doses
and 40 specimens per dose with 10 specimens per replicate.

     In all procedures, mosquitoes that have been exposed to insecticides
should be held in clean cages and supplied with 5-10% sugar water solution
on cotton pads or wicks.   These moistened pads will supply food, water
and additional humidity to the mosquitoes.

     Mortality observations should be made 24 hours or longer after exposure
to the insecticides.  Additional knockdown and mortality observations
can be made at various intervals, if desired.  Mosquitoes are counted as
dead if unable to walk when the cage is jarred.  The same criterion
should be used in determining knockdown.  Knockdown refers to short
interval post-treatment effects (one hour or less).  Quick  knockdown
is a desirable, but not essential, characteristic of mosquito adulticides.
With some insecticides, mosquitoes may recover from the knockdown effect,
therefore, the 24 hour mortality observation is considered essential.

     Each laboratory test should include cages of mosquitoes not exposed
to insecticides.  These mosquitoes should be exposed to the insecticide
solvents, the identical handling procedures and the same laboratory
environment as those exposed to insecticides.  The inclusion of mosquitoes
not treated with insecticides in each test will reveal any mishandling,
insecticide contamination or physiological weakness of the mosquitoes.
When untreated mosquitoes show more than 20% mortality, the test should be
discarded and repeated once the cause for excessive mortality has been
resolved.  Corrections for untreated mosquito mortality of 5-20% can
be made by Abbott's Formula.  Data from laboratory tests should be
reported as lethal concentrations at the 50% and 90% levels (LC^g and LC^Q)
as described under Data Analysis,

     The primary function  of laboratory  tests with  insecticides
against adult mosquitoes is to determine their relative toxicities.
Therefore, in testing and in reporting data, a standard insecticide
should be included for comparison,

Field Evaluation

     Ground Applications;—Ground applications include ultralow  volume
aerosols and high volume thermal and cold aerosols.  Regardless of the

application equipment used, the basic  methods of evaluation are essentially
the same.

     Commercial aerosol generators should be maintained and operated
according to recommendations of the manufacturer.  Exceptions to this
could be variance in atomization requirements for particular insecticide
formulations.  Testing with custom constructed generators should be preceded
by research showing adequate atomization capabilities.

     Aerosol generators should be calibrated  for liquid flow rates at
approximately the same ambient temperature at which actual applications
will be made.  This is especially critical when using generators equipped
with flowmeters calibrated with a float.  The float position can change
dramatically with changes in viscosity as influenced by temperature.  This
problem is solved in actual practice by use of temperature correction
data for each insecticide.

     It is essential to test the actual insecticide formulation to be
registered for use.  This is especially critical with ultralow volume
equipment since droplet size may change significantly with change in
viscosity and surface tension.

     When applying insecticides, it is important to monitor dispersal by
the use of remote instruments and/or correlate actual insecticide dis-
charge with application time.  It is also important to either monitor
vehicle speed or time each application with a stopwatch to insure
accuracy and even coverage.

     The efficacy of aerosols of insecticides is evaluated by exposing
either caged or natural infestations of adult female mosquitoes downwind
of the application.  Caged mosquitoes can be  from either a laboratory
colony of known insecticide susceptibility or wild mosquitoes collected
from target areas.  Laboratory reared mosquitoes should be  fairly uniform
in age within a range of two to eight days old.

     Tests with caged mosquitoes can be conducted over various types of
terrain.  Level, open terrain enchances even  coverage and thus minimizes
experimental variation.  Extremely hilly or densely vegetated terrain is
not suitable for this kind of testing.  Tests in urban or suburban areas
better simulate actual application conditions, but require somewhat
greater sampling than tests in open areas.

     Meteorological conditions needed for aerosol testing are usually
found during twilight or at night when adult mosquitoes are active.  Ideal
test condiditons include ground winds of 1-10 kilometers per hour and a
ground-based inversion.  Good application conditions also exist during
isothermals if winds are 5-10 kilometers per  hour.  These stronger winds
tend to drift the aerosol across the plot before a great deal of
vertical mixing takes place.  With regard to  temperatures, this type of
testing should be done "in season", i.e,, when mosquitoes are likely
to be prevalent in target areas.  This could  range from 15-40°C,
depending on latitude.

     Mosquito exposure cages should be constructed of 14-18 mesh screen
wire.  Specimens are anesthetized with either CC>2 or cold for sexing and
counting into cages.  One type of exposure cage is 15 cm diameter X 2,5 cm
deep (Rathburn and Boike 1975),  These are covered with 14 X 18 mesh
screen on both circular surfaces and hung vertically on poles with screen
surfaces facing into the wind,   With this type of cage, the exposed
mosquitoes are returned to the laboratory, anesthetized and then trans-
ferred to clean holding cages.

     A combination exposure-holding cage for aerosol tests is 4.5 cm
diameter X 15 cm long and is constructed entirely of 16-mesh, galvanized
screen wire (Mount et al. 1975),  This exposure cage is threaded into one
side of a plastic slide.  A 4,5 cm diameter X 14 cm long plastic tube
lined with clean paper is threaded into the opposite side for use as a
holding cage.  During exposure to the aerosols the holding tube is sealed
with masking tape.  Immediately after exposure, the tape is removed and
the mosquitoes are gently blown from the screen wire cage to the holding
cage through a hole in the movable slide separating the two cages.  This
type of cage has the advantage of immediate transfer of the mosquitoes
to a clean holding cage.  The need for transfer is to keep the mosquitoes
from being confined to a treated screen wire surface.  The object of this
type of space treatment is to kill by direct contact rather than indirect
contact on a treated surface.

     After aerosol exposure, the mosquitoes should be returned to the
laboratory for a holding period of 12-24 hours.  Mortality counts should
be made at the end of the holding period; additional knockdown and
mortality counts can be made at any time desired.  During the holding
period the mosquitoes should have access to 5-10% sugar water solution
on cotton pads placed on the screened tops of the holding cages.  The
sugar water provides food, water and added humidity to the mosquitoes.
While in the field, mosquitoes should be maintained in clean insulated
containers at ca. room temperature and provided moist cotton for
humidity.  Excessive heat and low humidity are very detrimental to adult

     Most aerosol applications are made with swaths of ca. 100 m  (one
city block).  Small-scale tests with caged mosquitoes consist of one
to three swaths of 300-400 m.  Cages of mosquitoes should be exposed
in two rows 50 m apart and at downwind distances of 45-50 m and 90-100 m
to demonstrate swath coverage.  A minimum of four cages of 24 mosquitoes
of each species should be exposed in each test (Mount and Pierce 1972,
Mount et al. 1975, Rathburn and Boike 1972, Taylor and Schoof, 1968).

     Wide swath (up to 1,600 m) applications are preferred for ground
aerosol applications against adult mosquitoes in the western United States,
Testing methods for extended swaths are the same except that additional
cages of mosquitoes and/or counting stations are needed.  A minimum of
two rows of cages at downwind intervals of 200 m are needed to
adequately demonstrate complete swath coverage.  Thus, 16 cages of
mosquitoes would be needed to sample a 1,600 m swath (Gillies et al,
1972, Sjogren et al, 1973, Womeldorf et al. 1973).

     Tests with natural infestations of mosquitoes are not usually
feasible on a small scale  (.< 4QQ hectares) ,  Exceptions to this can be
made, if short interval post-treatment evaluations are made (this can only
be done with insecticides  that produce rapid Rill) or rapid reinfestation
(overnight) will not occur.  Overnight reinfestation is likely to take
place in small plots with most species of mosquitoes.

     Methods of determining the efficacy of insecticidal aerosols
against natural infestations of mosquitoes include the use of caged wild
mosquitoes and a variety of pre- and post-treatment sampling methods.
These include landing and biting rates on humans, New Jersey light traps,
CDC battery traps, CO- traps, animal bait traps, suction traps and truck
traps.  Landing counts on humans are most commonly used since they yield
a direct indication of mosquito biting potential and can be made quickly,
thus providing a means of short interval pre- and post-treatment sampling.
A minimum of five counting stations should be established for each plot
to be treated and untreated check plots.  These stations should be
arranged in one or more rows perpendicular to swath lines.   Because of
natural fluctuations in mosquito activity due to time of day or night, it
is essential to count mosquitoes in the untreated check plots at ca. the
same time as for the treated plots.  In large-scale tests (400 hectares or
more), a minimum of 10 counting stations should be used (Mount et al. 1972,
Mount and Pierce 1974).

     In small-scale tests, each insecticide dosage should be replicated
at least three times and three or more discriminating doses are needed
to establish the minimum effective dose.  Minimum effective doses at the
90% or the 95% level should be based on either a probit analysis or an eye-
fitted line on logarithmic probability graph paper.

     Test series with new  insecticides should include tests with a
standard insecticide for comparison.  The inclusion of a standard yields
an additional check on mosquito sampling procedures and environmental
conditions present during  testing.

     Furthermore, each test should contain a replication of untreated
mosquitoes to reveal any mishandling or physiological weakness of the
mosquitoes.  Untreated mosquitoes should be exposed to the same handling
and environment (both laboratory and field) as those treated with
insecticides.  Caged mosquito tests in which untreated mosquitoes exhibit
more than 20% mortality should be discarded and repeated.  Corrections
for untreated mosquito mortality from 5-20% can be made by using Abbott's

     In tests with natural infestations, untreated samples should be
replicated equally with each insecticide treatment to reflect variance
in mosquito density due to natural causes.  The % control can then be
determined by the use of Formula II.

     Check plots may not be feasible in some large scale tests against
wild mosquitoes.  In these cases, the efficacy may be obtained by
comparing pre- and post-treatment observations using Formula I given in

Data Analysis, provided pre<- and post-treatment counts are made at the
same time of the day or very close to each other in time,

     Aerial Applications;-^Aerjal applications of aerosols include
ultralow volume and diluted sprays,  The test methods are the same for
both types of application,

     Aircraft nozzles must be maintained and operated according to
recommendations of the manufacturer.  Spray systems must be kept clean
and in excellent operating condition throughout the tests.  The spray
system should be calibrated for insecticide formulation flow rates at
ca. the same ambient temperature as anticipated during test applications.

     It is essential to test the same insecticide formulation to be
registered for use.  This is especially critical with ultralow volume
sprays since droplet size may change significantly with change in viscosity
and surface tension of the formulation.

     When applying insecticides by aircraft, it is essential to correlate
insecticide volume with the area treated to determine the actual dosage
applied.  Aircraft velocity and altitude should be monitored during
application.  Aircraft flight lines (swaths) should be marked to insure
even coverage.  In large scale tests electronic guidance systems could be
used to identify flight lines.

     The efficacy of aerial sprays of insecticides is evaluated by exposing
either caged of free-flying (natural infestations) adult female mosquitoes.
Caged mosquitoes can be from either a laboratory colony or from natural

     Aircraft tests may be conducted over various types of terrain.
Efficacy will be higher in tests over level, open terrain than in uneven,
densely vegetated terrain because of greater penetration and more even
coverage.  An attempt should be made to select test areas that either
resemble or are within actual target areas.

     The minimum test plot size should be 16 hectares for small fixed
wing and rotary wing aircraft flying at relatively low altitudes (15-25 m).
Tests with aircraft flying at an altitude of 46 m or more should be
conducted on plots of at least 400 hectares.

     Meteorological conditions needed for successful aircraft application
are usually found during twilight hours when many species of mosquitoes
are active.  Ground winds should be less than 10 kilometers per hour to
avoid excessive drift away from experimental plots.  Ambient temperatures
near ground level may range from 15-40°C, depending on latitude.  Tests
with caged mosquitoes should coincide with the mosquito season within a
geographic area.

     Mosquito cages and handling techniques used in aircraft spray  testing
are essentially the same as those given previously for ground aerosol
testing.  Cages of mosquitoes should be aligned perpendicular to flight
lines and parallel with prevailing winds.  One-half of the cages of
mosquitoes should be expos,ed at or near ground level,  The other half
of the cages should be exposed at levels of 1-2 m above the ground.  A
minimum of eight cages of 25 mosquitoes each should be exposed for
each replication.  Laboratory reared mosquitoes used in these tests
should be fairly uniform in age within a range of two to eight days old
(Mount et al. 1970, Rathburn et al, 1969, 1971).

     The most commonly used method of assaying aerial sprays has been
the use of pre- and post-treatment landing counts in treated and untreated
plots (Knapp and Gayle 1967, Knapp et al. 1976, Mount and Lofgren 1967,
Mulla et al. 1973, Stevens and Stroud 1967).  If small plots (16 ha) are
used, the applications must be made during the early morning so that post-
treatment counts can be made before possible nighttime reinfestation
takes place.  A minimum of five counting stations should be established
on small plots.  These stations should be aligned in one or two rows
perpendicular to flightlines.  Ten or more stations are needed on large
plots (440 ha or more) for adequate Campling.   Post-treatment counts
should be made at the same time of day as pre-treatment counts since land-
ing rates vary with the time of day.  These counts should be made at
1-12 hours post-treatment with additional counts at 24 hours or longer made
as needed to indicate duration of control.  Other sampling methods that
can be used in area-wide evaluations include New Jersey light traps,
CDC battery traps, C02 traps, animal bait traps, suction traps and
truck traps.

     Each insecticide dosage should be replicated at least two times
and three or more discriminating doses are needed to establish a
minimum effective dose.  At least five samples should be taken per plot.
Minimum effective dosages at the 90% or the 95% level should be based
on either a probit analysis or an eye-fitted line on logarithmic proba-
bility graph paper as given in Data Analysis.

     Tests in which untreated mosquitoes exhibit more than 20% mortality
should be discarded and repeated.  Corrections for untreated mosquito
mortality of 5-20% can be made by using Abbott's Formula,

     In tests with natural infestations, untreated plots should be
replicated equally with each insecticide treatment to reflect variance
in mosquito density due to natural causes.  In each plot, a minimum
of five samples should be taken.  The % control can be determined by
the use of Formula II.

     Check plots may not be feasible in some large scale tests against
wild mosquitoes.  In these cases, the efficacy may be obtained by
comparing pre- and post-treatment observations using Formula I,

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Bailey, D.L,, G.C. LaBrecque, and T.L. Whitfiled.  1970,  Slow-
  release and emulsifiab'le formulations of Dursban  and Abate
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Barnes, W.W., A.B. Webb, and L.B. Savage.  1976.  Laboratory
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Boike, A.H., Jr., and C.B. Rathburn, Jr.  1975.  Laboratory non-thermal
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Brooks, G.D., and H.F. Schoof.  1965.  Simulated field tests of new
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Chapman, H.C.  1955.   Tests with granulated and systemic insecticides
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Dame, D.A., R.E. Lowe, G.J. Wichterman, A.L. Cameron, K.F. Baldwin, and
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Elliot, R.  1955.  Larvicidal control of peridomestic mosquitoes.
  Roy. Soc. Trop. Med.Hyg. Trans.  49(6):528-42.

Gahan, J.B., W.W. Young, N.E. Pennington, and G.C. LaBrecque.  1965.
  Thermal  aerosol and larvicide tests with new insecticides to control
  two species of Culex mosquitoes on Okinawa.  Mosq. News  25(2):165-69.

Georghiou, G.P., and R.L. Metcalf,  1961.  A bioassay method and results
  of laboratory evaluation of insecticides against adult mosquitoes.
  Mosq. News  21(4):328-37

Georghiou, G.P., and F.E. Gidden.  1965,  Contact toxicity of insecticide
  deposits on filter paper to adult mosquitoes,  Mosq.. News  25(2) :204-08,

Gillies, P.A,, E.M, Fuasell, and D,J. Womeldorf,  1972.  Mortality of
  caged organophosphorus-resistant Culex topsails Coquillett using
  various adulticides applied as nonthermal aerosols,  Proc._ Calif.
  Mosq.. Contr. Assoo..  40;22r-25,

Glancey, B.M,, K,F, Baldwin, and C.S, Lofgren,  1969,  Laboratory tests
  of promising mosquito larvicides,  Mosq. News  29(1):41-43.

Hagstrum, D.W. , and M,S, Mulla..  1968,  Petroleum oils as mosquito
  larvicides  and pupicides.  I,  Correlation of physicochemical properties
  with biological activity.  J. Eoon. Entomol,  61:220-25.

Hsieh, M.Y.G., and C.D. Steelman.  1974,  Susceptibility of selected
  mosquito species to five chemicals which inhibit insect development.
  Mosq. News   34(3):278-82.

Jakob, W.L.   1972.  Additional studies with juvenile hormone-type
  compounds against mosquito larvae.  Mosq. News  32(4):592-95.

Jakob, W.L.,  and H.F. Schoof.  1971.  Studies with juvenile hormone-
  type compounds against mosquito larvae.  Mosq. News  31(4) :540-43.

        1972.  Mosquito larvicide studies with MON 585, a juvenile
  hormone mimic.  Mosq. News  32(1):6-10,

Knapp, F.W., and C.H, Gayle,  1967.  ULV aerial insecticide application
  for adult mosquito control in Kentucky.  Mosq. News  27(4):478-82.

Knapp, F.W., W. Carlson, and J. Olson.  1976.  Aerial application of
  propoxur for adult mosquito control.  Mosq. News  36(l):56-58.

LaBrecque, G.C., J.N. Noe, and J.B. Gahan.   1956.  Effectiveness of
  insecticides on granular clay carriers against mosquito larvae.
  Mosq.  News  16(1):1-3.

Ludvik.  G.F.  1953.  Topical application of  insecticide solutions to
  Anopheles quadri-maoulatus.  J. Eoon. Entomol.  45(2) : 364-65 .

Micks, D.S., G.V. Chambers, J. Jennings, and A. Rehmet.   1967.
  Mosquito control agents derived from petroleum hydrocarbons.  I.
  Laboratory effectiveness.  J. Eoon. Entomol.  60:426-29.

Micks, D.S., G.V. Chambers, J. Jennings, and K. Barnes.   1968.
  Mosquito control agents derived from petroleum hydrocarbons.  II.
  Laboratory evaluation of a new petroleum derivative, FLIT MLO.
  J. Eoon. Entomol.  61:647-50.

Mount, G.A., and C.S. Lofgren,  1967.  Ultra low volume and conventional
  aerial sprays for control of adult salt-marsh mosquitoes, Aedes
  sollioitans (Walker) and Aedes taenlorhynchus  (Weidemann) in  Florida.
  Mosq.  News  27(4):473-77-

Mount, G.A., R.E. Lowe, K.F. Baldwin, N.W. Pierce, and K.E. Savage.
  1970.  Ultralow volume aerial sprays of promising insecticides for
  mosquito control.  Mosq.  News  30(3) :342-46.

Mount, G.A., M.V. Meisch, J.T. Lee, N.W. Pierce, and K.F. Baldwin.
  1972.  Ultralow volume ground aerosols of insecticides for control
  of rice field mosquitoes in Arkansas.   Mosq. News  32(3):444-46.

Mount, G.A., and N.W. Pierce.  1972.  Adult mosquito kill and droplet
  size of ultralow volume ground aerosols of insecticides.  Mosq. News

Mount, G.A., and N.W. Pierce.  1974.  Ultralow volume ground aerosols
  of naled for control of Aedes taenioThynchus (Wiedemann) in the
  Florida Keys.  Mosq. News  34(3):268-69.

Mount, G.A., and N.W. Pierce.  1975.  Toxicity of pyrethroid and
  organophosphorus adulticides to  five species of mosquitoes.
  Mosq. News  35(1):63-66.

Mount, G.A., N.W. Pierce, and K.F. Baldwin.  1971.  Effectiveness of
  six promising insecticides for mosquito control.  Mosq. News  31(3):394-96.

Mount, G.A., N.W. Pierce, and K.F. Baldwin.  1975.  Ultralow volume
  ground aerosols of propoxur (Baygon MOS) for control of adult mosquitoes,
  Mosq. News  35(4):490-92.

Mount, G.A., N.W. Pierce, aid K.F.  Baldwin.  1976.  A new wind tunnel
  system for testing insecticidal  aerosols against mosquitoes and flies.
  Mosq. News  36(2):  (In Press)

Mulla, M.S.   1964.  Chemosterilization of the mosquito Culex p.
  quinquefassiatus.  Mosq.  News  24:212-17-

Mulla, M.S., J.R. Arias, R.D. Sjogren, and N.B. Akesson.   1973.
  Aerial application of mosquito adulticides in irrigated  pastures.
  Calif.  Mosq.  Contr. Proa.  41:51-56.

Mulla, M.S., and H. Axelrod.  1960a.  Effect of temperature on rate
  of release of toxicants from granules and on breakdown of certain
  insecticides in water.  Mosq. News  20(2):178-83.

	.  1960b.   Efficiency of granulated  insecticides influenced
  by solvents used for impregnation.  J. Boon. Entomol.  53(5):938-49,

 	.   1962.  The role of carriers in the performance of granular
  formulation of parathion for mosquito control.  J. Eoon. Entomol.
Mulla, M.S., and M.F.B. Chaudhury.  1968.  Ovicidal activity of al:
  tic amines and petroleum oils against two species of mosquitoes.
  J.  Eoon. Entnmnl.  61:510-15.

Mulla, M.S., and H.A. Darwazeh.  1971.  Field evaluation of aliphatic
  amines-petroleum oil formulations against preimaginal mosquitoes.
  Proe. Calif. Mosq. ContT. Assos.  39:120-36,

        1975a.  Field evaluation of aliphatic amines against immature
  mosquitoes.  Mosq. News  35(l):57-62.

	•  1975b.  Evaluation of insect growth regulators against
  Psorophora oonfinnis  (L-A) in southern California.  Mosq. News

Mulla, M.S., H.A. Darwazeh, and G. Majori.  1975.  Field efficacy of
  some promising mosquito larvicides and their effects on nontarget
  organisms.  Mosq. News  35(2):179-85.

Mulla, M.S., H.A. Darwazeh, and R.L. Norland.  1974.  Insect growth
  regulators:  evaluation procedure and activity against mosquitoes.
  J. Soon. Entomol.  67(3):329-32.

Mulla, M.S., L.W. Isaak, and H. Axelrod.  1960.  Laboratory and field
  evaluation of new insecticides against mosquito larvae.  Mosq. News

Mulla, M.S., G. Majori, and H.A. Darwazeh.  1975.  Effects of the insect
  growth regulator Dimilin  or TH 6040 on mosquitoes and some nontarget
  organisms.  Mosq. News  35(3):211-16.

Mulla, M.S., R.L. Metcalf, and A.F. Geib.  1966.  Laboratory and field
  evaluation of new mosquito larvicides.  Mosq. News  26(2):236-42.

Nelson, L.L., T.A. Miller, and W.W. Young.  1973.  Polymer formulations
  of mosquito larvicides.  V.  Effects of continuous low-level chlorpyrifos
  residues on the development of Culex pipiens quinquefasoiatus Say
  populations in the laboratory.  Mosq, News  33(3):396-402.

Raley, T.G., and E.D. Davis.  1949.  Observations on the use of toxic
  briquettes for mosquito control.  Mosq. News  9(2):68-71.

Rathburn, C.B., Jr.  1969.  A laboratory thermal aerosol generator for
  the testing of insecticidal aerosols.  Mosq. News  29(1):1-6.

Rathburn, C.B., Jr., and A.H. Boike, Jr.  1972.  Ultralow volume tests
  of SBP-1382 applied by ground equipment for the control of adult
  mosquitoes.  Mosq. News  32(3):334-37.

Rathburn, C.B., Jr., and A.H. Boike, Jr.  1975.  Ultralow volume tests of
 several insecticides applied by ground equipment for the control of adult
 mosquitoes.  Mosq. News  35(1):26-29.

Rathburn, C.B., Jr., A.J. Rogers, A.J. Boike, Jr., and R.M. Lee.  1969.
  Evaluation of the ultralow volume aerial spray technique by use of
  caged adult mosquoties.  Mosq. News  29(3):376-81.

	.  1971.  The effectiveness of aerial sprays for the control of
  adult mosquitoes in Florida as assessed by three methods.  Mosq. News

Schaefer, C.H., and W.H. Wilder.  1972.  Insect developmental inhibitors:
  a practical evaluation as mosquito control agents.  J. Eoon. Entomol.

Sjogren, R.D., M.S. Mulla, and J.R. Arias,  1973.  Evaluation of mosquito
  adulticides applied as nonthermal aerosols in irrigated pastures.
  Proa. Calif. Mosq. Contr. Assoo.  41:61-66.

Steelman, C.D., J.E. Farlow, T.P. Breaud, and P.E. Schilling.  1975.
  Effects of growth regulators on Psorophora columbiae  (Dyar and Knab)
  and nontarget aquatic insect species in rice fields.  Mosq. News

Stevens, L.R., and R.F. Stroud.  1967.  Control of mosquito adults and
  larvae with ultralow volume aerial applications of Baygon  and Baygon-
  Baytex  mixture.  Mosq. News  27(4):482-85.

Taylor, R.T., and H.F. Schoof.  1968.  Evaluation of thermal and nonthermal
  fogs against four species of mosquitoes.  Mosq. News  28(1):8-11.

World Health Organization.  1975.  Instructions for determining the
  susceptibility of resistance of mosquito larvae to insecticide.

Williams, C.M.  1967.  Third generation pesticides.  Saient. Am.  217:13-17.

Womeldorf, D.J., E.E. Lusk, K.R. Townzen, and P.A. Gillies.  1973.
  Evaluation of low volume nonthermal aerosols for mosquito control in
  California.  Proc. Calif. Mosq. Contr. Assoc.  41:67-73.

                      SIMULIIDAE  (Black Flies)
     Black flies are important vectors of disease and bloodsucking
pests of man and livestock in many parts of the world.  Although they
are not known to carry human disease in the United States, they are at
times abundant and annoying, especially in the forested mountainous
regions where the immature stages breed in clean, unpolluted streams.
They are occasionally pests near large rivers such as the Potomac in
Virginia and, before extensive pollution, in states bordering the
Mississippi River.  In some areas they are serious pests of turkeys
and ducks, interfering with commercial production of these birds
because they transmit lethal or debilitating protozoan parasites.  They
may also reduce milk or meat production of livestock or egg production
of hens when they are abundant (Jamnback 1973).  The construction of
extensive networks of irrigation channels and dams in recent years in
the western United States has provided new breeding sites for nuisance
species (Mulla and Lacey 1975).  The decreasing pollution of many
United States rivers also appears to be favoring a resurgence in
populations of some species of annoying black flies (Sleeper 1975).

     Black flies breed only in moving water of rivers, streams, brooks,
irrigation channels, etc.  The moving water automatically distributes
and transports the insecticide.  Under these conditions, many factors
influence the effectiveness of a given insecticide in addition to its
inherent toxicity.  These include the speed of the current, turbulence
of the stream, duration of exposure to the toxicant, specific gravity,
solubility and  irritant effect (as it influences larval feeding) of the
toxicant (WHO 1968, 1973).

     Black fly larvae are more susceptible to most insecticides than
the eggs or pupae.  They are also restricted to running water, and so
are far less widely distributed than the adults.  For this reason, most
control programs are directed against the larvae.
Laboratory Evaluation

     It should be noted that black flies have not been adapted to
laboratory culture and so studies of their susceptibility to chemicals
are hampered by the need for using wild larvae with all their inherent
variablility.  Laboratory tests, depending on their nature, may evaluate
the inherent toxicity of a pesticide or the effectiveness of a given
formulation.  There are 4 basic techniques:  jar, cloth bag, flushing
and draining, and the trough techniques.

     Jar Technique:—The jar test is especially adapted for testing the
inherent toxicity of a chemical in an acetone or alcohol solution.  It
is less useful for testing formulations which may settle or rise in the
exposure jar.  Clumps of grass or stones with black fly larvae attached
can be collected from streams.  These can be transported between plastic
sheets or in plastic bags.  The larvae must be kept moist, but they
survive poorly if transported in jars or bags partially filled with water.
If they are to be transported long distances in hot weather, they should
be kept in a portable cooler.  Although larvae will survive for several
days, if kept cool in a refrigerator, freshly collected specimens should
be used for testing.  Alternatively, field collected egg masses may be
brought into the laboratory for hatching and subsequent testing of the

     On arrival in the laboratory, the material collected in the field
is immersed in water in large containers.  Samples of the larvae are
gently picked up with forceps, eye dropper, brush or syringe, and
transferred to the test jars.  It is advisable to aerate the large
containers by directing a jet of air against the lower part of the inside
wall.  The larvae will migrate to this zone of maximum aeration, thus
permitting rapid collection.  Medium to large size larvae are required,
but not those of the last stage with dark gill-spots, which are liable to
pupate during the test.  Place approximately 250 ml of clean water,
preferably obtained from the breeding place, into each of 12 cylindrical
glass jars or beakers about 15 cm high and 5 cm in diameter.  The water
in each jar is aerated by a steady stream of compressed air directed
against the bottom of the jar through a glass tube, drawn out into a fine
point.  A standard sample of 25-50 larvae is placed in each test jar.
The larvae become attached to the walls of the jar and to the aeration
tube near its tip.  After about 30-60 minutes, any detached and damaged
larvae are removed with a pipet.

     The insecticide suspensions   are prepared by pipetting the required
quantity of standard solution into jars for a total volume of 250 ml of
solution.  The control should be prepared by the addition of 1 ml of
ethanol or acetone with 249 ml of water.  There should be two replicates
of each concentration and two  for  the  control.

     When the insecticide suspensions have been prepared, aeration of the
test jars is stopped, and the water is poured out *nd immediately replaced
by the insecticide suspension.  The aeration tubes are left in the test
jars during the exposure period (although aeration is not carried out)
since larvae have usually settled on them.  After a 30-minute or other
appropriate exposure period, each test solution is poured into a different
tray, and the jars are rinsed and refilled with water as before.
Detached larvae are collected from the trays and returned to the jars,
which are aerated again throughout a 24-hour recovery period.  At the
end of the 24 hours, the mortality is recorded.  Larvae are considered
dead if they do not move when probed with a needle.  Moribund larvae
may show discoloration, unnatural positions, tremors or inability to
stay attached.  Each dosage should be run in duplicates along with
duplicate controls on 3 different occasions.  The LC^g'3 an
     All larvae that have pupated during the test are discarded.  If
more than 10% of the control larvae have pupated, or if control mortality
is 20% or more, the test should be discarded.

     The living and dead larvae exposed to each insecticide concentration
should be preserved separately in 70% alcohol.  After they have been
identified by microscopic examination, the mortality for each species is
recorded.  The susceptibility level of any one species is regarded as
adequately tested when 100 or more larvae have been exposed to each
concentration (WHO 1975).

     Cloth Bags Technique:—A modification of the jar test involves
the use of cloth bags.  About 30 larvae are transferred individually
with a broad-tipped medicine dropper from a holding pan to a white organdy
cloth bag.  Only uninjured normal larvae are selected.  The bag
containing larvae is then closed and fastened at the top with a wire pipe
cleaner.  The bags are immersed for 20 minutes or other appropriate
period in jars containing 3,000 ml of nonchlorinated, nonaerated water
to which an appropriate amount of pesticide in acetone solution has been
added and mixed just prior to the test.  Acetone is added to the check
jar in a volume equal to the greatest amount of acetone solution added to
any of the treated jars.  Alternatively, small disposable paper cups may
be used (Jamnback and West 1970).  After 20 minutes, or longer period if
necessary, the bags are rinsed in fresh water and transferred to 3,000 ml
of untreated water.  The mortalities are determined after a 24-hour
holding period in the aerated water.  A minimum of two replicates of each
of three to five discriminating concentrations of insecticide are run
and the average are used in determining the dosage-response line and the
LCcQ and LC^Q with confidence limits.

     Laboratory tests with insect growth regulators can be carried out in
the same way as for conventional larvicides, except that after exposure,
larvae are reared to adults with larval, pupal and adult mortalities or
abnormalities recorded.  Troughs may be particularly useful for IGR's
because the larvae survive for long periods with less mortality than in
jars.  Testing to date has been confined to jars and long exposure periods
of 24 hours or more (Dove and McKague 1975).

     Flushing and Draining Technique:—This technique employs features
of both the jar technique and to some extent that of the trough technique.
The use of this technique approximates field conditions much more so than
the jar technique (Mulla and Lacey 1976).  This system can be used for
rearing of Si-mutium larvae, as well as bioassay of short-term and delayed
acting larvicides.

     The test units consist of glass tubing (10 cm diameter) connected to
pyrex glass funnels.  The capacity of the unit approximates 1 liter.
These glass units are provided with inflow arms (allowing water flow from
a water line or reservoir) and outflow arms situated about 2-5 mm below
the level of the inflow orifice on the opposite side.  An air stone is
affixed into the unit 10-15 mm above the edge of the funnel.  Stem of the
air stone is connected to air coming from an air pump or compressed air.

     An important feature of these units is drainage facilitated through
the stem of the funnel pointed downward.  A rubber tubing, with a hose-
cock clamp fitted over the funnel stem, can regulate the amount of
water drained out.  When not draining, the clamp can be tightened to shut
the water off from draining.

     If glass tubing is not available, the test units can be made from
pint-size polystyrene cups.  The bottom of the cups is removed, the
narrow end glued to polyethylene funnel using silicone caulking compound.
The overflow, inflow, aeration and drainage tubing are incorporated into
the units as described for the glass units.

     Water current in the units is provided with air passed through an
air stone.  The rearing chambers are placed in batteries of 4-6 units
in wooden racks in a room maintained at 20-25°C.

     For evaluation of quick and slow acting larvicides, 20-30 larvae of
a given instar are placed in each unit, provided with 35 mg of food
(2% aqueous suspension of ground up lab chow and brewer's yeast 3:1).
After 4-24 hours of equilibration, dead and diseased larvae and pupae
are removed.  The surviving larvae are counted, the units are flushed
with excess water for 5 minutes at the rate of 1 liter/min., in such a
way that water is drained both from the overflows orifice and the drain
stem of the funnel.  After flushing and draining (larvae are never exposed
out of water in this process), the larvae are provided with 35 mg of food.
One hour later, the units are treated with various concentrations or
different materials.  After appropriate exposure period (60 minutes for
quick acting larvicides, longer periods for slow acting agents), the
active agents are eliminated from the units by flushing with excess
water (3 liter/min) for 5 minutes.  After flushing, the larvae are
provided with 35 mg of food.  Once or twice daily, the units are flushed
and  drained (1 liter/min) for 5 minutes and provided with food.
Flushing and draining eliminates pollutants and toxic metabolites which
induce mortality in blackfly larvae.  It should be pointed out that in
the flushing and draining process, the units remain full all the time
and the larvae do not get exposed out of water  (Mulla and Lacey 1976).

     Mortality is assessed either 24 or 48 hours after treatment in the
case of quick acting compounds.  For the evaluation of Bacillus
thuringiensis and insect growth regulators and antimetabolites, the
mortality or level of emergence may be assessed 1-2 weeks after treatment.

     The dosage response line, using 3-4 discriminating dosages, is
established and the LCrgand LCgn levels are determined.  If check mortality
is more than 20%, the test should be discarded.  For check mortalities
greater than 5%, data should be corrected with Abbott's Formula.

     Trough Technique:—Trough tests are useful in evaluating the
effectiveness of formulations particularly in shallow water3 but require
modification for testing effectiveness in deeper water.  Because the
troughs are short, settling is minimal and so "carry", i.e., the distance
downstream that an insecticide remains effective is not evaluated.

     Sticks, stones, leaves, etc. with attached larvae are collected as
outlined above and placed in a trough, typically 1 m long, 0.5 m wide and
15 cm deep, through which running unchlorinated water is flowing.  The
larvae migrate to a lip at the downstream end of the trough about 15 cm
wide and 10 cm long.  Water flows (21 cm/sec.) over this lip in a narrow
sheet 0.5-0.8 cm deep through which the reactions of individual larvae to
insecticide can be closely monitored.  Known concentrations of larvicide
are poured or dripped into the upper end of the troughs over standardized
exposure periods.  Larval behavior and detachment rates are recorded.
Large numbers of formulations can be subjected to preliminary screening,
sufficient tests and replicates can be carried out to establish accurate
50% detachment levels using procedures commonly applied to mosquito
larvicide testing (Jamnback and Frempong-Boadu 1966, Frempong-Boadu 1966).

     A few larvicides have a knockdown effect that is followed by
subsequent recovery.  These can be detected by using two troughs in a
series with the lower trough (downstream) initially without larvae and
determining whether larvae that are dislodged from the upper trough
detached and carried to the second trough can recover and reestablish
themselves.  Alternatively, detached larvae can be captured in a cloth
bag or screen and held for 24 hours in aerated water to determine whether
or not they recover.  There are also a number of variations of the trough
test technique.

     Larvae can be tested in sloping v-shaped gutters supplied with water
by a pump or gravity.  These are typically about 2 m long and 4 cm deep,
made of sheet metal and are v or u shaped.  The rate of flow can be varied
by altering the angle of the gutters.  The gutter is lined with a sheet
of polyethylene film. This is covered on top with brown wrapping paper, a
good attachment material for larvae.  The polyethylene and paper are
disposable.  Water swirls through funnels into the upper end of the gutter,
insecticide is added by dripping into the funnels where it thoroughly
mixes with the water.  Larvae that detach are captured in a screen at the
bottom of the trough and held in a bag in a nearby stream for 24-48
hours mortality counts.  Larvae that do not detach are dislodged and
collected in nylon sieves which can be folded to form bags and held in a
stream for 24-48 hours evaluation (Wilton and Travis 1965).
Field Evaluation

     Single Stream Evaluations:—Promising larvicides can be tested in
single stream field tests.  The formulation is poured directly into the
stream, applied by dispenser or sprayed over the water.  In all cases,
application into a turbulent portion of the stream increases the likelihood
that it will mix well and become effective at, or slightly below, the
point of application.  Emulsions are widely used because of the ease
with which they can be dispersed through the water, wettable powders
disperse well, but tend to settle quickly and so have little "carry";
solutions must be sprayed as fine droplets into streams for maximum
effectiveness, either by hand or power sprayer from the ground or from

aircraft.  The cross-section and rate of flow of the stream must be
determined to calculate the concentration in parts per million per unit
time.  It should be noted that swiftly flowing turbulent streams may
require higher concentrations than those required for equivalent control
in slow streams with stretches of slack water.  In many cases not enough
larvicide will traverse a large pool to remain effective at the pool
outlet even at high concentrations (WHO 1968,1973).

     Ground application is typically used when there are one or a few
accessible streams to be treated and aerial application when there are
many and/or inaccessible streams which require treatment.  With the ground
application technique there is an initial overdose but the gradually
diminishing concentration remains effective for many miles below the single
point of application.

     Effectiveness can be evaluated by comparing larval populations above
and below the treatment point not more than one day before and one day
after treatment.  Longer periods allow larval drift, egg hatching and
pupation of black fly larvae to obscure the data.  Samples should be taken
at various distances below the point of treatment to determine how far
downstream below the point of application the toxicant becomes effective
and beyond that point how far downstream it remains effective (Jamnback
1969).  Effectiveness is evaluated by larval counts, either on a standardized
number of sticks, stones, trailing grass, etc. or preferably on
standardized attachment units, e.g., plastic tags, strips, cones or ceramic
tiles that are placed in suitable black fly attachment sites in the stream
several days prior to the pre-treatment evaluation (Lewis and Bennett

     Additional experimental single stream tests should be carried out
in a variety of stream sizes and types, e.g., rivers, creeks and even
small temporary brooks, both slow and fast, turbulent and less turbulent,
with and without pools, depending on the breeding habits of the black fly
species under attack.  In all cases, streams with relatively high and
uniform propulations should be used as this simplifies evaluation of
effectiveness, especially "carry".  If possible, a variety of formulations
should be tested to determine those which are the most effective.
Laboratory Evaluation

     Black fly adults typically have high mortality rates in captivity
and have not been tested for insecticide susceptibility although a
satisfactory method can undoubtedly be developed using a modified WHO
susceptibility testing kit for mosquitoes.  At present, systematic
research on adult control of black flies is a much neglected field,
however, there have been a number of unpublished observations that some
nonpersistent insecticides applied by aircraft as sprays or from
truck-mounted foggers may provide short term protection from black fly
attack, but these have not been systematically evaluated.

     Effectiveness of large scale adult control operations and area-
wide larvae control operations can be evaluated by population estimates
inside and outside of the control area before and after treatment.
Landing rates can be made using an 18-inch square piece of dark blue
flannel spread on the ground beside the observer (Davies 1951),   Where
black fly populations are high, a one-minute count beginning after a
five-minute waiting period is satisfactory.  Alternatively, 10 directed
continuous standardized sweeps about the head of the observer after a
five-minute wait will provide a useful index to the annoyance rate.
Counts should be taken simultaneously by observers outside and inside
margins of the treated plot.  Records of air temperature, wind velocity
and direction, and light intensity should be taken.  Low temperatures
below about 12°C and wind velocity above 10 kilometers greatly reduce
black fly activity.  After dusk black fly attack rates decrease quickly.
They do not attack man at night.  Light traps, sticky traps, C02 traps
and silhouette traps may catch black flies, sometimes in large numbers,
but require far more work and cost more than the landing rate or net
methods and have the disadvantage that nonnuisance species are collected
(sometimes in large numbers), as well as those that attack animals.
Davies, D.  1951.  Some observations of the number of black flies
  (Diptera, Simuliidae) landing on colored cloths.  Can. J. Zool.

Dove, R., and B. McKague.  1975.  Effects of insect developmental
  inhibitors on adult emergence of black flies  (Diptera:  Simuliidae).
  II.  Can. Entomol.  10.7(11): 1211-13.

Frempong-Boadu, J.   1966.  A laboratory study  of the effectiveness of
  methoxychlor, fenthion and carbaryl against blackfly  larvae  (Diptera:
  Simuliidae).  Mosq. News  26(4):562-64.

Jamnback, H.  1962.  An eclectic method of testing the  effectivess of
  chemicals in killing blackfly larvae (Simuliidae:  Diptera).  Mosj. News

Jamnback, H.  1969.  Field tests with larvicides other  than DDT  for
  control of blackfly (Diptera:  Simuliidae) in New York.  Bull,  VHO

Jamnback, H.  1973.  Recent developments in control of  blackflies.
  Ann. Rev. Entomol.  18:281-304.

Jamnback, H., and J. Frempong-Boadu.  1966,  Testing blackfly  larvicides
  in  the laboratory and in streams.  Bull.  I'lEO 34:405-21.

Jamnback, H.,  and A. West,  1970,  Decreased susceptibility of blackfly
  larvae to p,p  DDT in New York State and Eastern Canada.  J. Econ.
  Entomol*-  63:218-21,

Lewis, D., and G, Bennett.  1974.  An artificial substrate for the
  quantitative comparison of the densities of larvae Simuliid (Diptera)
  populations.  Can. J. Zool.   52(6):773-75.  RAE(B) 63(2).

Mulla, M., and L. Lacey.  1975.  Biting flies in the lower Colorado
  River Basin:  Economic and public health implications of Simuliwn.
  (Diptera-Simuliidae).  Proa.  Calif.  Mosq. Control Assoo. 43.:  (In

Mulla, M., and L. Lacey.  1976.  A rearing and bioassay technique for
  black fly larvae.  J. Eaon.  Entomol.  (In Press).

Peterson, D.,  and A. West.  1960.  Control of adult black flies (Diptera:
  Simuliidae)  in the forests of eastern Canada by aircraft spraying.
  Can. Entomol.   92(9): 714-19.

Sleeper, F.  1975.  Visit from a small monster.  Sports Illustrated.
  Aug. 25:  46-49.

Wilton, D., and B. Travis.  1965.  An improved method for simulated
  stream tests of blackfly larvicides.  Mosq. News  25:  118-23.

World Health Organization.  1968.  Joint US-AID/OCCGE/WHO Technical
  Meeting on the Feasibility of Onchocerciasis Control.  Tunis, 1-8,
  1968.  PD/68.8:  60 pp.

World Health Organization.  1973.  Onchocerciasis Control in the Volta
  River Basin.  OCP/73.1 Annex III-4:   8 pp.

World Health Organization.  1975.  Provisional Instructions for Deter-
  mining the Susceptibility or Resistance of Blackfly Larvae to Insec-
  ticides.  VBC/75.591:  5 pp.

                 CHIRQNOMIDAE AND CH/LQBORIDAE  (Midges)
      Aquatic  midges  include species  of  the  family  Chironomidae  and
 Chaoboridae.   The chironomids  form a large  group of  insects,  the  immature
 stages  are mostly aquatic  and  provide an ample  source  of  food for aquatic
 vertebrates and  invertebrates.   They breed  in lakes, reservoirs,  ponds,
 streams,  ditches and rivers.   The  chaoborids form  a  small group of insects,
 their immature stages are  also  aquatic,  they feed  on zooplankton  and in
 turn  are  used as food by many  aquatic organisms.

      The  aquatic midges have become  a serious nuisance problem  in some
 urban,  resort and industrial communities.   They pose a serious  economic
 and nuisance  problem in resorts, newer  sub-divisions encroaching  on
 eutrophic bodies of water (Grodhause 1963, Mulla  1974).  Their abatement
 in some communities  is an  essential  part of environmental health  programs.
 Control of larvae in the aquatic habitat is the most commonly used practice.
 Control of adults resting  in or on vegetation and  structures  is also practiced
 in some communities.
                        CHIRONOMID MIDGES
     Larvicides include  those compunds which act quickly on the larvae,
producing mortality at practical concentrations or dosages in 24-48
hours.  In contrast,  IGR's, materials which induce delayed mortality
or disrupt the developmental traits of the larvae, are slow acting
materials.  Their evaluation requires special and different techniques
which will be discussed  later.

Laboratory Evaluation

     Conventional Larvicides:—Biologically active compounds can be
evaluated against chironomid midges as technical or formulated materials.
The technical material should be dissolved in acetone or ethyl alcohol
as 1% solution and further serial dilutions should be made in the same
solvent if necessary.  Formulated materials such as WP  (wettable power),
EC  (emulsifiable concentrate) or flowable concentrate (FC) formulation
should be diluted with water or other appropriate diluents.  Solutions
of toxicants in organic  solvents should be kept refrigerated when not in
use.  Preparation of formulated materials in water should be made fresh.

     Larvae of uniform size (third or early fourth stage) should be
obtained from field populations or laboratory colonies.  Field larvae
may be obtained by taking bottom mud samples using an Ekman dredge or
another type of scoop.   The sample is stirred  in water, sieved through

a 50-mesh or another appropriate sieve to remove most of the mud and
detritus.  The larvae will remain on the top of the screen.  The larvae
are washed with water from the screens into cups, placed in cool ice
chest and brought into the laboratory,  Care should be taken to avoid
direct contact between larval containers and the ice in the ice chest
(Ali and Mulla 1976, Anderson et al. 1964, Mulla et al, 1973),

     In the laboratory, the sample is transferred to white enamel pans
or other appropriate containers.  Larvae of the desired stage and species
are picked up by a screen loop, tweezers, aspirator, eye dropper or pipet
and placed in the test units.  They may also be recovered by flotation
using saturated aqueous solution of Epsom salt (see below).

     Several types of containers may be employed as test units.  For
ease, disposable treated paper cups (ca. 100 ml) are normally employed,
as these are discarded after each test.  Custard cups, ointment jars,
beakers and other glass containers may be used, but these have to be
thoroughly washed and decontaminated after each use.

     The test units are provided with 100-200 ml (depending on the
size of the unit) of tap or field water.  It is essential that 5-10
g of dry standard masonry sand or other fine sand from a stream, river
or sand dune is added to each unit.  Most chironomid larvae are tube
builders and the sand will provide the necessary material for tube
building.  Provision of sand also reduces cannibalism in the tube-
building species (Mulla and Khasawinah 1969).  Twenty uniform size
larvae are transferred to each unit.  The units are then treated within
1-2 hours after addition of larvae with the desired concentrations of
the toxicants.  Several concentrations of each toxicant should be evaluated
for establishing dosage response line.  Each concentration should be
replicated at least two times and each treatment should be run on at least
three different occasions, yielding a total of six replicates per concen-
tration.  Check units receiving the highest amount of the solvent or
diluent should be run along with each test.  The test units should be
kept at temperatures approximating those to which they are accustomed.
Increased of 5-6°C over the normal temperature can be tolerated by many

     Mortality should be assessed 24 hours after treatment.  For this
short period, it is not necessary to feed the larvae.  The average
mortality after: correction (if needed) for each concentration is cal-
culated as given under Data Analysis.

     Care must be exercised in distinguishing dead and moribund larvae
from living larvae (Mulla and Khasawinah 1969) .  Some of the criteria
employed in distinguishing dead and moribund larvae are:

1.  Lack of movement when touched with a dissecting needle,

2.  Moribund larvae, although showing some indication of movement,
are unable to return to their natural position.  They usually are curved
and unable to make undulating movements.

3,  In pigmented red larvae, some areas become deeply colored., while
others develop light colored areas, thus showing blotchy appearance,

     Insect Growth Regulators and  Antimetabolites:—Insect growth
regulators and antimetabolitea, unlike standard larvicides, act slowly
and induce delayed response in the treated populations,  In the case
of some of these compounds, the effects may not be realized until the
treated larval population has either pupated or is undergoing emergence.
Therefore, the standard larval bioassay techniques will not provide
the desired information.  The assessment techniques discussed below will
be applicable to all bioactive compounds inducing delayed response in
the population.  In most assessment procedures, the development of the
treated or untreated population is scrutinized to the time of the emer-
gence of the test individuals.

     Technical or formulated materials may be evaluated.  The technical
material is dissolved in acetone or ethyl alcohol, making a 1% solution.
Formulated materials are diluted with water.  Serial dilutions are pre-
pared as needed.  The stock solutions should be stored in a refrigerator
when not in use.  In no case more than 1 ml of the solution should be
added to the test units.

     Since testing of IGR's and similar compounds takes longer (5-10
days)  than that of standard larvicides, only glass containers should
be used for testing.  Half-pint wide-mouth Mason jars or glass custard
cups are ideal units.  The units are filled with 200 ml water (tap or
field), provided with about 5-20 g of dry masonry quartz sand or fine
sand obtained from a stream, river or sand dune.  Twenty third to fourth
instar larvae from a laboratory colony or field collected material are
transferred to the units by means of screen loops, eye droppers or syringe.
The larvae are allowed to burrow into the substrate.  The units are
aerated by forcing air (50-60 ml/min.) through a hypodermic needle in
each unit.  The units are provided with food prior to treatment and then
once every other day.  TetramiriS1 fish food has been found satisfactory.
Many other types of high protein food may be substituted.  The food is
suspended in water and about 1 ml of the food suspension containing
50 mg of the dry food is added to each unit each time.  Each treatment,
including check, is replicated at least two times and the tests are
repeated on three different occasions (Mulla et al. 1974) yielding six re-

     The larval units are then treated with the dilute preparations
of the test materials.  The units are kept at temperatures approxi-
mating, or somewhat higher than that of normal populations and followed
until most of the population has emerged in the check units.  Emerging
adults are collected in small plastic cups provided with a screen patch
to allow for ventilation.  They are counted and removed daily or every
other day.  Only adults under-going complete eclosion are counted as
going through normal emergence.  In tests where emergence is less than
50% in the check units, the test results should be discarded.  In this
manner, the inhibition of emergence in the treated and check units can be

The inhibition of emergence for a given treatment is calculated by the
procedure presented under Data Analysis,
Field Evaluation

     Chironomid midges causing nuisance problems in urban situations
propagate in ponds, reservoirs, residential-recreational lakes, store
drains, flood control channels, sewage disposal systems and other water-
carrying structures.  Some of the midges are benthic where the larvae
reside in tubes constructed from detritus, sand and other particles,
or they swim freely in the bottom ooze.  Larvae of some species attach
to plants and other submerged substrates.  Therefore, different sampling
techniques have to be employed for these larvae, depending on where they
are found.

     Conventional Larvicides:—For field evaluation, chemical control
agents may be employed as liquid or solid formulations,  The liquid and
wettable powder formulations are diluted with water and sprayed onto
the water surface with hand or power sprayers,  Each treatment should
be replicated at least two times (if possible) and at least three to four
discrete larval samples taken from each treatment.  At the same time,
check plots should be run along with the treated plots.  If it is not
possible to run check plots, then the whole body of water should be treated
as one plot.

     Benthic larvae are sampled by means of an Ekman grab or dredge
(15 x 15 cm) from the deeper portions of water or by a scoop that can be
operated from the shoreline.  The mud samples are stirred in water and
sieved through a 50^mesh or other appropriate sieve.  The residue on
the screen is transferred with water to pint cups with lids and placed
in a cool ice chest for transport to the lab.  The cups containing larvae
should not come in direct contact with ice in the chest (Mulla et al.
1971, 1973).  The larvae are floated by adding 100-200 ml of saturated
aqueous solution of Epsom salt (see below).

     Free-swimming and drifting larvae (in flowing water) are sampled
by dipping (Norland and Mulla 1975) or tow nets provided with a col-
lection vial.  The dipping samples are taken close to vegetation where
larvae abound.

     In flowing water, the sampling technique consists of placing tubes
or cloth bags which are colonized by midge larvae, and these are
retrieved when needed for larval assessment  (Polls et al. 1976).  After
concentrating,  these larvae are transferred to cups and transported to
the laboratory in the manner described above,  Surber samplers or modified
Surber samplers have also been employed in sampling larvae in shallow
(less than 50 cm deep) flowing water (Ali et al, 1976),  The frame of the
sampler is placed on the substrate which is scraped and stirred in the
flowing water.   The larvae and substrate are collected into a fine-mesh

net, from where it is transferred into cups, transported to the labora-
tory.  The larvae are floated and counted in the manner described above,

     The larval samples, after transport to the laboratory, are placed
at room temperature for one hour.  The larvae are then floated by add-
ing 100-200 ml of saturated aqueous solution of Epsom salt (MgS04).
The larvae are counted under a magnifying lamp.  They should be iden-
tified to the genus or species, whichever identification is convenient.

     Larvae attached to macrophytes and other collectable substrates
can be sampled by removing a standard quantity (volume or weight) of
the substrate.  The substrate should be stirred vigorously in a bucket
of water and then discarded.  The dislodged larvae and detritus in the
bucket are poured through a 50-mesh or other appropriate sieve.  The
residues on the sieve are transferred with clean water into a cup.
The cups are transported in a cool ice chest to the laboratory where
the larvae are floated and counted in the manner described above.

     In semi-field evaluation utilizing small shallow ponds, larvae
are sampled by isolating bottom mud in 15 X 15 cm baking pans placed in
the natural habitat for 3 to 4 days.  At desired intervals, 3 to 4 of
these trays containing mud and larvae are retrieved.   The sample is washed
through the sieve and the larvae are transported to the laboratory in
the manner described above (Mulla and Khasawinah 1969).  In further
refinement of this technique, sheet metal enclosures (.6-1 m in diameter)
have been used for preliminary evaluation of larvicides against field
population.  In this regard, the enclosed water and bottom mud is considered
as one plot, wherein several trays of bottom mud are placed.

     For detailed and critical examination, the larvae and the residues
may be transferred to enamel pans or lit counting trays, where they are
counted and identified to genus or species.

     In assessing the biological activity, larval samples should be
taken prior to and 4, 7, 15 days or longer after treatment when the
effectiveness is worn out.  It is important that three to four days be
allowed between treatment and the first assessment, as it takes this
period of time for dead larvae to decompose or lose turgidity and form.
Extent of control in the treatments, run along with checks or without
checks, should be calculated in the manner described under Data Analysis.

     Insect Growth Regulators and Antimetabolites;—Two approaches
are used in semi-field and field evaluation of IGR's, antimetabolites
and other delayed acting bioactive compounds against chironomid midges
in aquatic habitats.  Since some of these compounds induce delayed
mortality in the larval stages, assessment of larval population, as
described under Conventional Larvicides will provide the needed infor-
mation on the action of these types of compounds on midge larvae.

     For assessment of the efficacy of most of these compounds, how-
ever, it will be necessary to determine the inhibition of emergence
of adults.  Emergence is assessed readily by setting submerged, emerged
or floating emergence straps.  For shallow waters ,3^6 m deep, cone
type sheet metal traps provided with a collection jar or chamber are
employed (Mulla et al, 1974, Mulla et al, 1975a, Norland and Mulla,
1975).  These will also work in shallow, flowing water.

     To evaluate the efficacy of a given IGR or similar acting compounds,
field plots should be treated with EC, WP, FC or granular formulations
of the compound.  The firs't three formulations should be diluted with
water and sprayed onto the surface of the water.  Granular formulations
are applied by a broadcaster mounted on-a boat or on an airplane.  If the
body of water lends itself to be divided into plots, check plots should
be run along with the treated plots.  In some situations, it may not be
possible to divide the habitat into plots.  In these situations, the whole
body of water is treated.

     For assessment of efficacy, emergence of adult midges is obtained
prior to treatment and then at intervals after treatment.  Emergence
traps should be placed in the treated as well as check plots if included
in the design of the experiment.  The emergence of midges as a function
of time is plotted and the trend of midge production as influenced by
the treatments can be ascertained until emergence level reaches pre-
treatment level.  If check plots are run along with treated plots and pre-
and post-treatment samples are taken, the inhibition of emergence (El)
for a given period is calculated by Formula II given under Data Analysis.
If no pre-treatment samples are taken, then, the El is calculated by
Formula III.  However, if the whole body of water is treated as a single
plot, then the extent of efficacy (El) is determined by comparing emergence
during post-treatment intervals with that during the pre-treatment
period, using Formula I.

     It is important that at least four traps be used per plot or 12
traps per whole body of water.  In general, plotting the emergence of
midges from check and treated plots yields the needed information.
Similarly, emergence during pre-treatment and post-treatment intervals
provides a basis for assessing effectiveness of IGR's and similar types of
compounds.  The emergence assessment should be continued until it reaches
to 30-50% of the emergence prior to treatment or the emergence in the check.
     As discussed earlier, adulticiding measures are employed on a
limited scale for the control of midges.  Nevertheless, many quick
knockdown and effective insecticides have become available recently
and these offer good tools for the control of adults.

Laboratory Evaluation

     No techniques have been developed specifically for the evalua-
tion of adulticides against chironomid adults.  The techniques and
protocol presented for the evaluation of mosquito adulticides will
serve well for midge adulticides.  It should be pointed out that due to
the short life span of midges, adults not older than 24 hours be used
in bioassays and that the mortality be read 4-8 hours after treatment.
Otherwise, the procedures given for mosquitoes should apply equally to
adult midges.
Field Evaluation

     Aerial and ground application of aerosols and mist sprays provide
temporary relief from nuisance midges.  Normally, the adult midges rest
in vegetation, on structures and other protective sites.  Discharge of
an adulticidal aerosol will yield high mortality of the resting adults.
The swaths should be applied in such a way that the aerosol cloud is
carried over the harborage sites by wind.

     Assessment of adult populations should be made before and at inter-
vals after treatment in the check area and the treated area.  After
population assessment, the level of reduction or control should be
calculated by Formula I if no check plots are used, or by Formula II
if check plots were established.

     Adult populations may be sampled by one of the following techniques:

1.  Insect sweep nets.  Resting sites, especially vegetation, lend
readily to this technique.  At least five samples of five sweeps each
should be taken at four to five locations at each sampling time.

2.  Counting on substrates.  Adult midges rest on leaves, stems of
plants and on surfaces of buildings and other protected niches.  Counts
should be made at the same time of the day and converted to numbers/
unit area (Patterson et al. 1966).

3.  Shelters.  Artificial standard shelters can be made and placed in
the infested area.  During the heat of the day, adult midges will enter
the shelters where they can be counted easily.

4.  Light traps.  The New Jersey light trap and other similar traps will
attract midges during the night.  Trap collections give valuable infor-
mation on the population activity and levels of adult midges.  Several
traps should be employed in the check and treated areas prior to and at
intervals after treatment.  The light traps should be located away from
other artificial lights.  The procedures outlined for mosquito adults
will be applicable here too.

                        CHAOBQRID MIDGES
     These midges breed in eutrophic lakes ?  ponds and reservoirs and
at times are co-existing with chironomid midges.  The chaoborid larvae
are relatively transparent and hard to see in water, and hence, they
are also known as "phantom larvae,u

     Chemical control measures are primarily aimed at larvae by treating
the aquatic habitat where the larvae are prevailing.  In temperate zones,
the midges become a nuisance problem in June'-September and treatments,
therefore, are necessary during this period.  At most, three to four
chemical treatments are needed during a given season.

     Larviciding is the most effective measure for the control of
Chaoborus species.  Adult control has not been found practical and
effective and, therefore, is not practiced except under certain cir-
cumstances.  Equipment and  methodologies employed for the control
of adult mosquitoes and chironomid midges will be equally applicable
here.  Assessment of adult populations should be made in the manner
described for those two groups.

     Eggs of these midges float mostly on the water surface and are
blown as drifts by wind.  Treatment of egg drifts with petroleum oils
(high aromatic content) has yielded good control of eggs (Dolphin and
Peterson 1960).  Notwithstanding this, ovicidal measures are not practical
as a good portion of the eggs sink and hatch in the deeper portions.
Laboratory Evaluation

     ChaoboTus species have not been colonized in the laboratory as
yet; therefore, larvae for bioassays are collected from their breeding
sources.  Young instar larvae (first and second) are pelagic, occupying
the upper portions of a lake, pond or reservoir, while the older larvae
(third and fourth) are benthic, occupying the deeper portions, primarily
residing at the mud and water interface during the daylight hours and in
winter months.  The  younger larvae are collected by plankton-tow nets
drawn vertically or horizontally through the water.  The older larvae
and pupae are sampled by means of an Ekman dredge or similar device.
Larvae collected by tow nets are transferred to the laboratory in the
lake water in a chilled ice chest.  Mud samples secured by the dredge
are washed through a SO^mesh screen, the residue containing larvae are
transferred in water to jars or cups and transported in a cool ice
chest to the laboratory (Brydon 1956, Cook 1965),

     For bioassay, the larvae from the mud residue are floated with
saturated solution of Epsom salt  (magnesium sulfate),  The larvae are
transferred to filtered or screened lake water in beakers or fruit jars,
Normally, lQOr-200 ml water is placed per unit, to which 20-25 larvae
are transferred by pipet, screen  loop or an aspirator.  Both first and
fourth instars are used in separate tests.

     Technical insecticides are dissolved in acetone as 1% solution,
serial dilutions when needed are made with water.  Formulated fluid
or wettable powder formulations are diluted with water and serial dilutions
made in water.  Aliquots of the solution to yield the desired concentra-
tions are added to the test units containing the larvae.  Each concentra-
tion should be replicated two times and each test repeated at least three
times on different occasions.  The units are held in a room with a tem-
perature of 20-25°C.  Checks are  run along each test to which the solvent
alone is added.  Chaoborid   larvae are susceptible to acetone and other
solvents; therefore, the quantity of solvent added per unit should not
be more than 0,1 ml.

     Larval mortality is assessed 24 hours after treatment under a
microscope; larvae with no movement are considered dead.  Mean larval
mortality is plotted against concentration on a probit log paper and the
points fitted with a straight line.  LC^Q and LC^Q concentrations are
read off this line or from a computer analysis of the data.

     Larvae of Chaoborus can survive for 24-48 hours without food.
Longer holding of larvae requires provision of food in the form of live
zooplankton, as the larvae are predacious.  For the evaluation of IGR's,
antimetabolites and other slow-acting bioactive compounds, evaluation
procedures in the laboratory are yet to be developed.  These procedures,
however, will be quite similar to those employed in mosquito and chironomid
midge studies, except that live food items have to be offered to the
larvae at regular intervals during the test period.
Field Evaluation

     Field evaluation of larvicides, IGR's and other bioactive compounds
against the larvae is quite similar to that presented for chironomid
midges.  For methods of application and dilutions, see those procedures.

     Larval samples are taken in the manner as described under Labora-
tory Evaluation for obtaining live larvae for laboratory bioassays.
For counting, the field larval samples are heated to 55-60°C for a few
minutes.  This temperature will kill the larvae recovered from the
plankton tow-nets or bottom mud samples.  The dead larvae float to the
top of  the water in the jar from where they can be decanted, aspirated
or screened out and counted in a dark photographic tray  (Cook 1967,
Hazeltine 1962, Lindquist et al. 1951, Snell and Hazeltine 1962),

     The larval counts prior to and after treatment are compared for
level of control.  The % reduction is calculated by Formula I? as reported
in Data Analysis,  If check plots are run along with the treated plots,
then the level of control or reduction is calculated by Formula II,

     Adult emergence can be assessed by a submerged emergence trap as
used for chironomid midges or by a floating type trap.  The level of
inhibition of emergence is to be calculated in the manner described for
chironomid midges,
Adulticiding measures are employed on a limited scale for the control
of midges.   Many quick knockdown and effective insecticides have become
available and these offer good tools for the control of adults.
Ali, A., and M.S. Mulla.  1976.  Chironomid midge productivity in relation
     to depth in a reservoir-percolation basin.  Environ, Entomol.
     (In Press).

Ali, A., M.S. Mulla, and F.W. Pelsue.  1976.  Removal of substrate for
     the control of chironomid midges in a concrete lined flood control
     channel.  Environ.  Entomol.  (In Press).

Anderson, L.D.,  E.G. Bay, and A.  A. Ingram.  1964.  Studies of chiro-
     nomid midge control in water-spreading basins near Montebello,
     California.   Calif. Vector Views  2:13-20.

Brydon, H.W.  1956.  The Clear Lake gnat and its control in Clear
     Lake, California during 1954.  J.  Econ. Entomol.  49:206-09.

Cook, S.F. Jr.  1965.  The Clear Lake Gnat:  its control, past,
     present and future.  Calif.  Vector Views  12:43-48.

        1967.  The increasing chaoborid midge problem in California.
     Calif. Vector Views  14:39-44.

Dolphin, R.E,, and R. N, Peterson.   1960,  Developments in the research
     and control program of the Clear Lake gnat  Chaoborus astictopus3
     DES. Proc. Papers Calif.  Mosq. Contr. Assoc.  28:90-94.

Grodhause, G,  1963..  Chironomid midges as a nuisance,  II,  The nature
     of the nuisance and remarks on its control,  Calif, Vector Views

Hazeltine, W, E. 1962,  The development of a new concept for control
     of the Clear Lake gnat,  J. Econ, Entomol..  56:621-26,

Lindquist, A.W,, A. R, Roth, and J, R. Walker.  1951.  Control of Clear
     Lake gnat in California,  J. Econ, Entomol.  572-77,

Mulla, M. S. 1974.  Chironomids in residential-recreational lakes,
     An emerging nuisance problem-measures for control.  Entomol. Tidskr.
     95 (Suppl.) :172-76.

Mulla, M. S., G. Majori, and H.A. Darwazeh.  1975b.  Effects of the
     insect growth regulator Dimilin or TH-6040 on mosquitoes and some
     nontarget organisms.  Mosq. News  35:211-16.

Mulla, M. S., D. R. Barnard, and R. L. Norland.  1975a.  Chironomid
     midges and their control in Spring Valley Lake, California.
     Mosq. News  35:389-95.

Mulla, M. S., and A. M. Khasawinah.  1969.  Laboratory and field evalua-
     tion of larvicides against chironomid midges.  J. Econ. Entomol.

Mulla, M. S., R. L. Norland, D. M. Fanara, H. A. Darwazeh, and D. W.
     McKean.  1971.  Control of chironomid midges in recreational lakes.
     J. Econ. Entomol.  64: 300-07.

Mulla, M. S., R. L. Norland, T. Ikeshoji, and W. Kramer.  1974.  Insect
     growth regulators for the control of aquatic midges,  J. Econ.
     Entomol.  67:165-70.

Mulla, M. S., R. L. Norland, W. E. Westlake, B. Dell, and J. St. Amant.
     1973.  Aquatic midge larvicides, their efficacy and residues in
     water, soil, and fish in a warm-water lake.  Environ. Entomol.

Norland, R. L., and M. S. Mulla.  1975.  Impact of Altosid on selected
     numbers of an aquatic ecosystem.  Environ. Entomol.  4:145-52.

Patterson, R. S., D. L. Von Windeguth, B, M. Clancy, and F. L. Wilson.
     1966.  Control of the midge Glyptotendipes paripes with low-volume
     aerial sprays of malathion.  J. Econ.. Entomol.  59:863-66,

Polls, I., B. Greenburg, and C, Lue-Hing,  1976,  Control of nuisance
     midges in a channel receiving treated municipal sewage,  Mosq.
     News  35:533-37.

Snell, J.  B,, and W, Hazeltine,  1962,  The use of insecticide  to  determine
     the, life history of an aquatic gnat Chaoborus ' ast-iotopus ^.  Ann.
     Entomol. Soo.. Amev.  56;816-18,

       CERATOPOGONIDAE   (Biting Midges, Punkies, Sand Flies)
     Although biting midges transmit a number of parasitic diseases
 (Kettle 1965), in the United States they are of particular importance
as bloodsucking pests of man.  Some species, adapted to brackish water,
are vicious pests near salt marshes and beaches; others are severe
pests in northern forests.  The immature stages of biting midges breed
primarily in moist to wet soil of salt marshes, beaches, swamps, margins
of ponds, small puddles, hoofprints, treeholes, decaying leaves, etc.
 (Hair et al. 1966).  These habitats are often shaded and protected by
living and decaying plants on the ground surface.

     Although it has not been rigorously demonstrated, the larvae are
probably more susceptible to insecticides than the eggs or the pupae.
Because the larvae live in a protected habitat, control usually is im-
practical unless persistent insecticides are used.  There has been a
general tendency to rely on environmental manipulation, e.g., filling,
flooding or draining larval breeding areas  (Linley and Davies 1971,
Foulk 1966).
Laboratory Evaluation

    1 Biting midges, with due allowance for their size, are handled much
like mosquitoes for laboratory larvicide screening tests.  Colonies of
a number of species have been established to facilitate laboratory testing
(Linley 1968).

     One ml of acetone solution of each of a number of insecticides in
serial dilutions is pipetted into 190 ml of distilled water in 250 ml
flasks with 1 ml of acetone serving as a check.  After 30 minutes, 10 ml
of distilled water containing 25 fourth instar larvae is added to each
beaker.  The flasks are held at about 25-30°C for 24 hours after which
time mortality readings are made.  A minimum of three tests should be
reported at each of three to four concentrations with mortalities between
10% and 90%.  Slopes, LCrQ's and LCgg's are then calculated as given
under Data Analysis.  Larvae are considered dead if they are incapable
of swimming normally or do not respond when probed with a needle.  Larvae
that pupate during the test are discarded.  If more than 10% of the
control larvae pupate, or the control mortality is more than 20%, the
test should be discarded (Harris and Jones 1962),
Field Evaluation

     Small plots experimentally treated with larvicides can vary in

size from 1 or 2 to 40 square m or more.  The larger sizes reduce the
likelihood of larval migration into treaced plots from adjacent untreated
areas effecting the results.  Small plots can be treated with hand
sprayers (or spreaders for granules) while larger plots usually require
power sprayers, and still larger plots aircraft.

     Effectiveness is much more difficult to evaluate for biting midge
larvae than for mosquito or black fly larvae since they live in soil
or decaying organic matter and are small and slender.  Species identi-
fication is necessary to determine which areas are to be treated bacause
many species that are nonbiting live in habitats superfically similar to
those inhabited by biting species.  Populations of larvae can be sampled
by collecting a standard area and volume of suspected breeding media.
These are taken to the laboratory where they are washed individually with
tap water through a 10-mesh sieve and a 20-mesh sieve into an 80-mesh
sieve.  Biting midge larvae are washed through the first two sieves but
retained by the 80-mesh sieve.  The material retained by the 80-mesh
sieve is placed in a low broadmouthed glass container which is filled
with a saturated solution of magnesium sulfate (Epsom salt) and stirred
(Kettle et al. 1956).  The container is then placed under a low-power
dissecting microscope and the surface of the solution examined'.  Because
of the high specific gravity of the solution, biting midge larvae,
pupae and other small organisms float to the surface along with plant
detritus.  The larvae or pupae can then be transferred to clean water
and counted or reared.  Variations of these methods are described by
Bidlingmayer (1957), Jamnback and Wall (1958), and Foulk (1966).

     A second method of evaluating the efficiency of larviciding is
emergence trapping (Davies 1966).  Traps can be made of inexpensive
cones of roofing felt joined with paper fasteners.  The basal opening
should be about 0.2 square m and the opening at the top about 10 cm.
A glass jar with the inside walls coated with castor oil is placed over
the top.  The emerging insects are attracted to light, enter the jar and
are trapped on the sides.  These are taken to the laboratory at regular
intervals and the biting midges removed, identified and counted.  The
traps should be removed after each count as the larvae tend to migrate
into the shaded areas beneath the trap from adjacent areas.
Laboratory Evaluation

     Biting midges are attracted to light and frequently enter houses.
They are small enough to pass through window screens and so can be a
serious nuisance at night.  Painting or spraying window screens with a
pesticide can prevent annoyance by killing the midges that alight on
the screen mesh in the process of entering a room.

     The toxicity of insecticides to adult biting midges can be measured
in the laboratory using standard WHO susceptibility kits in which the
standard screen through which the midges can pass is replaced with a fine
mesh screen or nylon stockings  (Service 1968).

     Formulation can also be tested in small chambers with one end of the
chamber covered by a transparent plastic and the other end opaque.  A
screen is inserted between the  light and dark ends of the container.
The adult midges are introduced into the container at the dark end.  They
are attracted to light and quickly alight on and pass through the treated
screen at which time they are exposed to insecticide.  The duration of
the effectiveness of such treatment can be measured by preparing a large
number of screens in the same way and exposing them to weathering.
Evaluation of effectivess is measured by determining the time from passing
through the screen (immediately after they are introduced) until mortality
(Jamnback 1961, Linley and Davies 1971).
Field Evaluation
     Adult populations estimates can be made using either landing rate
counts or light trap collections.  Whichever method is used, counts
should be taken simultaneously both inside and outside of the treated
area before and at regular intervals after treatment.  In this way, normal
hourly and seasonal changes in activity and populations levels can be
taken into account when evaluting the effectiveness of control.

     Landing rate counts preferably taken by two individuals at each site
at the same time should not begin until five minutes after arrival at
a given site as counts taken immediately on arrival are lower and less
uniform.  When biting midges are abundant, counts of the number landing
on one bared arm exposed to ca. 10 cm above the elbow over a two minute
period will provide a useful estimate (Jamnback and Watthews 1963).

     Adult biting midges are attracted to light and can be collected in
light traps in large numbers in the same way as mosquitoes to evaluate
the effectiveness of larval or adult control programs.  However, all
species are not equally attracted to light and biased counts may result.
Another drawback is that many species of insects other then nuisance
Culioo'tdes .are attracted to light, and extensive sorting is required
even when only aliquots of the collection are identified.  If light traps
are fitted with jars of alcohol rather than ethyl acetate, sorting,
handling and identification are simpler.  The larger moths, beetles,, flies,
etc. can  be  strained  out with 10- and 20-mesh sieves.  The larger insects
are retained by these sieves and the Culicoi-des by a 60-mesh sieve.

     Tests on insecticide aerosols have not been reported under field
conditions although they are routinely used in many areas.  The efficacy
can be evaluated by taking landing rate counts inside and outside of the
control area before and at intervals after treatment as described above.

Bidlingmayer, W.  1957-  Studies on CuUcoi-des furens  (Poey)  at  Vero
     Beach.  Mosq. News  17:292-24.

Davies, J.  1966.  An evaluation of the emergence or box  trap for
     estimating sand fly (Culicaides spp,:  Heleidae)  populations.
     Mosq. News  26(1):69-72.

Foulk, J.  1966.  Drainage of a desert spring creek for control  of
     Leptoconops kerteszi- (Diptera:  Ceratopogonidae).  Mosq.  News

Hair, J., E. Turner, and D. Messersmith.  1966.  Larval habitat  of
     some Virginia Culico-ides (Diptera:  Ceratopogonidae).  Mosq.
     News 26(2):195-204.

Harris, R., and R. Jones.  1962.  Larvicide tested with colony-reared
     Culi-coides vavi'ipenn-is.  J. Econ. Entomol.  55(4) :575-76.

Jamnback, H.  1961.  The effectiveness of chemically treated  screens
     in killing annoying punkies.  Culicoides obsoletus.  J.  Econ.
     Entomol.  54(1):578-80.

Jamnback, H., and W. Wall.  1958.  A sampling procedure for Cul'ico'ides
     mell&us (Coq.) (Diptera:  Heleidae) with observations on  the life
     histories of two coastal species.  Mosq. News 18(2):85-88.

Jamnback, H., and W. Watthews.  1963.  Studies of populations of adult
     and immature Cul-Loo'ides sanguisugs (Diptera:  Ceratopogonidae).
     Ann. Entomol. Soc. Amer.  56(6):728-32.

Kettle, D.  1965.  Biting ceratopogonids as vectors of human  and animal
     diseases.  Aota Trap.  22(4):356-62.

Kettle, D., R. Nash, and B. Hopkins.  1956.  Field tests with larvicides
     against CuUcoides impunotatus Goetghebuer in Scotland.   Bull.
     Entomol. Res.  47(3):553-73.

Linley, J.  1968.  Colonization of Culico-Ldes furens.  Ann. Entomol.
     Soc. Amer>.  61(6) :1486-90.

Linley, J., and J. Davies.  1971.  Sandflies and tourism  in Florida and
     the Bahamas and Caribbean area.  J. Econ. Entomol.   64(1):264-78.

Service, M.  1968.  The susceptibility of adults of Culico-ides impunctatus
     Goetghebuer and C. obsoletus (Meigen) to DDT and  Dieldrin.  Mosq.
     News  28(4):543-47.