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University Microfilms International 300 North Zeeb Road Ann Arbor, Michigan 48106 USA St. John’s Road, Tyler's Green High Wycombe, Bucks, England HP10 8HR 7619507 ESTENIK, JOHN FRANCIS TOXICITY STUDIES OF INSECTICIDES WITH LABORATORY COLONIES OF MIDGE LARVAE, CHIRONOMUS RIPARIUS (= CHIRONOMUS THUMMI), AND MOSOUITO LARVAE, AEDES AEGYPTI, AND THE IN VITRO CHARACTERIZATION OF ALDRIN EPOXIDATION IN THE MIDGE. THE OHIO STATE UNIVERSITY, P H #D,, 1978 TOXICITY STUDIES OF INSECTICIDES WITH LABORATORY

COLONIES OF MIDGE LARVAE, CHIRONOMUS RIPARIUS

(= CHIRONOMUS THUMMI), AND MOSQUITO LARVAE,

AEDES AEGYPTI, AND THE IN VITRO CHARACTERIZATION OF

ALDRIN EPOXIDATION IN THE MIDGE

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

By

John F. Estenik, B. S., M. S

The Ohio State University

1978

Reading Committee:

L. S. Putnam

N. Wilson Britt '/ Co-Advisors” William J. Collins Department of Zoology ACKNOWLEDGEMENTS

I would like to thank my co-advisors, Drs. N. W.

Britt and William J. Collins. Dr. Britt, whose help allowed me the freedom to define my graduate program, was appreciated. Dr. Collin's suggestions, interest, and help throughout all aspects of this research project and manuscript preparation is deeply appreciated.

I would like to thank Dr. L. S. Putnam for helping me return to graduate school and making this all pos sible.

I would like to thank Drs. W. B. Parrish and S. I.

Lustick, Department of Zoology, Dr. IV. Foster,

Department of Entomology, and Drs. G. S. Serif, D. H.

Ives, and Means, Department of Biochemistry, for the use of laboratory space and instrumentation during various aspects of this research project. I would also like to thank members of the Zoology and Entomology Departments who helped and encouraged me throughout my studies.

Finally, I would like to thank my parents and my grandmother, Mrs. F. Rose, for their help, encouragement, and financial aid throughout piy graduate studies. VITA

November 11, 1940 Born - Canton, Ohio

1962 B. S., John Carroll University, Cleveland, Ohio

1964 M. S., John Carroll University, Cleveland, Ohio

1964-1967 Research Assistant, Case Western Reserve University, Department of Radiation Biology, Cleveland, Ohio

1967-1970 Instructor, Cuyahoga Community College, Department of Biology, Cleveland, Ohio

1971-1975 Teaching Associate, Ohio State University, Department of Zoology

1975-1976 Teaching Associate, Ohio State University, School of Nursing

PUBLICATIONS

Estenik, J._F. and P. F. Federle, "The Susceptibility of an Adult Damselfly Ischnura verticalis to Selected Insecticides". The Ohio Jour, of Science, March, 19 75.

FIELDS OF STUDY

Major Field: Aquatic Biology - Toxicology TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS...... ii

VITA ...... iii

LIST OF TABLES...... vii

LIST OF F I G U R E S ...... ix

I. INTRODUCTION AND LITERATURE REVIEW ...... I

A. The Fate of Pesticides in an Aquatic Organism ...... I

B. The Biological Oxidation of Insecticides - 5

C. Chironomus riparius Meigen as a Test O r g a n i s m ...... 10

D. Objectives ...... 14

II. METHODS AND MATERIALS...... 16

A. Insect Culture ...... 16

B. Chemicals...... 17

C. Immersion Toxicity: Assay Procedure, Criteria of Response and Data Analysis . . 18

D. Midge and Mosquito Preparation for All Experiments Except Immersion Toxicity A s s a y s ...... 21

E. Homogenizing Procedure ...... 21

F. Extraction Procedure ...... 22

i v TABLE OF CONTENTS (CONTINUED)

Page

G. Chromatographic Analysis...... 23

H. Absorptive Uptake and Insecticide Loss. . 24

I. Assay of Whole Body Homogenate for Aldrin Epoxidation ...... 26

J. Microsome Preparation ...... 29

K. Assay of Subcellular Fractions for Aldrin Epoxidation ...... 30

L. Protein Analysis...... 31

III. R E S U L T S ...... 32

A. Toxicity Assays ...... 32 B. Absorptive Uptake and Insecticide Loss. . 41

C. Assay of Whole Body Homogenate for Aldrin Epoxidation...... 71

D. Assay of Subcellular Fractions for Aldrin Epoxidation...... 90

IV. DISCUSSION AND CONCLUSIONS...... 93

A. Toxicity Assays ...... 95

B. Absorptive Uptake and Insecticide Loss. . 107

C. Assay of lVhole Body Homogenates for Aldrin Epoxidation...... 115

D. Assay of Subcellular Fractions for Aldrin Epoxidation...... 12 7

E. Conclusions...... 134

v TABLE OF CONTENTS (CONTINUED)

Page

LITERATURE CITED...... 138

vi LIST OF TABLES

Table Page

1 Toxicity of Selected Insecticides With and Without Synergist to Fourth Instar Larval Chironomus riparius...... 33

2 Toxicity of Selected Insecticides With and Without Synergist to Late Third and Fourth Instar Larval Aedes aegypti...... 3?

3 The Absorption of Selected Insecticides by C. riparius Larvae and P u p a e ...... 43

4 The Absorption of Selected Insecticides by A. aegypti Larvae and Pupae...... 45

5 Depuration of Dieldrin by C. riparius Larvae . 72

6 The Uptake and Metabolism of Aldrin by Ligatured riparius Larvae 7 3

7 Aldrin Epoxidase Requirements of C. riparius Whole Body Homogenates...... 74

8 The Effect of Substrate Concentration (Aldrin) and BSA Upon in vitro Aldrin Epoxidase of C. riparius Whole Body Homogenates ...... g7

9 Aldrin Epoxidase Requirements of C. riparius Whole Body Homogenates Under Optimum Conditions...... 89

10 Effect of FMN, FAD, and PBO Upon in^ vitro C. riparius Aldrin Epoxidase Whole Body Homogenates...... 91

11 Subcellular Localization of Aldrin Epoxidase Activity in C. riparius Larvae ...... 92

via Li ST OF TABLES (CONTINUED)

Table Page

12 Reported 24 hr LCs q 's (ppb) of Susceptible Laboratory Strains of A. aegypti to Representative Insecticides...... 98

13 Comparative Rates of Aldrin Epoxidation by Insect Microsomes (pmoles of Dieldrin/min/ mg protein)...... 131 LIST OF FIGURES

Figure Page

1 The Fate of a Xenobiotic in an Aquatic Organism...... 3

2 The Metabolism of a Xenobiotic by an Aquatic Organism...... 4

3 * The Effect of Exposure Time on in vivo Insecticide Absorption and Metabolism by C_. riparius Larvae Immersed in 10 ppb Aldrin . . . 48

4 The Effect of Exposure Time on ill vivo Insecticide Absorption and Metabolism by A. aegypti Larvae Immersed in 10 ppb Aldrin . . . 51

5 The Effect of Exposure Time on the in vivo Hourly Rate of Insecticide Absorption and Metabolism/£. riparius Larva Immersed in 10 ppb Aldrin ...... 53

6 The Effect of Exposure Time on the in vivo Hourly Rate of Insecticide Absorption and Metabolism/A. aegypti Larva Immersed in 10 ppb Aldrin ...... 55

7 Dieldrin Production as a Function of Total Aldrin Absorbed for Larvae Immersed in 10 ppb A l d r i n ...... 58

8 The Effect of Aldrin Concentration on in vivo C. riparius Larval Absorption and Metabolism for a 1 Hour Immersion...... 60

9 The Effect of Aldrin Concentration on ill vivo A. aegypti Larval Absorption and Metabolism Tor a 1 Hour Immersion...... 62

10 The Effect of Aldrin Concentration on Percent of Aldrin Epoxidation for a 1 Hour Immersion. . 64

ix LIST 01; FIGURES (CONTINUED)

Figure Page

11 Dieldrin Production as a Function of Total Aldrin Absorbed for a 1 Hour Immersion. . . . 67

12 The Effect of Size on in vivo Insecticide Absorption and Metabolism by riparius Larvae Immersed in 20 ppb Aldrin for 2 Hours. 69

13 The Effect of pH on vitro Aldrin Epoxidation by Midges ...... 76

14 The Effect of Temperature on in vitro Aldrin Epoxidation by Midges ...... 78

15 The Effect of Buffer Concentration (Molarity) on in vitro Aldrin Epoxidation by Midges. . . SO

16 The in_ vitro Rate of Aldrin Epoxidation in Namomoles/Larva/Minute...... 83

17 The Effect of Larval Numbers (Enzyme Con­ centration) on in vitro Aldrin Epoxidation. , 85

x I. INTRODUCTION AND LITERATURE REVIEW

There is a growing concern regarding synthetic chemicals in aquatic systems which are potentially hazardous to living organisms, and of their possible role in the general deterioration of the environment.

Not only is there concern with direct primary effects upon the growth, survival, and reproduction of exposed organisms, but there is concern regarding secondary effects, or ecosystem changes which follow as a result of primary effects. A listing of xenobiotics, i.e. synthetic organic chemicals that have no nutritive or functional value in an organism's normal biochemical economy (Mason ejt al. 1965) , includes pesticides used in the chemical control of insect pests. Because of their widespread use, insecticide contamination_of freshwater systems and organisms has become an increasing problem.

A. The Fate of Pesticides in an Aquatic Organism

In general, pesticides have a very low' solubility in H2O (Gunther 1968). Aquatic organisms absorb pesticides through direct contact, or indirectly by ingestion of contaminated material. Once the pesticide 2 has been absorbed by the organism, various physiological processes occur depending upon the nature of the pesticide and the organism affected.

Absorbed compounds are translocated in an organism to a site of action (target), to a storage area, to a site of metabolic transformation, or may be immediately conjugated and/or excreted (Figure 1).

Metabolic transformations are generally classified according to the nature of the chemical reaction.

Various classifications for the metabolic transformations of xenobiotics have been proposed (Smith 1962, Shuster

1964, Gillete 1966, Smith 1968, Menzie 1969, Matsumura

1975). The following classification of reactions proposed by Parke (1968) can be used to describe the metabolic transformations of insecticides in living organisms. This scheme classifies reactions as micro­ somal and non-microsomal oxidations, reductions, and hydrolytic reactions, or miscellaneous reactions

(Figure 2) . These metabolic transformations may be activations, reactions which produce a metabolite(s) more toxic than the parent compound, or deactivations

(degradations), metabolites generally less toxic than the parent compound. Activated metabolites may undergo further metabolism (deactivations and/or conjugations), or be excreted (Figure 1). Parent compounds which undergo direct deactivation may be conjugated and/or 3

Xenobiotic in the Environment

Organism Integument

\/ AbsorptionI Target 1 Translocation • Unmodified Parent Molecule System Conjugation Organ and/or Tissue Excretion Molecule

Storage

* Metabolic Transformation

Activated Detoxification Forin Degraded Form

Excretion ^ Conjugation

Figure 1. The Interaction of an Organism and a Xenobiotic Metabolic Transformation ^ MISCELLANEOUS

OXIDATIVENON-OXIDATIVE

M F 0 < 1 ^ NON-MFO MFO — ^ NON-MFO

1. Hydroxylation 1, Deanimation Reduction 1. Reduction A. Acyclic 2. Oxidation A. Nitro A. Aldehydes B. Aromatic A. Alcohols B. Azo B. Ketones C. Alicyclic B. Aldehydes 2. Epoxidation 3. Aromatization 3. N-Hydroxylation A. Alicyclic (Amines) 4, Hydrolysis 4. N-Oxidation A. Esters (31 Amines) 5. S-Oxidation 6. Dealkylation 7. Deanimation 8. Desulphuration 9. Hydrolysis A. Esters B. Amides Activated Detoxification Form \ Degraded Form

Target Conjugation Excretion

Figure 2. Metabolism of a Xenobiotic (Parke 1968). 5 excreted (Figure 1). Any parent molecule or metabolite may be incorporated into an organism's tissues,

including the site of toxic action, at any time during

the metabolic process. The total dynamics of penetration,

activation, deactivation, and excretion determine an

organism’s tolerance to a toxic compound.

B. The Biological Oxidation of Insecticides

Generally, one of three different enzyme systems, microsomal oxidases, hydrolases, or glutathione S-trans-

ferases, is involved in an initial metabolic transforma­ tion of an insecticide by an organism (Matsumura 1975).

Mixed-function oxidase (MFO) enzymes are associated with microsomes, the fragmented form of the "endoplasmic reticulum" obtained by differential centrifugation for

in vitro experimentation (Siekevitz 1963, Siekevitz

1965, Mason et_ a^. “1965, Terriere 1968, Claude 1969).

MFO enzymes require NADPH, oxygen, and are inhibited by methylenedioxyphenyl compounds, or carbon monoxide

(Parke 1968, Casida 1969, Wilkinson and Brattsten 1973,

Kuhr 1974).

Biological oxidations catalysed by microsomes include acyclic, aromatic, and alicyclic hydroxylations, epoxidations, N-hydroxylations of amines, N-oxidations of tertiary amines, S-oxidations, dealkylations, deamina­ tions, desulphurations, and the hydrolysis of esters and 6 amides (Figure 2 ) . The types of reactions catalysed by

MFO enzymes found in organisms and/or tissues are studied by testing for oxidative activity with certain substrates. One common reaction used in screening for

MFO activity and for in-depth studies is epoxidation, and the substrate generally used is the insecticide aldrin. For a more extensive treatment of insecticide metabolism, the reader is referred to review articles

(Brooks 1969, Fukunaga et_ al. 1969, Bull 1972,

Schlagbauer and Schlagbauer 1972, Fukuto 1973, Menzer

1973, Menn and McBain 1974).

The activity of insect microsomal oxidase varies depending upon tissue source, preparatory procedure, and incubation conditions. The oxidative activity in some insect organs and tissues is greater than in others

(Krieger and Wilkinson 1969, Khan 1969, Benke and

Wilkinson 1971, Krieger and Lee 1973, Gilbert and

Wilkinson 1974, Turnquist and Brindley 1975). In general, gut, gastric caeca, Malpighian tubule7 and fat body microsome homogenates have a higher MFO activity than other tissue homogenates (Wilkinson and Brattsten

1973) . Because of the difficulty in isolating specific insect organs, homogenates prepared from whole insects or insect parts, i.e. headless insects, abdomens, 7 thoraxes, etc., have been used in many studies.

Physiological factors that affect MFO activity in­ clude stage of development (age), sex, strain, and diet.

Some immature insects have a low MFO activity during early development, but undergo a rapid increase in oxidative activity immediately prior to pupation, or development into the adult form (Hook et_ al^ 1968,

Krieger and Wilkinson 1969, Benke and Wilkinson 1971,

Yu and Terriere 1971, Gilbert and Wilkinson 1974,

Terriere and Yu 1976). Other insects have shown cyclic increases in MFO activity, with a loss of oxidative capacity during molting (Benke et_ aJL. 1972) . Little MFO activity is found during the pupal period of holometa- bolous insects (Krieger and Wilkinson 1971, Gilbert and

Wilkinson 1974, Yu and Terriere 1974a, Terriere and Yu

1976). MFO activity also changes during the adult stage of an insect's life cycle, usually increasing to a maximum early in adult development and then decreasing as the insect ages (Khan 1970, Benke and Wilkinson

1971, Yu and Terriere 1971, Hanson and Hodgson 1971,

Benke et al. 1972, Turnquist and Brindley 1975, Terriere and Yu 1976).

MFO activity is generally higher in female insects, when increases in enzyme action due to sexual differences 8

exist (Benke and Wilkinson 1971, Gilbert and Wilkinson

1974). However, MFO increases due to sexual differences become less pronounced when enzyme activity is expressed

in protein units, rather than as activity per insect

(Wilkinson and Brattsten 1973) .

MFO activity varies between laboratory strains of

insects having different degrees of insecticide

resistance (Terriere 1968, Khan 1970, Schonbrod ert al.

1973, Yu and Terriere 1974b). Increases in microsomal

activity can be one factor in the development of

resistance in an insect strain (Wilkinson and Brattsten

1973) .

Diet can influence the susceptibility of certain insects to insecticides, however, it has little or no effect upon gut tissue MFO activity in other insects

(Wilkinson and Brattsten 1973). The incorporation of certain drugs or insecticides into an organism's diet can substantially increase MFO activity (Hodgson and

Plapp 1970, Nasim and Brindley 1971, Brattsten and

Wilkinson 1973, Yu and Terriere 1974a, Yu and Terriere

1974b, Turnquist and Brindley 197 5, Terriere and Yu

1976). This temporary biochemical adaptation, or inductive effect, increases an organism's ability to survive potentially hazardous chemicals in its environment. 9

Physical and chemical factors which can affect in vitro MFO activity include homogenizing procedure and medium (Schonbrod and Terriere 1966, Hodgson and Plapp

1970, Wilkinson and Brattsten 1973), and the type, strength, and pH of the buffer used during incubation.

Generally, phosphate, Tris-HCl, KC1, sucrose, and sucrose-KCl combinations have been used as incubation media (Wilkinson and Brattsten 1973). The range for pH optima of MFO reactions is from pH 6.9 to pH 9.0; however, most MFO enzyme systems have pH optima of from pH 7.0 to pH 8.5 (Wilkinson and Brattsten 1973). The range for temperature optima of MFO activity is from

20°C to 33°C, with relatively high oxidative activity for certain insect species found between 25°C to 30°C

(Wilkinson and Brattsten 1973).

One of the most important characteristics of all animal mixed-function oxidase systems is their sensi­ tivity to methylenedioxypheny1 synergists (Matsumura

1975). Synergists enhance pesticide toxicity in insects by inhibiting detoxication reactions. On the other hand, these same synergists antagonize insecti­ cides which are activated by MFO enzymes. Such bio­ activations occur with certain chlorinated cyclodienes, some organophosphates, and some carbamates (Dahm and 10

Nakatsugawa 1968) .

The synergistic ratio (SR), i.e. the LD50 of the

insecticide alone divided by the LD50 of the insecticide-

synergist combination, provides a measure of detoxication ease of an insecticide (Metcalf 1967, Brattsten and

Metcalf 1970). Other terms used in place of SR include degree of synergism, synergistic activity, cotoxicity

coefficient, and synergistic effect (Metcalf 1967). A

SR less than 1.0 indicates inhibition of a toxification reaction and provides a measure of bioactivation ease of an insecticide.

C. Chironomus riparius Meigen (Diptera:Chironomidae)

as a Test Organism

Chironomus riparius was selected for the test organism in this research project for several reasons.

C. riparius is a common species having a wide distribu- ~ tion. Specimens have been collected throughout Europe,

Great Britain, and the United States and Canada, from

Alaska to Newfoundland, south to California and South

Carolina, and in Florida (Stone ert aJL. 1965, Credland

1973). In a recent review of the taxonomic status of

Chironomus riparius Meigen and Chironomus thummi Kieffer it has been concluded that iC. riparius and C. thummi are synonyms, the correct designation for the species 11 being the former, older name (Credland 1973). Taxonomic status is important because larvae with the names C. riparius and C. thummi have been used extensively in physiological, biochemical, and genetical research. This study is, to my knowledge, the first comprehensive report of midge-insecticide dynamics in this species.

The adult stage of various chironomid species

(midges) are considered pests in several areas of the

United States. Problems with mass emergences have occurred in California, Oregon, Massachusetts, Wisconsin,

Florida, and New Jersey (Gerry 1951, Grodhaus 1963a,

Hansens and Hagmann 1964). Immense numbers of midges swarming during periods of peak emergence are considered a nuisance. For a detailed discussion of midges as pests, the reader is referred to the articles by

Grodhaus (1963a, 1963b). Problems with midges are in­ creasing due to increases in urbanization and the use of recreational areas. However, very little information exists about midge-insecticide interactions and effects.

Most midge control studies using insecticides have been field studies where the effectiveness of various insecticides, or insecticide formulations, were eval­ uated. Exceptions to that are three papers dealing with laboratory studies on the acute toxicity of unfor­ mulated insecticides to midge larvae (Mulla and Khasawinah 1969, Karnak and Collins 1974, Ali and Mulla

1976) . Midges Tespond to insecticides in the part per billion (ppb) range. Such laboratory investigations of acute toxicity, plus additional experimentation on penetration and metabolism, may help in understanding the generally low tolerance of midges to insecticides.

They may also help determine the most effective insecticides and control measures against midges.

Midge larvae are usually found in great numbers in highly "polluted” waters, i.e. waters having very little dissolved oxygen. One reason for the survival of midge larvae in water having little dissolved oxygen is the presence of hemoglobin. The oxygen affinity of the hemoglobin found in riparius larvae is high. The oxygen pressure for 50% oxygemoglobin at 17°C was 0.6 mm

Hg for C.. riparius," compared to 2 7 mm Hg for human whole blood at 20°C (Fox 1945). Chironomids are able to live for extended periods under anoxic conditions, the length of survival time varying with species (Sturgess and Goulding 1968). The estimated LD50 for C. riparius larvae tested under anoxic conditions was 1.8-0.1 days

(Sturgess and Goulding 1968). Additional details con­ cerning oxygen consumption in relation to body size and temperature in C. riparius larvae are available in a 1 3

report by Edwards (1958). Since midge larvae tolerate

low oxygen conditions and are usually found in waters

having little dissolved oxygen and a high biological

oxygen demand (BOD), the importance of an enzyme system

that requires oxygen in order to detoxify insecticides

has special significance.

Chironomids are important as fish food organisms,

and may comprise a major portion of the diet for

certain fish species. Therefore, midge larvae may be

considered beneficial non-target organisms in pest

control programs. The examination of midge larvae-

insecticide dynamics in the laboratory will be helpful

in determining what effect these agents have upon

midges which are considered non-target organisms.

Chironomid larvae are usually found in large

numbers in aquatic environments. Population densities of

thousands and tens of thousands of larvae per square

meter are common. There have been reports of larvae

inhabiting and breeding in lagoons with population den­

sities in excess of 150,000/m^. Bloodworms (Tendipes

sp. = midge larvae) accumulated 1.4, 1.6, and 3.1 times

more insecticide in one week than Zygoptera, Notonectidae,

and Anisoptera respectively, and were 1.4 times more

important than backswimmers as a long term, potential pesticide source in a study using radioactive DDT in a 14

freshwater marsh (Meeks 1967, Eberhard et_ aJL. 1971).

More information about midge-insecticide interactions

may be helpful in determining potential problems and

secondary effects resulting from the insecticide con­

tamination of water systems.

Finally, of the estimated 2.5 to 10 million insect

species, only about 20 have been used for in-depth, biochemical studies of xenobiotic metabolism (Wilkinson

and Brattsten 1973). The majority of experiments have been qualitative in vivo studies comparing different species. There have been few in-depth, in vitro studies, and little correlation of in vivo and in vitro data. With the exception of a few in vivo insecticide uptake studies by mosquitoes, the midge larva C. tentans (Derr and Zabik 1972, Derr and Zabik

1974), the dragonfly nymphs T. cynosura and T. semiaquea

(Wilkes and Weiss 1971) , and a few' in_ vitro metabolic studies using mosquito and caddisfly larvae (Krieger and

Lee 1973), very little is known about aquatic species.

D. Objectives

The objectives of this study are:

1) to determine and compare the susceptibility of

Chironomus riparius larvae and Aedes aegypti larvae 3 5

(Dipter_a:Culicidae) to selected insecticides and

certain oxidative metabolites ;

2) to compare the rate of aldrin accumulation and meta­

bolism in C. riparius larvae and A. aegypti larvae as

a function of insecticide concentration and exposure

t ime;

3) to determine whether C. riparius larvae have an active

mixed-function oxidase system;

4) to determine the optimal assay conditions for aldrin

epoxidation using C. riparius larva whole body

homogenates;

5) to determine the specific activity of aldrin epoxida­

tion using C. riparius larval microsomes.

Such information will be helpful in explaining the high sensitivity of midge larvae to insecticides, and may be helpful in predicting the effectiveness of closely related chemical isnecticides for the control or protection of midges. II. METHODS AND MATERIALS

A. Insect Culture

A colony of Aedes aegypti CL.) was established from a susceptible strain maintained at the Department of

Entomology, The Ohio State University. The O.S.U. colony was obtained in 1960 from the U.S. Army Labora­ tories in Fort Detrick, Maryland. The latter colony was sub-cultured from the Rockefeller Strain (W. A. Foster, pers. comm., June, 1976).

Adult mosquitoes were kept in a 25 X 19 X 15 cm plastic cage fitted with a screen for ventilation. The colony was maintained at 27 t 2°C, a relative humidity of 80 - 5%, on a 16 hr light/8 hr dark photoperiod \vith incadescent lighting, and was provided with a vial of

10% glucose solution supplied on a cotton wick.

Adult females were blood fed on rats when eggs were required. An oviposition cup containing aged tap water and lined with brown paper towel strips was placed in the colony cage 3 days following blood feeding. The moist towel strips containing eggs were transferred to a small container, sealed with a lid, and placed in an incubator at 28 ± l°c for at least 4 days to allow for 17 embryonation. Newly hatched larvae were obtained by submerging the eggs in water and were immediately transferred, 200-300 larvae/450 ml of aged tap water, to 30 X 20 X 5 cm metal pans fitted with plastic lids.

The larvae were maintained at 28 * 1°C on a 16 hr light/8 hr dark photoperiod with incandescent lighting.

Larval food consisting of a 1:1:1 mixture of lactal- bumin, yeast, and Purina Lab Chow was added daily in the following amounts: day 1-25 mg, day 2 - 50 mg, days 3 and 4 - 100 mg, and day 5 - 17 5 mg.

Midge larvae were collected at the Jackson Pike sewage treatment plant in Columbus Ohio and reared to the adult stage in the laboratory. From them, a colony of Chironomus riparius (Meigen) was established and maintained in aerated tap water (21 - 2°C) in a bin,

48 X 37 X 23 cm, covered with a screen flight cage,

50 X 35 X 75 cm. The larvae were fed pulverized Hartz

Mountain Dog Yummies^ and reared according to a described method (Biever 1965) without the addition of substrate.

The larvae were maintained for 3 years as a laboratory culture prior to being used for experimentation.

B. Chemicals

Technical or analytical grade insecticides, at 18

_ least 95% pure, were prepared as acetone solutions:

p-p' DDT, lindane, parathion, paraoxon, malathion,

malaoxon, O-ethyl O-p-nitrophenyl phenylphosphonothioate

(EPN), propoxur, carbaryl, Landrin^, aminocarb, mexa-

carbate, allethrin (90%), piperonyl butoxide (PBO), and

sesoxane. Aldrin was 98.5% and dieldrin was 99+% pure.

The chemical definition and structure of these compounds

are available in Kenaga and End (1974).

All other chemicals and solvents employed were

analytical reagent grade. All solvents were redistilled

in glass prior to use.

C. Immersion Toxicity: Assay Procedure, Criteria of

Response and Data Analysis

The assay procedure used was a modification of

methods previously described (Mulla and Khasawina 1969_,

Karnak and Collins 1974). The toxicity assays were

conducted in narrow-mouth, quart glass jars containing

conditioned tap water (aged 24 hrs). The pH of aged,

Columbus tap water ranged from pH 7.5 to 8.5 and the

temperature was 21 - 2°C throughout all tests. The

insecticide assays were made without the addition of

food, substrate, or air during the test period.

Cannibalism was no problem with well fed larvae and

short term (24 hr) assays. The incidence of cannibalism 19

increased for test periods longer than 24 hrs.

All insecticide solutions or suspensions were pre­

pared by adding insecticide in 0.5 ml of reagent acetone

to 500 ml of conditioned tap water. The containers were

capped with a rubber stopper covered with aluminum foil,

and shaken thoroughly to insure complete mixing. New

acetone solutions of insecticide were prepared for each

experiment.

Each assay contained untreated controls, solvent

controls, and at least 3 to 7 insecticide concentrations

with predicted responses from 5%-95% mortality, based

upon preliminary assays.

Twenty fourth instar midge larvae, 15-18 days old,

or late third-early fourth instar mosquito larvae, 5

days old, were selected at random and placed in test

containers, 10 organisms/container, 2 containers/

insecticide concentration. Mortality was recorded after

24 hrs, moribund larvae being recorded as dead. Larvae

used in synergism experiments were pretreated for 1 hr

in 1 part per million (ppm) PBO, 1 ppm sesoxane, or

0.5 ppb EPN (sub-lethal doses). Pretreated larvae were

then transferred to jars containing insecticide with

synergist, and mortality was recorded after 24 hrs.

The synergistic ratio was obtained by dividing the lethal 20

concentration for the 501 mortality (LC^g) of the

insecticide alone by the LCgg of the insecticide-

synergist mixture (Metcalf 1967).

Data reported for toxicity assays represent mean

values of 3 replicate experiments performed on different

days. Organisms that pupated during the assay period,

or test concentrations where mortality was 0% or 100% were excluded from analysis. Toxicity assays were

corrected for control mortality (Abbott 1925), and the

data were pooled for analysis. The LC5q values,

regression coefficients (slope) , and associated

statistics at the 95% confidence level were determined using a computer program based on weighted probits

(Finney 1971).

Various criteria have been used to define mortality

in midges including lack of movement when touched with

a probe, inability to make undulating movements, and

color changes (Mulla and Khasawinah 1969, Augenfeld

1967, Sturgess and Goulding 1968, Karnak and Collins

1974). I used motility to determine midge mortality.

Midge larvae are capable of locomotion in a horizontal and vertical plane. The larva raises its anterior end slightly and moves its head from the midline laterally to the caudal region. The larva then moves its head 21

laterally through an arc of approximately 360°, bringing

its head to the caudal region on the opposite side.

The head then moves back to the midline to complete what

I define as one swimming cycle, which in a composite,

resembles a figure eight. Normal larvae continue these

movements, generating continuous figure eights. Any

larva which could not respond with 3 swimming cycles when

pinched with a tweezers in the region of the anal papillae

was considered moribund, and was included in mortality

counts. Mosquito larvae use a similar method to swim;

however, the movement is less pronounced. A 3 cycle

swimming response to tapping the container with a pencil

was also used to determine which mosquitoes were

unaffected.

D. Midge and Mosquito Preparation for All Experiments

Except Immersion Toxicity Assays

Fourth instars, 15-18 day old midge and 6 day old

mosquito larvae, or pupae were selected from large,

laboratory cultures and were weighed to the nearest 0.1 mg with a Mettler Balance.

E. Homogenizing Procedure

Larvae used in whole body homogenate assays and

subcellular fraction assays were homogenized in Tris-HCl 22 buffer unless otherwise noted. Test organisms used in absorptive uptake or insecticide loss experiments were removed from the exposure container, rinsed with water, and homogenized in 5 ml of distilled water. All homo- genates were prepared in a glass Potter-Elvehejm tissue grinder at the lowest speed for approximately 20 sec, 10 passes through the brie.

F. Extraction Procedure

A modification of an extraction procedure reported recently (Nelson £t al_. 1976) was used. Brie acidified with 2 ml of 5% trichloroacetic acid (TCA), was trans­ ferred to a separatory funnel and extracted 3 times with

20 ml of petroleum ether. The extracts were combined, reduced to 5 ml in a rotating evaporator (Buchi

Rotavapor-R), returned to the separatory funnel, and combined with 60 ml each of acetonitrile and distilled water. The acetonitrile-water-insecticide mixture was then extracted twice with 60 ml of petroleum ether.

Anhydrous Na2SO^ was added to the combined 120 ml petroleum ether extract. The extract was evaporated just to dryness and the residue was resuspended in a measured volume of benzene for analysis by gas-liquid chromato­ graphy (GLC). 23

Extraction efficiencies determined from spiked

samples were:

Organism % Insecticide Recovery

Aldrin Dieldrin Lindane

Midge Larvae 72.8 83.3 81.6

Midge Pupae 79.9 81. 5 79.6

Mosquito Larvae 70.5 80.1 -

Mosquito Pupae 76. 3 - -

Water samples were extracted 3 times with 50 ml of petroleum ether, and anhydrous Na2S04 was added to the

combined 150 ml extract. The extract was evaporated just to dryness and resuspended in a measured volume of benzene for GLC analysis. The extraction efficiency for dieldrin in water samples was 95%.

G. Chromatographic Analysis

The amount of aldrin, dieldrin, or lindane was determined by gas~liquid chromatography (GLC). A

Varian Aerograph GLC Model 1440 equipped with a 3HSc electron capture detector and a 150 cm X 2 mm Ci»d.) glass column packed with 3% (W/V) SE-30 on Gas Chrom Q was used. The following operating parameters were employed: injector 240°C, column 200°C, detector 260°C, 24

and a N 2 flow of 37.5 ml/min. The chromatogram was

recorded on a Model A-25 Varian Recorder with a chart

speed of 0.1 in/min. Insecticide concentrations were

determined by peak height using daily standard curves

under identical chromatographic conditions. Each

insecticide was determined from an average of three

inj ections.

H. Absorptive Uptake and Insecticide Loss

Insecticide solutions were prepared using the method

previously described for toxicity assays. Test organisms were selected and weighed prior to each experiment.

Fourth instar midge larvae or pupae, 2 0/container, were placed in 0.5 1 of solution. Fourth instar mosquito

larvae or pupae were assayed at the same organism/water volume ratio as midges, by exposing 1000 organisms in

2.5 1 of solution.

The absorptive uptake of aldrin, dieldrin, and

lindane was tested at 20 ppb for 2 hrs, and was compared

for midge and mosquito larvae and pupae under identical

conditions.

The effect of PBO on the absorptive uptake of aldrin

and dieldrin and the conversion of aldrin to dieldrin in larvae wTas also determined. Midges and mosquitoes 25

were pretreated for 1 hr in 1 ppm. PBO. The organisms

were then transferred to jars containing 20 ppb aldrin

or dieldrin with 1 ppm PBO and exposed for 2 hrs.

Mosquitoes were also tested in 20 ppb aldrin with 1 ppm

PBO without pretreatment and with a 24 hr pretreatment

of 2 ppm PBO.

Midges and mosquitoes were also tested under

"midge conditions", i.e. low identical insecticide

concentrations for short periods of time, in order to

establish the effects of time and aldrin concentration

upon insecticide absorption and conversion.

An experiment was designed to determine the rate of

insecticide loss from midge larvae in an insecticide

free environment. After exposing 60 midges for 1 hr

to 20 ppb dieldrin, 20 midge larvae were removed and

analysed for dieldrin content. The remaining 40 midges

were placed in 0.1 1 of insecticide-free water. After

3 hrs, 20 midges were removed and analysed. The

remaining 20 midges were transferred to a second 0.1 1

of clean water for another 3 hrs. The water of the

holding containers was also analysed for dieldrin.

An experiment was conducted to test for possible

differential insecticide uptake through anterior and posterior halves of midge larvae. A striction, a

tightly tied thread preventing the free movement of 26

materials between anterior and posterior halves, was used. Twenty prepared larvae were exposed for 1 hr

to 40 ppb aldrin. The pooled anterior and posterior

halves of the larvae were analysed for the absorption

and conversion of aldrin.

In all of the above experiments, test organisms were removed from insecticide solutions , rinsed with water one time, and were homogenized in distilled water.

The insecticide was extracted and analysed by GLC

(see the appropriate procedures).

All data for absorptive uptake and insecticide loss

experiments represent the mean value from 2 experiments performed on different days.

I. Assay of Whole Body Homogenate for Aldrin Epoxidation

The following experimental sequence was designed

to determine the optimum in vitro conditions for aldrin

epoxidation in larval whole body homogenates: (1) the

effect of component chemicals generally included in a

"standard" incubation mixture, (2) a pH profile, (3)

a temperature profile, (4) a molarity profile, (5) a reaction time profile, (6) a larval concentration

(protein concentration) profile, (7) a substrate concen­ tration profile, and (8) a restudy of the effects of component chemicals in the "standard" incubation mixture 27 upon aldrin epoxidation under optimum conditions as defined by steps 2-7 above. The effect of PBO, FMN, and FAD upon enzyme activity was also tested.

Midge larvae were weighed and homogenized in the appropriate medium. Distilled water, 8.3 X 10"^ M

Tris-HCl buffer, pH 7.5, or 8.3 X 10"1 M Tris-HCl buffer, pH 7.5, was used depending upon experimental requirements.

The effect upon enzyme activity of each chemical used in a slightly modified "standard" incubation mixture (Krieger and Wilkinson 1969) was tested using

20 midge larvae/3 ml, homogenized in 8.3 X 10”^ M Tris-

HCl buffer, pH 7.5, Each complete 5 ml incubation mixture contained 20 homogenized midges and the following chemicals: 5.0 X 10~2m Tris-HCl buffer, pH 7.5, 2.4

X 10"^M glucose 6-phosphate- (G-6-P), 1.6 units glucose

6-phosphate dehydrogenase (G-6-P dH), 5.1 X 10_^M

NADP, and 2.7 X 10"3m KC1. The foregoing was the standard mixture used by Krieger and Wilkinson (1969) .

The following chemicals were added to Krieger's and

Wilkinson's incubation mixture in the final concentration indicated: 5.1 X 1 0 " NADH, 1% (W/V) bovine serum albumin (BSA), and 1.0 mg aldrin in 0.1 ml ethanol.

All subsequent whole body homogenate experiments included 28

all of the previously listed chemicals and concentrations unless otherwise noted. Reaction mixtures were incubated

in test tubes at 30 * 1°C, and were swirled at 100 RPM

in a Metabolyte Water Bath Shaker CNew Brunswick

Scientific Co., Inc.). The reactions were stopped after

1 hr by the addition of 2 ml of TCA.

Modifications in larval homogenization procedures for specified experiments included the following. Midges used for the pH and molarity profiles were homogenized in distilled water. For the pH profile, the 5 ml volume reaction mixtures contained the desired pH and a final Tris-HCl buffer molarity of 5.0 X 10"2m ; for the molarity profile, the final molarity \^as attained with

Tris-HCl buffer at a pH of 7.5. Midges used in the temperature profile were homogenized in 8.3 X 10"2 M

Tris-HCl buffer, pH 7.5. Midge larvae used in the time profile, substrate profile, and the standard incubation mixture tested under optimum conditions were homogenized in 8.3 X 10"lM Tris-HCl buffer, pH 7.5, and were incubated for 15 min.

All data reported for whole body homogenate assays represents mean values from 2 experiments, with duplicate treatments/experiment, performed on different days. 29

J. Microsome Preparation

Fourth instar midge larvae, 1000/experiment, were weighed to the nearest 0.1 mg and homogenized in a 8.3 X

lO-iM Tris-HCl buffer, pH 7.5.

Preparatory centrifugations were performed in a

Sorvall RC2-B Model centrifuge with a SS-34 rotor. The

homogenate was sedimented at 2,400 g max (4,500 RPM)

for 15 min at 1 - 1°C to remove large cell fragments

and debris. The supernatant was removed with a pipette

and centrifuged at 20,000 g max (13,000 RPM) for IS min

at 1 - 1°C to isolate the mitochondrial pellet. The

post-mitochondrial supernatant was removed with a

pipette and placed in 10 ml cellulose nitrate tubes,

each tube containing microsome material equivalent to

100 larvae, and sedimented in a Beckman Model L centri­

fuge with a 50 rotor at 71,000 g max (30,000 RPM) for 1 hr at -12 - 1°C to isolate the microsomal pellet in the

initial experiment. The maximum centrifugal force was

increased to 127,000 g max (40,000 RPM) to isolate the microsomes in subsequent experiments because high epoxidase activity was found in the post-microsomal

fraction of the first experiment (Table 11).

The presence of hemoglobin, or hemoglobin components, may interfere with certain enzyme assays by binding with microsomes (Garfinkel 1958, Petermann and Pavlovec 30

1961, Maines and Kappas 1975). Three microsome pellets were transferred to a test tube and resuspended in 1.5 X

10_1M KC1, pH 7.5, using a glass rod and a vortex, in an attempt to reduce hemoglobin contamination and increase enzyme activity. The material was recentrifuged and the washed microsomes were re-isolated using the 127,000 g max force as previously described.

All equipment, solutions, and glassware were pre- cooled prior to larval homogenization and microsome isolation. All processed material was kept in crushed ice until incubated.

K. Assay of Subcellular Fractions for Aldrin Epoxidation

Mitochondrial and microsomal pellets were resuspended in Tris-HCl buffer, using a glass rod and a vortex. Each

5 ml volume incubation mixture contained the following:

2.4 X 10"3M G-6-P, 1.6 units G-6-P d H , 5.1 X 10"SM NADP,

1.0 mg aldrin in 0.2 ml ethanol when used alone or 1.0 mg in 0.1 ml ethanol when combined with PBO in the incubation mixture. The various fractions contained 150 larvae equivalents of mitochondrial pellet material, or

30 larvae equivalents of one of the following: whole body homogenate, microsome pellet material, or post-microsomal supernatant, each in 5.0 X lO'-^M Tris-HCl buffer, pH 7.5.

Incubation mixtures which included a synergist contained

1.0 mg PBO in 0.1 ml ethanol, and were pretreated for 3-5 31

min prior to the addition of substrate. Enzyme assays

were initiated by the addition of aldrin. The reaction

mixtures were agitated with a vortex, incubated in test

tubes at 30 - 1°C, and were swirled at 100 RPM in a

Metabolyte Water Bath Shaker for 15 min. Reactions were

stopped by acidifying with 2 ml of 51 TCA. The acidified

mixtures were extracted and analysed by GLC (see

extraction and chromatographic procedures).

All data reported for subcellular fraction enzyme

assays represents mean values from 2 experiments, with

duplicate treatments/experiment, performed on different

days.

L. Protein Analysis

Protein concentrations were determined with a

Spectronic 20 (Bausch and Lomb) spectophotometer

(Bramhall et_ al. 1969) employing BSA as a standard. Each

0.1 ml sample to be analyzed was spotted on 3 cm2

Whatman No. 42 filter paper and air dried. Samples were

stained with xylene brilliant cyanin G (K and K

Laboratories3 Cleveland, Ohio), and the absorbance at

610 nm was recorded against a blank containing distilled water. Samples were corrected using treated controls, blanks containing all components except protein. III. RESULTS

A. Toxicity Assays

The computed LCgg and its 95% confidence interval with and without synergist, the regression coefficient

(slope), and the synergistic ratios (SR) for each insecticide assayed with midges are contained in Table 1, and with mosquitoes in Table 2. The LC50 values for experiments with no control mortality were not signifi­ cantly different from those requiring correction for mortality in acetone or synergist controls. Control mortality for midge larvae was 5% or less in all experi­ ments. Most (^75%) experiments had no control mortality. There was no acetone or synergist control mortality in mosquito assays.

Dieldrin (LCso» 0.5 ppb) was the most toxic of all insecticides assayed with midges. The three other organochlorines tested, aldrin, lindane, and DDT, had

LC50 values of 0.8, 3.6, and 4.7 ppb respectively.

The LC5q values for the two organophosphate insecticides, parathion and malathion, and their initial oxidative metabolites, paraoxon and malaoxon, collectively

32 I

TABLE 1

Toxicity of Selected Insecticides With and Without Synergist to Fourth Instar Larval Chironomus riparius3

24 hr LC5q (95% C.I.), Regression Synergistic Insecticide ppb Coefficient Ratio

Ortrnnoch] orine

DDT 4.7 (4.2-5.3) 4.3

DDT+Sesamexb 2.9 (2.6-3.2) 5.4 1.63

Aldrin 0.8 (0.7-0.8) 10.9

Aldrin+PBOb 2.2 (1.9-2.6) 2.9 0.33

Dieldrin 0.5 (0.4-0.6) 4.0

Dieldrin+PBOb 0.3 (0.3-0.4) 2.4 1.39

h Lindane 3.6 (3.1-4.0) 3.0

Organophosphate

Parathion 2.5 (1.7-4.1) 2.1

Parathion+PBOb 17.1 (14.3-21.6) 2.5 0.15 TABLE 1 (CONTINUED)

24 hr LCrn (95% C.I.) , Regression Synergistic Insecticide ppb Coeffici cnt Ratio

Paraoxon 6.2 (5.8-6.7) 6.9

Paraoxon+FBO11 5.5 (4.8-6 .5) 3.9 1.13

Maiathion 1.9 (1.7-2.2) 5.5

Malathion+PBO^ 6.8 (6.0-7.8) 7.0 0.28

Malathion+EPN^ 2.4 (2.1-2.9) 3.1 0.81

Malaoxon (5.4 (4.9-5.9) 4.7

Malaoxon+Sesamex^ 1.9 (1.6-2.3) 6.3 2.78

Carbamate

Carbaryl 104.5 (83.3-151.7) 2.6

Carbaryl+PBO^ 62.4 (54.0-72.9) 2.9 1.68

Landrin^- 51.4 (45.0-59.3) 3.1

Landrin^+PBO^ 45.7 (37.5-54.5) 2.4 1.12

Aminocarb 376.6 (329.7-428.5) TABLE 1 (CONTINUED)

i 24 hr LC50 (951 C.I.), Regression Synergis Insecticide ppb Coefficient Ratio

Aminocarb+PBOb 1,172.2 (1,072.0-1,308.1) 6.0 0.32

Mexacarbate 12.2 (11.0-13.5) 5.3 -

Mexacarbate+PBOb 59.3 (52.9-66.2) 3.6 0.20

Propoxur 64.4 (59.5-69.4) 6.0 -

Propoxur+PBOb 29.2 (25.2-33.8) 2.9 2.21

Synthetic Botanical

Allethrin 41.9 (38.6-45.3) 5.6 -

Allethrin+PBOb 0.4 (0.3-0.5) 2.9 102.10

Synergist

PBO 2,700.0 6.3

t aValues represent 3 experiments, with 3-7 concentrations/experiment. TABLE 1 (CONTINUED)

^Larvae were given a 1 hour 1 ppm PBO, 1 ppm sesamex, or 0.5 ppb EPN pretreatment. Insecticide-inhibitor mixtures contained the pretreatment concentration of inhibitor, cThis value was determined by sight; all other values were determined by computer analysis. TABLE 2

Toxicity of Selected Insecticides With and Without Synergist to Late Third and Fourth Instar Larval Aedes aegypti3

24 hr LC50 (955 C.I.), Regression Synergistic Insecticide ppb Coefficient Ratio

Organochlorine

DDT 13.6 (11.7-15.9) 2.9

Aldrin 16.5 (14.7-18.3) 4.1 '1 Aldrin+PBOb 14.5 (9.8-18.2) 3.4 1.14

Dieldrin 16.5 (14.8-18.2) 5.1

Lindane 1 78.2 (72.8-84.5) 6.2

Organophosphate

Parathion 21.7 (19.1-24.2) 4.9

Parathion+PBOb 16.6 (14.5-18.5) 4.1 1.31

Malathion 373.8 (322.9-446.7) 4.9 TABLE 2 (CONTINUED)

24 hr LC50 (951 C.I.), Regression Synergistic Insecticide ppb Coefficient Ratio

Malathion+PBOb 305.2 (251.2-362.4) 7.0 1.22

Carbamate

Carbaryl 5,162.5 (4,691.6-5,788.5) 3.7

Carbaryl+PBOb 2.326.4 (2,075.5-2,563.7) 4.4 2.22

Mexacarbate 1.762.5 (1,470.8-2,026.0) 3.6

Mexacarbate+PBOb 1.420.6 (1,269.2-1,641.9) 4.1 1.23

Synthetic Botanical 1

Allethrin 176.2 (158.5-196.5) 4.4

Allethrin+PBOb 68.2 (60.8-73.9) 5.3 2.50 aValues represent 3 experiments, with 3-7 concentrations/experiment.

^Larvae were pretreated for 1 hour with 1 ppm PBO. Insecticide-inhibitor mixtures contained the pretreatment concentration of inhibitor. 39

ranged from LC50 1.9-6.2 ppb. Parathion was slightly

less toxic than malathion, and both oxidative metabolites

were slightly less toxic than their parent compounds.

Carbamate insecticides were generally less toxic to

midge larvae than organochlorine and organophosphate

insecticides. The toxicity of the five carbamate

insecticides can be grouped into two categories based on

closeness of LC50 values: mexacarbate, LandrinR , and

propoxur, with a LC50 range of 12.2-64.4 ppb, and

carbaryl and aminocarb, with LC50 values of 104.5 and

376.6 ppb.

The LC50 of allethrin, a synethic pyrethroid,

assayed with midge larvae was 41.9 ppb.

The range for the regression coefficients (RC)

of all insecticides assayed with midge larvae was

generally medium to high (3.0-10.9) except for parathion

and carbaryl which had RC values of 2.1 and 2.6

respectively.

The LC50 values for DDT, aldrin, dieldrin, and

lindane assayed with mosquito larvae were of the same

order of magnitude of toxicity; ranging from 13.6-78.2 ppb.

The organophosphate insecticides were more variable. 40

The LC50 values for A_. aegypti larvae was 21.7 ppb for parathion and 373.8 ppb for malathion.

The two carbamate insecticides carbaryl and mexa­

carbate, LC50 5,162.5 and 1,762.5 ppb respectively, were the least toxic of all the classes of insecticides assayed with mosquito larvae.

The LC50 value of allethrin for mosquito larvae was

17 6.2 ppb.

The range of the RC values for all insecticides assayed with mosquito larvae was 2.9-6.4.

The higher tolerance of mosquito larvae to insecti­ cides when compared to midge larvae tested under identical conditions can be grouped into three categories of insecticide tolerance, based on the ratio LC50 mosquito larvae/LCso midge larvae: mosquito larvae are 2.9, 4.2, and 8.7 times more tolerant to DDT, allethrin, and parathion; 20.6, 21.7, 33.0, and 49.4 times more tolerant to aldrin, lindane, dieldrin, and carbaryl; and, 144.5 and 196.7 times more tolerant to mexacarbate and malathion, respectively. Overall, LC50 ratios of mosquito larvae: midge larvae varied from approximately

3-200. Chironomus riparius larvae were more susceptible than A. aegypti larvae to all representatives of the major classes of insecticides. 41

Low synergistic ratios (less than 0.5 indicating an antagonistic response in midge larvae assays) resulted with aldrin (SR 0.33), parathion (SR 0.15), malathion

(SR 0.28), aminocarb (SR 0.32), and mexacarbate (SR 0.20), when the larvae were pretreated for 1 hr with 1 ppm PBO and then transferred to a 1 ppm PBO-insecticide combination. PBO synergized allethrin, a fast acting synthetic pyrethroid; the LC^q value for allethrin was

102 times greater than the LC50 value for allethrin +

PBO. Assays where synergism was weak or absent (ratios from 0.81-2.78) included: malathion + EPN, malaoxon + sesamex, propoxur + PBO, carbaryl + PBO, DDT + sesamex, dieldrin + PBO, paraoxon + PBO, and Landrin^ + PBO

(Table 1).

A synergistic response could not be demonstrated using PBO as the synergist for most mosquito larvae ~ assays. Allethrin and carbaryl were weakly synergized

(SR 2.50 and 2.22 respectively), while aldrin, parathion, malathion, and mexacarbate were not synergized by PBO

(Table 2).

B. Absorptive Uptake and Insecticide Loss

Midge larvae immersed in 20 ppb aldrin for 2 hrs absorbed 25.4 ng of insecticide per larva, converting 42

73% of it to dieldrin (Table 3). Larvae pretreated for

1 hr in 1 ppm PBO and transferred to 20 ppb aldrin with

1 ppm PBO absorbed 18.4 ng of aldrin per larva, but no aldrin was converted to dieldrin.

With or without PBO, midge larvae absorb dieldrin at the same rate as aldrin (Table 3).

Midge larvae absorbed 13.3 ng lindane/larva without further conversion (Table 3).

All insecticides affected the behavior of C. riparius larvae in a similar manner. The first symptom noted was an interference with larval motility. A normal

3 cycle swimming motion was generally reduced to 1 cycle at the onset of toxic symptoms. The loss of coordinated movement increased until the larva could not swim.

Larvae at this stage of intoxication were capable of jerky, spasmodic movements, and tended to curl into a coil when touched with a probe. The effect of the toxicant increased until the larva lost all ability to move; death soon followed. Changes in larval color vrere found to be unreliable for determining toxic affect, and were not considered when larvae were evaluated at the end of each assay. Similar symptoms of insecticide poisoning have been reported for stonefly naiads (Jensen and Gaufin 1964, Sanders and Cope 1968). TABLE 3

The Absorption of Selected Insecticides by C. riparius Larvae and Pupaea

Aldrin Dieldrin Total Insecticide ng/organism (ug/gm) ng/organism (ug/gm) ng/organism (ug/gm)

MIDGE LARVAE

Aldrin 10.7 (0.8) 14.7 (1.1) 25.4 (1.9)

Dieldrin 23.9 (1.9) 23.9 (1.9)

Aldrin+PBOb 18.4 (1.5) 18.4 (1.5)

Dieldrin+PBC)b 24.9 (2.0) 24.9 (2.0)

Lindane 13.3 (1.2)

MIDGE PUPAE

Aldrin 7.6 (0.9) 7.6 (0.9)

Dieldrin 10.2 (1.4) 10.2 (1.4)

Lindane 3.2 (0.4)

aMean values from 2 experiments, 20 ppb insecticide, 2 hour immersion.

^Larvae were given a 1 hr, 1 ppm PBO pretreatment and then immersed in a 1 ppm PBO-insecticide mixture. 44

Toxic symptoms were observed in approximately

50-75% of the midge larvae immersed in 20 ppb aldrin at the end of the 2 hr exposure and they were recorded as moribund. All larvae immersed in dieldrin, lindane, or dieldrin with PBO were moribund. Since all larvae pretreated in PBO and immersed in aldrin with PBO were unaffected, the toxic symptoms must be associated with the presence or accumulation of dieldrin for the test conditions used.

When tested under identical conditions, midge pupae absorbed insecticide at a slower rate than larvae: 7.6 ng aldrin/pupae, 10.2 ng dieldrin/pupae, and 3.2 ng lindane/pupae (Table 3) .

Mosquito larvae immersed in 20 ppb aldrin for 2 hrs absorbed 4.7 ng aldrin/larva, and converted 55% of it to dieldrin (Table 4). _

With or without PBO, mosquito larvae absorb dieldrin at the same rate. The amount of dieldrin absorbed by mosquito larvae was approximately 50% that of the aldrin absorbed under identical conditions (Table 4).

Mosquito larvae immersed in aldrin alone or pre­ treated in 1 ppm PBO for 1 hr and then transferred to a

20 ppb aldrin - 1 ppm PBO mixture for 2 hrs absorb equal amounts of insecticide, but the amount of aldrin con­ verted to dieldrin was reduced 15% in the presence of TABLE 4

The Absorption of Selected Insecticides by A. aegypti Larvae and Pupaea

Aldrin Dieldrin Total Insecticide ng/organism (ug/gm) ng/organism (ug/gm) ng/organism (ug/gm)

MOSQUITO LARVAE

SERIES 1

Aldrin 2.2 (0.6 2.6 (0.7) 4.7 (1.3)

Dieldrin 2.6 (0.7) 2.6 (0.7)

Aldrin+PBOb,e 2.8 (0.8) 1.9 (0.5) 4.7 (1.4)

Dieldrin+PBOb>e 2.4 (0.6) 2.4 (0.6)

SERIES 2

Aldrin 2.6 ( 0 . 6) 2.2 (0.5) 4.7 (1.0)

Aldrin+PBOc*e 1.0 (0 .2) 1.1 (0.3) 2.1 (0.5)

Aldrin+PBOd>e 1.1 (0.3) 1.1 (0.3) 2.2 (0.5)

MOSQUITO PUPAE

Aldrin 0.5 (0.2) Trace 0.5 (0.2) £ TABLE 4 (CONTINUED)

I aMean values for 2 experiments, 20 ppb insecticide, 2 hour immersion.

^Larvae were pretreated for 1 hour with 1 ppm PBO. cNo pretreatment was given.

^Larvae were pretreated for 24 hours with 2 ppm PBO. eTnsecticide-PBO mixtures always contained 1 ppm PBO. PBO (Table 4). Larvae pretreated for 24 hrs in 2 ppm

PBO or larvae without PBO pretreatment that were trans­ ferred to a 20 ppb aldrin - 1 ppm PBO mixture for 2 hrs absorbed less insecticide (55% and 53% respectively) than larvae immersed in aldrin alone. Although less aldrin was absorbed in the presence of PBO, larvae with no pretreatment and larvae pretreated for 24 hrs converted approximately 50% of absorbed aldrin to dieldrin, which was 50% less than larvae immersed in only aldrin (Table 4).

No toxic symptoms were observed in mosquito larvae immersed for 2 hrs in the insecticides or insecticide- synergist combinations used in this series.

Midge larvae immersed in 10 ppb aldrin for 6 hrs absorbed insecticide (aldrin + dieldrin) in proportion to exposure time during the first 4 hrs, reached a maximum between 4-5 hrs, and then decreased between hrs

5-6 (Figure 5). The amount of parent compound, aldrin, extracted from each larva increased rapidly for 3 hrs, reached a maximum at 4 hrs, and then decreased between hrs 4-6. The amount of converted aldrin, as dieldrin, extracted from.each larva increased in proportion to exposure time during the first 5 hrs and Figure 3. The effect of exposure time on fn vivo

insecticide absorption and metabolism by

C. riparius larvae immersed in 10 ppb

aldrin. Values represent mean values from

2 experiments: • , total insecticide;

aldrin;^ > dieldrin.

48 NG INSECTICIDE/LARVA 4.0 2.0 6.0 8.0 2 20 14 18 16 12 10

2 1 TI iue 3 Figure IA E : H O U H S 3 5 4 6 so

then decreased during hrs 5-6. Mosquito_larvae immersed

in 10 ppb aldrin for 6 hrs absorbed insecticide (aldrin

+ dieldrin) in proportion to exposure time throughout

the entire experiment (Figure 4). The conversion of

aldrin to dieldrin in mosquito larvae increased with

exposure time.

While the rate (ng/larva/hr) of aldrin absorption

decreased during 6 hrs in midge larvae (Figure 5), the

amount of aldrin converted to dieldrin increased to a

maximum rate at the five hour interval. These rate

values were computed for each time interval in the

experiment. The rates of aldrin uptake and conversion

to dieldrin were much less variable in mosquito larvae

(Figure 6). Mosquito larvae transformed 47% of their

absorbed aldrin to dieldrin at 1 hr, while midge larvae

converted only 20%. Mosquito larvae continued to

epoxidize a higher percentage of absorbed aldrin, 68%

at 3 hrs and 78% at 6 hrs, than midge larvae, 49% and

72% respectively.

The first toxic symptoms appeared in midge larvae

after 3 hrs and increased in severity until all larvae were moribund at 6 hrs. This may help explain the down­ ward deflection observed in Figures 3 and 5. The onset of toxic symptoms appeared in mosquito larvae at 6 hrs. Figure 4. The effect of exposure time on ill vivo

insecticide absorption and metabolism by

A. aegypti larvae immersed in 10 ppb

aldrin. Values represent mean values

from 2 experiments: • » total insecticide;

® , aldrin; Jk. , dieldrin.

51 i-o m 4^ 0) a> o o o o o • • • • i a NG INSECTICIDE/LARVA NG I 01 0)

• • • m X o c X

Figure Figure 5. The effect of exposure time on the iii vivo

hourly rate of insecticide absorption and

metabolism/^, riparius larva immersed in

10 ppb aldrin. Values represent mean

values from 2 experiments: • , total

insecticide; ■ , aldrin; ^ , dieldrin.

53 cn o> ro Q NG NG INSECTICIDE/LARVA/HOUR ro 0) 01 w m O c ^ 33 0)

Figure tn Figure 6 . The effect of exposure time on the iri vivo

hourly rate of insecticide absorption and

metabolism/A. aegypti larva immersed in

10 ppb aldrin. Values represent mean

values from 2 experiments: • , total

insecticide; ■ , aldrin; ^ , dieldrin.

55 to hour / larva / insecticide ng o -*> ro Ol

• * • m H 2 co x o 5 33 *

Figure 57

Even though mosquito larvae absorb less aldrin than midge larvae, they are more efficient in the epoxidation of aldrin (Figure 7). A similar relationship was obtained when conversion efficiencies were compared on a weight basis.

Midge larvae immersed in aldrin concentrations ranging from 10-100 ppb for 1 hr absorbed insecticide in proportion to concentration (Figure 8). The amount of aldrin and dieldrin extracted from each larva was also linearly related to concentration. Mosquito larval uptake and conversion was less than midges when immersed in 10-100 ppb aldrin for 1 hr, and was also proportional to concentration (Figure 9) .

During a 1 hr exposure, the conversion of aldrin to dieldrin in midge larvae was linear over all insecticide concentrations (Figure 10). The curve for dieldrin formation in mosquito larvae was biphasic, decreasing between 10-40 ppb, and remaining constant between 40-100 ppb (Figure 10).

Approximately 20% of the midge larvae immersed in the 80 ppb and 100 ppb aldrin solutions had toxic symptoms at the end of 1 hr. No toxic symptoms were observed in mosquito larvae.

As observed in the time-variable experiment Figure 7. Dieldrin production as a function of total

aldrin absorbed for larvae immersed in 10

ppb aldrin. Each point represents

different exposure times over a six hour

period. Values represent mean values from

2 experiments: (— ■), £. riparius larvae;

(----), A. aegypti larvae.

58 o to o to r NG NG DIELDRIN PRODUCED/ LARVA log scale) I h o to o o cn oo

NG INSECTICIDE ABSORBED/LARVA Hog scale! Figure 8. The effect of aldrin concentration on

in vivo C. riparius larval absorption and

metabolism for a 1 hour immersion. Values

represent mean values from 2 experiments:

• , total insecticide; ■ , aldrin;

A. > dieldrin.

60 ng insecticide / larva 0 8 70 0 6 4 4 0 0 5 0 3 20 10 20 n i r d l a iue 8 Figure

0 4 n o i t a r t n e c n o c 0 6 0 8 n : B P P 100 B r> i Figure 9. The effect of aldrin concentration on in

vivo A. aegypti larval absorption and

metabolism for a 1 hour immersion. Values

represent mean values from 2 experiments:

• , total insecticide; ■ , aldrin;

a . , dieldrin.

62 NG INSECTICIDE/LARVA 4.0 6.0 8.0 10 20 LRN OCNRTO: PPB CONCENTRATION: ALDRIN iue 9 Figure 6040 80 100 fi3 Figure 10. The effect of aldrin concentration on

percent of aldrin epoxidation for a 1

hour immersion. Values represent mean

values from 2 experiments: ( ■) > C..

riparius larvae; (---- ), A. aegypti larvae.

64 9 O _j. _j. o ro o co o A o ALDRIN CONVERTED/LARVA %

Figure 10 66

(Figure 7), mosquito larvae are more efficient in the

conversion of aldrin to dieldrin than midge larvae over

the range of concentrations tested (Figure 11). Similar

results were obtained when corrections were made for weight.

The amount of insecticide absorbed by midge larvae

is dependent upon larval size, larger larvae absorbing

more than smaller larvae. Late second or early thrid

instars with an average weight of 2 mg/larva immersed

in 20 ppb aldrin for 2 hrs absorbed 14.2 ng/larva,

converting 9.8 ng/larva to dieldrin. Larger late third

or early fourth instars with an average weight of 6 mg/

larva absorbed 24.5 ng/larva and converted 14.1 ng/

larva to dieldrin. The largest larvae tested in this

series, late fourth instars with an average weight of

9 mg/larva, absorbed the greatest amount of insecticide~

31.8 ng/larva, and converted 19.9 ng/larva to dieldrin.

The amount of aldrin epoxidized by all larva ranged

from 591-69%. Positive regression coefficients result when the amount of insecticide/larva is plotted against average wet weight/larva (Figure 12). Negative regression coefficients result when corrections are made for weight differences and ng/g of larva are plotted against average wet weight (Figure 12). On a larval Figure 11. Dieldrin production as a function of

total aldrin absorbed for a 1 hour

immersion. Values represent mean values

from 2 experiments: (1 — ), C. riparius

larvae; (----), A. aegypti larvae.

67

00 ) ) • • I •• 0 o o o I log scalel log I \ a \ o T - p - m j Q o ro larva / -fc -fc o o 9 o CO • • • ■ O O TTTT produced O A

ro o dieldrin NG NG 2 2 4 6 810 20 40 60 80 100

INSECTICIDE ABSORBED/LARVA [log scale] Figure 12. The effect of size on iii vivo insecticide

absorption and metabolism by C. riparius

larvae immersed in 20 ppb aldrin for 2

hours. Values represent mean values from

2 experiments: (— , and left ordinate),

insecticide/gm larvae; (-----, and right

ordinate), insecticide/larva; • , total

insecticide; ■ , aldrin; , dieldrin.

69 NG 1NSECTI C1DE/GM LARVAE llog scalel 10000 0 0 0 8 2000 0 0 0 4 0 0 0 6 1000 1 AVERAGE WET WEIGHT: MG WEIGHT: WET AVERAGE 2 Figure o scale] Hog 4 12 6

8

10

0 4 20 10

NG INSECTICIDE/LARVA llog scale] 70 71 basis, large larvae accumulate more insecticide, but on a weight basis they accumulate progressively less.

A small amount of insecticide was released by midge larvae containing dieldrin when they were trans­ ferred to clean water. Larvae placed in 20 ppb dieldrin for 1 hr and then transferred to clean water for 3 hrs released 0.5 ng of dieldrin/larva/hr. A final group of larvae transferred to a second volume of clean water for an additional 3 hrs, or a total of 6 hrs, continued to release dieldrin to the water at approximately the same rate, 0.4 ng/larva/hr (Table 5).

Midge larvae immersed in 40 ppb aldrin for 1 hr where anterior and posterior halves were separated by a striction absorbed and metabolized approximately equal amounts of insecticide (Table 6).

C. Assay of Whole Body Homogenate for Aldrin

Epoxidati on

Preliminary tests with whole body homogenates and component chemicals included in the incubation mixture resulted in an increase in dieldrin production from

15.2 pmoles/insect/minute for homogenate only to

52,7 pmoles/insect/minute for the complete incubation mixture (Table 7). TABLE 5

. Depuration of Dieldrin by C. riparius Larvaea

Time Dieldrin Dieldrin in Water Depuration Rate Hours ng/Larva ng ng/Larva/Hour

0 19.0

3 19.5 1.3 0.5

6 17.8 1.0 0.4

aLarvae were immersed in 20 ppb dieldrin for 1 hour and then transferred to insecticide-free water; values represent mean values from 2 experiments, 20 larvae/treatment. TABLE 6

The Uptake and Metabolism of Aldrin by Ligatured C. riparius Larvaea

Total Insecticide Aldrin Dieldrin Larval Half ng/section ng/section ng/section

Anterior 13.6 3.7 9.9

Posterior 14.4 2.4 12.0

aLarvae .with strictures were immersed in 40 ppb aldrin for 1 hour; values represent mean values from 2 experiments, 20 pooled larval halves/experiment. 74

TABLE 7

Aldrin Epoxidase Requirements of C, n p a n u s Whole Body Homogenatesa

Activity Percent of nmoles dieldrin/ Maximum Incubation Medium^5 insect/minute Activity

Crude Homogenate 0.015 28

+ 1% BSA (W/V) 0.016 30

+ 1% BSA (W/V) 0.03 8 72

G-6-P (2.4 X 10"3M)

G-6-P dH (1.6 units)

NADH (5.1 X 10-5m )

NADP (5.1 X 10"SM)

+ KC1 (2.7 X 10-3M0 0.053 100 and all of the above

aMean values from 2 experiments, with 2 treatments/ experiment. Optimum conditions were not used.

^Larvae were homogenized in 8.3 X 10“2m Tris-HCl, pH 7.5 buffer. Complete incubation medium (final concentrations): 3 ml of homogenate (20 larvae); 5.0 X 10"2m Tris-HCl, pH 7.5 buffer; 1.0 mg aldrin in 0.5 ml ethanol; total volume, 5 ml. Mixtures were incubated at 30°C for 1 hour. 75

The effect of pH on in vitro aldrin epoxidase activity was established over a pH range 6 .5-8.5

(Figure 13). Epoxidase activity increased from a pH

6.5 until reaching a maximum at a pH 7.5. Further increases in alkalinity resulted in a slight decrease in activity until pH 8.0, followed by a rapid loss in activity for pH values above 8.0. Since C. riparius whole body homogenates had an optimum pH range of pH

7.5-8.0, a pH of 7.5 was used for all subsequent experiments.

The effect of temperature on in^ vitro aldrin epoxidase activity was determined for a temperature range 20°-40°C (Figure 14). Epoxidase activity increased as incubation temperature increased, reaching a maximum at 30°C. Increasing the temperature above the

30"^ optimum resulted in a rapid decrease in enzyme activity. £. riparius whole body homogenates had an optimum incubation temperature of 30°C, and this incubation temperature was used in all subsequent experiments.

Epoxidase activity increased as the Tn vitro molar concentration of Tris-HCl buffer increased; the maximum activity was attained at a buffer concentration of

5.0 X 10“1m (Figure 15). An increase in buffer Figure 13. The effect of pH on iii vitro aldrin

epoxidation by midges. Values represent

mean values from 2 experiments, with 2

treatments/experiment.

76 MAXIMUM ACTIVITY 100 0 8 0 4 0 6 20 6.5 iue 13 Figure 7.0 .7.5 H p 8.0 8.5 Figure 14. The effect of temperature on in vitro

aldrin epoxidation by midges Values

represent mean values from 2 experiments,

with 2 treatments/experiment

78 MAXIMUM ACTIVITY 100 0 8 0 6 0 4 20 5 3 0 3 5 2 0 2 EPRTR: ° TEMPERATURE: V JL iue 14 Figure c X 0 4 79 Figure 15. The effect of buffer concentration

(molarity) on In vitro aldrin epoxidation

by midges. Values represent mean values

from 2 experiments, with 2 treatments/

experiment. MAXIMUM ACTIVITY’ 100 80 0 6 20 - / . ' ' ' 0.0 5 < / UFR OCNRTO: MOLAR CONCENTRATION: BUFFER ■ / / iue 15 Figure Nf / 025 0.5 5 1.0 5 7 . 0 0 5 . 0 5 .0.2 I 1 a R1 82

concentration above 5.0 X 10"^M produced a rapid

decrease in epoxidase activity. The optimum 5.0 X 10"^M

Tris-HCl buffer concentration was used in all subsequent

experiments.

The amount of dieldrin formed per insect increased

as incubation time increased from 15 min to 30 min and

then approached a plateau over a following 45 min

interval (Figure 16). The time rate analysis of dieldrin

production/insect/minute, however, showed a negative

linearity when plotted against time for the 75 minute

test period (Figure 16). Because of the decrease in

dieldrin production observed in the time rate analysis,

a 15 minute incubation time was used in all subsequent

experiments.

The iri vitro aldrin epoxidase activity was linearly correlated with "homogenate concentration (enzyme concentration) over a range of 1 larva/ml to 5 larvae/ ml (Figure 17).

The rate of dieldrin production increased from 25.7 to 141.3 pmoles/insect/minute when the substrate concen­ tration was increased, respectively, from 0.01 mg to

1.0 mg in a final volume of 5 ml of incubation mixture

(Table 8) .

After the completion of preliminary experiments Figure 16. The in_ vitro rate of aldrin epoxidation

in nanomoles/larva/minute ( , and left

ordinate), and total aldrin epoxidation/

larva (---- , and right ordinate). Values

represent mean values from 2 experiments,

with 2 treatments/experiment.

85 co r o mJk « ■ o o o o N-MOLES DIELDRIN PRODUCED/LARVA scalel Hog N-MOLES DIELDRIN PRODUCED/LARVA/MINUTE scale log * o I o U1 w o o H c m co

Figure 16 Figure 17. The effect of larval numbers (enzyme

concentration) on iii vitro aldrin

epoxidation. Values represent mean

values from 2 experiments, with 2

treatments/experiment.

85 NG DIELDRIN PRODUCED/TUBE 0 0 0 0 3 20000 10000 0 0 0 5 2 0 0 0 5 1 0 0 0 5 1 iue 17 Figure 2 LARVAL NU&ESR 3 4 j L*L L 8 6 87

TABLE 8

The Effect of Substrate Concentration (Aldrin) and BSA Upon iii vitro Aldrin Epoxidase of C. riparius Whole Body Homogenatesa

Activity Substrate Concentration nmoles dieldrin/ mg Aldrin/Incubation % BSA (W/V) insect

1.0 1.0 1.91

1.0 0.1 2.12

0.1 1.0 1 . 20

0 . 01 1.0 0.39

aMean values from 2 experiments, with 2 treatments/ experiment. ~

^Optimum preparatory and incubation conditions of Table 9 were used. 88

establishing reaction conditions, the effect of certain

chemical supplements upon in vitro aldrin epoxidase

activity was re-examined under optimum conditions. There

was a significant increase in dieldrin production

(Table 9) compared with conditions used prior to

determining optimum conditions (Table 7). Several

chemicals used in preliminary incubation mixtures, and

throughout experiments used in determining optimum

conditions, had no effect upon dieldrin production when

optimum conditions were used. These are BSA, KC1, and

NADH (Table 9).

Optimizing assay conditions produced approximately

a fourteen-fold enhancement, when the aldrin epoxidase

activity of whole body homogenate using initial

conditions (Table 7) is compared to homogenate activity

using optimum conditions (Table 9). This conclusion

is based upon the following assumptions: 1) dieldrin production/larva for 30 min equals that for 60 min, or

0.015 nmoles/insect; 2) dieldrin production/larva for

30 min approximates two times the dieldrin production for

15 min. Therefore, an estimated 0.008 nmoles of dieldrin was produced/insect under initial conditions compared to 0.116 nmoles of dieldrin produced/insect under optimum 89

TABLE 9

Aldrin Epoxidase Requirements of £. riparius Whole Body Homogenates Under Optimum Conditionsa

Activity Percent of nmoles dieldrin/ Maximum Incubation Medium** insect/minute Activity

Crude Homogenate 0.116 36

+ (electron generator) 0. 324 100

G-6-P (2.4 X 10"3M)

G-6-P dH (1.6 units)

NADP (5.1 X 10"5M)

+ KC1 (2.7 X 10-3M) and electron generator 0.314 97

+ 0.1% BSA (W/V), KC1, and electron generator 0. 279 86

+ NADH (5.1 X 10“5M) and all of the above 0. 275 85

^ean values from 2 experiments, with 2 treatments/ experiment. Optimum conditions were used.

^Larvae were homogenized in 8.3 X 10”-*M Tris-HCl, pH 7.5 buffer. Complete incubation medium (final concentrations): 3 ml of homogenate (20 larvae); 5.0 X 10“lM Tris-HCl, pH 7.5 buffer; 1.0 mg aldrin in 0.5 ml ethanol; total volume, 5 ml. Reaction mixtures were incubated at 30OC for 15 minutes. 90

conditions. The addition of G-6-P, G-6-P dH, and NADP

increased dieldrin production to 0,324 nmoles/insect

(Table 9), an approximate forty-fold increase, when

compared to the aldrin epoxidase activity found in the

homogenate under initial experimental conditions (Table

7) .

The addition of FMN decreased dieldrin production,

while the addition of FAD resulted in a slight increase

in ill vitro aldrin epoxidase activity (Table 10) . The

addition of PBO to whole body homogenates prevented the

in vitro oxidative transformation of aldrin to dieldrin

(Table 10).

D. Assay of Subcellular Fractions for Aldrin Epoxidation

In the initial experiment of this series, mito­

chondrial and microsomal subcellular fractions had

similar epoxidase activity, while the greatest activity was found in the post-microsomal supernatant (Table 11).

Ir is also evident that washing the microsomes in KC1 prior to incubation increased epoxidase activity and that PBO prevented the formation of dieldrin by the microsomal fraction (Table 11). A modification of the preparatory procedure in subsequent experiments resulted in a change in the distribution of epoxidase activity 91

TABLE 10

Effect of FMN, FAD, and PBO Upon iji vitro C. riparius Aldrin Epoxidase Whole Body Homogenatesa

Activity nmoles dieldrin/ Incubation Medium insect/minute Percent

Complete^ 0.081 100c

FMN (10'5M) 0.075 92

FAD (10_3M) 0.089 110

PBO (6.0 X 10"4M)d N.D.e 0

aMean values from 2 experiments, with 2 treatments/ experiment.

^Optimum preparatory and incubation conditions of Table 9 were used. cStandard. dHomogenates were pretreated for 15 minutes with PBO in 0.25 ml ethanol prior to the addition of aldrin. eNot detected. TABLE 11

Subcellular Localization of Aldrin Epoxidase Activity in C. riparius Larvaea

Specific Activity^ Specific Activity0 pmoles/mg protein/min; pmoles/mg protein/min; Fraction (pmoles/insect equiv./min) (pmoles/insect equiv./min)

Homogenate (80.0) 236.0 (86.7)

Mitochondrial 1.334.0 (9.0) 757.0 (6.6)

Microsomal 779.0 (9.0) 798.0 (19.4)

Microsomal (washed) 1.033.0 (13.0) 1,303.5 (23.8)

Microsomal+PBO^ N.D.e N.D.

Post-microsomal Supernatant 77.0 (23.0) N.D. a0ptimum preparatory and incubation conditions of Table 9 were used.

^Values from 1 experiment, with 2 treatments; post-mitochondrial supernatant centrifuged at 71,000g max for 1 hour.

°Mean values from 2 experiments, with 2 treatments/experiment; post-mitochondrial supernatant centrifuged at 128,000g max for 1 hour. cVlicrosomes were pretrcated with 1 mgPBO in 0.25 ml ethanol added to5 ml reaction mixtures for5 minutes prior to the addition of aldrin. TABLE 11 (CONTINUED)

eNot detected of subcellular fractions (Table 11). Low apoxidase activity was limited to the mitochondrial fraction, the remaining high epoxidase activity being restricted to the microsome fraction. Washed microsomes were slightly more active than unwashed microsomes in contrast to the initial experiment. No epoxidase activity was detected in the post-microsomal supernatant, or when PBO was added to incubation mixtures containing microsomes or post-microsomal supernatant. iy, d i s c u s s i o n AND conclusions

A. Toxicity Assays

The larvae of Chironomus riparius are highly suscep­ tible to the representatives of the major classes of insecticides assayed in this study. Previously reported

24 hr LC5o's for midge larvae determined by laboratory assay include several organophosphate insecticides and

Neopynamin for three chironomid species (Mulla and

Khasawinah 1969), representatives from each major class of insecticide assayed with Chironomus tentans (Karnak and Collins 1974) , and seven organophosphate insecticides for three chironomid genera (Ali and Mulla 1976). The corrected LCs q 's for C. tentans with two organochlorines, dieldrin CO-9 ppb) and DDT (19.5 ppb), approximated the values I recorded for £. riparius (Table 1). The ranges of literature values for organophosphate insecticides and midge larvae were variable. The midge larvae assays which most closely approximated the 1.9-6.2 ppb range

I found for C. riparius were: Tanypus grodhausi (field culture) 3.8-11.0 ppb, T. grodhausi (laboratory culture)

0.5-1.5 ppb (Mulla and Khasawinah 1969), and £. tentans

95 96

2.0-6.4 ppb (Karnak and Collins 1974). Other chironomid

species and genera had greater ranges for LC^q values

of organophosphate insecticides: Chironomus sp. 51, 0.42-

8.0 ppb, Goeldichironomus holoprassinus, 0.97-54.0 ppb,

Chironomus sp., 0.60-58.0 ppb, and Tanytarsus sp.., 0.72-

44.0 ppb (Mulla and Khasawinah 1969, Ali and Mulla 1976).

The range for organophosphate insecticides assayed with

Cricotopus sp., 33.0-500.0 ppb, was the highest reported

(Ali and Mulla 1976). Turning to specific organophos-

phates, the toxicity of parathion to C. s p . 51, LC50

5.5 ppb, G. holoprassinus, LC50 1.7 ppb, and T. grodhausi

(field culture), LC50 3.8 ppb (Mulla and Khasawinah 1969) was similar to the LC50 of 2.5 ppb that I recorded for

£.• riparius. Similarly, reported LC^q's were also near

the value I found for allethrin, LC50 13.8 ppb (Karnak

and Collins 1974J and LC50 41.9 ppb (Table 1) and for malathion: C_. sp. 51, LC50 2.1 ppb (Mulla and Khasawinah

1969), C. tentans, LC50 1.7 ppb (Karnak and Collins

1974), and C. riparius, LC50 1.9 ppb (Table 1). The greatest differences in reported values were between

C^. tentans and C^. riparius for carbamate insecticides, with ranges of 1.6-7.0 ppb (Karnak and Collins 1974) and

12.2-376.6 ppb (Table 1), respectively. The LCso's for

C. riparius assayed with mexacarbate, propoxur, and 97

carbaryl were 7, 38, 65-fold greater, respectively, than

the values reported by Karnak and Collins (1974). The

range for the regression coefficients of all insecticides

assayed with C. tentans were low, 0.8-2.7 (Karnak and

Collins 1974), while the majority of values reported for

the three chironomid species assayed with organophosphate

insecticides, 3.1-5.6 (Mulla and Khasawinah 1969), were

in closer agreement to my values for C. riparius (Table

1). The foregoing suggests that the susceptibility of

C^. riparius to insecticides is comparable to other

chironomid species which have been used in laboratory

toxicity assays, allowing for differences due to species

variability, subjective differences in the factors which

determine mortality in midge larvae, and differences

in assay techniques.

The assay technique used to determine LC50 values

for C. riparius larvae was also used to assay nine

insecticides with Aedes aegypti larvae. The following ranges have been determined from reported LC5q's in a review of susceptible laboratory strains of A. aegypti

(Table 12): DDT 3.0-80.0 ppb, lindane 52.0-230.0 ppb, aldrin 3.5-50.0 ppb, dieldrin 4.2-200.0 ppb, parathion

4.3-35.0 ppb, malathion 60.0-300.0 ppb, and carbaryl

1,800.0-4,400.0 ppb. For a more extensive review of TABLE 12

Reported 24 hr LCSg's (ppb) of Susceptible Laboratory Strains of A. aegypti to Representative Insecticides

Reference DDT Lindane Aldrin Dieldrin Parathion Malathion

Sutherland (1964) 5.0 55.0 3.5 4.2 10.1

Shidrawi (1957) 30.0 140.0 50.0 40.0 20.0 230.0 1 \ Matsumura and Brown (1963) 3.0 - 8.0 60.0

Klassen et al. (1965) ' - - - - 14.0 250.0

Brown and Abedi (1960) 80.0 230.0 - 28.0 35.0 300.0

Wharton (1955) 8.0 60.0 - 14.0

Boike and Rathburn (1968) - - - - 25; 30

Pass and Knapp (1966) - 4.3 47.5 TABLE 12 (CONTINUED)

Reference DDT Lindane Aldrin Dieldrin Parathion Malathion

i Fay (3359) 4.0 280.0 80.0 70.0

Estenik (This Study) 13.6 72.8 16.5 16.5 21.7 373.8 100 base-line LCso’s of susceptible A. aegypti larvae, which includes field strains, see Brown and Pal (1971).

Thus, the LC50 values for the O.S.U. strain of A. aegypti

(Table 2) fall within the ranges of LCgg values for

DDT, lindane, aldrin, dieldrin, parathion, and equals a reported LC50 value for allethrin (Granett et^ aJL. 1951) .

The LCso's that I reported for malathion and carbaryl are slightly higher than reported values (Table 2) . A reported LC50 value for mexacarbate (Kutz and Burbutis

1966) was approximately six-fold greater than my value

(Table 2).

The LC50 values for selected insecticides (Table 1) assayed with C. riparius larvae equal the LC50 values reported for other midges, while the LC5Q values for selected insecticides (Table 2) assayed \vith an O.S.U. strain of A. aegypti larvae equal or approximate the

LC50 values reported for other susceptible strains of

A. aegypti. Therefore, the differences in LC5q values

I reported for riparius and A. aegypti larvae when identical assay techniques were used are real. A. aegypti larvae are more tolerant to the insecticides assayed than C. riparius larvae; the LC50 values

(mosquito:midge) over all isnecticides range from 3 to

200-fold greater. ]01

C_. riparius larvae are generally more susceptible to insecticides than certain other non-target, aquatic insects. Three species of stonefly naiads (Pteronarcys californica, Pteronarcella badia, and Claassenia subulosa) and two species of damselfly naiads (Ischura sp. and Ischnura verticalis) are more tolerant to organo- chlorine and/or organophosphate insecticides based upon the ranges of their 24 hr LC50 values (Sanders and Cope

1968, Macek and Sanders 1970, Schoettger 1970). The 24 hr LC50 values for DDT assayed with two species of stonefly (Paragnetina media and Peltoperla maria) , two species of mayfly (Ephemerella subvaria and IE. aurivilli) , Hydrophyche sp., and Isonychia sp. ranged from 20-17,300 ppb (Hitchcock 1965). However,

Hitchcock's LC50 values may be only estimates for these organisms, since assay containers were aerated during the assay procedure and moribund insects were not included in dosage-morta]ity curves. Abate, an organo­ phosphate insecticide, assayed with the stonefly naiad

Togoperla media gave a 24 hr LC50 of 560 ppb (Swabey and Schenk 1967). californica and C. subulosa have the same tolerance to carbamates as (E. riparius , while

P. badia is 21 times more susceptible to carbaryl than

C. riparius (Sanders and Cope 1968). Lestes congener, 102

Notonecta undulata. and Peltodytes sp. adults have LCc;g

values greater than C. riparius for all insecticides

reported, and have a very high tolerance to organophos-

phates and carbamates (Roberts et^ aj. 1973, Federle and

Collins 1976). Chlorpyrifos, an organophosphate

insecticide, assayed with Laccophilus fasciatus adults

and Chaoborus punctipennis larvae gave 24 hr LC50 values

of 2.1 ppb and 5.4 ppb, respectively (Roberts £t al.

1973) , values which are within the range of LC^g values

for C. riparius with organophosphate insecticides

(Table 1) .

Active MFO enzymes may be demonstrated iji vivo if

an MFO inhibitor plus insecticide produces a low

synergistic ratio (interference with an activation) or a

high synergistic ratio (interference with a detoxifica­

tion). -While a significant change in toxicity in

inhibitor-insecticide experiments can be used as evidence

for an active MFO system, the absence of a significant

change may not be conclusive proof that an organism does

not possess an active MFO system.

The PBO-insecticide toxicity assays suggest that

C. riparius larvae have an active MFO system. The

oxidation of aldrin to dieldrin, the desulfuration of parathion and malathion to their oxygen analogs, and the 3.03

N-demethylations of mexacarbate by MFO produce meta­ bolites that are more toxic or more potent acetyl­ cholinesterase inhibitors than the parent compound

(Metcalf and March 1949, Perry 1960, Sun and Johnson

1960, Brooks and Harrison 1963, Metcalf 1967, Oonnithan and Casida 1968). The reduction in toxicity of the above parent compounds in the presence of PBO in

C. riparius assays (Table 1) is evidence of an active

MFO enzyme system. The rapid detoxification of allethrin in the house fly is strongly inhibited by PBO (Bridges

1957). An SR of 102.1 for allethrin with PBO is an indication of a similar inhibitory response occurring in C. riparius assays. Additional evidence for the presence of an active MFO system is indicated by the minor differences between LC^q 's of aldrin, parathion, and malathion and the LCso’s of their oxidative meta­ bolites dieldrin, paraoxon, and malaoxon respectively

(Table 1), i.e. there is sufficient MFO activity to activate each of these parent compounds to more toxic forms. An SR of 1.13 for paraoxon+PBO suggests a lack of significant, additional detoxifying oxidations in C. riparius. Marginal SR values for malaoxon+sesamex and malathion+EPN, 2.78 and 0.81 respectively, suggests that additional oxidative transformations and carboxyesterase 1 04

activity may occur in C. riparius.

The low SR values for mexacarbate and aminocarb, a

carbamate which differs from mexacarbate only by lacking

a methyl group in the 5-position, are probably the

result of N-demethylations by MFO enzymes in both com­

pounds, since both compounds are metabolized in a similar

manner (Schlagbauer and Schlagbauer 1972, Fukuto 1973).

Landrin^, a carbamate with a methyl group replacing the

4-dimethylamino group of mexacarbate or aminocarb, was

not synergized or antagonized by PBO. These synergistic

data suggest that oxidative demethylation of phenyl

N-dimethyl amino groups in carbamate insecticides occurs

as an activation reaction in C. riparius.

A suggestion that the SR of carbaryl with PBO as

an inhibitor could be used as a criterion of MFO activity

in Insecta has been proposed (Brattsten and Metcalf

1970). The SR of 1.68 for carbaryl and PBO assayed with

C. riparius would have provided evidence for no or minimal

activity in this species. Caution should be used when

screening for MFO activity indirectly; a variety of

compounds should be used.

A synergistic response in C. tentans assays with PBO

and carbaryl or allethrin, SR 0.8 and SR 2.3 respectively, was not conclusive in providing evidence of an active 105

MFO in another chironomid species (Karnak and Collins

1974). Reasons for the disparity in SR values for

allethrin may reside in techniques or species differences;

one would expect a high SR value in C. tentans too.

In another aquatic insect, Peltodytes sp. adults,

allethrin and mexacarbate were synergized with PBO

(SR 15.0 and SR 3.0 respectively), while propoxur

and carbaryl were not synergized (Federle and Collins

1976). Similar SR values were obtained in carbaryl and propoxur assays with C. riparius and Peltodytes sp., while opposite results were obtained with mexacarbate

(SR 0.32) in riparius assays (Table 1). Therefore,

all aquatic insects are not alike in metabolic disposi­ tion.

A synergistic response could not be conclusively demonstrated when insecticides and PBO were assayed with

A. aegypti larvae (Table 2). Aldrin, parathion, malathion, and mexacarbate were not synergized with PBO

(SR values ranged from 1.14-1.31), while allethrin and carbaryl were only weakly synergized (SR 2.50 and SR

2.22 respectively).

Generally, synergistic ratios for similar PBO interaction studies in A. aegypti reported in the literature were similar to my data. For example, two 106 carbamates and PBO were assayed with the Trinidad-R strain (propoxur SR 1.51 and carbofuran SR 1.51), while a slightly greater response w'as obtained with the pyrethroid Dimethrin (SR 3.45) (Klassen et^ al_. 1965).

The Rutgers strain of A. aegypti (allethrin LC50, 190 ppb) had SR values of 3.80 and 1.06 respectively, for pyrethrins and allethrin with PBO (Granett et^ al.

1951), which were very close to the values of the O.S.U. strain (allethrin LC5Q, 176 ppb; allethrin SR, 2.50).

The use of a second synergist, Sulfoxide, a PBO relative, had low SR values with allethrin (SR 1.11) and pyrethrins (SR 1.47) (Granett e^t a_l. 1951).

Variations in SR ratios and the effectiveness of an insecticide-PBO combination varies with the mosquito species and strain assayed. The organophosphate insecticides, methyl parathion, methyl paraoxon, and chlorpvrifos, each combined with PBO and assayed with a susceptible strain of Culex £. quinquefasciatus has SR values ranging from 1.0-1.2 (Georghiou ejt al. 1975), while a susceptible strain of Culex tarsalis has SR values from 0.024-0.82 for a group of organophosphates, including the previous three insecticides (Apperson and

Georghiou 1975). 107

B. Absorptive Uptake and Insecticide Loss

The amount of insecticide absorbed and metabolized

varied with the insecticide, insect species, size of the

insect, stage of development, insecticide concentration,

exposure period, and presence or absence of a synergist, when C. riparius and A. aegypti were assayed under

identical conditions (Tables 3-4, Figures 3-12).

£. riparius larvae absorbed aldrin and dieldrin at

approximately the same rate on an individual or body weight basis, while less lindane (48% larval basis;

63% weight basis) was absorbed than either of the above insecticides under identical assay conditions (20 ppb insecticide for 2 hrs) (Table 3).

Dieldrin and lindane did not undergo further de­ tectable metabolism by C. riparius larvae and pupae or

A* aegypti larvae with the experimental and analytical methods used.

Both organisms accumulated and metabolized aldrin in proportion to insecticide concentration (Figures 8-9) and exposure time (Figures 3-4). However, C. riparius larvae absorbed aldrin at a faster rate than A. aegypti larvae with increasing insecticide concentration or exposure. C. riparius larvae absorbed 81% more aldrin,

80% more dieldrin, when immersed in 20 ppb insecticide ] 08 for 2 hrs (Tables 3-4). The differences in aldrin and dieldrin accumulation between C. riparius and A. aegypti larvae were less pronounced when corrections were made for weight; 32% and 63% more, respectively, for C. riparius larvae (Tables 3-4). C. riparius larvae immersed in 100 ppb aldrin for 1 hr absorbed 6.5 times more insecticide, and contained 2.9 times more dieldrin per individual than A. aegypti larvae (Figures 3-4).

Similar results were obtained when both organisms were immersed in 10 ppb aldrin for 5 hrs; C. riparius larvae absorbed 4.1 times more insecticide and contained 4.3 times more dieldrin per larva (Figures 8-9).

Aldrin metabolism also varied with experimental conditions. The larvae of both organisms converted approximately 55% of absorbed aldrin to dieldrin when immersed in 20 ppb aldrin for 2 hrs (Tables 3-4). The percent aldrin epoxidized to dieldrin in 1 hr in A. aegypti larvae decreased from 40% at 10 ppb to 29% at _

40 ppb and then remained constant between 40 ppb to 100 ppb, while the percent aldrin epoxidized in 1 hr by C. riparius larvae was approximately 13% for all aldrin concentrations tested (Figure 10). The amount of aldrin epoxidized also increased with time, when both organisms were immersed in 10 ppb aldrin: C^. riparius larvae 109

epoxidized 205 at 1 hr and 725 at 6 hrs, while A. aegypti

larvae epoxidized 475 at 1 hr and 785 at 6 hrs (Figures

8-9) .

Hourly rate analysis of uptake for each experimental period (Figures 5-6) suggests total insecticide/larva decreased more rapidly in midges than mosquitoes, when larvae were immersed in 10 ppb aldrin for 6 hrs (Figures

5-6). The hourly rate of dieldrin converted/larva was relatively constant for mosquitoes, but increased from 1 to 5 hrs and decreased between hrs 5 and 6 in midge larvae.

Although C. riparius larvae absorbed more aldrin and contained more dieldrin/larva than A. aegypti larvae, mosquito larvae were more efficient in the epoxidation of aldrin than midge larvae with constant exposure time (Figure 11) and constant insecticide concentration

(Figure 7). The results were unchanged when corrections were made for organism weight.

The amount of aldrin absorbed and metabolized by midge larvae is dependent upon larval size. The largest larvae accumulated the highest amount of aldrin on an individual basis (Figure 12). According to Derr and

Zabik (1974) a high degree of correlation was found between cuticular surface area and DDE absorption by C^. 110 tentans.. Larger C. riparius larvae accumulated pro- progressively less aldrin on a weight basis as average wet weight increased (Figure 12). Differences in insecti­ cide tolerance have been reported for the following stonefly naiads, the larger forms of each species being more tolerant than small ones: Acroneuria, Paragnetina media (Hitchcock 1965), and Pteronarcys californica

(Saunders and Cope 1968).

Variations in the amount of insecticide absorbed by aquatic insects due to differences in the insecticide used, species, insecticide concentration, and exposure time have been reported for mosquitoes (Korte 1962;

Schmidt and Weidhaas 1958, Schmidt and Weidhaas 1961),

Chironomus tentans (Derr and Zabik 1974), and dragonfly nymphs (Wilkes and Weiss 1971).

The amount of insecticide absorbed increased with increasing insecticide concentration and increasing exposure time for mosquitoes (Schmidt and Weidhaas 1958,

Schmidt and Weidhaas 1961), C. tentans (Derr and Zabik

1972), and dragonfly nymphs (Wilkes and Weiss 1971). I obtained similar results with C. riparius and A. aegypti larvae (Figures 8-9, 3-4). A. aegypti larvae immersed in 10 ppb aldrin for 6 hrs absorbed increasing amounts of insecticide over the entire test period, while C. Ill riparius larvae, under identical conditions, absorbed a maximum amount of insecticide at 5 hrs, the body concen­ tration decreasing between hrs 5 and 6 (Figures 3-4).

Dragonfly nymphs immersed in 13.3 ppb DDT absorbed increasing amounts of insecticide over 9 days, while the

DDE absorbed by tentans immersed in 1.1 ppb insecticide increased for 30 days (Wilkes and Weiss 1971,

Derr and Zabik 1972). In a survey of freshwater inverte­ brates for iji vivo aldrin epoxidation, A. aegypti larvae immersed in 100 ppb aldrin for 2 hrs absorbed 164.9 ng of aldrin/larva and converted 42.4% of it to dieldrin

(Khan _et _al. 1972). A direct comparison can not be made between Khan’s values and my values, since metabolic rate changes with exposure time at a given concentration

(Figure 10). However, if one assumes a constant rate of insecticide accumulation and metabolism, each mosquito larva would have accumulated 47.3 ng of aldrin/larva and

34.9 ng of dieldrin/larva in 1 hr. These values are 4.6- fold greater for aldrin, 17-fold greater for dieldrin, and 15.4% greater for aldrin metabolized/larva than my values. The differences in aldrin absorption and metabolism may be due to variations in insect strain or experimental technique.

The rate of dieldrin depuration for midge larvae was 112

0.5 ng/larva/hr, or 2.6% of the total dieldrin absorbed

(Table 5). No dieldrin was excreted in A. aegypti

larvae (Gerolt 1965).

No conversion products of dieldrin were detected in

midge larvae with the experimental conditions used. A

susceptible strain of 450-500 A. aegypti larvae exposed

to 8.0 ppb dieldrin for 24 hrs metabolized 3% of their

absorbed dieldrin to unidentified hydrophilic material(s)

(Gerolt 1965). This may explain the lack of additional

metabolites in my experiments in which 1,000 larvae were

exposed to low concentrations (20-100 ppb) for short

immersion times (2-6 hrs). Another strain of A. aegypti

larvae exposed to 15 ppb dieldrin for 24 hrs metabolized

371 of their absorbed dieldrin to hydrophilic materials

(Korte et^ A l . 1962) . Gerolt (1965) suggested that the higher percentage of conversion product was probably due to the large number of larvae used, since each larva produces 0.2 ng of hydrophilic product/larva.

The presence of PBO did not affect the total uptake of either aldrin or dieldrin by midge larvae (Table 3); however, the epoxidation of aldrin to dieldrin was prevented by PBO. The MFO enzyme system of C. riparius was completely inhibited by PBO; no detectable dieldrin was found in any iri vivo synergism experiments throughout 113

this study. An important difference between C. riparius

and A. aegypti larvae became evident when identical PBO

interaction experiments were done with mosquito larvae.

PBO reduced but did not prevent aldrin epoxidation in

the mosquito (Table 4). In the presence of PBO,

mosquito larvae converted 40$ of absorbed aldrin to

dieldrin (Series 1, Table 4); even a 24 hr 2 ppm pre­

treatment with PBO was ineffective in stopping a 50$

conversion of aldrin to dieldrin (Series 2, Table 4).

The effect upon the rate of aldrin uptake by PBO was

not consistent in mosquitoes. In one series of

experiments the total insecticide absorbed was decreased, but in a second series of experiments the total insecti­

cide absorbed was unchanged. The ineffectiveness of

PBO when combined with insecticides in A. aegypti larval

assays may help to explain the reported, low synergistic

ratios for A. aegypti (Granett et a_l. 1951, Klassen et^ al.

1965) . The amount of dieldrin absorbed by A. aegypti

larvae was unaffected by PBO (Table 4).

C. riparius and A. aegypti pupae absorbed less aldrin, dieldrin, and lindane than their respective larvae when immersed in 20 ppb insecticide for 2 hrs;

70$, 57$, and 74$ less/organism, or 53$, 26$, and 77$ less/gm of organism for midges, and 89$ less aldrin/ ] 14 organism, or 853 less aldrin/gm of organism for mosquitoes (Tables 3-4). Midge pupae also had an inact­ ive MFO enzyme system; no epoxidized aldrin (dieldrin) was detected. Generally, aldrin epoxidase activity is reduced during the pupal period. Reduction or loss of activity has been reported for house flies (Yu and

Terriere 1974), honey bees (Gilbert and Wilkinson 1974), and for flesh flies and blow flies (Terriere and Yu 1976).

An additional indication of the general sensitivity of C. riparjus larvae to insecticides was evident when the effects of the toxicants upon the test organism were compared for larvae and pupae. Approximately 753 of the midge larvae immersed in aldrin and 1003 of the larvae immersed in dieldrin, lindane, and dieldrin with

PBO were moribund after 2 hrs in 20 ppb insecticide.

C. riparius larvae immersed in aldrin with PBO under the same experimental conditions were unaffected; the toxic symptoms were associated with the presence or accumula­ tion of dieldrin. Toxic symptoms also appeared in midge larvae when they were immersed in 10 ppb aldrin for 3 hrs, or when they were immersed in 80 ppb and

100 ppb aldrin for 1 hr. The decrease in the absorptive uptake of total insecticide and aldrin at 3 hrs, and dieldrin at 5 hrs in midge larvae was probably the result 115

of the cumulative toxic effects of dieldrin (Figure 3).

The onset of toxic symptoms appeared in A. aegypti larvae

at 6 hrs, when immersed in 10 ppb aldrin. C. riparius

pupae and A. aegypti larvae and pupae exhibited no

symptoms of poisoning in all other insecticide uptake

assays.

Differences between insecticide tolerances of the

larvae and pupae of aquatic species have been previously

reported for mosquitoes, pupae being the more tolerant

developmental stage (Mitchener 1953a, Mitchener 1953b,

Thevasagayam 1957) . This corresponds to my observations

made with C. riparius.

C. Assay of Whole Body Homogenates for Aldrin

Epoxidation

An optimum pH of 7.5-8.0 (Figure 13) for aldrin

epoxidase activity in whole body homogenates of £.

riparius larvae is similar to the slightly alkaline pH

values reported for crude homogenates of other insect

species: Sarcophaga bullata Parker and Phormia regina

(Meigen) whole body homogenates minus heads, pH 7.2

(Terriere and Yu 1976); Acheta domesticus (L)

Malpighian tubules, pH 7.4 (Benke and Wilkinson 1971);

Gromphadorina portentosa crop, gastric caeca, Malpighian

tubules, hindgut, and fat. body crud homogenates, pH 7.6 116

and midgut crude homogenate, pH 8.0 (Benke ejt al. 1972);

Prodenia eridania gut homogenate, pH 7.8 (Krieger and

Wilkinson 1969); Musea domestica vicina whole body

homogenate, pH 8.2 (Rey 1967, Brooks and Harrison 1968);

and Limnephilus sp. fat body and gut homogenates, pH 8.2

(Krieger and Lee 1973). Epoxidase activity in C.

riparius homogenates decreased rapidly as pH increased

above the optimum value (Figure 13), which resembles

the pH profiles of epoxidase activity in several other

insects (Krieger and Wilkinson 1969, Benke and Wilkinson

1971, Terriere and Yu 1976) .

An optimum incubation temperature of 30°C (Figure

14) for C. riparius whole body homogenate epoxidase activity is the same as optimum temperatures for house fly abdomens (Tsukamoto and Casida 1967), eridania larval gut microsomes (Krieger and Wilkinson 1969), house cricket Malpighian tubules (Benke and Wilkinson

1971), and for whole blowflies, P. regina, and flesh- flies, Sarcophaga sp. (cited in Wilkinson and Brattsten

1973). Considerable epoxidase activity, 80 to 85% of optimum, was observed at 20°-25°C in C. riparius homogenates. These lower temperatures reflect epoxidase activity at temperatures in which this organism normally lives. Microsomal oxidative activity reported for two aquatic insects also show high enzyme activity at

temperatures below 30°C. The caddisfly larva, Limnephilus sp., has a temperature optimum of 20°-25°C for aldrin epoxidase activity in gut and fat body homogenates

(Krieger and Lee 1973). The oxidative metabolism of propoxur by the mosquito larva Culex £. fatigans increased as the whole body homogenate incubation temperature was lowered from 30°C to 2S°C; the activity profile was not determined for temperatures below 25°C (Shrivastava et_ al. 1971). The highest optimum incubation temperature for aldrin epoxidase activity in insects, 40°C, has been reported for drone honey bee larvae microsomes, Apis mellifera (L) (Gilbert and Wilkinson 1974) .

Very few studies include an activity profile for buffer concentration (ionic strength) when optimum assay conditions are determined. The optimum Tris-HCl concentration for C. riparius larvae was 5.0 X 10 “*M

(Figure 15). Many researchers used published methods for their tissue homogenization and incubation. In _ n general, this includes a 5.0 X 10 M buffer concentration and the addition of KC1 without establishing the optimum ionic strength (Brooks and Harrison 1969, Krieger e_t al.

1970, Krieger and Wilkinson 1971, Benke ejt al. 1972 ,

Krieger and Lee 1973, Reed 1974). The change in aldrin 118 epoxidase activity by different buffer concentrations probably reflects the effect of ionic strength on enzyme activity. House fly microsomal N- and O-demethylases have been shown to be sensitive to changes in the ionic strength of the initial homogenizing media; the maximum activity was reported at ionic strength 0.57 (pH 7.8) and ionic strength 0.71 (pH 6.9) (Hodgson and Plapp

1970, Hansen and Hodgson 1971). An increase in the ionic strength of sodium potassium hydrogen phosphate buffer, pH 7.5, from 5.0 X 10‘2m to 1.0M increased the amount of

N-demethyl propoxur produced by a propoxur resistant strain in Culex p. fatigans larval homogenates; an increase from 1.0M to 1.6M decreased activity (Shriva- stava et^ aJL. 1971).

The amount of dieldrin produced on an individual larval basis in C. rip~arius homogenates increased with increasing incubation time (Figure 16). There was only a slight increase in the amount of dieldrin produced per larva after 30 minutes. Similar biphasic curves were reported for aldrin epoxidation by southern Armyworm gut homogenates (Krieger and Wilkinson 1969) and house cricket Malpighian tubule homogenates (Benke and

Wilkinson 1971). Dieldrin production also increased linearly for 30 minutes in housefly whole body ] 19

homogenates (Ray 1967).

A time rate analysis of dieldrin produced/larva/

minute showed a constant decrease over 75 minutes, when

incubated at 30°C (Figure 16). Such decreases in enzyme

activity may indicate instability of the aldrin

epoxidase enzyme at 30°C, a loss in activity due to

endogenous inhibitors, or a combination of both factors.

Consequently, a 15 minute incubation period was used'

for both whole body homogenates and microsomes. A time

rate analysis for dieldrin produced by housefly micro­

somes showed a rapid decrease over a 40 minute period

(Brooks and Harrison 1969). Crude homogenates and microsomal suspensions of cricket Malpighian tubules

showed a rapid, immediate decrease in aldrin epoxidase

and dihydroisodrin hydrolase activity following prepara­ tion; storage in buffer resulted in the loss of approxi~ mately 30% of the initial activity within 3 hrs (Benke and Wilkinson 1971). A decrease in N- and O-demethylase enzyme activity has been reported for propoxur resistant

Culex fatigans homogenates at 25°C; the rate of enzyme loss was reduced but not prevented by storing homogen­ ates at 0°C (Shrivastava et^ al. 1971) .

The in vitro production of dieldrin increased linearly in proportion to larval concentration (enzyme ] 20

concentration) as the number of_larvae was increased from

1 to 5 larvae/ml (Figure 17). Linear increases in

dieldrin production in proportion to increasing protein

concentration, from approximately 1 to 2.5 mg/incubation

mixture, have been reported for house cricket Malpighian

tubules (Benke and Wilkinson 1971) and Southern Armyworm

gut preparations (Krieger and Wilkinson 1969). Aldrin

epoxidase activity was essentially linear up to five

flesh fly or ten blow fly microsome equivalents, while

further increases resulted in decreases in the rate of

dieldrin production (Terriere and Yu 1976). A similar

decrease in N- and O-demethylase enzyme activity has

also been reported for propoxur resistant Culex

fatigans homogenates, when larval equivalents were

increased from 400 to 800 mg/incubation (Shrivastava

et al . 1971).

The rate of dieldrin production by C. riparius whole

body homogenates increased linearly as aldrin concen­

tration was increased from 0.01 mg to 1.0 mg per

incubation mixture. A 1.0 mg aldrin concentration was

used in reaction mixtures for midge crude homogenate and microsomal studies. Contradictory results have been

reported when incubation mixtures with different volumes

containing the same concentration of aldrin/ml were 121

compared to incubation mixtures with the same 5 ml

volume but different aldrin concentrations/ml (Lewis

et al. 1967). Varying the reaction mixture volume and

keeping aldrin concentration/ml constant had no effect

upon epoxidase activity, while varying the aldrin con­

centration/ml in a 5 ml volume reaction mixture increased

epoxidase activity. A 0.1 mg aldrin concentration per

reaction mixture has generally been used in iji vitro

studies (Krieger and Wilkinson 1969, Brooks and Harrison

1969, Krieger and Wilkinson 1970a, Krieger ejf aJL. 1970,

Benke and Wilkinson 1971, Benke et_ aJL. 1972 , Krieger and

Lee 1973, Reed 1974). A 20 mg aldrin concentration per

incubation mixture was used in housefly microsome reaction mixtures (Rey 1967).

A preliminary estimate of the Michaelis Constant

(Km) for crude C. Tiparius homogenate aldrin epoxidase, derived from a double reciprocal Lineweaver-Burk plot

(Dawes 1972) with substrate (aldrin) concentration data

(Table 8) was Km = 2.0 X 10“^M. This value corresponds to P. eridania, Km = 5.9 X 10“^M (Krieger and

Wilkinson 1969), and the housefly Musca domestica, Km =

2.0-2.5 X 10~^M (Rey 1967). Reports where aldrin epoxidase had a greater affinity for the substrate include: Acheta domesticus Km = 5.8 X 1 0 " (Benke and 122

Wilkinson 1971), H. virescens Km = 6.03 X 10"^M and H. zea Km = 6.66 X 10"^M (Williamson and Schiechter 1970), and Apis mellifera Km = 1.43 X 10-?M (Gilbert and

Wilkinson 1974) .

The incorporation of BSA (1-2% W/V) in reaction mixtures has been used to reduce the effect of endogenous inhibitors and increase epoxidase enzyme activity in flesh fly microsomes (Terriere and Yu 1976), caddisfly gut and fat body microsomes (Krieger and Lee 1973), P. eridania gut microsomes (Krieger and Wilkinson 1970b)» and housefly abdomen microsomes (Tsukamoto and Casida

1967b, Khan et^ al_. 1970). The addition of 1.0% and 0.1%

(W/V) BSA reduced activity in C. riparius homogenates,

10%, 13%, and 19% respectively (Tables 7, 9). Similar reductions in aldrin epoxidase activity have been reported when 20 mg BSA/incubation was used with flesh fly and blow fly microsomes (Terriere and Yu 1976), or when 2% (W/V) BSA was used to study N- and O-demethylase activity with house fly microsomes (Hansen and Hodgson

1971).

The reduction in epoxidase activity by BSA in midge whole body homogenates suggests the absence of BSA- neutralized endogenous inhibitors similar to that reported for P. eridania larval gut preparations (Krieger 123 and Wilkinson 1970a).

The effect upon enzyme activity of chemicals in­ cluded in "standard" mixtures was re-examined after optimum conditions were determined for C. riparius whole body homogenates. Maximum epoxidation activity was obtained with the addition of an electron generating system which included G-6-P, G-6-P dH, and NADP

(Table 9), In general, the deletion of NADPH from microsomal preparations produced substantial reductions in aldrin epoxidation, while the deletion of G-6-P dH produced a less pronounced decrease in enzyme activity. Microsome preparations in which reductions in enzyme activity occurred when NADPH or G-6-P dH were not added include P. eridania gut microsomes (Krieger and Wilkinson 1969) , Acheta domesticus Malpighian tubule microsomes (Benke and Wilkinson 1971), G. portentosa midgut, caeca, and fat body microsomes (Benke al .

1972), and Apis mellifera drone larvae microsomes

(Gilbert and Wilkinson 1974). The deletion of NADPH from caddisfly gut and fat body microsomes (Krieger and

Lee 1973) and housefly microsomes (Rey 1967) resulted in a total loss of enzyme activity, while the deletion of

G-6-P dll had no effect upon aldrin epoxidation. The removal of NADPH from flesh fly and blow fly microsome 124 preparations (Terriere and Yu 1976) and from lepidop- terous larvae preparations (Williamson and Schechter

1970) reduced enzyme activity 84% +; the effect of G-6-P dll upon reaction mixtures was not examined. In another aquatic species, the addition of G-6-P and NADPH increased propoxur metabolism in Culex p. fatigans larval homogenates (Shrivastava ep al. 1971).

The deletion of KC1 from the reaction mixture had no effect upon enzyme activity when optimum preparatory and incubation conditions were used (Table 9). A possible explanation for this has already been discussed in a previous section.

The addition of 0.11 BSA (W/V) resulted in a slight decrease in enzyme activity, while the deletion of

NADH had no effect upon aldrin epoxidation when optimum conditions were used (Table 9).

The addition of FMN or FAD generally inhibits aldrin epoxidase activity, or produces little effect.

The addition of FMN or FAD inhibited aldrin epoxidase activity in midgut, caeca, and fat body microsomes of the cockroach Gromphadorhina protentosa (Benke ep al.

1972) and Malpighian tubule microsomes of the cricket

Acheta domesticus (Benke and Wilkinson 1971). The addition of FMN also decreased enzyme activity in Southern 125

Arm/worm gut microsome homogenates (Krieger and

Wilkinson 1969) and C. riparius crude whole body homo­

genates (Table 10), while the addition of FAD produced

a slight increase in aldrin epoxidase activity, 12% in

eridania (Krieger and Wilkinson 1969) and 10% in C^. riparius crude whole body homogenates (Table 10) . t The addition of 1.0 mg (6.0 X 10-4m ) of PBO to the incubation mixture prevented the epoxidation of aldrin in C. riparius whole body homogenates (Table 10) .

Increases in the inhibition of aldrin epoxidase activity with corresponding increases in PBO molarity have been reported for flesh fly and blow fly microsomes (Terriere * and Yu 1976) and for microsomes of vairous house fly strains (Yu and Terriere 1974a). The addition of sesamex (10"^M) reduced aldrin epoxidase activity 78% and 75%, respectively, in caddisfly gut and fat body microsomes (Krieger and Lee 1973). Reductions in aldrin epoxidase activity have also been reported with increasing molar concentrations of 1,3-Benzodioxole derivatives for housefly and pig liver microsome prepara­ tions (Lewis et_ a_l. 1967) .

Based upon the results of the previous series of experiments, I conclude that the aldrin epoxidase system of C. riparius whole body larval homogenates is a typical 126 microsomal mixed-function oxidase. Essentially, there

is not much difference from other aquatic or terrestrial

insects. To summarize, maximum aldrin epoxidase

activity occurred in larval whole body homogenates when

20 larval equivalents were incubated with substrate (1.0 mg aldrin added in 0.5 ml of ethanol) at 30°C for 15 minutes in 5.0 X 10“^M Tris-HCl, pH 7.5 buffer. Reaction mixture requirements included 2.4 X 10~^M G-6-P, 1.6 units G-6-P dH, and 5.1 X 10“5M NADP added in a final

incubation volume of 5 ml'. The rate of aldrin epoxidase activity decreased as incubation time increased at

30°C. The amount of aldrin epoxidized increased expo­ nentially as substrate (aldrin) concentration was increased from 0.01 mg to 1.0 mg/incubation mixture. A

- s very rough estimate of the Km, 2.0 X 10 M, was similar to two previously reported Km values for aldrin epoxidase systems using insect microsomes. When optimum incubation conditions were used, BSA was not needed to protect aldrin epoxidase activity. The addition of KC1 and NADH had no effect upon dieldrin formation. The addition of the cofactor FMN to reaction mixtures had no effect, while the addition of FAD only slightly increased aldrin epoxidase activity. The addition of 1.0 mg

(6.0 X 10"^M) of the classical MFO inhibitor, PBO, 127

prevented aldrin epoxidation. C. riparius larval homo^

genates have no unique properties, when compared to

other insect species. However, C. riparius homogenates

have a high aldrin epoxidase activity compared to values

reported for other insect species.

D. Assay of Subcellular Fractions for Aldrin Epoxidation

The specific activity of aldrin epoxidase of midge

microsomes was determined using optimal preparatory and

incubation conditions that were ascertained with whole

body homogenate studies. In the initial experiment, the

post-mitochondrial supernatant was centrifuged at

71,000g max to isolate microsomal material. Approximately

29% (dieldrin produced/organism) of the original

epoxidase activity was found in the post-microsomal

supernatant (Table 11). In subsequent experiments,

128,000g max was used and epoxidase activity was no

longer detected in the post-microsomal supernatant

(Table 11). The morphological characteristics and

composition of microsomal pellets have been shown to be

dependent upon the homogenizing medium and centrifugation

speeds used to isolate the material (Cassidy et^ al .

1969). The g-forces I used to isolate microsomal material evidently was a factor in the different 12 8

distributions of biochemical activity in the various

subcellular fractions (Table 11).

In another series of experiments, a 12,000g max

post-mitochondrial supernatant from material prepared in

0.2M K2HPO4 - Na2HPC>4 , pH 7.8, buffer was centrifuged

at 105,000g max at 4°C for 1 hr to isolate microsomal

material. An electron microscopic examination showed

that the microsomal pellet was composed of a homogenous

mixture of fragmented vesicles derived from rough and

smooth endoplasmic reticulum, ribosomes, and a very few mitochondria (J. F. Estenik, unpublished observations).

P_. eridania larval fat body and gut microsomal pellets

prepared by centrifuging a 15,000g supernatant for 15 minutes at 1 0 0 ,000g for 1 hr in a 0 .1M potassium phos­ phate buffer, pH 7.4, haye a similar morphological

composition (Cassidy e_t a_l. 1969) . _

The average specific activity for aldrin epoxidase

in C. riparius larval whole body homogenates was 236 pmoles/mg protein/minute, while the activity for washed microsomes was 1,303.5 pmoles/mg protein/minute,

a 5 fold increase (Table 11). The removal of inactive protein and/or endogenous MFO inhibitors by washing microsomes would explain the increase in specific

activity of washed microsomes compared to unwashed pellet 129 material (Table JL1) . The light pink color of a midge

larval microsomal pellet is probably due to the binding of hemoglobin to microsome material. The binding of hemoglobin, porphyrins, and heme compounds has been reported in mammalian microsome studies (Garfinkel 1958,

Petermann and Pavlovec 1961, Maines and Kappas 197 5).

The presence of certain porphyrins and heme compounds were also found to be effective inhibitors of MFO activity (Maines and Kappas 1975). Binding strength between hemoglobin and ribonucleoprotein decreased as pH increased from 6.5 to 7.5, or as ionic strength increased from 1.0 X 10"2M to 5.0 X 10~2M (Petermann and Pavlovec

1961). Consequently, midge microsomal pellets were resuspended in 1.5 X l O ^ M KC1, pH 7.5, and recentrifuged, a method reported to ensure removal of adventitious hemoglobin (Remmer £t^ aJL. 1966) .

Microsomal epoxidase activity was not detected when microsome suspensions were pretreated for 15 minutes with 1.0 mg of PBO prior to the addition of aldrin to the microsome-PBO mixture (Table 11). The MFO system of

C. riparius larvae is evidently very sensitive to inhibition by PBO. Throughout all experiments, i.e. in vivo immersion, in. vivo absorptive uptake, In vitro whole body homogenate and microsomal incubations, the 130 presence of PBO completely1 inhibited aldrin epoxidase

activity.

A comparison of C. riparius MFO specific activity with other insect species or insect tissue preparations may be misleading. Few comprehensive studies have been made of insect microsomal activity for various species

or tissues. Many studies were general surveys or had

experimental objectives which did not include determining optimum in vitro conditions for maximum enzyme activity.

However, with this in mind, a comparison was made of

specific activity for midge aldrin epoxidase with other

insects (Table 13).

The aldrin epoxidase activity of C. riparius micro­ somes is much higher than most other insect species or insect tissues (Table 13). Higher aldrin epoxidase activity has been reported for adult Mu sea domest ica whole fly preparations (Lewis et a_l. 1967) and abdomen preparations (Khan et_ a^. 1970); for P. eridania whole larvae preparations (Krieger and Wilkinson 1970) and larval gut preparations (Krieger and Wilkinson 1969); susceptible and resistant H. yirescens and H. zea (Reed

1974) .

The following conclusions can be made regarding C_. riparius larval whole body homogenates and microsomes: TABLE 13 Rnte of Aldrin Fipoxitlation by Insect MicTosomcs nnd tlonogenates3 Specific Activity Reference Species Source proles dieldrin/irj pretcin/r.in

£' d . ' 1P o 7) ’in sen unrest lea f\) if I ? 41

•H'ftTJ-Lii region (A) !\’I ? 3 ef nl_. f 1 f>071 Musc.i donestjen fA} I'.'I 300,000c Rey iliUrf) Musca dnnest ica (A) Id 210'- Rrleyer nr.2 Wilkinson f 1000) Prodeni:: eridanln (L) G 2,145 fhnr. ILIiothis con a) AC, MT, FB 2.0-0.1 Antlmcren polvphcrcug f|.) AC, MT, F3 0.P3-0.nr”: Khnt. 'Jl ill- (1'1:0} Mnscn donesticn (A,FI Abd RSl.TOO i inr.son nr.d ?chcc'it''r 11970) H e liothis :e.n (L) -Hd 330-110 I tie 1 i ot ’.i s vi resccns (L) -Hd 1,090-130 Ostrinin nubilnlis (L) -ltd SO Aryvrornen.in vclutin.nnn (1,1 -lid 4 0 Pcctinophom gossvpielIn (I.) -Hd 20 TAHLF. 15 (CO.VTINHF!)) Specific Activity pnoles JielJrin/r..; nrotcir./njp Reference Species Source

Kr it r ot al. lIJTAl Mncrer.nhv:as varianus CL) If I 2c5 ‘!r::-'.?r and Wilkinson f!!'7n) Prodcni ;t eridania CL.PP.P) KI 1,140-40

.,t'i!i,v :ir. ! i 1 i r.sbn 1) j^c'ut:l doi.csticus CM) CR. MG, !!G, M T , PR 24-;c (F) Clt, MG, HC, MT. FB 92*3

Benke ct a_l_. flfi7 2) (i. portent os a CM) CR, CC, MG, HG, MT, FB 215-16 (F) CR, CC, MG, HG, MT, FB SS-15

!\rie*cr anJ Lee (1973) Linncnhiltis sp. (L) G, FB 0.P3-P.P1 Gilbert and Wilkinson Anis -rc 11 i Tern (M,I.) KI 251 Feed riii:4) HeliotLij^ vires cons (LI G llcliothis ten (L) G 4.65 i' Terrier.' a-,j \ii ■Scrconhatja hullata CA,F) KI, -Hd, Th, AbJ 23-11 I’horria recina CA.F) WT, -ltd, Th, Abd 14-7

t-J TABLE 13 (CONTINUED) aAll numbers rounded.

^Abbreviations: (A)=adult, (L)=larvae, (PP)=prepupae, (P)=pupae, (M)=male, (F)=£emnle WI=whole body, -lkl=headless, Abd=abdomen, Th=thorax, AC=alimentary canal, G=gut, GC=gastric caeca, CR=crop, MG=midgut, HG=hindgut, MT=Malpighian tubule, FB=fat body 1. The aldrin epoxidase activity of C. riparius

larval homogenates and the specific aldrin

epoxidase activity of C. riparius larval micro-

some preparations are as high or higher than

many terrestrial insects and most aquatic insects,

2. Microsomal preparations and biochemical

characteristics are generally similar to other

insects, even though differences in details

exist,

3. The. specific activity of microsomes is higher

than whole body homogenates, as expected. Whole

body homogenates are a simple system to prepare;

there are no endogenous inhibitors to contend

with.

Conclusions

1. Chironomus riparius lsrrvae were susceptible to

insecticides in the part per billion (ppb) range.

2. Aedes aegypti laryae tested under identical

conditions were 3 to 200-fold more tolerant than

midges.

3. £. riparius larvae immersed in insecticide-

synergist combinations produced results that

were typical of an insect with an active MFO

system, i.e. aldrin, parathion, malathion, 1 3 5

aminocarb, and mexacarbate were antagonized by

a synergist (SR ratios<0.5); allethrin was

synergized by PBO

4. A. aegypti larvae immersed in insecticide-PBO

combinations produced inconclusive results, i.e.

there was little evidence for a PBO-sensitive

MFO system.

5. Midge pupae absorbed less aldrin, dieldrin, and

lindane than larvae on an individual or on a

weight basis. Pupae did not convert insecticide

to any detectable metabolites.

6 . £. riparius larvae epoxidized aldrin to dieldrin

without additional detectable conversion products

of aldrin or dieldrin. The LCgg’s for dieldrin

with and without PBO were not significantly

different. Aldrin conversion to dieldrin was an

activation reaction in midges based upon

differences in LC50 values of aldrin with and

without PBO.

7. C. riparius larvae absorbed and metabolized more

aldrin than A. aegypti larvae under identical

conditions; the amount absorbed and metabolized

increased in proportion to insecticide concentra­

tion and exposure time. 1 3 6 8 . The conversion efficiency for aldrin epoxidase

* was higher in £. aogypti larvae than C. riparius

larvae, i.e. more dieldrin was produced/amount

of aldrin absorbed in mosquitoes on an indivi­

dual basis or on a weight basis.

9. The amount of aldrin absorbed and metabolized by

C. riparius larvae depended upon organism size;

larger larvae absorbed and metabolized more

insecticide per individual; however, they

accumulated progressively less on a body weight

basis.

10. The rate of dieldrin depuration for C. riparius

larvae during a six hour period of 0.5 ng/larva/

hr, or approximately 2.5% of the total insecti­

cide absorbed.

11. The characteristics of the aldrin epoxidase system

of £. riparius whole body homogenates were

essentially the same as for other aquatic and

terrestrial insects. Although there were

differences in details, the optimum conditions

for rn vitro mixed-function oxidase of C.

riparius larvae were typical of other insects.

12. The aldrin epoxidase activities of larval £.

riparius homogenates and microsomes were higher 1 3 7

than many other insects , terrestrial or

aquatic.

13. The MFO system of C. riparius larvae was very

sensitive to MFO inhibitors. Treatment with

insecticide-PBO combinations resulted in the

total inhibition of 111 vivo and in vitro aldrin

epoxidation.

14. C. riparius whole body homogenates and micro­

somes were simple to prepare; there were no

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