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61 L-

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Quaestiones

entomologicae

VOLUME XI

A periodica! record of entomological investigations,

published at the Department of Entomology,

University of Alberta, Edmonton, Canada.

1975

11

CONTENTS

Editorial 1

Book Review — Rohdendorf, B. B. 1974. The Historical Development of Diptera .... 3

Whitehead — Additions to “Annotated Key to Platynu s' ' (Coleoptera: Carabidae:

Agonini) 6

Rempel - The Evolution of the Insect Head: the Endless Dispute 7

Heming — Antennal Structure and Metamorphosis in Frankliniella fusca (Hinds)

(Thripidae) and Haplothrips verbasci (Osborn) (Phlaeothripidae)

(Thysanoptera) 25

Rosenberg — Fate of Dieldrin in Sediment, Water, Vegetation, and Invertebrates of a

Slough in Central Alberta, Canada 69

Rosenberg — Food Chain Concentration of Chlorinated Hydrocarbon Pesticides in

Invertebrate Communities: A Re-evaluation 97

Book Review — Nachtigall, Werner. 1974. Insects in Flight: A glimpse behind the

scenes in biophysical research Ill

Steiner — Description of the Territorial Behavior of Podalonia valida (Hymenoptera,

Sphecidae) Females in Southeast Arizona, with Remarks on Digger Wasp

Territorial Behavior 113

Griffiths — Studies on Boreal Agromyzidae (Diptera). IX. Phytomyza Miners of

Boraginaceae in North America 129

Ball — Pericaline Lebiini: Notes on Classification, A Synopsis of the New World

Genera, and a Revision of the Genus Phloeoxena Chaudoir (Coleoptera,

Carabidae) 143

Book Review — Hayat, Principles and Techniques of Scanning Electron Microscopy

and Wells, Scanning Electron Microscopy 243

Larson — The Predaceous Water Beetles (Coleoptera: Dytiscidae) of Alberta:

Systematics, Natural History and Distribution 245

Book Notice - Walker, E. M., and P. S. Corbet. 1975. The Odonata of Canada

and Alaska 499

Book Review — Nachtigall, W. 1974. Biological Mechanisms of Attachment 501

Adisoemarto and Wood - The Nearctic Species of Dioctria and Six Related

Genera (Diptera, Asilidae) 505

Lyneborg — The First Record of an Authentic Dialineura Species in North

America (Diptera: Therevidae) 577

Madge — The Type-Species of Bonelli’s Genera of Carabidae (Coleoptera) 579

Evans — Wax Secretions in the Infrared Sensory Pit of Melanophila acuminata

(Coleoptera: Buprestidae) 587

Whitehead and Ball - Classification of the Middle American Genus Cyrtolaus

Bates (Coleoptera: Carabidae: Pterostichini) 591

Editor’s Acknowledgements 621

INDEX

Abacidus, 594

Abax, 579

aberti, Ammophila, 121

Acercaria, 61

Acilius sernisculatus, 84

acraea, Estigmene, 113, 115

Acricotopus, 81

Actenonyx, 147

bembidioides, 147

Aculeata, 126

acuminatus, Carabus, 580

adelgids, 59

Adephaga, 240, 242

Adisoemarto, S., 499, 505-576

aduncus, Drepanocladius, 71

Aeolothripidae, 26, 54, 55, 60, 64, 65

Aeolothrips fasciatus, 60, 62

intermedins, 65

Aeshna interrupta, 83, 89, 90

aethiops, Carabus, 585

affinis, Ammophila, 127

affinis, Dialineura, 577

affinis, Gambusia, 99, 101

affinis, Podalonia, 122

“Africanae spuriae” species group,

Phloeoxena, 179

Agabus, 84

antennatus, 84

agilis, Carabus, 581

agilis, Hanseniella, 19

Agoni, 593

Agonina, 593, 594, 595

agonines, 145, 152, 154

Agonini, 6, 239, 593, 619

Agonochila, 145

Agonum, 579

convexulum, 1

Agromyza, 131

Agromyzidae, 113, 129-142

alaskensis, Callicorixa, 79

alb at a, Dialineura, 577

albicornis, Dioctria, 509, 514

albicornis, Dioctria (Nannodioctria), 514,

515, 516, 519, 521, 553, 559, 560, 563,

565, 566, 567, 569

albicornis, Dioctria (Neodioctria), 514

albipes, Carabus, 579

albius, Dioctria, 537, 539

albius, Dioctria (Eudioctria), 539

iii

albius, Eudioctria, 537, 538, 539, 540, 545,

548, 549, 553, 559, 560, 564, 565, 566,

567, 568, 571, 574

albius f. auri faces, Dioctria ( Eudioctria),

539, 540

albius f. xanthopennis, Dioctria (Eudioctria),

539, 540

albius group, Eudioctria, 537, 539, 541, 547,

548

alder, 523

Alexiopogon, 555

Aleyrodidae, 59, 64

algae, 74, 98, 101, 105, 109

alligator juniper, 1 1 7

Allolobophora, 103

caliginosa, 105

Allotriopus, 595, 596

Alluaud, C., 180, 239,

Alpaeus, 580

angusticollis, 580

castaneus, 580

ferrugineus, 580

gagates, 580

tibialis, 580

alpestris, Myosotis, 134, 136

alpinus, Carabus, 580

Amara, 580

lunicollis, 580

americana, Hackelia, 134

americana, Periplaneta, 1 8

Ammophila, 115, 117, 126, 127

aberti, 121

affinis, 127

hard, 124

pictipennis, 124

procera, 121

tydei, 127

Amphipoda, 89

Anchomenus, 580

convexulus, 6

melanarius, 162

Anchusa, 132

Anchuseae, 130

Anderson, D. T., 60, 65

Anderson, H. W. (see Petrocelli, S. R.), 104,

109

Andrewes, H. E., 579, 582, 585

Andrews, A. K. (see Bridges, W. R.), 69, 70,

85, 86, 87, 92, 93, 98, 107

IV

angulatum, Coenagrion, 82, 83

angusticollis, Carabus, 580, 582, 584

anilis, Dialineum, 577

anisodactylines, 242

Anisoptera, 81, 89, 499

Annelida, 16

anomalus, Ithytolus, 591, 598, 599, 600

anomalus, Pterostichus (Ithytolus), 600, 605

anomalus, Pterostichus ( Ophryogaster), 600

Antarctiina, 595

antennatus, Agabus, 84

anthracinus, Carabus, 585

ants, 121, 125

harvester, 1 1 7

anurans, 234

Apache plume, 116, 117, 118

aphids, 59

Aphilanthops, 126

Aphoebantus, 511

obtectus, 576

Apioceridae, 506

Apocrita, 60

Applegate, R. L. (see Hannon, M. R.), 98,

108

apricarius, Carabus, 580

Aptinus, 580

aquatilis, Carex, 71

Archaeodiptera, 4

Archidiptera, 3, 5

Architipula radiata Rohd., 5

Arctiidae, 113, 115

Arellano, A. R. V. (see Durham, J. W.), 24

argentatus, Dicolonus, 524

Aristus, 582

Arizona cypress, 1 1 7

grape, 1 1 7

sycamore, 1 1 7

walnut, 1 1 7

arizonica, Cupressus, 1 1 7

arizonicus, Vitus, 1 1 7

Armstrong, A. E. (see Frank, R.), 102, 10^

105, 108

Arthropoda, 8, 10, 11, 13, 14, 15, 16, 17,

66, 501, 502

asiatica, Myosotis, 134, 136

Asilidae, 499, 505-576

Asilinae, 506

Asilomorpha, 4, 506, 507

Asilus oelandicus, 5 1 6

aspen, 523

Asperugo, 132

assimilis, Carabus, 580, 584

Astata, 114, 126

occidentalis, 122

Astatinae, 127

aterrimus, Carabus, 583, 584

atherodes, Carex, 71

Atlantic herring, 102

atopodonta, Helisoma, 79

atratus, Prionyx, 121

atricallipus, Carabus, 581

atricornis, Phytomyza, 137

atrocyanea, Podalonia, 122

audeni, Callicorixa, 78, 79, 85

aulicus, Carabus, 580

aurata, Dialineura, 577

australis, Dioctria (Nannodioctria), 505, 514

515, 516, 519, 521, 553, 559

austriacus, Carabus, 579

azteca, Hyallela, 92

Aztecarpalus, 239

azureus, Carabus, 581, 582

azureus. Dinodes, 581

Back, E. A. 518, 524, 525, 551, 554

Bactridothrips brevitubus, 55, 58, 66

Badonnel, A. 61, 63, 65

Bagnalliella yuccae, 28, 61, 62

Bailey, S. F., 39, 65

Baker, E. A. (see Skerrett, E. J.), 76, 95

Ball, G. E., 1-2, 143-242, 499-500, 591-619

Ball, R. L. (see Hamelink, J. L.), 104, 105,

106, 108

balsamifera, Populus, 71

Banks, N., 517, 541, 543, 544, 554

banksi, Dioctria, 544

banksi, Dioctria (Eudioctria), 544

Barry, H. C., 16, 92

Barton, J. R. (see Bradshaw, J. S.), 16, 92

Basilewsky, P., 147, 148, 152, 166, 240,

579, 581, 582, 585

Bates, H. W., 6, 148, 154, 162, 163, 166,

18, 172, 174, 175, 176, 178, 194, 195, 199,

201, 215, 240, 593, 598, 599, 600, 605,

606, 607, 610, 611, 616, 619

baumhaueri, Dioctria, 519

baumhaueri, Dioctria (Dioctria), 513, 517,

519, 553

Bayrock, L. A., 71 , 92

V

beameri, Dioctria ( Eudioctria), 540

beameri, Eudioctria, 537, 546, 547, 548,

553, 572, 575

beckii, Lepidosaphes, 59

Bedel, L., 582, 585

Bedford, J. W. 104, 107

beetles, 102, 103, 165, 194, 208, 210, 238,

239, 240, 241, 586, 593, 594, 617

Beiger, M., 129, 130, 137

bembidioides, Actenonyx, 147

bembidioides, Ochropisus, 165

Bembix, 1 1 7

beringiana, Phytomyza, 129, 130, 131, 135-

136, 140, 142

Bibionomorpha, 4

bicolor, Lelis, 174

Bicyrtes, 1 1 7

Bigot, J. M. F., 528, 554

biguttatus, Laccophilus, 84

Billberg, G. J., 582, 583, 586

bimaculatus, Gryllus, 63, 66, 67

Biomphalaria glabrata, 93

birch, 523

Bischoff, A. I. (see Hunt, E. G.), 70, 94, 98,

108

blackfly, 95, 99

Blackwelder, R. E., 6, 156, 159, 162, 163,

166, 172, 174, 175, 176, 178, 240

Blanchard, C. E., 176, 240

Blaps spinipes, 584

Blephariceridae, 501

Blephericeromorpha, 4

Blethisa, 580

blister beetle, 9

bloodworm, 88

bluebottle, 1 1 1

bluegills, 94, 102, 104

bluegrass, Kentucky, 71

Bodenstein, D., 60, 65

Boelens, R. G. (see Frank, R.), 102, 104, 105

108

Bohart, R. M., 114, 115, 121, 126

Bohartia, 505, 507, 508, 510, 51 1, 512, 513,

523, 529, 530, 531, 532, 533, 534, 535,

553

bromleyi, 529, 530, 531, 533, 534, 535

553, 570

isabella, 505, 530, 531, 532, 533, 534,

535, 553, 559, 560, 561, 564, 565,

566, 567, 568, 570, 574

Bohartia (continued),

martini, 505, 530, 531, 533, 534, 535,

553, 564, 566, 570, 574

mimda, 505, 530, 531, 533, 534, 535, 553

nitor, 505, 530, 532, 533, 534, 535, 553

senecta, 505, 530, 532, 533, 534, 535, 553

tenuis, 505, 530, 533, 534, 535, 553

Bombyliidae, 121, 123, 507, 51 1

Bombyliiis, 1 1 1

Bonelli, F. A., 579-585

bonellii, Carenum, 581

Bonner, F. L., (see El Sayed, E. I.), 102, 108

Booth, G. M., 104, 107

Boothe, P. N., 104, 107

Boraginaceae, 113, 129-142

Borago, 132

borealis, Carabus, 580

Borkhausen, M. B., 582, 586

Borovicka, R. L. (see Terriere, L. C.), 98, 1 10

Bowser, W. E., 71, 92

Brachinus mutilatus, 580

Brachycera, 3, 509

Brachyninae, 241

Bradshaw, J. S., 76, 92

Brady, U. E. (see Wallace, J. B.), 99, 100, 104,

105, 110

Bradycellina, 594

brassicae, Pieris, 65

Braulomorpha, 4

Braun, H. E. (see Frank, R.), 102, 104, 105, 108

Breidenbach, A. W., 70, 75, 92

Breitkreitz, W. E.,(see Kadis, V. W.), 91, 92,

94

brevis, Dioctria, 543,

brevis, Dioctria (Eudioctria), 543

brevis, Eudioctria, 537, 538, 543, 546, 547,

548, 553, 559, 573, 575

brevispina, Cyrtolaus, 591, 598, 599, 600,

602, 604, 606, 608, 609, 612, 613, 615,

616, 617, 618

brevitubus, Bactridothrips, 55, 58, 66

Bridges, W. R., 69, 70, 85, 86, 87, 92, 93,

98, 107

Britton, E. B., 147, 154, 240

Brocksen, R. W. (see Chadwick, G. G.), 104,

105, 106, 107

bromeliads, 145, 183

Bromley, S. W., 515, 540, 551, 554

bromleyi, Bohartia, 529, 530, 531, 533, 534,

535, 553, 554, 570

VI

bromleyi, Dioctria (Bohartia), 530

Broscus, 581

Bruce, W. N., (see Leland, H. V.), 103, 109

Brulle, A., 580, 582

bucephalus, Scarites, 582

Buescher, C. A., 70, 93

Buffa, P., 60, 65

buffalo, 102

bullhead, black, 102

Bullock, T. H., 62, 65

bulrush, great, 71

Buprestidae, 499, 587-589

buprestoides, Carabus, 582

burbot, 102

Burke, J. A. (see Barry, H. C.), 76, 92

Butler, P. A., 104, 107

Butt, F. H., 8, 10, 15, 16, 17, 20, 21

buxi, Psylla, 59, 68

Bycanistes cristatus, 1 54

cabbage butterfly, 13

Caccinia, 132

caddisfly, 99

Calathus, 239, 580, 594, 619

caliginosa, Allolobophora, 105

Callicorixa alaskensis, 79

audeni, 78, 79, 85

Callistinae, 585

Callistus, 580, 581

calydonius, Scarites, 582

Campbell, R. S. (see Johnson, B. T.), 88, 92,

94, 97, 105, 106, 108

Camponotus vagus, 61

Cancer poguras, 100

cane cholla, 1 1 7

Carabidae, 6, 102, 103, 104, 113, 143-242,

199, 579-586, 591-619

Carabinae, 148

Carabus acuminatus, 580

aethiops, 585

agilis, 581

albipes, 579

alpinus, 580

angusticollis, 580, 582, 584

anthracinus, 585

apricarius, 580

assimilis, 580, 584

aterrimus, 583, 584

atricallipus, 581

aulicus, 580

Carabus (continued),

austriacus, 579

azureus, 581 , 582

bombarda, 580

borealis, 580

buprestoides, 582

cephalotes, 581

cinctus, 583

circumscriptus, 583

cisteloides, 580

communis, 580

confluens, 584

coriaceus, 585

croesus, 583

cupreus, 585

cyanocephalus, 583

dimidiatus, 585

dorsalis, 580

elatus, 584

elevatus, 169

ericeti, 580

eurynotus, 580

fasciatopunctatus, 583, 585

fasciolatus ( = Buprestis connexus), 585

ferrugineus, 580

festivus, 581

flavicornis, 582

flavipes, 579

fuliginosus, 579

fulvus, 580

fuscipes, 580

halensis, 582

helipiodes, 584

hellwigii, 580

holosericens, 581

illigeri, 584

impiger, 579

impressus, 579

interruptus, 582

janthinus, 583

jurine, 585

kugelanni, 585

laeivgatus, 584

leucopthalmus, 584

longicornis, 582

lunatus, 581

lutescens, 579

marginatus, 579, 581

melanocephalus, 580

Cambus (continued),

metallicus, 585

multipunctatus, 580

niger, 584, 585

nigricornis, 581

nigrita, 583, 584, 585

oblongopunctatus, 585

ovalis, 579

pallipes, 580

panzeri, 585

parallelepipedus, 579

parumpunctatus, 580

pavidus, 579

paykullii, 584

picipes, 579

pmsinus, 580

quadrimaculatus, 581, 582

rotundatus, 579

scrobiculatus, 580

sexpunctatus, 579

spoliatus, 581

striola, 579

striolatus, 579

subcyaneus, 583

terricola, 583, 584

torridus, 580

truncatellus, 582

vernalis, 585

vestitus, 581

viduus, 579

vivalis, 579

vulgaris, 580

zonatus, 581

Carausius morosus, 1 8

Carcinus maenas, 100

Cardium edule, 1 00

Carenum, 580

bonellii, 581

Carex aquatilis, 71

atherodes, 71

rostrata, 71, 74, 85

carolinensis, Echthodopa, 550,

554

carp, 102

carp suckers, 102

Casey, T. L., 6

castaneum, Tribolium, 154

castaneus, Alpaeus, 580

Catapiesini, 619

vii

Catapiesis, 595

Catascopellus, 143, 152, 153, 155, 169, 171,

239

crassiceps, 170, 171

Catascopi, 147

Catascopidius, 166

Catascopina, 143, 147

Catascopus, 143, 145, 147, 149, 152, 153,

154, 155, 156, 166-168, 169

(sensu latissime), 166-168

(Catascopus), 153

chontalensis, 158, 167, 169

hardwickii, 166

mexicanus, 169

obscuroviridis, 169

validus, 167

caterpillar, 115, 125, 127

catfish, channel, 102, 104, 107

Catostomidae, 523

Catostomus catostomus, 523

catostomus, Catostomus, 523

cattail, 71

caudalis, Ochropisus, 163, 165

Cepaea hortensis, 104, 107

Cephalotes, 580

cephalotes, Carabus, 581

Ceratophyllum demersum, 71, 74, 85

Ceraturgus lobicornis, 528

Cerceris, 1 1 7

cerealium, Limothrips, 55, 58, 63, 65

Cerinthe, 132

Chadwick, G. G., 104, 105, 106, 107

chalybea, Podalonia, 122

championi, Percolaus, 594

Chandler, J. H. (see Sanders, H. O.), 88, 95

Chapman, R. F., 9, 17, 60, 61, 62, 65

Chaudoir, M. de, 147, 148, 156, 159, 161,

166, 172, 174, 175, 176, 178, 179, 180,

200, 201, 208, 209, 215, 240, 579, 583,

584, 586, 600, 605, 619

chaudoiri, Phloeotherates, 158

551, 552, chaudoiri, Stenognathus, 143, 162, 163, 164

Chaudonneret, J., 12, 17, 20, 21

Chelicerata, 8

Chelonodema, 175

Chesters, G., 70, 93

Chilopoda, 19

Chironomidae, 79, 80, 81, 89, 90, 92, 104, 107

Chironomus, 80, 8 1 , 90

vin

Chironomus (continued),

ten tans, 105, 107

Chlaenius, 581

cholla cactus, 1 1 7

chontalensis, Catascopus, 158, 169

Chromatomyia, 131

horticula, 132

Chrysopilus, 511, 576

Chrysopogonini, 507

Church, N. S. (see Rempel, J. G.), 8, 10, 15,

16, 17, 18, 20, 22

cicada killer, 1 27

Cicindela, 619

Cicindelidae, 239, 241, 242

Cicindelinae, 619

cinctus, Carabus, 583

circumscriptus, Carabus, 583

cisteloides, Carabus, 580

civile, Enallagma, 82, 83

Clairville, J. P. de, 583

clam, 101

clypeadon, Laticinctus, 114

Coats, J. R. (see Kapoor, I. P.), 108

Coccina, 60

Coccinea, 66

Coccoidea, 59, 63

cockle, 100

Cocks, J. A., 70, 93

cod, 102

Coenagrion angulatum, 82, 83

resolutum, 82, 83

resolu turn— Enallagma cyathigerum, 83

Coenagrionidae, 83

Cole, F. R., 51 1, 517, 555, 577, 578

Coleoptera, 6, 7, 17, 18, 19, 60, 63, 65, 1 13,

143-242, 245-498, 579-586, 587-589,

591-619

collaris, Coptodera, 215

Collins, H. L., 99. 107

Colpodes, 240, 594, 619

tine tip ennis, 6

Colyrnbetes, 84

sculp tilis, 84

communis, Carabus, 580

communis, Podalonia, 1 22, 1 24

complanata, Glossiphonia, 81

complanatus, Pristonychus, 583

concolor, Ochropisus, 143, 158, 165-166,

167

confluens, Carabus, 584

congener, Lestes, 83

consirnilis, Rhantus, 84

consularis, Platynus, 6

convexulum, Agonum, 6

convexulus, Anchomenus, 6

convexulus, Platynus, 6

Cooke, A. S., 90, 93

Coon, F. B., 70, 94, 98, 108

Cope, O. B., 70, 86, 88, 93, 104, 105, 106, 107

Coptodera, 143, 149, 152, 153, 154, 156, 163

168, 172, 176, 234, 240

collaris, 215

(Coptodera), 153, 176-178

elongata, 173, 177

emarginata, 176

fasciatopunctata, 175

signata, 205, 213

Coptoderides, 147

Coptoderina, 143, 177, 241

Coptoderitae, 147, 152

Coquillett, D. W., 516, 536, 555, 577

Corbet, P. S., 69, 93,499-500

Corbicula manilensis, 101

Corduliidae, 499

coriaceus, Carabus, 585

coriaceous, Phlaeothrips, 60, 62

Corixidae, 78, 79, 81, 88, 90, 91, 99

cornuta, Corydalis, 99, 100

cornu tus, Laemostenus, 583

corsicus, Harpalus, 584

Corydalis cornuta, 99, 100

costata, Phloeoxena (Phloeoxena), 182, 184,

187, 191, 192, 193, 222, 225, 227, 229,

231, 232

costatus, Helluo, 583

costiferum, Sympetrum, 82

cotton, 65

Cottus perplexus, 105

Coulson, J. C., 88, 93, 99, 106, 1 10

crab, 98, 101

marsh, 104

shore, 100

Crabtree, A. N. (see Robinson, J.), 88, 93, 99,

106, no

Craig, D. A., 1 1 1-1 12, 501, 243-244, 621

Crampton, G. C., 508, 509, 51 1, 555

crappie, black, 102

white, 102

IX

cmssiceps, Catascopellus, 170, 171

Cratocerina, 595

crayfish, 89, 98, 99

Creutzer, C., 583, 585

crickets, 103

Cricotopus, 81

cristatus, Bycanistes, 154

Croker, R. A., 85, 86, 87, 93

Crosby, D.,G., 88, 93, 105, 107

Cross, P. M.,(see Hearle, J. W. S.), 28, 66

Crustacea, 8, 13, 19, 100

Cryobius, 594

Csiki, E., 6, 147, 148, 156, 159, 162, 163,

166, 172, 174, 175, 176, 178, 240, 599,

600, 606, 607,610,611,619

Ctenodactylini, 148

Culex pipiens, 88

pipiens quinquefasciatus, 101

Cummins, K. W., 98, 107

cuneata, Rhangia, 109

Cupressus arizonica, 1 1 7

cupreus, Carabus, 585

Cursorius, 582

Curtis, J., 580. 581, 583, 585, 586

Cutten, F. E. A., 4

cutworms, 115, 122

cyanellus, Lepomis, 99

cyaneus, Scarites, 581

cyanocephalus, Carabus, 583

cyathigerum, Enallagma, 82, 83

Cyclops, 11

Cyclorrhapha, 3, 4, 59, 60

Cymindina, 147

Cynoglossum, 132

Cyrtolaina, 591, 595

Cyrtolaus, 499, 591-619

brevispina, 591, 598, 599, 600, 602, 604,

606, 608, 609, 612, 613, 615, 616,

617, 618

(Cyrtolaus), 598, 599, 606, 607, 609,

614, 616

furculifer, 591, 598, 599, 600, 602, 605,

606, 608, 609,610,611, 612,613,615

617, 618

grumufer, 591, 598, 599, 600, 602, 604,

606, 608, 609, 610, 611, 612, 613,

615, 617, 618

(Ithytolus) orizabae, 591, 598, 599, 600,

601, 603, 605, 607, 612, 613, 615, 617

lobipennis, 591, 598, 599, 600, 601, 603,

Cyrtolaus, (continued),

lobipennis, (continued), 606, 607, 610, 613,

615, 616, 617

newtoni, 591, 598, 599, 600, 602, 606, 608,

609, 612, 613, 614, 615, 616, 618

orizabae ( = anomalus), 6 1 6

rieardo, 591, 597, 598, 599, 600, 602, 603,

606, 608, 609. 611, 613, 615, 617, 618

spinieauda, 591, 598, 599, 600, 601, 603,

606, 607, 609, 610, 611, 613, 615, 616,

617

subirideseens, 591, 598, 599, 600, 601, 603,

606, 607, 608, 609. 612, 613, 615, 616,

617, 618

Daborn, G. R., 74, 93

Dacus tryoni, 65

Damalini, 507

Damalus, 507

dama, Searites, 582

Damerothrips, 65

Daphnia, 77, 101, 105

magna, 88, 93, 95, 107, 1 10

Darlington, P. J., Jr., 154, 166, 169, 176,

184, 187, 188, 189, 190, 223, 224, 234,

235, 238, 240, 523, 555, 594, 617, 619, 621

Dasypogoninae, 506, 507, 51 1, 554, 556

Davies, R. G., 26, 55, 58, 60, 65

Davis, B. N. K., 99, 102, 104, 107

Davis, J. R. (see Collins, H. L.), 99

davisi, Plenoeulus, 114

dealata, Phloeoxena ( Phloeoxena), 181, 188,

189, 191, 192, 193, 225, 227, 229, 231,

232, 238

Deans, L R. (see Coulson, J. C.), 88, 93

deeipiens, Harpalus, 582

deeoratella, Seirpus, 79

Degeer, C., 580

Dejean, P. F. M., 166, 169, 176, 178, 205, 213,

240, 241, 580, 581, 582, 583, 584, 586

de la Cruz, A. A. (see Naqvi, S. M.), 99, 101,

109

de Lappe, B. (see Risebrough, R. W.), 102,

110

Demerec, M., 8, 18

demersum, Ceratophylum, 71, 74, 85

Demetrias, 581

dentifer, Pristolomus, 158, 162

dentifer, Stenognathus, 162, 164

denuda, Eudioetria, 537, 539, 542, 543, 545,

548, 549, 553, 572, 575

X

depressus, Scarites, 584

Derbeneva, N. N., 55, 64, 65

Deroceras, 103,

Derr, S. K., 97, 104, 105, 107

Des Gozis, M., 582

Deuterophlebiomorpha, 4

Dialineura, 499, 577-578

af finis, 577

albata, 577

anilis, 571

aurata, 577

gorodkovi, 577, 578

intermedia, 577

lyneborgi, 577

mongolica, 577

nigrofemorata, 577

Diaptomus, 11

Diaspididae, 59

Dicaelus, 581

elongatus, 581

purpura tus, 581

teter, 581

violaceus, 581

Dicolonus, 505, 506, 508, 510, 51 1, 512,

513, 523, 524, 526, 527, 553

argent at us, 524

medium, 505, 524, 525, 526, 527, 553,

561, 567

nigricentrum, 505, 524, 525, 526, 527,

553, 567

pulchrum, 505, 524, 525, 526, 527, 553

simplex, 524, 525, 526, 527, 553, 559,

561, 565, 573, 575

sparsipilosum, 504, 524, 525, 526, 527,

553

Dicrotendipes, 81

digger wasp, 113-127

dimidiatus, Carabus, 585

Dimond, J. B., 69, 70, 93, 103, 107

Dindal, D. L., 104, 107

Dinkelman, M. G. (see Malfait, B. T.), 234,

235, 242

Dinodes, 581

rotundicollis, 581

rufipes, 581

Dioctria, 499, 505-576

albicornis, 509, 514, 553

albius, 537, 539

banksi, 544

Dioctria, (continued),

baumhaueri, 519

(Bohartia) bromleyi, 530

brevis, 543

(Dioctria), 507, 513, 518

baumhaueri, 513, 517, 519, 553

henshawi, 517, 520, 521, 522, 553, 560,

563, 569

pleuralis, 517, 518, 520, 521, 522, 553,

559, 560, 563, 566, 567, 568, 569

pusio, 517, 518, 520, 521, 522, 553,

559, 560, 563, 565, 574

Seminole, 515, 553

vera, 517, 518, 520, 521, 522, 553, 569,

574

wilcoxi, 505, 517, 518, 520, 521, 522,

553, 569, 574

doanei, 540

(Eudioctria) albius, 539

f. aurifacies, 539, 540

f. xanthopennis, 539, 540

banksi, 544

beameri, 540

brevis, 543

doanei, 540

media, 541

monrovia, 542

nitida, 542

f. denuda, 542

propinqua, 540

sackeni, 544

sackeni rivalis, 544

tibialis, 544

flavipes, 517

henshawi, 517

longicornis, 544

longicornis var. tibialis, 544, 545

media, 541

(Metadioctria) rubida, 536

rubida atripes, 536

rubida nigripilosa, 536

(Nannodioctria) albicornis, 514, 515, 516,

519, 521, 553, 559, 560, 563, 565, 566,

567, 569

australis, 505, 514, 515, 516, 519, 521,

553, 559

Seminole, 514, 515, 516, 519, 521, 553,

559, 560, 563, 565, 566

(Neodioctria) albicornis, 514

XI

Dioctria (continued),

nitida, 542

parvula, 538

pleuralis, 517

propinqua, 540

pusio, 518

nibida, 535

rubidus, 536

sackeni, 544

sackeni f. rivalis, 544

(sensu latiore), 530

vera, 518

Dioctriini, 505, 506, 507, 509, 510, 51 1,

512, 513, 523, 536, 553

(sensu stricto), 505, 508, 51 1

Diodontus virginiana, 121

Diplocardia, 103

Diplocheila, 619

Diplopoda, 19

Diptera, 3-5, 59, 60, 95, 107, 1 1 1-1 13, 121,

129-142, 499,501, 505-576, 577-578

disjuncta, Eudioctria, 537, 538, 543, 544,

546, 547, 548, 553

disjunctus, Lestes, 83

dispar, Hy grot us, 84

dispar, Myelaphus, 528

dissimilis, Eudioctria, 509, 538, 541, 542,

545, 548, 549, 553, 571, 574

Ditomus, 582

Dixit, P. N., 8, 18

Dixus, 582

doanei, Dioctria, 540

Dioctria (Eudioctria), 540

Eudioctria, 537, 539, 541, 546, 547, 548,

553, 572, 575

doanei group, (Eudioctria), 537, 540, 547,

548

Doeksen, J., 26, 60, 61, 65

Dolichus, 582

dorsalis, Carabus, 580

Dougherty, J. H. (see Buescher, C. A.), 70, 9

Douglas, C. W. (see Frank, R.), 102, 104,

105, 108

Downe, A. E. R. (see Wallace, R. R.), 69, 95

dracaenae, Parthenothrips, 28

dragonfly, 99, 110

Drepanocladius aduncus, 7 1

Dromius, 582

Drosophila, 8, 18, 65

Drosophila, (continued),

melanogaster, 59, 67

Dryudella, 114

duck weed, 7 1

Duellman, W. E., 234, 238, 241, 617, 618,619

Dufour, L., 582

Duftschmid, C. E., 579, 581, 582, 583

Duggan, R. E.,(see Barry, H. C.), 76, 92

Duponchel, D. A. J., 584, 586

DuPorte, E. M., 10, 15, 17, 20, 21

Durham, J. W., 241

Dustman, E. H., 88, 93, 103, 104, 106, 107

Dyschromus, 595, 596

Dytiscidae, 81, 84, 88, 89, 90, 91, 245-498

Index to Generic Names, 497

Index to Keys, 497

Index to Species Names, 497

Dytiscus, 84

ooligbucki, 84

Eassa, Y. E. E., 60, 65

Eastham, L. E. S., 13, 14, 15, 16, 18, 20, 21

Eastop, V. F. (see Woodward, T. E.), 59, 64,

68

ebenius, Platynus, 6

Eberhardt, L. L., 86, 90, 93

Echinus esculentis, 100

Echium, 132

Echthodopa, 505, 506, 508, 510, 51 1, 512,

513, 523, 548, 549, 550, 551, 552, 553

carolinensis, 550, 551, 552, 553

formosa, 550, 551, 552, 553

pubera, 549, 550, 551, 552, 553, 559, 562,

565, 573, 575

Echthodopini, 505, 506, 507, 508, 509, 510,

51 1, 512, 513, 520, 523, 536, 538, 553

Echthopoda, 549

Echtodopa, 549

Echtopoda, 549

Eddy, C. 0.,28, 65

edule, Cordium, 100

edulis, Mytilis, 100

Edwards, C. A., 70, 88, 93, 98, 99, 103, 104,

105, 106, 108

Edwards, R. W., 69, 86, 93

Egan, H. (see Edwards, R. W.), 69, 86, 93

Eichelberger, J. W., Jr., (see Breidenback, A. W.),

75, 92

Einfeldia pagana, 81

pectoralis, 81

Xll

Elateridae, 102

elatus, Carabus, 584

elevatus, Carabus, 169

elevatus, Scaphinotus, 169

Eller, L. L. (see Kennedy, H. D.), 69, 94

Ellipsoptera, 619

Elliptoleus, 594

Elodea, 101

elodes, Lymnaea, 78, 79, 90

elongata, Coptodera, 173, 177

elongatus, Dicaelus, 581

El Sayed, E. I., 102, 108

emarginata, Coptodera, 176

Emden, F. L, van, 149, 241, 509, 511, 555

emoryi, Quercus, 1 1 7

Enallagma civile, 82, 83

-Coenagrion, 82, 83

cyathigerum, 82, 83

hageni, 82, 83

encelioides, Verbesina, 116, 117

Enceladus, 583

gygas, 583

Endochironomus, 81

endomychids, 176

Engel, E. O. 513, 523, 555

equestris, Lygaeus, 502

Epomis, 583

ericeti, Carabus, 580

Ernst, K. -D., 63, 65

Erotidothrips, 65

Erotylidae, 154, 176

Erpobdella punctata, 81, 89

Erpobdellidae, 81, 82, 89, 90

Erwin, T. L., 593, 594

(see Ball, G. E.), 595, 616, 619, 621

esculentis. Echinus, 100

Estigmene acraea, 113, 115

Euchroina, 595

Eudioctria, 505, 506, 507, 508, 510, 511,

512, 513, 523, 537, 538, 545, 546, 547,

548, 549, 550

Eudioctria (continued),

denuda {conimuQd), 549, 553, 572, 575

disjuncta, 537, 538, 543, 544, 546, 547,

548, 553

dissimilis, 509, 537, 538, 541, 542, 545,

548, 549, 553, 571, 574

doanei, 537, 538, 541, 546, 547, 548, 553,

572, 575

media, 509, 537, 538, 541, 545, 547, 548

571, 574

Monrovia, 509, 537, 538, 541, 545, 548

549, 553, 571, 574

nitida, 537, 539, 542, 545, 548, 549, 553,

572, 575

propinqua, 538, 539, 540, 541, 545, 548,

549, 554, 571, 574

sackeni, 537, 538, 543, 544, 546, 547,

548, 554, 559, 565, 573, 575

tibialis, 537, 538, 543, 544, 546, 547, 548,

554, 564

unica, 537, 538, 543, 545, 548, 549, 553,

554, 556, 566, 572, 575

Eudiptera, 4

Euoplothrips, 28

Eurycoleoids, 143, 153, 154, 171-172

Eurycoleus, 143, 152, 153, 154, 156, 172,

174, 175-176, 242

macularis, 173, 177

Eurydera, 145, 148, 156

lugubrina, 150

eurynotus, Carabus, 580

Evans, H. E., 114, 122, 123, 124, 126, 621

Evans, J. W. (see Woodward, T. E.), 59, 64, 68

Evans, W. G., 499, 587-589, 621

Evarthrus, 596

Exoprosopa, 121

Fabre, J. H. 122, 126

Fabricius, J. C., 169, 241, 579, 580, 581, 582,

583, 584, 585

Fallugia paradoxa, 116, 117

fasciatopunctata, Coptodera, 175

albius, 537, 538, 539, 540, 541, 545, 548, fasciatopunctatus, Carabus, 583, 585

549, 553, 559, 560, 562, 564, 565, 566, fasciatus, Aeolothrips, 60, 62

567, 568, 571, 574 fasciolatus, Carabus ( = Buprestis connexus),

beameri, 537, 538, 540, 546, 547, 548, 585

553 Fattig, P. W., 215, 241

brevis, 537, 538, 543, 546, 547, 548, 553, fedorovi, Taeniothrips, 65

559, 573, 575 Fernald, H. T., 1 1 5, 1 26

denuda, 537, 539, 542, 543, 545, 548, Feronia, 586, 594

Xlll

Feronia, (continued),

lacertosa, 584

navarica, 584

robusta, 584

Ferris, G. F„ 8, 1 1, 15, 18, 20, 21

ferrugineus, Carabus, 580

Ferus, 162

quadricolUs, 143, 162

fervida, Mooreobdella, 81

festivus, Carabus, 581

flavicincta, Thereva, 576

flavicornis, Carabus, 582

flavipes, Carabus, 579

flavipes, Dioctria, 5 1 7

Flickinger, E. L., 98, 99, 108

Foehrenbach, J., 88, 93, 99, 103, 108

Formicidae, 67

formosa, Echthodopa, 550, 551, 552, 554

Foudia sakalava, 155

Fourcroy, A. F., 585

Fox, A. C. (see Hannon, M. R.), 98, 108

Frank, R. 102, 104, 105, 108

Frankliniella fusca, 25-68

tritici, 28, 61, 62, 68

Franklinothrips, 55

Fraxinus velutina, 1 17

Fredeen, F. J. H., 69, 70, 76, 93, 102, 108

Frey, P. J., 69, 93

frontalis, Rhantus, 84

Fuhriman, D. K. (see Bradshaw, J. S.), 76, 92

fuliginosus, Carabus, 579

fulva, Mytilaspis, 59

fulvus, Carabus, 580

fungi, 175

furculifer, Cyrtolaus, 591, 598, 599, 600,

602, 605, 606, 608, 609, 610, 611, 612,

613, 615, 617, 618

fusca, Frankliniella, 25-68

fusca, Helobdella, 81

fuscipes, Carabus, 580

Gabouriaut, D., 62, 63, 67

gagates, Alpaeus, 580

Gambusia affinis, 99, 101

Gammarus, 99, 100

Gaskin, L. J. R, 579, 586

Gastropoda, 77, 78, 79, 89, 90, 91

Gaul, J. A. (see Barry, H. C.), 76, 92

geniculata, Phloeoxena (Oenaphelox), 157,

166, 182, 206, 207, 208, 209, 210, 212,

geniculata, Phloeoxena ( Oenaphelox), (continued),

213, 214, 227, 229, 231, 232, 233, 235

Gerlach, A. R. (see Terriere, L. C.), 98, 1 10

Germar, E. F., 579, 584, 586

gib bus, Scarites, 583

Gillaspy, J. E., 121, 126

Gish, C. D., 88, 93, 103, 108

glabrata, Biomphalaria, 93

Glanodes, 239

Glossiphonia complanata, 8 1

Glossiphoniidae, 81, 82, 89, 90, 104

Glyptotendipes, 80, 81, 90

Gnatzy, W. (see Schmidt, K.), 63, 66, 67

Godsil, P. J., 98, 108

Goeze, J. A. E., 580

goldeye, 102

goldweed, 1 17

gorodkovi, Dialineura, 511 , 578

Gouin, F., 60, 66

Goulet, H. 234, 238, 241, 617, 619, 621

graminis, Haplothrips, 28, 43

graphiptera, Phloeoxena (Phloeoxena), 199,

201

graphiptera var. limbicollis, Phloeoxena (Phloe-

oxena), 143, 199

Graphoderus, 84

occidentalis, 84

perplexus, 84

Grasse, P. P., 61 , 66

grasshopper, 121, 123

grass shrimp, 99

Graves, J. B. (see El Sayed, E. E), 102, 108

Green, R. S., 70, 94

Greichus, Y. A. (see Hannon, M. R.), 98, 108

Griffiths, G. C. D., 3-5, 129-142

grumufer, Cyrtolaus, 591, 598, 599, 600, 602,

604, 606, 608, 609, 610, 611,612, 613,

615, 617, 618

Grundmann, E. (see Basilewsky, P.), 581, 585

Gryllidae, 63, 66, 67

Gryllus bimaculatus, 63, 66, 67

Grzenda, A. R., 69, 94

Guerin, F. E., 583, 586

Gunnerson, G. C. (see Green, R. S.), 70, 94

Gunther, F. A. (see Westlake, W. E.), 70, 71,

96

guppy, 95, 1 10

guttatus, Pericalus, 150

gygas, Enceladus, 583

XIV

Gynaikothrips uzeli, 28

Habu, A., 147, 152, 154, 156, 166, 169,

171, 176, 241

Hackelia, 133, 134

americana, 134

haemorroidalis, Heliothrips, 65

Haga, K., 26, 55, 58, 66

hageni, Enallagma, 82, 83

halensis, Carabiis, 582

Halffter, G., 617, 619

Hall, T. F. (see Wojtalik, T. A.), 88, 96, 100

Hamelink, J. L., 104, 105, 106, 108

Hanks, A. R. (see Petrocelli, S. R.), 104, 109

Hannon, M. R., 98, 108

Hansen, D. J. (see Booth, G. M.), 104, 107

Hansen, W., 147, 153, 241

Hanseniella agilis, 1 9

Hanstrom, B.,10, 18

Haplothrips gmminis, 28, 43

niger, 66

statices, 60, 62

verbasci, 25-68

hardwickii, Catascopus, 166

Harpalinae, 148, 240, 619

Harpalini, 153, 239, 241, 242, 619

Harpalus, 239

corsicus, 584

decipiens, 582

Harrison, R. B. (see Davis, B. N. K.), 99, 102

107

harti, Ammophila, 124

Hartland-Rowe, R. 74, 94

Hartman, C. 114, 122

Hartmann, C. G., 126

Hatfield, C. T., 69, 94

Hayat, M. A., 243, 244

Hearle, J. W. S., 28, 66

Hedgpeth,! W., 11, 18

Heleomyzidae, 5

Heleomyzinae, 5

Helianthus petiolaris, 116,117

Heliothripinae, 64

Heliothrips haemorrhoidalis, 65

Heliotropium, 132

heliopioides, Carabus, 584

Helisoma atopodonta, 79

trivolvis, 78, 79, 90

hellgrammite, 99

Helluo, 583

Helluo (continued),

costatus, 583

Helluonini, 153, 242

hellwigii, Carabus, 580

Helm, J. M. (see Moubry, R. J.), 99, 109

Helobdella fusca, 81

stagnalis, 81, 90

Helodrilus, 103

Hemimetabola, 16

Heming, B. S., 25-68, 501-502

Hemiptera, 68, 77, 99

hemipteroids, 61, 66, 67

Henke, C. F. (see Breidenback, A. W.), 75, 92

Hennig, W., 4, 60, 61, 66, 222, 241, 506, 507, 509

51 1, 555, 593, 619

henshawi, Dioctria, 517, 560

henshawi, Dioctria (Dioctria), 517, 520, 521,

522, 553, 569

Herbst, J. F. W., 583, 584

herculeano, Phloeoxena (Tacana), 143, 157,

180, 182, 183, 184, 185, 202, 224, 227,

229, 231, 232, 233, 235

herculeano, Tacana, 181

Hering, E. M., 130, 137

Hershkovitz, P., 235, 238, 241, 618, 619

Heteroptera, 61, 64, 67

Heterothripidae, 54, 55, 60, 64

Hickey, J. J., 70, 94, 98, 108

Hicks, C. H., 122, 124, 126

Hill, L. O. (see Wojtalik, T. A.), 88, 96

Hippuris vulgaris, 71

hirsuta, Podalonia, 122

Hirudinea, 76, 79, 81, 82, 89, 90, 91

Hirwe, A. S. (see Kapoor, I. P.), 108

Hitchcock, S. W., 69, 94

hoegi, Phloeoxena ( Oenaphelox), 2 1 5

Hogan, J. W. (see Macek, K. J.), 105, 109

Holcoderus, 145

Holdaway, G. G. (see Loan, C.), 55, 66

Holmgren, N., 10, 18

holosericens, Carabus, 581

Holz, D. D. (see Macek, K. J.), 105, 109

Homalomorpha, 595

Homarus vulgaris, 1 00

Homoptera, 59, 61, 63, 64, 66, 67, 68

Homo sapiens, 500

honestus, Pterostichus ( Gastrellarius), 594

honey tail, 502

Hope, F. W., 581, 582, 584, 586

XV

Hoplothrips, 28

Hopkins, D. M., 523, 555

Horn, G. H., 148, 176, 178, 215, 241

hornbills, 1 54

silvery-cheeked, 240

hornwort, 71

Horridge, G. A., 8, 9, 18, 62, 65

horsemint, 1 1 7

hortensis, Cepaea, 104, 107

horticula, Chromatomyia, 132

Howse, P. E., 62, 63, 66

Hughes, G. M. (see Bayrock, L. A.), 71, 92

Hughes, N. F. (see Smart, J.), 8, 19

Hughes, R. A., 88, 94, 102, 105, 108

Hull, F. M., 505, 506, 507, 523, 529, 530,

555

Hundley, J. C. (see Barry, H. C.), 76, 92

Hunt, E. G., 70, 94, 98, 108

Hyallela azteca, 92

Hydropsyche, 99, 100

Hy grot us, 84

dispar, 84

sayi, 84

hylid frogs, 234, 238, 241, 618, 619

Hymenoptera, 60, 67, 113-127

Hynes, H. B. N., 69, 94, 95

Ide, F. P., 69, 94

Illiger, J. C. W., 579, 580, 583

niybius subaeneus, 84

imitatrix, Phloeoxena (Phloeoxena), 180,

188, 189, 190, 191, 192, 193, 225, 227,

229, 231, 232, 234

Imms, A. D., 12, 13, 18, 19, 20, 21, 59, 60,

61, 66

impiger, Carabus, 579

impressus, Carabus, 579

inquinatus, Stenopogon, 576

insculpta, Lelis, 173, 174

Insecta, 8, 14, 18, 19, 66, 68, 242, 619

intacta, Leucorrhinia, 82

intermedia, Dialineura, 577

intermedius, Aeolothrips, 65

internum, Sympetrum, 82

interrupta, Aeshna, 83, 89, 90

interruptus, Carabus, 582

Isaacson, P. A. (see Woodwell, G. M.), 98,

110

isabella, Bohartia, 505, 530, 531, 532, 533,

534, 535, 553, 559, 561, 564, 565, 566,

isabella, Bohartia (continued), 567, 568, 570,

574

Ithytolus, 591, 593, 594, 598, 599, 606, 609,

614, 616, 617

anomalus, 591, 598, 599, 600

lobipennis, 607

Janvier, H., 114, 126

Japygidae, 18

janthinus, Carabus, 583

Jeannel, R., 147, 148, 152, 154, 166, 169,

176, 180, 241, 583, 584, 586

Jedlicka, A., 147, 241

Johnson, B. T., 88, 94, 97, 105, 106, 108

Johnson, C. W., 517, 519, 544, 555

Johnson, L. G. (see Morris, R. L.), 70, 95,

102, 104, 105, 109

Johnson, L. Y. (see Barry,JI. C.), 76, 92

Johnson, W. C. (see Godsil, P. J.), 98, 108

Jonasson, O. A. (see Kadis, V. W.), 91, 92, 94

Jones, B. R., 69, 94

Jones, J. R. E., 108

jozanus, Myelaphus, 528

Juglans major, 1 1 7

Juniperus deppeana, 1 1 7

jurine, Carabus, 585

Kadis, V. W., 91, 92, 94

Kallman, B. J. (see Bridges, W. R.), 70, 85, 86,

87, 93, 98, 107

Kamal, A. (see Khan, M. A. Q.), 103, 109

Kapoor, I. P. (see Metcalf, R. L.), 101, 103,

108, 109

Karl, E. 555

Kawatski, J. A., 88, 94, 104, 106, 108

Keith, J. A., 70, 94, 98, 108, 109

Kenaga, E. E., 85, 88, 94, 98, 99, 103, 105,

106, 109

Kennedy, H. D., 69, 94

Kertesz, C., 555

Kevan, D. K. (see Cutten, F. E. A.), 4

Khalsa, M. S. (see Kapoor, I. P.), 108

Khan, M. A. Q., 103, 104, 109

Kiigemagi, U. (see Terriere, L. C.), 98, 110

King, K. A., (see Flickinger, E. L.), 98, 99, 108

Kirby, W., 166, 241,606,619

Kirk, V. M., 215, 241

Kjearsgaard, A. A. (see Bowser, W. E.), 71, 92

Knauer, G. A. (see Boothe, G. M.), 104, 107

Konrad, J. G. (see Chesters, G.), 70, 93

Korn, S, (see Macek, K. J.), 105, 109

XVI

Koteja, J., 62, 66

Krbber, O., 577

Krombein, K. V., 1 14, 1 15, 121, 122, 126,

127

kugelanni, Carabus, 585

Kurczewski, F. E., 1 14, 121, 124, 125, 127

Labiatae, 129

Labocephalus, 145, 148, 156

longipennis, 150

striatus, 151

Laccophilus, 84

bigut tatus, 84

lacertosa, Feronia, 584

Lachnophorina, 595

Lacordaire, J. R., 147, 156, 161, 166, 172,

176, 241

Laemostenus, 579, 583,

cornu tus, nomen nudum, 583

laevigatus, Carabus, 584

Lamprias, 583,

Lange, A. B., 55, 66

Laphriinae, 506, 556

Laphystiini, 507, 530

Lappula, 132

largemouth bass, 102, 104

Larink, O., 12, 16, 18

Larrinae, 120, 127

Lasiopogon, 555

Latham, J., 582

Larson, D. J., 243, 245-498

laticinctus, Clypeadon, 114

latifolia, Typha, 71

Latreille, P. A., 516, 555, 579, 581, 582,

583, 584, 585, 586

Lauer, G. J. (see Grzenda, A. R.), 69, 94

Learner, M. A. (see Edwards, R. W.), 69, 86,

93

Lebia, 154, 175, 221

Lebiidae, 147, 242

Lebiina {sensu Habu), 169

Lebiinae, 147, 169, 241

Lebiini, 145, 147, 149, 152, 153, 169, 240,

619

Lebistes reticulatus, 95, 110

LeConte, J. L. 215, 241

Lee, G. F. (see Hughes, R. A.), 88, 94, 102,

105, 108

leeches, 89

Leland, H. V., 103, 109

Lelis, 143, 152, 153, 154, 156, 172, 174-175,

177

bicolor, 174

insculpta, 173, 174

Lemna minor, 1 1

triscula, 71, 74, 85

Leng, C. W., 176, 178, 215, 241

Lepidoptera, 60, 65

Lepidosaphes beckii, 59

Lepisma, 12

saccharina, 18

Lepomis cyanellus, 99

Leptogastrinae, 506, 554

Lestes congener, 83

disjunctus, 83

Lestidae, 83

Leucophaea maderae, 67

leucopthalmus, Carabus, 584

Leucorrhinia Intacta, 82

Lewis, E. (see Gaskin, L. J. P.), 579, 586

Lewis, T. R., 26, 55, 63, 64, 66

Libellulidae, 81, 82, 83, 89, 90, 104, 499

Libelluloidea, 499

Lichnasthenini, 155, 169, 242

Lichnasthenus, 169

Lichtenberg, J. J. (see Green, R.S. ), 70, 75,

92, 94

Lieftinck, M. A., 149,241

Ligura, 121

Umax, 103

limbicollis, Phloeoxena ( Phloeoxena), 181, 194,

195, 196, 198, 199, 202, 203, 204, 224, 226,

227, 229, 231, 232

Limnephilus rhombicus, 99, 100

Limothrips cerealium, 55, 58, 63, 65

limpet, 100

Lin, N. 114, 127

Lindroth, C. H., 147, 241, 523, 555, 580, 586,

594, 619

lineata, Stennoglossa, 172, 173, 174

Linnaeus, C., 516, 555, 579, 580, 581, 582,

583, 584, 585

Linne, C., von, 577

Lioptera, 176

Liothrips, 54

oleae, 55, 58, 63, 61

Lithospermeae, 130

Lithospermum, 132

Liris nigra, 1 20, 1 27

XVll

lithospermi, Phytomyza, 129, 130, 131, 132

liverwort, 71

Livingstone, E. M. (see Eddy, C. O.), 28, 65

Loan, C., 55, 66

lobicomis, Ceraturgus, 528

lobicornis, Myelaphus, 528, 529, 553, 559,

561, 565, 573, 575

lobipennis, Cyrtolaus, 591, 598, 599, 600,

601, 603, 606, 607, 610, 613, 615, 616,

617

lobipennis, Ithytolus, 607

lobipennis, Pterostichus (Ithytolus), 607

Lobodontus, 145

lobster, 100

Loding, H. P., 215, 241

Loew, H., 524, 525, 549, 551, 555

longicornis, Cambus, 582

longicornis, Dioctria, 544

var. tibialis, 544, 545

longipennis, Labocephalus, 1 50

Loricerini, 619

Loveridge, E. L. (see Bradshaw, J. S.), 76, 92

Lovett, A. L. (see Cole, E. R.), 555

Lu, P. -Y. (see Kapoor, I. P.), 108

lucida, Oculobdella, 81

Luckmann, W. H. (see Moye, W. C.), 69, 95

Loxandms, 594

luctuosa, Podalonia, 122, 124, 126

lugubrina, Eurydem, 150

Lumbricus, 103,

terrestris, 105

lunatus, Cambus, 581

lundbecki, Trichophthalma, 576

lunicollis, Amam, 580

lutescens, Cambus, 579

Lycopsis, 132

Lygaeus equestris, 502

Lymnaea, 88, 89

elodes, 78, 79, 90

stagnalis, 77, 78, 85, 90, 104

Lyneborg, E., 499, 577-578

lyneborgi, Dialineum, 577

Lyperopherus, 594

Lytta viridana, 7, 9, 15, 16, 17, 18, 20

Macek, K. J., 104, 105, 109

MacGinitie, H. D., 523, 555

Mackay, D., 70, 94

Mackerras, I. M., 61, 66, 51 1, 555

Macromiidae, 499

Macquart, D., 556

macrozooplankton, 99, 100

macularis, Eury coleus, 173, 177

maculicollis, Phloeoxena ( Oenaphelox), 215

mademe, Leucophaea, 67

Madge, R. B., 499, 579-586

rnaenas, Carcinus, 100

magna, Daphnia, 88, 93, 95, 107, 1 10

Maindron, M., 156, 159, 162, 242

Maitra, N. (see Khan, M. A. Q.), 103, 104, 109

major, Juglans, 1 1 7

Makel, M., 60, 66

Malacostraca, 12

Maldonaldo-Koerdell, M., 234, 242

Malfait, B. T., 234, 235, 242

Mallis, A., 9, 11, 18

Malzacher, P., 8, 16, 18

manilensis, Corbicula, 101

Manton, S. M. (see Tiegs, O. W.), 8, 10, 11,

13, 18, 19, 20, 21

(see Imms, A. D.), 1 2

Marchal, P., 122, 127

mare’s tail, 71

marginatus, Carabus, 579, 581

marginatus, Scarites, 584

Maris, P. J.,(see Edwards, R. W.), 69, 86, 93

Markin, G. P. (see Collins, H. E.), 99, 107

Marsh, P. M. (see Bohart, R. M.), 1 14, 121, 126

Martin, D. H., 507, 514, 515, 517, 518, 519,

530, 535, 536, 537, 538, 539, 540, 541,

542, 543, 544, 556

Martin, P. S., 208, 242

martini, Bohartia, 505, 530, 531, 533, 534,

535, 553, 564, 566, 570, 574

Mascarenhia, 180

Maslin, T. P., 222, 242

Masson, C., 62, 63, 67

Mateu, J., 147, 155, 169, 242

Matsuda, R., 8, 9, 12, 16, 18, 19, 20, 21, 60,

61, 67

mealy bugs, 60

Mecoptera, 60

media, Dioctria, 541

media, Dioctria, (Eudioctria), 541

media, Eudioctria, 509, 538, 545, 547, 548,

553, 571, 574

group, 537, 541, 547

medium, Dicolonus, 505, 524, 525, 526, 527,

553, 561, 567

XVlll

Meeks, R. L., 85, 86, 87, 88, 89, 90, 93, 94,

95, 97, 109

megalops, Phloeoxena ( Phloeoxena), 181,

193, 194, 195, 198, 199, 223, 224, 226,

227, 229, 231, 232, 233

chiriquina, 181, 195, 198, 202, 203, 204,

205

complex, 228, 233

erwinorum, 143, 181, 194, 197, 198, 202,

203, 204

megalops, 181, 194, 196, 198, 202, 203,

204

Megaloptera, 60

Meigen, J. W. 516, 517, 519, 544, 556

melanarius, Anchomenus, 162

Melander, A. L., 540, 544, 556

Melanius, 583

melanocephalus, Carabus, 580

melanogaster. Drosophila, 59, 67

Melanophila acuminata, 587-589

melas, Myelaphus, 528, 529, 553

Melin, D., 509, 556

Melis, A., 26, 55, 58, 60, 67

Meloidae, 7, 17, 18

Menke, A. S. (see Bohart, R. M.), 115, 126

Menzie, C. M., 70, 95

Merothripidae, 54, 55, 60, 64, 65

Merothrips morgani, 54

Mertensia, 133, 135, 136

paniculata, 131, 134, 135, 136, 141, 142

mertensiae, Phytomyza, 129, 130, 131, 132,

134, 135, 136, 139

Me table tus, 582

Metadioctria, 505, 506, 507, 508, 510, 51 1,

512, 513, 523, 535, 538, 553

resplendens, 535, 538

mbida, 534, 536, 559, 560, 563, 565, 566

567, 568, 570, 574

metallicus, Carabus, 585

Metaphragma planiceps, 576

Metcalf, R. L., 101, 103, 108, 109

mexicanus, Catascopus, 169

Mexisphodrus, 6, 593, 619

Mickoleit, E., 26, 58, 60, 61, 62, 64, 67

mice, 108, 619

Microvelindopsis, 1 69

microzooplankton, 100

midge, 107

minax, Uca, 101

Minkiewicz, R., 114, 127

minor, Lemna, 71

Minuthodes, 145

Miscelus, 145

Mitterpacher, L., von Mitterburg (see Filler, M.),

579

Moctherus, 145

molitor, Tenebrio, 19

Mollusca, 104, 109, 502

Molops, 584

Monarda pectinata, 1 17

mongolica, Dialineura, 577

monrovia, Dioctria ( Eudioctria), 542

monrovia, Eudioctria, 509, 538, 542, 545,

548, 549, 553, 571, 574

group, 537, 541, 547, 548

montana, Phloeoxena ( Phloeoxena), 152, 181,

187, 188, 191, 192, 193, 222, 225, 227,

229, 231, 232, 238

Moore, N. W., 69, 70, 95, 98, 103, 104, 106,

109

Mooreobdella fervida, 8 1

morgani, Merothrips, 54

Moriarty, F., 85, 95, 98, 99, 103, 104, 105,

106, 109

Mormolyce, 143, 147, 148, 149, 154, 241

phy nodes, 148, 151

Mormolycini, 148

morosus, Carausius, 18

Morris, R. L., 70, 95, 102, 104, 105, 109

mosquito, 101

moss, 71

Moubry, R. J., 99, 100, 109

Mougeotia, 71

Mound, L. A., 65

mountain ash, 1 1 7

, Moye, W. C., 69. 95

Moyle, J. B. (see Jones, B. R.), 69, 94

Muesebeck, C. F. W., 115, 127

Muirhead-Thomson, R. C., 70, 85, 88, 95

Muller, G., 580

multipunctatus, Carabus, 580

munda, Bohartia, 505, 530, 532, 533, 534,

535, 553,

Murphy, P. G., 105, 109

Murray, F. T., 60, 67

Murray, W. D., 115, 127

Musidoromorpha, 4

mussel, 100, 104, 107

XIX

mutilatus, Brachinus, 580

Myadina, 595

Mydidae, 506, 51 1

Myelaphus, 505, 506, 508, 510, 51 1, 512,

513, 523, 527, 528, 529, 553

dispar, 528

jozanus, 528

lobicornis, 528, 529, 553, 559, 561, 565,

573, 575

melas, 528, 529, 553

rufus, 528

Myiomorpha, 4

myosotica, Phytomyza, 129, 130, 131, 133

136

Myosotidae, 130

Myosotis, 131, 132, 133

alpestris, 134, 136

asiatica, 134, 136

sylvatica, 131

Myrdal, G. R. (see Moubry, R. J.), 99, 100,

109

Myriapoda, 8, 10, 14, 66

Mytilaspis fulva, 59

Mytilis edulis, 100

Nachtigall, W., 11 1-1 12, 501-502

Nannodioctria, 505, 506, 507, 508, 510, 51 1

513, 514, 515, 516, 520, 521, 553, 556

Naqvi, S. M., 99, 101, 109

navarica, Feronica, 584

Necrophorus, 65

vespilloides, 63

Negre, J. (see Ball, G. E.), 146, 239, 592,

593, 594, 616, 619

Nematocera, 3, 507, 51 1

Nemomydas, 5 1 1

pantherinus, 576

Neodioctria, 507, 514, 556

Neuroptera, 60

Newcomer, E. L., 122, 124, 127

Newsom, L. D., 28, 67, 95

newtoni, Cyrtolaus, 591, 598, 599, 600, 602,

606, 608, 609, 612, 613, 614, 615, 616,

618

Nicholson, H. P. (see Grzenda, A. R.), 69, 88,

nidicola, Paulianites, 1 54

niger, Haplothrips, 66

nigra, Liris, 120, 127

nigricollis, Phloeoxena ( Phloeoxena), 143, 181,

194, 195, 196, 198, 199, 200, 202, 203, 204,

224, 226, 227, 229, 231, 232

nigricornis, Carabus, 581

nigrita, Carabus, 583, 584, 585

nigrofemorata, Dialineura, 577

sensu Lyneborg, 577

nigrohirta, Podalonia, 122

nigropiceus, Phloeotherates, 162

nitida, Dioctria, 542

nitida, Dioctria (Eudioctria), 542

f. denuda, 542

group, 537, 542, 547, 548

nitida, Eudioctria, 537, 539, 545, 548, 549,

553, 572, 575

nitor, Bohartia, 505, 530, 532, 533, 534, 535

553

Noctuidae, 1 1 5

Nodularia, 71

Nonea, 132

Noonan, G. R., 153, 234, 242, 621

Notogonia pompiliformis, 127

Notonecta, 88, 89

undulata, 78, 80

Notonectidae, 78, 81, 88, 89, 90, 91, 99

Nowakowski, J. T., 129, 130, 131, 137

nowakowskiana, Phytomyza, 129, 130, 132

Nycteis, 145, 153, 176

Nycteribiomorpha, 4

Nymphomyiidae, 4

Nystrom, R. F. (see Kapoor, I. P.), 108

oak, 117, 210

obliquus, Scenedesmus, 95, 109

oblongopunctatus, Carabus, 585

O’Brien, R. D., 95

obscura, Phytomyza, 137

obscuroviridis, Catascopus, 169

obtectus, Aphoebantus, 576

occidentalis, Astata, 122

occidentalis, Graphoderus, 84

Ochropisus, 143, 152, 153, 155, 156, 161,

94, 95 163-164, 166, 234

niger, Carabus, 584, 585 bembidiodes, 165

nigricentrum, Dicolonus, 505, 524, 525, 526, caudalis, 163, 165

527, 553, 567 concolor, 143, 158, 165-166, 167

nidicola, Oecornis, 1 54, 240 Oculobdella lucida, 8 1

XX

Odacanthinae, 241

Odacanthini, 148

Odonata, 81, 83, 89, 90, 92, 499-500

Oecornis, 169

nidicola, 1 54, 240

Oedogonium cardiacum, 1 0 1

oelandicus, Asilus, 516

Oenaphelox, 153, 155, 156, 179, 180, 205,

206, 208, 209, 212, 213, 226, 228, 235

236

geniculata, 182

pluto, 182, 223, 237

signata, 180, 182, 223, 237

undata, 180

oleae, Liothrips, 55, 58, 63, 67

Olivier, A. G., 582, 585

Omaseus, 583

Omphalodes, 132

O’Neill, K. (see Mound, L. A.), 65

Onychophora, 8, 13, 18

Ony pterygia, 595

Oodes, 584

ooligbucki, Dytiscus, 84

Ophryogaster, 596

Opuntia spinosior, 1 1 7

Oreodicastes, 143, 152, 153, 155, 156, 159,

234

subcyaneus, 158, 160

Orestes, Rhagio, 576

orizabae, Cyrtolaus (Ithytolus), 591, 598,

599, 600, 601, 603, 605, 606, 612, 613,

615, 616, 617

orizabae, Pterostichus (Ithytolus), 600, 605

ornatus, Pericalus, 1 50

Orthocladiinae, 81

Orthoptera, 63

Oscillatoria, 71

Osten Sacken, C. R., Baron von, 518, 528,

556

ostracods, 108

ovalis, Carabus, 579

ovalis, Phytomyza, 130, 131, 132, 133-134,

135, 136, 138, 142

Oxybelus, 114, 121

sericeum, 126

Oxyglossus, 156, 159

subcyaneus, 156

oysters, 104

Pachy teles, 194

pagana, Einfeldia, 81

Palaeodipteron walkeri, 4

pallipes, Carabus, 580

paniculata, Mertensia, 131, 134, 135, 136,

141, 142

pantherinus, Nemomydas, 576

Panzer, G. W. F., 579, 580, 581, 582, 583, 584,

585, 586

panzeri, Carabus, 585

Papavero, N., 506, 556

Parachironomus, 81

paradoxa, Fallugia, 116, 117

parallelepipedus, Carabus, 579

Paraneoptera, 6 1

parasitoids, 121

Par atany tarsus, 81

Paratrichocladius, 8 1

parkeri, Prionyx, 1 24

Parthenothrips dracaenae, 28

parumpunctatus, Carabus, 580

parvula, Dioctria, 538

Pasimachus, 584

Patella vulgata, 100

Paulianites, 169

nidicola, 154

Pauropoda, 14, 19

Pauropus silvaticus, 19

pavidus, Carabus, 579

Paykull, G., 579-580, 581, 584, 585

paykulli, Carabus, 584

Peck, J. V. (see Durham, J. W.), 241

pectinata, Monarda, 1 1 7

pectinatus, Potamogeton, 71

pectoralis, Einfeldia, 81

Pelecypoda, 89

Pelmatellus, 238, 241, 619

Pelor, 584

Pennak, R. W., 89, 95

Pentagonicini, 240

Pephrica, 169

Percolaus, 594, 595, 596

championi, 594

Percus, 584

Pericali, 147

Pericalides, 147

Pericalina, 1 13, 143, 147, 149, 153, 154, 155,

156, 157, 158, 159, 161, 168, 169, 170, 171,

173, 174, 175, 176, 179, 180, 182, 227, 234,

238

XXI

Pericalitae, 147

Pericaloids, 143, 153, 154, 176

Pericalus, 145, 153, 154, 176

guttatus, 150

ornatus, 150

Perigonini, 240, 593

Peripatopsis, 18

Periplaneta americana, 1 8

Peripristus, 145

perplexus, Cottus, 105

perplexus, Graphoderus, 84

Pesson, P., 59, 60, 64, 67

Peterle, T. J., 86, 90, 93, 95

Peters, T. W. (see Bowser, W. E.), 71, 92

Peterson, A., 60, 63, 64, 67, 508, 556

Peterson, J. L. (see Bradshaw, J. S.), 76, 92

petiolaris, Helianthus, 116, 117, 139

petiolaris, Phytomyza, 129, 130, 131, 132,

134, 135, 136, 141

Petrocelli, S. R., 104, 109

Pfefferkorn, G. E., 589

Phasmida, 18

Phellini, 507

Philophloeus, 145

Phlaeothripidae, 25, 26, 28, 45, 54, 55, 58, 60,

61, 63, 64, 66

Phlaeothrips coriaceous, 60, 62

Phloeotherates, 143, 153, 155, 162, 163

chaudoiri, 158

nigropiceus, 162

Phloeoxena, 113, 143-242,619

''Africanae spuriae'' species group, 179

( Oenaphelox)

geniculata, 157, 166, 206, 207, 208, 209,

210, 212, 213, 214, 227, 229, 231,

232, 233, 235

hoegi, 215

maculicollis, 215

pluto, 206, 207, 210, 212, 213, 227, 229,

231, 232, 233, 235

signata, 206, 208, 209, 210, 212, 213,

214, 215, 216, 217, 218, 219, 220,

226, 227, 229, 231, 232, 233, 234,

235

var. collaris, 2 1 5

hoegi, 221

nigripennis, 2 1 5

signata, 221

undata, 157, 180, 206, 207, 208, 209,

Phloeoxena (continued),

undata (continued), 210, 211, 212, 213,

227, 229, 231, 232, 233, 235

undulata, 208

(Phloeoxena), 153, 180, 184, 235, 236, 237

costata, 182, 184, 187, 191, 192, 193, 222,

225, 227, 229, 231, 232

dealata, 181, 187, 188, 189, 191, 192, 193,

225, 227, 229, 231, 232, 238

graphiptera, 199, 201

xsLT. limbicollis, 143, 199

imitatrix, 180, 188, 189, 190, 191, 192,

193,225,227,229, 231,232,234

limbicollis, 181, 194, 195, 196, 198, 199,

202, 203, 204, 224, 226, 227, 229, 231,

232

megalops, 181, 193, 195, 198, 199, 223,

224, 226, 227, 228, 229, 231, 232, 233

chiriquina, 181, 195, 198, 202, 203,

204, 205

erwinorum, 143, 181, 194, 197, 198,

202, 203, 204

megalops, 181, 194, 196, 198, 202,

203, 204

montana, 152, 181, 187, 188, 191, 192,

193, 222, 225, 227, 229, 231, 232, 238

nigricollis, 143, 181, 194, 195, 196, 198,

199, 200, 202, 203, 204, 224, 226, 227,

229, 231, 232

picta, 143, 178, 184, 186, 190, 195, 198,

199, 201, 202, 203, 204, 223, 224, 226,

228, 229, 233, 234, 237, 238, 239

apicalis, 143, 181, 186, 200, 201, 202,

203, 204, 205, 226, 227, 229, 231,

232

batesi, 143, 181, 186, 195, 199, 200,

201, 202, 203, 204, 205, 226, 227,

229, 231, 232, 233

franiae, 143, 181, 195, 198, 199, 200,

201, 202, 203, 204, 205, 226, 227,

229, 231, 232

picta, 181, 186, 195, 199, 200, 201,

202, 203, 204, 205, 226, 227, 229,

231, 232

unicolor, 157, 181, 186, 200, 202, 203,

204, 205, 224, 226, 227, 229, 231,

232, 233

plagiata, 180, 188, 189, 191, 192, 193, 225,

227, 229, 231, 232

XXll

Phloeoxena (continued),

( Phloeoxena)

portoricensis, 182, 188, 189, 190, 191

192, 193, 225, 227, 229, 231, 232,

238,

picta, Phloeoxena (Phloeoxena) (continued),

picta, 181, 186, 195, 199, 200, 201, 202,

203, 204, 205, 226, 227, 229, 231, 232

unicolor, 157, 181, 186, 200, 202, 203, 204,

205, 224, 226, 227, 229, 231, 232, 233

schwarzi, 143, 182, 184, 186, 187, 188, pictipennis, Ammophila, 124

189, 190, 191, 192, 193, 222, 223, Pieris brassicae, 65

225, 227, 228, 229, 231, 232, 233,

234, 237, 238

''species verae"\ 179

unicolor, 143, 200

(Tacana) herculeano, 143, 157, 180, 182,

183, 184, 185, 186, 202, 224, 227, 229,

231,-232, 233, 235

Phoromorpha, 4

Phthiraptera, 61

phyllodes, Mormolyce, 148, 151

Physa, 101

physapus, Thrips, 60, 62, 67

Phytomyza, 113, 129-142

atricornis, 137

rapae, 13, 16, 18

pike, northern, 102

Piller, M., 579

Pillmore, R. E. (see Rudd, R. L.), 70, 98

pipiens, Culex, 88

quinquefasciatus, 101

plagiata, Phloeoxena (Phloeoxena), 180, 188,

189, 191, 192, 193, 225, 227, 229, 231, 232

planiceps, Metaphragma, 576

plankton, 98, 102

Planorbidae, 88, 89

Platynella, 595, 596

Platynus, 6, 154, 584, 593, 594, 595, 619

consularis, 6

beringiana, 129, 130, 131, 132, 135-136, 140 convexulus, 6

lithospermi, 129, 130, 131, 132

mertensiae, 129, 130, 131, 132, 134, 135,

136, 139

myosotica, 129, 130, 131, 133, 136

nowakowskiana, 129, 130, 132

obscura, 129, 137

ovalis, 130, 131, 132, 133-134, 135, 136,

138, 141

petiolaris, 129, 130, 131, 132, 134-135,

136, 139, 141

pulmonariae, 129, 130, 131, 132

symphyti, 129, 130, 131, 132, 135, 137

syngenesiae, 134

Picard, F. 122, 127

piceus, Scarites, 584

picipes, Carabus, 579

picta, Phloeoxena ( Phloeoxena), 143, 178,

184, 198, 199, 202, 203, 204, 223,224,

226, 233, 234, 237, 238, 239

apicalis,\^3, 181, 186, 200, 201, 202, 203,

204, 205, 226, 227, 229, 231, 232

batesi, 143, 181, 195, 199, 200, 202, 203,

204, 205, 226, 227, 229, 231, 232, 233

franiae, 143, 181, 195, 198, 199, 200, 201,

202, 203, 204, 205, 226, 227, 229, 231,

232

group, 233, 234, 235

ebenius, 6

porrectus, 6

teter, 6

tinctipennis, 6

tlamayensis, 7

wrightii, 117

Platysma, 584

Plenoculus davisi, 114, 127

pleuralis, Dioctria, 517, 553

(Dioctria), 517, 518, 520, 521, 522, 559,

560, 563, 566, 567, 568, 569

pluto group, Phloeoxena, 206, 233, 235

pluto, Phloeoxena (Oenaphelox), 182, 206,

207, 210, 212, 213, 224, 227, 229, 231,

232, 235, 237

Poa pratensis, 1 1

Podalonia, 123, 126, 127

af finis, 122

atrocyanea, 122

caucasica, 122

chalybea, 122

communis, 122, 124

hirsuta, 122

luctuosa, 122, 124, 126

nigrohirta, 122

robusta, 122, 124

tydei, 122

XXlll

Podalonia (continued),

valida, 113-127

violaceipennis, 1 22, 1 24, 1 26

Podocarpus, 208

Podonosma, 132

Poecilus, 585

Pogonomyrmex, 117, 118

poguras, Cancer, 100

Poisson, R., 64, 67

Polistichus, 585

politum, Trypoxylon, 126

pompiliformis, Notogonia, 127

Pontoppidan, E., 580

Poodry, C. A., 59, 67

Populus balsamifera, 1 1

porrectus, Platynus, 6

portoricensis, Phloeoxena (Phloeoxena), 182,

188, 189, 190, 191, 192, 193, 222, 225,

227, 229, 231, 232, 238

Postlethwait, J. H., 60, 67

Potamogeton pectinatus, 71

Potts, G. R. (see Robinson, J.), 88, 93, 99,

106, 110

Powell, J. A. 121, 127

prasinus, Carabus, 580

pratensis, Poa, 71

Psectrotanypus, 81

Pseudococcidae, 60

Pseudococcus, 66

Pseudomaseus, 583

Pseudomorphini, 240

Psilocephala variegata (sensu latiore), 577

Psocoptera, 61, 65

Psylla buxi, 59, 68

Psyllidae, 59, 63

Pterostichi, 593, 616

Pterostichina, 593, 594, 595

Pterostichini, 147, 153,499, 591-619

Pterostichus, 585, 594, 595, 600, 605

(Gastrellarius) honestus, 594

(Ithytolus), 599

anomalus, 600, 605

orizabae, 600, 605

lobipennis, 607

( Ophryogaster) anomalus, 600

pubera, Echthodopa, 549, 550, 551, 552, 554,

559, 560, 565, 573, 575

pubidorsus, Prionyx, 1 24

pulchrum, Dicolonus, 505, 524, 525, 526, 527,

553

pulmonariae, Phytomyza, 129, 130, 131, 132

punctata, Erpobdella, 81, 89

Priesner, H., 26, 39, 54, 55, 60, 61, 62, 64, 61purpuratus, Dicaelus, 581

Prionyx atratus, 122

parkeri, 124

pubidorsus, 124

thomae, 1 22

Pristolomus, 143, 153, 156, 162

dentifer, 158, 162

Pristonychus, 583

complanatus, 583

Pristosia, 594

Pristosiae, 593

Pristosiina, 594, 595

procera, Ammophila, 121

Procrustes, 585

Proctacanthus, 507

propinqua, Dioctria, 540

(Eudioctria), 540

propinqua, Eudioctria, 538, 539, 540, 541

545, 548, 549, 553, 571, 574

Prosphodrus, 593

Protozoa, 501

Psammophila, 126

Psectrocladius, 81

pusio, Dioctria (Dioctria), 517, 518, 520, 521,

522, 553, 559, 560, 562, 563, 565, 574

quadricollis, Ferus, 143, 162

quadricollis, Stenognathus (Stenognathus),

143, 158, 160, 162, 164

quadrimaculatus, Carabus, 581, 582

Quercus emoryi, 1 1 7

radiata Rohd., Architipula, 5

Rana temporaria, 93

Rangia cuneata, 1 09

rapae, Pieris, 13, 16, 18

Raphidioidea, 60

Rau, N. (see Rau, P.), 1 2 1 , 1 24, 1 27

Rau, P., 121, 124, 127

Razvyazkina, G. M. (see Lange, A. B.), 55, 66

redhorse, northern, 102

Refonia, 594

Reichardt, H., 153, 166, 169, 174, 175, 176,

242, 51 1, 556, 595, 619,621

Reiche, E., 175, 242

Reinert, R. E., 88, 95, 105, 109

Rempel, J. G., 7-25

XXIV

resolutum, Coenagrion, 82, 83

resplendens, Metadioctria, 535, 538

reticulatus, Lebistes, 95, 110

Reyne, A., 55, 60, 63, 64, 67

Rhadine, 145

Rhagio, 51 1

orestes, 576

Rhagionidae, 5 1 1

Rhantus, 84

consimilis, 84

frontalis, 84

wallisi, 84

Rhicciocarpus natans, 71

rhombicus, Limnephilus, 99, 100

ricardo, Cyrtolaus, 591, 597, 598, 599, 600,

602, 603, 606, 608, 609, 611, 613, 615,

617, 618

Ricardo, G., 514, 556

rice, 108

Richards, O. W., 114, 127

Richardson, A. (see Robinson, J.), 99, 100,

106, 110

Rippee, K. P. (see Bradshaw, J. S.), 76, 92

Risebrough, R. W.,'t02, 110

Risler, H., 26, 58, 60, 61, 62

robber flies, 554, 555

Robinson, J., 88, 93, 99, 100, 106, 1 10

robusta, Feronia, 584

robusta, Podalonia, 1 22, 1 24

Rohdendorf, B. B., 3-5

Romoser, W. S., 8, 1 8

Roonwal, M. L„ 12, 18, 20, 21

Rondani, C., 577

Rosen, J. D. (see Khan, M. A. Q.), 103, 104,

109

Rosenberg, D. M., 69-96, 97-1 10

Ross, H. H., 234, 235, 238, 242

Rossi, P., 581, 582, 583, 584

rostrata, Carex, 71, 74, 85

rotundatus, Carabus, 579

rotundicollis, Dinodes, 581

Rousseau, E., 148, 242

Roussel, J. S. (see Newsom, L. D.), 28, 61

Royer, L. M. (see Fredeen, F. J. H.), 70, 76,

93, 102, 108

rubida atrip es, Dioctria (Metadioctria), 536

rubida, Dioctria, 535

(Metadioctria), 536

rubida, Metadioctria, 535, 536, 553, 559, 560,

rubida, Metadioctria (continued), 563, 565,

566, 567, 568, 570, 574

rubida nigripilosa, Dioctria (Metadioctria), 536

rubidus, Dioctria, 536

rubrocinctus, Selenothrips, 28

rufipes. Dinodes, 581

Rudd, R. F., 70, 95, 98, 110

rufus, Myelaphus, 528

Runnels, J., 109

Sabienus, 582

saccharina, Lepisma, 18

sackeni, Dioctria, 544

(Eudioctria), 544

sackeni f. rivalis, 544

sackeni rivalis, 544

sackeni, Eudioctria, 537, 539, 543, 544, 546,

547, 548, 553, 565, 573, 575

group, 537, 543, 544, 547, 548

Sago pondweed, 71

Saha, J. G. (see Fredeen, F. J. H.), 70, 76, 93,

102, 108

sakalava, Foudia, 155

Salonen, F. (see Vaajakorpi, H. A.), 85, 86, 87,

88, 89, 95, 97, 1 10

Saltatoria, 66, 67

Samouelle, G., 579, 582, 586

Sanborn, J. R. , 101, 110

Sanders, FI. O. (see Johnson, B. T.), 88, 89, 94,

95, 97, 105, 106, 108

Sangha, G. K. (see Metcalf, R. L.), 101, 109

sapiens. Homo, 500

saugers, 102

Saunders, C. R. (see Johnson, B. T.), 88, 94,

97, 105, 106, 108

Savage, J. M., 234, 242

Saxifragaceae, 137

sayi, Hy grot us, 84

Scaphinotus elevatus, 169

Scarabaeidae, 103

Scarites bucephalus, 582

calydonius, 582

cyaneus, 580

dama, 582

depressus, 584

gib bus, 583

marginatus, 584

piceus, 584

sphaerocephalus, 582

thoracicus, 583

XXV

Scaritini, 242, 619

Scaurus sulcatus, 582

Sceliphronini, 1 26

Scenedesmus obliquus, 95, 109

Scenopinidae, 506

Schafer, R., 59, 67

Schaller, J. G., 582, 584

Schaupp, F. G., 215, 242

Schiner, J. R., 556

Schi0dte, J. C., 580

Schizogenius, 242, 619

Schmidt, K., 63, 66, 67

Schmidt-Goebel, H. M., 582

Schmulbach, J. C. (see Kawatski, J. A.), 88,

94, 104, 106, 108

Schneider, D., 60, 61, 62, 63, 67

Schneiderman, H. A. (see Poodry, C. A.), 59,

67

(see Postlethwait, J. H.), 60, 67

Scholl, G., 8, 12, 15, 16, 17, 18, 20, 21

schwarzi, Phloeoxena (Phloeoxena), 143, 181

184, 186, 187, 188, 189, 190, 191, 192,

193, 222, 223, 225, 227, 228, 229, 231,

232, 233, 234, 237, 238

group, 238

Scirpus decoratella, 79

validus, 71

scrobiculatus, Carabus, 580

Scrophulariaceae, 39

Scudder, G. G. E., 8, 18

sculptilis, Colymbetes, 84

sea urchin, 1 00

sedges, 71

Sehgal, V. K., 129, 137

Seidlitz, G. von, 169, 242

Sekhon, S. S. (see Slifer, E. H.), 28, 61, 62,

63, 68

Selenothrips mbrocinctus, 28

Seminole, Dioctria (Dioctha), 515

(Nannodioctria), 514, 516, 519, 521, 553,

559, 560, 563, 565, 566

serniscidatus, Acilus, 84

senecta, Bohartia, 505, 530, 532, 533, 534,

535, 553

sericeum, Oxybelus, 126

Sericoda, 594

Serville, J. G. A., 584

sexpunctatus, Carabus, 579

Sharov, A. G., 11, 17, 18, 19, 21

shellfish, 99

Slump, N. F. (see Leland, H. V.), 103, 109

Shuckard, W. E., 114, 127

Sialis,99, 100

Slewing, R., 10, 15, 16, 18

signata, Coptodera, 205, 213

signata, Oenaphelox, 180, 182, 223, 237

signata, Phloeoxena ( Oenaphelox), 206, 208,

209, 210, 212, 213, 214, 215, 216, 217,

218, 219, 220, 226, 227, 229, 231, 232,

233, 234, 235

var. collaris, 2 1 5

group, 206, 207, 209, 223, 233, 235

hoegi, 221

var. nigripennis, 2 1 5

signata, 221

Silphidae, 63

silvaticLis, Pauropiis, 1 9

Silvestri, F., 12, 18

simplex, Dicolonus, 524, 525, 526, 527, 553,

;, 559, 561, 565, 573, 575

simplex, Taeniothrips, 28

Simpson, G. G., 523, 555

Simuliidae, 95

Simulium vittatum, 99, 100

Singilini, 147, 155, 169, 242

Singilis, 169

Sinurus, 145

Siphonaptera, 60

Skerrett, E. J., 76, 95

Skrinde, R. T.,(see Buescher, C. A.), 70, 93

Slifer, E. H., 28, 61, 62, 63, 67, 68

slugs, 102, 103, 104

Smart, J., 8, 17, 19

Smith, C. E. (see Newsom, E. D.), 28, 67

Smith, D. J. (see Breidenback, A. W.), 75, 92

snail, 101, 103, 104

Snodgrass, R. E., 7, 9, 10, 1 1, 15, 16, 19, 20,

21, 58, 59, 60, 61, 68

Somotrichini, 147, 169, 242

somotrichoids, 143, 153, 155, 169

Somotrichus, 143, 147, 152, 153, 155, 169,

unifasciatus, 152, 154, 169, 170

vadoni, 169

Sparrow, J. T. (see Hearle, J. W. S.), 28, 66

sparsipilosum, Dicolonus, 504, 524, 525, 526,

527, 553

"''species verae'\ Phloeoxena, 179

speciosus, Sphecius, 127

XXVI

Sphaerium, 89

sphaerocephalus, Scarites, 582

Sphecidae, 113-127

Sphecinae, 126

Sphecini, 126

Sphecius speciosus, 114, 127

Sphecoidea, 126

Sphex, 114, 121

tepanecus, 121, 126

Sphodri, 593

Sphodrides, 586

Sphodrina, 594, 595

Sphodrus venust us, nomen nudum ( = Pristo-

nychus venustus), 583

spinicauda, Cyrtolaus, 591, 598, 599, 600,

601, 603, 606, 607, 609, 610, 611, 613,

615, 616, 617

spinipes, Blaps, 584

spinosior, Opuntia, 117

spoliatus. Carabus, 581

stagnalis, Helobdella, 81

stagnalis, Lymnaea, 77, 78, 85, 88, 90, 104

Stannard, L. J., 58, 64, 68

Stanton, R. H. (see Khan, M. A. Q.), 103,

104, 109

Staphylinidae, 102, 609

statices, Haplothrips, 60, 62

Steiner, A. L., 113-127

Steinmann, H., 8, 19

Stenoglossa, 143, 152, 153, 156, 163, 171,

172, 177, 194

lineata, 172, 173, 174

variegata, 172

Stenognathus, 143, 152, 153, 155, 156, 159-

161, 162, 163, 166

chaudoiri, 143, 162, 163, 164

dentifer, 162, 164

quadricollis, 143, 162

(Stenognathus), 143, 153, 155, 161-162

quadricollis, 158, 160, 164

Stenopogon inquinatus, 576

Stenotelus, 145

Sternorrhyncha, 61

Shekel, L. F., 70, 88, 93, 95, 103, 104, 106,

108, 110

Stomina, 595

Stone, A., 577, 578

Straneo, S. L., 171, 242

Streblomorpha, 4

striatus, Labocephalus, 151

striola, Carabus, 579

striolatus, Carabus, 579

subaeneus, Illybius, 84

subappendiculatus, Thyreopterus, 180

subeyaneus, Carabus, 583

subeyaneus, Oreodicastes, 158, 160

subeyaneus, Oxyglossus, 156

subiridescens, Cyrtolaus, 591, 598, 599, 600,

601, 603, 606, 607, 608, 609, 612, 613,

615, 616, 617, 618

suckers, longnose, 102

white, 102

sulcatus, Scaurus, 582

Sutherland, D. J. (see Khan, M. A. Q.), 103,

104, 109

sylvatica, Myosotis, 131

Sympetrum costiferum, 82

internum, 82

Symphoromyia, 511, 576

Symphyla, 14

Symphyta, 60

symphyti, Phytomyza, 129, 130, 131, 132,

135, 137

Symphytum, 130, 132

syngenesiae, Phytomyza, 137

Synuchi, 593

Synuchina, 594, 595

Syrphidae, 556

t abaci, Thrips, 68

Tabanidae, 511, 555

Tabanomorpha, 506, 507

Tacana, 143, 152, 153, 155, 156, 180, 182,

223, 228, 233, 235, 236, 237

herculeano, 181, 182-183

Tachalus, 594

Tachymenis, 194

Tachysphex, 1 14, 1 17, 121, 124, 125, 127

terminatus, 121

Tachytes, 114

tadpoles, 93

Taeniothrips fedorovi, 65

simplex, 28

Tarzwell, C. M., 70, 95

temporaria, Rana, 93

Tendipes, 88

Tenebrio, 16, 19

molitor, 19

XXVll

tentans, Chironomus, 105, 107

tenuis, Bohartia, 505, 530, 533, 534, 535,

553

tepanecus, Sphex, 121, 126

Terebrantia, 26, 63, 67

terminatus, Tachysphex, 121

Termitoxeniomorpha, 4

terrestris, Lumbricus, 105

ferric ola, Carabus, 583, 584

Terriere, L. C, 98, 110

teter, Dicaelus, 581

teter, Platynus, 6

Tetragoneuria, 1 1 0

thapsus, Verbascum, 28

Thereva flavicincta, 576

Therevidae, 499, 506, 507, 577-578

Thermobia, 12

Theromyzon, 81

thomae, Prionyx, 122

Thompson, A. R., 70, 93, 103, 108

thoracicus, Scarites, 583

Thorpe, W. H., 13, 19

Thripidae, 25, 26, 28, 54, 55, 60, 61, 63-66,

68

Thripinae, 64

thrips, 63, 64, 65, 66, 67, 68

red clover, 66

tobacco, 66, 67

Thrips physapus, 60, 62, 67

tabaci, 68

Thunberg, G., 580

Thurman, E. B., 9, 19

Thyreopterides, 147, 159, 240

Thyreopterinae, 147, 148

thyreopterines, 143, 148

Thyreopteritae, 147

thyreopteroids, 143, 153, 154, 156

Thyreopterus, 145, 156

subappendiculatus, 1 80

Thysanoptera, 25, 26, 54, 55, 60, 61, 63, 64,

65, 66, 67, 68

Thysanotini, 147

Thysanura, 18, 61

tibialis, Alpaeus, 580

tibialis, Dioctria (Eudioctria), 544

tibialis, Eudioctria, 537, 538, 543, 546, 547,

548, 554, 564

Tiegs, O. W., 8, 14, 15, 17, 19, 20, 21, 60, 67

tiger beetles, 1 54

tinctipennis, Colpodes, 6

tinctipennis, Platynus, 6

Tipulidea, 5

Tipulomorpha, 4

tlarnayensis, Platynus, 6

torridus, Carabus, 580

Trialeurodes vaporariorum, 59

Triboleum castaneum, 1 54

Trichodesma, 132

Trichophthalma lundbecki, 576

Trichoptera, 60, 89, 93, 242

triscula, Lemna, 71, 74, 85

tritici, Frankliniella, 28, 61, 62, 68

trivolvis, Helisoma, 77, 78, 79, 88,90

trout, brook, 105

truncatellus, Carabus, 582

Truncatipennes, 241, 593

tryoni, Dacus, 65

Trypoxylon, 114, 127

politum, 126

Tsuneki, K., 1 15, 1 16, 122, 127

Tubulifera, 26, 63, 68

Tucker, R. K. (see Crosby, D. G.), 88, 93, 105, 107

tydei, Ammophila, 127

tydei, Podalonia, 122

Typha lati folia, 71

Uca minax, 101

Ullmann, S. L., 10, 19

undata, Phloeoxena (Oenaphelox), 157, 180,

206, 207, 208, 209, 210, 21 1, 212, 213, 227,

229, 231, 232, 233, 235

undulata, Notonecta, 78, 80

undulata, Phloeoxena ( Oenaphelox), 208

unica, Eudioctria, 537, 538, 543, 545, 548, 549,

553, 554, 556, 566, 572, 575

unicolor, Phloeoxena ( Phloeoxena), 143, 201

unifasciatus, Somotrichus, 152, 154, 169, 170

urothripines, 54

uzeli, Gynaikothrips, 28

Vaajakorpi, H. A., 85, 86, 87, 88, 89, 95, 97, 1 10

vadoni, Somotrichus, 169

vagus, Camponotus, 67

valida, Podalonia, 113-127

validus, Catascopus, 167

validus, Scirpus, 71

Van Middelem, C. H.,71,95

vaporariorum, Trialeurodes, 59

variegata, Psilocephala (sensu latiore), 577

variegata, Stenoglossa, 172

XXVlll

Velindomimus, 169 Westwood, J. O., 579, 580, 581, 583, 584, 586

Velindopsis, 169 White, D. A. (see Bradshaw, J. S.), 76, 92, 106

velutina, Fraxinus, 1 17 White, M. J. D., 4

venustus, Sphodnis, nomen nudum ( = Pristo- whiteflies, 59

nychus venustus), 583 white grubs, 103

vem, Dioctria, 518 Whitehead, D. R., 7, 183, 222, 234, 242, 499,

(Dioctria), 517, 518, 520, 521, 553, 569, 591-619

574

verbasci, Haplothrips, 25-68

Verbascum, 39

thapsus, 28

Verbesina encelioides, 116, 117

vernalis, Carabus, 585

vespilloides, Necrophorus, 63

vestitus, Carabus, 581

violaceipennis, Podalonia, 122, 124, 126

violaceus, Dicaelus, 581

viduus, Carabus, 579

virginiana, Diodontus, 121

viridana, Lytta, 7, 9, 15, 16, 17, 18, 20

Vitis arizonicus, 1 1 7

vittatum, Simulium, 99, 100

vivalis, Carabus, 579

vulgaris, Carabus, 580

vulgaris, Hippuris, 71

vulgaris, Homarus, 100

vulgata. Patella, 100

Walker, E. M., 499-500, 539, 556

walkeri, Palaeodipteron, 4

Wallace, R. R., 69, 95, 99, 100, 104, 105,

110

walleye, 102

wallisi, Rhantus, 84

Walsh, D. F. (see Kennedy, H. D.), 69, 94,

105, 109

Walton, W. R., 508, 556

Watts, J. G., 28, 68

Waybrant, R. C. (see Hamelink, J. L.), 104,

105, 106, 108

weaver-birds, 154

Weber, H., 14, 15, 17, 19, 20, 21, 59, 60, 63

64, 68

Weiss, C. M. (see Wilkes, F. G.), 97, 103, 104,

105, 106, 110

Wells, O. C., 243, 244

Wells, R. E. (see Bowser, W. E.), 71, 92

Welsh, P. S., 74, 96

West, A. W. (see Wallace, R. R.), 69, 95

Westlake, W. E., 70, 71, 96

Wiebes, J. T. (see Lieftinck, M. A.), 149

Wigglesworth, V. B., 12, 19

Wilcox, J., 507, 51 1, 514, 515, 517, 518, 519,

530, 535, 536, 537, 538, 539, 540, 541, 542,

543, 544, 555, 556

wilcoxi, Dioctria (Dioctria), 505, 517, 518, 520,

521, 522, 553, 569, 574

Wilkes, F. G., 97, 103, 104, 105, 110

Williams, F. X., 114, 122, 127

Williams, T. R. (see Hynes, H. B. N.), 69, 94

Willis, H. L., 616, 619

Williston, S. W., 528, 542, 544, 556

willow, 523

Wilson, A. J., 85, 86, 87,93

Wojtalik, T. A., 88, 96

Wolin, R. J. (see Khan, M. A. Q.), 109

Wolkoff, A. W. (see Mackay, D.), 70, 94

Wood, D. M., 499, 505-576

Woodward, T. E., 59, 64, 68

Woodwell, G. M.,98, 110

wrightii, Platynus, 117

Wurster, C. F., 88, 96, 98, 103, 1 10

Wurzinger, K. -H. (see Dindal, D. L.), 104, 107

Xylocelia, 121

Yu, C. -C. (see Sanborn, J. R.), 101, 104, 107,

110

yuccae, Bagnalliella, 26, 61, 62

yellowfish, 102

Zabik, M. J. (see Derr, S. K.), 97, 104, 105, 107

Zaitzev, V. F., 577, 578

zonatus, Carabus, 581

zooplankton, 70, 75, 77, 88, 90, 91

Zygoptera, 81, 82, 83, 85, 89, 90

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Quaest

lones

Entomologicae

1975 )

A periodical record of entomological investigations,

published at the Department of Entomology,

University of Alberta, Edmonton, Canada.

VOLUME n

NUMBER 1

JANUARY 1975

I

1

c

QUAESTIONES ENTOMOLOGICAE

A periodical record of entomological investigation published at the Department of

Entomology, University of Alberta, Edmonton, Alberta.

Volume 11 Number 1 January 1975

CONTENTS

Editorial 1

Book Review - Rohdendorf, B. B. 1974. The Historical Development of Diptera 3

Whitehead - Additions to “Annotated Key to Platynus"' (Coleoptera: Carabidae:

Agonini) 6

Rempel - The Evolution of the Insect Head: the Endless Dispute 7

Heming - Antennal Structure and Metamorphosis in Frankliniella fusca (Hinds)

(Thripidae) and Haplothrips verbasci (Osborn) (Phlaeothripidae) (Thy-

sanoptera) 25

Rosenberg — Fate of Dieldrin in Sediment, Water, Vegetation, and Invertebrates of a

Slough in Central Alberta, Canada 69

Rosenberg — Food Chain Concentration of Chlorinated Hydrocarbon Pesticides in

Invertebrate Communities: A Re-evaluation . 97

Book Review — Nachtigall, Werner. 1974. Insects in Flight: A glimpse behind the

scenes in biophysical research Ill

Editorial

With sorrow, I record in these pages of Volume 1 1 that Brian Hocking, founder and first

editor of this journal is with us in spirit, only. His time as a living mortal ended on May 23,

1974. His last bed-ridden days were devoted to writing and to visits with the members of

his family and his friends including colleagues and students. How intensely he suffered we

will never know, for his courage and interest in life sustained him, and even as his last bit of

strength ebbed, he completed an advanced draft of a biology textbook that he had wanted to

write. Thus, his thoughts were of life, even as the specter of death loomed large. We have no

record of self-pity, no words of bitterness or disappointment. If Brian Hocking felt these

emotions, he did not share them with his associates. He left us, knowing that he had done his

best in all things, and that we would try to follow his example. We can do no less. So it is

that I take up his pen officially, as the second editor of Quaestiones Entomologicae. Though

the editorial section will lack the wit, brilliance, and clever turn of phrase that characterized

Brian’s writing, the main body of the publication will be of the same high level as was achieved

during the days of his editorship. To this end, I wish to state a few points of editorial policy,

and to explain a few changes.

First, is the matter of policy. As previously, we will accept papers on all aspects of entomol-

ogy. Overall, we prefer comprehensive treatments of subjects: results of original research; ex-

tensive, analytical book reviews; and broadly based comprehensive and critical reviews of the

literature dealing with various aspects of entomology. We want extensive and intensive system-

atic treatments of any group of insects — but not unintegrated collections of descriptions of

new taxa. We prefer that the integration be in terms of phylogenetic and zoogeographic con-

siderations, but we will accept good Adansonian-type treatments. We will make exceptions.

Being in western Canada, we have a special obligation to extend knowledge of the poorly

known local fauna. Thus, for this part of the world, we will consider simple descriptive treat-

ments, but preferably extensive ones.

As in the past, all submitted manuscripts will be examined by competent reviewers, prior

to acceptance for publication. In the last issue of each volume, names of reviewers will be

listed as a public, formal acknowledgement by the editor.

We regret that page charges have to be levied. This is mainly because our list of subscribers

continues to be short, so income is restricted. We also regret that prices have had to be increased,

and that they will have to be increased still more, so long as inflation continues to erode the

buying power of money.

Examination of the cover of this issue will reveal that the “Quaestiones entomologicae”

has been replaced by “Quaestiones Entomologicae”. We are departing from the “World List”

style of reference citation, and hence from its type of notation. It takes little extra space to

provide bibliographic citations with the name of the journal spelled out fully, rather than in

abbreviated form. This additional information can make easier the task of finding a given paper,

should one wish to locate a listed citation.

We are also abandoning the use of Roman numerals as a designation for each volume, and

using instead arabic numbers. This is hardly in keeping with our Latin title, but we expect

to be publishing for a long time, and in the year 2041 it will be easier to think of and write

“Volume 77” than “Volume LXXVII”. So much for editorial housekeeping.

This journal is fortunate to count among its contributors Dr. J. G. Rempel, author of the

first major paper in this volume, and one of the distinguished biologists of Canada. Now re-

tired from his long career of teaching and research, in the Department of Biology, University

of Saskatchewan, Dr. Rempel and his wife Greta continue to enjoy life among friends of

many years, in Victoria, British Columbia. Dr. Rempel’s daily round of activity includes study

of insect embryos, and from this research and his extensive knowledge of the literature comes

his masterful synthesis “The Insect Head: the endless dispute”. I am sure the spirit of Brian

Hocking is pleased that his “Quaest. ent.” contains this paper, for it is the quintessence of the

sort of publication that he valued, and at a more personal level, he valued Dr. Rempel as a

scientist and friend.

George E. Ball

3

Book Review

ROHDENDORF, B. B. 1974. The Historical Development of Diptera (English Edition). Uni-

versity of Alberta Press, Edmonton, xv + 360 pp. 85 figures. Size 9 inches by 6 inches, cloth

covers. Price: $13.50 Canadian, postage free. (Translation of Russian Edition published by

“Nauka” as Volume 100, Transactions of the Institute of Paleontology, Academy of Sciences

of the USSR, Moscow, 1964).

I find it difficult to review this work as 1 disagree fundamentally with some of the author’s

methods and principles. Rohdendorf is clearly a representative of those orthodox Marxists who

believe in historical explanations in the form of the dialectical triad (thesis, antithesis, synthesis).

In Rohdendorf s presentation the theses and antitheses are called “conflicts” and the syntheses

“solutions”. Following the orthodox dialectical schema, evolutionary progress is seen as the

successive solution of conflicts each produced by the solution of some previous conflict. These

methodological concepts are presented in Part IV of Rohdendorf’s book as if they were some-

how conclusions drawn from his studies. In fact they are preconceptions through which he

attempts to interpret his data. Dialectics is not an empirical theory, but has its roots in idealist

philosophies (notably that of Hegel) utterly opposed to empirical science. I refer the reader to

an excellent review of this subject by Karl Popper (1940. What is dialectic? Mind 49: 403-426).

This is not to say that Rohdendorf s explanations are all valueless, and without interest.

There are undoubtedly points of importance in his analysis. But by his rigid adherence to the

dialectical pattern of explanation, he denies himself the possibility of subsuming evolutionary

changes under covering laws in accordance with the generally accepted concept of scientific

explanation. Let us consider one example, Rohdendorf s first “concrete example of internal

conflicts” (p. 319). Here we are offered an explanation of the evolution of apodous (legless)

laivae in Diptera, in the form of a solution to the following conflict:

A. Need for a solid substrate for B. Need for nutrition in a moist,

locomotion of the larva, pro- semiliquid substrate, forming

vided by three pairs of walking comparatively large masses,

legs.

Obviously there is some truth in this. An insect with thoracic walking legs which penetrated

a mass of viscous substrate would find its legs an obstacle, not an aid, to locomotion. There-

fore it is reasonable to hypothesize that leglessness was evolved in one of the ancestors of the

Diptera whose larvae came to feed in a viscous substrate, such as wet soil or decomposing or-

ganic matter of some kind. However, the covering laws under which this morphological change

can be subsumed must surely involve the mechanics of locomotion through a viscous medium,

which Rohdendorf does not discuss. Moreover, it makes little sense to talk of a conflict between

locomotion and nutrition in the larval stages of holometabolous insects, since their need for

locomotion is largely determined by, not in conflict with, their need for food and shelter. But

the invalidity of the supposed conflict is not the most important point. What is more serious,

and applies even in those examples where the stated conflicts seem valid, is that Rohdendorf s

concern to discover a “conflict” has diverted his attention from the possibility of genuine nomo-

logical explanation.

Throughout his book Rohdendorf emphasizes the need for functional and ecological explan-

ations of evolutionary change. This emphasis seems to me well justified. But I think we will

make more progress in providing such explanations if our thinking is not rigidly constrained

by the dialectical schema.

Now to more specific matters. Part I of Rohdendorf s book deals with the characterization

and classification of the Diptera. He rejects the conventional suborders Nematocera, Brachy-

cera and Cyclorrhapha, but makes his primary division into the suborders Archidiptera

Quaest. Ent, 1975, 1 1 (1)

4

(Nymphomyiidae only in the Recent fauna) and Eudiptera (all the rest). The first name is in

error (Archaeodiptera conveys the intended meaning), as has already been pointed out by

Cutten and Kevan (1970. The Nymphomyiidae [Diptera] , with special reference to Palaeo-

dipteron walked Ide and its larva in Quebec, and a description of a new genus and species from

India. Can. J. Zool. 48: 1-24). The Eudiptera are then divided into 12 infraorders (Deutero-

phlebiomorpha, Blephariceromorpha, Tipulomorpha, Bibionomorpha, Asilomorpha, Musidoro-

morpha, Phoromorpha, Termitoxeniomorpha, Myiomorpha, Braulomorpha, Nycteribiomorpha

and Streblomorpha).

The validity of Rohdendorf s separation of Nymphomyiidae from the rest of the Diptera

remains unsettled. Cutten and Kevan (loc. cit.) are inclined to support Rohdendorf s view,

but other authors have advocated alternatives. A cytological study might well settle this dis-

pute, since White’s (1949. Cytological evidence on the phytogeny and classification of the

Diptera. Evolution 3: 252-261) work has shown that the major subgroups of Diptera have

quite different genetic systems. (Rohdendorf s failure to consider White’s work was a serious

omission).

Rohdendorf s subdivision of the Eudiptera is highly arbitrary and has found no acceptance

outside the Soviet Union. It is evident from his phytogeny diagrams (Figs. 81-85) that many

of his group concepts are not intended to represent monophyletic groups in the sense of phylo-

genetic (Hennigian) systematics. Rohdendorf believes in the evolution of new high-ranking

groups by rapid “qualitative transformation”, and sees the gaps between existing groups as

evidence of such transformations. Consequently he identifies systematic groups with “biotypes

. . . associated with definite conditions of existence, which are reflected in similarities of struc-

ture, function and ontogentical development” (p. 17). Rohdendorf s nomenclatural scheme

is thus not intended to represent the branching pattern of phytogeny, but rather constitutes a

typological classification superimposed on phytogeny. It is easy to misunderstand his views on

phylogenetic relationships, as these are not consistently represented in the classification and

there is an inherent ambiguity in his use of “relationship” and similar terms in discussion (some-

times typological, sometimes phylogenetic relationship is meant). Now that the complete text

of this work is available to me, I find that both Hennig and myself have succeeded in misunder-

standing certain points on this account. The only monophyletic groups among Rohdendorf s

infraorders are the Deuterophlebiomorpha, Blephariceromorpha, Musidoromorpha, Termitox-

eniomorpha, Braulomorpha and Streblomorpha. All these are small, highly specialized groups

usually given family, or even lower, rank by Western authors. We are told that the first four are

“relicts” (a term used liberally in Rohdendorf s evolutionary discussions), but not offered the

slightest evidence that they were ever more diverse or more widely distributed than at present.

A detailed commentary on Rohdendorf s classification would take up much space, and is

scarcely necessary since such commentary has already been published. Rohdendorf s infraorder

and superfamily classification of the Eudiptera has been discussed in some detail by Hennig

(1968. Kritische Bemerkungen fiber den Bau der Flugelwurzel bei den Dipteren und die Frage

nach der Monophyhe der Nematocera. Stuttg. Beitr. Naturk. no. 193. 23 pp.). His treatment

of the Cyclorrhapha has been discussed in my recent book (Griffiths, G. C. D. 1972. The

phylogenetic classification of Diptera Cyclorrhapha, with special reference to the structure

of the male postabdomen. Junk, The Hague. 340 pp.). Neither of us were of the opinion that

the changes proposed by Rohdendorf are improvements from the viewpoint of phylogenetic

systematics.

Part II of Rohdendorf s work is entitled Diptera of the Geological Past. Most of this Part

is devoted to descriptions and discussion of the Upper Triassic Issyk-kul and Middle Jurassic

Karatau faunas of Central Asia. The work of Rohdendorf and his collaborators (the information

in the present work has since been supplemented by a series of further publications) remains

5

the main primary source of information on Mesozoic Diptera, and should be studied by all

interested in the history of the Diptera. Again I will not attempt to give a detailed commentary,

but refer the reader to the discussion in Hennig’s recent book on fossil insects (1969. Die Stam-

mesgeschichte der Insekten. Waldemar Kramer, Frankfurt am Main. 436 pp.). However, certain

major points of criticism do deserve mention here.

Like Hennig, I am not satisfied that the wing fragments referred by Rohdendorf to the

“Archidiptera” are Diptera at all. More complete fossils are needed for the relationships of

these insects to be reliably assessed. It seems to me quite unwarranted to base elaborate evol-

utionary hypotheses on the available fragments, as Rohdendorf does later (p. 289).

I also share Hennig’s opinion that there must be inaccuracies in certain of the wing figures,

which show cross-veins and vein stubs in positions where these cannot be found in any existing

Diptera. For instance, if we consider the Triassic fossils ascribed to the Tipulidea, there is only

one (Architipula radiata Rohd.) whose wing venation is entirely convincing. Rohdendorf was

doubtless faced with technical problems in distinguishing impressions made by wing veins from

other impressions in the rock. Such problems may not be entirely soluble with present-day

paleontological techniques. But what is more difficult to understand is that Rohdendorf seems

to take little account of these problems and frequently uses doubtful features as defining char-

acters for his fossil taxa. For this reason I fear that future workers without access to the orig-

inal material will find it difficult to interpret many of these taxa.

If Rohdendorf reads this review, his reaction will likely be that I am trying to “abolish the

dialectic” (a shocking and unthinkable thing to the orthodox Marxist). I openly admit that

this is my intention. The privileged position of dialectics in Marxist countries (due to the

acceptance of Hegelian dialectics by Marx) is a constant threat to the progress of the empirical

sciences there. Most Soviet scientists, while paying lip service to dialectics, have the good sense

not to allow it to influence their practical work. Rohdendorf is an exception. He really tries to

implement the official party line. The results serve, in my opinion, merely to show that dialectics

has nothing to contribute to empirical science except an empty play of words.

In view of these weighty criticisms, do I think it was worthwhile publishing an English trans-

lation of this work? Yes, marginally. Rohdendorf s work is the main source of information on

Mesozoic Diptera, and buried in the dialectical verbiage are occasional points of genuine interest

and importance. However, only those with extensive knowledge of the Diptera will be able to

separate the grain from the chaff. Keep this book away from inexperienced students; it will only

confuse them.

The University of Alberta Press may be complimented for an attractive production, printed

on much better paper than the Russian original. Printing errors are few, and mostly unimportant.

But note that in the third paragraph on page 111, “Heleomyzidea” should replace “Heleomyzidae”

in the first hne, and “Helcomyzinae” should replace “Heleomyzinae” in the last line.

Graham C. D. Griffiths

Department of Entomology

University of Alberta

Edmonton, Alberta T6G 2E3

Quaest. Ent., 1975, 11 (1)

ADDITIONS TO “ANNOTATED KEY TO PLATYNUS^'

(COLEOPTERA: CARABIDAE; AGONINI).

DONALD R. WHITEHEAD

Organization for Tropical Studies

c/o Department of Entomology

U.S. National Museum Quaes tiones Entomologicae

Washington, D. C. 20560 11:6 1975

Since publication of my “Annotated key to Platynus'" (1973), I have discovered that two

important entries were inadvertently dropped from early drafts and hence omitted in final

proof.

Page 192: replace discussion under entry for/*, consularis and add entry fox P. convexulus,

as follows.

Platynus consularis is known only from various localities in the state of Guerrero. This

species has been confused in collections and in the literature with P. ebeninus but is well dis-

tinguished by characteristics cited in the key, and the two species probably are not closely

related.

Platynus convexulus (Casey), new combination.

Anchomenus convexulus Casey 1920: 38. Holotype female (?), labelled “Tamaulipas Mex.

. . .” (script), “TYPE USNM 47412” (USNM). Probably a female, but not sexed definitely

since both front tarsi as well as all other claw bearing tarsal articles are lost.

Agonum convexulum, Csiki 1931: 862 (subgQxms Anchomenus); Blackwelder 1944: 41.

This form is evidently quite closely related to P. porrectus, and perhaps is conspecific with

it. Specimens seen from the state of Hidalgo agree with the holotype of P. convexulus. Speci-

mens from various localities in the Trans- Volcanic Sierra differ by having dorsoapical setae on

the hind femora, and may represent a distinct species.

Page 212: insert following between entries forP. teter andP. tlamayensis.

Platynus tinctipennis (Bates), new combination.

Colpodes tinctipennis Bates 1891: 257. Lectotype female, here designated, labelled “TYPE

H. T.”, “Ciudad, Durango. Hoge.” (BMNH).

Colpodes tinctipennis, Csiki 1931: 764; Blackwelder 1944: 40.

This species is known only from the original localities in Durango, Ciudad and Refugio.

REFERENCE

Whitehead, D. R. 1973. Annotated key to Platynus, mchxdmg Mexisphodrus and most “Col-

podes”, so far described from North America including Mexico (Coleoptera: Carabidae:

Agonini). Quaestiones Entomologicae 9: 173-217.

THE EVOLUTION OF THE INSECT HEAD :

THE ENDLESS DISPUTE*

J. G. REM PEL

Department of Biology

University of Saskatchewan

Saskatoon, Saskatchewan

Quaes tiones Entomologicae

11: 7-25 1975

It would be too bad if the question of head segmentation ever should be

finally settled; it has been for so long such fertile ground for theorizing

that arthropodists would miss it as a field for mental exercise.

Snodgrass, 1960

Head segmentation in insects has long been a subject of dispute. The controversy concerns

possible pregnathal segments, namely intercalary, antennal and preantennal, and the presence

or absence of an acron. The total number of segments has been placed as low as three and as

high as seven as seen in a brief review of the theories by Snodgrass, DuPorte, Butt, Ferris,

Sharov, Roonwal, Chaudonneret, Matsuda, Imms and Manton, Eastham, Tiegs and Weber,

Scholl, Rempel and Church. In the embryo o/Lytta viridana (Coleoptera: Meloidae) an inter-

calary segment is recognized by the presence of coelomic sacs, a neuromere ( trito cerebrum),

and apodemes (anterior tentorial arm); an antennal segment by a pair of appendages (antennae),

coelomic sacs, a neuromere (deutocerebrum), and apodemes (mandibular extensor apodemes);

a preantennal segment by a pair of appendages (labrum), coelomic sacs, and apodemes. The

proposal is made that the ancestor of the insect was annelid-like, that the prostomium became

the acron and that the six originally postoral segments joined the acron to form the head tagma.

The theory differs from the classical theory of a six-segmented insect head in that the labrum

is considered to be appendiculate innervated originally by the nervus connectivus from a pre-

antennal neuromere (prosocerebrum).

La segmentation de la tete des insectes a ete depuis longtemps le sujet d’une longue dispute.

Le probleme concerne les segments preoraux: Tintercalaire, Tantennaire et le preantennaire,

et la presence ou Tabsence dun acron. Le nombre de segments varie entre trois et sept selon

les courtes revues de ces theories par Snodgrass, DuPorte, Butt, Ferris, Sharov, Roonwal,

Chaudonneret, Matsuda, Imms et Manton, Eastham, Tiegs et Weber, Scholl, Rempel et Church.

Chez Tembryo de Lytta viridana ( Coleoptere: Meloidae) un segment intercallaire est reconnu

par la presence de sacs coelomiques, dun neuromere ( tritocerebrum) et d’apodemes ( embranche-

ment anterieure, du tentorium); un segment antennaire est reconnu par la presence d’une paire

d’appendices (antennes), de sacs coelomiques, d’un neuromere (deutocerebrum) et d’apodemes

(apodemes des extenseurs des mandibules); un segment preantennaire est reconnu par la pre-

sence d’une pair d’appendices (labre), de sacs coelomiques et d’apodemes. Nous proposons que

I’ancetre des insectes ressemblaient les annelides, que le prostomium est devenu I’acron et que

les six segments postoraux originaux ont joint I’acron pour former la tete. Cette theorie differe

de la theorie classique de la tete de I’insecte composee de six segments en ce que le labre est

considere comme innerve par le nervus connectivus d’un nuromere preantennaire (prosocere-

brum).

* Based on a paper read at a seminar of the Departments of Entomology and Zoology, Univer-

sity of Alberta, Edmonton, September 26, 1974.

8

Rempel

INTRODUCTION

The problem of head segmentation in insects has been a lively topic of discussion since the

turn of the last century. Numerous papers have appeared since then, each one outlining, often

at great length, the history of evolutionaiy thought with respect to head segmentation. One

of the latest and most detailed reviews is given by Matsuda (1965). When the latter work ap-

peared in print it seemed that the problem was largely solved and little would be gained in

reopening the debate. But new information has been published lately which suggests that part

of the protocerebrum is a preantennal neuromere (Malzacher, 1968; Scholl, 1969) and that

the labrum, which has been the center of the controversy, is indeed an appendicular structure

(Rempel and Church, 1971).

Of further significance is that recent publications still feature the theories expounded by

North American workers, notably Snodgrass, Ferris and Butt, although the weight of evidence

is against these theories. Of concern is the adoption of the Snodgrass (1935) theory by the

author of a monumental work on the structure and function in the nervous systems of in-

vertebrates (Horridge, 1965) and by the author of a recent textbook in entomology (Romoser,

1973). The Ferris (1950) theory which has never received serious consideration was featured

by Demerec (1950), in his “Biology of Drosophila''. Two recent papers (Steinmann, 1970;

Dixit, 1972) follow Butt (1957, 1960) in assigning the labrum to the intercalary or tritocere-

braf segment and are thus subject to the same criticism as the papers by Butt.

THE INSECT HEAD

The Problem of Head Metamerism in Insects

In the study of animal evolution various approaches have been used. The most direct evi-

dence is afforded by the fossil record, but in the case of insects, palaeontology has not given

us the information that we require. The secret of head metamerism should be looked for in

early Devonian fossils for the primitive insect stock appeared in the Devonian (Smart and

Hughes, 1972), but such material is not available. Indeed, we even lack sufficient later palae-

ontological specimens to enable us to reconstruct an undisputed phylogenetic tree for the

insectan world (Scudder, 1973).

Similarly, problems have been encountered in studies of comparative morphology. First,

most insect morphologists who have been concerned with head segmentation assume that the

present day arthropods are the product of a monophyletic line of evolution. On this assumption

any theory of head segmentation for insects must be equally applicable to the Chelicerata and

the Crustacea. That this approach is questionable is evident from the extensive studies of Tiegs

and Manton (1958) and Manton (1964). These authors advance evidence that in the case of

arthropods we are confronted with a di- or polyphyletic origin. For example, Manton has

shown that the mandibles have evolved independently in (i) the Crustacea; (ii) the Chelicerata;

(iii) the Onychophora, Myriapoda, Insecta. In the last the mandible is a modified whole limb,

while in the first two the jaw arises from a part of the limb.

Of additional significance is the arrangement of cephalic apodemes. The Crustacea possess

no common basic plan, while in the myriapods and hexapods the apodemes are arranged in

an intersegmental series that is similar in the two groups - although the posterior tentorial arm

is absent in the myriapods. On this thesis the relationship between Crustacea and Insecta is a

distant one. For this reason I believe that study of evolution of the arthropod head should be

done independently in the three major groups in the hope that such studies will indicate the

extent of phylogenetic relationships or the extent of convergent evolution.

A second difficulty from studies of comparative morphology arises from the fact that these

studies are often based on structural features of the adult. Bearing in mind the great morpholog-

ical changes that many insects undergo in their ontogeny, such studies seem to be doomed from

Evolution of the Insect Head

9

the beginning. It is a study of a ‘finished’ product and this should make it suspect. Morpholo-

gists now generally agree that sutures in the larva and adult with the possible exception of the

postoccipital suture cannot be used in determining segmental boundaries (Chapman, 1969).

Further, in morphological studies too much emphasis has been placed on muscle arrangement

and muscle innervation. Muscles may be composite structures derived from more than one

mesodermal somite, and muscles can also traverse two or more segments before reaching in-

sertion. Some authors have emphasized muscle innervation in head segmentation, but motor

neurons do not invariably innervate muscles of their own segment and sensory neurons do not

invariably lead to the ganglion of their own segment. The above statement is based on the find-

ings of Horridge (1965) who has made an unusually detailed study of the nervous system of

invertebrates.

Because of the inadequate fossil record and the inadequacy of the morphological approach

in studies of head segmentation, I agree with Matsuda (1965) that “a concept of the constitu-

tion of the insect head should be based primarily on embryological data”. My views are based

mainly on studies of the embryonic development of Lytta viridana, a blister beetle. Behavioral

and physiological studies of this species were first undertaken by Dr. Norman Church, then a

member of the Federal Department of Agriculture at Lethbridge, Alberta. Later he continued

this work at Saskatoon and collaborated with me in the developmental studies.

The term segment is treated here in terms of a metamere which is defined by Snodgrass

(1935) as a body division of the embryo; an embryonic somite or primary body segment. This

definition is quite insufficient for our purpose. In defining a metamere four criteria are now

generally used, namely: the presence of a pair of mesodermal somites; a pair of appendages;

a pair of apodemes, and a neuromere. It must be understood that during the long period of

evolution a component may be greatly reduced or even be lost. A controversy has arisen over

the relative importance of the different components. Some authors have emphasized coelomic

sacs, while others have minimized them. Some have stressed innervation and muscle arrange-

ment. But the chief difficulty has arisen because of the frequent neglect of embryological

studies.

The controversy in regard to head segmentation in arthropods concerns exclusively possible

pregnathal segments, namely intercalary, antennal, and preantennal. There has never been any

disagreement with regard to the status of the mandibular, maxillary, and labial segments. The

total number of segments has been placed as low as three (with no pregnathal segments) and

as high as seven. There has also been disagreement regarding the presence or absence of an

acron (= prostomium).

A brief review of the theories that have been proposed from time to time will now be given.

The selection was made to demonstrate the extent of difference in thought. Detailed accounts

of the history of evolutionary thought regarding head segmentation are extant and readily

available (Matsuda, 1965).

Theories of Head Segmentation

The Snodgrass Theory (Fig. 1). - Robert Evans Snodgrass was one of the world’s leading

insect morphologists and anatomists, a very productive researcher, a prominent teacher, an

artist and philosopher. He had a colorful and eventful career. For nearly a quarter of a century

he held a dual post in the Federal Bureau of Entomology and lecturer in entomology at the

University of Maryland (Mallis, 1971).

A biographer (Thurman, 1959) has this to say of Snodgrass. He was a “dignified, erect,

gracious unassuming gentleman. . .”. A man with “a phenomenal memory for facts and events,

a wealth of basic knowledge at his ready command, thorough training in the use of the Classical

and Romance languages, and an unlimited vocabulary in English and German. . .”.

Quaest. Ent, 1975, 11 (1)

10

Rempel

Snodgrass (1935, 1960) adopted the theory originally proposed by two European workers,

Holmgren (1916) and Hanstrom (1928). In accordance with this theory the insect head con-

sists of an acron and four metameres. The acron is large and encompasses the ocular, labral,

and antennal regions. The labrum is believed to be a mere outgrowth over the stomodaeum.

It is not appendiculate and there is no labral segment. The presence of labral coelomic sacs

recorded in some forms is not believed to indicate a segment. The antennae are not homolo-

gous with other appendages, and the deutocerebrum is not a separate ganglion but with the

protocerebrum constitutes the primitive brain, the archicerebrum.

The theory has had widespread acceptance in North America, but it has been criticized by

European workers, notably Manton (1949) and Slewing (1963) and others. Although generally

considered to be the leading North American insect morphologist, Snodgrass repeatedly be-

trayed unfamiliarity with insect embryology. Thus he assigned the origin of segmentation

phylogenetically to the ectoderm when developmental studies have clearly shown that the

mesoderm segments before the ectoderm and thus sets the pattern of segmentation. His claim

that coelomic sacs of the acronal region (as defined by him) are best developed in higher arthro-

pods is contradicted by embryology. Thus some coleopteran embryos have well developed

coelomic sacs in the head (Ullmann, 1964; Rempel and Church, 1969; Church and Rempel,

1971), while they are totally absent from dipteran embryos. His denial of appendiculate status

to the antennae must be rejected by every modern insect embryologist. The antennae arise in

the embryo as appendage-like outgrowths; they are provided with large coelomic sacs, apodemal

invaginations are present and the deutocerebrum is the neuromere. Although many workers in

the past, following Snodgrass, have denied segmental status to the labrum, recent studies pro-

vide ample evidence that the labrum represents the fused appendages of a labral segment (see

below).

The DuPorte Theory (Fig. 2). — Dr. Ernest Melville DuPorte rightfully deserves the title

Dean of Canadian entomology. But his reputation flows beyond the border of this country

and we can refer to him as one of North America’s leading insect morphologists and teachers

of entomology. Dr. DuPorte was bom in the West Indies in 1891 and in 1910 came to Canada

to enrol in entomology at McGill University. Upon receiving the Ph.D. he joined the faculty of

McGill and served that institution with great distinction until his retirement in 1957. At the

time of his retirement it was said that half of Canada’s practicing entomologists had at one

time or another studied under Dr. DuPorte. On a more personal basis one might say that this

outstanding researcher and teacher is noted for the warmth of his personality and fondness

for his students. It is no wonder that he enjoys the high esteem and affection of all. Dr. DuPorte,

now in his 80’s, is presently completing a textbook on insect morphology. One might well ask

“Why has there not appeared ere now an extended biography of this outstanding Canadian

biologist?”

The DuPorte theory is an extension of the Snodgrass theory and suffers from the same weak-

ness. Embryological evidence is underrated and coelomic sacs are not regarded as a valid criter-

ion of metamerism. The acron is large bearing the eyes, the antennae and the labrum. That is,

the preoral region is entirely unsegmented. Like Ferris he does not recognize an intercalary

segment. But embryological evidence contradicts DuPorte’s theory. The presence of an inter-

calary and antennal segment has been established beyond doubt by a wealth of developmental

studies.

The Butt Theory (Fig. 3). — Dr. F. H. Butt was for many years Professor of Insect Morphology

and Embryology at Cornell University. He is co-author (with O. A. Johannsen) of the Textbook

of Embryology of Insects and Myriapods. Dr. Butt, now retired at Friday Harbor on the Pacific,

was for many years interested in the segmentation of the insect head and published several pa-

pers on the subject. With Snodgrass he identified the acron as the region that bears the compound

Evolution of the Insect Head

11

eyes and the antennae, but excludes the labrum. The latter is looked upon as the fused labral

appendages, and since the labrum is innervated from the tritocerebrum, it is considered to be

part of the premandibular or intercalary segment. The antennae are not considered to be ap-

pendicular and the deutocerebrum is not a segmental ganglion but with the protocerebrum

corresponds to the archicerebrum of the annelid ancestors. The theory has been criticized by

Snodgrass ( 1 960) on the ground that the labral nerve is largely if not entirely a sensory nerve

and thus cannot be used in determining segmentation. The theory has also received strong

criticism by Manton (1960), a criticism which I believe to be intemperate. To settle the dis-

pute we need further detailed studies of the nerve pathways in the labral nerve and the nervus

connectivus.

The Ferris Theory (Fig. 4). - Gordon Floyd Ferris was born in Kansas in 1893. Early in

his life he moved west and was for nearly a half a century associated with Stanford University,

first as a student 1912-1917, and then as a faculty member 1917-1958. As professor he had

a distinguished career (Mallis, 1971). According to one biographer he was one of the giants

of taxonomic entomology in this century and this is borne out by his publications which

number 217. He was also an accomplished teacher of entomology and a field naturalist. Ferris

was an independent and forceful personality; he stated what was on his mind in energetic lang-

uage. Students held him in awe, but were attracted to him and inspired by him. They came

from various states outside California and from foreign lands. But in a university an overpow-

ering force can exert a negative influence. And so it was with Ferris. The publications of many

of his students betray too strong a Ferris stamp.

Ferris’ theory of head segmentation is most unusual. It is based on two main assumptions,

namely that the principal transverse sutures of the adult are intersegmental lines marking the

limits of the labral, clypeal, oculo-antennal, mandibular, maxillary, and labial segments, and

that a ganglion innervates a segment to which it originally belonged. The theory is based large-

ly on the findings of one of his co-workers. Miss Henry. Since the theory is based largely on

innervation, a difficulty is encountered in that the labrum is innervated from the tritocerebrum.

A ready explanation is found by assuming that the tritocerebral ganglion has phylogenetically

been shifted posteriorly. There is no room for the intercalary segment which Ferris claims was

‘invented’. A weakness of the theory is that it is based on features of the adult insect; on the

belief that the nervous system can be used to determine homology although no attempt is made

to trace nerve pathways. But the greatest drawback is the total disregard of embryological

evidence. The theory is now merely a historical curiosity that has no further place in the

entomological literature.

The Sharov Theory (Fig. 5). — One of the leading Russian authorities to take part in the

controversy of head segmentation was A. G. Sharov. A brief outline of his biography indicates

a remarkable career. He was bom near Moscow in 1922. At the age of 17 he entered the Uni-

versity of Moscow, but his education was interrupted by the war. Upon graduation in 1950

he joined the Institute of Animal Morphology and later transferred to the Paleontological

Institute, Academy of Sciences. Here he rose quickly from instmctor to senior scientist. He

was the leader of a series of expeditions to Asiatic Russia to collect vertebrate and invertebrate

fossils. As a scientist he was very productive and in his short life published 67 scientific papers

and monographs. Of added interest is that he appeared to have complete mastery of English.

Sharov believes in the monophyletic origin of the arthropods and therefore maintains that

a theory of head segmentation must fit the entire phylum. He postulates a seven-segmented

head. The first or anteriormost segment is the labral, and the labmm is appendiculate. This is

followed by the ocular or preantennal segment. The compound eyes are considered to be mod-

ified limbs. Then follow the segments in the usual way. Sharov’s book Basic Arthropodan Stock

in which he outlines his theory has been severely criticized by Hedgpeth ( 1 967) who calls the

Quaest. Ent., 1975, 11 (1)

12

Rempel

book an “unnecessary and misleading contribution.”

The Roonwal Theory (Fig. 6). — M. L. Roonwal published his theory of head segmentation in

1938. He postulates a seven segmented insect head. The acron, and with it the archicerebrum,

has disappeared. The first segment is the labral and its neuromere is part of the protocerebrum.

It is followed by the preantennal, with a second part of the protocerebrum as the neuromere.

The protocerebum is thus composed of two pairs of ganglia. Then follow in succession the

antennal, intercalary, mandibular, maxillary and labial. Roonwal is aware that the labrum is

innervated from the tritocerebrum and not from the protocerebrum, but considers this to be

a secondary feature. He thus seems to anticipate the view expressed later by Scholl (1969).

The Chaudonneret Theory (Fig. 7). — Jean Chaudonneret is a morphologist and embryolo-

gist of note. He was born in Dijon, France and received his education at the Universities of

Dijon and Paris. He has been associated with the University of Dijon since 1943. His publica-

tions consist of a series of papers dealing with the morphology and embryology of the insect

head and the gnathal region of the Malacostraca.

According to Chaudonneret (1966) the acron is small, well hidden. The first segment is the

protocephalic or preantennal bearing the eyes. This is followed in succession by the antennal,

tritocerebral, superlingual, mandibular, maxillary and labial. The uniqueness of this theory is

the presence of the superlingual segment, the superlinguae being considered appendicular.

The labrum is composite in structure derived from several originally postoral segments. The

acron is overgrown by parts of these segments. Matsuda (1965) points out the weakness of

this theory in that the superlinguae cannot be appendages of a separate segment for according

to Silvestri (1933) they arise embryologically from the posterior region of the mandibular seg-

ment and the anterior region of the maxillary segment. Moreover, since Chaudonneret’s theory

is based mainly on a study of Thermobia, it is of interest to note that Larink (1970) found

no evidence for the presence of a superlingual segment in the embryonic development of

Lepisma.

The Matsuda Theory (Fig. 8). — Dr. Matsuda studied at Stanford under Ferris. He is present-

ly associated with the Federal Department of Agriculture in Ottawa. On the basis of his ex-

cellent publications in insect morphology, we must look upon Dr. Matsuda as one of the lead-

ing insect morphologists on this continent.

Dr. Matsuda’s (1965) theory of head segmentation is outlined in his book the “Morphology

and Evolution of the Insect Head”. I briefly summarize his views. The acron is large and bears

the eyes and labrum. The neuromere is the protocerebrum. There is no preantennal segment

and the bilobed labrum is not appendicular. The first definitive segment in insects is the an-

tennal, with the deutocerebrum as it neuromere. The antennae are appendicular. The head is

thus made up of an acron and five segments.

The Imms-Manton Theory (Fig. 9). — Angustus Daniel Imms is one of the best known Eng-

lish entomologists. His fame rests largely on his widely used General Textbook of Entomology

and he must be looked upon as being first and foremost a teacher, rather than a researcher.

There is no photograph of Imms in existence, but a sketch by an artist is in the possession of

Sir Vincent Wigglesworth who very kindly sent me a copy and a biography of Imms.

Imms was born in 1 880. He received his early education, which was frequently interrupted

by ill health, in Birmingham. In 1903 he graduated from London and two years later entered

Cambridge as an 1851 Exhibition Science scholar. At Cambridge he came in contact with some

of the best entomologists of the day. He graduated in 1907 with a B.A. degree and was award-

ed the Darwin Prize of Christ’s College. In the same year Birmingham bestowed upon him the

D.Sc.

After graduation Imms spent six years in India, first as professor of biology at the Univer-

sity of AUahabad and later as Forest Entomologist of the Government of India. For reasons

Evolution of the Insect Head

13

of health he returned to England and accepted a readership in Agricultural Entomology at

Manchester. During the war years 1914-1918 he tried repeatedly to enlist but was rejected

on account of ill health. In 1918 Imms moved to the Experimental Station at Rothamsted

and remained there until 1931. While at Rothamsted he wrote the General Textbook of Ent-

omology, the book being published in 1925. This immediately brought him widespread rec-

ognition at home and abroad. He travelled widely and in 1925 and again in 1928 made ex-

tensive tours in Canada and the United States. In 1929 he was elected a Fellow of the Royal

Society, and in 1931 accepted a readership in entomology at Cambridge. Here he established

a fine record as a teacher, but he also ‘produced some valuable memoirs bearing particularly

on morphological problems fundamental to questions of the ancestry of insects and phylo-

genetic relationships of the main orders’ (Thorpe, 1949). His theory of head segmentation in

insects appears in his textbook and I shall briefly outline his views as found in the 1957 re-

vised edition. But we must first introduce another British authority whose views are in close

agreement with those of Imms.

Sidnie M. Manton, the holder of an impressive list of degrees and honours:

M.A., Ph.C., Sc. D. (Cambridge)

Fellow, Royal Society of London

Hon. Doctor Filosophie (Univ. Lund)

Fellow, Linnean Society

Linnean Gold Medalist.

The above academic achievements speak clearly of a scientist and scholar of a high order.

Academically Dr. Manton has been associated with Kings College, with Queen Mary College,

and the British Museum of Natural History. She has to her credit numerous publications deal-

ing with the morphology, embryology and evolution of the Arthropoda, especially Crustacea

and Onychophora. She has an incisive mind, a great background of knowledge and a great

facility in writing. As a consequence the word ‘monumental’ can very well be ascribed to her

work. But she can be intolerant of the views of others and I believe it is in order to call to

Dr. Manton’s attention the words of the philosopher Sir Thomas Browne, written in his

Religio Medici some three centuries ago:

“I should never divide myself from any man upon the difference

of an opinion or be angry with his judgement for not agreeing

with me.”

Imms accepts the classical theory of head segmentation and recognizes an acron and six meta-

meres: preantennary, antennary, intercalary, mandibular, maxillary, and labial. He associates

the preantennary segment with the protocerebrum although the latter may also comprise the

archicerebrum or primitive prostomial ganglion. The labrum is simply an unpaired sclerite over-

hanging the mouth. It is not appendiculate, and there is no labral segment.

Manton’s views are in close accord with those of Imms, and we need not elaborate further.

The Eastham Theory (Fig. 10). — Dr. Eastham is associated with the Department of Zoology,

Cambridge University. His theory is based on a detailed study of the embryology of Pieris rapae,

the cabbage butterfly, published in 1930.

In the early embryo Eastham made the important observation that there are six pairs of ap-

pendages in the regions that ultimately form the head tagma. The first pair fuse early and

develop into the labrum; the second pair are large and become the antennae; the third pair are

rudimentary. The fourth, fifth and sixth pairs form the mandibles, the maxillae and labium

respectively. Of special significance are the following. The labrum arises as two appendages

and each is provided with a somite. The protocerebrum is considered to be the neuromere.

Of interest too are the rudimentary appendages of the tritocerebral segment. Their presence

clearly indicates that the labrum cannot be the appendage of this segment. According to

Quaest. Ent, 1975, 11 (1)

14

Rempel

Eastham then, the insect head is made up of six segments, which is in agreement with the

classical theory of head segmentation. It differs from the view of others in that he omits the

acron. The ocular region is thus part of the labral segment. Eastham must be accorded the

credit for calling attention to the importance of the cephalic apodemes as an indicator of the

segmental constitution of the embryonic head.

The Tiegs and Weber Theory (Fig. 1 1). — In November 1956 the entomological world lost

two of its most illustrious members. The first was Oscar Werner Tiegs. The second was Hermann

Weber. Tiegs was bom in Brisbane, Australia, on March 12, 1897. His parents were of German

origin and had migrated to Australia at an early age. When still quite young he showed a great

interest in insects which seemed to foreshadow the remarkable contribution he was later to

make in entomology. Educated at the University of Queensland and Adelaide University he

joined the faculty of the University of Melbourne in 1925 and served that institution until

his death .

Tiegs’ early work involved the study of striated muscle in both vertebrates and invertebrates,

and in these studies he was far ahead of his times. One wonders what this man would have

achieved had he been able to draw on modern knowledge of biochemistry and had he had at

his disposal modern equipment. In the field of entomology he made truly outstanding con-

tributions in the embryology and phylogeny of the Myriapoda and Insecta. These studies

were undertaken in the hope that they would throw light on the evolution and stmctural

relationships of the Arthropoda. His work is characterized by attention to minute detail which

gives it lasting value. He achieved international acclaim, especially in Britain where formal rec-

ognition of his work was to be accorded him at the Darwin celebrations in 1958. He died un-

fortunately in 1956 at the age of 59.

Herman Weber was the ‘pride’ of German entomology and zoology. But his fame extended

far beyond the borders of his homeland, and other countries vied with each other to honour

this illustrious man. His outstanding scientific achievements were recognized by academicians

in German universities generally and there was keen competition for his services — Bonn, Dan-

zig, Freiburg, Munster, Vienna, Strassburg, Tubingen. During the war he suffered internment

and this seriously undermined his health which cut short his career at the age of 57. Weber

made brilliant contributions in three spheres. He was the author of numerous outstanding

scientific papers; he was involved in the editorship of prestigious scientific journals such as

Zoologische Jahrbucher, Fortschritte der Zoologie and others; he was the author of several

textbooks. His Lehrbuch der Entomologie was written in one year and it is now regarded as

a rare jewel in zoological literature. I consider his Grundriss der Insektenkunde written in

1938 still as the finest textbook of entomology. Unfortunately to Canadian students with

knowledge limited to English or French it is a closed treasure.

Tiegs’ (1941, 1947) theory of head segmentation is based largely on his detailed studies of

the embryology of Symphyla and Pauropoda. Weber (1952) made a lengthly review of the

various theories of head segmentation and his views largely coincide with those of Tiegs. We

may summarize these as follows:

The acron is homologous with the annelid prostomium. It bears the archicerebrum which

originally was made up of a small median unpaired nervous element. To this were later added

a pair of ganglia which form the larger part of the archicerebrum and innervate the eyes. The

acron is followed by 6 segments. The first is the preantennal segment, or prosocephalon. Its

neuromere is the prosocerebrum (Nebenlappen; Accessory Lobe). It is closely associated with

the archicerebrum to form what is commonly called the protocerebrum. The second metamere

is the antennal. Its appendages, the antennae, are innervated by the deutocerebrum. This is

followed by the premandibular or intercalary segment with the tritocerebrum as its neuromere.

The labrum, according to this theory, is composite in structure and is the product of the three

Evolution of the Insect Head

15

preoral segments. There is thus no labral segment. The statement is significant in view of what

many other researchers believe.

In 1 963 Rolf Slewing of the Zoological Institute of the Christian-Albrechts University at

Kiel made an extensive analysis of the problem of head segmentation in arthropods based on

a review of the literature. His conclusions are in rather close agreement with those of Tiegs

and Weber. He too concludes that there is an acron and six metameres, namely: preantennal,

antennal, premandibular, mandibular, maxillary and labial. The labrum is considered to be

solely part of the preantennal segment but he is not prepared to assign it appendiculate status.

The Scholl, Rempel and Church Theory (Fig. 12). — During the last few years Dr. Church

and I have made a rather detailed study of the embryonic development of Lytta viridana. Our

studies completely confirm the findings of Tiegs, Weber and Slewing. But we go a step further.

We find that the labrum must be considered appendiculate. We conclude that the insect head

consists of the annelid prostomium (referred to as the acron) and six metameres, the anterior-

most three having assumed a preoral position.

I now want to outline the proof for our theory. May I point out again that there has never

been any disagreement regarding the status of the mandibular, maxillary and labial segments.

I therefore limit the discussion to the status of the premandibular or intercalary segment, the

antennal segment, and the preantennal or labral segment.

First the intercalary segment. Its existence has been denied by only two workers, DuPorte

and Ferris. Ferris called the segment an ‘invention’. What does embryology show? In 60 h

embryos of Lytta, parasagittal section (Fig. 14), the intercalary segment is as distinct as the

succeeding segments. Note the coelomic sac. At 70 h (Fig. 15) a neuromere (tritocerebum)

is formed; at 80 h (Fig. 16) the apodemal invagination (the anterior tentorial arm) is promi-

nent. At this stage the tritocerebrum has joined the brain. Eastham (1930) (Fig. 13) recorded

rudimentary premandibular appendages. Thus the four criteria of a metamere are met.

What about the antennal segment? Here the North American workers (Snodgrass, Ferris,

DuPorte and Butt) are in a separate class from the European workers in that they reject ap-

pendiculate status for the antennae and deny the presence of an antennal segment. Let us

recall that we have four criteria for a segment. In an early embryo, a whole mount with all

embryonic membranes removed, appendage-like outgrowths are found from the labral to the

first abdominal segment (Fig. 1 3). Note that the largest among them are the antennae. In other

words, the latter arise like all other appendages. The statement has been made that the antennae

in structure differ markedly from the other appendages. In reply we may point to the great

plasticity of the arthropod appendage. It can be changed in various ways. For example, in

Lytta the appendages of the first abdominal segment change into glands, the pleuropodia.

That antennae generally have large coelomic sacs is well known from embryology (Fig. 16).

That satisfies two criteria. Antennal apodemes have been reported by a number of workers.

We observed them in an early embryo of Lytta (Rempel and Church, 1971, Figs. 2, 7, 8). In

this species, these apodemes extend backward into the mandible (Fig. 18). As the latter rotate

in a posteromedian direction through an angle of 120° (Fig. 19), the apodemes become located

at the outer edge of the mandible and later serve as support for the mandibular extensor mus-

cles. Our view of the ultimate fate of the antennal apodeme differs from that of Scholl (1969)

who maintains that it forms the dorsal arm of the tentorium.

That leaves us the problem of an antennal neuromere. The four leading American workers

mentioned above believe that the archicerebrum, the ancient brain, divided secondarily into

protocerebral and deutocerebral lobes. In other words, according to them the deutocerebrum

is not a segmental ganglion. This view is rejected by modern embryologists, for it is well known

that the deutocerebrum arises in the embryo apart from the protocerebrum. In the illustrations

(Figs. 20 and 21) the deutocerebrum with the sensory and motor neurons of the antennae are

Quaest. Ent., 1975, 11 (1)

16

Rempel

indicated.

That brings us to the final, the central problem^ the crucial point of the long-lasting contro-

versy. With few exceptions the entomologists of the world who have concerned themselves

with head segmentation look upon the labrum as merely an outgrowth of the body wall in

front of the stomodaeum. The view is well expressed by Snodgrass (1960) when he says, “it

seems much simpler to accept the labrum for what it appears to be in all arthropods from

trilobites to insects, namely, a preoral lobe of the head”. But you will recall that when I dis-

cussed theories of head segmentation that Eastham as early as 1930 considered the labrum

the fused labral appendages and the protocerebrum their neuromere. Butt in 1957 and again

in 1960 advanced the theory that the labrum resulted from the fused appendages of the inter-

calary segment. Slewing in 1 963 accepted the presence of a preantennal or labral segment but

was not prepared to assign appendiculate status to the labrum. There are numerous references

in the literature to the early appearance of the labrum as two appendage-like outgrowths.

This is especially well seen in Pieris (Fig. 13), in Tenebrio (Ullmann, 1964) and in Lytta (Rem-

pel and Church, 1969). That each appendage may have a somite laid down in situ is well docu-

mented. In 1969 Scholl made a detailed study of the embryonic development of the head of

the stick insect. He set out to investigate first whether there is a distinct region between the

acron and the antennal segment which can be demarcated from the two, and which has not

been translated there secondarily, and, second, whether one can assign segmental status to this

area. On the bases of the development of the mesoderm Scholl could delimit three preoral

regions: labral, antennal and premandibular. He is inclined to look upon the labrum as an

appendicular structure.

Reference has been made to the intersegmental arrangement of cephalic apodemes. In 1965

Matsuda made a careful survey of the literature in regard to cephalic apodemes. His findings

are summarized in Fig. 17. According to Matsuda no one had ever found apodemes in associa-

tion with the labrum. But in 1971 we discovered these apodemes in early Lytta embryos

(Figs. 18, 19). Also Scholl (1969) lists the following cephalic apodemes: labral (Frontalleisten?),

antennal, intercalary, mandibular, maxillary and labial. Scholl believes that the arrangement

of the apodemes can be used as a criterion for head segmentation if used in association with

other criteria.

This leaves us the last hurdle. If there is a labral segment, what is its neuromere and what

is its innervation? If the labral segment is a preantennal segment, then we must look for its

neuromere in front of the deutocerebrum and for a nerve that leads from here to the clypeo-

labral muscles. According to Scholl ( 1 969) the corpora pedunculata and the optic centres are

common to the Annelida and Arthropoda. But the central body and the protocerebral bridge

are found in arthropods only. Hence we may look upon the optic centres and the corpora

pedunculata as the archicerebrum. The rest of the protocerebrum may then represent the

preantennal neuromere, or the prosocerebrum. This is in agreement with the view of Larink

(1970) and the view of Malzacher (1968) who lists the following sequence:

Archicerebrum — Optic ganglia

corpora pedunculata

Preantennal neuromere - Neurosecretory cells of the pars intercerebralis

Protocerebral bridge

Central body

Accessory lobe (Nebenlappen)

Antennal neuromere — Deutocerebrum

Intercalary neuromere — Tritocerebrum

I have illustrated this diagrammatically in Fig. 20.

In many orders of the Hemimetabola innervation of the clypeolabral muscles is via the

Evolution of the Insect Head

17

nervus connectivus, the frontal ganglion, and the nervus procurrens. Scholl ( 1 969) believes

that this was the original mode of innervation and that innervation via the labral nerve is a

secondary acquisition. I have illustrated this in Fig. 21.

I now want to summarize my argument. The ancestors of the insects were annelid-like.

The prostomium housed the archicerebrum, or primitive brain. All other ganglia were postoral

(Fig. 22). The next forward step involved the acquisition of a haemocoel, a cuticle, segment-

al appendages, and apodemes for improved muscle attachment (Fig. 23). Over a period of

milhons of years cephalization involved a progressive shift of ganglia forward into a preoral

position. The first one to join the archicerebrum was the labral (Fig. 24). While the archi-

cerebrum began to shift dorsad and caudad, the labrum (a pair of fused appendages) shifted

caudad to form a preoral cavity. We observe this phenomenon today in embryonic develop-

ment. The labral nerve is the nervus connectivus. The next ganglion to shift into a preoral

position is the antennal (Fig. 25). Finally the tritocerebral ganglion did phylogenetically what

it does today ontogenetically. It too shifts from a postoral into a preoral position. The man-

dibular, maxillary and labial ganglia move forward, fuse, but retain their postoral position.

The insect head thus consists of an acron and six metameres (Fig. 26). The appendages of

the first metamere fuse to form the labrum; the appendages of the second become the anten-

nae; those of the third are lost, although occasionally present in the early embryo. The ap-

pendages of the fourth, fifth and sixth segments develop into the mandibles, maxillae and

labium respectively.

The theory proposed here is in line with the classical theory of a six-segmented insect head,

and I beheve it is in close agreement with the facts as we know them today.

ACKNOWFEDGEMENTS

I want to thank the following for providing me with biographical material of authorities

discussed in this paper: Sir Vincent Wigglesworth and Dr. John Smart, Cambridge, England

(Imms); Dr. F. H. Drummond, University of Melbourne, Parkville, Australia (Tiegs); Dr. G.

Mickoleit, Tubingen, Germany (Weber); Professor B. Rohdendorf, Academy of Sciences,

Moscow, U.S.S.R. (Sharov); Professor J. Chaudonneret, Dijon, France; Professor F. O. Mor-

rison, McGill, Montreal (DuPorte). J. S. Scott, chief technician. Department of Entomology,

University of Alberta, provided expert assistance in the tedious process of preparing the il-

lustrations for reproduction. Publication costs were met by grants from the Strickland Memo-

rial Trust Fund and the National Research Council of Canada.

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Butt, F. H. 1957. The role of the premandibular or intercalary segment in head segmentation

of insects and other arthropods. Transactions of the American Entomological Society 83:

1-30.

Butt, F. H. 1960. Head development in the arthropods. Biological Reviews, Cambridge 35:

43-91.

Chapman, R. F. 1969. The Insects Structure and Function. The English Universities Press,

London.

Chaudonneret, J. 1 966. La construction phylogenetique de la tete des Insectes. 1 . Le Squelette.

Bulletin Scientifique de Bourgogne. 24: 241-263.

Church, N. S. and J. G. Rempel. 1971. The embryology of Lytta viridana Le Conte (Coleop-

tera: Meloidae). VI. The appendiculate 72-h embryo. Canadian Journal of Zoology 49 (12):

1563-1570.

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Rempel

Demerec, M. 1950. Biology of Drosophila. John Wiley & Sons, New York.

Dixit, P. N. 1972. Morphology of the Arthropod Labrum and its Bearing on the Head Problem.

Unpublished Thesis for Ph.D., Faculty of Science, Nagpur University.

Eastham, L. E. S. 1930. The Embryology of Pieris rapae. Organogeny. Philosophical Trans-

actions of the Royal Society of London, B. 219: 1-50.

Ferris, G. F. 1950. In Biology of Drosophila by Demerec. John Wiley & Sons, New York.

Hanstrom, B. 1928. Vergleichende Anatomie des Nervensystems der wirbellosen Tiere unter

besonderer Beriicksichtigung seiner Funktion. Springer, Berlin, 628SS.

Hedgpeth, J. W. 1967. New Biological Books. Nature, March 1967, p. 69.

Holmgren, N. 1916. Zur vergleichenden Anatomie des Gehirns von Polychaeten, Onychophoren,

Xiphosuren, Arachniden, Crustacean, Myriapoden und Insekten. Kunglica Svenska Veten-

skapsakademiens Handlingar, Uppsala, Stockholm 56: 1-303.

Horridge, G. A. 1965. Structure and Function in the Nervous Systems of Invertebrates. Vol.

II. W. H. Freeman & Co. San Francisco & London.

Imms, A. D. 1957. A General Textbook of Entomology. Ninth Edition, Methuen & Co.,

London.

Larink, O. 1970. Die Kopfentwicklung von Lepisma saccharina L. (Insecta, Thysanura).

Zeitschrift fuer Morphologie der Tiere 67: 1-15.

Malhs, A. 1971. American Entomologists. Rutgers University Press, New Brunswick, New

Jersey.

Malzacher, P. 1968. Die Embryogenese des Gehirns paurometaboler Insekten. Untersuchungen

an Carausius morosus und Periplaneta americana. Zeitschrift fuer Morphologie der Tiere

62: 103-161.

Manton, S. M. 1 949. Studies on the Onychophora VII. The early stages of Peripatopsis, and

some general considerations concerning the morphology and phylogeny of the Arthropoda.

Philosophical Transactions of the Royal Society of London, B. 233: 483-580.

Manton, S. M. 1960. Concerning head development in the arthropods. Biological Reviews 35:

265-282.

Manton, S. M. 1964. Mandibular mechanisms and evolution of arthropods. Philosophical

Transactions of the Royal Society of London, B. 247: 1-183.

Matsuda, R. 1965. Morphology and evolution of the insect head. Memoirs of the American

Entomological Institute 4, Ann Arbor, Michigan.

Rempel, J. G. and N. S. Church. 1969. The embryology of Lytta viridana Le Conte (Coleop-

tera, Meloidae). V. The blastoderm, germ layers, and body segments. Canadian Journal of

Zoology 47 (6): 1157-1171.

Rempel, J. G. and N. S. Church. 1971. The embryology of Lytta viridana Le Conte (Coleop-

tera: Meloidae). VII. Eighty-eight to 132 h: the appendages, the cephalic apodemes, and

head segmentation. Canadian Journal of Zoology 49 (12): 1571-1581.

Romoser, W. S. 1973. The science of entomology. Macmillan Publ. Co., N. Y.

Roonwal, M. L. 1938. Some recent advances in insect embryology, with a complete biblio-

graphy of the subject. Journal of the Asiatic Society of Bengal, Science 4: 17-105.

Scholl, G. 1969. Die Embry onalentiwicklung des Kopfes und Prothorax von Carausius morosus

Br. (Insecta, Phasmida) 2. Zeitschrift fuer Morphologie der Tiere 65: 1-142.

Scudder, G. G. E. 1973. Recent advances in the higher systematics and phylogenetic concepts

in entomology. Canadian Entomologist 105 (9): 1251-1263.

Sharov, A. G. 1966. Basic arthropodan stock. Pergamon Press, Oxford.

Siewing, R. 1963. Zum Problem der Arthropodenkopfsegmentierung. Zoologischer Anzeiger

170: 429-468.

Silvestri, F. 1933. Sulle appendici del capo degli “Japygidae” (Thysanura Entotropha) e rispet-

Evolution of the Insect Head

19

tivo confronto con quelle dei Chilopodi, dei Diplopodi e dei Crostacei. Travaux 5® Congress

International Entomologique, Paris (1932). p. 329-343. Not seen. As quoted by Matsuda.

Smart, J. and N. F. Hughes. 1972. The insect and the plant: progressive palaeoecological in-

tegration. Symposium on insect/plant relationships of the Royal Entomological Society

of London. Number six. Blackwell Scientific Publication, Oxford.

Snodgrass, R. E. 1935. Principles of Insect Morphology. McGraw-Hill, N. Y.

Snodgrass, R. E. 1960. Facts and theories concerning the insect head. Smithsonian Miscel-

laneous Collections 142 (1): 1-61.

Steinmann, H. 1970. A comparative Anatomy of the Insect Head. I. Muscles and Nerves

of the Regio verticalis, R. antennalis and R. labralis. Folia Entomologica Hungarica 23 (3):

113-124.

Thorpe, W. H. 1949. Dr. A. D. Imms, F. R. S. Nature, Volume 163, p. 712, May 7, 1949.

Thurman, E. B. 1959. Robert Evans Snodgrass, Insect Anatomist and Morphologist. Smith-

sonian Miscellaneous Collections 137: 1-17.

Tiegs, O. W. 1941. The embryology and affinities of Symphila, based on a study of Hansen-

iella agilis. Quarterly Journal of Microscopical Science 82: 1-125.

Tiegs, O. W. 1947. The development and affinities of the Pauropoda, based on a study of

Pauropus silvaticus. Quarterly Journal of Microscopical Science 88: 165-267; 275-336.

Tiegs, O. W. and S. M. Manton. 1958. The evolution of the Arthropoda. Biological Reviews

33: 255-337.

Ullmann, S. L. 1964. The origin and structure of the mesoderm and the formation of coelomic

sacs in Tenebrio molitor L. (Insecta, Coleoptera). Philosophical Transactions of the Royal

Society of London, B. 747, 248: 245-277.

Weber, H. 1952. Morphologie, Histologie und Entwicklungsgeschichte der Articulaten. II.

Die Kopfsegmentierung und die Morphologie des Kopfes iiberhaupt. Fortschritte der

Zoologie 9: 18-231.

Wigglesworth, V. B. 1949. Angustus Daniel Imms 1880-1949. Obituary Notices of Fellows

of the Royal Society. Volume 6, November 1949.

Quaest. Ent, 1975, 1 1 (1)

20

Rempel

FIGURE LEGENDS

Fig. 1. The theory of Head Segmentation by Snodgrass.

Fig. 2. The DuPorte Theory.

Fig. 3. The Butt Theory.

Fig. 4. The Ferris Theory.

Fig. 5. The Sharov Theory.

Fig. 6. The Roonwal Theory.

Fig. 7. The Chaudonneret Theory.

Fig. 8. The Matsuda Theory.

Fig. 9. The Imms-Manton Theory.

Fig. 1 0. The Eastham Theory.

Fig. 11. The Tiegs-Weber Theory.

Fig. 1 2. The Scholl-Rempel-Church Theory.

Fig. 13. The embryo to show appendages and somites. Redrawn from Eastham, 1930, modified.

Fig. 14. Parasagittal section of 60h embryo of Lytta viridana to show coelomic sacs. Note

intercalary segment.

Fig. 1 5. Parasagittal section of 70h embryo to show formation of ganglia. Note tritocerebral

ganglion.

Fig. 16. Parasagittal section of 80h embryo. Note: tritocerebrum as part of the brain; antennal

coelomic sac; intercalary apodeme or anterior tentorial arm; maxillary apodeme or

posterior tentorial arm; suboesophageal ganglion.

Fig. 17. Hypothetical arrangement of cephalic apodemes according to Matsuda, 1965.

Fig. 1 8. Diagram to show arrangement of cephalic apodemes in embryo of Lytta viridana.

Fig. 1 9. Diagram to show tentorium, labral apodemes, antennal apodemes (mandibular ex-

tensor apodeme), mandibular apodeme (mandibular flexor apodeme), labial apo-

demes (labial diverticula). 132h embryo of Lytta viridana.

Fig. 20. Diagram to show archicerebrum and segmental cephalic ganglia and important

cephalic nerves, ventral view.

Fig. 21. As in Fig. 20, lateral view.

Fig. 22. A. Annelid-like ancestor of insects to show postoral arrangement of future cephalic

ganglia. Note prostomium with archicerebrum. B. As above, dorsal view of archi-

cerebrum and nerve cord.

Fig. 23. Same as Fig. 22 A, but with cuticle, segmental appendages, and apodemes.

Fig. 24. A. Hypothetical arthropod in which the first ganglion has shifted into a preoral

position to form the prosocerebrum, ganglion of the labral segment. B. As above,

dorsal view of brain and nerve cord.

Fig. 25. A. Arthropod with primary syncerebrum (Weber) (= archicerebrum, prosocerebrum,

deutocerebrum). B. As above, dorsal view.

Fig. 26. A. Insect head. The tritocerebrum (ganglion of intercalary segment) has moved into

a preoral position to become part of the brain. The ganglia of the gnathal segments

fuse to form the suboesophageal ganglion. B. As above, dorsal view.

Evolution of the Insect Head

21

THEORIES OF HEAD SEGMENTATION IN INSECTS

Snodgrass, 1960

HEAD OF INSECT

Acron + 4 segments

1- intercalary

2- mandibular

3- maxillary

4- labial

Labrum and Antenna

not appendiculate,

part of acron

Du Porte, 1963

Butt, 1960

Ferris, 1950

HEAD OF INSECT

Acron + 4 segments

1- intercalary

2- mandibular

3- maxillary

4- labial

Labrum appendiculate,

part of intercalary segment

Antenna not appendiculate, part of acron

HEAD OF INSECT

Acron absent.

6 segments

1- labral

2- clypeal

3- oculo- antennal

4- mandibular

5- maxiliary

6- labial

intercalary absent

Sharov, 1966

Roonwal, 1938

HEAD OF INSECT

Acron absent.

7 segments

1- labral

2- preantennal (ocular)

3- antenna I

4- intercalary

5- mandibular

6- maxillary

7- labial

HEAD OF INSECT

Acron absent,

7 segments

1- labral

2- preantennal

3- antenna I

4- intercalary

5- mandibular

6- maxillary

7- labial

Labrum appendiculate

Quaest. Ent., 1975, 11 (1)

22

Rempel

THEORIES OF HEAD SEGMENTATION IN INSECTS

Chaudonneret, 1966

HEAD OF INSECT

Acron small, hidden

7 segments

1- preantenna I

2- antenna I

3- intercalary

4- superlinguai

5- mandibular

6- maxillary

7- labia I

8

Matsuda, 1965

HEAD OF INSECT

Acron large,

5 segments

1- antennal

2- intercalary

3- mandibular

4- maxillary

5- labial

9 lmms,1957;Manton, I960, 1964 10

Eastham,1930

HEAD OF INSECT

Acron + 6 segments

1- preantenna I

2- antennal

3- intercalary

4- mandibular

5- maxillary

6- labial

HEAD OF INSECT

Acron absent,

6 segments

1- labral

2- antennal

3- intercalary

4- mandibular

5- maxillary

6- labia I

Labrum appendiculate,

Premandibular appendages present, vestigial

11 Tiegs, 1940, 1947; Weber, 1952

HEAD OF INSECT

Acron + 6 segments

1- preantennal

2- antennal

3- intercalary

4- mandibular

5- maxillary

6- labial

12 Scholl, 1969; Rempel & Church, 1971

HEAD OF INSECT

Labrum composite, hence not appendiculate

Labrum appendiculate

Evolution of the Insect Head

23

13

bbrum

14

15

16

17 Apodemes

18 Apodemes

19

Tentorium

antenna!

-intercalary

mandibular

maxillary-

labial

20

labral

intercalary

antennal

mandible

tentorium

-maxillary

labial diverticula

21

corpus pedunculatum

archicerebrum

prosocerebrum

deutocerebrum

tritocerebrum

archicerebrum

prosocerebrum

deutocerebrum

tritocerebrum

commissure

labral nerve

nervus connectivus

nervus recurrens

frontal ganglion

nervus procurrens

commissure

Quaest. Ent., 1975, 1 1 (1)

24

Rem pel

22

arc hi cere brum

proslomium ‘ ^

mouth

prosocerebrum appendage

2> A

apodeme

^ labrum

^ ^ apodeme anterior tentorial arm

deutocerebrum

antennal segment

/

nervus connectives

labral apodeme

ANTENNAL STRUCTURE AND METAMORPHOSIS IN

FRANKLINIELLA FUSCA (HINDS) (THRIPIDAE) AND

HAPLOTHRIPS VERBASCI (OSBORN) (PHLAEOTHRIPIDAE)

(THYSANOPTERA).

BRUCES. HEMING

Department of Entomology

University of Alberta Quaes tiones Entomologicae

Edmonton, Alberta T6G 2E3 11: 25 - 68 1975

Larval and adult antennae, in both Frankliniella fusca^^c? Haplothrips verbasci, consist of

seven and eight segments respectively. In adults of both species and in larvae o/F. fusca, the

antennae are raised and lowered by levator and depressor muscles inserting into the bases of

the scapi and originating on the anterior tentorial arms. Both these muscles, in larvae o/H.

verbasci, have additional branches originating on the vertex of the head.

Propupal antennae in F. fusca are short, weakly segmented and forward-directed; those of

pupae unsegmented and flexed dorsally over the head and prothorax. During the quiescent

stages, both the extrinsic and intrinsic antennal muscles maintain their myofibrils but cease

to function because they are no longer attached to cuticle. They increase slightly in diameter

and considerably in length, but are unchanged otherwise.

During the larva Il-propupal apolysis o/H. verbasci, all larval head muscles contract max-

imally; their myofibrils degenerating shortly thereafter. This contraction, plus changes in cell

shape probably cause the complete withdrawal of epidermis from within the larva II antennae.

At ecdysis, the propupal antennae are evaginated from epidermal pockets as short, unsegment-

ed stubs. In the two pupal stages, they remain unsegmented but lengthen posteriorly along

either side of the head. Imaginal segmentation and myofibrils begin to differentiate in the

exuvial pharate adult stage.

Sense organs are similar in type, position, and number in larvae of both species but are very

different in adults. No obvious sexual differences exist in the imaginal antennae of either

species except for their smaller size in males.

The Johnston s Organ, in larvae o/F. fusca, consists of three chordotonal organs having

two scolopidia each; in H. verbasci of four having two or three each. In both species, the

Johnstons Organ is carried through metamorphosis, two (H. verbasci) or three fF. fusca)

chordotonal organs and additional scolopidia being added to each during the quiescent stages.

Antennal structure and development in thrips is compared with that occurring in other

insects and an hypothesis is offered to explain the origin of the differences in antennal meta-

morphosis existing between the two species. It is concluded that the drastic events occurring

in H. verbasci other phlaeothripids have probably evolved in conjunction with the adoption,

by this family, of a primarily cryptophilous existence.

Les antennes de Frankliniella fusca et de /’Haplothrips verbasci aux stades larvaires et adultes

consistent respectivement de sept et huit articles. Chez les adultes de chaque espece et la larve

de F. fusca les antennes sont elevees et abaissees par les muscles levateurs et depresseurs. Ces

muscles sont attaches a la base des scapes antennaires et sur Vembranchement anterieur du

tentorium. Ces deux muscles chez la larve de H. verbasci ont des embranchements additionnels

allant sur le vertex de la tete.

Les antennes de la propupe de F. fusca sont courtes, faiblement articulees et dirigees anter-

ieurement; celles de la pupe ne sont pas articulees et sont orientees dorsalement au-dessus de

la tete et du prothorax. Lors des stades inactifs, les muscles intrinseques et extrinseques

des antennes maintiennent leurs myofibrilles mais cessent toutes fonctions car Us ne sont plus

26

Heming

attaches a la cuticule. Elies out un diametre legerement plus grand et sont tres allongees, mats

demeurent inchangees autrernent.

Lors de Vapolyse de la seconde larve-propupale de /’H. verbasci tons les muscles larvaires

de la tete se contractent au maximum; les myofibrilles degenerent peu apres. Cette contraction

avec un changement dans la forme des cellules epidermiques probablement provoque le de-

tachement complet de Vepiderme a Vinterieur de Vantenne de la seconde larve. Lors de la

metamorphose les antennes de la propupe apparaissent comme des petites projections de Vepi-

derme et nest pas articulees. Durant les deux stades de la pupe les antennes demeurent in-

articulees mais s’allongent posterieurement de chaque cote de la tete. Chez Vimago la segment-

ation et les myofibrilles commencent a se differencier lors de la formation de Vimago a Vinterieur

de la pupe (Pharate stage).

Les organes des sens sont similaires au point de vue de la forme, de la position et du nombre

chez les larves des deux especes mais Us sont differents chez les adultes. II n’y a aucune difference

sexuelle evidente entre les antennes des adultes des deux especes excepte qu’elles sont plus

court es chez les males.

L’organe de Johnston, chez les larves de F. fusca consiste de trois organes chordotonaux

pourvus chacun de deux scolopidia; chez cefles d’R. verbasci de quatre organes chordotonaux

ayant chacun deux ou trois scolopidia. L’organe de Johnston est preserve durant les metamor-

phoses, mais deux fH. verbascij ou trois (F. fuscaj organes chordotonaux et des scolopidia

additionels sont adjoutes durant les stades inactifs.

La structure et le development des antennes chez les thysanopteres sont compares aux autres

insectes et une hypothese est offerte pour expliquer Vorigine des differences observees entre les

deux especes lors de la metamorphose des antennes. Nous concluons que les evennements im-

portants se deroulant chez /’H. verbasci et les autres phlaeothripides ont probablement evolues

conjointement avec Vadoption d’un mode de vie principalement cryptophile chez cette famille.

CONTENTS

Introduction 26

Methods 28

Observations 28

Discussion 54

Acknowledgements 64

References 65

INTRODUCTION

The antennae of adult Thysanoptera are their best known structures because of their impor-

tance in taxonomy. This is due to the diversity of shapes, lengths and numbers of their seg-

ments and to the numbers and kinds of sense organs present on these segments in different

taxa (Doeksen, 1941 ; Priesner, 1960). The two larval stages have smaller antennae differing

structurally from those of the adults (Priesner, 1960), while those of the quiescent instars

(propupa and pupa in Terebrantia; propupa, pupa I and pupa II in Tubulifera) have reduced

segmentation or lack it completely and are flexed dorsally over (Terebrantia) or laterally be-

side (Tubulifera) the head (Priesner, 1960; Lewis, 1973).

Meiis (1934b), Risler (1957) and Mickoleit (1963) have described the structure and muscu-

lature of the adult antennae of several species of Aeolothripidae, Thripidae and Phlaeothripidae,

while Meiis (1934b), Davies (1969) and Haga (1974) have briefly discussed larval structure

and metamorphosis of the antennae in representative species of Phlaeothripidae and Thripidae.

Antennal Structure and Metamorphosis in Thysanoptera

27

Abbreviations Used in Figs. 1-87

Quaest. Ent., 1975, 1 1 (1)

28

Heming

The ultrastructure of the imaginal, antennal sense organs of Bagnalliella yuccae (Hinds) (Phlaeo-

thripidae) and Frankliniella tritici (Fitch) (Thripidae) has recently been described by Slifer and

Sekhon (1974) (reprint was received after this study was completed).

As part of a continuing study of metamorphosis in thrips (see Heming (1973) for a summary

of previous work), I here describe the structure and postembryogenesis of the antennae of two

species, each representing one of the thysanopteran suborders. Development is compared with

that occurring in other insects and an attempt is made to explain the origin of the differences

between terebrantian and tubuliferan antennal metamorphosis.

METHODS

Cleared and uncleared whole mounts (Heming, 1969) and serial sections (Heming, 1970,

1971) were prepared of all stages of both sexes of Frankliniella fusca (Hinds) (Thripidae) and

Haplothrips verbasci (Osborn) (Phlaeothripidae). Unmacerated whole mounts were studied

under phase contrast and polarized light to show internal details, while sense organs were ex-

amined with phase contrast in cleared specimens mounted in Hoyer’s medium (this medium

has optical properties superior to those of Canada balsam). Illustrations were drawn with the

aid of a drawing attachment through a Wild M20 microscope.

Specimens of all stages of H. verbasci were killed in hot water, washed in detergent, and

prepared for scanning electron microscopy using the critical point drying technique (Hearle

et.al., 1972, pp. 197-198). Observations on living//, verbasci were made at low and high (x500)

magnifications as previously described (Heming, 1971, 1972). Representatives of this species

were reared on potted mullein {Verbascum thapsus L.) in the laboratory.

I also examined Canada balsam whole mounts of specimens of all stages of Taeniothrips

simplex (Morison) (Thripidae), Selenothrips rubrocinctus (Girard) (Thripidae), Par theno thrips

dracaenae (Heeger) (Thripidae), Haplothrips graminis Hood (Phlaeothripidae), Gy naiko thrips

uzeli Zimmemiann (Phlaeothripidae), Haplothrips sp. (Phlaeothripidae) and Euoplothrips sp.

(Phlaeothripidae).

Illustrations in this paper are arranged by instar. Antennal postembryogenesis in F. fusca

is illustrated in Figs. 1-33 and in H verbasci in Figs. 34-87. Positions and angles of sections

are indicated by numbered arrows on drawings of the entire antenna. Arrows having numbers

in parentheses indicate that the sections in question were made from specimens at different

stages in development than were the whole mounts used to make the drawings. Those with

underlined numbers mark the positions of frontal sections.

OBSERVATIONS

Frankliniella fusca

Under summer conditions in South Carolina, F. fusca has a generation time of about 16.7

days with 6.7, 2.8, 3.5, 1.1 and 2.6 days spent, respectively in the egg, first and second larval,

propupal and pupal stages (Watts, 1934). The species is polyphagous and specializes on seed-

lings. Larvae congregate at the growing points of plants, adults there or in flowers and propupae

and pupae within the leaf sheaths (Heming, 1970) or in the soil (Eddy and Livingstone, 1931 ;

Newsom et.al., 1953).

Larval Stages ( Figs. 1-11).

Structure. — Antennae of the two larval stages are practically identical except for size (Figs.

1, 9). Each consists of a scape (scp.), a pedicel (ped.), and five flagellar segments (fl.) (usually

called subsegments because they lack muscles). The scape of each antenna is inserted into a

Antennal Structure and Metamorphosis in Thysanoptera

29

socket (ant. s.) in the head capsule and is articulated to it by means of weakly-developed lat-

eral and median condyles (antennifers) extending inwards from the socket rim (not shown in

figures because of their position). Large areas of membranous cuticle are present in each socket

dorsal and ventral to the scape, allowing each antenna to be raised and lowered by levator (lev.

ant.) and depressor (dep. ant.) muscles originating on the anterior tentorial arms (at.) of the

head capsule and inserting, respectively, into the dorsal and ventral margins of the scape base

(Figs. 6-8). Each of these muscles consists of three fibres (Fig. 8).

Figs. 1-5. F. fusca. Larval: Fig. 1. Right antenna, dorsal aspect. Fig. 2. Same, optical section. In this and many other illustra-

tions, the hatching represents epidermal and sensory ceUs. Fig. 3. Transverse section through pedicel and Johnston’s Organ

taken at point indicated by arrow in Fig. 1. Pharate larva II: Fig. 4. Right antenna, dorsal aspect, showing developing larva

II antenna (Note that most larva I sensilla omitted from drawing). Fig. 5. Transverse section through pedicel and Johnston’s

Organ taken at point indicated by arrow in Fig. 4.

The pedicel (ped.) articulates with the scape (scp.) by means of a single dorsal condyle

(Figs. 1, 9) in the apex of the latter and is flexed laterally and extended anteriorly by flexor

(fix. ant.) and extensor (ext. ant.) muscles within the scape (Figs. 2, 6, 10). These muscles

each consist of three fibres (Fig. 14), are inserted into the base of the pedicel on either side

of the condyle, and arise proximally on the lateral and medial walls of the scape. The narrow

Quaest. Ent., 1975, 11 (1)

30

Heming

ant. s.

Figs. 6-14. F. fusca. Larva II: Fig. 6. Ventral aspect of head, showing origins and insertions of intrinsic (fix. ant.; ext. ant.)

and extrinsic (lev. ant.; dep. ant.) antennal muscles (all setae have been omitted). Fig. 7. Oblique, parasagittal section taken

at point indicated by arrow in Fig. 6., showing origins and insertions of antennal levator (lev. ant.) and depressor (dep. ant.)

muscles. Fig. 8. Transverse section taken at point indicated by arrows in Figs. 6 and 7. Note the 3 fibres of each muscle.

Fig. 9. Right antenna, dorsal aspect. Fig. 10. Same, optical section. Fig. 11. Transverse section through pedicel taken at

point indicated by arrow in Fig. 9. Pharate Propupa: Fig. 12. Right antenna, dorsal aspect (Note: Most larva II sensilla omitted

from drawing). Fig. 13. Transverse section through pedicel taken at point indicated by arrow in Fig. 12. Fig. 14. Same, through

scape and antennal flexor (fix. ant.) and extensor (ext. ant.) muscles. Section taken at point indicated by arrow in Fig. 12.

Antennal Structure and Metamorphosis in Thysanoptera

31

base of the first flagellar segment (usually called the “pedicel” by thysanopterists) articulates

with the pedicel via dorsal and ventral condyles in the latter’s apex (Figs. 1, 9). Therefore, each

antenna has two points of flexion and extension: the scape-pedicellar and the pedicel-flagellar

joints.

Antennal segments three and four in both larval instars consist respectively of five and six

lightly-sclerotized rings of cuticle (Figs. 1, 9). In segment four, rings two to five each subtend

a whorl of delicate microtricheae (m. t.).

Sense Organs. — The antennal sense organs of larval F. fusca are illustrated in part in Figs.

1-3 and 9-1 1 and are listed in Table I for the second-stage larva. They are identical in larvae

of both sexes and instars except where indicated. Not all sensilla listed are illustrated because

they are either ventrally located or are too small to show up at the magnification of the draw-

ings.

Table I. Sense organs on the antenna of the larva II of F. fusca (N = 8).

Grand Total 42

The Johnston’s Organ (J. O.) in larvae of F. fusca (Figs. 2, 3, 5, 10, 11) consists of three

chordotonal organs each containing two scolopidia (sco. J. O.). Some of these appear to be

innervated by one and others by two sensory neurons according to whether they contain one

or two ciliary dilations (note dots inside each scolopale), but this requires verification by

transmission electron microscopy. The cap cells of each organ insert into the articular mem-

brane at the base of the first flagellar segment; the ligament of each arising basally on the walls

of the pedicel (ped.) (Figs. 2, 10). The cell bodies of the sensory neurons innervating the organ

(s. cl. J. O.) are situated basally within the pedicel with their axons joining the antennal nerve

(ant. nv.) proximally (Figs. 2, 10). This nerve continues posteriorly through the small larval

head and eventually enters the deutocerebrum of the brain, the latter being situated in the

prothorax.

The three apical sense organs (ap. s. o.) of antennal segment three (Figs. 1, 9) appear to

be universal in thrips as they are present in larvae and adults of both F. fusca and H. verbasci

Quaest. Enl, 1975, 11 (1)

32

Heming

(see below).

That portion of the antennal lumen occupied by epidermal and sensory cells (s. cl.) is shown

in optical section (by hatching) in Figs. 2 and 10. Details are difficult to observe, even with

phase contrast, because of the presence of obscuring yellow pigment granules within the epi-

dermal cells. The antennal nerve bifurcates at the base of the first flagellar segment, each

branch penetrating the basement membrane of the antennal epidermis laterally (Figs. 2, 10).

Each branch then continues distally periodically receiving axons from the sensory neurones

of the antennal sensilla.

Larva I- Larva II Moult. — Shortly before the end of the first instar, the mitotic rate increases

in the epidermis of each antenna. Apolysis then occurs, including the origins and insertions

of Johnston’s Organ and the extrinsic and intrinsic antennal muscles. This is followed by secre-

tion of a new, larger, second-instar cuticle inside the first (Figs. 4, 5). As soon as this secretion

begins, the ends of Johnston’s Organ and muscles attach to the new cuticle. All sense organs

are duplicated exactly except that two additional tactile hairs (t. h.) develop on each scape.

Each sensillum develops as a cytoplasmic extension of a trichogen cell around which cuticle

is subsequently deposited. If a particular developing sensillum is unable to fit into the space

available for it within the first stage cuticle, it is bent at the socket only. The cuticle of the

second-instar antennal segments is deposited in a folded state (Fig. 4) and space is left apically

in each first-stage antenna for the formation of the second-stage apical hairs. A dendritic sheath

(d. sh. s. o.) is observed in some preparations extending through a central pore in the second-

instar campaniform sensillum (L. II. c. s.) to the dome of that of the first (L. I. c. s.) (Figs. 4,

5), indicating that the first-instar sensilla function in the pharate second-instar. Such sheaths

are probably associated with all sensilla but were not visible in either whole mount or section-

ed preparations. The pharate larva II can move its antennae only slightly because their muscles

detached from the flrst-instar cuticle at apolysis.

Shortly before ecdysis, the larva becomes quiescent. At ecdysis, the first-instar head capsule

splits medially, with each new second-instar antenna being pulled out of the base of the first.

This requires that the new flagellar segments of each antenna be pulled through the very nar-

row base of the first-instar flagellar segment (Fig. 4).

As soon as its old cuticle is shed, the larva swallows air, contracts its abdominal muscles,

increases the blood pressure in its appendages, and stretches out the new second-instar cuticle.

Sclerotization follows shortly thereafter.

Propupa (Figs. 12-1 7).

Larva Il-Propupal Moult. — The details of apolysis and ecdysis at this moult are similar to

those of the previous one just described, except that in the cuticular pharate phase, the ex-

trinsic and intrinsic antennal muscles do not attach to the propupal cuticle as it is deposited

(Figs. 12-14). In addition, whereas the larval moult always takes place on the host plant, this

one usually occurs in the pupation site, either in a protected place on the host, or in the soil,

both areas having been entered by the larva before moulting takes place.

Structure. — The propupal antennae of F. fusca, although directed anteriorly, are short, are

of membranous cuticle, and lack well defined segmentation (Fig. 15). Just distal to the pro-

pupal campaniform sensillum (pro. c. s.), a deep, circular, annulus separates pedicellar and

flagellar regions. Three additional, weakly-developed, annuli suggest additional segmentation.

The homologies of these “segments” with their larval counterparts are best seen in pharate

propupae (Fig. 12).

The extrinsic (lev. ant.; dep. ant.) and intrinsic (fix. ant.; ext. ant.; scp. m.) muscles of the

larval antennae are unaltered in the propupal stage and maintain their birefringence (Figs. 14,

16, 17, 19). However, they do not function because they originate and insert in epidermis

Antennal Structure and Metamorphosis in Thysanoptera

33

SCO. J.O.

10jjm

Figs. 15-19. F. fusca. Propupa: Fig. 15. Right antenna, dorsal aspect. Fig. 16. Same, optical section. Note presence of scape

muscles (flx. ant; ext. ant). Fig. 17. Transverse section through head taken at point indicated by arrow in Fig. 6. Note

presence of extrinsic antennal muscles (lev. ant.; dep. ant.). Pharate Pupa: Fig. 18. Right antenna, dorsal aspect (Note: Most

propupal sensilla omitted from drawing). Fig. 19. Oblique sagittal section of base of antenna taken at point and angle indicated

by arrow in Fig. 18. Note J.O. scolopidia (sco. J. O.) and intrinsic (scp. m.) and extrinsic (lev. ant.; dep. ant.) antennal muscles.

rather than on propupal cuticle (Figs. 16, 19). Propupae are probably unable to move their

antennae because of this and because these appendages lack functional articulations.

Sense Organs. — Sensilla of the propupal scape and pedicellar regions are similar to those

of larvae, although the tactile hairs are longer and more delicate, and arise from raised sockets

(Fig. 1 5). Only two, well-developed thin-walled chemoreceptors (t.-w. chr.) are visible on the

flagellum, and many of its tactile hairs (t. h.) are either absent or much reduced (compare

Fig. 9 and 1 5).

The larval Johnston’s Organ (J. O.; sco. J. O.; s. cl. J. O.) is carried through unaltered into

Quaest. Ent., 1975, 11 (1)

34

Heming

the propupal stage (Figs. 13, 16, 19) but, like the musculature, has its origins and insertions

in epidermis rather than in propupal cuticle. Thus, it probably ceases to function in this instar

too.

Pupa ( Figs. 1 8-21 ).

Propupal- Pupal Moult. — The details of apolysis, cuticle deposition and ecdysis in this

moult are similar to those of the previous one (compare Figs. 12 and 18), and, again, take

place in the hidden pupation site of the insect. The main events occurring in each antenna are

(1) an increase in its length, (2) an increasing complexity in its Johnston’s Organ, and (3) its

dorsal flexion over the head and prothorax. Events (1) and (3) are both evident in the pharate

pupal stage (Figs. 18 and 19). The larger pupal cuticle (p. c.), developing within the propupal

cuticle (pro. c.), is deeply folded, particularly in the flagellar region and on the ventral side

of the scape and pedicel. When the pupa emerges and expands its new cuticle, these folds

straighten out. The expansion of the ventral folds causes each antenna to bend slowly back

over the head, and of the flagellar ones to almost double the length of the flagellar region

(compare Fig. 1 5 and 20).

The Johnston’s Organ (sco. J. O.; J. O.) in the propupa has its long axis slanted forward

and upward at an angle of about 30° (Fig. 19). At pupal emergence it is flipped over vertically

through an arc of about 90° with the result that its apex eventually points posteriorly at an

elevation of about 60° (Figs. 21, 22). This movement is passive and is due completely to the

expansion of pupal cuticle carrying along the underlying epidermis into which the apex of the

organ is inserted.

During the pharate pupal and young pupal stages, three new chordotonal organs develop

(for a total of six) and additional sensory elements are added to those already present (for a

total of from three to five scolopidia per scoloparium) (compare Figs. 13 and 25). Since these

have their imaginal form by the exuvial pharate adult stage (Fig. 25), the differentiative div-

isions responsible for their formation must occur earlier in the exuvial pharate pupal to newly-

emerged pupal periods. Unfortunately, most preparations of animals at this stage in their

development do not show the antennae to best advantage (frontal sections of the whole ani-

mal are the only ones of use) and those that do (e.g. Fig. 19 of a pharate pupa) show no evi-

dence of increased mitotic activity. Therefore, more detailed study at this period of develop-

ment, preferably with the transmission electron microscope, is required before positive con-

clusions can be drawn.

Structure. — The pupal antennae of F. fusca are of membranous cuticle, lack segmentation

completely, and are bent back dorsally over the head and prothorax (Fig. 20 — a lateral view).

As in the previous stage, both the extrinsic and intrinsic antennal muscles maintain their bire-

fringence, and, presumably, their ability to function, even though they are unable to do so

productively. As in the previous stage, they originate and insert in epidermis rather than in

cuticle (Figs. 19, 23, 24).

Sense Organs. — It is difficult, except for those of the pedicellar region (pupal campaniform

sensillum and Johnston’s Organ), to homologize the sensilla of this stage with those of the

propupa and adult. There are three, sub-apical, thin-walled chemoreceptors (t.-w. chr.). Most

tactile hairs (t. h.) arise from sockets that are higher than those of the propupa (Fig. 20). The

apex of each antenna bears six or seven protuberances, each topped by a tiny seta (t. h.). Some

setae which were short in the propupa are long in the pupa and vice versa. Numerous addition-

al small sense organs, just visible at the maximum resolving power of the light microscope, are

scattered throughout the length of the flagellar region.

Although difficult to see because of its position, a pedicellar campaniform sensillum (p. c. s.)

is situated at the base of each pupal antenna just behind its point of dorsal flexion (Figs. 20-

Antennal Structure and Metamorphosis in Thysanoptera

35

Figs. 20-27. F. fusca. Newly- emerged Pupa: Fig. 20. Right antenna, lateral aspect. Fig. 21. Same, optical section. Note groups

of imaginal sensory neurons (s. cl.). Exuvial Pharate Adult: Fig. 22. Right antenna, optical section. Note developing adult

antennal segments (d. seg.). Fig. 23. Oblique frontal section through developing scape and pedicel taken at point shown by

arrow in Fig. 20. Note scape muscles (scp. m.), blood cells (b. c.), and degenerating epidermal nuclei (p. n.). Fig. 24. Trans-

verse section through developing scape taken at point indicated by arrow in Fig. 22. Fig. 25. Same, through pedicel taken

at point shown by arrow in Fig. 22. Compare with Fig. 13 and note additional chordotonal organs (J. O.) and scolopidia

(SCO. J. O.). (Note that pupal cuticle omitted). Pharate Adult: Fig. 26. Right antenna, lateral aspect. Note that bases of

antennal segments 3-5 are telescoped, into their respective segments- and also the positions of the imaginal tactile hairs (t. h.).

(Most sensiUa are omitted from pupal cuticle in the drawing). Fig. 27. Transverse section through head taken at point in-

dicated by arrow in Fig. 28. Compare with Figs. 8 and 17 and note similarity in extrinsic antennal muscles (lev. ant.; dep.

ant.) in these stages.

Quaest. Ent., 1975, 11 (1)

36

Heming

22, 26). In the pharate pupa this sensillum is connected by a dendritic sheath (Fig. 18; d. sh.

s. o.) to its propupal equivalent (pro. c. s.).

Adult (Figs. 22-33).

Pupal-Adult Moult. — The actual pupal stage in F. fusca is very short, most time within the

pupal cuticle being spent as a pharate adult. Shortly after pupal emergence (Fig. 21), apolysis

occurs, the epidermal and sensory cells quickly assuming the configuration of the adult anten-

na (Fig. 22; hatching). Before imaginal cuticle deposition begins, a lengthened dendritic sheath

(d. sh. s. o.) is associated with the pupal campaniform sensillum (p. c. s.) (Fig. 22). In whole

mounts, many other pupal sensilla show similar, though finer sheaths. By this stage in develop-

ment, most internal structures of the antenna have assumed their final position (Figs. 22-25),

even though muscle fibres and scolopophorous organs have not yet connected to cuticle.

Adult tactile hairs (t. h.) and chemoreceptors (t.-w. chr.) develop and differentiate in the

same way as those of the larva II. The apices of each of the long, apical, tactile hairs (t. h.)

of the adult fit singly into apical protuberances available to them in the pupal antennal cuticle

(Fig. 26), suggesting that the two are homologous.

The cuticle of antennal segments two, six, seven, and eight is deposited in its fully expanded

configuration; whereas that of the scape (one) is thrown into folds, and that of the stalks of

segments three to five laid down telescoped into the bases of their respective segments (Fig.

26). The fully-expanded parts start to harden before adult emergence, the process continuing

in teneral individuals. When the adult emerges and expands its new cuticle, the folded and

telescoped portions straighten out, remaining lighter in colour than the remainder of the an-

tenna.

As imaginal cuticle deposition begins, the extrinsic and intrinsic antennal muscles, for the

first time since the second larval instar, re-attach to the cuticle. At first, this attachment is by

epidermal tendons; later, the myofibrils attach directly. Although they maintain their birefrin-

gence throughout metamorphosis, both groups of antennal muscles undergo some developmen-

tal changes in the pupa and pharate adult. The levator (lev. ant.) and depressor (dep. ant.)

muscles increase in length from about 43 jum in the larva II to about 75 pm in the adult female

and also in diameter (compare Figs. 27 and 28 with 6-8). Similar, though smaller changes occur

in the scape muscles (fix. ant; ext ant.; scp. m.) (compare Figs. 31 and 33 with 14 and 24).

Shortly after adult emergence, most antennal epidermal cells (as is true of those elsewhere

in the body) degenerate, leaving a thin epidermis with widely scattered nuclei in the mature

adult (compare Fig. 25 with 32 and 24 with 33).

Structure. - The imaginal antennae of F. fusca are similar in the two sexes but are smaller

in males. Each consists of a scape (scp.), a pedicel (ped.) and six flagellar segments (fl.) (Fig.

29). The scape of each is inserted into a socket (ant. s.) in the head capsule (Fig. 28) and is

articulated to it by means of small median and lateral antennifers extending inwards from the

socket rim (not shown because of their position). As in the larva, each antenna is raised and

lowered by levator (lev. ant.) and depressor (dep. ant.) muscles originating ventrally on the

anterior tentorial arms (at.) of the head capsule and inserting, respectively, into the dorsal

and ventral margins of the' scape base via short (9.0 pm) tendons (Figs. 27, 28). Each consists

usually of three (rarely of two) fibres (Fig. 27).

The base of the pedicel articulates with the scape by means of a large dorsal and a very small

ventral condyle borne at the apex of the latter. It is flexed laterally and extended anteriorly

by the scape muscles (fix. ant.; ext. ant.) (Figs. 28, 30, 31, 33). These consist of three fibres

each, as in the larva, and have the same origins and insertions.

Unlike that of larvae, the apex of the adult pedicel lacks dorsal and ventral condyles. In-

stead, a cuticular ring (c. r.) is present through which pass the cap cells of the Johnston’s Organ

Antennal Structure and Metamorphosis in Thysanoptera

37

Figs. 28-33. F. fusca. Adult: Fig. 28. Dorsal aspect of head (9), showing origins and insertions of intrinsic (fix. ant; ext.

ant) and extrinsic (lev. ant.; dep. ant.) antennal muscles (All setae omitted). Fig. 29. Right antenna, dorsal aspect. Fig. 30.

Same, optical section. Fig. 31. Frontal section through scape (scp.) and pedicel (ped.) of left antenna taken at point indicated

by arrow in Fig. 29. Note position of Johnston’s Organ sensory neurons (s. cl. J. O.). Fig. 32. Transverse section through

pedicel taken at point shown in Fig. 30. Fig. 33. Same, through scape taken at point shown by arrow in Fig. 29. Compare

with Fig. 14.

Quaest. Ent., 1975, 11 (1)

38

Heming

scolopidia (Figs. 29, 30).

Antennal segments two, three, four, five, and six bear, respectively, three, four, five, three,

and three whorls of micro trichia (m. t.) (Fig. 29). In females, all antennal segments are dark

brown, with segments three to five having yellowish stalks. Male antennae are yellow with

some light brown color in segments one to five.

Sense Organs. — The sense organs of the antenna of an adult female are illustrated in Figs.

26, 29, 31 and 32 and are listed in Table II. As can be seen by comparing Tables I and II, there

has been an increase in the number of tactile hairs (t. h.; from 29 to 53) and thin-walled

chemoreceptors (t.-w. chr.; from five to nine), but an apparent replacement of the apical

sense organs (ap. s. o.) of segment three of the larva by those of segment five of the adult. In

addition, most adult chemoreceptors, including the coeloconic pegs (c. pg.), differ in size,

shape and/or position from those of the larval stages (compare Fig. 29 with 1 and 9).

As already indicated, the imaginal Johnston’s Organ of F. fusca consists of six chordotonal

organs, each containing three to five scolopidia (Figs. 30-32). Since the latter are small and

not in register, a single transverse section (Fig. 32) does not show them all so that counts are

difficult to make. Since five were clearly visible in many of the sectioned organs, this is prob-

ably the number most characteristic.

As in larvae, some scolopidia appear to be innervated by one and others by two sensory

neurones according to the number of ciliary dilations visible inside each scolopidium (note

dots in Figs. 25 and 32). Insertions and origins of the chordotonal organs and the positions

occupied by their sensory cells (s. cl. J. O.) are approximately the same as those of larvae (Figs.

30, 31). The antennal nerve (ant. nv.) of each antenna (Fig. 30, 31) proximally, enters the

deutocerebrum of the brain; the latter now occupying the much larger adult head rather than

the prothorax. As in larvae, the nerve of each antenna bifurcates within the base of the first

flagellar segment (Fig. 30), the two branches coursing distally within the basement membrane

of the antennal epidermis.

Table II. Sense organs on the antenna of the adult female of F. fusca (N = 1 5).

Grand Total 71

Antennal Structure and Metamorphosis in Thysanoptera

39

Haplothrips verbasci

H. verbasci is confined to plants of the genus Verbascum (Scrophulariaceae) (Priesner, 1928;

Bailey, 1939), the entire life cycle being spent on the host. The two larval stages congregate

near the growing apex of the flower stalk or in the flowers, where they feed on the inner sur-

faces of the sepals. The quiescent propupae, pupae I and pupae II are usually hidden between

the seed capsules and the sepals in the lower, older part of the inflorescence. Teneral adults

stay in their pupation sites but mature ones are found wandering and feeding everywhere,

particularly among the flowers.

Larval Stages (Figs. 34-43; 76-78).

Structure. — Antennae of the two larval stages of H. verbasci (Figs. 34, 35, 40, 41) contain

the same number of segments (seven) as those of F. fusca, but differ from them in shape. The

scape (scp.) of each is inserted into a socket (ant. s.) in the head capsule (Fig. 39) but is artic-

ulated to it by a single median antennifer only. Each antenna is raised and lowered by two

groups of extrinsic muscles. The first are identical to those of F. fusca and consist of levator

(lev. ant.) and depressor (dep. ant.) fibres originating ventrally on the anterior tentorial arms

(at.) and inserting respectively into the dorsal and ventral margins of the scape base (Figs.

38, 39). The second set (lev. ant. c.; dep. ant. c.) have the same insertions; but converge post-

erodorsally to a common origin mid-dorsally on the vertex of the cranium (Figs. 38, 39).

Proximally, the pedicel (ped.) articulates on dorsal and ventral condyles borne by the apex

of the scape (scp.) (only the dorsal one is present in larvae of F. fusca). It is flexed and ex-

tended by 3-fibred scape muscles (fix. ant.; ext. ant.; scp. m.) with origins and insertions

identical to those of F. fusca (Figs. 35, 37-39, 41, 42). As in F. fusca, the first flagellar seg-

ment articulates laterally and mesally on dorsal and ventral condyles borne by the apex of

the pedicel (Figs. 34, 40).

In hving larvae of both sexes and stages, mounted in water under a coverslip (Heming, 1972)

and viewed at X500, irregular pumping movements can be seen occurring just proximal to the

base of each antenna. These movements vary in rate in fresh specimens observed at room temp-

erature, from about 32 to 80 X per minute and are asynchronous in the two antennae. Usually,

one contracts more rapidly than the other, with the pump of either antenna beating the faster.

In specimens mounted venter uppermost, there appear to be two fibres to this pumping organ;

one inserted dorsally, the other ventrally into the basal margins of the scape. I am unsure of

their origins. In sections (Figs. 37, 38, 42), I have been unable to see muscular tissue in the

site of the “pump” other than that of the extrinsic antennal muscles (lev. ant.; dep. ant.;

lev. ant. c.; dep. ant. c.). Since they have approximately the same orientation as the antennal

tentorial muscles, I, at first, thought them to be the same. However, the “pumps” continue

to beat without interruption throughout the quiescent stages until the pharate adult, even

during the pharate propupal period when massive changes occur in the structure of the an-

tennae (see below). Since the contractile elements of the antennal muscles degenerate in the

propupa, the antennal “pumps” cannot be these muscles. The “pumps” are not apparent in

adults of either sex.

The cuticle of all seven larval segments is smooth and sclerotized and lacks the whorls of

microtrichea present in those of F. fusca. (Figs. 34, 40). The apical portions of segments two

to six, bearing the sensilla, are of unpigmented, thinner cuticle (mb. c.) (Figs. 34, 37, 40, 42,

43).

Sense Organs. — The antennal sense organs of larval H. verbasci are identical in both instars

and sexes. They are illustrated in part in Figs. 34, 35, 37, 40-43, 76-78 and are listed in Table

III. Despite superficial differences in antennal structure existing between larvae of H. verbasci

and F. fusca (compare Figs. 34 and 40 with 1 and 9), their sensilla are remarkably similar, both

Quaest. Ent., 1975, 11 (1)

40

Heming

Figs. 34-38. H. verbasci Larva I: Fig. 34. Right antenna, dorsal aspect. Note the tiny coeloconic peg (c. pg.) on segment 6.

Fig. 35. Same, optical section. Pharate Larva II: Fig. 36. Right antenna, dorsal aspect, showing developing larva II antenna

(Note: Most larva I sensUla omitted). Fig. 37. Frontal section of scape (scp.) and pedicel (ped.) of right antenna taken at

point indicated by arrow in Fig. 36. Note that the origins of the Johnston’s Organ chordotonal organs (sco. J. O.) and the

scape muscles (fix. ant.; ext. ant.) are detached from the larva 1 cuticle (L. 1. c.). Fig. 38. Oblique parasagittal section of

head taken at point indicated by arrow in Fig. 39. Note the common insertions of the dorsal (lev. ant. c.; dep. ant. c.) and

ventral (lev. ant; dep. ant.) branches of the extrinsic antennal muscles and that these are detached at both ends from the

larva 1 cuticle (L. 1. c.).

Antennal Structure and Metamorphosis in Thysanoptera

41

ext. ant.

Figs. 39-43. H. verbascL Larva II: Fig. 39. Dorsolateral aspect of head, showing origins and insertions of intrinsic (fix. ant.;

ext. ant.) and extrinsic (lev. ant.; lev. ant. c.; dep. ant; dep. ant. c.) antennal muscles. Compare with Fig. 6 of F. fusca,

noting the additional, dorsal set of extrinsic antennal muscles (lev. ant c.; dep. ant. c.) of this species. (All setae are omitted).

Fig. 40. Right antenna, dorsal aspect. Fig. 41. Same, optical section. Fig. 42. Frontal section of scape (scp.) and pedicel

(ped.) of left antenna, taken at point indicated by arrow in Fig. 41. (Note: The structure labelled trachea (tra.) may be a

blood vessel). Fig. 43. Transverse section through pedicel taken at point indicated by arrow in Fig. 40. (Note: Johnston’s

Organ scolopidia (sco. J. O.) are not in register, so that not all are figured).

Quaest. Ent., 1975, 11 (1)

42

Heming

in number and in relative position in each segment (compare Tables I and III). The only dif-

ferences are an apparent absence of three apical sense organs (ap. s. o.) from segment five of

larval F. fusca (these are very small and may well be present in both species), and a shifting

of the chemoreceptor (t.-w. chr.) of segment three to the dorsolateral side in H. verbasci (it

is ventral in F. fusca; note absence in Figs 1 and 9).

Table III. Sense organs on the antenna of the larva II of H. verbasci (N = 5).

Grand Total 44

The Johnston’s Organ (J. O.) in larvae of H. verbasci consists of four chordotonal organs

each containing two or three scolopidia (sco. J. O.) (Figs. 35, 37, 41-43 - contrasting with

three containing two in larvae of F. fusca). Scolopidial counts are difficult to make in larvae

of this species because they are not in register as they are in F. fusca. All scolopidia are prob-

ably associated with two sensory neurons as most contain two ciliary dilations (Fig. 43 — note

dots). The origins and insertions of the Johnston’s Organ and the disposition of its sensory

neurons are similar to those of F. fusca (Figs. 35 and 41). As in that species, the larval head

is relatively small so that the brain is situated in the prothorax. The nerve of each antenna

likewise bifurcates within the base of the first flagellar segment (Figs. 35, 41).

Larva I- Larva LI Moult. - Replacement of the first by the second-instar antennae in H.

verbasci transpires in the same way as described forF. fusca (Figs. 36-38). In the pharate

larva II (Fig. 36) the old (L. I. c. s.) and new (L. II. c. s.) pedicellar campaniform sensilla are

likewise connected by a dendritic sheath (d. sh. s. o.). During apolysis and ecdysis, the anten-

nal “pumps” continue to beat.

Propupa (Figs. 44-53; 79, 80).

Larva Il-Propupal Moult. — The events occurring during replacement of the second larval

by the propupal antennae in H. verbasci are far more complex than those taking place in F.

fusca. At the larva Il-propupal apolysis, all cells within each larval antenna detach from the

cuticle and withdraw caudally down the length of the antenna into the head capsule (Fig. 44).

Antennal Structure and Metamorphosis in Thysanoptera

43

Figs. 44-48. H. verbascL Exuvial Pharate Propupa: Fig. 44. Right antenna, optical section, showing larval antennal epidermis

(L. II. e.) being withdrawn proximally. Pharate Propupa: Fig. 45. Dorsal aspect of head, showing propupal antenna (pro. ant.)

invaginated into a pouch (invg. pro. ant.) in the head epidermis. Note that the larval eye (L. II. omtd.) is situated in the pro-

thorax far caudad of the larva II cornea (L. II. cr.). (Note: Propupal sensilla and dendritic sheaths of larva II sensiUa (Fig. 47)

are omitted from the drawing). Fig. 46. Sagittal section through antenna taken at point indicated by arrow in Fig. 45. Note

that the sarcolemma (sarc.) of the contracted, larval muscles is thrown into folds and also the posterior position of the

Johnston’s Organ scolopidia (sco. J. O.). Fig. 47. Larva II right antenna, dorsal aspect. Notice the long dendritic sheath of

each sensillum and also that those of the scape and pedicel run proximally to the propupal antenna (Fig. 45).Emerging

Propupa: Fig. 48. Dorsal aspect of head, showing right propupal antenna (pro. ant.) evaginating from its pouch (Note that

this drawing is of a specimen of Haplothrips graminis Hood not H. verbasci).

Quaest. Ent., 1975, 1 1 (1)

44

Heming

Figs. 49-54. H. verbasci Newly-emerged Propupa: Fig. 49. Right antenna, dorsal aspect. Fig. 50. Same, optical section. Note

the thick epidermis. Fig. 51. Oblique sagittal section of antennal base taken at point indicated by arrow in Fig. 49. Note

that the extrinsic antennal muscles (L. II. lev. ant; L. II. dep. ant.) of the larva have lost most of their myofibrils (white).

Exuvial Pharate Pupa I: Fig. 52. Transverse section through antenna taken at point indicated by arrow in Fig. 49. Note shape

of propupal campaniform sensiUum (pro. c. s.), the large size of epidermal nuclei (compare with Fig. 46), and the dividing

cells (c. m.; c. t.). Fig. 53. Same, taken at point indicated by arrow in Fig. 50. Note that the Johnston’s Organ (J. O.) still

consists of 4 chordotonal organs. Pharate Pupa I: Fig. 54. Right antenna, dorsal aspect, showing developing pupa I antenna

(Note: Propupal sensilla omitted from drawing).

Antennal Structure and Metamorphosis in Thysanoptera

45

This withdrawal appears to have two causes. First, all extrinsic and intrinsic antennal muscles

contract maximally, the sarcolemma (sarc.) of each fibre being thrown into folds (Fig. 46)

(this contraction involves all head muscles (Heming, in prep)). Since all head muscles have

separated from the larval cuticle, this contraction serves to bunch epidermal and sensory cells

together in the centre of the head capsule (compare Fig. 46 with 38). In addition, there must

also be some change in shape in individual antennal epidermal cells, since muscle contraction

alone cannot account for complete withdrawal of all cells from within each antenna, and be-

cause these cells are bunched, when completely retracted, even distal to the intrinsic muscles

(L. II. scp. m.) (Fig. 46). At the completion of withdrawal, the ommatidia of the larval eyes

(L. II. omtd.) are situated anteriorly in either side of the prothorax (Fig. 45) and the John-

ston’s Organ (sco. J. O.) and scape muscles (L. II. scp. m.) far caudad of the base of each

larval antenna (Fig. 46).

Although I have not watched epidermal withdrawal in living specimens in its entirety (all

larvae drowned before completion), I know that it takes place rather slowly. Posterior move-

ment of the cells is imperceptible in live specimens and there is sufficient time for the tormo-

gen cell of each sense organ to produce a long dendritic sheath (Fig. 47). In spite of these,

at the completion of withdrawal, only the sensilla of the larval scape and pedicel appear to

maintain contact with the propupa within (Fig. 47).

A small number of degenerating cells (p. n.), as is usual for a moult (Heming, 1973), are

visible (Fig. 46) in each antenna during and after withdrawal, but cell death does not play a

major role in metamorphosis. However, individual cell nuclei are smaller (Fig. 46) than they

are later (Figs. 51-53).

As soon as antennal withdrawal is complete, the epidermis begins to deposit propupal cu-

ticle (Figs. 45, 46). At this stage, one can see that each antenna (pro. ant.) is sunk to its tip

in a pocket (invg. pro. ant.) in the head epidermis (Figs. 45, 46). As the long, propupal tactile

hairs (t. h.) (Fig. 49) develop, they poke apically out of these pockets where they are easily

confused with the elongated dendritic sheaths of the larval scape and pedicellar sensilla (Fig.

47).

All head muscles, except for two delicate fibres inserting into the anterior wall of the cib-

arium, begin to regress shortly after propupal cuticle deposition begins. Loss of birefringence

in their myofibrils is much more rapid than that occurring in the pretarsal depressor muscles

of this species (Heming, 1973), and is complete by the middle of the propupal stage. The

tentorial extrinsic antennal muscles (L. II. lev. ant.; L. II. dep. ant.) are the last to lose it.

(Fig. 51). As occurs in the pretarsal depressor muscles, regression in the head muscles involves

only their contractile elements, their sheaths remaining recognizable throughout metamorpho-

sis. However, both the scape muscles and the dorsal contingent of antennal extrinsic muscles

disappear completely.

A pharate propupa is easy to recognize under the stereoscopic microscope because it is

sluggish, is very fat, cannot move its larval antennae, and has its larval maxillary stylets extrud-

ed (Heming, in prep.). At ecdysis, the larval head capsule (L. II. c.) fractures behind the eyes

(L. II. cr.) and the propupal antennae are everted out of their pouches by pressure of haemo-

lymph (Fig. 48). The antennal “accessory hearts” maintain their beating throughout the period

described above.

Structure. — The propupal antennae of H. verbasci and other phlaeothripids are short,thick,

unsegmented, membranous stubs (Figs. 49, 50, 79). In newly-emerged individuals, the antennal

epidermis is very thick and consists of numerous closely-packed nuclei having large nucleoli

(Figs. 50, 51-53). Many cells are in various stages of mitosis (c. a.; c. m.; c. t.). These divisions

are probably both multiplicative and differentiative, presumably giving rise to the numerous

sensory, trichogen and tormogen cells of the imaginal array of sensilla as well as to additional

Quaest. Ent., 1975, 11 (1)

46

Heming

epidermal cells contributing to the longer antennae of subsequent instars. Some birefringence

is still evident in the larval antennal levator (L. II. lev. ant.) and depressor (L. II. dep. ant.)

muscles until the propupal-pupa I apolysis (Fig. 51).

Sense Organs. — Each propupal antenna bears three large, two medium, and three small

tactile hairs (t. h.), at least four thin-walled chemoreceptors (t.-w. chr.), and a pedicellar

campaniform sensillum (pro. c. s.) (Figs. 49, 52, 79, 80). There are also a few, very fine sen-

silla at the antennal apex just visible at the maximum resolving power of the light microscope.

The propupal campaniform sensillum (pro. c. s.) is smaller than that of the larva II and is more

convex (Figs. 52, 80).

The Johnston’s Organ in propupae still contains four chordotonal organs and is embedded

in thickened epidermis (Figs. 50, 53) proximal to the campaniform sensillum. I was unable

to determine the number of scolopidia (sco. J. O.) contained in each chordotonal organ, but

the larval number (three) is probably still present.

Antennal nerve (ant. nv.) and trachea (tra.) persist, with the latter thrown into loops (Fig.

50).

Pupa I (Figs. 54-58; 81, 82).

Propupal-Pupa I Moult. — During the propupal-pupa I apolysis, the epidermis separates

from the cuticle and begins to secrete a larger, folded, pupa I cuticle within (Fig. 54). The

pupa I campaniform sensillum (p. I. c. s.) develops some distance proximal to the propupal

one (pro. c. s.), so that the dendritic sheath (d. sh. s. o.) connecting the two is quite long. The

new, long tactile hairs (p. I. t. h.) develop with their long axes parallel to that of the propupal

antenna (Fig. 54). The increase in antennal length arises in the apical half of the propupal

antenna between the bases of the long tactile hairs (t. h.) proximally, and the apical chemo-

receptors (t.-w. chr.) (Figs. 49, 54, 55, 79, 81).

That the pupa I antenna is bent back posteriorly alongside the head (Figs. 55, 81) is due

to the way that its cuticle is deposited in the pharate pupa I (Fig. 54). More cuticle is produced

anteriorly than posteriorly and more folds develop here. At ecdysis, the fragile, propupal

cuticle remaining splits between the antennae and the new antennae are pulled out of the

bases of the old. On expansion of the new cuticle, these folds stretch out and the antennae

slowly assume their characteristic flexed position. Although not as drastic, the process is very

similar to that occurring after the propupal-pupal ecdysis of F. fusca.

Structure. — Pupa I antennae (Figs. 55, 81) are very similar to propupal ones (Figs. 49, 79)

except for their greater length, reflexed orientation and thin, attenuated epidermis (Fig. 56).

Sense Organs. — Antennal sensilla of this instar are practically identical to those of the

previous one, including their relative positions (compare Figs. 55, and 56 with 49 and 50).

The pupa I antenna lacks two chemoreceptors apically, and there are additional small setae

(not illustrated because of their size) scattered throughout. Sense organ positions indicate

that most of the increase in antennal length is interposed in its middle.

The high mitotic rate (c. p.; c. m.) characteristic of the antennal epidermis of the propupal

stage, continues during this instar (Figs. 57, 58). When the epidermis is attenuated in newly-

emerged individuals (Fig. 56), sensory cells (s. cl.) are easy to recognize because they adhere

together in groups (Figs. 56, 57). The mitoses taking place within these bunches are undoubted-

ly differentiative (Fig. 57).

In the pharate pupa II (Fig. 60), each Johnston’s Organ comprises six chordotonal organs

containing three (sometimes four) scolopidia each. These, again, are difficult to see because

they are embedded in epidermis and are not in register. As only four chordotonal organs are

present in the young propupa (Fig. 53), the additional ones must have been added sometime

between the pharate pupa I and the exuvial pharate pupa II stages. The many mitotic divisions

Antennal Structure and Metamorphosis in Thysanoptera

47

s cUO.

lOjjm

Figs. 55-60. H. verbasci. Newly- emerged Pupa I: Fig. 55. Right antenna, dorsal aspect. Fig. 56. Same, optical section. Note

attenuated epidermis and haemocytes (b. c.). Fig. 57. Frontal section, through antenna, taken at point indicated by arrow

in Fig. 55, showing differentiative mitotic divisions in a developing sensory cell group. Fig. 58. Transverse section, through

antenna taken at point indicated by arrow in Fig. 55. Note cells (c. p.; c. m.) in various stages of mitosis. Pharate Pupa II: Fig.

59. Right antenna, dorsal aspect, showing developing pupa II antenna. (Note; Most pupa I sensilla omitted from drawing).

Notice that the folds in the pupa II cuticle delimit 8 “segments”. Fig. 60. Transverse section through pedicellar region taken

at point indicated by arrow in Fig. 59. Compare with Fig. 53 and note the additional 2 chordotonal organs comprising the

Johnston’s Organ (J. O.).

Quaest. Ent, 1975, 1 1 (1)

48

Heming

occurring in this area during this period probably contribute to the development of these new

structures. Details of their development are difficult to follow because only perfect sagittal

sections of the animal can be used.

Pupa II (Figs. 59-63; 83-86).

Pupa I-Pupa II Moult. — The first clear indication of adult segmentation appears in the

pharate pupa II (Fig. 59) where, new cuticle, as it is deposited, is thrown into folds. These

are not dispersed evenly throughout each antenna but, instead, are spaced in such a way that

they delimit eight segments. When the pupa II emerges, these folds stretch out with the result

that segmentation again becomes less distinct (Figs. 61 ; 83, 85). In other respects, events are

the same as those occurring in the previous moult.

Structure. — Except for their increased length, pupa II antennae (Figs. 61 ; 83-86) are simi-

lar to those of propupa (Figs. 49, 79) and pupa I (Figs. 55, 81). They are about 1.5 fold long-

er than those of the pupa I, are more tightly fitted to the sides of the head (which now has

the shape and size of the adult head) and are arched dorsally at their bases and bent ventrally

and medially at their apices under the prothorax (Figs. 83-85).

Sense Organs. — Sensilla in pupae II are similar in number and placement to those of pupae

I (compare Figs. 61 and 62, with 55 and 56), but only one large chemoreceptor (t.-w. chr.)

is present on each antenna. There are also many small sensilla scattered throughout that I

have not illustrated. Most increase in antennal length again has arisen distal to the pedicellar

campaniform sensillum (p. II. c. s.).

Mitosis in epidermis and in sensory cell groups practically ceases in this instar and most

subsequent development involves the movement and differentiation of already-present cells.

The epidermis is, again, greatly attenuated in teneral forms (Figs. 62, 63), with groups of

sensory cells clearly defined.

Adult (Figs. 64-75:87).

Pupa II- Adult Moult. — Most time within the pupa II cuticle is spent as a pharate adult,

(Figs. 64-69). At the pupa Il-adult apolysis (Fig. 64), the epidermis contracts away from the

cuticle and almost immediately assumes the configuration of the imaginal antennal segments.

Trichogen cells (t. cl.) begin to send out protoplasmic extensions around which cuticle is de-

posited in the cuticular phase to form the external parts of adult sense organs. (Figs. 64, 66).

All tactile hairs are at first protoplasmic too, but I have not drawn them in. Their disposition

from the beginning, is as is shown in Fig. 69. By the exuvial pharate adult stage, most internal

structures have their imaginal positions (Figs. 64-68).

Figure 69 shows the position of adult antennal segments shortly before adult eclosion. This

drawing is fore-shortened proximally and apically because of the arched condition of the pupa

II antennal sheath (foreshortening is also present in Figs. 61, 62, and 64). Notice that the

cuticle of antennal segments two, seven and eight is deposited fully-expanded, whereas that

of segment one is folded, and that of the stalks of segments three to six is telescoped into the

bases of their respective segments. As in F. fusca, the fully expanded parts start to preharden

before adult emergence. After the pupa II cuticle is shed, the scape and the reflexed portions

expand and straighten, the antenna assuming its mature length (Fig. 71 — drawn at same mag-

nification as Fig. 69). Most epidermal cells degenerate soon thereafter.

In the exuvial pharate adult stage, the cells of the remnants of the extrinsic antennal muscles

(lev. ant.; dep. ant.) begin to show signs of renewed physiological activity. Their nuclei and

nucleoli enlarge and their cytoplasm increases in volume and in susceptibility to basic dyes

(Fig. 65). The cells begin to synthesize new myofibrillar material at about the same time as

adult cuticle deposition begins and birefringence begins to become apparent in these muscles

Antennal Structure and Metamorphosis in Thysanoptera

49

s.cl. JO.

Figs. 61-63. H. verbasci. Newly- emerged Pupa II: Fig. 61. Right antenna, dorsal aspect. Compare with Figs. 49 and 55 and

note the common chaetotaxy of the antenna in all 3 quiescent stages. Fig. 62. Same, optical section. Notice attenuated

epidermis and groups of sensory cells (s. cl.). Fig. 63. Frontal section through antenna taken at point indicated by arrow in

Fig. 61. Notice the haemocytes (b. c.) and the groups of sensory neurons (s. cl.) and their support cells.

Quaest. Ent., 1975, 11 (1)

50

Heming

66 68

Figs. 64-68. H. verbasci Exuvial Pharate Adult: Fig. 64. Riglit antenna, optical section. Notice developing segments (d. seg.)

and thin-walled chemoreceptors (d. t.-w. chr.). Fig. 65. Oblique sagittal section of antennal base taken at point indicated by

arrow in Fig. 64. Notice the developing, imaginal intrinsic (scp. m.) and extrinsic (lev. ant.; dep. ant.) antennal muscles.

Fig. 66. Frontal section of antennal segment 4, taken at point indicated by arrow in Fig. 64. Trichogen cells (t. cl.) have sent

out protoplasmic extensions around which cuticle will subsequently be deposited. Compare with Fig. 63 and notice the re-

arrangement of the epidermal cells. Fig. 67. Frontal section of scape and pedicel taken at point indicated by arrow in Fig. 64.

Notice the developing extensor muscle (ext. ant.), and the position of the sensory cells of the Johnston’s Organ (s. cl. J. O.).

Fig. 68. Transverse section through pedicel taken at point indicated by arrow in Fig. 64.

Antennal Structure and Metamorphosis in Thysanoptera

51

Figs. 69-71. H. verbasci Pharate Adult: Fig. 69. Right antenna, dorsal aspect. The adult cuticle of the scape is folded, while

that of the bases of segments 3-6 is telescoped into their respective segments. Notice that the pediceUar campaniform sensilla

of the adult (c. s.) is larger than that of the pupa II (p. II. c. s.). (Note: Other pupa II sensilla omitted from drawing). Adult:

Fig. 70. Dorsal aspect of head ( (5), showing origins and insertions of the intrinsic (fbc. ant.; ext. ant.) and extrinsic (lev. ant;

dep. ant.) antennal muscles. Compare with Fig. 39 (of the larva) and notice the absence of the dorsal contingent of extrinsic

muscles (All setae are omitted). Fig. 71. Right antenna, dorsal aspect.

Quaest. Ent., 1975, 11 (1)

52

Heming

d

m_|_

74

10jjm

Figs. 72-75. H. verbasci. Adult: Fig. 72. Right antenna, optical section. (Note: The sensory cells of the Johnston’s Organ

(s. cL J. O.) are actually situated in the base of the pedicel). Figs. 73- 75. Transverse sections through antenna taken at points

indicated by arrows in Fig. 72. Fig. 73. Through junction of pedicel and stalk of first flagellar segment, showing dendritic

terminals (d. ter.) of Johnston’s Organ. Fig. 74. Through pedicel, Johnston’s Organ (J. O.), and pedicellar campaniform

sensillum (c. s.). Fig. 75. Through scape and intrinsic antennal muscles (flx. ant.; ext. ant.).

Antennal Structure and Metamorphosis in Thysanoptera

53

in specimens examined under polarized light. Birefringence increases throughout the remain-

der of the pharate adult stage with banding appearing shortly before adult emergence. In the

exuvial period (Fig. 65), the rejuvenating fibres are attached to the epidermis. When cuticle

begins to appear, they attach to it, first by epidermal tendons; later directly.

The description above holds true also for most other head muscles (Heming, in prep.) and

for the pretarsal depressor muscles of the legs (Heming, 1973). No myoblasts have been ob-

served to take part in their development.

The scape muscles of the larva, unlike the tentorial complement of extrinsic muscles, ap-

peared to disappear completely during the propupal stage. In the exuvial pharate adult, these

muscles reappear as short strings of cells arising and inserting in the developing scape epider-

mis (Figs. 64, 65, 67; scp. m.; ext. ant.; fix. ant.). I have not observed myoblasts in this region

of the head (although haemocytes (b. c.) are present in large numbers shortly after pupa II

emergence (Fig. 62)), although these cells do take part in the de novo formation of other,

solely imaginal, muscles (Heming, 1973). Their subsequent development is identical to that

described above for the extrinsic muscles.

Structure. — Except for their slightly smaller size in males, antennae of the two sexes are

very similar. Each consists of a scape (scp.), a pedicel (ped.) and six flagellar segments (Fig.

71). The scape of each is inserted into a socket in the head capsule between the eyes (e) and

is articulated to it by well-developed median and lateral antennifers (antf.) extending inwards

from the socket rim (Fig. 71). As in adults of F. fusca, each antenna is raised and lowered by

levator (lev. ant.) and depressor (dep. ant.) muscles inserting respectively, by tendons into the

dorsal and ventral margins of the scape base and originating ventrally on the anterior tentorial

arms (at.) (Fig. 70). The cranial branches of these muscles, present in larvae (Fig. 39), are ab-

sent in adults.

Each pedicel (ped.) articulates with the apex of the scape (scp.) by means of dorsal and

ventral condyles borne by the latter, and is flexed and extended by muscles situated, respect-

ively, laterally (fix. ant.) and medially (ext. ant.) within the scape (Figs. 69, 70, 72, 75). As

in F. fusca, these consist of three fibres each (Fig. 75).

The imaginal pedicel lacks the dorsal and ventral condyles of the larval one and has, instead,

a cuticular ring (c. r.) through which pass the cap cells (d. ter.) of the Johnston’s Organ chor-

dotonal organs (Figs. 69, 71-73).

In adults of both sexes, segments one and two are dark brown, three to six yellow and seven

and eight light brown. Adults have no antennal “pumps”.

Sense Organs. - The sense organs on the antenna of a female of H. verbasci are shown in

Figs. 71, 72, 74 and 87 and are listed in Table IV. As can be seen by comparing Tables III

(larva) and IV, there has been an increase in the number of tactile hairs (t. h.; from 28 to 97),

thin-walled chemoreceptors (t.-w. chr.; from five to 14), and coeloconic pegs (c. pg.; from

three to four), but the three apical sense organs (ap. s. o.) of segment three of the larva are

absent in adults. In addition, all adult chemoreceptors differ in size, shape and position from

those of the larval stages.

The imaginal Johnston’s Organ of H. verbasci comprises six chordotonal organs each con-

taining three (sometimes four) scolopidia (Figs. 12,1 A) (compared with four chordotonal or-

gans of three scolopidia in the larva). Each scolopale is probably innervated by two sensory

neurons since most contain two ciliary dilations (indicated by dots in Fig. 74). The position

of the Johnston’s Organ and its sensory cells (s. cl.) in the adult antenna, is similar to that of

the larva (Fig. 72; sense cells in base of pedicel not in scape as shown). The dendritic terminals

(d. ter.) of the Johnston’s Organ fit into grooves in the stalk of the first flagellar segment

(Fig. 73).

The pedicellar campaniform sensillum (c. s.) of the adult is much larger than that of the

Quaest. Ent., 1975, 1 1 (1)

54

Heming

three quiescent stages, is not as convex, and differs in structure (compare Figs. 69, 71, 72

and 87 with 49, 52, 55, 61, 68, 80, 82, 86).

Proximally, the nerve from each antenna (ant. nv.) enters the deutocerebrum of the brain.

This is now, as in adults of F. fusca, situated mostly in the much larger adult head. As in

larvae, the nerve of each antenna (ant. nv.) bifurcates within the base of the first flagellar

segment (Fig. 72).

Table IV. Sense organs on the antenna of the adult female of H. verbasci (N = 11).

Grand Total 120

DISCUSSION

Larval Stages

Thysanoptera

Structure and variation in the antennae of larval aeolothripids, thripids and phlaeothripids

are briefly discussed by Priesner (1960). They are usually seven-segmented, as in the two spec-

ies of this paper, but those of Merothrips morgani Hood (Merothripidae) are six-segmented,

and there is a greater (in urothripines) or lesser (eg. Liothrips spp.) amount of fusion in the

distal segments of some phlaeothripids (Priesner, 1960; Heming, in prep.). Microtrichia are

usually present on the antennal segments of larval aeolothripids and most thripids, but are

absent from those of merothripids and phlaeothripids (the antennae of larval Heterothripidae

remain undescribed).

The numbers, kinds and positions of sense organs on the larval antennae of F. fusca and

H. verbasci are very similar (compare Tables I and III), even though these two species are not

closely related (see Fig. 50 in Heming, 1973). When larvae of Aeolothripidae, Heterothripidae

and Merothripidae are similarly investigated, their sensillar arrays will therefore, probably be

Antennal Structure and Metamorphosis in Thysanoptera

55

shown to have this common pattern too. Because of this similarity, larval antennae will prob-

ably be shown not to have the usefulness in taxonomic studies that imaginal antennae have

had — at least not at the generic and specific level.

Other Insects

With the exception of some sternorrhynchous Homoptera (see below), the antennae of

immature exopterygotes usually resemble those of their adults except for their reduced num-

ber of segments. In young endopterygotes, larval antennae are usually greatly simplified and

in some taxa (e.g. Diptera - Cyclorrhapha) are practically absent (Grasse, 1951 ; Snodgrass,

1954; Imms, 1957; Peterson, 1960, 1965); Mackerras, 1970).

Metamorphosis

Thysanoptera

The propupal and pupal antennae of thrips are briefly described by Priesner ( 1 960) and

have been illustrated for some species that have been the subject of detailed life history stud-

ies (eg. Lange and Razvyazkina, 1953; Loan and Holdaway, 1955; Derbeneva, 1962, 1967;

Lewis, 1973 (refs.) and Haga, 1974). In most aeolothripids, both propupal and pupal antennae

are unsegmented and are flexed dorsally over the head and prothorax (Priesner, 1960; Derben-

eva, 1967). Members of the genus Franklino thrips Back, however, apparently lack a propupal

stage (Reyne, 1920). In the pupa, the antennae are very long and reach posteriorly to the

second abdominal segment. This suggests, that in the species of this genus, there is no telescop-

ing of the adult antennal segments as they are forming. In thripids and phlaeothripids the

antennae are, respectively, as described here for F. fusca and H. verbasci. The quiescent stages

of Heterothripidae and Merothripidae, so far as I know, remain undescribed. Since the Hetero-

thripidae is probably a sister group of the Thripidae (see Fig. 50 in Heming, 1973), the an-

tennae of the quiescent stages of species in this family will probably be shown to be similar to

those of F. fusca and other thripids. Similarly, evidence from many sources (Heming, in prep.),

suggests that the Merothripidae and Phlaeothripidae are sister groups (see Fig. 50 in Heming,

1973). Therefore, the quiescent stages of merothripids, when discovered, may well prove to

have some antennal characteristics in common with those of Phlaeothripidae. It will be par-

ticularly interesting to learn whether species of this family have one or two pupal stages.

Events occurring during antennal metamorphosis have been briefly described in Limothrips

cerealium Haliday (Thripidae) by Davies (1969), in Liothrips oleae (Costa) (Phlaeothripidae)

by Melis (1934) and in Bactrido thrips brevitubus Takahashi (Phlaeothripidae) by Haga (1974).

In L. cerealium, the head musculature, as in F. fusca, is carried through practically unaltered

from larva to adult (Davies, 1969), although “one or two additional fibres” (p. 213) are present

in the extrinsic antennal muscles of the adult that are absent in larvae. During the propupal

and early pupal periods in this species, “. . . a hollow tube of densely packed nuclei” develops

on each side of the head, having an orientation parallel to that of the larval antennal levator

and depressor muscles. These nuclear aggregates are temporary however, and have disappeared

by the middle of the pupal stage without appearing to have contributed to the imaginal mus-

culature (p. 220). I have observed nothing in F. fusca resembling these aggregations except for

the antennal nerves which are shorter and thicker in the quiescent stages, and the antennal

tracheae which have a thicker epidermis when they are secreting a new intima during the

pharate portion of each instar. Davies ( 1 969) may have interpreted these structures as myo-

blast aggregations.

Davies (1969) also mentioned that the larval extrinsic muscles of the antennae seem to

undergo “some kind of reconstruction” during the quiescent stages, even though they main-

Quaest. Ent., 1975, 1 1 (1)

56

Heming

Figs. 76-81. H. verbascL Steroscan micrographs. Larval: Fig. 76. PediceUar campaniform sensillum. Fig. 77. Apical sense

organ of antennal segment 3. Larva II: Fig. 78. PediceUar campaniform sensiUum. Propupa: Fig. 79. Head and prothorax,

dorsal aspect. Fig. 80. PediceUar campaniform sensiUum. Pupa I: Fig. 81. Head and prothorax, dorsal aspect. Notice the

sUght indication of segmentation in the, antennae.

Antennal Structure and Metamorphosis in Thysanoptera

57

Figs. 82-87. H. verbasci. Stereoscan micrographs. Pupal: Fig. 82. Pedicellar campaniform sensillum. Pupa II: Fig. 83. Head

and prothorax, dorsal aspect. Fig. 84. Same, lateral aspect. Fig. 85. Same, ventral aspect. In these three figures, notice tlie

slight segmentation evident in the antennae. Fig. 86. Pedicellar campaniform sensillum. Adult: Fig. 87. Pedicellar campani-

form sensillum. Compare with Figs. 76, 78, 80, 82, and 86 and notice the large size of the imaginal organ.

Quaest. Ent., 1975, 1 1 (1)

58

Heming

tain their susceptibility to eosin. This also occurs in these muscles in F. fusca, giving rise to

the longer and more robust fibres of the adult.

Early in the propupal stage of L. oleae, there occurs, just as in H. verbasci, “una dissoluzione

completa dei muscoli intrinseci” (p. 288-289) of the head, including those of the antennae

(Melis, 1934b). From the dense masses of nuclei and cytoplasm remaining, the imaginal mus-

cles are reconstructed by secretion of new fibrillar material during the second pupal stage.

These events are described in detail in Melis’ text, but are illustrated inadequately in his photo-

micrographs (Fig. 3, plate 16; Fig. 6, plate 17 and Fig. 2, plate 19). Flis discussion is weakened

further by his failure to discriminate between the events occurring during the two pupal stages.

Nevertheless, the events taking place in H. verbasci and L. oleae appear to be very similar.

Haga figures accurately the heads of the pharate propupa (Fig. 1 1), pharate pupa I (Fig.

12), pharate pupa II (Fig. 13) and pharate adult (Figs. 14-16) of B. brevitubus. As in H. ver-

basci, imaginal segments three to six in this species form in a telescoped manner (Haga was

the first to describe this). Because the adult antennal segments of this megathripine are, rela-

tively, very much longer than those of H. verbasci, the degree of telescoping is very much

greater (compare his Figs. 15 and 16 with Fig. 69). However, his interpretation of what goes

on during metamorphosis is inaccurate because of his apparent complete reliance on whole

mount preparations (in spite of a statement to the contrary in his “Method” section). Accord-

ing to Haga (1974), “The histolysis of the antennal tissue begins after the fourth ecdysis, and

two days later the segmented blocks of the tissue appear again. These blocks are transformed

into each of the antennal segments of the adult.” (p. 22). I have found no evidence of histolysis

occurring at any time during metamorphosis of H. verbasci other than the usual isolated cel-

lular degenerations accompanying and following moults (Heming, 1973). His interpretation

of events occurring in the pharate propupa is similarly “off base” (p. 21).

Many of the discrepancies in musculature and metamorphosis that exist between L. cerealium

and L. oleae are not due to the inadequacies of Melis’ (1934b) investigation, as Davies (1969)

sometimes implied, but, instead are related to fundamental differences between the two in-

sects: the head muscles of thripids {F. fusca and L. cerealium) are carried through practically

unaltered, whereas those of phlaeothripids {H. verbasci and L. oleae) degenerate almost com-

pletely and are rebuilt.

The presence of a dorsal group of antennal extrinsic muscles (lev. ant. c.; dep. ant. c.) in

larvae of L. oleae, H. verbasci (Figs. 38, 39) and other phlaeothripids, may be required for

successful retraction, into the head, of the larval antennal epidermis at the end of the second

instar. This withdrawal does not occur in larvae of F. fusca and L. cerealium, and these mus-

cles are correspondingly absent. They are absent also from adults of species in both families

(Risler, 1957; Mickoleit, 1963; Davies, 1969; here).

The Phlaeothripidae is probably the most recently derived (Stannard, 1957) and the most

apomorphic of the five thysanopterous families (Stannard, 1957; Heming, 1970, 1973; see

Fig. 50). Therefore, the additional extrinsic antennal muscles of the larva must be derived

secondarily along with the type of head metamorphosis undergone by insects of this family.

Metamorphosis of the Musculature and Epidermis

Remarks made previously (Heming, 1973; pp. 1227-1229) concerning the metamorphosis

of musculature and epidermis in thysanopteran legs, are equally pertinent to the antennae.

The antennal muscles of F. fusca are midway between Snodgrass’ (1954) types (1) and (2)

in terms of their metamorphosis. Although larval fibres are reconstructed into adult ones,

they maintain their contractile elements throughout the process.

In H. verbasci, the dorsal extrinsic antennal muscles of the larva degenerate completely

and, therefore, belong in Snodgrass’ (1954) category (3) (larval muscles destroyed and not

Antennal Structure and Metamorphosis in Thysanoptera

59

replaced). The tentorial extrinsic antennal muscles undergo a reconstruction similar to that

followed by the pretarsal depressor muscles (Heming, 1973), differing only in that they lose

their contractile elements more quickly (completed by the mid-propupal stage instead of the

early pupa I stage). These muscles likewise are assigned to Snodgrass’ (1954) category (2) (lar-

val muscles reconstructed into adult muscles).

The developmental history of the intrinsic antennal muscles in this species differs from

that of the extrinsic muscles in that they apparently degenerate completely and then regen-

erate de novo. They thus fit in Snodgrass’ (1954) category (4) (larval muscles that degenerate

completely and are replaced by equivalent adult muscles).

Withdrawal of epidermis from within the larval antennae of H. verbasci at the end of the

second-instar, appears to involve both muscular contraction and changes in cell shape. Before

withdrawal, the epidermal cells are attenuated;after withdrawal, columnar or cuboidal. Sim-

ilar changes in shape, in a reverse sequence, occur during appendage eversion in the exuvial

pharate pupa of Drosophila melanogaster Meigen (Diptera-Cyclorrhapha) (Poodry and Schnei-

derman, 1971 — and refs.). These authors have experimental evidence that this change in shape

is caused by a change in intercellular adhesivity between cells of the imaginal discs.

Other Insects

The amount of reorganization occurring in the antennae of insects during metamorphosis

varies, depending on how far the young stages have diverged in form and habit from the adults

(Snodgrass, 1954). In the “segmented” antennae (ie. those having muscles in the flagellar

segments) of entognathous ametabolous apterygotes, additional segments are added through-

out postern bryogenesis by continuous subdivision of the apical segment (Imms, 1940). In

Thysanura and the orthopteromorph insects, new segments are added by division of the first

flagellar segment (meriston) and sometimes also by those situated immediately distal to it

(Imms, 1940; Schafer, 1973 — and refs.).

In certain sternorrhynchous Homoptera, there is a tendency for the juveniles to develop

special characters of their own that are absent from their adults (Snodgrass, 1954). Such dif-

ferences often show up on their antennae. Psylla buxi L. (Psyllidae) has five nymphal stages

(Wilcke, 1941). Antennae of the first stage are 1 -segmented; of the second and third, 3-seg-

mented (but quite different); of the fourth, 5-segmented; of the fifth, 9-segmented and of

the adult, 1 0-segmented. As additional segments are added, the shapes of other segments

change and new kinds of sensilla appear (his Figs. 26-30). Similar changes occur also in the

antennae of adelgids and some aphids (Weber, 1930; Pesson, 1951a; Woodward et al., 1970).

In Trialeurodes vaporariorum (Westwood) (Aleyrodidae), there are four nymphal stages

(Weber, 1934). First-stage nymphs are free-living “crawlers” with long, 3-segmented antennae.

The sessile second-, third-, and fourth-stage nymphs all have greatly reduced antennae which

are, respectively, annulated, unsegmented but with curled apices, and 2-segmented (his Fig. 1).

In the pharate adult, the long, 6-segmented adult antennae develop beneath their stubby,

fourth-stage counterparts in a looped configuration (his Fig. 9). Thus, whiteflies resemble

F. fusca, and H. verbasci in having a simplification of antennal structure in their intermediate

instars but, in the former insects, this reduction is correlated with the sessile habits of these

instars.

Depending on the family to which they belong, male Coccoidea undergo a variable amount

of antennal metamorphosis, (see table p. 569 in Weber (1933) for a comparison of life histories).

InMytilaspis fulva Targioni (= Lepidosaphes beckii Newman) (Diaspididae), the first-instar

“crawlers” have well developed, 5-segmented antennae (Fig. 1456 in Pesson, 1951a). In the

second-instar, these are atrophied; whereas in male propupae and pupae they reappear as suc-

cessively longer but unsegmented appendages bent back laterally on either side of the head

Quaest. Ent., 1975, 1 1 (1)

60

Heming

(Fig. 1456). Imaginal antennae are moniloform and 10-segmented.

Similar changes occur in the antennae of male Pseudococcidae, except that those of the

intermediate stages also are well developed (Makel, 1942). In these mealy bugs, the four ex-

trinsic antennal muscles are carried through the quiescent stages (Makel’s table p. 506) just

as they are in F. fusca, even though the propupal and pupal antennae are unsegmented, in-

flexible and flexed posteriorly on either side of the head. Although she did not discuss them

in the text, Makel illustrated also the scape muscles for second-stage nymphs (Fig. 4) and

adults (Figs. 14 and 15), but not for propupae (Figs. 6-8) or pupae (Figs. 10-12). Perhaps

they degenerate and then regenerate as do those of H. verhasci.

A detailed study of antennal metamorphosis in scale insects is lacking, so far as I know,

but events resolved up to now seem to be similar to those described here for F. fusca and

to a lesser extent, for H. verbasci. Although Thysanoptera and Coccina are quite closely re-

lated (see Fig. 64 in Hennig, 1969), the similarities in antennal metamorphosis summarized

above have probably arisen through convergent evolution.

In the less derived endopterygote insects (ie. Hymenoptera-Symphyta, most Coleoptera,

Raphidioidea, Megaloptera, Neuroptera, Mecoptera, Trichoptera, and Lepidoptera), each

adult antenna arises through a proliferation and reorganization of epidermal cells at the base

of the larval antenna (Eassa, 1953, Snodgrass, 1954). This new appendage then everts onto

the surface of the developing head at the larval-pupal apolysis and in the pupa is unsegmented,

is flexed ventrally or laterally, and is either free (exarate pupae) or fused to the body (obtect

pupae).

The antennae of those holometabolous insects (Hymenoptera-Apocrita, some Coleoptera,

Siphonaptera, Diptera-Cyclorrhapha) having apodous larvae, usually differentiate from anten-

nal disc cells that are invaginated into the larval head shortly before or after the insect hatches

from the egg (Murray and Tiegs, 1935; Bodenstein, 1950; Snodgrass, 1954; Anderson, 1963;

Postlethwait and Schneiderman, 1971). Their subsequent eversion at the larval-pupal apolysis

is similar to that mentioned in the previous paragraph as is their later development.

Adult

Structure

Thysanoptera. — The imaginal structure of thysanopterous antennae has been described

in greater or lesser detail; for representatives of all five families of the order (Buffa, 1898;

Peterson, 1915;Reyne, 1927; Priesner, 1928, 1960;Melis, 1934a, b; Doeksen, 1941;Pesson,

1951b; Risler, 1957; Mickoleit, 1963; and Davies, 1969). The most detailed studies are those

of Risler (1957) on Thrips physapus L. (Thripidae) and of Mickoleit (1963) on Aeolothrips

fasciatus (L.)(Aeolothripidae), Haplothrips statices Haliday (Phlaeothripidae) and Phlaeothrips

coriaceous Haliday (Phlaeothripidae). Both gave details of antennal structure, musculature and

innervation that are supported completely by my own observations. In addition, Risler (1957)

described the origin of each antennal nerve in the deutocerebrum of the brain (Figs. 41-43),

and the innervation of the extrinsic and intrinsic antennal muscles by branches of this nerve

(Fig. 9).

The anterior tentorial arms, upon which the extrinsic antennal muscles originate, are more

or less developed according to whether the tentorium is complete (Aeolothripidae, some

Heterothripidae) somewhat reduced (Merothripidae), or greatly reduced (most Heterothripidae,

all Thripidae and Phlaeothripidae). (Doeksen, 1941 ; Risler, 1957; Mickoleit, 1963; Heming,

in prep.).

Other Insects. — The structure of insect antennae is reviewed by Imms (1939), Schneider

(1964), Matsuda (1965) and Gouin (1968), is summarized in Weber (1933), Snodgrass (1935)

Antennal Structure and Metamorphosis in Thysanoptera

61

and Chapman (1969) and is described for insects in all orders by Imms (1957), and by the

various authorities in Grasse (1951) and Mackerras ( 1 970).

All thrips have the “annulated” antennae characteristic of Thysanura and the pterygote

insects, in which intrinsic muscles are limited to the scape. Their antennae are unusual in

that each scape has a dicondylar articulation with the head capsule. Usually, only a single,

ventral antennifer is present (Snodgrass, 1935; Chapman, 1969). Although the dicondylar

articulation is considered by some (eg. Matsuda, 1965) to be primitive, this condition in

Thysanoptera is probably secondarily derived since the closely related Psocoptera have a

single ventral condyle (Heming, per. obser.).

Matsuda (1965) has proposed that primitively, the extrinsic antennal muscles of insects

consisted of two dorsal levator and two ventral depressor muscles, all originating on the dor-

sal or anterior tentorial arms. In psocids, one of the ventral muscles has been lost and in thrips

only single levator and depressor muscles remain.

The primitive number of intrinsic muscles in insects is also considered to be four, comprising

single depressor, flexor, extensor and levator muscles (Matsuda, 1965). All investigated thrips

have the usual derived condition of two scape muscles functioning in flexion and extension

(although, in many insects, these are levators and depressors instead). Each consists of three

fibres, a characteristic shared with those of Psocoptera (Badonnel, 1951, his Fig. 1 159).

In Thysanura and the orthopteromorph insects, the antennae are usually many-segmented

and filiform (Imms, 1957). In hemipteroids (= Acercaria of Hennig, 1969), there is a strong

tendency towards reduction, the Psocoptera having 13 to 50, the Phthiraptera three to five,

the Thysanoptera six to ten, the Homoptera three to 20 and the Heteroptera four or five. The

Thysanoptera are considered to have arisen from a psocopteroid ancestor and are usually

placed after the Psocodea (Psocoptera + Phthiraptera) and before the Homoptera-Sternorrhyncha

in phytogenies of the Paraneoptera (see Fig. 64, Hennig, 1969). The number of antennal seg-

ments in adult Thysanoptera supports this placement.

All insects that have been thoroughly studied, including the Psocoptera (Badonnel, 1951;

his Fig. 1153), have been shown to have a blood vessel in each antenna, often associated with

a more or less well-developed, basal, “accessory heart” (Schneider, 1964). Slifer and Sekhon

(1974) have described such a vessel in antennae of Bagnalliella yuccae (Hinds) (Phlaeothripidae)

and Frankliniella tritici (Fitch) (Thripidae) three to four jam in diameter, this being “best seen

in electron micrographs of the entire cross section of the antenna” (p. 446). Neither I (Figs.

32, 33, 73-75), nor Risler (1957; Figs. 7, 8), norMickoleit (1963; Fig. 12) have found such a

vessel in our sections through scape and pedicel, but all of us have described an antennal trachea

that we may have confused with the blood vessel since “The trachea is small and difficult to

identify in sections with the light microscope.” (Slifer and Sekhon, 1974; p. 447). Although

my transverse sections of antennal flagella are rather poor, the only structure possibly a blood

vessel appears to me to be the basement membrane of the antennal epidermal cells. I have

already mentioned that immature H. verbasci have a “pumping organ” at the base of each an-

tenna.

Sense Organs

Thysanoptera. — Sense organs on the antennae of adult Thysanoptera, particularly those

of segments three and four are important key characters because of their diversity in different

taxa (Doeksen, 1941 ; Priesner, 1960). Thin-walled chemoreceptors (called “sensory areas” or

“sense cones” by thysanopterists) are used most frequently with the result that their variation

within the order is well known.

Slifer and Sekhon (1974) have recently described the ultrastructure of the adult sense organs

of antennae of B. yuccae and F. tritici, using, for the first time, a terminology conforming

Quaest. Ent., 1975, 11 (1)

62

Heming

with that developed for other insects. My observations on antennal sensilla agree with theirs

with the following exceptions:

(1) Using crystal violet stain and electron microscopy, Slifer and Sekhon (1974) have shown

that six of the 1 3 “tactile hairs” of segment eight (= subsegment six) in B. yuccae and four

of six in F. tritici are, in fact, thick-walled chemoreceptors. Such hairs are round-tipped,

double-walled, and curved or wavy. A similar number of terminal setae in larvae and adults

of F. fusca and H. verbasci are probably also of this kind.

(2) I have observed a single, small coeloconic peg (c. pg.) in each of antennal segments

three, five, six and seven in F. fusca (Fig. 29; Table II) and in segments three, five, seven and

eight in H. verbasci (Fig. 71 ; Table IV) that are not mentioned by these authors. I have since

seen them in antennae of F. tritici, and they are undoubtedly present in B. yuccae as well.

They are rather small (1-4 X 0.5- 1.0 jum) and are easily overlooked, even in macerated speci-

mens mounted in Hoyer’s medium and examined with phase contrast. Similar structures have

recently been described by Koteja (1974) from the antennae of certain scale insects (Fig. IB).

(3) Slifer and Sekhon (1974) also failed to notice the three “apical sense organs” of an-

tennal segment five, that I have found in adults of both F. fusca (Table II) and H. verbasci

(Fig. 71 ; Table IV). I have since located them on this same segment in F. tritici and they are

probably present also inB. yuccae. Slifer {in litt.), after reading my description of them, sug-

gests that these organs may be campanifomi sensilla as the latter are found in the antennal

intersegmental membrane of insects of several orders. They need additional study by trans-

mission electron microscopy. Figure 77 is a scanning electron microscope photo of one of

these structures from segment three of a larva I of H. verbasci.

(4) The single, large sensillum on the dorsal apex of the pedicel that I have called the

“pedicellar campaniform sensillum” (see Figs. 29, 69, 71, 74, 86) was shown by Slifer and

Sekhon (1974) in B. yuccae (but not in F. tritici) to be “a typical coeloconic chemoreceptor”.

They based their conclusion on the uptake, by the “peg” of this structure, of crystal violet

stain. These sensilla are present in all species of thrips in all life stages (Priesner ( 1 960) termed

each an “areola”) and are presumably homologous. Their failure to stain this organ in F. tritici

is probably due to its small size in this species. It is unfortunate that these authors provided

no electron micrographs of the organ, since, in immature stages (Figs. 5, 1 1, 43, 47, 52, 68,

76, 78, 80, 82, and 85), it has more resemblance to a campaniform sensillum.

(5) In F. tritici and B. yuccae “The axons from the sensory neurons of the flagellum join

to form two nerves that pass proximally and unite in subsegment 1 (= antennal segment three)

to form the antennal nerve.” (p. 446; Slifer and Sekhon, 1974). These nerves “are embedded

in the epidermis one opposite the other, and are separated from the lumen by the basement

membrane that lines the cavity”. This aspect was overlooked by me until I read Slifer and

Sekhon’s paper and also by Risler (1957) and Mickoleit (1963). Such details are difficult to

see in sections because they are at the maximum resolving power of the light microscope. I

have since re-examined sections of both larvae and adults of F. fusca and H. verbasci and

found a similar disposition of the antennal nerve in both species (see Figs. 2, 10, 30, 35, 41,

72).

The Johnston’s Organ of thrips has been previously described in the imaginal pedicel of

T. physapus by Risler (1957; Fig. 8a, b) and in those of A. fasciatus, H. statices and P. coria-

ceous by Mickoleit (1963). In all these species it consists “aus 5-6 Gruppen von je 3 Scolopidien”

(p. 113 in Mickoleit). That of H. verbasci is the same, but the Johnston’s Organ of F. fusca

contains up to five scolopidia per chordotonal organ.

Other Insects. — The sense organs of insects are presently being studied intensively by

electron microscopists and electrophysiologists (see reviews of Schneider, 1964; Bullock and

Horridge, 1965;Howse, 1968; Chapman, 1969; Slifer, 1970;Masson and Gabouriaut, 1973;

Antennal Structure and Metamorphosis in Thysanoptera

63

and numerous others). Those of Thysanoptera are similar to those of other insects with the

exception of the pedicellar coeloconic chemoreceptor (= pedicellar campaniform sensillum

of this paper). According to Slifer and Sekhon (1974), “This is the only instance known to

us of the presence, on the pedicel, of this type of chemoreceptor.” (p. 450). Similar organs

on the pedicels of psocids (Badonnel, 1951) and ants (Masson and Gabouriaut, 1973; Fig. 3)

may also prove to be chemoreceptors.

The scolopidia of the Johnston’s Organ of insects can each be innervated by one, two or

three sensory neurons (Howse, 1968; Masson and Gabouriaut, 1973), with two probably being

the characteristic number for Thysanoptera. Ultrastructural details are now known of the

Johnston’s Organ of several species in several different orders (Masson and Gabouriaut, 1973;

and references).

Most male insects have more complex sensilla arrays on their antennae than do females

(Schneider, 1964). This complexity is usually correlated with the ability of their females to

release sex attractants. Such differences are usually absent or are small in insects which live

gregariously or use auditory or visual organs for sexual orientation (Schneider, 1964). Since

most Thysanoptera are gregarious (Lewis, 1973), it is not surprising that the sensilla of the

adults of both sexes are similar. There is no evidence that thrips produce pheromones nor

that “the sexes find each other by means of sense cones on the antennae” (p. 13, Lewis,

1973).

Sense Organ Development. — Most sense organs in insects develop from solitary, epidermal

“mother cells’ (Stammzellen) after a characteristic number of mitotic “differentiative” divisions

(see Fig. 28 in Weber, 1954). Much of the mitotic activity observed in propupal (F. fusca,

H. verbasci) and pupa I {H. verbasci) antennal epidermis is probably of this kind. The number

of divisions giving rise to each antennal sensillum of F. fusca and H. verbasci is unknown as

it is for those of most insects. The ultrastructural details of sensillum differentiation have been

followed in the tactile hairs, thick-walled chemoreceptors and campaniform sensilla of the

cerci of Gryllus bimaculatus Degeer (Orthoptera, Gryllidae) (Schmidt and Gnatzy, 1971 ;

Gnatzy and Schmidt, 1972a, b) and in a thin-walled chemoreceptor of the antennae of Necro-

phorus vespilloides Herbst (Coleoptera, Silphidae) (Ernst, 1972). Details described in these

papers are probably similar to those occurring during sense organ development in F. fusca, H.

verbasci and other insects since the fully-developed structure of insect sensilla is basically the

same in all groups (Slifer, 1970).

Evolutionary Considerations

Three closely related evolutionary questions are raised by this study: (1) Why is a more

(//. verbasci) or less {F. fusca) drastic metamorphosis required of the antennae of thrips, when

the larval and imaginal structure of these appendages is no more divergent than that of insects

in which less drastic changes occur (eg. Psyllidae)? (2) How can one account for the origin of

the very different antennal metamorphosis of thripids (F. fusca, L. cerealium) and phlaeo-

thripids {H. verbasci, L. oleae)‘l (3) Why are antennae of the quiescent stages of thripids and

aeolothripids flexed dorsally over the head while those of phlaeothripids are flexed laterally?

My answers to these questions are best understood in the context of some of my previous

conclusions. In two papers on the thysanopteran pretarsus (Heming, 1972, 1973), I suggested

that some structural differences between larvae and adults can be accounted for by the usually

more cryptophilous habits of larvae. I hypothesized also that the quiescent stages of thrips

arose as a consequence of structural divergence between the young and adults of some ances-

tral form (Heming, 1973). Finally, I proposed that many of the structural differences existing

between adults of Terebrantia and Tubulifera, had arisen as a result of the adoption, by the

latter insects, of a mostly cryptophilous existence (Heming, 1970, 1972, 1973).

Quaest. Ent., 1975, 1 1 (1)

64

Heming

Many of the changes occurring during metamorphosis in thrips do not seem to be necessary.

A particularly good example of this is provided by the mouthparts. These are very similar in

larvae and adults (Peterson, 1915; Heming, in prep.) but are greatly reduced and lack function-

al stylets in the quiescent stages (Reyne, 1920, 1927; Heming, in prep.). In Homoptera (except

for male Coccoidea whose adults lack mouthparts) and Heteroptera, functional mouthparts

are usually present in all stages (Weber, 1930; Pesson, 1951a; Poisson, 1951 ; Woodward et al.,

1970), even in those taxa (eg. Aleyrodidae) having widely divergent juveniles and adults (Weber,

1934). I suggest here that the metamorphosis of the antennae may be another example of

this. Therefore, one answer to question (1) is that the quiescent stages of thrips are relicts of

an ancestral situation in which differences between larvae and adults were greater and pupal

stages more necessary, than is true of present forms. An indication of this is that species in

the more primitive families (Aeolothripidae, Heterothripidae and possibly Merothripidae),

still pupate within a cocoon spun by the second-stage larvae (see Heming, 1973 for a fuller

discussion). Also, members of the primitive, aeolothripid genus Franklinothrips have only a

single pupal stage (Reyne, 1920), just like holometabolous insects. Finally, the quiescent

stages of at least some aeolothripids, unlike those of thripids and phlaeothripids, are unable

to move when disturbed (Derbeneva, 1967), suggesting that greater structural changes occur

during metamorphosis of these insects. Another possibility, of course, is that there are reasons

for holometabolism in present-day thrips that we have not yet discovered.

The answers to questions (2) and (3) may be related to the more cryptophilous life habits

of both larvae and adults of Phlaeothripidae. Aeolothripids, heterothripids and Thripinae

usually pupate in soil (Priesner, 1960; Lewis, 1973) or in cocoons on the leaves (some aeolo-

thripids). Heliothripinae, which are mostly leaf feeders, pupate on the underside of leaves

(Lewis, 1973). In these pupation sites, dorsally flexed antennae are no problem to the bearer.

Phlaeothripids, however, usually pupate in crevices, either on their host plants (gall formers

and other phytophagous species) or under bark and litter (mycophagous forms) (Lewis, 1973).

Because their antennae are either short (propupae) or flexed back tightly against their heads

(pupae I and II), they would offer little hindrance to the movement of the animal if it were

disturbed. The prothorax is usually as wide or wider than the reflexed antennae (Figs. 79, 81,

83-85), and, together, they form a wedge, enabling these stages to further push themselves

into crevices. Their antennae would also not keep them from backing out, whereas the dor-

sally-flexed ones of aeolothripids and thripids would.

The Aeolothripidae is probably the most primitive family of the Thysanoptera (Stannard,

1957; Mickoleit, 1963; Heming, 1973, in prep.). Therefore, the dorsally flexed position of the

antennae in both propupae and pupae is probably primitive in this order. That they are oriented

thus rather than laterally or ventrally as they are in most insect pupae (Peterson, 1960, 1965),

is further evidence that the pupal stages of Thysanoptera are independently derived.

The reason that the antennal (and most other head) muscles of Phlaeothripidae undergo

almost complete dissolution followed by regeneration during metamorphosis, whereas those

of thripids do not, is probably associated with mouthparts and feeding and I prefer to leave

speculation on this topic to a future contribution on the metamorphosis of the mouthparts.

ACKNOWLEDGEMENTS

Much of the practical work and all of the writing of this paper were done in the Laborator-

ium voor Entomologie, Landbouwhogeschool, Wageningen, The Netherlands. I thank Dr. R.

H. Cobben for reading an early draft of the manuscript and for his constructive criticism and

discussion. I am indebted also to Prof. Dr. J. de Wilde, Director of the Laboratory, for pro-

viding space and facilities and to J. W. Brangert for reproducing copies of the stereoscan

Antennal Structure and Metamorphosis in Thysanoptera

65

micrographs. These were taken by G. D. Braybrook. J. S. Scott prepared the photographic

A

plates. H. Goulet translated the abstract into French and Mrs. S. Hamilton typed the manu-

script. This study was supported by a grant (A5756) from the National Research Council of

Canada.

Note added in proof: “A recent paper by Mound and O’Neill (1974) (Mound, L. A., and

K. O’Neill. 1974. Taxonomy of the Merothripidae, with ecological and phylogenetic consid-

erations. (Thysanoptera). Journal of Natural History 8: 481-509.) received after this manu-

script was submitted, contains a convincing discussion of the phytogeny of the Thysanoptera

that differs somewhat from that illustrated in my 1973 paper (Fig. 50, Heming, 1973). These

authors show a fully-developed tentorium to be present in the heads of adult Damerothrips

Hood and Erotidothrips Hood (Merothripidae) and also illustrate representative antennae for

most species in the Merothripidae. They conclude that the Merothripidae, not the Aeolothripidae,

is the most plesiomorphous of the thysanopteran families.”

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FATE OF DIELDRIN IN SEDIMENT, WATER, VEGETATION,

AND INVERTEBRATES OF A SLOUGH IN CENTRAL ALBERTA, CANADA

D. M. ROSENBERG*

Department of Entomology

University of Alberta Quaestiones Entomologicae

Edmonton, Alberta T6G 2E3 11:69-96 1975

Sufficient dieldrin was applied in July, 1967 to a slough in central Alberta, Canada to give

a concentration of approximately 1 ppb in water. Dieldrin concentrations in mud, water, veg-

etation, and invertebrates were monitored by gas chromatography until fall, 1968. Residues

were undetectable in mud, water, and vegetation by the spring of 1968 but persisted at low

levels in resident invertebrate populations throughout the remainder of the study, final con-

centrations varying from 1.8 to 44.6 ppb. Resident primary and secondary invertebrate con-

sumers carried similar levels of dieldrin in their tissues.

The fate of chlorinated hydrocarbon insecticides in mud, water, vegetation, and invertebrates

as determined in this study is compared with the results of similar studies.

Une concentration approchant une partie par milliard de dieldrin dans I’eau d’un marais du

centre de V Alberta tut appliquee en juillet, 1967. Les concentrations dans le lit beueux Veau,

la vegetation et les invertebres tu rent analysees par chromatographic gazeuse fusqu’d Tautomne

de 1968. Aucun residu ne tut decele dans la boue, Veau et la vegetation au printemps de 1968,

mais une concentration tres basse (concentrations finales variant entre 1.8 et 44.6 parties par

billions), a persiste dans les populations des invertebres residants pendant le reste de Vetude.

Les invertebres consomateurs primaires et secondaires residant dans le marais ont demontre

des taux similaires de dieldrin dans leurs tissues.

Le sort des insecticides d ’hydrocarbons chlorines dans la boue, Veau, la vegetation et les

invertebres comme determine dans cette etude est compare aux resultats d’etudes similaires.

INTRODUCTION

Community studies dealing with the effects of chlorinated hydrocarbon^ insecticides on

the diversity of freshwater invertebrates are common. Studies on flowing water systems in-

clude those of Ide (1957), Corbet (1958), Bridges and Andrews (1961), Frey (1961), Hynes

and Williams (1962), Moye and Luckmann (1964), Hitchcock (1965), Dimond (1967), Hat-

field (1969), Fredeen (1972), and Wallace et al. (1973). Lentic studies include those of Grzenda,

Lauer, and Nicholson (1962), Jones and Moyle (1963), Edwards et al. (1964), and Kennedy,

Eller, and Walsh (1970).

In general, these studies deal with relatively high concentrations of pesticides and the inver-

tebrates are not analyzed for residue levels. Unfortunately, studies of the effects of pesticides

on the diversity of freshwater invertebrates are separate from studies of residue levels. In ad-

dition, the effects of dieldrin, a once common agricultural insecticide, on aquatic communities

have not been studied.

* Present Address: Fisheries & Marine Service, Freshwater Institute, 501 University Crescent,

Winnipeg, Manitoba R3T 2N6.

1. I use “chlorinated hydrocarbons” to designate DDT and its related compounds, the

cyclodienes (excluding toxaphene which is not a true cyclodiene; O’Brien, 1967), poly-

chlorinated biphenyls (PCB’s), and the hexachlorocyclohexanes.

70

Rosenberg

With the advent of more sensitive analytical techniques (Moore, 1967; Edwards, 1970;

Chesters and Konrad, 1971 ; Cope, 1971 ; Muirhead-Thomson, 1971) and the creation of

national pesticide surveillance programs (Breidenbach and Lichtenberg, 1963; Stickel, 1968;

Edwards, 1970; Chesters and Konrad, 1971) pesticide residues have been detected in concen-

trations of tens of parts per billion and less in water (Breidenbach and Lichtenberg, 1963;

Tarzwell, 1965; Westlake and Gunther, 1966; Edwards, 1970; Chesters and Konrad, 1971;

Muirhead-Thomson, 1971); bottom sediments (eg. Morris and Johnson, 1971); and fish (e.g.

Fredeen, Saha, and Royer, 1971). Of these residues, dieldrin is widespread and common (Bues-

cher, Dougherty, and Skrinde, 1964; Green, Gunnerson, and Lichtenberg, 1967; Moore, 1967;

Stickel, 1968; Edwards, 1970; Chesters and Konrad, 1971). The persistence of dieldrin in the

environment has long been known from field studies. Mackay and Wolkoff (1973) have shown

mathematically that the persistence of dieldrin in water was the longest of several compounds

examined (two alkanes, six aromatics, four pesticides, four PCB’s, and one mercury). Its po-

tential evaporation half-life in 1 m^ of water at 25 C was 723 days. Mackay and Wolkoff (1973)

also noted that evaporation rates in the environment may be substantially slower, making

dieldrin one of the most persistent environmental contaminants.

Unfortunately, the literature on the effects of pesticides on fauna is fraught with general-

ities. There exists a belief that the use of pesticides automatically implies an effect on diversity

(e.g. Moore [ 1 967] ; p. Ill, 113, 114, 1 25). If such is the case, the need to determine the

levels of pesticides in fauna that will cause diversity changes is obvious. Tarzwell (1965),

Moore (1967), and Stickel (1968) discussed this point but only indirectly. However, with re-

spect to DDT in Maine forests, Dimond (1969, p. 5) stated: “. . . thresholds for such [sub-

lethal] effects are not known. This aspect of the problem needs thorough study in the imme-

diate future.” Cocks (1973, p. 149) noted that “The biological effects of low doses of pesticides

are at present poorly understood”. Edwards and Thompson (1973) noted that only sparse

evidence exists on the effects of sublethal doses of pesticides on soil inhabiting invertebrates.

Menzie (1972, p. 216) stated: “We have only scratched the surface in understanding the sig-

nificance and the effects of low levels of pesticides.”

Trophic level effects (i.e. the concentration of pesticides along food chains) are a commonly

reported result of pesticide use (e.g. Hunt and Bischoff, 1960; Fillmore, in Rudd, 1964; Bridges,

Kallman, and Andrews, 1963; Hickey, Keith, and Coon, 1966; see also reviews by Moore, 1967;

Newsom, 1967; Stickel, 1968; Edwards, 1970; and Cope, 1971) and are an important aspect

of studies of the effects of chlorinated hydrocarbon insecticides on faunal diversity. However,

the study of trophic level effects solely in invertebrates has been neglected.

In 1966, an experiment was begun to examine the effect of a single small addition of dieldrin

to a slough in central Alberta, Canada. The major objectives were to determine the persistence

of dieldrin in the aquatic ecosystem, whether the chemical became concentrated in higher troph-

ic levels of the invertebrate community, and whether such an application had a significant

effect on the diversity of the aquatic invertebrates. The results of the first two objectives are

presented in this report.

METHODS

Introduction.

The occurrence of dieldrin residues in the water, substratum, plants, zooplankton, and

benthic invertebrates was measured in two sloughs in central Alberta, Canada: a control slough

to which no dieldrin was added and an experimental slough to which enough dieldrin was

added in July, 1967 to give a concentration of approximately 1 ppb in water.

Dieldrin was used because of its widespread and common occurrence in freshwater ecosystems.

Fate of Dieldrin in Invertebrates

71

and because of its persistence. A concentration of 1 ppb in water was chosen because it was

well below dieldrin LC^q’s for a number of aquatic invertebrates (Rosenberg, 1973). This

concentration fulfilled the requirement of low level dieldrin contamination and ensured that

mass kills of fauna did not occur. Sloughs were chosen as the experimental sites because they

are a common feature of the Edmonton area, they sometimes receive runoff containing pesti-

cides (see Westlake and Gunther, 1966; Van Middelem, 1966), and because their relatively

small size makes community studies more manageable in a practical sense.

Description of study site.

As dieldrin was at no time during the study detected in any of the samples from the control

slough, this slough will not be discussed further here. A detailed description of the control

slough is given in Rosenberg (1973).

The experimental slough (hereinafter called “D”) is located in the parkland area of central

Alberta in the County of Strathcona approximately 14.5 km southeast of the city of Edmon-

ton and in an area of mixed farming. It is at 113° 22' 08" W, 53° 25' 27" N, approximately

738 m above sea level, and has an area of approximately 1 ha. The surrounding terrain has

rolling moraines and kettlehole sloughs. Bayrock and Hughes (1962) described the geology of

the area and Bowser et al. ( 1 962) the soils. For size, shape, and depth contours of D slough

as taken at the start of the study see Fig. 1.

Algal blooms occurred during the period May to October over the 3 years of the study. The

commonest algae were Oscillatoria sp., Nodularia sp., and Mougeotia sp. In 1966 and 1967

the main blooms occurred during July and August and were over by September. However, in

1968 the main blooms occurred during June and July with a minor bloom (mainly Oscillatoria

sp.) in August.

The main submergent vegetation was the hornwort {Ceratophyllum demersum L.). It was

dispersed fairly evenly throughout the slough. Sago pondweed {Potamogeton pectinatus L.)

was present in 1967. It became more common in 1968. Ivy-leaved duckweed {Lemna trisulca

L.) was the dominant species of floating vegetation and lesser duckweed {L. minor L.) was

also present. The emergent vegetation was composed of stands of common cattail {Typha

latifolia L.), great bulrush {Scirpus validus Vahl), and mare’s tail {Hippuris vulgaris L.) which

were discontinuously distributed around the periphery of the slough and faded shoreward

into the sedges (Carex rostrata Stokes, C. aquatilis Wahlenb., and C. atherodes Spreng.) and

Kentucky blue grass {Poa pratensis L.) which in turn continued from the water-land transition

onto the land. Among the sedges and grass along the shore were moss, Drepanocladius aduncus

(Hedw.) Warnst., and liverwort, Rhicciocarpus natans (L.) Corda. A typical hydrarch succession

was evident in the transition from water to land with Populus balsamifera L. being the climax

species (Rosenberg, 1973).

There was an overall decline in the level of water in the slough during the 3 years of the

study, probably caused mainly by the below-average precipitation over the study period.

Maximum and minimum and surface water temperature regimes are shown in Figs. 2 and

3 respectively. Maximum and minimum temperatures generally increased and decreased re-

spectively throughout the study, probably reflecting the inability of the reduced water vol-

ume in the slough to buffer air temperature change. Stratification did not occur.

Since free water was observed in D slough past the end of December in 1966 and 1967, the

slough possibly did not freeze to the bottom in the first (1966/1967) or second (1967/1968)

winters. Almost certainly, however, D slough totally froze in the third winter (1968/1969)

probably because of the much reduced water level.

A Secchi disc could always be seen at the bottom of the deepest part of the slough on a

clear day and turbidity levels (measured by the Model DR-EL Direct Reading Engineer’s

Quaest. Ent, 1975, 1 1 (1)

METERS

72

Rosenberg

Sd3i3^

Fig. 1. Size, shape, and depth contours of D slough at the start of the study (end of May, 1966). (Methods according to Welch, 1948).

TEMPERATURE (®C) 2 TEMPERATURE (°C)

Fate of Dieldrin in Invertebrates

73

35n

LEGEND

X 1966

• 1967

A 1968

V DIELDRIN APPLIED

MAX. TEMP

MIN. TEMP

30-

25-

15-

10-

5-

MAY I JUNE ^ JULY I AUGUST I SEPTEMBER I OCTOBER I

TIME OF YEAR

2. Maximum and minimum water temperatures in D slough for May to October 1966, 1967, and 1968.

30-1

25-

20-

15-

10-

LEGEND

X 1966

• 1967

A 1968

V DIELDRIN APPLIED

MAY ^ JUNE ^ JULY ^ AUGUST I SEPTEMBER I OCTOBER I

TIME OF YEAR

Fig. 3. Mean surface water temperatures in D slough for May to October 1966, 1967, and 1968.

Quaest. Ent., 1975, 1 1 (1)

74

Rosenberg

Laboratory Hach Kit; Hach Chemical Co.; Ames, Iowa) were in the 0 to 25.0 ppm range.

Oxygen concentrations (Winkler iodometric method: American Public Health Association,

1960) generally did not exceed 20 ppm. Anaerobic conditions were found in the slough in

November and December, September, and February of 1966, 1967, and 1968 respectively.

The only evidence of oxygen stratification was during the open-water period of 1967. Oxy-

gen concentrations usually varied among depth classes. Many times during winter sampling

H2S was evident.

Dissolved solids, and other dissolved gases were measured by the Hach Kit. Total hardness

was in the 300 to 400 ppm range and the overall mean calcium to magnesium ratio was 1.84.

Alkalinity was mainly due to the bicarbonate ion. The pH varied from 7.65 to 9.30 with an

overall mean of 8.55. Free CO2 concentrations were generally within the 0 to 10 ppm range.

Values for orthophosphate usually remained below 1.0 ppm throughout the study. Sulphate

was the most abundant anion in the slough. Concentrations of sulphate were generally in the

200 to 350 ppm range. Nitrite and nitrate nitrogen were generally in the 0 to 0.02 ppm and

10 to 25 ppm range respectively. Physical and chemical parameters of the water of D slough

were similar to those of the slough described by Daborn (1969) and within the ranges report-

ed by Hartland-Rowe (1966) for permanent ponds in the Canadian prairies.

Dieldrin application.

To estimate the volume of water for dieldrin application, the slough was re-surveyed and

re-mapped two days before the pesticide was added. Four depth classes were used, and from

the formula in Welch (1948), the volume was calculated to be close to 4.0 X 10^ liters. As

the 0 to 30.5 cm depth class was impossible to delimit in the field, a volume of 4.0 X 10^

liters was used.

The dieldrin used (Shell Canada Ltd.) was an emulsifiable concentrate with an average

concentration of 0.2 X 10^ ppm (Rosenberg, 1973). The amount required to yield a concen-

tration of 1 ppb in 4 X 10^ liters of water was calculated to be 22.7 ml.

The pesticide was applied using a compressed air sprayer (Leigh Products Inc., Universal

Metal Products Div., Sarnac, Mich.) calibrated to ensure an even flow. The boat was rowed

over the slough in patterns meant to yield an even application of the pesticide (Rosenberg,

1973; Fig. 2). The nozzle of the sprayer was held about 5 cm below the water surface to en-

sure that all the pesticide was added to the slough.

Sampling program for gas chromatography.

Eleven sets of dieldrin analyses were done on mud, water, vegetation, and invertebrates:

once before application and eight times after application during the open-water period (at

approximately monthly intervals after application until September, 1967, from May to Sept-

ember, 1968, and in June, 1969) (Table 1). Mud and ice were collected on February 9, 1968

and on March 6, 1969.

Mud and water samples were collected from each slough according to a random sampling

plan and in proportion to the area being sampled. For the collection of mud, core samples

were taken from each quarter of the slough, using a Moore (Phleger) sampler with a glass tube.

The corer sampled the top 7.6 to 10.2 cm of the slough bottom. Samples were pooled by

quarter and a subsample taken for analysis. Water samples were taken, from the shallowest to

the deepest of the four depth classes, with a Kemmerer bottle which was emptied into a 20

liter pesticide-free pail designated for each depth class. A subsample was removed from each

pail for analysis.

Four kinds of vegetation were collected for dieldrin analysis: submergent (C. demersum),

floating {L. trisulca), emergent (C. rostrata), and algae. Algae samples were collected whenever

Fate of Dieldrin in Invertebrates

75

possible in 1968 (usually during a bloom). The sampling plan for invertebrates (Rosenberg,

1973) was used to establish the areas from which each type of vegetation was collected. The

samples of each type and from each quarter were put into separate plastic freezer bags for

return to the laboratory and sub-samples removed from the bags for analysis.

Table 1. Samples taken from D slough during the open-water season for dieldrin analysis.

* Sub. = submergent; Float. = floating; Em. = emergent.

** Mixed with algae.

The mud, water, and vegetation samples were stored at 7 to 8 C and gas chromatographic

analysis was usually completed within a few days after their collection.

Sampling of invertebrates was done with a dip net at predetermined locations in the littoral

area of each slough (Rosenberg, 1973). Zooplankton was collected with a Wisconsin plankton

net. Invertebrates were sorted in the field and were put immediately into a portable freezer

containing dry ice. Vials were stored in the laboratory in a -23 C freezer room prior to dieldrin

analysis. Chironomid immatures were also collected from unpreserved benthic samples taken

with a 15.2 cm^ Ekman grab (Rosenberg, 1973). Gastropods were collected at random in

addition to those collected as described above. While it was unnecessary to make quantitative

collections of invertebrates, it was important that enough biomass be collected for gas chro-

matographic analyses.

Methods for the gas chromatographic analysis of mud, water, vegetation, and invertebrates.

Full descriptions of apparatus, reagents, and procedures are given in Rosenberg (1973).

Briefly, water samples were extracted basically according to Breidenbach et al. (1966, p. 22-

25) and injected into the gas chromatograph without cleanup. Extraction of mud was done

Quaest. Ent, 1975, 11 (1)

76

Rosenberg

according to Tyo (unpublished)^ and cleanup was basically as described in Barry et al. (1968,

section 211.15). Vegetation was extracted according to Barry et al. (1968, section 212.13b

and 212.1 5). Cleanup was the same as for the mud samples. Recovery studies using a 1 ppb

spike in water, 10 ppb in mud, and 100 ppb in vegetation gave average recoveries (3 replicates)

of 86.5, 81.1, and 83.0% respectively.

The confirmatory procedure used involved chemical conversion of dieldrin to a ketone by

boron trifluoride etherate (Skerrett and Baker, 1959; J. Singh^, personal communication).

Published accounts of extraction and cleanup procedures specifically for the gas chroma-

tographic analysis of chlorinated hydrocarbon residues in aquatic invertebrates are rare in the

literature. Therefore, a description of the method developed for invertebrates and used in

this study is given in Appendix I. To provide maximum sensitivity, invertebrate specimens

were pooled wherever necessary and possible (e.g. see Fredeen et al., 1971 ; Bradshaw et al.,

1972). In such instances, attempts were made to keep lower ranking taxa separate but where

it was obvious that it would be impossible to detect any residue unless a larger sample was

used, lower taxa were combined. Primary and secondary consumers were kept separate (ex-

cept for one sample of Hirudinea, see Table 9).

An SE30 column in the Varian 2100 gas chromatograph was used as a confirmatory pro-

cedure for invertebrate samples additional to a micro-coulometric system and the boron tri-

fluoride conversion (Rosenberg, 1973).

Concentrations of dieldrin in mud, vegetation, and invertebrates are on a wet-weight basis.

RESULTS

No dieldrin was detectable in mud, water, vegetation or invertebrates prior to application.

Dieldrin concentrations in the mud and water of D slough after application are shown in

Table 2. Residues in mud went below detectable levels between 47 days after application

(September 6, 1967) and the next date of sampling, 203 days after application (February 9,

1968). The actual time was probably closer to the first date given because concentrations

were less than 1 ppb at that time.

Table 2. Dieldrin concentrations (ppb) in mud and water of D slough°°.

* Solid underlining = confirmed with boron trifluoride etherate.

** Broken underlining = boron trifluoride etherate conversion unclear due to excessively dirty sample or sample with

insufficient dieldrin for confirmatory procedure.

+ Five times usual volume injected to get value.

++ Below experimental limits of detection.

Corrected to 100% recovery.

2. R. M. Tyo: Pacific Northwest Water Laboratory, Federal Water Pollution Control Admin-

istration, U. S. Department of the Interior, Corvallis, Oregon. See Rosenberg (1973) for

method.

3. Scientific Services Laboratory, Canada Department of Agriculture, Ottawa, Ontario.

See Rosenberg (1973) for method.

Fate of Dieldrin in Invertebrates

77

Maximum concentrations of dieldrin in water were reached the day after dieldrin applica-

tion and were below detectable levels shortly after 20 days after application (Table 2).

Residues in the submergent vegetation went below detectable levels some time between

47 days after application (September 6, 1967) and the first sampling of the following spring,

315 days after application (May 31, 1968) (Table 3). As in the mud samples, the time was

probably closer to the former because of the relatively low levels present at that time.

Table 3. Dieldrin concentrations (ppb) in vegetation of D slough°°.

* Solid underlining = confirmed with boron trifiuoride etherate.

+ Broken underlining = boron trifiuoride etherate conversion unclear because of excessively dirty sample or sample with

insufficient dieldrin for confirmatory procedure.

++ Below experimental limits of detection.

°° Corrected to 100% recovery.

Dieldrin concentrations in zooplankton were at a maximum on the day after treatment and

thereafter they declined steadily (Table 4). The initial concentration of dieldrin was the high-

est recorded for any ecosystem component for the study. Dieldrin residues were still detectable

at the end of the study, 424 days after dieldrin application.

*

Table 4. Dieldrin concentrations in zooplankton of D slough.

* The samples analyzed were composed mainly of Daphnia sp., Diaptomus sp., and Cyclops sp.

+ These samples combined to have sufficient weight of dieldrin for confirmation.

Dieldrin concentrations in Gastropoda of D slough are shown in Table 5. A number of sam-

ples showed no detectable residues because an insufficient weight of snail tissue was originally

used for extraction and cleanup. The negative confirmation for L. stagnalis sample number 2

for 348 and 349 days after application and H. trivolvis (Say) for 424 days after application

was due to the injection of an insufficient volume of sample (and hence weight of dieldrin)

into the microcoulometric apparatus. Highest residues generally occurred 2 days after dieldrin

apphcation and were still detectable in sufficiently large samples at the end of the study.

Tables 6 and 7 show residue concentrations in the Hemiptera of D slough. Residues in

Quaest Ent, 1975, 11 (1)

78

Rosenberg

adult Corixidae and Notonectidae were highest 2 days after dieldrin application. Residues were

below detectable levels in the immature corixids 349 days after application (July 4, 1968)

probably because the sample used was too small (Table 6). The larger sample of N. undulata

Say immatures 376 days after application (July 31, 1968) gave positive results (Table 7). How-

ever, the absence of detectable levels of dieldrin in adult C. audeni Hungerford and TV. undulata

in 1968 probably reflects a true absence because some of the samples analyzed were relatively

quite large (see Table 6, 349 days after application; and Table 7, 376 days after application).

Adult corixids and notonectids are capable of leaving and entering habitats and so the absence

of detectable dieldrin in 1968 is not surprising.

Table 5. Dieldrin concentrations in Gastropoda of D slough.

424 L. stagnalis

positive

Fate of Dieldrin in Invertebrates

79

Table 5. (concluded). Dieldrin concentrations in Gastropoda of D slough.

* Microcoulometric.

+ Varian Aerograph 2100 using 3% SE-30 on 100/120 mesh Aeropak 30 column packing.

** Below experimental limits of detection.

Table 6. Dieldrin concentrations in Corixidae of D slough.

* Microcoulometric.

+ Varian Aerograph 2100 using 3% SE-30 on 100/120 mesh Aeropak 30 column packing.

** Below experimental limits of detection.

++ C. audeni, C. alaskensis, H. atopodonta, S. decoraiella.

Dieldrin residues in larval Chironomidae are given in Table 8. Highest concentrations were

reached 21 days after application. Residues were still detectable in primary consumers at the

end of the study. It is likely that residues were not detected in the secondary consumers

(Tanypodinae) because of the small sample sizes.

Species of Hirudinea analyzed are shown in Table 9 and the results of the analyses in Table

10. The sample for 349 days after application (July 4, 1968) was too small to yield a detectable

dieldrin level. Maximum residue concentrations were reached 21 days after application, the

first date after application that leeches could be collected, and were still present when the

study ended.

Quaest. Ent, 1975, 1 1 (1)

80

Rosenberg

Table 7. Dieldrin concentrations in Notonecta undulata of D slough.

Fate of Dieldrin in Invertebrates

81

Table 8. (concluded). Dieldrin concentrations in larval Chironomidae of D slough.

+ Chironomus, Dicrotendipes, Einfeldia pagana & pectoralis grps., Glyptotendipes, Paratanytarsus.

++ Acricotopus, Chironomus, Glyptotendipes, Parachironomus, Cricotopus (“Paratrichocladius” type^, Psectrocladius,

unidentified Orthocladiinae genus.

+++ Chironomus, Dicrotendipes, Endochironomus, Glyptotendipes, Psectrocladius.

f Below experimental limits of detection.

Table 9. Species of Hirudinea used in gas chromatographic analyses of dieldrin concentrations

in D slough.

Names of Zygoptera and Libellulidae used in gas chromatographic analyses for each date

are given in Tables 1 1 and 12. Results of the analyses of Odonata are shown in Table 13.

Residues reached maximum concentrations in Anisoptera in 2 1 days and in Zygoptera 2 days

after application. Residues were still present at the end of the study.

Names of larval and adult Dytiscidae used in gas chromatographic analyses for each date

are given in Tables 14 and 1 5. The results of these analyses are given in Table 16. As with the

adult Corixidae and Notonectidae, adult Dytiscidae are capable of moving in and out of habi-

tats. Therefore, the origin of the high residue value in the sample from 376 days after applica-

tion (July 31, 1968) is questionable. Maximum residues occurred in the larvae 2 days after

application and in the adults 48 days after application. Residues were still present in both

life stages at the end of the study.

Quaest. Ent., 1975, 1 1 (1)

82

Rosenberg

Table 10. Dieldrin concentrations in Hirudinea of D slough.

Fate of Dieldrin in Invertebrates

83

Table 12. (concluded). Species of Zygoptera used in gas chromatographic analyses of dieldrin

concentrations in D slough.

Table 13. Dieldrin concentrations in larval Odonata of D slough.

Quaest. EnL, 1975, 11 (1)

Days after Dieldrin

Application Sample Size Concentration Confirmation

(g) (ppb) BF^ Etherate

Fate of Dieldrin in Invertebrates

85

95% confidence limits for sampling dates which have three or more replicates of the same

type of sample (for all environmental components of the study) are given in Table 17. The

reader’s attention is drawn to the excellent reviews of Kenaga (1972, 1973) and Moriarty

(1972) for a discussion of the factors that can cause variability in determinations of pesticide

residues.

Table 17. 95% confidence limits for dieldrin concentrations in mud, water, vegetation, and

invertebrates from sampling dates with three replicates or more.

C. rostrata

C. demersum and L. trisulca

DISCUSSION

Table 18 summarizes some parameters of this and other studies done on the fate of the

chlorinated hydrocarbons added to whole ecosystems. It compares times at which maximum

concentrations are reached and pesticide is no longer detectable in mud and/or sediment.

“Length of Time until Maximum Concentration” should be interpreted with caution because

the different authors did not use equivalent time intervals in their analyses. Maximum con-

centration has been used here because, for most studies, the actual amount of pesticide that

entered the water after application is not known^. Data are insufficient to correlate maximum

concentration with length of time until residues were undetectable. However, an indication

exists that lower concentrations became undetectable faster (see Croker and Wilson, 1965).

Again, caution should be used in drawing such conclusions because sensitivity of most of the

analytical methods used is unknown.

It is apparent from Table 19 that maximum concentrations and the time until residues are

detectable are reached sooner in water than in any other ecosystem component and these

rates would seem to be independent of initial concentration. The maximum mean concentra-

tion of 0.35 ppb in the water of D slough was below the concentration intended (1.0 ppb).

However, Bridges et al. (1963) and Meeks (1968) showed that the maximum concentration

of DDT in the water samples studied by them was reached in the first hour after treatment

4. In both Vaajakorpi and Salonen (1973) and this study the insecticide was added below

the water surface.

Quaest. Ent., 1975, 11 (1)

86

Rosenberg

and by the following day, levels were approximately 1/25 and 1/2 the maximum concentra-

tion respectively. Thus, almost certainly the maximum concentration of dieldrin in the water

of D slough was greater than 0.35 ppb. The rapid decline of chlorinated hydrocarbon concen-

trations is due to two major processes: their relative insolubility in water, compared to organ-

ic matter, which causes them to preferentially attach to organic matter and be removed from

the water (e.g. by settling to the bottom) (Edwards, 1970; Muirhead-Thomson, 1971 ; Cope,

1971) and co-distillation (Eberhardt, Meeks, and Peterle, 1971).

Table 18. Some parameters of and decline of chlorinated hydrocarbon pesticide residues in

mud and/or sediment of studies cited.

Mud not sampled until 16 days after application of endrin.

Bridges et al. (1963) and Meeks (1968) continued to detect residue levels in submergent

and emergent vegetation in the year following application whereas I did not (Table 20). [The

maximum concentrations given for Meeks (1968) and Vaajakorpi and Salonen (1973) are

the highest of the species of submergent and emergent vegetation analyzed. For Meeks (1968),

the length of time until residues could not be detected is the time of the final sample for the

species which had the lowest level of pesticide] . This could be a function of the low initial

concentration of my study. As in Meeks (1968), submergent vegetation of my study had

higher levels of residues than emergent. This is a function of the surface area available for

the pesticides to be adsorbed (Meeks and Peterle, 1967). Among the studies in Table 20

Fate of Dieldrin in Invertebrates

87

Table 19. Decline of chlorinated hydrocarbon pesticide residues in water of studies cited.

* No vegetation samples taken until 16 days after endrin applied.

** Sample not taken until 3 days after application,

t See discussion in text.

Quaest. Ent., 1975, 1 1 (1)

88

Rosenberg

“Time to Maximum Concentration” varies but indications are that it occurs within the first

day after application. Also, there are indications that the higher initial concentrations persist

longer than the lower ones.

Undoubtedly, the initial high concentration of dieldrin in zooplankton was due to the high

surface to volume ratio of zooplankton (Kenaga, 1973). Crosby and Tucker (1971) reported

accumulations of DDT from dilute suspensions in water of 16,000 to 23,000 times by Daph-

nia magna Strauss. Wojtalik, Hall, and Hill (1971) reported uptake of high concentrations of

2, 4-D by plankters. Johnson et al. (1971) reported the rapid direct uptake of aldrin and

DDT from water by the freshwater invertebrates of their study and that D. magna and Culex

pipiens L. showed the greatest degree of magnification. Sanders and Chandler (1972) found

that D. magna had the highest magnification factor of the eight species of invertebrates ex-

posed to PCB’s in the ppb range. Hughes and Lee (1973) reported that net plankton accumu-

lated large amounts of toxaphene. The differential solubility of chlorinated hydrocarbon in-

secticides causes them to move directly to organic matter as soon as they are applied to water

(Dustman and Stickel, 1969; Edwards, 1970; Muirhead-Thomson, 1971;Cope, 1971) and it

has been shown that these hydrophobic compounds adhere to suspended organic particles in

the water (Nicholson, 1967; Stickel, 1968; Wurster, 1969; Edwards, 1970). Vaajakorpi and

Salonen (1973) showed that 3 p diameter seston had higher DDT concentrations than 8 p

diameter seston and had the highest DDT concentrations of any of the ecosystem components

they analyzed. Reinert (1967) showed that Daphnia accumulated more dieldrin from water

than from food. Plankton, therefore, merely serve as immediately accessible suspended or-

ganic particles to which chlorinated hydrocarbons move directly when applied to water. In

fact, because of their lipid content, greater chlorinated hydrocarbon deposition may occur in

plankton than in other organic seston (Cope, 1971 ; see also Kawatski and Schmulbach, 1971).

Residues in the planorbids and Lymnaea sp. of Meeks’ (1968) study reached a maximum

concentration 1 and 3 days respectively after application of the DDT-Cp^. The snails of my

study reached maximum observed residue concentrations on the first sampling date, which

was 2 days after application. Maximum concentrations in Meeks’ study were among the low-

est of all the invertebrates. Those of the L. stagnalis of my study were second only to the

zooplankton. I used only soft tissues for analysis whereas Meeks probably used the whole

body. Because the shell is such a large percentage of the total weight, and because later analyses

by Meeks of planorbids showed the shells had no residues, the concentrations reported by

Meeks are likely underestimates. Meeks reported the presence of residues in both kinds of

snails at the end of the study (13 months after application). Eevels were still detectable in

L. stagnalis and H. trivolvis at the end of my study (approximately 14 months after applica-

tion).

Residues in notonectids of Meeks’ (1968) and Vaajakorpi and Salonen’s (1973) studies

were at a maximum 1 week and 2 to 4 days respectively after application of the DDT. In my

study, maximum concentrations were reached in adults on the second day after application.

Meeks recorded no residue levels in the notonectid samples he studied in the second and twelfth

months after application, and a relatively low level in the thirteenth month. Vaajakorpi and

Salonen continued to detect DDT in Notonecta sp. until their last sampling (30 days after

application). Neither works indicated whether the notonectids were adults or immatures.

Because adult corixids and notonectids can fly, their reliability as indicators of pesticide levels

in habitats is questionable. Similar rationale (i.e. that movement of the indicator organism be

restricted to the area under study) was used by Gish (1970), Foehrenbach (1972), and Coulson

et al. (1972) in choosing the indicator species of their studies.

The maximum concentration of residues in the bloodworm (Tendipes sp.) of Meeks’ (1968)

study was reached 1 week after application. In my study, maximum concentration was reached

Fate of Dieldrin in Invertebrates

89

in 3 weeks. This delay was also shown by the Libellulidae, Aeshna interrupta Walker, and

adult Dytiscidae of my study (see below). Unfortunately, analysis of chironomids in Meeks

(1968) was ended 1 month after application. The residues in the chironomid primary consumers

of my study were still detectable at the end of the study (approximately 14 months after ap-

plication).

Meeks (1968) analyzed Erpobdella punctata, a species of Hirudinea present in D slough

and used in my analyses of residues (see Table 9). He reported that a maximum concentration

of residue was reached 2 weeks after application of the pesticide and that E. punctata had

the highest residues of all the invertebrates analyzed. In my study, the Erpobdellidae, the

family to which E. punctata belongs, had lower residue levels in general than the Glossiphoni-

idae and the leeches did not have the highest residues of the invertebrates. Residues were still

present in leeches of both studies when the studies were terminated (Meeks’ — 1 5 months;

and mine — approximately 14 months after application).

Residues in the Anisoptera and Zygoptera larvae of Meeks’ (1968) study reached maximum

concentrations 1 week after pesticide application. Vaajakorpi and Salonen (1973) reported

maximum concentrations in Odonata larvae 2 weeks after application of the DDT. In my study,

maximum concentrations were detected in Anisoptera in 3 weeks and in Zygoptera in 2 days.

Residues were still present in the Odonata at the termination of the studies (Meeks’ — 1 3

months; Vaajakorpi and Salonen’s — 59 days; and mine - 14 months after application).

Neither Meeks (1968) nor Vaajakorpi and Salonen (1973) presented any data for Dytiscidae.

Tables 21 and 22 show maximum and final pesticide concentrations of primary and secon-

dary consumers in Meeks (1968), Vaajakorpi and Salonen (1973), and this study. (I hesitate

to accept the results of the adult Dytiscidae analyses shown in Table 22 because they are the

only group that showed a dramatic rise at the end of the study; and because they are capable

of entering and leaving the slough so their role as indicators of pesticide relationships in the

slough is open to question). It is evident from these tables that similar ranges of pesticide con-

centrations exist between primary and secondary consumers. The reality of food chain con-

centration of chlorinated hydrocarbon insecticides in invertebrate communities is discussed

in Rosenberg (1975).

Table 21. Maximum and final DDT concentrations (ppm) in invertebrates of Meeks (1968)

and Vaajakorpi and Salonen (1973).

Arbitrarily called primary consumers. They are scavengers or feed on aquatic invertebrates (Pennak, 1953; A. R. Smith,

Alberta Department of Lands and Forests, Fish and Wildlife Division, Edmonton, Alberta; unpubUshed). That is, they

are “opportunistic” feeders.

From Vaajakorpi and Salonen (1973). The other data are from Meeks (1968).

90

Rosenberg

Table 22. Maximum and final dieldrin concentrations (ppb) in invertebrates of this study.

Below experimental limits of detection.

The decline of dieldrin concentrations in mud, water, vegetation, and invertebrates of D

slough is a result of complex interactions of physical and biological processes. This subject

is reviewed in Rosenberg (1973). In general, the declines of dieldrin residues in mud, water,

vegetation, and invertebrates in D slough follows the form of the first order kinetics model

described for acute applications by Eberhardt et al. (1971). [See Rosenberg (1973) for graphs

of dieldrin declines in the ecosystem components discussed above]. Cooke (1973) noted that

repeated (chronic) dosing would cause a greater proportion of the pesticide to be stored in

fatty tissues from which residues would be lost slowly whereas in a single (acute) dose the

residues are lost quickly because only low concentrations are accumulated in the fat. Residues

in the ecosystem components of this study, therefore, are mostly in the “fast compartment”

of Eberhardt et al.’s (1971) model.

SUMMARY AND CONCLUSIONS

1. Levels of dieldrin were below detection in mud, water, and vegetation of D slough after

47 days after application. Dieldrin levels were detectable in zooplankton. Gastropoda,

larval Chironomidae, Hirudinea, larval Odonata, and larval and adult Dytiscidae at the

end of the study (= 1 year) but were below detection in adult Corixidae and Notonec-

tidae before the end of the study.

2. Zooplankton had the highest initial concentration of dieldrin of any ecosystem constitu-

ent.

3. Mud, water, vegetation, and primary and secondary invertebrate consumers carried

similar levels of dieldrin in their tissues.

Fate of Dieldrin in Invertebrates

91

4. Dieldrin residues in mud, water, vegetation, and invertebrates of D slough declined

according to a first order kinetics equation, as found by other authors. These declines

were influenced by the hydrophobic nature of chlorinated hydrocarbons and the inter-

action of physical and biological processes.

5. The fate of chlorinated hydrocarbon insecticides in other small, standing water ecosys-

tems was similar to the fate of dieldrin in the ecosystem of this study.

ACKNOWLEDGEMENTS

I would like to thank W. G. Evans whose original concept formed the basis of this study;

for his advice, continual encouragement, and the many helpful discussions we have had. Thanks

are also due to the other members of my advisory committee for their help in producing the

thesis which formed the basis of this paper: G. E. Ball, Department of Entomology, University

of Alberta; H. F. Clifford and J. R. Nursall, both of the Department of Zoology, University

of Alberta; and W. F. Allen, Department of Chemistry, University of Alberta. F. A. J. Arm-

strong, M. Healey, and R. D. Hamilton of the Freshwater Institute, Winnipeg, G. R. B. Web-

ster, Pesticide Research Unit, University of Manitoba, and D. Pimentel, Cornell University,

critically read the manuscript. I have used many of their suggestions.

I am indebted to the people who helped me identify the invertebrates of this study: J. R.

Morris (Gastropoda), Department of Zoology, University of Alberta; A. R. Smith (Hirudinea),

Alberta Department of Lands and Forests, Fish and Wildlife Division, Edmonton, Alberta;

J. S. Carr (Coleoptera), R.R. No. 4, Calgary, Alberta; A. L. Hamilton and O. A. Saether

(Chironomidae), both of the Fisheries Research Board of Canada, Freshwater Institute, Win-

nipeg, Manitoba; P. S. Corbet (Odonata), Department of Biology, University of Waterloo,

Waterloo, Ontario.

V. W. Kadis, O. J. Jonasson, W. Currie, and W. Breitkreitz, all of the Alberta Department

of Agriculture, Dairy and Food Laboratory, Edmonton, Alberta, gave me considerable help

and advice with the gas chromatographic aspects of the study.

L. L. Kennedy and M. Dumais, both of the Department of Botany, University of Alberta,

identified plants; and D. Christophel, Department of Botany, University of Alberta, helped

describe the study site from an eco-botanical aspect.

H. Hall, L. Hill, T. McMahon, K. Morgan, W. Skrlac, and D. Wales provided technical help.

Special thanks are due to R. Henschel and D. Swanek for letting me work on their proper-

ties.

G. Forth and C. Plumridge, Freshwater Institute, Winnipeg, typed drafts of the manuscript.

A. P. Wiens and C. Madder helped proofread the drafts.

This research was supported by National Research Council of Canada and Canadian Wild-

life Service grants to Dr. W. G. Evans. I am pleased to acknowledge my appreciation to the

National Research Council of Canada for scholarships awarded.

APPENDIX I.

METHOD FOR THE GAS CHROMATOGRAPHIC ANALYSIS OF INVERTEBRATES

A: Apparatus and reagents - See Kadis, Jonasson, and Breitkreitz (1968) and Rosenberg

(1973).

B: Procedure — Invertebrates were divided into two types for purposes of maceration:

those with hard exoskeletons (e.g. Corixidae, Notonectidae, and adult Dytiscidae) and those

with soft exoskeletons (e.g. Hirudinea, Gastropoda removed from their shells, zooplankton, and

Quaest. Ent., 1975, 11 (1)

92

Rosenberg

Chironomidae and Odonata immatures). After surface moisture was removed from each sam-

ple by blotting with paper toweling, the former were combined with sand and anhydrous

Na2S04 (for 2 g invertebrate tissue, approximately 5 g sand and 10 g Na2S04) and ground

with the Omni-Mixer at medium speed for about 5 min or until a visually homogeneous mix-

ture resulted (Jonasson and Rosenberg, 1969). The mixture was then added to 50 g deactivated

florisil in a mortar and pestle, the Omni-Mixer cup and blades were rinsed with petroleum

ether, and the rinsings were added to the mortar and pestle. The invertebrates with soft exo-

skeletons were ground directly in 50 g of deactivated florisil in the mortar and pestle. For

soft and hard types, the mixture from the pestle was then added to a chromatographic column

containing 50 g deactivated florisil prewashed with 150 ml of 1 : 1 methylene chloride-petroleum

ether. The column was eluted with 800 ml 20:80 methylene chloride-petroleum ether. The

eluant was evaporated to near dryness, made up in a suitable volume, and injected

(Kadis, et al., 1968). When necessary, samples were further cleaned up on a magnesium

oxide-celite column as described in Barry et al. (1968, section 21 1.16c) and/or by the follow-

ing method: 10 g deactivated florisil sandwiched between two 2.5 cm portions of anhydrous

Na2S04 prewashed three times with 25 ml petroleum ether. The sample was added to the

column and the column was then eluted with 1 50 ml 5% benzene in petroleum ether and then

200 ml of 25% benzene in petroleum ether, the second fraction containing the dieldrin.

Recovery studies were done using Hyallela azteca (Saussure) from the control slough. Sam-

ples spiked with 0.003 jug dieldrin (0. 1 2 X 10’^ ppm in 2.42 g tissue) gave an average recovery

of 95.2% (3 replicates).

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FOOD CHAIN CONCENTRATION OF CHLORINATED HYDROCARBON

PESTICIDES IN INVERTEBRATE COMMUNITIES: A RE-EVALUATION

D. M. ROSENBERG*

Department of Entomology

University of Alberta Quaestiones Entomologicae

Edmonton, Alberta T6G 2E3 11: 97- 110 1975

Concentration of chlorinated hydrocarbon pesticides in invertebrate food chains is not

shown by results reported in the literature. It is proposed that habitat and mode of life rather

than trophic level are likely the most important ecological characteristics determining uptake

and final concentration of chlorinated hydrocarbon pesticides in aquatic invertebrates and

that the idea of “food chain concentration” or “trophic level effect” should be replaced by

the idea of “bioconcentration”.

II nest pas encore demontre au travers des nombreuses publications qu’il y a un rapport

chez les invertebres entres niveau trophique et la concentration des pesticides en hydrocar-

bures chlorines. Nous proposons que Vhabitat et le mode devie plustot que le niveau trophique

est a la base de Vabsorption et de la concentration finale des hydrocarbons chlorines chez

les invertebres aquatiques et que Videe de “la concentration par niveau trophique” ou “Veffet

du niveau trophique” soit remplacee par Videe de “bioconcentration”.

INTRODUCTION

A study of the fate of dieldrin in ecosystem components of a slough in central Alberta

revealed that the range of concentrations of the pesticide was similar in primary and secondary

consumer invertebrates over the duration of the study (Rosenberg, 1975; Table 22). The

results of similar studies (Meeks, 1968; Vaajakorpi and Salonen, 1973) revealed the same

lack of trophic level effect (Rosenberg, 1975; Table 21).

These results were surprising in view of the generalizations that have existed in the pesti-

cide literature over the last decade or longer regarding the inevitability of food chain concen-

tration of pesticides, especially chlorinated hydrocarbons. Thus, I undertook a literature sur-

vey to determine whether or not the results referred to above were atypical.

Because of the immensity of the literature dealing with the effects of pesticides on fauna,

the survey dealt almost entirely with the chlorinated hydrocarbons, supposedly the chief

offenders in food chain concentration, and mainly considered freshwater invertebrate com-

munities. However, I have given some consideration to pesticide uptake by fish and terrestrial

and marine invertebrate communities. In general, only those papers which contained a level

of identification sufficient to designate the trophic level to which the animal belonged were

used. Other papers are only briefly mentioned.

I have also limited the review to an analysis of the results of field studies and those labora-

tory studies which have allowed feeding to occur. The use of studies which have allowed

feeding is obviously necessary for a consideration of trophic level effects. Studies which have

not allowed feeding have already shown that aquatic invertebrates readily concentrate pesti-

cides from water (e.g. Johnson et al., 1971 ; Wilkes and Weiss, 1971 ; Derr and Zabik, 1972,

* Present Address: Fisheries & Marine Service, Freshwater Institute, 501 University Crescent,

Winnipeg, Manitoba R3T 2N6.

98

Rosenberg

1974). Also, the analysis of studies which have allowed normal trophic relations gives this

review a degree of comparability with the studies from which the generalizations about food

chain concentration of pesticides originated (e.g. see Rudd, 1964). Admittedly, this approach

introduces a degree of circumstantiality into the interpretation. However, in the absence of

experiments which quantify the relative contributions made by food and other factors (see

below) to the pesticide levels in aquatic invertebrates, I believe the approach is useful.

Variability of residue concentrations in field-collected samples analyzed by gas chromato-

graphy is well known (e.g. see Moriarty, 1972; Kenaga, 1972; Rosenberg, 1975, Table 17).

Therefore, I have considered differences in residue concentrations from primary to secondary

consumer trophic levels to be significant only when these differences are at least an order of

magnitude (10-fold).

Finally, I have found that studies of food chain concentrations of pesticides have the fol-

lowing shortcomings which make interpretation of the results difficult and which should be

identified at the outset of this review:

1. Changing food habits during a particular life stage and over the lifetime of an animal

are usually not considered. For example, different life stages of invertebrates are analyzed

simultaneously for pesticide residues. Because specific information on the extent of an

animal’s mode of feeding prior to residue analysis is usually unavailable, we assume that

its major mode of feeding is its only one.

2. Most studies do not correlate a predator with its actual prey. Rather, we assume that

of the animals designated as primary consumers some will be eaten by those designated

as secondary consumers.

3. The grouping of invertebrates into primary and secondary consumer trophic levels is

arbitrary (as are other classification systems of trophic relationships; see Cummins,

1973) and oversimplifies these relationships.

LITERATURE SURVEY

Despite the findings of Godsil and Johnson (1967) that a low concentration of endrin in

the lake of their study did not result in food chain concentration, the works of Hunt and

Bischoff (1960), Fillmore (in Rudd, 1964), Hunt (in Rudd, 1964), Bridges, Kallman, and

Andrews (1963), and Hickey, Keith, and Coon (1966), among others, have reported a trophic

level effect resulting from pesticide applications to aquatic ecosystems. Rudd (1964) has

discussed instances of trophic level effects in terrestrial ecosystems. However, in each of these

studies, invertebrates have been used as a single step in the food chain and usually they are

of a single species or are zooplankton. Other studies (Terriere et al., 1966; Keith, 1966; and

those reviewed and summarized in Table IV of Moore, 1967) which have also used aquatic

invertebrates as a single step, have presumably lumped a number of species of aquatic inver-

tebrates of different trophic levels. Of course, aquatic invertebrates have their own trophic

interrelationships (e.g. see Jones, 1949).

Hannon et al. (1970) separated the aquatic invertebrates of their study into three unlikely

groups: plankton-algae, crayfish, and aquatic insects (composed of midge larvae and Gyrinidae)

so no information on trophic distribution of the chlorinated hydrocarbons is available. Wood-

well, Wurster, and Isaacson’s (1967) study of the DDT residues in an east coast estuary gives

an extensive list of residue data for various species of invertebrates but, unfortunately, none

can be classed as secondary consumers. The same is true of the review and summary presented

in Table 12 of Edwards (1970) except for the 1964 United States Department of the Interior

study which gives a concentration factor for a crab that is the lowest for the entire study. None

of the aquatic invertebrates listed in Table 4 of Flickinger and King (1972) are predators except

Food Chain Concentration of Pesticides

99

for the Notonectidae which have been combined with Corixidae under the heading “Aquatic

Hemiptera”. The range of dieldrin concentrations in the crayfish from Brazoria County in

Flickinger and King (1972) spans the concentrations given for all the aquatic invertebrates

in Table 4 (none detectable to 17.0 ppm). Also, no secondary consumers are present among

the terrestrial invertebrates in Table 10 of Edwards (1970) except for Davis and Harrison’s

(1966) work which will be considered below.

Moubry, Helm, and Myrdal (1968) reported similar DDT, DDT-metabolites, dieldrin, and

endrin levels in Gammarus sp., Limnephilus rhombicus (L.) larvae, and Sialis sp. larvae (Table

1). The last, of course, are predators. Dieldrin residues in some invertebrates exposed to a

dieldrin industrial effluent entering the Rocky River, South Carolina, are shown in Table 2

(Wallace and Brady, 1971). It can be seen that the predaceous hellgrammite larvae, Corydalis

cornuta, had lower levels than the filter-feeding blackfly (Simulium vittatum) and caddisfly

larvae {Hydropsyche sp.). Robinson et al. (1967) reported similar concentrations of DDE and

dieldrin in primary and secondary consumer marine invertebrates except in macrozooplankton

which they classed as a secondary consumer and which had extraordinarily high concentrations

(Table 3). Robinson et al.’s results are shown diagramatically in Fig. 3 of Edwards (1970).

The similarity in pesticide levels between trophic levels 2 (= primary consumer: 0.02 ppm)

and 3 (= secondary consumer: 0.03 ppm) is striking. Although Foehrenbach (1972, p. 622)

claimed higher dieldrin concentrations existed in invertebrates listed in Table III of his paper

than those of the shellfish listed in Table II “. . . probably because the organisms in Table III

are higher in the food chain. . .” the range of concentrations is similar (Table II: 0 to 0.132

mg/kg; Table III: 0.004 to 0.236 mg/kg; the 0.236 value was for a non-predaceous grass shrimp).

Also, Table III contains invertebrates that are primary consumers. Naqvi and de la Cruz (1973)

analyzed mirex residues in a variety of aquatic, terrestrial, and estuarine invertebrates and

vertebrates in Mississippi. In order to calculate mean levels of mirex in herbivore, carnivore,

and omnivore trophic levels, the authors combined animals from pond, lake, creek, grassland,

and estuarine habitats and lumped invertebrate carnivores with vertebrate carnivores. The

mean residue levels presented for these three trophic levels (Table 2 of Naqvi and de la Cruz)

and the conclusions reached are, therefore, difficult to interpret. The authors implied food

chain concentration occurred even though the increase in mean residues for the three trophic

levels was not in the expected order (0.23, 0.30, and 0.35 ppm for herbivores, carnivores, and

omnivores respectively). Furthermore, these residue levels are virtually identical in view of

the precision possible for samples from the field analyzed by gas-liquid chromatography (see

Kenaga, 1972; Moriarty, 1972). However, it is possible to examine whether or not food chain

concentration has occurred by using the residue data given for species of aquatic invertebrates.

Only residue data for the simultaneous presence of the three trophic levels of aquatic inver-

tebrates is used (Table 4). It can be seen that residue levels in carnivores were highest in only

one of four locations, The number of replicates in this location was low for all three trophic

levels. Fish {Gambusia affinis and Lepomis cyanellus) from Bluff Lake had a mean mirex

residue of 0.19 ppm (range: 0.07-0.38 ppm; 3 replicates). Fish (same two species as above)

from a pond in the Louisville and Noxapater areas had a mean residue of 0.39 ppm (range:

0. 17-1.00 ppm; 2 replicates) whereas a single mirex residue in an aquatic invertebrate herbi-

vore was 0.05 ppm and a mean mirex residue of 0.16 ppm (range: 0.07-0.26 ppm; 4 replicates)

was present in aquatic invertebrate carnivores in the same area. The single relatively low herbi-

vore residue value is difficult to interpret in view of the ranges of residues reported for aquatic

invertebrates in Naqvi and de la Cruz. Collins, Davis, and Markin (1973) reported similar mir-

ex residues in crayfish (range: 0.01 to 0.40 ppm) and dragonfly nymphs (<0.01 to 0.70

ppm).

Quaes t.Ent., 1975, 11 (1)

100

Rosenberg

Table 1. Chlorinated hydrocarbon pesticide residues in invertebrates of Moubry et al.’s (1968)

study.

Table 2. Dieldrin residues in invertebrates of Wallace and Brady’s (1971) study*.

Organism Residue (ppm)+

Position in Relation to Effluent

Upstream Downstream

* Data is for the April 18, 1970 collection, the only date with residue levels for more than a

single trophic level.

+ Values shown are means of replicates and the two gas chromatographic columns used.

Table 3. Concentrations of organo chlorine compounds in marine invertebrate samples taken

off the Northumberland Coast, 1965-1966 (adapted from Robinson et al., 1967).

Food Chain Concentration of Pesticides

101

Table 4. Mirex residues in aquatic invertebrates of Naqvi and de la Cruz’s (1973) study.

+ All Mississippi.

* Mean value is followed by range in parentheses, and number of replicates.

Thus far, only the results of field studies have been discussed. Many laboratory studies

have been done on the uptake and accumulation of chlorinated hydrocarbon pesticides by

invertebrates but most of these have not allowed feeding. Only the model ecosystem studies

(Metcalf, Sangha, and Kapoor, 1971) attempt to duplicate a field situation. Of the many

model ecosystem studies consulted, only that of Sanborn and Yu (1973) has used more than

one trophic level of invertebrates. Concentrations of dieldrin were highest in the snail {Physa

sp. — 229.87 ppm), followed by alga (Oedogonium cardiacum — 14.96 ppm), fish {Gambusia

affinis — 12.29 ppm), Daphnia sp. (5.07 ppm), Elodea sp. (2.56 ppm), clam {Corbicula mani-

lensis — 2.03 ppm), and crab {Uca minax — 0.495 ppm). In fact, results of the model eco-

system, studies of Kapoor et al. (1972, 1973), using an alga {Oedogonium cardiacum) - snail

{Physa sp.) — mosquito {Culex pipiens quinquefasciatis) - fish {Gambusia affinis) food chain

do not follow the classical concept of food chain accumulation (Table 5).

Table 5. Concentrations of chlorinated hydrocarbons in food chain elements of Kapoor et

al. (1972, 1973) model ecosystem studies.

Reference Compound Concentration (ppm)

* Concentrations are total values for each compound.

Quaest. Ent., 1975, 11 (1)

102

Rosenberg

There is some evidence that findings of an apparent lack of food chain concentration of

chlorinated hydrocarbons (and closely related pesticides) can be extended to fish occupying

primary and secondary consumer trophic levels. A number of relatively recent papers have

reported that fish do not exhibit food chain concentrations of chlorinated hydrocarbon

pesticides. Fredeen, Saha, and Royer (1971) found no difference in the concentration of

organochlorine residues in fishes at the end of the food chain (walleyes, saugers, and northern

pike) and those lower down (white and longnose suckers, northern redhorse, burbot, goldeye,

and yellow perch). Levels of DDT and DDD ranged from < 0.01 to 0.05 ppm, of DDE from

< 0.01 to 0.06 ppm, and of dieldrin from 0.002 to 0.006 ppm in fish from all trophic levels.

Morris and Johnson (1971) found concentrations of 1600 ppb dieldrin in channel catfish. Other

rough fish (buffalo, carp, and carp suckers) had concentrations ranging from 1 5 to 840 ppb

dieldrin while the pan and gamefish (largemouth bass, black and white crappie, black bullhead,

bluegill, walleye, and northern pike) had the lowest concentrations (11 to 175 ppb). This latter

group, of course, contains several “top predators”. Hughes and Lee (1973) concluded that ei-

ther a clear delineation of toxaphene levels was absent between prey and predator fish or prey

fish accumulated higher toxaphene concentrations than the predators (e.g. results from Fox

Lake, 6 months after stocking: bluegill and sucker: 9.4 to 10.6 jug/g; bass, northern pike, and

walleye: 2.2, 2.3, and 1.2 jug/g)- The results of the study of Frank et al. (1974) remind us that

factors other than feeding habits are involved in the accumulation of chlorinated hydrocarbons

in fish. DDT and dieldrin levels in fish taken from a variety of locations in Ontario, Canada

tended to depend on fat content and weight (age) of the fish which, in turn, are related to

feeding habits. Risebrough and deLappe (1972) reported that polychlorinated biphenyl (PCB)

levels on a fresh weight basis in Atlantic herring were an order of magnitude higher than levels

in cod although the latter occupies a higher trophic level. PCB concentrations were comparable

on a fat basis (Table 6). Risebrough and deLappe (1972, p. 43) concluded that “the amounts

and kinds of lipids may affect the retention of PCB’s, modifying the trophic accumulation

predicted by the classical food chain concentration theory. Consistent with this hypothesis

are the higher PCB residues measured in extractable lipids of the North Atlantic plankton

than in the lipids of fish from the same area.”

Table 6. Levels of PCB in Atlantic herring and cod (from Risebrough and deLappe, 1972).

The number of adequate field studies on uptake of pesticides in different trophic levels

of terrestrial invertebrates is equally as low as for aquatic invertebrates. The residue values

presented by El Sayed, Graves, and Bonner (1967) show conflicting patterns probably be-

cause of a lack of collection consistency more than anything else. However, the studies of

Davis and Harrison (1966) and Davis (1968) have shown that “. . . worms and slugs usually

contain higher amounts and a greater range of organochlorine compounds than beetles”

(Davis, 1968; p. 43 to 44). The beetles he referred to were mostly Carabidae as well as some

Staphylinidae and Elateridae. Carabidae and Staphylinidae are predatory. Korschgen (in

Food Chain Concentration of Pesticides

103

Dustman and Stickel, 1969) reported similar levels of dieldrin in earthworms, crickets, and

carabids in a Missouri field which had had long-term aldrin applications. Gish (1970) analyzed

earthworms of the genera Allolobophora, Diplocardia, Helodrilus, and Lumbricus; white grubs

(Scarabaeidae larvae); slugs belonging to the genera Deroceras and Umax', and unidentified

terrestrial snails. Unfortunately, he did not analyze any predators. Average concentrations

of chlorinated hydrocarbons were 0.6 ppm for the Scarabaeidae larvae, 3.5 ppm for snails

(shells included), 13.8 ppm in earthworms, and 89.0 ppm in slugs. Highest levels were 180

times the lowest levels, all for non-predatory forms. Gish (1970) credited the differences in

chlorinated hydrocarbon levels to dissimilar feeding habits. Table VII of the review paper by

Edwards and Thompson (1973) summarized the results of a large number of pesticide analyses

of earthworms, slugs, and beetles (mostly Carabidae). The results are summarized in Table 7.

It can be seen that the range of concentrations for the beetles never exceeds that of the earth-

worms and slugs, something expected of a trophic level effect.

Table 7. Range of concentrations of DDT and dieldrin in earthworms, slugs, and beetles in

Table VII of Edwards and Thompson (1973).

Kenaga (1972) has noted that chlorinated hydrocarbon pesticides best illustrate the mech-

anisms of bioconcentration of heavy metals and other environmental contaminants. Nonethe-

less, Moriarty (1972) stated that similar conclusions regarding the lack of a trophic level effect

have sometimes been reached for mercury and radionuclides. The literature on uptake and

accumulation of heavy metals (e.g. mercury and arsenic) is inconsistent on this point and a

review is outside the scope of this paper. By way of speculation, however, it would be sur-

prising if uptake and accumulation of trace and heavy metals by invertebrates was mainly

due to a trophic level effect.

Thus, it appears that of the studies which have adequately dealt with chlorinated hydro-

carbon pesticide residues in different trophic levels of aquatic or terrestrial invertebrates, a

trophic level effect or food chain concentration has not been adequately demonstrated. Yet,

unqualified generalizations about food chain concentration of pesticides keep appearing in

the literature (e.g. Moore, 1967, p. 113;Dimond, 1969, p. 2, 6; Wurster, 1969, p. 125; Wilkes

and Weiss, 1971, p. 223; Foehrenbach, 1972, p. 619, 622, 623, 624; Khan et al., 1973, p.

159, 166; Leland, Bruce, and Shimp, 1973, p. 833; Metcalf, 1973, p. 512).

In his review, Kenaga (1973, p. 80) concluded that “Maximum pesticide residues may

sometimes be accumulated by algae or by similar ‘first link’ organisms in the chain-of-life

organisms and do not necessarily result in increasing concentrations in each succeeding link

of the chain.” Moriarty (1972), in his review of the effects of pesticide residues on wildlife

has severely criticized the concept of food chain accumulation of pesticides and other toxic

chemicals. He concluded (p. 267): “It is unlikely that predators accumulate the insecticide

contained in their prey. All the evidence suggests that aquatic predators acquire their insect-

icide directly from the water, not from their food.” He questioned the validity of a trophic

level effect in any food chain. The results and discussion here would lend support to his

Quaest. Ent, 1975, 11 (1)

104

Rosenberg

contention.

Aquatic organisms acquire pesticides from their surroundings and through their food according

to Moore ( 1 967), Chadwick and Brocksen ( 1 969), Dustman and Stickel ( 1 969), Macek ( 1 969),

Edwards (1970), Cope (1971), Hamelink, Waybrant, and Ball (1971), Kawatski and Schmul-

bach (1971), Wilkes and Weiss (1971), Moriarty ( 1 972), and Booth, Yu, and Hansen ( 1 973).

To speak of a trophic level effect automatically assumes that food is the more important of

the two. It seems likely that habitat and mode of life rather than trophic level are the most

important ecological characteristics determining uptake and final concentration of chlorinated

hydrocarbon pesticides in aquatic invertebrates. For example, invertebrates leading a plank-

tonic existence usually accumulate the highest concentrations because they provide a lipid

source in the water column on which the hydrophobic chlorinated hydrocarbons can adsorb.

Once the chlorinated hydrocarbon pesticide has left the water and entered into an organic

reservoir it would then be available largely to those organisms associated with the particular

substrates for which the chemical has the greatest affinities (Macek, 1969). Invertebrates

occupying or contacting a substrate that is high in organic matter favoring partitioning of

non-polar pesticides will likely carry high concentrations of chlorinated hydrocarbons (Wallace

and Brady, 1971 ; Derr and Zabik, 1972; Frank et al., 1974). Thus the chemical would not

be equally available to all trophic levels in an aquatic community (Macek, 1969). The relatively

high concentrations of dieldrin in the Lymnaea stagnalis, Chironomidae, Glossiphoniidae, and

Libellulidae in Rosenberg (1975) can be explained in this way although other reasons must

be sought for the relatively high residues detected in invertebrates not associated primarily

with the bottom sediments. Nowhere is the influence of habitat on pesticide uptake better

illustrated than in fish. For example, Morris and Johnson (1971) found that bottom-dwelling

fish (channel catfish) contained much higher concentrations of dieldrin than non-bottom-

dwelling fish (largemouth bass and bluegill). It is not hard to imagine why terrestrial inverte-

brates such as earthworms and slugs which are constantly in contact with the soil — a poten-

tially large pesticide reservoir — can accumulate higher concentrations than an invertebrate

predator which may live in leaf litter and/or scurry around above the soil (e.g. Carabidae).

In terms of the influence mode of life has on pesticide uptake, bivalve molluscs such as

oysters, mussels, and marsh clams are well known for their abilities to accumulate high

concentrations of chlorinated hydrocarbons (e.g. see Butler, 1969; Khan et ah, 1972; Bedford

and Zabik, 1973; Petrocelli, Hanks, and Anderson, 1973). Bivalves filter large volumes of

water and if the water contains chlorinated hydrocarbons and suspended organic particles to

which chlorinated hydrocarbons are adsorbed, the bivalves will contact large amounts of

pesticide. However, Bedford and Zabik (1973) have noted that, for mussels, other ecological

factors such as: (1) previous conditioning and insecticide residue burden of the mussel; (2)

food content and temperature of the water; (3) water quality (including the presence of chem-

ical pollutants and suspended sediment load); and (4) type of pesticide are all involved in the

final concentration achieved. Some evidence exists that several trace elements can be magnified

in passing from the food to the feces of marine primary consumers (Boothe and Knauer, 1972).

Contamination of coastal waters by trace and heavy metals apparently may be as widespread

as by pesticides and since fecal material is important to the trophic relationships of coastal

benthic communities, the concentrating mechanism may have a significant influence on levels

of trace and heavy metals in coprophagous and other members of detrital food webs (Boothe

and Knauer, 1972). There is every likelihood that similar processes occur among freshwater

invertebrates with chlorinated hydrocarbon pesticides. Dindal and Wurzinger (1971) using

the terrestrial snail Cepaea hortensis (Muller), showed that highest DDT residues occurred in

the feces. They pointed out that since snails frequently re-ingest their own feces, the pesticide

can be recycled. Davis (1971) has further illustrated how habitat and mode of life affects

Food Chain Concentration of Pesticides

105

uptake and accumulation of chorinated hydrocarbons in invertebrates in his consideration

of DDT and dieldrin dynamics in two species of earthworms (Lumbricus terrestris L. and

Allolobophora caliginosa Sav.)- The main factors affecting uptake and accumulation of dieldrin

and DDT were: (1) different soil types — Organic matter content will influence the amount

of pesticide stored; soil moisture and pH will affect the physiological state of the earthworm

and hence its ingestion activity; (2) feeding habits of different species — A. caliginosa ingests

relatively more soil than L. terrestris and at a greater depth. L. terrestris is thus more likely

to accumulate residues remaining on the vegetation and soil surface. Unfortunately, a paucity

exists of this kind of precise habitat and mode of life information as related to pesticides in

freshwater invertebrates. It would be more profitable to attempt such studies rather than

doing interminable monitoring studies which are already over-abundant in the literature.

Hamelink et al. (1971) have proposed that exchange equilibria control the degree of ac-

cumulation of chlorinated hydrocarbons by organisms in lentic environments. This is supported

by the earlier findings of Reinert (1967) and Chadwick and Brocksen (1969) that the major

uptake of DDT by Daphnia and dieldrin by Cottus perplexus Gilbert and Evermann respective-

ly was from water and not food. (See also Edwards, 1970). Crosby and Tucker (1971) mini-

mized the importance of ingestion as a route by which Daphnia are exposed to suspended

chemicals. Derr and Zabik (1974) proposed an adsorption-diffusion mechanism was respon-

sible for the uptake and concentration of DDE by Chironomus tentans Fabricius. Macek

(1969) and Macek and Korn (1970) reported that brook trout accumulated 10 times more

DDT from food than from water in their laboratory studies using approximately 3 pptr. They

concluded that since a higher concentration of DDT exists in the food web than in water in

natural conditions that the food web is the major source of DDT contamination in fish.

Macek (1969) suggested that this was true of lower trophic levels (presumably invertebrates)

as well. Macek and Korn’s (1970) conclusion that more DDT is available from the food web

than from water is not only unsubstantiated by them but also ignores other non-food web

reservoirs of DDT in aquatic ecosystems (e.g. suspended organic matter and bottom sediments).

Moreover, Murphy (1971) has shown that the results of Macek and Korn (1970) were an arti-

fact of the size of the test fish used. Complexity of the relationship between feeding habits,

fat content, and age of fish is illustrated by the study of Frank et al. (1974).

Exchange equilibria would depend on a number of factors according to Hamelink et. al.

(1971): (1) original concentration of the pesticide in the water (see also Macek et al., 1972;

Derr and Zabik, 1972); (2) the water or fat solubility of the pesticide — an increase in water

solubility or decrease in fat solubility should reduce the degree of accumulation; (3) the fat

content of the animal (see also Morris and .lohnson, 1971 ; Hughes and Lee, 1973; Frank et.

al., 1974); (4) the species sampled; (5) the time available for exchange (see also Chadwick

and Brocksen, 1 969; Cope, 1971; Johnson et. al., 1971; Wilkes and Weiss, 1971; Morris and

Johnson, 1971 ; Derr and Zabik, 1972); and (6) habitat — persistent pesticides would be tied up

in various reservoirs (e.g. algae, bottom sediments of high organic matter content) in a eutrophic

lake moreso than in an oligotrophic one and thus be less available for accumulation. Size

(body weight and surface to volume ratio) of the organism would also be involved (Morris

and Johnson, 1971; Murphy, 1971;Kenaga, 1973; Frank et al., 1974). Moriarty (1972) has

criticized Hamelink et al.’s (1971) model as being too simplistic and has quite rightly pointed

out that it is not the final explanation to the observed phenomena. Nevertheless, in my view,

it has been an important contribution.

Other factors, not all of which can be named here, influence the accumulation of pesticide

residues by aquatic invertebrates^. Kenaga (1972, p. 195) has emphasized that the phenomenon

1. For example, see Derr and Zabik’s (1974) remarks about the possible role of the epicuticular

lipid layer in invertebrates. Also see Wallace and Brady (1971) for possiblity of seasonal effects.

Quaest. Ent., 1975, 11 (1)

106

Rosenberg

is hard to define because of the “many variable inputs, interpretations, and lack of definition

in the literature.” In trying to explain the lack of consistent food chain buildup in their study,

Robinson et al. (1967) wrote of the possibility of a differential ability of vertebrates and

invertebrates to metabolize and excrete the pesticide. Pharmacokinetics greatly influence the

pesticide levels in organisms (Moore, 1967; Stickel, 1968; Chadwick and Brocksen, 1969;

Dustman and Stickel, 1969; Moriarty, 1969, 1972; Edwards, 1970; Cope, 1971 ; Hamelink

et al., 1971; Kawatski and Schmulbach, 1971; and Wilkes and Weiss, 1971). Some pesticide

accumulation undoubtedly results from ingestion (Kenaga, 1973). Physical and chemical

properties of the pesticides, in addition to solubility and partitioning coefficients already dis-

cussed, are involved (see Kenaga, 1972, 1973). Finally, extrinsic factors must affect pesticide

levels in aquatic invertebrates: (1) varying concentrations (e.g. see Chadwick and Brocksen,

1969; Hamelink et al., 1971; Wilkes and Weiss, 1971); and (2) whether exposure is acute or

chronic(e.g. see Moore, 1967; Stickel, 1968; Chadwick and Brocksen, 1969; Moriarty, 1969;

Edwards, 1970; Johnson et al., 1971).

It is important to realize that a literature survey which shows an apparent absence of a

trophic level effect in invertebrate communities is only indirect evidence that food chain up-

take is not as important as other ecological factors in determining the concentrations of pest-

icides in invertebrates. Determination of the relative, quantitative contributions made by the

factors discussed above to the pesticide levels achieved in invertebrates depends on carefully

controlled experimentation and is a critical research need in the study of pesticide-fauna in-

teractions. Until such research is done, and perhaps even after, we should not be using the

terms “food chain concentration” or “trophic level effect”. Rather, we should talk of “bio-

concentration” which Kenaga (1973, p. 75) has defined as “the amount of a pesticide residue

accumulated by an organism by adsorption, and by absorption via oral or other route of entry,

which results in an increased concentration of the pesticide by the organisms or specific tissues.”

SUMMARY AND CONCLUSIONS

1. Concentrations of chlorinated hydrocarbon pesticides in aquatic invertebrates as reported

in the literature do not reveal a trophic level effect.

2. Uptake and accumulation of chlorinated hydrocarbon pesticides in aquatic invertebrates

is more likely a function of habitat, mode of life, and exchange equilibria than food but

is also affected by size of the organism, pharmacokinetics, physical and chemical proper-

ties of the pesticides, and various extrinsic factors.

3. Until adequate research is done, the relative contributions of the factors listed above to

pesticide levels in invertebrates will remain unknown.

4. The idea of “food chain concentration” or “trophic level effect” is inaccurate and should

be replaced by the more accurate idea denoted by the term “bioconcentration”.

ACKNOWLEDGEMENTS

I would like to thank W. G. Evans for his advice, continual encouragement, and the many

helpful discussions we have had. Thanks are also due to the other members of my advisory

committee for their help in producing the thesis which formed the basis of this paper: G. E.

Ball, Department of Entomology, University of Alberta; H. F. Clifford and J. R. Nursall, both

of the Department of Zoology, University of Alberta; and W. F. Allen, Department of Chemistry,

University of Alberta. F. A. J. Armstrong, M. Healey, and R. D. Hamilton of the Freshwater

Institute, Winnipeg, G. R. B. Webster, Pesticide Research Unit, University of Manitoba, and

D. Pimentel, Cornell University, critically read the manuscript. I have used many of their

suggestions.

Food Chain Concentration of Pesticides

107

G. Forth and C. Plumridge, Freshwater Institute, Winnipeg, typed drafts of the manuscript.

A. P. Wiens and C. Madder helped proofread the drafts. E. Marshall and his staff (Freshwater

Institute Library) provided help obtaining some of the literature.

This research was supported by National Research Council of Canada and Canadian Wild-

life Service grants to Dr. W. G. Evans. I am pleased to acknowledge my appreciation to the

National Research Council of Canada for scholarships awarded.

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Ill

Book Review

NACHTIGALL, WERNER. 1974. Insects in Flight; A glimpse behind the scenes in biophy-

sical research. Translated by Oldroyd, R., R. H. Abbott and M. Biederman-Thorson. George

Allen and Unwin, Ltd., London. 153 p. $14.00

The title of this book is very appropriate as Nachtigall writes not only about insect flight,

but gives a glimpse into the ways in which physicists think. Never, however, does he forget

that the object of his research is an animal and a very complex one at that.

His opening statement sets the stage for this unusual book, when he asks “a blue bottle

irritates us with its buzzing and we squash it. What have we destroyed?” The remainder of

the book answers his question.

There are 44 chapters in this book of only 1 53 pages and consequently some chapters are

very short — often less than one page. This large number of short chapters unfortunately at

times gives the impression that the book is only a series of loosely connected ideas. However,

this does not detract from the book’s worth.

Nachtigall, in his forward, states that the book is aimed at the general reader and student.

But, althougli he is meticulously careful in his explanation of the physics involved, once past

the initial chapters on general entomology a more than rudimentary knowledge of physics is

a decided advantage.

Nachtigall first explains the forces that act on a gliding body, illustrating his discussion with

everyday examples, such as gliders and powered flight. Almost without realizing it the reader

is then introduced to the way in which these forces act on an insect and its wings.

Explained at length is the difficulty of observing wing beats with frequencies of only a few

hundredths of a second. He almost apologizes for spending $30,000 and many hours of com-

puter time just in studying “a tiny fly”, but explains that this amount of money is not large

if it serves to add meaningful information to man’s reservoir of knowledge.

Chapters 15 and 16 which tell of the Vv^ays insect wings generate lift and thrust are superb,

and for me are one of the high spots of the book. His explanation of how the wings generate

lift and thrust on the upstroke is particularly clear and easily understood (provided the read-

er understands a parallelogram of forces).

Having considered the forces that keep the insect in the air, Nachtigall next considers the

mechanics of power transfer to the wing. He gives an average treatment of the “click mech-

anism” and then considers the thorax as a resonating system that stores and releases energy.

He has an interesting series of chapters (a single chapter would serve as well), on the way

oxygen and fuel are supplied to the flight muscles, making comparisons with everyday objects.

He refers to one athletic fly that flew on a flight mill for six days, being regularly “tanked up

with a sugar solution”. The fly stopped only when its wings were torn and tattered and Nachti-

gall marvels at the magnificent “motor” that drove the wings up and down some eight million

times without breaking down. Another particularly good section of the book are chapters 28

to 31 where he explains how fast and slow flight muscles work, and how the muscle contrac-

tions are initiated and maintained. Also well done are the chapters on flight initiation, main-

tenance and landing. In chapter 34 he explains effectively how one pair of mesothoracic di-

rect flight muscles initiates the downstroke of the wing at take-off. This can be shown by

making a fly take off from smoked paper, and observing that the mesothoracic legs leave a

larger white mark since they pushed hardest, and thereby giving the dorsolongitudinal flight

muscles a strong stimulus for contraction.

The book is lavishly supplied with extraordinarily good photos of insects in flight. There is

a particularly good series of photographs of Bombylius flies illustrating the stylized flying

attitude of this insect.

Quaest. Ent., 1975, 1 1 (1)

112

Nachtigall is not as effective in some chapters covering topics peripheral to insect flight.

This shows towards the end of the book where he considers direction finding and wing-beat

frequency as a mating stimulus. It is puzzling that earlier in the book in chapter 32 on “Steering

gear”, he is also rather poor and the chapter does not seem well-placed.

The book is well-produced and bound and contains few mistakes. However, occasionally

there appears to have been layout problems, as on page 87 where there is a peculiar hiatus in

the double columns.

There are no references given in the text, but the pertinent literature is listed by chapter in

a bibliography at the end of the book, along with an adequate index.

This book is essential reading for all entomologists. Not just for the information that it

contains on the physics of insect flight, but for the appreciation of Nachtigall’s wonder and

amazement of how all the many facets of that “small blue-black troublesome buzzing creature”

have evolved together to produce a highly efficient animal, a fly.

D. A. Craig

Department of Entomology

University of Alberta

Publication of Quaestiones Entomologicae was started in 1965 as part

of a memorial project for Professor E. H. Strickland, the founder of the

Department of Entomology at the University of Alberta in Edmonton

in 1922.

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charges are normally levied, the rate determined by printer’s charges. For

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uaestiones

Entomologicae

JUK 91975

A periodical record of entomological investigations,

published at the Department of Entomology,

University of Alberta, Edmonton, Canada.

VOLUME 11

NUMBER 2

APRIL 1975

QUAESTIONES ENTOMOLOGICAE

A periodical record of entomological investigation published at the Department of

Entomology, University of Alberta, Edmonton, Alberta.

Volume 1 1 Number 2 April 1975

CONTENTS

Steiner — Description of the Territorial Behavior of Podalonia valida (Hymenoptera, Sphecidae)

Females in Southeast Arizona, with Remarks on Digger Wasp Territorial Behavior

113

Griffiths — Studies on Boreal Agromyzidae (Diptera). IX. Phytomyza Miners of Boraginaceae

in North America 1 29

Ball — Pericaline Lebiini: Notes on Classification, A Synopsis of the New World Genera, and

a Revision of the Genus Phloeoxena Chaudoir (Coleoptera: Carabidae) . 143

DESCRIPTION OF THE TERRITORIAL BEHAVIOR OF

PODALONIA VALIDA (HYMENOPTERA, SPHECIDAE) FEMALES IN

SOUTHEAST ARIZONA, WITH REMARKS ON DIGGER WASP TERRITORIAL

BEHAVIOR

ANDRE L STEINER

Department of Zoology

University of Alberta Quaestiones Entomologicae

Edmonton, Alberta T6G 2E1 11: 113-127 1975

Each female Podalonia valida concentrates many consecutively built nests on small, often

discrete, defended and frequently visited nesting sites in densely vegetated areas, over periods

of days or weeks. A special “ tail-up /wings-sp read” (and wing-fluttering) posture is assumed

on the nesting site, possibly a territorial display (and/or signal of non-receptivity to males;

a deterrent to ant invaders). Contrary to many or most Podalonia females, those of P. valida

dig a burrow before prey capture. This behavior pattern and territoriality require accurate

medium-range homing ability. Territoriality and nest-clumping might have evolved in relation

with the scarcity of suitable nesting sites in densely vegetated habitats where the heavy prey

(larvae of the arctiid Estigmene acraea) live and from which they probably cannot be carried

over great distances. Territoriality probably also reduces the incidence of re-excavation of

provisioned nests by conspecifics. Territoriality probably also ensures that one female only

will occupy the small site if several daughter-females emerge from the clumped nests, thereby

encouraging dispersal and reducing local crowding. The special posturing might have evolved

from activities characteristic of some wasps excitedly engaged in nesting activities such as

wing fluttering, running about, holding the abdomen high.

Chaque femelle de Podalonia valida concentre de nombreux nids successifs sur des lieux de

nidification restreints, quelle defend et visit e frequemment, jour apres jour, en adoptant une

posture speciale avec les ailes etendues, vibrant continuellement, et Tabdomen releve. Cette

posture territoriale (?) pent etre aussi un signal de non-receptivite pour les males ou meme une

posture d’intimidation envers les fourmis qui envahissent son territoire. Contrairement a Thabi-

tude chez les Podalonia, le terrier est creuse avant la capture de la proie. Ceci exige, avec le

comportement territorial, des facultes d’orientation et de retour au nid. Ces comportements

114

Steiner

inhabituels sont peut-etre adaptes d la rarete des lieux de nidification en des endroits d vegetation

dense, oil vit la lourde proie (une chenille d'arctiide: Estigmene acraca) qui ne pent etre trans-

portee sur de grandes distances. Le comportement territorial reduit aussi probablement les

chances de re-ouverture, par dautres femelles, de nids approvisionnes, et assure Vexclusivite

du lieu de nidification d une seule femelle en cas de competition entre plusieurs femelles-filles.

La posture speciale est peut-etre derivee d’elements caracteristiques de guepes febrilement

engagees dans des activites nidificatrices, tels que la vibration des ailes, f elevation de Vabdomen,

les deplacements rapides sur le sol.

INTRODUCTION

Territoriality and Site Attachment in Digger Wasps

In this paper, a first detailed description of the territorial behavior of female digger wasps

is presented. Intermittent observations were made on Podalonia valida (Cresson) wasps which,

in the area considered, were found to nest mostly singly on small, discrete and defended pockets

of bare soil surrounded by large areas of tall, dense, herbaceous vegetation, a rather unusual

nesting situation for digger wasps.

Descriptions of territorial behavior and site defense in female digger wasps are almost

entirely lacking, even for gregarious species, which often nest at very close quarters and seem

to ignore or tolerate each other. For example: nest entrances of 9 different females of Plenoculus

davisi were located in an area of 1.1 x 1.2 m, each separated by distances of from 18 to 90

(mean 38) cm (Kurczewski, 1968) (distances as small as 2-5 cm were found in an aggregation

of Clypeadon laticinctus, Evans, 1962). When present, defense reactions are usually restricted

both in space (the immediate vicinity of the nest) and in time (only during performance of

nesting activities) in sharp contrast to P. valida females which exhibit extensive site defense

behavior both in space (defense of a sizeable area) and even more so in time (over days or

weeks and essentially throughout the day, if only intermittently). However, some apparently

density-dependent fighting is reported among females of Sphecius speciosus (Lin, 1963b).

Much more information is available on male digger wasp territorial or related behavior and/

or site attachment. Examples are as follows: Astata (Shuckard, 1837; Minkiewicz, 1934; Wil-

liams, 1946; Evans, 1957 and 1970; and Steiner, \913);Dryudella (Steiner, 1973); Tachysphex

(Kurczewski, 1966); Tachytes (Lin, 1963); Trypoxylon (Richards, 1934; Hartman, 1944;

Krombein, 1954); Sphex (Janvier, 1928); Oxybelus (Bohart and Marsh, 1960). These males

often select a specific perch where they return consistently (after threatening or pursuit flights

directed at male conspecifics or even other insects). In some cases (some observations on

Astata, Dryudella, for instance) observations on active defense are lacking, suggesting the pos-

sibility of mere “site attachment” or “maintaining a station”. Defense is particularly vigorous

in the species Sphecius speciosus, studied in detail by Lin ( 1 963a), where male territory holding

extends over periods of 8, 9 and 1 2 days (the females, which are non-territorial, were not at-

tacked when trespassing). Most or all of this behavior seems, however, essentially related to

copulatory, pre-copulatory behavior, or courtship and seems to promote male spacing and

undisturbed male/female contacts.

Habitat and Microhabitat

Since the territorial behavior described here might well be habitat-dependent, and the species

extends over a wide range of habitats, climates, and geographical areas, habitat characteristics

must be defined in some detail, for future comparative purposes.

The study was conducted during the late summer and fall of 1972, only a few miles north-

west of the Chiricahua Mountains (one mile west of the Chiricahua National Monument en-

Behavior of Podalonia valida

115

trance, Cochise County, Arizona, U. S. A.), in the surrounding arid lands and deserts (Arizona

section of the Sonoran Desert), mainly desert-grassland in the Upper Sonoran and Transition

Zones. Rain is highly seasonal with maxima recorded during July, August, and September.

Xeric vegetation and cacti are common in the area.

The nesting sites of female Podalonia valida were located on a much lusher riparian and more

mesic habitat (along Bonita Creek, a temporary water course dried up most of the time) covered

with diverse vegetation consisting of large trees and an abundant understory of shrubs and vine

tangles; and after the period of heavy rains, during the nesting season, a carpet of lush, dense,

herbaceous vegetation also covered most areas of alluvial soil (Fig. 1).

A few bare patches remained in the looser sections of the alluvial deposits along paths, wild-

Ufe runways, or around ant hills and mounds made by burrowing rodents (Fig. 2; open arrows

1, 2, and 3, and solid arrow).

Behavior and Biology of Podalonia Wasps

Insofar as known, Nearctic Podalonia females paralyze lepidopterous larvae that are often

of large size, i.e. cutworms (Noctuidae). This they do usually before digging a short, L-shaped

unicellular nest. After the latter is prepared, the caterpillar is dragged to the nest, where it is

stored and is provided with the egg of the wasp. The nest is then permanently closed (see for

instance Murray, 1940; Muesebeck et al., 1951 ; Krombein et al., 1958, 1967;Bohart and

Menke, 1963).

NESTING BEHAVIOR AND TERRITORIALITY OF PODALONIA VALIDA

Podalonia valida, variously named in the literature, is a wide ranging species in western

North America (Canada to Mexico) most common in arid areas (details in Fernald, 1927;

Murray, 1940; Muesebeck et al, 1951 ; etc.). The general biology and behavior of P. valida

wasps (not described in the current literature) is described elsewhere (Steiner, in preparation).

Briefly, the biology of this species is unusual among Podalonia wasps in that the females

studied dug their burrows before rather than after hunting (see, however, Tsuneki, 1968) and

left the burrow open during prey hunting. They preyed on large, very hairy caterpillars of

Estigmene acraea (Drury) (Arctiidae) or on those of very closely related forms (Steiner, 1974),

and did not have to first excavate them from underground shelter like the cutworm-hunting

species do. Nest-clustering is probably less exceptional, since it has already been recorded at

least for females of some other species.

Like some other Podalonia species studied, nest closure involves first wedging one or a few

pebbles, earth clods, etc. in the tunnel, then kicking in sand and tamping it tight with blows

of the head and/or occasionally rubbing the dirt with a stone or a twiglet held in the mandibles

(but not “hammering” the dirt as do some wasps). Bits of debris are often accumu-

lated on the burrow site.

Individual Nesting Sites

Four such widely separated nesting sites were discovered in an area approximately 70 by

100 meters. Two of them were located on a relatively large area of bare or sparsely vegetated

soil (e.g.. Fig. 4); and are called “open type”. The other two sites were very small, discrete

pockets of bare soil: old mounds made by burrowing rodents and/or ant hills flattened out

by weathering (e.g., Fig. 3); they are called “closed type”. They were completely isolated in

a matrix of dense vegetation, mostly tall grasses (see Fig. 1 : the small nesting site of closed

type, shown enlarged in Fig. 3, is indicated by an arrow).

Each of these four nesting sites was occupied throughout the day, if only intermittently.

Quaest. Ent., 1975, 11 (2)

116

Steiner

and for a period of several days (at least 10 for site 1, and probably much more) by only one

female with her successive nests. Occasional short-duration visits to such areas by other females

were observed. However, the latter were driven away as soon as detected by the owner. Individual

identification of the wasps was made easy by considerable size differences, different wear pat-

terns of the wing-tips (see Fig. 5, the wing-tip on the right, for instance) and behavioral idio-

syncrasies; some also had missing leg or antennal segments.

All four nesting sites were away from large trees or shrubs, although the open type ones

were close to tall herbaceous plants (Helianthus petiolaris? and other sunflower-related plants;

Verbesina encelioides?) and small shrubs (Apache plume: Fallugia paradoxa) (Fig. 4). All four

sites were on fine alluvial soil, rather sandy and compact. The surface layer for about an inch

deep was dry and loose, except at times after rain. Damper soil was usually soon found in deeper

layers. During some short-term dry periods within the rainy season, the damp layer sank to about

2 inches or more below the surface, making digging more difficult for the wasp because of col-

lapse of the entrance or upper wall of the burrow, or because of sand slides.

Most observations were concentrated on one closed type nesting site (site 1) (Fig. 1, 2, 3)

because it was occupied by the most active female (Female 1) and the one that showed territorial

behavior most frequently and typically. Site 1 was only about 30-40 cm across (in the “open

type” no evident border was visible, but the nests were concentrated on a slightly larger total

surface, about 50-60 cm across; see Fig. 4, area delimitated by dotted Ijne). Site 1 was occupied

by Female 1 at least from September 10, 1972 (beginning of observation) to September 20,

1972, when she switched to another nest site.

Behavior of Individual Females on Their Nesting Sites

Occupancy of the nest site. — Long-term occupancy of the same restricted nesting site, to

the exclusion of other females (an unusual feature for digger wasps), amounts to a quasi-

monopolization of the site.

When not engaged in nesting activities right on the site, the wasp typically pays frequent

or spaced visits to the site, staying for relatively short periods of time, even giving the impres-

sion of “guarding” and/or “advertising” the site. This contrasts sharply with most mass-pro-

visioning digger wasp females, who leave the nesting site immediately after completion of the

nest and pay no further attention to it.

A typical, idealized daily activity pattern can be summarized as follows. The wasp arrives

on the site some time in the morning (apparently not spending the night there), usually not

very early (see Table 1) and basks in the sun. [Tsuneki (1968) reports that in East Mongolia

some Podalonia wasps dig burrows apparently for the sole purpose of overnighting; these

burrows were not subsequently used for nests.] She then becomes active, soon excitedly run-

ning in all directions over the nesting site, while assuming a peculiar tail-up/ wings-spread (and

wing-beating) posture (described later). When she leaves the site, either in flight or on foot,

for variable periods of time, for feeding on flowers, etc., she abandons the special posture

described above, re-assuming it when she returns to the site, most often on foot.

The wasp progressively engages in nesting activities. Depending on whether or not a burrow

had been dug the previous day, in the evening or late afternoon (see Table 1), she under-

takes a series of prey-hunting trips (flying away) if a burrow is available (first hunting trips

are mostly unsuccessful), or if not, she shows appetitive digging on the site, followed sooner

or later by actual burrow digging. These activities usually take place late in the morning or

around midday and into early afternoon.

The rhythm of hunting trips and visits accelerates as time proceeds, until the wasp captures

a caterpillar, stings it, brings it back on foot, stores it in the terminal cell, oviposits, then closes

the burrow. During the performance of these nesting activities (nest digging and closure), the

Behavior of Podalonia valida

117

Fig. 1. General view oi Podalonia valida habitat (photo taken looking west). Arrow indicates location of small, discrete, nesting

site = site 1 in text (old pocket gopher hill?) isolated in an area of dense herbaceous vegetation, a: Apache plume (Fallugia paradoxa);

b: Helianthus spp. (e.g., petiolaris‘1) and “go Id weed” (Verbesina encelioides); c: Arizona sycamore (Platanus wrightii); d: moun-

tain ash (Fraxinus velutina); e: oak (Quercus emoryi); f: Arizona cypress (Cupressus arizonica); g: alligator juniper (Juniperus

deppeana). The line of trees in the distance indicates the location of a temporary creek (Bonita Creek). Fig. 2. Podalonia valida

habitat (photo taken looking east). Arrow indicates location of same small nesting site 1 shown in Fig. 1. The dense shrubby and

herbaceous vegetation on the right of photo is the “hunting ground” of the wasp, where the latter caught most prey, a: Apache

plume; b: Helianthus spp. (e.g., petiolaris?) and Verbesina encelioides); c: Arizona sycamore; h + i: vegetation tangle, with

mainly Arizona grape (Vitis arizonicus) (h) and young Arizona walnut (Juglans major) (i), etc.;/.' cholla cactus (cane choUa?

Opuntia spinosior). Note bare patches = potential nesting sites (“open” type in text) (open arrows: 1, path; 2, area cleared by

harvester ants: Pogonomyrmex sp.; 3, cattle “dusting place”). Some of these areas were occupied by other digger wasps too

{Ammophila spp., Bicyrtes spp., Bembix spp., Tachysphex spp., Cerceris sp., etc.). (Note: Horsemint {Monarda pectinata),

'another characteristic plant of the area, is not visible on this photo.) Fig. 3. Detail of the “closed” type of nesting site (site

1 in tex-t) that was indicated by an arrow in Fig. 1 and 2. It is about 30-40 cm across and probably represents an old mound

made by a burrowing rodent, weathered away and isolated in dense herbaceous vegetation.

Quaest. Ent, 1975, 11 (2)

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Steiner

Fig. 4. Detail of an “open” type of nesting site (about 50-60 cm across; approximate delimitation: dotted line), located on

a relatively large area of bare soil. Arrow indicates position of burrow of the wasp, left open after digging. The darker areas

of soil in the center of the nesting site represent nest fill left on the area after digging and closing the previous nests. The large

circular area of bare soil, more or less concentric with the nesting site (dotted line), represents an old site previously occupied

and cleared by a harvester ant colony {Pogonomyrmex sp.). a: Apache plume bush. Fig. 5. Special tail-up/wings-spread

(fluttering) posture (territorial display?) assumed by the “owner” of a nesting site (nesting territory?), while active on the

latter, or while engaged in nesting activities there, or chasing intruders away. The wasp also walks excitedly all over the site

while exhibiting this behavior. Note the tattered wing tips, characteristic of wasps near the end of their reproductive season.

Behavior of Podalonia valida

119

wasp also exhibits the special posture particularly apparent during the early (appetitive) stages.

Several nests can be completed in one day. As evening approaches, the wasp visits the nesting

site less and less frequently until she fails to reappear (sometimes as late as 1815, 10 Sept.;

1 822, 1 1 Sept. ; 1730, 15 Sept. ; 1717, 17 Sept. ; and even 1 900, at which time the wasp just

finished nest closure, 16 Sept.).

Table 1. Early occupancy of nesting site 1 in the daily cycle (in daylight saving time).

+ an open burrow (dug the previous day) is visible on the nesting site.

As an illustration of daily occupancy of a nesting site, the following summary of field notes

is given. On September 1 1, 1972, the wasp owner (Female 1) was first seen on nest site 1 at

1020 (beginning of observation) and last seen there at 1822 (end of observation: 1830). From

1020 to 1058 the wasp visited the site frequently and also chased away an intruding female

who tried to re-open an already provisioned nest (re-excavation behavior). A copulation attempt,

by a visiting male, was also recorded. From 1058 to 1225, the wasp again frequently came back

to the nest site (between hunting trips?) and ran excitedly over it, while assuming a special

tail-up/ wings-spread (and vibrating) posture. She also performed some digging in a burrow that

had been dug the previous day. Another copulation attempt was recorded. Prey capture and

storage probably occurred during an interruption of the observation (from 1225 to 1309) since

at 1309 the wasp was found in the process of closing the nest (early stage). This ended at ap-

proximately 1346, but from 1330 on, nest closure activities became interspersed with increas-

ingly frequent and long lasting bouts of rest. At 1348, again an intruding female was noticed

but left at once. At 1350, the owner started digging a new burrow, very close (7-8 cm) to the

nest just completed, and later chased (1490) a visiting male who tried to copulate. Digging

proceeded, intermittently, until 1446, when suddenly the wasp started to fill the non-provisioned

nest with sand, apparently in response to an invasion of the burrow by tiny ants. The burrow

was abandoned at 1450 and the wasp left, but re-visited the nest site repeatedly (also another

Quaest. Ent., 1975, 11 (2)

120

Steiner

copulation attempt was observed) until 1505 when she started reopening the burrow filled

before (!) only to fill it again (1508), apparently in response to ant intrusions. More intermit-

tent visits and rests followed, until approximately 1600-1615 when digging of a new burrow

started and ended between 1631 and 1635. Intermittent and increasingly spaced visits to the

nest site followed, accompanied by the special posturing mentioned earlier, except during the

resting bouts (an intruding female was also vigorously driven away at 1659). The wasp left the

sitq for good at 1 822.

Proportions of time spent by the wasp on her nesting site appear, however, to vary consider-

ably from day to day, and probably from individual to individual, and according to circum-

stances. Thus disturbances and intrusions (of ants, conspecifics, etc.) appear to exacerbate

the wasp into staying longer on the site, and/or into coming back more frequently. In a larrine

wasp, Liris nigra, studied in captivity, frequent visits of intruders to an occupied burrow ap-

parently often resulted in the owner staying inside it most of the time — a highly unusual be-

havior in this species (Steiner, 1962), suggesting “nest-guarding”.

Behavior of the owner toward intruders. — Whenever the wasp detects an insect intruder or

a conspecific, either male or female (but particularly the latter, early in the season), on the

nesting site, she runs excitedly toward it, threatens it by using a frontal threat display (Fig. 6b),

and if this is not enough, darts at it on foot, sometimes in a short flight, mandibles open, and

may even grapple with the intruder.

H

Fig. 6. Non-displaying wasp (a, c) and displaying wasp (b, d) in frontal view (a, b) and seen from above (c, d).

Behavior of Podalonia valida

121

Frequent intrusions of ants or even columns of them also elicit the same hostile reactions,

but the wasp often appears to experience great difficulties in expelling these persevering

visitors.

Males of Podalonia valida also visit these sites, pounce on nesting females and try to mount

» them. In the absence of a female, a male often remains on the site sunning himself. A male

may “stalk” a female. For example, Gillaspy (1969) found a male Sphex tepanecus near the

entrance of a nest while the female was filling the burrow. The reaction of the owner to such

male intrusions varies from some measure of tolerance (early in the morning, at basking time,

or during the pauses between nesting activities, etc.) to threatening or even violent expulsion,

often necessitating much grappling since the male comes back repeatedly. Such attempted

copulations were also particularly frequent early in the season. Thus between 1630 and 1700

(September 10) three attempts were witnessed (one of which was apparently successful); between

approximately 1 130 and 1 500 (September 1 1), five such attempts were observed and nine

between 1045 and 1410 (September 12). Male interference with nesting females is reported

from many species, for instance in various Tachysphex (Kurczewski, 1966). Females usually

resisted copulation by moving to one side, by raising the wings nearly vertically and beating

them continuously. Krombein (1958) also reports disruption by males of nesting activities

in Diodontus virginiana (under the name Xylocelia v. ) and Powell ( 1 964) reports the same

in Ammophila aberti. Female Sphex (Gillaspy, 1962) and Oxybelus (Bohart and Marsh, 1960),

however, were seen interrupting their nesting activities to mate.

An occasional grasshopper that lands on the nesting site is also expelled at once. Such

persistent hostile (or defense?) behavior effectively prevented other insects (except an under-

ground colony of tiny ants) and conspecifics from establishing themselves on the site.

Somewhat oddly, some fly parasitoids such as large bombyliids (perhaps Ligura sp. or

Exoprosopa sp.) with striking black markings on the wings (a device that perhaps increases

the apparent size of its bearer) when hovering over the site, burrow, or owner, did not elicit

hostile reactions, and sometimes rather elicited fear reactions and even flight of the owner

from its own “territory”! These flies had then unobstructed access to the open burrow, in

which they would almost invariably oviposit at once.

Conspicuousness of the owner on her nesting site (motions and postures). — As mentioned

earlier, the wasp owner (but not an intruding conspecific) exhibited a very characteristic

behavior and assumed a striking posture whenever she was on her nesting site (but not on a

strange nesting site). She ran excitedly over the nesting site, her brilliant red abdomen held

upwards at an angle with the rest of the body which was more or less parallel with the ground.

The wings, which vibrate or flutter continuously, were held horizontal and outspread, at right

angles with the body axis (Fig. 5 and 6). Often the wasp appeared to walk in a tilted way,

almost “on tip toes”. At this time the wasp was ready to attack anything moving on or near

the nesting site.

This tail-up/ wings-spread (fluttering) posture is striking and makes the wasp very conspicuous

on the nesting site, even at several meters from the site, both at ground level (Fig. 6a, b) and

from above (Fig. 6c, d), to an animal flying over the site.

DISCUSSION AND INTERPRETATION

Monopolization and defense of a small nesting site; nest clustering

Use of small areas for nesting sites and/or clustering of consecutive nests in the same re-

stricted location are reported for some other wasps. Thus females of Ammophila procera

studied by Rau and Rau (1918) were seen “digging in the friable soil of a small, barren area

amid low vegetation”. Rau and Rau (1918) believe that a female Tachysphex terminatus ob-

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Steiner

served for some period of time “builds a chain of nests”. Clustering of consecutive nests is

recorded for instance ^ox Prionyx atratus (Lepeletier) (Williams \9\3 In Evans 1958) and

Prionyx thomae (Fabricius), both in Kansas (Evans, 1958); Hartman (1905) found nine nests

of the latter species in Texas in an area only a few centimeters square, all apparently made by

the same wasp over a period of several days. The latter observation fits remarkably with the

data reported here for Podalonia valida. There seems to be also some degree of monopolization

(but no defense?) of the area by one female over a period of at least several days.

Similar trends seem also to be at least indicated in some other species of Podalonia. Thus

Krombein (1936) found in New York a female P. robusta (see \xndQv Podalonia violaceipennis

(Lepeletier)) nesting only in one situation — in little pockets of soil formed between two roots

of several uprooted stumps. In this instance, no other Podalonia females were in sight during

excavation and filling in. Later, Krombein found provisioned nests that were re-excavated by

the wasp or some other one. Re-excavation of provisioned nests by other females is also re-

ported by Newcomer (1930), being common also in the Palearctic species P. hirsuta (unpub-

lished personal observations). In the latter species, re-excavation was followed by “stealing”

of the stored prey. What appear to be unsuccessful attempts at re-excavation were also observed

in P. valida (pers. obs.: see above). The owner, however, expelled the intruder during a visit to

the nesting site before the prey, stored in the terminal cell, had been reached, “stolen” or thrown

away (if using burrow instead of prey for this latter case). This observation suggests that ter-

ritorial defense, monopolization, and/or frequent visits might be deterrents of considerable

selective advantage in preventing the nests from being re-excavated or accidentally destroyed

by conspecifics, particularly in a situation where there are only a few limited nesting sites iso-

lated in large areas of unsuitable (densely vegetated) areas. Such sites are easily detected vis-

ually.

Still another factor might be of relevance in this respect: Podalonia females often prey on

agrotine, noctuid larvae (cutworms) that bury themselves during the daytime (examples are

as follows. Nearctic species: P. robusta (Cresson), Evans, 1963; P. luctuosa (Smith) and/or

P. communis (Cresson), Hicks, 1931a and b, and 1932. Palaearctic species: P. hirsuta (Scopoli),

Fabre, 1879, 1882;P. affinis (Kirby), Marchal, 1892;P tydei (Le Guillou), Fabre, 1879, and

Picard, 1903; P. atrocyanea Eversmann (= P. chalybea Kohl), Tsuneki, 1968; P nigrohirta

Kohl and/or P. caucasica Morawitz, Tsuneki, 1968). Digging out these prey larvae (after ol-

factory? detection, by tapping the antennae on the ground) is then part of the normal hunting

procedure in these species and could be a latent propensity in the whole genus Podalonia that

facilitates re-excavation of previously stored prey (particularly if concentrated on a small area).

Moreover, many wasps return to areas where they have made successful hunting trips, for in-

stance Astata occidentalis, (see Evans, 1957). Thus the danger of re-excavation or accidental

destruction of nests by competitors could be particularly serious in P. valida, due to the con-

centration of stored nests. Visual advertisement by the wasp (if any) might therefore function

as an indicator of occupancy. One may even wonder if presence of a large open burrow on the

site (Fig. 4, arrow) does not contribute to the same function, particularly while the owner is

away.

Such a possible selective pressure could have favored a reversal of the usual sequence, the

nest being dug before rather than after hunting. Furthermore, presence of a burrow could

also motivate the owner to return repeatedly, and thus to visit the nesting site more often

and persistently than would otherwise be the case. If so, territorial behavior per se and return

to the previously dug nest would promote better occupancy of the nest site throughout the

day.

Besides, both behaviors seem to work synergistically in that they both require (and promote?)

accurate medium-range homing abilities and topographical memory, which are unnecessary to

Behavior of Podalonia valida

123

non-territorial wasps that also use a single prey for each nest, dug after prey capture (females

of most Podalonia species of known biology), and if no nest clumping occurs.

The unusual nesting behavior of P. valida might also have been imposed on the females, at

least to some extent, by the necessity of finding suitable (bare) nesting sites, in short supply,

as close as possible to densely vegetated areas, or even inside the latter. This seems desirable

first because these areas are likely to be the best (or only?) hunting grounds (see Fig. 1 ; also

Fig. 2; the area on the right of the photo) and second because it would probably be uneconom-

ical, if at all possible, to carry such very large and heavy prey organisms over great distances

through dense vegetation (away from which nesting sites would not be in such short supply).

Besides, long distance transport would also require a long range homing ability (on foot) that

the wasp perhaps does not possess. The advantage of close proximity of the prey (and of ex-

ploiting a new food niche where there is no or less interspecific competition) might have been

acquired at the cost of some measure of intraspecific competition for scarce suitable nesting

sites. Dominant individuals are perhaps at an advantage in such a competition. This situation

is rather unusual for the Sphecidae, because most females of this family prey on small insects

that can be carried over great distances, on the wing.

Nest clustering probably also results in several sister-females emerging later on the same

small area. This could lead to local crowding if all these wasps attempt to stay on or near their

birthplace. Territorial behavior might then be selected for as a spacing-dispersal mechanism

by which the surplus wasps (if any) are driven away from a site that is sufficient for one nest-

ing wasp only. In fact, most territorial contests, trespassing and attempts at re-excavating pro-

visioned nests were observed at early stages of this study, particularly during the first three

days of observation (September 10-12). Thus within an observation period of approximately

two hours on September 10, between 1545 and 1745 hours, six instances of trespassing by

other females were observed (of which one was apparently an attempt at re-excavating a stored

nest); at one time as many as ioux Podalonia (males and females) were seen on nesting site 1.

Each time if on the site, the owner vigorously expelled the intruders. On September 1 1, during

a six-hour period of observation (approximately 1 100-1700) eight instances of female trespas-

sing were recorded (of which two were re-excavation attempts). Again, at one point, four

Podalonia (males and females) were sighted on the small nesting area.

Unfortunately, this study did not begin early enough to include the emergence period of

the wasps. Later in the season trespassing became much less frequent, possibly indicating that

the period of intensive search for nesting territories was over, or that spacing had indeed taken

place and that the wasps (including the males) had become progressively conditioned to the

position of the different “territories”, perhaps on the basis of indicators of occupancy as men-

tioned earlier. Thus between September 17 and 20 only two instances of trespassing were re-

corded, one of which was a grasshopper that was chased away and followed by the wasp in

flight as he jumped away. In fact, trespassing became so exceptional later in the season that

the territorial nature of such nesting sites could then easily have been overlooked.

A conspicuous, easily detectable, system of territories, territory owners, and indicators of

occupancy has potentially advantages, since it can attract males, and repels competitors, and

disadvantages, since it can make detection by parasites (particularly some rax ’’-looking

bombyliids) easier. There is no doubt, that the latter are attracted by and oviposit in openings,

including the “accessory burrows” dug by some wasps around their real nest (Evans, 1966),

and also including the all-too-conspicuous burrows of P. valida females.

More information is required in order to know if the unusual nesting behavior of P. valida

described in this paper is characteristic of the species as a whole, or only of some populations

in some geographical and/or habitat (local) situations. At any rate, the clustering of nests does

not appear to be always imposed on the wasp, since this was observed even when rather large

Quaest. Ent., 1975, 11 (2)

124

Steiner

bare areas were available (see Fig. 4).

Furthermore, each individual female did not appear to select exclusively one of the two types

of nesting sites (closed or open). Thus the female most observed (Female 1) selected first a

“small pocket” site (site 1, Fig. 3) of closed type, then another one of same type, and finally

a site in a more open, bare area (Fig. 2, open arrow 3) about 50-70 m from site 1. More observa-

tions are needed. Unfortunately, simultaneous observation of more than one female is very

difficult or impossible because of the wide scattering of the wasps on their discrete nesting

sites, and of the disturbance that would result from visiting the sites too frequently. In con-

clusion, if nesting on small pockets of bare soil is the cause of nest clustering and/or territorial

behavior, it appears to be a mediate rather than a proximate cause.

Conspicuousness of the wasp, posturing and movements on the nesting site

At least some elements of such behavior are found in other wasp species. Thus a female

Prionyx parkeri (under the name P. pubidorsus (Costa)), observed in the process of digging

her nest . . ., “flicked her wings constantly while outside the nest and made a slight buzzing

noise while digging” (Evans, 1958: 183); a female Ammophila harti was seen holding the

abdomen high while searching for small stones used in the temporary closure of the nest

(Evans, 1959); the female Podalonia robusta (under the name P. violaceipennis) mentioned

earlier was observed “running nervously” about on the surface of a pocket of soil . . . also the

buzzing was very pronounced during digging (Krombein, 1936). This might well be a behavior

comparable to the one observed in Podalonia valida. Podalonia luctuosa and/or P. communis

females were also described as feverishly running from one grass clump to another, inspecting

the ground carefully while hunting (Newcomer, 1930; Hicks, 1931a and b, 1932). Rau and

Rau (1918) describe an '"Ammophila pictipennis'' wasp closing her burrow and packing the

fill with her head. They add: “To accomplish this she almost stood on her head, bracing her-

self with her hind legs.” Kurczewski (1966) in his behavioral study of some male Tachysphex

(that maintain fixed “stations”) saw some females spreading their wings vertically or at right

angles and beating them continuously. This he interpreted as being a possible signal for males

that insemination occurred.

Such behavior might be, in part, a manifestation of the excitation of wasps actively engaged

in nesting activities and/or in “appetitive” stages of them, but in Podalonia valida it is not

reducible to it, since: 1) the “owner” of the territory exhibited this behavior during “pauses”

in the nesting cycle (except during basking and resting), for instance after nest digging and

before going on a hunting trip, and even outside the nesting cycles, for instance during the

short “visits” paid to the nesting site; 2) during the hunting trips, as soon as the wasp had left

her nesting site, this behavior stopped at once, although the wasp was obviously still engaged

in a nesting activity (hunting). The wasp was then even remarkably inconspicuous (no wing-

fluttering, no “nervous walking about”, etc.) in sharp contrast with the situation when she

resumed this behavior upon returning to the nesting area with or without prey; 3) intruding

females did not show such behavior (except on their own nesting site), even if apparently

engaged in nesting activities as indicated by attempts at re-excavating provisioned nests, trying

to dig a nest, etc.; 4) in one instance, a switch to another nesting site by one female was wit-

nessed after she had already made a large number of nests on the first site, possibly to full

capacity. The transition was gradual. The wasp first “visited” another, apparently vacant,

nesting site (but previously occupied by another female) at a distance of about 50-100 m from

the other site, behaving like an “intruder” and abstained from showing the special behavior

and posture, although the wasp was evidently “interested” in the burrows found there and

even performed a few digging (or re-excavating?) motions in some of them. Progressively, the

number of visits to this new site increased as the wasp came less and less frequently to the

Behavior of Podalonia valida

125

first site. At one point, not determined with precision, the wasp was found to exhibit again

the special behavior on the new site (as well as at the old site when visiting it infrequently).

The new site then became a new nesting site (nesting territory?).

Thus territory “ownership” seems to be an important prerequisite to or correlate of such

behavior which appears to be a territorial visual display or advertising device, with threat

components. Ethologists have noticed that territorial or threatening animals often make them-

selves as conspicuous, large or frightening as possible on their territories (in sharp contrast to

“trespassers”, who remain as inconspicuous, “cryptic” and small as possible). Conspicuousness

can be achieved by special movements (here running excitedly about and beating the extended

wings), coloration (here the bright red abdomen) and special posturing (here holding the brilliant

red abdomen erect), and often increasing their apparent size - “bluff” behavior (here full ex-

tension of the dark wings increasing the apparent size). If this is indeed a display, it could be

derived from pre-existing components mentioned in other nesting wasps. Or at least such

components could be the raw material used and incorporated into the display.

Holding the red abdomen high seems particularly well adapted for a frontal display (Fig. 6b),

when the wasp threatens an intruder on the ground, face to face. It makes the otherwise in-

visible abdomen (Fig. 6a) visible to the opponent (Fig. 6b) (some jumping spiders bend then-

abdomen sideways, at a right angle with the body axis, rather than upwards, when they per-

form frontal displays). This also considerably increases the apparent size of the performer

(“bluff’ component?), along with the vibrating, spread out wings and the “tilted” appearance.

The very dark, spread out wings, however, are particularly conspicuous when seen from above,

for instance by an airborne conspecific (compare Fig. 6c and 6d).

Beating the wings continuously might be also or instead a signal of non-receptivity to the

males, as suggested for some Tachysphex females observed by Kurczewski (1966), or a reaction,

a deterrent, to ant invasion (or even an attempt to “fan away” the ants). The wasp under study

(Female 1) was plagued by such invasions which were the source of considerable disturbance

(such as filling up freshly dug burrows after invasion by the ants), particularly when the ants

were very small and could not be picked up or driven away individually by the usual means.

Wing beating was, however, also exhibited by a female occupying a nesting site (Fig. 4) devoid

of ants. This makes the ant repelling explanation less likely than the territorial one.

The territorial display might have incorporated, enhanced and “ritualized” behavioral and

postural elements (e.g., wing vibration, running about, abdomen up) already present in the

repertoire of other wasps, perhaps “emancipating” them in part or totally from their original

(non-territorial) motivation(s).

ACKNOWFEDGEMENTS

The study was supported in part by a grant (A3499) from the National Research Council

of Canada.

I would like to express my gratitude to the Directors, staff and guests of the Southwestern

Research Station, Portal, Arizona, of the American Museum of Natural History (New York),

and of the Chiricahua National Monument, Arizona, for the facilities, help and suggestions

provided and for their hospitality at various stages of this project.

The Podalonia valida specimens were identified by R. M. Bohart, University of California,

Davis, and the caterpillar-prey by D. M. Weisman, Systematic Entomology Faboratory, ARS,

USDA, c/o U. S. National Museum, Washington. Much help was also provided by S. Mouat, and

V. D. Roth, in becoming familiar with the flora and fauna of Arizona.

Special thanks go to B. and E. Erickson and L. Riggs for generously opening their ranch and

property to me during this study.

Quaest. Ent, 1975, 11 (2)

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Steiner

Thanks are also due to D. A. Boag, University of Alberta (Zoology), and H. E. Evans,

Colorado State University, for critically reading and editing the manuscript.

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STUDIES ON BOREAL AGROMYZIDAE (DIPTERA). IX.

PHYTOMYZA MINERS OF BORAGINACEAE IN NORTH AMERICA

GRAHAM C. D. GRIFFITHS

Department of Entomology

University of Alberta Quaestiones Entomologicae

Edmonton, Alberta T6G 2E3 U: 129 - 142 1975

Four species of the Phytomyza symphyti group are recorded as leaf-miners of Boraginaceae

in northwestern North America, as follows: Phytomyza mertensiae Sehgal, P. petiolaris n. sp.

(type-locality Walker Fork, Alaska), P. ovalis sp. (type-locality Lake Teslin, Yukon Territory)

and P. beringiana n. sp. ( type-locality Salcha River, Alaska). Revised keys are given to the males

and leaf-mines of this group, as well as a discussion of phylogeny and host-plant relationships.

Quatre especes du groupe Phytomyza symphyti sont signalees comme mineuses des feuilles

des Boraginacees dans le nord-ouest d’Amerique du nord, tel que: Phytomyza mertensiae

Sehgal, P. petiolaris^. sp. (localitNtype Walker Fork, Alaska), P. ovalis n. sp. (localite-type

Lac Teslin, Territoire de Yukon) et P. beringiana n. sp. (localitFtype Riviere de Salcha, Alaska).

Des clefs nouvelles sont pourvues pour les males et les mines de ce groupe, aussi quune dis-

cussion de la phylogenie et les relations avec les holes vegetals.

Folgende vier Artender Phytomyza symphyXi-Gruppe werden als Blattminierer von Boraginaceae

aus dem nordwestlichen Nordamerika besprochen: Phytomyza mertensiae Sehgal, P. petiolaris

n. sp. (Fundort des Typus: Walker Fork, Alaska), P. ovalis /r. sp. (Fundort des Typus: Lake

Teslin, Yukon Territorium) und P. beringiana n. sp. (Fundort des Typus: Salcha River, Alaska).

Neue Bestimmungstabellenzum Mdnnchen und Blattminen fur dieser Gruppe werden gegeben,

sowie Erorterungen iiber Phylogenie und Wirtspflanzenbeziehungen.

The European species of the Phytomyza symphyti group have been described and discussed

in much detail by Nowakowski (1959), who treats this group of Boraginaceae-miners as a sub-

group of the Phytomyza obscura group sensu lato (the other subgroups being miners of Labiatae).

In that paper Nowakowski distinguished four European species of the P. symphyti group: P.

lithospermi Now., P. pulmonariae Now., P. symphyti Hendel and P. myosotica Now. I am

satisfied on the basis of my European material that Nowakowski’s species concepts are correct.

Very recently, Beiger (1975) has described a fifth European species, P. nowakowskiana Beiger,

so far known only from south-east Poland. Since no further revision of the European species

seems needed, I describe only the North American species in this paper. My rearings on this