LEPEOPHTHEIRUS SALMONIS (KROYER, 1837) (COPEPODA; CALIGIDAE) Examining Committee: Chairman: Dr
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(Krayer, 1837) (Copepoda: Caligidae)
by
Stewart Charles Johnson
B.Sc., University of Victoria, 1978
M.Sc., Dalhousie University, 1386
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@ Stewart Charles Johnson 1991
SIMON FRASER UNIVERSITY
December 1991
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ISBN 0-315-78163-7 APPROVAL
Name: STEWART CHARLES JOHNSON Degree: Doctor of Philosophy
Title of Thesis: THE BIOLOGY OF LEPEOPHTHEIRUS SALMONIS (KROYER, 1837) (COPEPODA; CALIGIDAE) Examining Committee: Chairman: Dr. C.L. K~F,Associate Professor
- Dr. L.J. Mbgght, Professor, Senior Supervisor, Dept. B&&ical Sciences, S&
J.M. Webstkr, Professor, t. Biologica/Sciences, SFU
- Dr. 2. Kabata, Adjunct Professor, Dept. Fisheries and Oceans Pacific, Nanaimo, B.C.
External Examiner
r(
Date Approved / 7 3c~E~.tZh/mI, /q4/ Fisheries Pdches .1*1 and Oceans et (ream 26 NOV 1991
Pacific Biological Station Nanaimo. B.C. V9R 5K6
November 20, 1991
Dr. L. Maddock Executive Editor Journal of the Marine Biological Association The Laboratory Citadel Hill Plymouth, U.K. PL1 2PB Dear Dr. Maddock: I am writing to request a letter of permission to use the material published in "Johnson, S. C. and Albright, L. J. 1991. Development, growth, and survival of Lepeophtheirus salmonis (Copepoda: Caligidae) under laboratory conditions. JMBA 71: 425-436. (Ref. JMBA 2304)" as part of my PhD thesis. The completed thesis will placed in the Simon Fraser University Library. It will be also catalogued and microfilmed by the National Library in Ottawa. I require this letter before December 17, 1991 so that I can complete the requirements for my degree this semester. If there is any problem with this request could you please inform me by Fax. Thank you for your assistance in this matter.
Yours truly,
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Dear Dr- Eales: I an writing to request a letter of permission to use the materials published in -Johnson, S. C. and Albright, L. J. 1991, Developmental stages of Lepeopfitheirus salmonis (Kroyer, 1837) (eopepoda: Caligidae) . Can. J. Zaal. 69: 929-950 (Ref. E 923) and " Johnson, S. C, and Albright, t. J. 1991. Lepeophtheirus c;d?tei f er Kabata , 1974 (Copepoda : ~aligidae)f roaa saltwater reared rainbow trout, and Atlantic salmon, in the Strait of Georgia, British Columbia, Canada. Can. J, 2-1. 69: 1414-1416. (Ref. E. 969)** am part of my PhD thesis- The completed thesis will placed in the Simon Fraser University Library. It will be almo catalogued and microfilmed by the National ~ibraryin Ottawa. I require this letter before December 17, 1991 so that I can complete the requirements for my degree this semester. If there is any problem with this requect could you please inform ma by Fax. Thank you for your asefstancr in this matter.
Ypurs truly,
Stewart C. Johbson
The Matima1 Research Co~~ci.1cf Canada is pleased to grant per~ission to include in ycur thesis the article by S.C. 30hnsnn am3 L.J. Aibright, published in the Canadian Journal of Zoolcgy, pages 1414-1416, 1491. Kindly acknowledge the source.
LV December 1991 Joan Hill i Manager, Research Journals 1 Nationa.1 Research Council Canada PART 1.A L COPYR I GHY h l CENSE . '.-. . . ., . T.:
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TIt l e of Thes 1s/Pro Ject/Extended Essay The Biology of ~epeophtheirussalmonis (Kroyer, 1837)
(Copepoda: Caligidaej
Author: (signature)
Stewart Charles Johnson
Dec. 17, 1991 (date) Lepeophfheims sulmonis (Copepoda: Cal ig idae) is an economical 1y important ectoparasite of wild and ]sen-reared salmonids. The morphology of its deyelopmental stages is described and compared with that of other species of
L~peophtheirrrsand Caligus. Development of L. sulmonis is very similar to that reported for other Lepeophtheirus species. In general the appendages attain the adult condition and sexes become distinguishable later in development than in
Culigrs species.
Experimentally determined development, growth, and survival data are provided. Development from eggs to infectious copepodid stage took 27 days at 5"
C, 12 days at 10" C, and 7 days at 15" C. Development from egg to adult male took 40 days, and from egg to adult female 52 days at 10" C. Average survival times for the copepodid stage ranged between 2 and 8 days at salinities of 15, LO,
25, and 30 %O and temperatures of 5, 10, and 15" C.
Lepeophtheirus sulmonis feeds on host mucous, skin, and blood. The morphology and dtrastructure of the alimentary tract are described and compared with those of other parasitic and free-living crustaceans. Five cell types are described from the midgut. The most abundant cell types are: E-cells (embryonic cells) which replace cells lost through degeneration or holocrine secretion, B-cells
(vacuolar cells) which produce digestive enzymes, and C-cells which function as absorptive cells and sites of lipid storage and metabolism. - iv - Naive Atlantic, chinook, and coho salmon are equally susceptible, but differ in their resistance to infection with L. saltnonis. Histological examination of
L. sulmonis feeding sites on the less resistant species, Atlantic and chinook salmon, revealed little tissue response. In contrast, the most resistant species, coho salmon, showed well-developed inflammatory responses and epithelial hyperplasias which may account for their higher resistance.
Administration of hydrocortisol increased the susceptibility and decreased the resistance of naive coho salmon to infection with L. salnwnis. Cortisol- implanted fish showed reduced inflammatory responses and suppression of epithelial hyperplasias. These data support the hypothesis that non-specific host defence mechanisms are important in resistance of coho salmon to infection with
L. salmonis. - v - ACKNOWLEDGEMENTS
i wouid like to express my gratitude iu my s~ipervisor,Dr. L. J. Albright for his guidance and support throughout the period of this study. I am also grateful to the other members of my supervisory committee, Dr. 2. Kabata and
Dr. J. Webster, for their support.
The technical assistance and advice of Mr. J. Bagshaw, Dr. V. Bourne,
Dr. M. Kent, and Dr. L. Margolis is gratefully acknowledged.
Finally, I would like to thank D. Clark and my parents for their concern and support throughout my graduate program.
This study was funded by a Natural Sciences and Engineering Research
Council of Canada grant to L. J. Albright, the Department of Fisheries and
Oceans' Biological Sciences Branch at Nanaimo, a postgraduate sch&rship from the British Columbia Science Council, and a Simon Fraser University Graduate
Fellowship. - vi - TABLE OF CONTENTS
TPTLEPAGE ...... i .. APPROVAL PAGE ...... 11 ... ABSTRACT ...... 111
ACKNOWLEDGEMENTS ...... v
TABLEOFCONTENTS ...... vi
LIST OF TABLES ...... ix
LIST OF FIGURES ...... x
GENERAL INTRODUCTION ...... 1
CHAPTER ONE:
Taxonomic Position of Lepeophtheirus sulmonis ...... 7
Comparison of the Geographic and Host Ranges of Sea Lice Species ...... 8
Identification of Sea Lice Species ...... 1 1
Key to Preadult and Adult Sea Lice of Salmon and Trout ...... 13
CHAPTER TWO: The developmental stages of Lepeophtheirus salmonis.
Introduction ...... 19
Materials and Methods ...... 21
Results ...... 22
Discussion ...... -42 - vii - CHAPTER THREE: Development, growth, and survival of Lepeophtheirus sulmonis under laboratory conditions.
Introduction ...... 68
Materials and Methods ...... 70
Results ...... 74
Discussion ...... 78
CHAPTER FOUR: The morphology and ultrastructure of the alimentary tract of Lepeophtheirrrs sulmotzis.
Introduction ...... 95
Materials and Methods ...... 100
Results ...... I02
Discussion ...... I11
CHAPTER FIVE: Comparative susceptibility, resistance, and histopathology of the response of naive Atlantic, chinook, and coho salmon to experimental infection with Lepeophtheirus sulmonis.
Introduction ...... I34
Materials and Methods ...... 136
Results ...... 138
Discussion ...... I46
CHAPTER SIX: Effects of cortisol implants on the susceptibility, resistance, and tissue responses of naive coho salmon ?o experimental infection with Lepe~phtheims satinonis.
Introduction ...... I73
Materials and Methods ...... 175 .viii .
Results ...... 178
Discussion ...... 183
GENERAL DISCUSSION ...... 196
REFERENCES ...... 201 - ix - LIST OF TABLES
Chapter Three
Table C1. Mean time in hours to first moult of the second nauplius and copepodid stages of Lepeophcheirus sulmon.is, maintained at three temperatures and ambient salinity......
Table C2. Cumulative development time, time of first and last appearance, and duration of each Lepeophtheirus salrnonis developmental stage at 10" C......
Table C3. Hatching success of egg strings of Lepeophrheirus salmonist and the mean percentage of total eggs that produced active nauplii and copepodids at 10" C and salinities of 10 to 30 %O ......
Chapter Four
Table Dl. Summary of species of parasitic copepods in which the gross morphology and/or the structure of the alimentary tract has been described based on light microscopy observations...... -X-
LIST OF FIGURES
Ch 3ter One
Figure A 1. Adult stages of L . scrln~onis.C . cllrn~s. C. clenzensi. and C. elonyan(s ...... 14
Figure A2 . Basal spine of exopod of the adult third leg of L . cuneif~rand L . su1n1oni.s...... I5
Figure A3 . Adult stages of L . cuneifc.r and L . sc11n~oni.s...... 16
Figure A4 . Structure of the adult fourth leg of C . ci~ims. C. cletnmsi. and C. elongarus ...... 17
Figure A5 . Distal margin of the exopsd of the adult first leg of C . currus. C. clemensi. and C . elorzgatm ...... 18
Chapter Two
Figure B1 . Active first nauplius ...... 53
Figure 332 . First and second nauplius. first antenna ...... 53
Figure B3 . First and second nauplius. tip of first antenna ...... 53
Figure B4 . First nauplius. second antenna ...... 53
Figure B5 . Detail of antenna seta ...... 53
Figure B6 . First and second nauplius. mandible...... 53
Figure B7 . Second nauplius. second antenna. tip of endopod .....53
Figure B8 . Newly molted copepodid ...... 54
Figure B9 . Copepodid. rostrum ...... 54
Figure B10 . Copepodid. first antenna ...... 54 .X1 .
Figure B 1 1. Copepodid, second antenna ...... 54
Figure B12 . Copepodid. mandible ...... 54
Figure B 13 . Copepodid. first maxilla ...... 54
Figure B14 . Copepodid. second maxilla ...... 54
Figure B15 . Copepodid. maxilliped ...... 54
Figure B16 . Copepodid. first leg ...... 55
Figure B17 . Copepodid. endopod of first leg ...... 55
Figure B18 . Copepodid. second leg ...... 55
Figure B19 . Copepodid. exopod of second leg ...... 55
Figure B20 . Copepodid. third leg ...... 55
Figure B2 1. Copepodid. caudal ramus ...... 55
Figure B22 . Early first chalimus...... 55
Figure B23 . First chalimus. first antenna ...... 55
Figure B24 . First chalimus. second antenna ...... 55
Figure B25 . First chalimus. mandible...... 55
Figure B26 . First chalimus. first maxilla ...... 55
Figure B27 . First chalimus. second maxilla ...... 55
Figure B28 . First chalimus. maxilliped ...... 55
Figure B29 . First chalimus. first leg ...... 56
Figure B30 . First chalimus. second leg ...... 56
Figure B3 1. First chalimus. third leg ...... 56
Figure B32 . First chalimus. fourth leg ...... 56 .X11 .
Figure B33 . First chalimus, caudal ramus ...... 56
Figure B34 . Second chalimus...... 56
Figure B35 . Second chalimus. first leg ...... 56
Figure B36 . Second chalimus. second leg ...... 56
Figure B37 . Second chalimus. third leg ...... 56
Figure I338 . Second chalimus. fourth leg ...... 56
Figure B39 . Third chalimus ...... 57
Figure B40 . Third chalimus. first antenna ...... 57
Figure B4 1. Third chalimus. second antenna ...... 57
Figure B42 . Third ckalimus. postantennary process ...... 57
Figure B43 . Third chalimus. first maxilla ...... 57
Figure B44 . Third chalimus. second maxilla ...... 57
Figure B45 . Third chalimus. maxilliped ...... 57
Figure B46 . Third chalimus. first leg ...... 57
Figure B47 . Third chalimus. second leg ...... 57
Figure B48 . Third chalimus. third leg ...... 57
Figure B49 . Third chalimus. fourth leg ...... 57
Figure B50 . Third chalimus. fifth leg ...... 5'7
Figure B5 1 Third chalimus. caudal ramus ...... 57
Figure B52 . Fourth chalimus...... 58
Figure B53 . Fourth chalimus. second antenna ...... 58
Figure B54 . Fourth chalimus. postantennary process ...... 58 .xiii . Figure B55 . Fourth chalimus. mandible ...... 58
Figure B56 . Fourth chalimus. first maxilla ...... 58
Figure B57 . Fourth chalimus. sternal furca ...... 58
Figure B58 . Fourth chalimus. first leg ...... 58
Figure B59 . Fourth chalimus. third leg ...... 59
Figure B60 . Fourth chalimus. fourth leg ...... 58
Figure B6 1. First preadult female ...... 59
Figure B62 . First preadult female. first antenna proximal segment ...... 59
Figure B63 . First preadult female. second antenna and postantennary process ...... 59
Figure B64 . First preadult female. first maxilla ...... 59
Figure B65 . First preadult female. second maxilla ...... 59
Figure B66 . First preadult female. sternal furca ...... 59
Figure B67 . First preadult female. first leg ...... 59
Figure B68 . First preadult female. first leg. tip of exopod ...... 59
Figure B69 . First preadult female. second leg ...... 59
Figure B70 . First preadult female. exopod of second leg ...... 60
Figure B7 1. First preadult female. third leg ...... 60
Figure B72 . First preadult female. raini of third leg ...... 60
Figure B73 . First preadult female. fourth leg ...... 60
Figure B74 . First preadult female. fifth leg ...... 60
Figure B75 . First preadult female. caudal ramus ...... 60 .xiv . Figure B76 . First preadult male ...... 60
Figure B77 . First preadult male. fifth and sixth legs ...... 60
Figure B78 . Second preadult female...... 61
Figure B79 . Second preadult female. first antenna proximal segment ...... 61
Figure B80 . Second preadult female. postantennary process ...... 61
Figure B8 1. Second preadult female. posterior process of first maxilla ...... 61
Figure B82 . Second preadult female. second leg ...... 61
Figure B83 . Second preadult female. exopod of second leg ...... 61
Figure B84 . Second preadult female. third ieg ...... 61
Figure B85 . Second preadult female. ralni of third leg ...... 61
Figure B86 . Second preadult female. fourth leg ...... 61
Figure B87 . Second preadult female. fourth leg tip of exopod .....61
Figure B88 . Second preadult female. fifth leg ...... 62
Figure B89 . Second preadult male ...... 62
Figure B90 . Second preadult male. fifth and sixth legs ...... 62
Figure B9 1. Adult female...... 62
Figure B92 . Adult female. genital field ...... 62
Figure B93 . Adult female. second antenna ...... 62
Figure B94 . Adult female. first maxilla ...... 62
Figure B95 . Adult female. second maxilla ...... 62
Figure B96 . Adult female. armature of second maxilla ...... 62 .xv .
Figure B97 . Adult female, maxilliped ......
Figure B98 . Adult female. sternal furca ...... 63
Figure B99 . Adult female. exopod of second leg ...... 63
Figure B180 . Adult female. exopod of third leg ...... 63
Figure B101 . Adult female. fourth leg ...... 63
Figure B102 . Adult female. tip of exopod of fourth leg ...... 63
Figure B103 . Adult female. fifth leg ...... 63
Figure B104 . Adult female. caudal ramus ...... 63
Figure B105 . Adult male ...... 63
Figure B106 . Adult male. second antenna. posterior ...... 63
Figure B107 . Adult male. second antenna. anterior ...... 63
Figure B108 . Adult male. first maxilla ...... 64
Figure B109 . Adult male. maxilliped ...... 64
Figure B 1 10. Adult male. sternal furca ...... 64
Figure B 1 1 1. Adult male. fifth leg ...... 64
Figure B112 . Adult male. sixth leg ...... 64
Figure B 1 13. Adult male. caudal ramus ...... 64
Figure B114 . Copepodid. ventral surface of cephalothorax...... 65
Figure B 115 . Copepodid. mouth cone. tip ...... 65
Figure B116 . Copepodid. tip of second maxilla ...... 65
Figure B117 . Copepodid. first maxilla and tines ...... 65
Figure B 1 18. First chalimus. postantennary process ...... 66 - xvi -
Figure B119. First chalirnus, moutli cone......
Figure B120. First chaiimus, tip of second maxilla...... 66
Figure B121. Third chalimus, tip of second tnaxilla...... 66
Figure B122. First preadult female, frontal organ...... 67
Figure B123. First preadult female, tip of second maxilla...... 67
Figure B124. First preadult female, cuticular ridges of maxilliped...... 67
Figure B125. Adult male, postoral adhesion pad...... 67
Chapter Three
Figure C 1. Mean development time of eggs of L. salmonis at various temperatures arid ambient salinity...... 89
Figure C2. Development sequence of a cohort of L. scrlrmnis at 10" C and ambient salinity...... 90
Figure C3. Percentage moult increments of total length for L. sal~nonisraised at 10" C...... 9 1
Figure C4. Percentage moult increments of cephalothorax width for L. sulrnonis raised at 10" C...... 92
Figure C5. Distribution of copepodids and chalimus larvae of L. sulrnonis on body surfaces of Atlantic salmon smolts...... 93
Figure C6. Mean (+ SD) survival time of newly moulted copepodids of L. sdmonis at various temperztures and salinities...... 94
Chapter Four
Figure D 1. Female, mouth cone oesophagus and anterior midgut. . . 120
Figure D2. Male, oesophagus anterior to nerve ring...... 120
Figure D3. Female, oesophagus junction with the midgut...... 120 .xvii .
Figure D4 . Female. mouth cone ...... 221
Figure D5 . Female. mouth cone tip ...... 121
Figure D6 . Female. external surface of the anterior midgut ...... 121
Figure D7 . Female. inner surface of the anterior midgut ...... 121
Figure D8 . Female. oesophageal cuticle and epithelium near the nerve ring ...... 122
Figure D9 . Female. higher magnification of oesophageal cuticle...... 122
Figure Dl0. Female. oesophageal cuticle and epithelium near the junction with the midgut ...... 122
Figure D 1 1 . Male. midgut near the junction with the oesophagus ...... 123
Figure D 12 . Female. anterior midgut. showing a cluster of type E cells ...... 123
Figure Dl3. Female. anterior midgut. showing several type B cells ...... 123
Figure Dl4 . Female. posterior midgut. showing several type C cells that contain large lipid deposits ...... 123
Figure D 15 . Female. type E cells ...... 124
Figure Dl 6 . Female. higher magnification of the apical region of a type E cell ...... 124
Figure 1917. Female. early type C cell ...... 124
Figure D 18 . Female. median sa~ittalsection of the anterior rnidgut ...... 125
Figure Dl9. Female. late stage type A cell ...... 126
Figure D20 . Female. apical region of a late stage type A cell ...... 126 - xviii - Figure D2 1. Female, higher magnification of the region surrounding the nucleus of a type A cell...... 126
Figure D22. Female, apical region of a late stage type B cell...... 127
Figure D23. Female, basal region of a middle to late stage type B cell...... 127
Figure D24. Female, apical region of a type A and typeBcel1...... 127
Figure D25. Female, middle to late stage type C cell of a mucus-feeding copepod...... 128
Figure D26. Female, late stage type C cell of a mucus- feeding copepod...... 128
Figure D27. Female, higher magnification of the apical region of an early to middle stage type C cell of a mucus-feeding copepod...... 128
Figure D28. Female, anterior midgut of a blood- feeding copepod...... 129
Figure D29. Female, apical region of a middle stage type C cell from the anterior midgut of a blood-feeding copepod...... 129
Figure D30. Female, degenerating nucleus of a late stage type C cell from the anterior midgut of a blood-feeding copepod...... 129
Figure D3 1 Female, type C cells of the posterior midgut of a blood-feeding copepod...... 130
Figure D32. Female, apical region of a late stage type C cell from the posterior midgut of a bld- feeding copepod...... 13 1
Figure D33. Female, higher magnification of the subapical region of a late stage type C cell from the posterior midgut of a blood-feeding copepod...... 131 - xix -
Figure D34. Female, whorls of roikgh endoplasmic reticulum associated with the nucleus of a type C cell from posterior midgut of a blood-feeding copepod...... 131
Figure D35. Female, anterior hindgut...... 132
Figure D36. Female, apical region of an electron- transparent cell of the anterior hindgut...... 132
Figure D37. Female, apical region of an epithelial cell from the midregio;] of the hindgl~t...... 132
Figure D38. Female, higher magnitication of the anterior hindgut showing mineral concretions...... 133 Chapter Five
Figure E 1. Mean (+SE) intensity of L. scrlrnonis on naive Atlantic, coho, and chinook salmon at various times post-infection...... 157
Figure E2. Distribution of L. sulrnonis on naive coho salmon...... 158
Figure E3. Distribution of L. scrlmonis on naive Atlantic salmon, ...... 159
Figure E4. Distribution of L. sultnonis on naive chinook salmon...... 160
Figure E5. Distribution of L. sal~nonison the fins of naive Atlantic, chinook, and coho salmon...... 161
Figure E6. Developmental stages of L. scrlrnonis present on naive Atlantic and chinook salmon at 20 days post-infection...... 162
Figure E7. Developmental stages of L. satrnonis present on different body regions of naive Atlantic salmon at 20 days post-infection...... 163
Figure E8. Copepodid on a gill of a naive coho salmon, 1 day post-infection...... 164 Figure E9. Inflammatory response of a gill of a naive coho salmon to L. safnionis, 1 day post-infection...... 164
Figure E10. Copepodid on a gill of a naive coho salmon, 5 days post-infection...... 164
Figure El 1. Copepodid on a gill of a naive coho salmon, 5 days post-infection...... 164
Figure E12. Copepodid on a gill of a naive Atlantic salmon, 1 day post-infection...... 165
Figure El 3. Copepodid on a gill of a naive Atlantic salmon, 5 days post-infection...... 165
Figure E14. Copepodid on a gill of a naive Chinook salmon, 5 days post-infection...... 165
Figure El5. Copepodid on a gill of a naive chinook salmon, 5 days post-infection...... 165
Figure E16. Chalimus larva on a gill of a naive Atlantic salmon, 10 days post-infection...... 166
Figure E17. Tissue response of a gill of a naive Atlantic salmon to the frontal filament of L. soltm~tiis, 20 days post-infection...... 166
Figure E18. Chalimus larva on a gill of a naive Chinook salmon, 10 days post-infection...... 166
Figure E19. Tissue response of a gill of a naive chinook salmo.; to L. sulmonis, 10 days post-infection...... 166
Figure E20. Chalimus larva on a gill of a naive Atlantic salmon, 20 days post-infection...... 167
Figure E2 1. Tissue response of a gill of a naive Atlantic salmon to L. salmonis, 20 days post-infection...... 167
Figure E22. Chalimus larva feeding site on a gill of a naive chinook salmon, 15 days post-infection...... 167 - xxi - Figure E23. Chalimus larva on a gill of a naive chinook salmon, 20 days post-infection...... 167
Figure E24. Copepodid on a fin of a naive coho salmon, 1 day post-infection...... 168
Figure E25, Copepodid on a fin of a naive coho salmon, 5 days post-infection...... 168
Figure E26. Copepodid on a fin of a naive chinook salmon, 1 day post-infection...... 168
Figure E27. Copepodid on a fin of a naive Atlantic salmon, 5 days post-infection...... 168
Figure E28. Chalimus larva on a fin of a naive coho salmon, 10 days post-infection...... 169
Figure E29. Inflammatory response of a fin of a naive coho salmon to L. sulmonis, 10 days post-infection...... 169
Figure E30. Chalimus larva on a fin of a naive coho salmon, 15 days post-infection...... 169
Figure E3 1. Inflammatory response of a fin of a naive coho salmon to a frontal filament of L. sulmonis, 15 days post-infection...... 169
Figure E32. Chalimus larva on a fin of a naive Atlantic salmon, 10 days post-infection...... 170
Figure E33. Higher magnification of a lesion caused by L. sulmotris on a fin of a naive Atlantic salmon, 15 days post-infection...... 170
Figure E31. Chalimus larvae on a fin of a naive chinook salmon, 10 days post-infection...... 170
Figure E35. Higher magnification of a chalimus larva on a fin of a naive chinook salmon, 10 days post-infection...... 170
Figure E36. Chalimus larva on a fin of a naive chinook salmon, 20 days post-infection...... 171 - xxii - Figure E37. Higher magnification of a lesion caused by L. salmnis on a fin of a naive chinook salmon, 20 days post-infection...... 17 1
Figure E38. Frontal filament of L. salmonis, 10 days post-infection...... 172
Figure E39. Fully formed frontal filament within the anterior cephalothorax of a premolt chalimus larva, 10 days post-infection...... 172
Figure E40. Higher magnification of a fully formed frontal filament within the anterior cephalothorax of a premolt chalimus larva, 10 days post-infection...... 172 Chapter Six
Figure F 1. Mean (+SE) intensity of L. salmonis on cortisol-implanted and control naive coho salmon at various time post-infection...... 187
Figure F2. Distribution of L. salmonis on naive coho salmon controls...... 188
Figure F3. Distribution of L. salmonis on naive cortisol- implanted coho salmon...... 1 89
Figure F4. Copepodid on a gill of a naive control coho salmon, 1 day post-infixtion...... 190
Figure F5. Inflammatory response of a gill of a naive control coho salmon to L. salmonis, 1 day post-infection...... 190
Figure F6. Copepodid on a gill of a naive control coho salmon, 5 days post-infection...... 190
Figure F7. Inflammatory response of a gill of a naive control coho salmon to L. salmonis, 5 days post-infection...... 190
Figure F8. Copepodid on a gill of a naive cortisol -implanted coho salmon, 5 days post -infection...... 19 1 - xxiii -
Figure F9. Feeding site of L. sulnlonis on a gill of a naive cortisol-implanted coho salmon, 5 days post-infection...... 191
Figure F10. Chalimus larva on a gill of a naive cortisol- implanted coho salmon, 15 days post-infection...... 191
Figure Fl 1 . Attachment site of L. sulinonis on a gill of a naive cortisol-implanted coho salmon, 15 days post-infection...... 191
Figure F12. Chalimus larva on a gill of a naive cortisol- implanted coho salmon, 20 days post-infection...... 192
Figure F 13. Feeding site of L. sult~~onison a gill of a naive cortisol-implanted coho salmon, 20 days post-infection...... 192
Figure F14. Copepodid on a fin of a naive control coho salmon, 1 day post-infection...... 193
Figure F15. Copepodid on a fin of a naive cortisol- i~nplantedcoho salmon, 1 day post-infection...... 193
Figure F16. Chalimus larva on a fin of a naive control coho salmon, 10 days post-infection...... 193
Figure F 17. Edge of a lesion caused by L. sulmonis feeding on a fin of a naive control coho salmon, 10 days post-infection...... 193
Figure F18. Attached preadult on a fin of a naive control coho salmon, 20 days post-infection...... 194
Figure F 19. Higher magnification of the inflarninatory infiltrate of a naive control coho salmon, 20 days post-infection...... 194
Figure F20. Chalirnus larva on a fin of a naive cortisol- implanted coho salmon, 10 days post-infection...... 194
Figure F2 1. Attachment of the frontal filament to a fin of a naive cortisol-implanted coho salmon, 10 days post-infection...... 194 - xxiv -
Figure F22. Feeding site of a chalimus larva on the fin of a cortisol-implanted coho salmon, 20 days post-infection...... 195 GENERAL INTRODUCTION
The salmon louse Lepeophtheim salmonis (Copepoda: Caligidae) is a common marine ectoparasitic copepod of salmonids throughout the northern hemisphere. This species along with four other caligid copepod species Caligus clememi, Caligus curtus, Caligus elongatus, and Lepeophtheim cuneifer which also parasitize salmonids can be collectively referred to as sea lice.
All species of sea lice studied have a direct life cycle with ten developmental stages. These stages include two free-living planktonic nauplius stages, one free-swimming infectious copepodid stage, four attached chalimus stages, two free-moving preadult stages, and one free-moving adult stage (Kabata,
1972; Johannessen, 1974; present study).
Attached copepodids, chalimus larvae, preadults, and adults feed on host mucus, skin, and blood (Kabata, 1970, 1974; Brandal et al., 1976; Jones et al.,
1990; present study). Damage to the host by the copepodid and chalimus larvae is limited to a small area around their point of attachment where the epidermis and sub-epidermis is commonly eroded. Damage by preadults and adults which are larger and capable of moving on the surface of the host, is generally more severe and widespread. Heavily infected salmonids commonly show grey patches
(extensive areas of skin erosion and hemomhaging) on the head and back, and a distinct area of erosion, dark coloration, and sub-epidermal hemorrhages in the perianal region (Wootten et a!., 1982). Heavily infected salmonids may die as the - 2 - result of secondary bacterial infections (e.g., vibrissis), fungal infections when fish are re-introduced to fresh water, or, in severe cases, by osmotic stress
(Wootten et al., 1982).
Sea lice have been reported on wild and sea-farmed salmonids in the North
Atlantic and North Pacific oceans and adjacent seas (Kabata, 1979, 1988;
Margolis and Arthur, 1979; Wootten et al., 1982; Nagasawa, 1987; Hogans and
Trudeau, 1989 a,b; Tully, 1989). They have rarely been reported to cause serious disease in wild salmonids (Huntsman, 1918; White, 1940, 1942a; Parker and
Margolis, 1964; Kabata, 1970; unpublished data for British Columbia), although localized tissue damage commonly occurs through the grazing activities of adult L. salmonis (cf. Nagasawa, 1987). However, in sea-farmed salmonids, sea lice are commonly reported to cause serious disease in many seawater net pen facilities and in some seawater-fed landbased facilities (Brandal and Egidius, 1979; Wmtten et al., 1982; Hogans and Trudeau, 1989b; Pike, 1989; Tully, 1989; unpublished data for British Columbia).
At present the only effective method for control of sea lice on farmed salmonids is the organophosphorus insecticide, dichlorvos, marketed as 'Nuvan
500EC' or in its related trichlorphon form as 'Neguvon'. Brandal and Egidius
(1977) reported the first use of trichlorphon for the treatment of salmonids infected with sea lice. In their study trichlorphon which was administered orally resulted in a decline in the number of sea lice present and a high level of mortality in the treated fish. Dichlorvo's and trichlorphon have been used since the 1960's as - 3 - a bath treatment for parasites in pond fish culture (reviewed in Schmahl et al.,
1989). Currently these compounds are administered to salmonids as a bath
treatment following the methods described by Brandal and Egidius (1979). These
treatments effectively remove both the preadult and adult stages of sea lice but not
the chalimus larvae, therefore, successive treatments usually at two to four week
intervals are required to control infections (Wootten et al., 1982). Physical
damage and high levels of stress imposed during treatment commonly results in
the development of secondary diseases (eg. vibriosis, furunculosis) in the fish. In
addition, production levels of treated stocks are lower due to lowered growth and
feed conversion rates.
Currently about 12,000 litres of dichlorvos per year are used to control sea lice in Scotland (Pike, 1989). Estimates for other countries are not available.
Although, these compounds are recognized as toxic to a wide variety of marine organisms, they are released into the surrounding waters after treatment is completed, where it is argued that their impact on non-target species will be minimal due to the extensive dilution and their rapid degradation (Egidius and
Moster, 1987; Cusack and Johnson, 1990; Dobson and Tack, 1991). This practice has resulted in conflict between environmental groups and salmon farmers over the use of these chemicals and has led Norway, Scotland, and Ireland to consider banning their use. These compounds are not registered for use in Canada.
There is little published information on alternative treatment methods for sea lice. Palmer et al. (1987) report the results of preliminary studies on the -4- efficacy of oral doses of Ivermectin for the control of sea lice on Atlantic salmon.
Ivermectin is effective in reducing populations of sea lice, however, the drug has a narrow margin of safety. Currently studies are under way to: (I) further investigate the toxicity of Ivermectin to salmonids, (2) to determine the minimum dosage required to control sea lice, and (3) to determine tissue clearance times.
These studies are being conducted in Ireland and British Columbia but as yet no results have been published.
Bjordal (1988, 1990) reported the results of both laboratory and field studies involving the use of cleaner-fish (wrasse) for controlling L. salmonis on
Atlantic salmon. In these studies wrasse were demonstrated to remove sea lice from salmonids but not always in a predictable manner. Furthermore, the wrasse species used in these studies do not occur naturally in regions where salmonids are cultured and therefore they must be imported and cultured.
Scottish researchers are investigating the possibility of producing a vaccine to control sea lice. Norwegian researchers are investigating the use of pyrethrias administered as a bath treatment. To date no information on the exact nature of these projects or results obtained has been released.
There is a serious lack of basic information on the biology of sea lice
(reviewed in Pike, 1989). Without this information we are unable to: (1) understand the epizootiology of sea lice on salmonids, (2) optimize our use of available treatment methods to obtain a maximal effect, (3) propose modifications -5- of current husbandry practices to reduce sea lice infections, and (4) propose alternative treatment methods.
This thesis presents the results of a series of investigations undertaken to provide basic information on the biology of L. salmonis. This species was chosen as it is the most common sea lice species present on farmed salmonids in British
Columbia, Scotland, and Norway (Wootten et al., 1982; Pike, 1989; unpublished data for British Columbia).
In chapter one, the taxonomic position, geographical range, and host range of L. salmonis is reviewed and compared to other sea lice species. A key which enables the identification of the preadults and adult stages of L. salmonis from those of other sea lice species of salmon and trout is given. The life cycle and morphology of developmental stages of L. salmonis are described in chapter two.
The pattern of development seen in L. salmonis is compared to other
Lepeophtheirus and Caligus species. Development, growth, and survival data derived from laboratory experiments conducted under various conditions of temperature and salinity are provided for L. salmonis in chapter three. These data are compared to other species of parasitic and free-living copepods. The functional morphology and ultrastructure of the alimentary tract of L. salmonis is described in chapter four. Function of the different cell types is inferred from their morphology and comparisons with cell types reported in other species of parasitic copepods and free-living crustaceans. The comparative susceptibility and resistance of naive coho, chinook and Atlantic salmon to infection with L. -6- salmonis is reported in chapter five. The histopathology of infection sites is
described for each salmon species and used to explain the observed differences in
susceptibility and resistance. The effects of cortisol implants on the susceptibility
and resistance of naive coho salmon to infection with L. salmonis is reported on in
chapter six. Differences in the histopathology of infection sites between the control
and cortisol-implanted groups is used to explain observed differences in
susceptibility and resistance. CHAPTER ONE
Taxonomic Position of Lepeophtheirus salmonis
The classification scheme used in this thesis is that of Kabata (1979). By this classification L. salmonis and all other species of sea lice are members of the suborder Siphonostomafoida and family Caligidae. The suborder
Siphonostornatoida comprises about 1500 described species all characterized by having a siphonostome mouth. The siphonostome mouth is tube- or siphon-shaped and formed by the partial or complete fusion of the labrum and labium (see
Kabata, 1979; Boxshall, 1990; present study). The nistory, systematics, and definition of the family Caligidae is presented in detail by Kabata (1979) and has not been repeated here.
Berland and Margolis (1983) reviewed the early history of the salmon louse
Lepeophtheirus salmonis (Krayer, 1837) and concluded that the earliest binomem applied to this species was Binoculus salmoneus (Miiller, 1785). However the name Binoculus salmoneus was judged to be unavailable by the International
Commission on Zoological Nomenclature and Lepeophtheincs salmonis (Krqer,
1837) remains the valid name for this species (see Margolis and Berland, 1984).
Other scientific synonyms for L. salmonis as listed in Kabata (1979) include:
Caligus salmonis Kmryer, 1838; Caligus vespa Edwards, 1840; Caligus strtimii,
Baird , 1 848; tepeophtheim strtimii (Baud, 1850); Caligus pac@cus Gissler , - 8 - 1883; Lepeophtheirus vesper? of Milne Edwards, in Bassett-Smith (1899); Caligus vespef? M-E, of Bassett-Smith (1899); Lepeophtheirus pac@cus (Gissler, 1883). of Wilson (1905), and Lepeophtheirus uenoi Yamaguti, 1939. In current literature there is no confusion in the use of L. salmonis.
Comparison of the Geographic and Host Ranges of Sea Lice Species
Lepeophtheim salmonis has a circumplar distribution in the northern hemisphere ( Margolis, 1958; Kabata, 1979, 1988). This species is essentially limited to salmonid hosts including: Oncorhynchus clarki (= Salmo clurki)
(coastd cutthroat trout), Oncorhynchus gorbuscha @ink salmon), Oncorhynchus keta (chum salmon), Oncorhynchus kisutch (coho salmon), Oncorhynchus masou
(cherry or masu salmon), Oncorhynchus mykiss (= Salmo gairdneri) (rainbow or steelhead trout), Oncorhynchus nerka (sockeye salmon) , Oncorhynchus tschawytscha (chinook salmon), Sulmo salar (Atlantic salmon), Sulvelinus fontinalis (brook trout) (Kabata, 1979, 1988; Wsotten, et al., 1982; Nagasawa, et al., 1987; Hogans and Trudeau, 1989b; Tully, 1989). Kabata (1979) lists four records of non-salmonid hosts, but notes that such occurrences should be considered unusual. Bruno and Stone (1990) report low numbers of preadult L. salmonis on saithe (Pollachinus virens) collected in the vicinity of sea farms. The infestation most likely arose from farmed salmonids as saithe collected away from the sea farms were not infected with L. salmonis. Chalimus stages were not recorded therefore it is impossible to determine if L. salmonis had matured to the preadult stage on saithe or if transfer of preadults from salmon hosts occurred. - 9 - Lepenphaheirus sulmonis occurs on sea-farmed salmonids dong the Pacific
coast of Canada (present study), along the Atlantic coast of Canada (Hogans and
Trudau, 1989b), in Ireland (Tully, 1989), in Scotland (Wootten et ul., 1982), and in Norway (Brandal and Egidius, 1979).
Although L. salmonis is a marine parasite it has been reported to occur on salmonids which have recently entered fresh water (Hutton, 1923; White, 1940;
Kabata, 1981; Hahnenkamp and Fyhn, 1985).
In contrast to L. salmonis, the other species of sea lice occur naturally on a wider range of host species, mainly non-salmonids.
The distribution of Caligus clemensi extends throughout the North Pacific
(Parker and Margolis, 1964). Salmonid hosts of this species include: 0. gorbuscha, 0. keta, 0. kisutch, 0. mykiss, 0. nerka, and S. salar (Kabata, 1988; unpublished data for B.C.). This species likely occurs also on 8. tshawytscha.
Kabata (1988) lists six non-salmonid host species from the west coast of Canada
(British Columbia) including: Clupea harengur pallasi (Pacific herring),
Gasterosteus aculeatus (threespine stickleback), Hexagrammos sp. (greenling),
Hydrolagus colliei (ratfish), Sebastes caurinus (copper rockfish), and neragra chalcogramma (walleye pollock).
Caligus clemensi occurs on sea-farmed salmonids on the Pacific coast of
Canada (British Columbia) (unpublished data) and probably the northwest coast of the United States (Washington). - 10- The distribution of Caligus curtw extends throughout the North Atlantic
(Parker et al., 1968; Kabata, 1979). This species is generally considered a parasite of gadid fishes, but has a broad host range, including more than 35 species of teleosts and elasmobranchs (Parker et al., 1968; Margolis et al., 1975; Kabata,
1979, 1988). This species has been found in low abundance on S. salar at sea-farm sites on the Atlantic coast of Canada (Bay of Fundy) (Hogans and
Trudeau, 1989b). It is not considered to be a economically important parasite of farmed salmonids.
Caligus elongatus is common in the North Atlantic and has also been reported in the South Atlantic and in Australian waters (Parker, 1969; Margolis et al., 1975). This species has a very broad host range, with more than eighty species of teleost and elasmobranch hosts reported (Parker, 1969; Margolis ef ul.,
1975; Kabata, 1979, 1988). Salmonid hosts include: 0. mykiss, S. fontinulis, S. salar, and Salmo tmtta (brown trout) (Kabata, 1979, 1988; Wootten et ul., 1982;
Hogans and Trudeau, 1989a,b; Tully, 1989). Caligus elongatus is reported as common on sea-farmed salmonids along the Atlantic coast of Canada (Hogans and
Trudeau, 1989a,b), in Ireland (Tully, 1989), and in Scotland (Wootten ef al.,
1982). This species is an economically important parasite of farmed salmonids in
Atlantic Canada, Ireland, and portions of Scotland.
Lepeophtheiw cuneifer is distributed along the northern Pacific coast of
North America (Alaska and British Columbia) (Kabata, 1974; present study). Two known and eight possible non-Amonids hosts of L. cuneij2r have been identified. - 11 - Known hosts include Raja binoculata (big skate) and Hexagrammos lagocephalus
(rock greenling). Kabata (1974) listed other possible hosts as Leptocottus armatus
(Pacific staghorn sculpin), Theragra chalcogramma (walleye pollock), Zsopsetta isoZepis (butter sole), Hippoglossus stenolepis (Pacific halibut), Gadur mucrocephulus (Pacific cod), Microgadus proximus (Pacific tomcod), Lumpenus
'?, sagitta (snake prickleback), and Squalus acanthias (spiny dogfish). Lepeophtheim cuneifer has been found in low abundance on 0.mykiss and S. salar cultured in sea water in the southern Strait of Georgia, British Columbia (present study). This species is not considered an economically important parasite of farmed salmonids.
Identification of Sea Lice Species of Salmon and Trout
In generd, the preadult and adult stages of Caligus species can be easily distinguished from those of Lepeophtheim species by the presence of lunules (1,
Fig. Al).
On the Pacific coast of Canada the preadult and adult stages of C. clemensi can be distinguished from those of L. salmonis and L. cuneifer by the presence of lunules (1, Figs. A1 and A3). Identification of preadult and adult C. clemensi can be confirmed by reference to Kabata (1972, 1988) and Parker and Margolis
(1964).
Preadult and adult L. salmonis can be distinguished from L. cuneijier by differences in the structure of the basal spine of the exopod of the third leg (Fig.
82). Furthermore, L. salmonis adults can be easily recognized from L. cuneifer - 12 - adults by the shape and relative size of the genital complex and abdomen (Fig.
A3).
Identification of preadult and adult stages of L. salmonis can be confirmed by reference to Kabata (1973, 1979, 1988) and the present study. Identification of the adult stage of L. cuneifer can be confirmed by reference to Kabata (1974).
In Atlantic waters and adjacent seas the preadult and adult stages of C. curtus and C. elongatus can be distinguished from L. salmonis by the presence of lunules (Fig. Al). Preadult and adult C. currus can be distinguished from C. elongatus due to differences in the number of setae on the exopod of the fourth leg (Fig. A4). The fourth leg has 4 setae in C. currus and 5 setae in C. elon,gatus.
Furthermore, C. curtus adults can be easily distinguished from C. elongurus adults by differences in the shape of the genital complex and abdomen (Fig. Al). Species identification can be confirmed by reference to Parker et ul. (1968) and Kabata
(1979, 1988).
The following key serves to distinguish the preadults anci adults of the four species of sea lice reported from sea-farmed salmon and trout. Key to Preadult and Adult Sea Lice of Salmon and Trout.
1. Lunules absent...... 2
Lunules present...... 3
2. Basal spine of third exopod some distance from tip of basal swelling (Fig.
A2A) ...... L. cuneifer (Figs. A3A and A3B) (Pacific).
Basal spine of third exopod at tip of basal swelling (Fig. A2B) ......
...... L. salmonis (Fig. A3C and A3D) (Atlantic and Pacific)
3. Exopod of fourth leg with 4 setae (Fig. A4A); distal margin of exopod of
first leg with 4 undivided setae, seta 4 longer than others (Fig. A5A) ....
...... C. curtus (Figs. A1C and AID) (Atlantic)
Exopod of fourth leg with 5 setae (Fig. A4B and A4C) ...... 4
4. Distal margin of exopod of first leg with setae 1 and 4 undivided and
unarmed, setae 2 and 3 bifid (Fig. A5B) .....C. clemensi (Figs.
A 1E and A1F) (northeast Pacific)
Distal margin of exopod of first leg with setae 1 and 4 undivided and
armed, setae 2 and 3 appearing chelate due to presence of secondary
process arising near midlength (Fig. A5C) ...... C. elongatw
(Figs. AlG and AlH) (Atlantic) FIG. Al. Adult stages of sea lice: (A) Lepeophtheirus salmonis, female;
(B) same, male; (C) Caligus curtus, female; (D) same, male; (E) Cdgm clemensi, female; 0;) same, male; (G) Caligus elongatus, female; (H) same, male
(A-D, G, H, redrawn from Kabata 1979; E, F, redrawn from Parker and Margolis
1964). (abd, abdomen; e, egg sac; f, fourth leg; gc, genital complex; 1, lunule)
FIG. A2. Basal spine of exopod of the adult third leg: (A) Lepeophtheirm cuneifer; (B) Lepeophtheirus salmonis. Note: In both species the basal spine of the third leg exopod has essentially attained the adult form by the first preadult stage.
FIG. A3. Adult stages: (A) Lepeophtheirus cuneuer, female; (B) same, male; (C) Lepeophtheirus salnitonis, female; (D) same, male (A,B, redrawn from
Kabata 1974; C,D, original). (gc, genital complex; abd, abdomen)
FIG. A4. Structure of the adult fourth leg: (A) Culigus curtus; (IS) Caligw clemensi; (C) Caligus elongatus (A, redrawn from Kabata 1979; B, redrawn from
Kabata 1972; C, original). Note: Setation of the fourth leg in the first and second preadult stages is the same as the adult. However, its segmentation may be less distinct in the earlier stages.
FIG. A5. Distal margin of the exopod of the adult first leg: (A) Culigus curtus; (B) Caligus clemensi; (C) Caligus elongatus (A,C, modified from Kabata
1979; B, modified from Kabata 1972). Note: Armature of the terminal setae attains the adult condition by the first preadult stage.
CHAETER TWO
The developmental stages of Lepeophtheims salmonis (Kreyer,
1837) (Copepoda: Caligidae).
Introduction
The life cycle of caligid copepods consists of five phases and ten stages.
These include two free-swimming naupliar stages, one free-swimming infective copepodid stage, four attached chalimus stages, two preadult stages, and an adult stage (Kabata, 1972).
The morphology of of Lepeophtheirus salmonis and other caligid copepods varies among species considerably less during the early developmental stages than at the adult stage. Therefore, detailed morphological descriptions are required to ensure correct identification of stage as well as species.
Although L. salmonis is an economically important parasite of salmonids, the morphology of its developmental stages is poorly known. The nqupliar stages
(Johannessen, 19781, the copepodid, and some of the chalimus stages (White,
1942b) of L. salmonis have been previously described. However, these descriptions are not adequate enough to ensure correct identification. - 20 - The morphology of the adult stage of L. salmonis has been described by numerous authors and most recently by Kabata (1973, 1979). It is redescribed below, to enable a comparison to be made with the earlier developmental stages.
The objective of this part of the study was to describe the developmental stages of L. salmonis, studied with the aid of light and scanning electron microscopy. These descriptions include unique features that enable identification of all developmental stages of L. salmonis. They also help to distinquish between
L. salmonis and C. clemensi at all development stages, except the nauplius. Materials and Methods
The developmental stages were obtained by rearing from eggs in the laboratory at 10" C using chinook (Oncorhynchus tshawyrscha) and Atlantic salmon (Salmo salar) as hosts. For light microscopy the specimens were initially fixed in 10% formaiin in sea water and transferred to 70% ethyl alcohol for storage. Whole mounts as well as dissected appendages were mounted in Hoyer's mounting medium and examined using phase contrast and Nomarsky's interference phase illumination. The terminology used in the descriptions is that of Kabata
(1979). All measurements are in mm (mean + 1 standard deviation) and were made on relaxed individuals prior to fixation. Pinnules have been omitted on some figures for clarity.
Specimens for scanning electron microscopy were initially fixed in 3% gluteraldehyde at 4O C overnight. Specimens were then prepared following the procedure of Felgenhauer (1987) using 0.22 gm filtered sea water as the buffer and acetone as the transitional fluid. After critical point drying in CO,, the specimens were observed in an Autoscan scanning electron microscope operating at an accelerating voltage of 20 Kv. Results
Descriptions of developmental stages
First Nauplius
Actively swimming first nauplius (Fig. B1) irregularly oval, tapering posteriorly, external segmentation not evident, long and slender (length 0.54 rt
0.04; width 0.22 & 0.01; based on 25 specimens) compared with inactive nauplii
(length 0.47 f 0.03; width 0.25 + 0.02; based on 25 specimens). Inactive nauplii deformed; few molt to the second nauplius stage. Body translucent, with distinct pigment regions, pair of eyespots on dorsal surface. Globular deposits (yolk?) and bands of longitudinal muscles visible within the body. Posterior margin with two balancers, diverging laterally. Balancers nonsegmented, with cylindrical bases and flattened tips.
First antenna (Figs. B2 and B3) uniramous, subcylindrical, divided into two segments of approximately equal length; proximal segment with two short unarmed setae on ventral surface; distal segment (Fig. B3) with short aesthete, two longer armed setae at apex, two short spines and cuticular ridge on dorsal surface, one short spine on ventral surface. Setae armed with serrated membranes
(omitted in drawings) on opposite margins. Second antenna (Fig. EM): sympod unsegmented, fused to the proximal segments of the rami, making it appear distinctly bifid. Endopod indistinctly two-segmented, shorter than exopod; proximal segment short without armature, distal segment longer with one short unarmed spine, a short blunt spiniform outgrowth, and two large armed setae each - 23 - with serrated membrane and row of pinnules on opposite surfaces (Fig. BS).
Exopod indistinctly five-segmented, segments decreasing in size distally; proximal segment unarmed, segments two through four with small distal processes on inner
(facing the endopod) surfaces each with single large armed seta; distal segment with small spiniform process on outer surface and single large armed seta on apex. Endopod setae armed with serrated membranes directed mediad; serrated membranes of exopod setae directed laterally. Mandible (Fig. B6) biramous; sympod unsegmented, indistinctly delimited from the exopod. Endopod one-segmented; two large armed setae at apex, single unarmed seta on ventral surface. Exopod four-segmented; segments one to three with distinct processes on inner surfaces, each with one large armed seta; distal segment smdl with large armed seta on apex. Setae armed as in second antenna.
Second Nauplius
Active second nauplius similar in shape to first nauplius, but longer and narrower (length 0.56 + 0.01; width 0.20 + 0.01; based on 16 specimens).
Appendages unchanged except for endopod of second antenna (Fig. B7); size of apical spiniform outgrowth about three times size of that of first nauplius.
Copepodid
Newly molted copepodids (Fig. B8) longer and wider (length 0.70 f 0.01; width 0.28 + 8.01;based on 25 specimens) than nauplii. Body with two distinct regions: cephalothorax and a posterior region consisting of four segments.
Cephalothorax formed by fusion of cephalic segments with first two thoracic - 24 - segments. Dorsal shield present; two eyespots, 10 pairs of setules arranged
symmetrically about median longitudinal axis. Six pairs are simple setules, four
pairs bifurcate above point of origin. Rostrum (Fig. B9) with posteriorly directed
tine, two simple setules near base and two small pores (< 1 um diameter) on
adjacent margin of dorsal shield. Posterior region comprising third through fifth
thoracic segments and elongate segment. Latter segment formed by fusion of
genital segment and abdomen, with caudal rami on posterior margin. Pair of
simple setules on dorsal surface of fourth thoracic segment; pair of bifurcating
setules on dorsal surface of genitallabdominal segment. Ventral surface of
cephalothorax accommodating antennae, mouthparts and first thoracic legs (Fig.
B114). First antenna (Fig. B10) uniramous, two-segmented, cylindrical in cross
section; proximal segment with three unarmed setae on medial margin; distal
segment with thirteen elements: five setae (two arising from common base) with
irregularly branching tips on posteroventral and posterodorsal surfaces, seven
unarmed setae on anterior and ventral surfaces and one aesthete on anterior
margin. Second antenna (Fig. B 11) uniramous, three-segmented; proximal
segment small with small posteriorly directed spiniform process; middle segment
large, tapered distally with distinct cuticular ridge; distal segment (terminal claw)
sickle-shaped with short spine on base. Mouth cone (Fig. B115) similar to that of
adult, lacking strigil. Mandible (Fig. B12) uniramous, indistinctly
three-segmented; proximal segment inflated; middle segment cylindrical, slender; distal segment blade-like, armed with 10 teeth. First maxilla (Fig. B13) bipartite, - 25 - consisting of posteriorly directed blunt conical process with anteroventrally
directed palp; latter with two short and one longer unarmed setae. Second maxilla
(Fig. B14 and B116) brachiform; lacertus and brachium about equal in length;
calamus short, indistinctly delimited; canna approximately half length of calamus;
brachium with membrane on medial margin; calamus with fine membranes on
medial, anterior, and posterior surfaces; canna with one serrated margin.
Maxilliped (Fig. B15) three-segmented; corpus maxillipedis robust, unarmed;
subchela shorter, consisting of indistinctly separated shaft and claw; shaft conical,
about twice as long as claw, with branching barb near base of latter; claw
unarmed, tapering, gently curving. Tines (Fig. B117) present near base of the first
and second maxillae, projecting posteromedially to base of maxillipeds.
First thoracic legs (Figs. B16 and B17) biramous, connected by interpodal
bar; sympod two-segmented with plumose seta on lateral margin; both rami
one-segmented; exopod armed with eight elements: lateral margin with unarmed
spine, distolateral margin with two spines, both armed with fine pinnules, distal
margin with one short seta armed with strips of marginal membrane, and one
longer semipinnate seta, distal through medial margins with three long pinnate
setae; endopod (Fig. B17) with small, distolateral spiniform projection and seven long pinnate setae on distal and medial margins. Second thoracic leg (Figs. B18 and B19) similar to first leg, but with sympod indistinctly segmented, with papilliform outgrowth on medial margin; exopod (Fig. B19) lacks one spine on
apical lateral margin, medial margin with fine setules; endopod with six pinnate - 26 - setae and a smaller spiniform process. Third thoracic leg (Fig. B20) consists of
bulbous outgrowth of body wail with one short unarmed spine and one longer seta
armed with strips of serrated membrane. Caudal ramus (Fig. B21) with two short
stout pinnate setae on psterolateral margin, short pinnate seta and aesthete on distal margin and two very long pinnate setae on posteromedial margin.
First Chalimus
Early first chalimus larvae (Fig. B22) longer and wider than copepodid
(length 1.21 f 0.05; width 0.50 _+ 0.02; based on 12 specimens). Cephalothorax includes third thoracic segment; anterior region with frontal filament; posterior margin with two small indentations (posterior sinuses). Dorsal shield with simple and bifurcating setules on dorsal and lateral surfaces.
Appendage segmentation and armature generally reduced compared with those of copepodid, First antenna (Fig. B23) two-segmented; proximal segment with seven unarmed setae, six on ventral surface, one on dorsal surface; armature of distal segment similar to that of copepodid, but with shorter armature; branching setae absent, two elements with blunt tips (possibly aesthetes). Second antenna (Fig. B24) indistinctly two-segrr nted; proximal segment squat and unarmed; terminal claw smaller and more robust than in copepodid, armed with small robust seta.
Postantennary process (Fig. B118) rudimentary, psterolateral to second antenna, consisting of subtriangular process bearing bifurcating setule at base and simple setule on adjacent body wall. Mouth cone (Fig. B119) similar to that of adult (see also Kabata, 1974); formed by partial fusion of labium and labrum.
Labium with marginal membrane (mm),labial fold (If), strigil (s), and mandibles
(m) lying in labial groove (Ig). Labrum with marginal membrane on tip and buccal stylets (bs) on inner surface. Mandible (Fig. B25) indistinctly four-segmented; distal segment with stronger curve and finer point than that of copepodid, armed with 12 teeth. First maxilla (Fig. B26): posterior process more elongated; palp arising from wall closer to base, armature reduced. Second maxilla (Figs. B27 and
B120) lacertus shorter and more robust than in copepodid, basal process present; marginal membrane of brachiurn replaced by row of setules; calamus and canna more slender and curved, marginal membranes on calamus replaced by rows of spinules (Fig. B 120). Maxilliped (Fig. B28): subchela unsegmented, shorter and more robust than in copepodid, bearing smooth and slightly curved spine. Sternal furca absent.
First leg (Fig. B29) without interpodal bar; sympod indistinctly segmented, with robust naked seta on medial margin, small seta on lateral margin; exopod increased in length, with armature similar to that of copepodid, although unarmed and reduced in length; endopod narrower with two unarmed setae at apex. Second leg (Fig. B30): sympod indistinctly segmented, bearing robust naked seta on medial margin, small seta on lateral margin; exopod armed with eight setae; endopod with seven unarmed setae, generally reduced in length, when compared to copepadid. Third leg (Fig. B31) biramous; sympod unsegmented, with single unarmed seta on lateral margin; both rami one-segmented with greatly reduced - 28 - armature; exopod with seven setae, endopod with four. Some setae without distinct bases. Fourth leg (Fig. B32) represented by short bulbous outgrowth with two small naked setae. Caudal ramus (Fig. B33): setae reduced in length, unarmed; aesthete broader with tapered tip.
Secorui Chu1imu.r
Faly second chalimus larvae (Fig. B34): total length 1.52 + 0.06, maximum width 0.59 + 0.02 (based on 13 specimens). Cephalothorax with partially incorporated fourth thoracic segment, boundaries between cepkalothorax and body segments indistinct.
Only minor differences between appendages of second and first chalimus larvae. Most appendages flaccid in appearance, with obscure segmental boundaries, reduced and unevenly shaped armature. First leg (Fig. B35): exopod and endopod indistinctly separated from sympod; endopod further reduced in size relative to exopod. Second leg (Fig. B36): exopod and endopod indistinctly separated from sympod. Third leg (Fig. B37): sympod broader, expanded medially. Fourth leg (Fig. B38) indistinctly two-segmented; distal segment with three (occasionally two) minute unarmed setae. Border between caudal ramus and abdomen indistinct, medial margin with scattered pinnules.
nird Chalirnm
Third chalimus larvae (Fig. B39): total length 2.20 $ 0.08, maximum width 1.0 + 0.06 (based on 11 specimens). Cephalothorax includes fourth thoracic segment; posterior sinuses distinct; H-shaped suture lines on dorsal - 29 - shield. Ebrder between fifth thoracic segment and genital complex distinct; border between genital complex and abdomen indistinct.
First antenna (Fig. B40): proximal segment with I I setae on ventral surface, 2 on dorsal surface, some unevenly pinnate; distal segment as in adult, with 14 unarmed elements, 1 seta on posterior margin, 11 setae and 2 possible aesthetes at apex. Second antenna (Fig. B4 1) indistinctly segmented; terminal claw large relative to basall segment, with two unarmed setae and small projection on lip. Postantennary process (Fig. B42) blunt, conical, directed posterolaterally, with multiple setule on base and adjacent body wall. Mouth cone and mandible unchanged from preceding stage. First maxilla (Fig. B43): posterior process increased in size relative to palp; medial surface with minute outgrowth; palp arising from body wall anterior to posterior process, with setae reduced in length relative to posterior process; rudimentary anterior sclerite present. Second maxilla
(Figs. B44 and B121) calamus and canna longer, more strongly curved and tapered to finer points than that of previous stage; calanus with second row of spinules on anterior and posterior surfaces. Maxilliped (Fig. B45) with structure essentially unchanged.
First leg (Fig. EM): sympod indistinctly segmented, both lateral and medial setae pinnate; exopod indistinctly two-segmented, proximal segment larger with short unarmed seta on distolateral margin, distal segment with four short setae on distal margin and three longer sparsely pinnate setae on medial margin; endopod and its two setae further reduced in size. Second leg (Fig. E347) with - 30 - broad interpodal bar; syrnpod unsegmented, medial seta larger; rami indistinctly
separated from sympod; exopod indistinctly two-segmented, proximal segment
with two unarmed setae on distolateral and distomedial margins, distal segment
with four short spines on lateral margin and five long lightly armed setae on distal
through medial margins; endopod indistinctly two-segmented, with scattered
setules on lateral margin, proximal segment with one long lightly armed seta on
medial margin, distal segment with seven long lightly armed setae on medial
through lateral margins. Third leg (Fig. B48): sympod unsegmented, fused to
opposite member by broad interpodal bar, small seta on lateral margin, stout seta
on medial margin near interpodal bar, both medial and lateral setae lightly armed;
rami unsegmented; exopod with one robust seta on lateral margin and seven setae
near apex; endopod with one long robust seta on medial margin and four shorter
setae on apex; all setae unarmed. Fourth leg (Fig. B49) uniramous; sympod squat, indistinctly separated from smaller exopod, bearing lightly armed seta on distolateral margin; exopod with five minute setae on lateral and apical margins.
Fifth leg (Fig. B50) represented by outgrowth of body wall bearing two small unarmed setae on apex. Caudal ramus (Fig. B51): armature longer tha' that of preceding stage; distolateral and distomedial setae lightly armed, setules present on medial margin.
Fourth Chalimus
Fourth chalimus larvae (Fig. B52): total length 2.77 + 0.18, maximum width 1 34 f 0.1 1 (based on 12 specimens). Similar in general appearance to - 31 - third chalimus larvae, but posterior sinuses more developed, rudimentary thoracic valves present, genital complex and abdomen indistinctly separated.
Minor differences in structure of appendages between fourth and third chalimus larvae include the following: second antenna (Fig. B53): terminal claw highly sclerotized, tapering to blunt point; postantennary process (Fig. B54) longer, tapering to finer point; mandible (Fig. B55) distinctly four-segmented; first leg (Fig. B58) with interpodal bar; third leg (Fig. B59) with both rami indistinctly two-segmented, setules on lateral margin of endopod's proximal segment; fourth leg (Fig. B60) with sympod larger than exopod. More setae with scattered pinnules.
Major differences in structure of appendages between fourth and third chalimus larvae include the following: first maxilla (Fig. B56) with posterior process divided to form small medial and large lateral tines; sternal furca (Fig.
B57) present, consisting of two widely spaced tines arising from common base
(cuticular ridges anterior to insertion may represent the furcal box).
First Preadult
First preadult female (Fig. B61): total length 3.70 rf: 0.16, maximum width 2.14 + 0.20 (based on 1 1 specimens). Generally free-moving, a few remaining attached by frontal filament to host. Similar in general morphology to adult female except for differences in shape of genital and abdominal segments.
Cephalothorax: frontal plates fully developed, anterior and lateral margins with broad marginal membrane, distinct setule on both lateral margins posterior to - 32 - midpoint; posterior margin with fully developed posterior sinuses and thoracic
valves. Numerous setules present on cephalothorax, free thoracic segments,
thoracic appendages, abdomen and caudal rami, consisting of three main types:
simple (unbranched, generally short); bifurcating (two setules arising from a common base); multiple (up to eight setules arising from a common base).
Thoracic appendages, abdomen and caudal rarni generally with simple and bifurcating setules. Dorsal shield generally with multiple setules. Sensory organ
(sensu Kabata, 1981) (Fig. B122) present on ventral surface near anterior margin of cephalothorax, at point of origin of frontal filament. Posterior margin of third legs extends beyond posterior margin of fifth thoracic segment, posterior and lateral margins with broad marginal membranes. Genital complex ovoid with distinct cuticular folds on anterolateral margins. Abdomen distinctly separated from genital complex; lateral margins diverging posteriorly.
First antenna (Fig. B62): proximal segment with 20 pinnate setae, 18 on ventral surface, (15 positioned on or near anterior and apical margins), 2 on dorsal surface; distal segment unchanged from preceding stage. Second antenna
(Fig. B63) three-segmented; basal segment short, squat, with well developed posterior process; second segment robust with reniform adhesion pad on anterior surface near base of terminal claw; terminal claw strongly curved, tapered to fine point, armed with small blunt outgrowth near its base and longer unarmed seta arising at about its midlength. Sclerite on ventral body wall between base of second antenna and postantennary process (Fig. B63). Latter longer and more - 33 - tapered than in preceding stage, distinctly curved in posterornedial direction, one multiple setule on base and one on adjacent body wall. Mouth cone and mandible unchanged from preceding stage. First maxilla (Fig. B64): posterior prxess broader than that of preceding stage, medial tine smaller than lateral tine; distinct mterior sclerite between base of posterior process and palp. Second maxilla (Figs.
B65 and B123) calamus with sharper curve, tapering to finer point than in preceding stage, with additional row of spinules on anterior and posterior surfaces; brachium with small spine and patch of setules. Maxilliped (Fig. B124) anterior and lateral surfaces with regions of minute cuticular outgrowths, otherwise unchanged from preceding stage. Sternal furca (Fig. B66): base more robust than in preceding stage, tines short relative to total length; furcal box present as uneven ovoid depression.
First leg (Fig. B67): sympod distinctly two-segmented, proximal segment with setule on lateral margin, distal segment with one pinnate seta on both lateral and medial margins; exopod two-segmented, distinctly separated from sympod, proximal segment with small unarmed spine on dktolateral margin and row of setules on medid margin, distal segment with three large pinnate setae on medial margin, one shorter pinnate seta on distomedial margin and three short curving setae on apex (Fig. B68), the largest armed with row of fine denticles on concave margin, other two armed with short membrane and spine on concave margins and short row of denticles on convex margin near tips; patens on anterior apical surfaces near their bases; endopod vestigial, indistinctly separated from sympod, - 34 - with two setae on apical margin. Two anteriorly directed lobes, each with simple
setule, on ventral body wall between interpodal bars of first and second legs.
Second leg (Figs. B69 and B70) fused to opposite member by a broad interpodal
bar, with broad membrane along posterior ventral margin; sympod distinctly
two-segmented, proximal segment with long pinnate seta on medial margin and small setule on anterior surface, distal segment with small pinnate seta on lateral surface nea exopod base and setule on anteromedial margin, posterolateral and medial margins fringed with broad membrane; both rami distinctly two-segmented; exopod (Fig. B70): proximal segment over one half total length of ramus, with stout armed spine on distolateral corner, long pinnate seta and row of setules on medial margin; apical lateral margin of distal segment with two unarmed spines, one spine armed with membranes, one seta armed with strip of membrane and row of pinnules, apical through medial margins with five long pinnate setae and short row of setules; posterior surface of proximal segment and part of distal segment covered with broad membrane; endopod: proximal segment shorter than distal, with long pinnate seta on medial margin and row of setules on lateral margin, distal segment with eight long pinnate setae, its medial margin with fringe and lateral margin with broader strip of setules. Third leg (Figs. B71 and B72): sympod well developed, unsegmented, fused to opposite member by a broad interpodal bar, with stout pinnate seta on medial margin near bar, small pinnate seta on Iateral margin near base of exopod, posterolateral and posteromdial margins of sympod and posteroventral margin of interpodal bar with broad - 35 - membranes, fine denticles on posterolateral margin of sympod; both rami distinctly two-segmented; proximal segment of exopod with pinnate seta on medial margin and stout spine with membrane on lateral surface on distolateral corner, distal segment with four short unarmed spiniform setae on lateral margin and tive long pinnate setae on apical through medial margins; proximal segment of endopod with large pinnate seta on medial margin and continuous row of setules on lateral margin, distal segment with five long pinnate setae on medial through apical margins and row of setules on lateral margin. Fourth leg (Fig. B73): sympod and exopod distinct, sympod with single pinnate seta on distolateral margin, exopod with two naked setae on lateral margin and three setae armed with fine pinnules on apical margin. Fifth leg (Fig. B74) represented by bulbous outgrowth of ventrolateral surface of genital complex, with three pinnate and one naked setae. Caudal ramus (Fig. B75) essentially unchanged from previous stage, minor differences include abundant pinnules on setae, sensory setules on surfaces and more setules on medial margin.
First Preadult Mule
First preadult male (Fig. B76) similar to corresponding female, but smaller
(total length 2.90 + 0.45 and maximum width 1.8 1 + 0.16; based on 10 specimens); without cuticular folds on genital complex and diverging lateral margins of abdomen. Appendages similar to those of corresponding female, except sixth leg (Fig. B77) present, consisting of a bulbous outgrowth of posteroventral surface of genital complex, with two unarmed setae. Second Prectclult
Second preadult females (Fig. B78): tow length 5.40 ) 0.51, maximum
width 3.24 + 0.16 (based on eight specimens). General morphology similar to
that of adult female except for differences in size and shape of genital and
abdomind segments. Genital complex relatively large, with distinct cuticular
folds, lateral margins diverging posteriorly to form distinct lobes on posterolateral
comers, projecting past anterior margin of abdomi~alsegment. Lateral margins of
abdomen diverge posteriorly.
First antenna (Fig. B79) proximal segment with 27 setae, 25 (pinnate) on
ventral and 2 (unarmed) on dorsal surface; distal segment unchanged from
preceding stage. Second antenna unchanged from preceding stage. Postantennary
process (Fig. B80) with two multiple setules on base and one on adjacent body wall. First maxilla (Fig. B81): posterior process lateral and medial tines almost equal in length. Structure of mandibles, mouth cone, second maxillae,
maxillipeds, sternal furca and first legs unchanged from preceding stage. Second
leg (Figs. B82 and B83) in adult condition; rami three-segmented; proximal segment of exopod (Fig. B83) with pecten at base of distolateral seta, middle segment with large pinnate seta on medial margin and naked spine on distolateral margin, lateral margin of distal segment with one unarmed spine, one spine armed with opposing membranes and one seta armed with membrane and pinnules, five long pinnate setae on apical through medial margins; medial margins of all three segments with rows of setules; endopod: medial margin of proximal and middle - 37 - segments with one and two long pinnate setae, respectively, distal segment lateral through medial margins with six long pinnate setae, lateral margins of all three segments and medial margin of middle segment with rows of setules. Third leg
(Figs. B84 and B85) iri adult condition; synlpod with two small simple setules on anteroventral margin; exopod (Fig. B85) three-segmented, proximal segment with long pinnate seta on medial margin and stout seta armed with membrane on distolaterd margin, two small simple setules on anterior surface of segment near base of stout seta, middle segment with long pinnate seta on distomedial margin and short spine on distolateral margin, distal segment with three short unarmed setae on lateral margin and four longer pinnate setae on apical through medial margins, lateral margins of middle and distal segments and medial margin of distal segment with rows of setules; endopod two-segmented, proximal segment with large pinnate seta on distomedial margin, distal segment with six pinnate setae on lateral through medial margins, lateral margins of both segments and medial margin of distal segment with rows of setules. Fourth leg (Fig. B86): sympod with pinnate seta on lateral margin and row of four simple setules on anterior surface; exopod (Fig. B86 and B87) indistinctly three-segmented, distolateral margin of proximal segment bearing unarmed short seta with a pecten at base, distolateral margin of middle segment with armed seta with pecten at base, distal segment apical margin with three armed setae increasing in length towards rnedial margin. Fifth leg (Fig. B88), on posteroventral surface of genital complex, consists of two processes, a larger medial process with three pinnate setae and a - 38 - smaller lateral process with one pinnate seta. Surfaces of caudal rami with additional simple setules, otherwise unchanged from previous stage.
Second Prewfult Male
Second preadult male (Fig. B89): total length 4.27 + 0.52, maximum width 2.60 + 0.40 (based on six specimens). Cephalothorax similar in morphology to that of corresponding female. Genital complex different, ovoid in shape, without cuticular folds on anterolateral comers. Lateral margins of abdomen diverge posteriorly.
Appendages similar to those of female with the following differences: fifth leg (Fig. B90) consisting of bulbous outgrowth of genital complex with four pinnate setae; sixth leg (Fig. B90), on posterolateral corner of genital complex, consists of bulbous outgrowth with three pinnate setae.
Addt Femak
Additional descriptions of adult body and appendage morphology are given in Kabata (1973, 1979). Ovigerous and postovigerous females (Fig. B91): total length 9.96 f 1-55, maximum width 4.16 + 0.35 (based on 26 specimens).
Cephalothorax ovoid, with shallow posterior sinuses and posterolateral lobes projecting posteriorly to level of posteromedial margin. Fifth thoracic segment shorter and narrower than genital complex. Genital complex of young adult females with conspicuous cuticular fold on anterolateral comers, similar in shape to that of second preadult female. Genital complex of ovigerous and postovigerous females markedly larger than that of young adult females, without cuticular folds, - 39 - with rounded anterolateral comers, parallel lateral margins and prominent posterolateral lobes. Genital region (Fig. B92) on psteroventral surface of genital complex near insertion of abdomen. In inseminated females consists of cuticular girdle (cg), two spermatophores (s) attached to vaginal openings, and two oviduct orifices (ov). Spermathecae and shell glands visible within the genital complex.
Abdomen one-segmented, cylindrical in cross section, in ovigerous and postovigerous females approximately same length as genital complex.
First antenna unchanged from preceding stage. Second antenna (Fig. B93): posteromedial process on proximal segment more pronounced than in preceding stage; dorsal surface of middle segment with two small adhesion pads; terminal claw more robust than in preceding stage. Postantennary process, mandible and mouth cone unchanged. First maxilla (Fig. B94): medial and lateral tines of posterior process almost equal in length; setae of palp further reduced in size cornpared to posterior process; anterior sclcri te well developed, with cuticularized process on posterolateral corner. Second maxilla (Figs. B95 and B96) brach ium longer relative to lacertus, more tapered towards tip, midregion setules replaced by strips of serrated membrane (flabellum); calamus and canna more slender and strongly curved, canna with small curved barb on inner margin near base.
Maxilliped (Fig. B97): medial margin of corpus with two small sensory processes.
Sternal furca (Fig. B98) with robust base and blunt spatulate tines, older specimens commonly with tines unequally eroded, furcal box present as oval depression about base. - 40 - First leg unchanged from preceding stage. Second leg: exopod (Fig. B99) with small pecten near base of distolateral seta of proximal segment; all previously unarmed setae of the lateral margins of the middle and distal segment now armed with opposing strips of membrane. Third leg: exopod (Fig. B100): large lateral spine of proximal segment armed with opposing strips of membrane, basal swelling of this spine with membrane and two simple setules. Fourth leg (Figs.
BlOl and B102): sympod and exopod approximately equal in length, sympod more slender than in previous stage; segmental boundaries of exopod more distinct, medial margins with distinct membranes, all armature with pectens at bases. Fifth leg (Fig. B103) on ventral surface of genital complex lateral to oviduct orifices; consists of large medial outgrowth with three pinnate setae and smaller lateral outgrowth with one pinnate seta. Caudal ramus (Fig. B104): distal setae with distinctly inflated bases, relatively short compared with other elements.
Adult Male
Adult males (Fig. B105): total length 5.40 $ 0.48, maximum width 3.25
+ 0.35 (based on 15 specimens). Cephalothorax similar to that of female. Genital complex ovoid, approximately same width as lateral margins of fifth thoracic segment. Abdomen one-segmented, cylindrical, shorter than genital complex.
All appendages similar to those of female except as follows. Second antenna (Figs. B106 and B107) more robust; proximal segment elongate with distinct adhesion pad on posterior surface; middle segment with two adhesion pads on posterior surface and three smaller adhesion pads on anterior surface; terminal - 41 - claw sharply flexed in distal half with two small flanges on posterior and anterior surfaces, one short robust seta on base and one longer slender seta at midpoint.
First maxilla (Fig. B108): posterior process with additional small thin-walled tine near apex. Postoral adhesion pad (Fig. B125) on body wall, posteromedial to base of first maxilla, consisting of distinct outgrowth of body wall with series of distinct cuticular ridges. Maxilliped (Fig. B109): anterior surface of corpus with distinct region of cuticular outgrowths; subchela longer and more slender, with larger basal seta. Sternal furca (Fig. Bl10): tines longer relative to total length, tapering to finer points, tips commonly unequally eroded.
Fifth legs (Fig. B111) on lateral margins near midpoint of genital complex, consisting of outgrowth of body wall armed with four pinnate setae. Sixth legs
(Fig. 31 12) on psterolateral corners of genital complex, consisting of outgrowth of body wall armed with three pinnate setae. Caudal ramus (Fig. B113): terminal setae longer than in female, lacking inflated bases. Discussion
Within the family Caligidae three other species of Lepeophtheirus and six
species of Culigus have had all or most of their devetopmental stages described.
These species include Lepeophtheirus dissimulurus (cf. Lewis, l96?),
Lepeophtheirus hospitulis (c f. Vot h , 1972) Lepeophtheirus pectoralis (c f. Boxshall,
1974a), Cu&igtucentrodonti (cf. Gurney, 1934), Culigtcs curtus (cf. Heegaard,
1947), Culigus orientulis (cf. Hwa, 19651, Culigus spinosus (cf. Izawa, 1969),
Culigw cl~merzsi(cf. Kabata, 1972) and Culigus pqeti (cf. Ben Hassine, 1983).
Of these, the descriptions of C. cenrrodonri (cf. Gurney, 1934) and C. currus (cf. Heegaard, 1947) lack many of the morphological details necessary for comparative purposes. Furthermore, the structure of many of the appendages of
C. curtus appears to be at variance with those described for other species. The source of this vanance is most commonly a misinterpretation of appendage segmentation. These misinterpretations may have been due to the translucent nature of the exoskeleton, which commonly allows muscles and the structure of the develsging appendages to be visible within, or to folds of the exoskeleton which commonly occur after molting.
In the description of C. spinosus (cf. Izawa, 2969) only three chalimus stages are reported. Kabata (1972) suggested that the fourth chalimus stage was either not present or simply missed by Izawa. However, based upon the level of development seen in the structure of the thoracic legs of Izawa's second and third chalimus stages, I feel that the second chalimus stage was missed. Therefore, - 43 - throughout this discussion I refer to Izawa's second and third chalimus stages as the third and fourth chalimus stages, respectively.
The development of C. pageti consists of two naupliar, one copepodid, four chalimus, one preadult, and an adult stage (Ben Hassine, 1983). The fourth chalimus stage although attached by a frontal filament has fillly developed lunules and a marginal membrane on the cephalothorax. Fully developed lunules and the presence of the marginal membrane are features of the first preadult shge of other
Caligus species. A portion of the first preadults of C. ckmensi, C. spinosus, and most species of Lepeophrheirus are reported as attached by a frontal filament
(Izawa, 1969; Lewis, 1963; Voth, 1972; Boxshall, 1974a; this study), For these reasons I feel that the first preadult stage has been misidentified by Ben Hassine as the fourth chalimus stage. The level of development seen in the appendages of the second and third chalimus stages is similar to that reported for the third and fourth chalimus stages of other Culigus species. Therefore, it is likely that either the first or second chalimus stage was missed by Ben Hassine. For the purposes of this discussion I refer to Ben Hassine's second, third, and fourth chalimus stages as the third and fourth chalimus and the first preadult stages, respectively.
Lepeophtheim sulmonh and other caligids undergo a major metamorphosis with the molt from the second naupliar to the copepodid stage. Development from the copepodid stage to the adult stage is more gradual. The rnoit from the copepodid to the first chalimus stage is generally accompanied by a loss of distinct segmentation of both the body and the appendages, as well as a reduction in the - 44 - length and complexity of the elements of appendage armature. In L. salmonis and most other species further reductions occur with the molt to the second chalimus stage. With the molt to the third chalimus stage and continuing through to the adult stage there is generally a progressive development of body and appendage segmentation, as well as a general increase in the complexity and number of its components. As reported by Boxshall (1974a) for L. pectoralis, the different appendages of L. scslmonis undergo this process of reduction and then development at different rates. In general, the adult condition of the appendages in L. sulmonis and other Lepeophtheirus species is reached later in development than in Caligus species.
Female L. sulmotiis, like L. pectoralis (cf. Boxshall, 1974a), undergo a considerable change in the size and shape of the genital complex with the change from the pre-ovigerous to ovigerous condition as well as smaller changes during the preadult stages. The folds of cuticle reported at the anterior end of the genital complex in newly molted preadult and adult females may serve as a reservoir of cuticle, enabling these changes to occur without molting. Folding of the cuticle has been shown to enable the 'elongation phase' of the abdomen in adult female
Lernaocm brunchidis (cf. Smith and Whitfield, 1988).
In L. salmonis and other Lepeophtheirus species, sexes can be first distinguished on the basis of morphology in the first preadult stage. In Caligus species, differences between the sexes are commonly apparent earlier in development. Differences have been reported as early as the third chalimus stage - 45 - of C. clemensi (cf. Kabata, 1972) and the fourth chalimus stage of 41'. oriuncta1i.s
(cf. Hwa, 1965).
As early as the fourth chalimus stage, L. salnzonis and other
Lepeophtheirus species can be differentiated from Caligus species by their lack of lunules.
The developmental pattern of the first antenna in L. strlttmis and L. pectoralis (cf. Boxshall, 1974a) is identical up to the second preadult stage. In both of these species the number of elements on the proximal segment generally increases throughout development. In L. sulmonis the adult condition of 27 setae is reached by the second preadult stage. Peculiarly in L. pectorulis, 29 setae are reported on the proximal segment in the adult (Boxshall, 1974b). The armature of the distal segment attains the adult condition in the third chalimus stage of both species.
The setal numbers reported during the development of the first antenna of
L. dissimulutus (cf. Lewis, 1963) and L. hospitdis (cf. Voth, 1972) appear to be incorrect.
In C. clenzensi the armature of the distal and proximal segments of the first antenna attains the adult condition by the third and fourth chalimus stages respectively (Kabata, 1972). Unfortunately, the descriptions of the first antenna in
C. orientalis (cf. Hwa, l965), C. spinosus (cf. Izawa, 1969), and C. pugeti (cf.
Ben Hassine, 1983) are inadequate for comparative purposes.
In L. salmonis, L. dissimulatus (cf. Lewis, 1963), L. hospitulis (cf. Voth, - 46 -
1972), and C. spinow (cf. Imwa, 1969) the second naupliar stage may be
distinguished from the first by an elongation of the apical process of the endopod
of the second antenna. This process gives rise to the third segment (terminal claw)
of the copepodite and later stages (Izawa 1969). In L. salmonis and other
Lepeophtheim species the structure of the second antenna is similar to that of the
copepodid up to the third chalimus stage, at which time the terminal claw is
modified into an elongate process. With the molt to the first preadult stage the
second antenna of both males and females closely resembles that of the adult
female. In the adult stage the structure of the second antenna becomes sexually
dimorphic, the males bearing large adhesion pads and modified claws.
In Culigm species the terminal claw of the second antenna is reduced
throughout the chalimus stages. However, by the preadult stages the second
antenna resembles that of the adult. Sexual dimorphism in the structure of the
second antenna is apparent earlier in development of these species. This has been
noted in the third chalimus stage of C. clemensi (cf. Kabata, 1972), in the fourth
chalimus stage of C. orienraiis (cf. Hwa, 1965), and in the first preadult stage of
C. spinosm (cf. Izawa, 1969) and C. pageri (cf. Ben Hassine, 1983).
Rudimentary forms of the postantennary process are present in the copepodite stage of C. ciememi (cf. Kabata, 1972), the first chalimus stage of L. salnzonis (this study) and L. pectoralis (cf. Boxshall, 1974a), and the second chalimus stage of C. orientalis (cf. Hwa, 1965). However, it is not until the third chalimus stage of all species studied that the postantennary process appears in a - 47 - form similar to that of the adult.
The mouth cone of the copepodid stage of L. salmonis lacks the strigil seen in later developmental stages. Variation in the structure of the mouth cone during ontogeny has not been reported in the other species studied.
In all Lepeophtheirus and Caligus species the mandible essentially attains the adult condition by the first chalimus stage, although its segmentation may become more distinct at latzr stages.
Postoral adhesion pads are present in the adult male of L. salmonis as well as in the second preadult male, adult male and adult female of L. pectoralis (cf.
Boxshall, 1974a; 1974b). Although not described by Voth (1972), postoral adhesion pads are also present on the male of L. kareii (= L. hospitalis (cf.
Yamaguti, 1936; Lopez, 1976)). They have not been reported in L. dissimulutus
(cf. Lewis, 1963) or species of Caligw.
The pattern of development seen in the first madla of L. salmonis is identical with that described for L. pectoralis (cf. Boxshall, 1974a; 1974b). In L. hospitalis (cf. Voth, 1972) the posterior process is not reported as bifid until the first preadult stage. Although Voth (1972) reports no difference in the structure of the first maxilla between adult males and females, both Yamaguti (1936) and
Kabata (1973) have reported a trifid posterior process in the adult male.
Lepeophtheim dissimulam differs from the other Lepeophtheirus species as its posterior process is bifid in the first and second chalimus stages and simple in all other stages (Lewis 1963). In C. clemensi (cf. Parker and Margolis, 1964; - 48 - Kabata, 1972), C. orienrulis (cf. Hwa, 1965), and C. pugeti (cf. Ben Hassine,
1983) the posterior process remains simple throughout development. In C.
spinosus (cf. Izawa, 1969) the posterior process is simple up to the fourth
chalimus stage and bifid in both sexes thereafter.
In L. sulmonis the first maxilla of the copepodid is similar in form to those
of adults of more primitive siphonostome genera (e.g. Dissonus) (cf. Kabata,
1966; 1979). By the first preadult stage the first maxilla has attained the form characteristic of addt caligid copepods.
Development of the second maxilla in L. sulmonis is similar to that reported for other species. In general, changes involve differences in the relative proportions of its lacertus and brachium as well as differences in the shape and curvature of the calarnus and canna.
The maxilliped develops in a similar manner in L. sulinonis and other
Lepeophtheirus and Culigus species. The most dramatic change in maxilliped structure occurs with the molt to the first chalimus stage. In C. clemensi (cf.
Kabata, 1972) and C. orientalis (cf. Hwa, 1965) the spiniform processes on the corpus maxillipedis and subchela of the copepodite maxilliped are lost with this molt. The presence of these processes in C. clemensi is one feature that allows it to be distinguished from L. salmonis in the copepodite stage.
Posteriorly directed tines have been reported on the ventral body surface anterior to the maxillipeds in the copepodite stages of L. sulmonis (present study),
L. pecmralis (cf. Boxshall, 1974a), C. clemensi (cf. Kabata, 1972), and C. - 49 - orientalis (cf. Hwa, 1965). It is not clear whether these structures are homologous to the true sternal furca of later developmental stages.
The true sternal furca may first be seen developing beneath the exoskeleton of the third chalimus stage (Lewis, 1963; Boxshall, 1974a; present study) or as ridges of cuticle at the point from which it will arise fHwa. 1965; Kabata, 1972).
In L. salmonis, as in other Lepeophrheim and Caligus species except C. pugvri and C. spinosus, the sternal furca first appears in the fourth chalimus stage. In C. pageti the sternal furca is first reported in the third chalimus stage (Ben Hassine,
1983). However, it is not clear whether the sternal furca is above or beneath the larval exoskeleton in this stage. In C. spinosus it is first reported in the first preadult stage (Izawa, 1969). The presence of the sternal furca is an important feature for differentiating the fourth chalimus stage from earlier stages.
The pattern of development seen in the first leg of L. sulmonis is virtually identical with that described for other Lepeophtheirus and Culigus species. Culigus clemensi differs from all other species as it has only one seta on the endopod of the frrst chalimus through adult stages (Parker and Margolis, 1964; Kabata, 1972), whereas all other species have two.
Except for a difference in the number of endopod setae in the second chalimus stage, the development of the second leg of L. ~ulmonisis identical with that reported for t.pectoralis (cf. Boxshall, 1974a; 1974b). Lepeophtheim dissirnulatm (cf. Lewis, 1963) and L. hospitalis (cf. Voth, 1972) have a similar pattern of development of the second leg. However, the numbers of setae reported - 50 - in the first through third chalimus stage of both species and in the first preadult stage of L. rlissirnulunis differ. In all Lepeoplztheirus species the broad marginal and surface membranes first appear in the first preadult stage and the adult condition of this appendage is reached by the second preadult stage.
In C. clemensi (cf. Kabata, 1972) and C. orientulis (cf. Hwa, 1965) the second leg is essentially the same as that of the adult in the fourth chalimus stage, although the broad surface and marginal membranes are not present until the preadult stage. Unfortunately, only one preadult stage has been described for these species. In C. spinosus (cf. Izawa, 1969) and C. pugeti (cf. Ben Hassine, 1983) the adult form of the second leg is reached in the first preadult stage.
Differences in setation of the first and second legs enable us to distinguish between the copepodid and chalimus stages of L. salmonis and C. clemensi.
However, caution should be exercised when counting the endopod setae of the first leg as they become very tiny and undifferentiated from the segmental margin.
In some individuals of L. salmonis additional setae are present on the endopod of the first leg.
The development of the third leg in L. salrnonis is very similar to that seen in other Lepeophrheim species. With respect to setation, L. salmonis differs from
L. pectorcalis (cf. Boxshall, 1974a; 1974b) only in the first chalimus stage. In
L. dissimuiarus and L. hospiralis there are generally fewer setae reported on the first through third chalimus stages (Le.wis, 1963; Voth, 1972). In L. salrnonis the rami become two-segmented in the first preadult stage whereas in other - 51 - Lepeophtheirus species they are described as two-segmented from the third chalimus stage. In all Lepeophtheirus species membranes tht appear on the margins of the third leg in the first preadult stage and the adult condition of the appendage is reached by the second preadult stage. In species of Culigus the third legs are essentially the same as those of the adult by the fourth chalimus stage, although the broad surface and n-arginal membranes are not present until the preadult stages.
The fourth leg first appears in the first chalimus stage of all described
Lepeophtheivus species and in the second chalimus stage of all described Cu1igu.s species except C. pugeti (sf. Ben Hassine, 1983). Development of the fourth leg is more rapid in Culigus species. All species report a fourth leg similar in structure to that of the adult by the fourth chalimus stage.
The fifth legs first appear in the third chalimus stage of L. salmonis, L. dissimulutus (cf. Lewis, 1963), L. hospitalis (cf. Voth, 1972), L. pcctorcrlis (cf.
Boxshall, 1974a), C. orientulis (cf. Hwa, 1965), and C. pugeri (cf. Ben Hassine,
1983). In C. clemensi (cf. Kabata, 1972) and C. spinosus (cf. Izawa, 1969) the fifth leg first appears in the first preadult stage.
The sixth legs first appear in the first preadult males of L. sulmonis and all other species studied. Hwa's (1965) description of the sixth leg in the third chalimus stage of C, oriemdis is an error resulting from his improper interpretation of the structure of the rudimentary fifth leg.
In summary, the development of L. sulrnonis is almost identical with that - 52 - rejxxted for L. pectoralis (cf. Boxshall, 1974a) and very similar to that reported
for L. dissimulutus (cf. Lewis, 1963) and L. hospitalis (cf. Voth, 1972). The appendages of L. sulmonis and other Lepeophtheirur species generally develop at a
slower rate than those of Culigus species. The sexes are generally distinguishable earlier in the development of Culigus species. Culigus species can be distinguished from Lepeophrheirus species as early as the fourth chalimus stage, owing to the presence of rudimentary lunules. With the exception of the naupliar stages, L. safmonis can be distinguished from the co-occuring C. clemensi by differences in appendage morphology . FIGS. B1 to B7. Lepeopfzrheirus sulmonis, nauplius stages. Fig. B1. Active
first nauplius, dorsal. Fig. B2. First and second nauplius, first antenna, anterior.
Fig. B3. Same, tip, anterodorsal. Fig. B4. First nauplius, second antenna, posterior. Fig. B5. Detail of antenna seta. Fig. B6. First and second nauplius, mandible, anterior. Fig. B7. Second nauplius, second antenna, tip of endopod, posterior. (ae, aesthete; b, balancers; cr, cuticular ridge; ed, endopod; e, exopod).
FIGS. B8 to B 15. Lepeclphtheinrs suln~onis,copepodid. Fig. B8. Newly molted copepodid, dorsal. Fig. B9. Rostrum, ventral. Fig. B10. First antenna, ventral. Fig. B11. Second antenna, lateral. Fig. B12. Mandible, lateral. Fig. 1313.
First maxilla, lateral. Fig. B14. Second maxilla, lateral. Fig. B 15. Maxilliped, lateral. (ae, aesthete; cr, cuticiilar ridge; p, palp; pp, posterior process; , simple setule; , bifurcated setule).
FIGS. 3 16 to B28. Lepeophtheirus salmonis, copepodid and first chalimus.
Fig. B16. Copepodid, first leg, posterior. Fig. B17. Same, endopod, posterior.
Fig. B18. Seccnd leg, anterior. Fig. B19. Same, exopod, anterior. Fig. R20.
Third leg, ventral. Fig. B21. Caudal ramus, ventral. Fig. B22 Early first chalimus, dorsal. Fig. B23. First antenna, ventral. Fig. B24. Second antxna, lateral. Fig. B25. Mandible, lat~ral.Fig. B26. First maxilla, medial. Fig. 827.
Second maxilla, lateral. Fig. B28. Maxilliped, anterior.
FIGS. 829 to B38. Lepe~~phtheinrssulmonis, first and second chali mus.
Fig. B29. First chalimus, first leg, posterior. Fig. B30. Second leg, posterior.
Fig. B31. Third leg, posterior. Fig. B32. Fourth leg, dorsal. Fig. B33. Caudal ramus, ventral. Fig. B34. Second chalimus, dorsal. Fig. B35. First leg, posterior.
Fig. B36. Second leg, posterior. Fig. B37. Third leg, posterior. Fig. B38. Fourth leg, dorsal.
FIGS. B39 to B5 1. Lepeophtheirus salmonis Third chalimus. Fig.
B39. Third chalimus, dorsal. Fig. B40. First antenna, ventral. Fig. 841. Second antenna, lateral. Fig. B42. Postantennary process, ventral. Fig. 843. First maxilla, anterior. Fig. B44. Second maxilla, lateral. Fig. B45. Maxilliped, posterior. Fig. B46. First leg, anterior. Fig. B47. Second leg, posterior. Fig. B48.
Third leg, posterior. Fig. B49. Fourth leg, dorsal. Fig. B50. Fifth leg, ventral.
Fig. B51. Caudal ramus, ventral.
FIGS. B52 to B60. Lepeophtheirus sulmonis Fourth chalimus. Fig. B52.
Fourth chalimus, dorsal. Fig. B53. Second antenna, lateral. Fig. B54.
Postantennary process, ventral. Fig. B55. Mandible, lateral. Fig. B56. First maxilla, lateral and medial surfaces. Fig. B57. Sternal furca, anterior. Fig. B58.
First leg, anterior. Fig. •’359.Third leg, anterior. Fig. B60. Fourth leg, anterior.
FIGS. B61 to B69. Lepeophtheirus salmonis First preadult. Fig. F61. First
preadult female, dorsal. Fig. B62. First antenna proxinaal segment, ventral. Fig.
B63. Second antenna and postantennary process, ventrd. Fig. B64. First maxilla,
ventral and medial surfaces. Fig. B65. Second maxilla, anterior. Fig. B66. Sternal
furca, anterior. Fig. B67. First leg, anterior. Fig. B68. Same, tip of exopod, anterior. Fig. B69. Second leg, posterior. (s, sclerite).
FIGS. B70 to B77. Lepeophrheirus salmonis First preadult. Fig. B70.
Second leg exopod, anterior. Fig. B71. Third leg, anterior. Fig. B72. Same, rami,
anterior. Fig. B73. Fourth leg, anterior. Fig. B74. Fifth leg, ventral. Fig. B75.
Caudal ramus, ventral. Fig. B76. First preadult male, dorsal. Fig. B77. Male fifth and sixth legs, ventral.
FIGS. B78 to B87, Lepeophrheirus salmonis Second preadult. Fig. B78.
Second preadult female, dorsal. Fig. B79. First antenna proximal segment, ventral. Fig. BSO. Postantennary process, ventral. Fig. B81. Posterior process of first maxilla, ventral. Fig. B82. Second leg, posterior. Fig. B83. Second leg exopod, anterior. Fig. B84. Third leg, anterior. Fig. B85. Third leg rami, anterior. Fig. B86. Fourth leg, anterior. Fig. B87. Fourth leg tip of exopod, anterior.
FIGS. B88 to B96. Lepeophtheirus salmonis Second preadult and adult.
Fig. 888. Fifth leg, ventral. Fig. B89. Second preadult male, dorsal. Fig. B90.
Second preadult male fifth and sixth legs, ventral. Fig. B91. Adult female, dorsal.
Fig. B92. Genital field, ventral. Fig. B93. Second antenna, posterior. Fig. B94.
First maxilla, anterior. Fig. B95. Second maxilla, anterior. Fig. B96. Second maxilla armature, anterior. (cg, cuticlular girdle; ov, oviduct orifice; s, spermatophore)
FIGS. B97 to B 107. Lepeopkrheirus salmonis Adult female and male. Fig.
B97. Maxilliped, anterior. Fig. B98. Sternal furca, anterior. Fig. B99. Second leg exopod, anterior. Fig. B100. Third leg exopod, anterior. Fig. B101. Fourth leg, anterior. Fig. B102. Fourth leg tip of exopod, anterior. Fig. B103. Fifth leg, ventral. Fig. B104. Caudal ramus, ventral. Fig. B105. Adult male, dorsal. Fig.
B106. Second antenna, posterior. Fig. B107. Second antenna, anterior.
FIGS. B 108 to B 1 13. Lepeophlheirus salmonis Adult male. Fig. B108.
First maxilla, anterior. Fig. l3109. Maxilliped, anterior. Fig. B110. Sternal furca, anterior. Fig. B111. Fifth leg, ventral. Fig. B112. Sixth leg, ventral. Fig. B113.
Caudal ramus, ventral. (pap, postoral adhesion pad).
FIGS. B 114 to I31 17. Lepeophtheirus salmonis, copepodid. Fig. B 114.
Cephalothorax, ventral. Scale bar = 50 pm. Fig. B115. Mouth cone, tip, ventral.
Scale bar = 10 pm. Fig. B116. Second maxilla, tip, anterior. Scale bar = 5 pm.
Fig. B117. First maxilla and tines, ventromedial. Scale bar = 20 pm. (bs, buccal stylets; fa, first antenna; fm, first maxilla; If, labial fold; lg, labial groove; m, mandible; mc, mouth cone; mm, marginal membrane; mxp, maxilliped; sa, second antenna; sm, second maxilla; t, tine).
FIGS. B 118 to B12 1. Lepeophrheirus salmonis, first and third chalimus.
Fig. B118. First chalimus, postantennary process, ventral. Scale bar = 5 pm.
Fig. B1119. First chalimus, mouth cone, ventral. Scale bar = 20 pm. Fig. B120.
First chalimus, tip of second maxilla, anterior. Scale bar = 10 pm. Fig. B121.
Third chalimus, tip of second maxilla, anterior. Scale bar = 20 pm. @s, buccal stylets; If, labial fold; lg, labial groove; m, mandible; mm, marginal membrane; s, strigil).
FIGS. B122 to B125. Lepeophtheirw salmonis First preadult and adult male. Fig. B122. First preadult female, frontal organ, dorsal. Scale bar = 20 pm.
Fig. B123. First preadult female, tip of second maxilla, anterior. Scale bar = 20 pm. Fig. B124. First preadult female, maxilliped cuticular ridges, anterior. Scale bar = 20 pm. Fig. B125. Adult male, postorai adhesion pad. Scale bar = 20 pm.
CHAPTER THREE
Development, growth, and survival of Lepeophtheirus salnonis
(Copepoda: Csligidae) under laboratory conditions.
Introduction
The life cycle of Lepeophtheirus salmonis consists of ten stages. These
stages include two free-living planktonic nauplliar stages, one free-swimming
infectious copepodid stage, four attached chalimus stages, two preadult stages, and an adult stage (Chapter 2). Both of the naupliar stages and the copepodid stage prior to its attachment to the host are non-feeding. Attached copepodids, chalimus larvae, preadults, and adults feed on host mucus, skin, and blood (Kabata, 1974;
Brandal et al., 1976; Jones et al., 1990; present study).
Although L. salmonis is an economically important parasite, most aspects of its basic biology have been poorly documented. The effects of temperature on the duration of the egg bearing period (Johannessen, 1978), the duration of the naupliar stages, and survival of the infectious copepodid stage (Johannessen, 1978;
Wmtten et al., 1982) have been previously studied. However, these investigations were limited by the poor survival of L. salmonis lawae in the laboratory. The effects of temperature on the duration of the copepodid, chalimus, preadult, and the - 69 - salmonis nauplii and adults, and surprisingly reports that nauplii are more tolerant to changes in salinity than are the adults.
This chapter reports the effects of temperature and salinity on development, growth, and survival of L. salmonis under labratory conditions. This information is important to our understanding of the dynamics of epizootics of L. salmonis. - 70 - Materiais and Methods
Ovigerous L. salmonis were collected from sea-farmed and wild chinook
(Omrhynchus tschawytschu (Walbaum)) and sea- farmed Atlantic salmon (Salmo
sular (Linnaeus)) from Quadra Island and Departure Bay on the east coast of
Vancouver Island, Canada. Where possible copepods were collected when the
water temperature corresponded to the experimental temperature at which they
were to be incubated. Possible maternal effects were minimized by using eggs or
nauplii from several females in each stage of my investigations.
In most experiments eggs and developing nauplii were cultured in glass jars
(250 ml to 3.5 1) covered with 100 pm Nitex mesh. The jars were suspended in
tanks of flowing seawater (28.5 to 30.5 %O salinity) and set temperatures (f 0.5"
C). Water exchange and circulation within the jars was maintained by gentle aeration.
Egg development times were determined using the indirect regression technique of Edmondson (1965). Entire collections of ovigerous females were divided into groups of five and cultured at either 5, 10, or 15" C in flowing sea water. Cultures were monitored daily from 8:00 to 23:OO hours at two hour intervals. Time to hatching for each female was recorded and the number of ovigerous females remaining at each time plotted against time from the start of the experiment. The least-squares regression of numbers of ovigerous females against time was calculated for each temperature. The point of intersection on the time axis is taken to be the mean development time. This technique is based on the - 71 - assumption that the age distribution of the eggs is uniform within the sample.
Although selective predation on females with eggs or embryos has been shown to result in a violation of this assumption for some free-living zooplankton species
(see Threlkeld, 1979), I feel that this assumption is valid for the eggs of L. salmonis as the females are not free-swimming and no natural predators are known.
To determine the duration of the naupliar stages, eggs were natched in tanks of flowing sea water with a temperature of 5, 10, or 15" C. Actively swimming first nauplii that hatched over the preceding two hour period were pipetted into 250 ml culture jars and returned to their respective tanks. Moulting activity was monitored every two hours from 8:00 to 23:00 hours. Time of development is defined as the time from hatching to the first observation of either the second nauplius or copepodid stage.
Copepodids that developed at 10" C and had moulted within the previous
12 hours were used to infect previously uninfected Atlantic salmon. Infections were carried out in dark aerated tanks with no water flow. Exposure times ranged between 8 and 12 hours. The fish were maintained in 10" C flowing sea water, killed at intervals and examined for parasites. All developmental stages of the copepod were identified and their position on the host recorded. For each copepod, measurements were made of total length (anterior margin to the base of furca), and maximum eephalothorax width (excluding marginal membrane). Time of development is measured from the time at which the eggs were extruded. The - 72 - duration of the chdimus and preadult stages was determined from observed changes in stage frequency following the methods of Landry (1983). Time of development is defined as the time when 50% of the population had moulted to the next stage, as estimated from least-squares regression of the proportion of the population which had completed a given moult versus time. The data were arcsine square root transformed to meet the assumptions of the regression model (Zar,
1984).
To determine the effect of salinity on hatching success and survival to the coppodid stage, ten ovigerous females with non-pigmented eggs were pipetted in individual 250 rnl culture containers and incubated at 9 to 10" C in static seawater baths of 10, 15, 20, 25, and 30 950 salinity, and a flowing seawater bath of 30 to
30.5 750 salinity. Water in the static baths was changed every second day.
Salinities were adjusted by adding glass-distilled water. Cultures were checked at
1 day post-hatchinit when unhatched eggs, and dead and moribund nauplii were counted and removed. At 5 days post-hatching the remaining nauplii and copepodids were counted.
Actively swimming copepodids which had molted within the previous twelve hours were transferred to individual test tubes containing 20 mls of .45 prn filtered sea water of various salinities ( 15, 20, 25, or 30 %o) and the temperature at which they were reared (5, 10, or 15" C). Tubes were incubated at their respective temperatures and mortality was assessed daily. Copepodids were considered dead when they failed to respond to mechanical stimulation. Best - 73 - transformation of the survival data was estimated using the procedure of Box et al., (1978). All data were log (x+ 1) transformed and differences in survival investigated by a two-way analysis of variance. Multiple comparisons of survival times at each temperature were made using a Scheffe's test (Zar, 1984). Results
For each of the 3 experimental temperatures, the number of ovigerous
females plotted against time, the least squares regression equation, and the
calculated mean egg development times are presented in Fig. C 1. Mean egg development times were 4 19.1 hours (17.5 days) at 5" C, 207.1 hours (8.6 days) at 10" C, and 130.8 hours (5.5 days) at 15" C.
The time from first to last nauplius hatched was highly variable, ranging from 18 to 65 hours (mean = 3 1.7 _+ 13.0 hours; n = 16) for egg strings maintained at 10" C in 30 Y6 static water. Egg numbers ranged between 251 and
423 (mean = 344.6 + 79.8; n = 16). There was no correlation with number of eggs and duration of hatching. Egg strings attached to the female began hatching from the posterior end. Those released by the female began hatching at any point along their length.
The duration of the first nauplius stage was shorter than that of the second nauplius stage at all temperatures (Table Cl). The average duration of the first nauplius stage varied from 52 hours (2.2 days) at 5" C to 9.2 hours at 15' C. The average duration from hatching to the copepodid stage varied from 222.3 hours
(9.3 days) at 5" C to 44.8 hours (1.9 days) at 15" C.
The time to hatch, naupliar development times, and the calculated median development times for the chalimus and preadult stages of L. salmonis raised at
10" C are summarized in Table C2 and Figure C2. The two naupliar stages developed quickly compared with the copepodid stage, which required almost 7 - 75 - days for 50% to moult to the first chalimus stage. The third chalimus and both preadult stages also had protracted stage durations. The first adult males were obtained 40 days after egg extrusion and two adult females were obtained at 52 days after egg extrusion.
There is a large range of variability in developmental rates between individual copepods, with the time between the first and last appearance of the stages far exceeding the initial 12 hour difference in ages (Table C2, Fig. C2).
Although both male and female first preadults first appeared at 32 days after egg extrusion, males reach maturity before females. Both adult males and second preadult females first appeared on day 40. Of the 123 preadults and adults obtained 61 % were males and 39 % were females.
The percentage moult increments (sensu Hartnoll, 1982) calculated from the mean sizes demonstrate no consistent trends over development (Fig. D3 and
D4). There are marked increases in total length and cephalothorax width with the moults to the copepodid and third chalimus stages. With the moult from the second preadult to adult female there is a marked increase in total length and a relatively small increase in cephalothorax width.
In the growth experiments approximately 53 % of the copepodid and chalimus larvae were attached to the tips of the gill filaments and 33% to the fins
(Fig. D5). Of those on the fins the majority were on the pelvic (14 %), pectoral
(9%),and anal (7%) fins. Larvae attached to the body were most commonly found dong the margin of the operculum. Of the preadults collected, 15 % had retained - 76 - their frontal filament. Unattached preadults and adults were found on the body
surfaces, but most commonly on the posterodorsal surface of the head and posterodorsal to the dorsal fin.
Egg strings maintained at 10" C and 10 %O in static water failed to develop
(Table C3). In static water of 15 and 20 %O salinity, high proportions of the egg strings hatched, but active nauplii were obtained only at 20 %o. At 20 %O from 0 to 89.9%(mean = 19%)of the eggs developed to the copepodid stage, but only one active copepodid was obtained. At higher salinities 100% of the egg strings hatched and the percentage of eggs that produced active nauplii ranged from 9.7 to
95%. At 25 %O from 0 to 2.9%(mean = 0.9%)of the eggs developed to the copepodid stage, but only one active copepodid was obtained. At 30 %O the percentage of eggs that produced active copepodids ranged from 0 to 80.6%.
In preliminary experiments at 10" (3 newly moulted copepodids survived for less than 3 hours when transferred to 5 %O salinity water, and less than 1 day when transferred 10 %O salinity water. At higher salinities (15 to 30 %o)and temperatures of 5, 10 and 15" C survival was prolonged. Maximum survival time was 17 days at 10" C and 25 ko salinity, with average survival times ranging between 2 and 8 days (Fig. D6). A comparison of survival times (2-way ANOVA) indicated that temperatures (p _( 0.001) and salinities @ I0.001) had significant effects on survival and that there was a significant interactive effect of these two factors @ 5 0.001). (A significant interaction indicates that the direction of difference among salinities varied among temperatures.) The results of multiple - 77 - range tests (Scheffk's test; p 5 0.05) over each temperature showed survival times to be significantly higher at 30 YM when compared to 15 %O at all temperatures. At IO" C survival time at 25 %O was significantly higher than survival at 15, 20, or 30 %. At all temperatures copepodids at low salinities (15 and 20 %o) were generally less active than those maintained at higher salinities
(25 and 30 %o).
At 9 to 10" C adult female copepods which were removed from their fish hosts survived for a maximum of 12 days (mean = 9.5 + 2.5; n = 5) and 13 days (mean = 9.3 + 2.3; n = 10) in static water baths of 10 %O and 15 %O salinity, respectively. At higher salinities survival was prolonged with a maximum survival of 18 days (mean = 13.9 + 3,8; n = 16) at 30 to 31 %o. Discussion
This is the first comphrehensive study of development, growth, and
survival of L. salmonis under controlled laboratory conditions. In this discussion I
cite results obtained for the pennellid copepds Lemaeocera branchialis and
Lernueenicus sprattae and for the lernaeopodid copepod Sdminicola californiensis.
These species represent the only other siphonostome copepods outside of the
family Caligidae for which development, growth or survival data are available. I
have included them to supplement the limited data available for caligid copepods.
Our time to hatch of 8.6 days at 10" C corresponds poorly with previous determinations for L. salmonis of 10 to 14, 25, and 33 to 33 days at llSO,9S0, and 9" C, respectively (Johannessen, 1978). At 15" C the time to hatch of 5.5 days corresponds well with the 5.7 to 6.1 days reported for Lepeophtheirus kareii (=
Lepeophtheirus hospitalis) at 15" C (Lopez, 1976). Other determinations of time to hatch for Lepeophtheim species have been made at higher temperatures. These include: 2.7 days for Lepeophtheim hospitulis at 20•‹ C (Voth, 1972), and 1.3 to
1.7 days for Lepeophtheirus dissimularus at 23" C (Lewis, 1963). The eggs of L. salmonis develop faster than those of pennellid copepods at comparable temperatures. Egg development takes approximately 12.7 days at 10" C for L. branchialis (cf. Whitfield et al., l988), and 16 days at 7 to 9" C for L. spruttae
(Schram, 1979).
In my experiments the hatching period of the egg strings was highly variable and longer than previously reported. Johannessen (1978) reports that the - 79 - hatching period of L. salmonis egg strings containing 100 to 500 eggs was less than 40 hours, with the majority of the nauplii hatching within 5 to 10 hours. It is difficult to compare the hatching rates of L. sulmonis with other species of parasitic copepods due to differences in egg number and rearing temperature.
Izawa (1969) reports the egg strings of Caligus spinosus, which carry between 10 to 20 eggs per string, hatch in 3 to 8 hours at approximately 20" C. The hatching period for a pair of egg strings of Lernueoceru branchiulis, commonly containing an average of 1145 eggs per pair, was 12 days at 10" C, although most hatched within the first 3 days (Whitfield et al., 1988). Egg strings of Lernueenicus sprattae, which contain between 50 and 600 eggs, hatch in 4 to 5 hours at 15" C
(Schram & Anstensrud, 1985).
In my experiments, the moult to the second nauplius stage took 30.5 hours at 10" C and 9.2 hours at 15" C. Development to the copepodid stage took 87.4 hours at 10" C and 44.8 hours at 15" C. These durations correspond well with previous studies on L. salmonis: 35 hours at 9.2" C, 12 hours at 15.5" C
(Johannessen, 1978), and 18 hours at 12" C (Wootten et ul., 1982) for the moult to the second nauplius stage, and 77 hours at 9.2" C, 63 hours at 1 lo C
(Johannessen, 1978), and 46 hours at 12" C. (Wootten et ul., 1982) for development to the copepodid stage. In L. hospitulis, the moults to the second nauplius and copepodid stages take approximately 24 hours and 64 hours, respectively, at 15" C (Voth, 1972). In contrast, Lopez (1976) reported deveIopment to the copepodid stage to take 120 to 150 hours at 15" C in L. kureii - 80 -
(= L. hospitalis). Other determinations of naupliar development in Lepeophtheirus species have been made at higher temperatures. For L, hospitalis, the average duration of the first nauplius stage is 7.5 hours (range = 6 to 9 hours), and the average duration of the second nauplius stage is 11 hours (range = 10.5 to 12 hours) at 20" C (Voth, 1972). For L. dissimulurus, the average duration of the first nauplius stage is 6.5 hours (range = 4.5 to 13 hours), and the average duration of the second nauplius stage is 14.5 hours (range = 9.5 to 19 hours) at approximately 23" C (Lewis, 1963).
Determinations of naupliar development times have been made for several
Caligus species. Hogans & Trudeau (1989) reported that the duration of the second naupliar stage of Caligus elongatlcs is 35 hours at 10' C. Ben Hassine
(1983) reports naupliar development times for Caligus pageti over a wide range of temperatures (16 to 26" C). With one exception the duration of both the first and second naupliar stages is reported to be 24 hours at each of these temperatures.
Based upon my studies at 15" C I feel that the observation period in Ben Hassine's study was too long to determine accurately the naupliar development times, and that development may be faster than reported.
In the pennellid copepod L. sprattae, development from the first to the second nauplius stage takes 23 to 27 hours and development from the first nauplius to the copepodid stage 48 to 51 hours at 15" C (Schram & Anstensrud,
1985). - 81 - With the exception of C. elongalus (cf. Hogans & Trudeau, 1989), the duration of the first nauplius stage is shorter than that of the second nauplius stage in all caligid species studied. In both L. salmonis and L. hospitulis the duration of the copepodid stage is relatively long. Moulting to the first chalimus stage occurred approximately 6 to 8 days after host contact in L. hospitalis at 15O C
(Voth, 1972). The duration of the copepodid stage in L. dissinnulutus is unknown.
The relatively long duration of the copepodid stage may be due to the need to recover energy lost during development of the non-feeding naupliar stages, or the requirement of additional time or energy for completion of development prior to filament production and moulting. A similar trend is seen in the free-living copepods where the duration of the pre-feeding naupliar stages is relatively short
(possibly due to energetic considerations), and the duration of the first feeding instars relatively long (Landry , 1983).
The earliest adult male was obtained at 32 days and the earliest adult female at 40 days after egg hatching for L. salmonis at 10' C. For L. pecrorulis,
Anstensrud (1990) reports development to adult males in 24 to 27 days and adult females in 29 to 32 days from egg hatching at 10 to 12" C and 30 to 31 %O salinity. Other studies on Lepeophtheirus species have been conducted at higher temperatures. Development from the first chalimus stage to the second preadult stage took 21.5 days at 20" C for L. hospitulis (cf. Voth, 1972) and 8.5 days at
23" C for L. dissimulatus (cf. Lewis, 1963). - 82 - Development rates have been investigated for several species of Caligus. In
C. elongutus, development from the first chdimus to newly moulted adult takes
approximately 3 weeks at 12" C (Hogans & Trudeau, 1989). In C. pageii
development from first nauplius to second preadult takes 23.5 days at 16 to 18" C,
16 days at 20 to 22" C, and 10.5 to 11.5 days at 5.4 to 26O C (Ben Hassine, 1983).
I estimate the generation time of L. salmanis to be 7.5 to 8 weeks at 10" C.
Other estimates of the generation time of L. salmonis have been made based on field data. Wootten et al., (1982) reported a generation time of approximately 6 weeks at 9 to 12" C, and Tully (1989) reported generation times between 7 to 13 weeks depending on water temperature and the development stage used for the calculation.
In the caligids L. salmonis, Lepeophtheirus pectoralis, the pennellid L. branchiulis, the lernaeopodid S. californiensis, and many free living copepods, males mature faster than females (present study; Anstensrud, 1989, 1990; Kabata
& Cousens, 1973; Landry, 1983). Adult males of L. salmonis commonly establish precopula with first and second preadult females (present study; Wootten el al.,
1982). In both L. salmonis a?d L. pecioralis copulation occurs upon moult to the adult female and results in the female genital openings being sealed by the spermatophores. It is therefore advantageous for the males of a cohort to mature first and develop reproductive products.
The percent moult increments calculated from the size data for L. suhmis do not decrease with increasing size as suggested for a wide variety of Crustacea - 83 - (reviewed in Hartnoll, 1982). The large increase in total length with t!!e moult to the adult female occurs after the moult when the genital complex undergoes a considerable change in size and shape with the change from the pre-ovigerous to ovigerous condition (Chapter 2).
In my experiments the majority of the copepodid and chalimus larvae were collected from the gills. Copepodids and chalimus larvae of L. salmonis have not been previously reported from the gills of salmon. Wootten et a!. (1982) reported that chalimus larvae of L. salmonis are most commonly found attached to the dorsal and pelvic fins and the area around the anus, although it is not clear whether the gills were examined. Gills may serve as an important site for initial attachment of copepodids.
Eggs of L. salmonis fail to develop in 10 9~ salinity water at temperatures of 18 and 12" C @resent study; Wwtten et al., 1982). Johannessen (1978) reported that most eggs of L. salrnonis aborted, and that the few nauplii produced only lived for a short time at 11.5 Y'i salinity and temperatures of 5 to 12" C.
Berger (1970) reported that the nauplii of L, salmonis survived for 32 and 48 hours when transferred to water with salinities of 8 and 12 %o, respectively. All nauplii of the pennellid L. sprattae died when transferred to 10 %O salinity water at temperatures between 5 and 25" C (Schram & Anstensrud, 1985). At 15 960 eggs of L. salmonis developed and hatched, but produced few active naupiii.
Schram & Anstensrud (1985) report that some nauplii of L. spruttue moulted to the second nauplius stage, but none survived to the copepodid stage when - 84 - transferred to water of 15 760 salinity. However, when transferred to salinities of
20 to 30 YW most nauplii (60 to 80 % of the first nauplii) survived to the copepodid stage. The eggs of L. salmnis that developed and hatched at higher salinities (20, 25, and 30 %) produced active nauplii. At 20 and 25 %O the majority of the active nauplii died at the copepodid moult. With exception of several individuals, active coppodids were obtained only at 30 9ik salinity.
Additional stresses imposed in the laboratory (eg. handling, less than optimal water quality) may limit survival to the copepodid stage at less optimal salinities.
Lepeophtheirus salmonis, like Lemaeenicus sprattat?, may be excluded from low salinity areas (< 15 760)due to reduced hatching success and survival of the naupliar and copepodid stages.
In my experiments, average survival of the copepodid stage ranged between 2 and 8 days, depending on temperature and salinity. Similar results were obtained by Wootten et al. (1982) who reported that copepodids of L. salmonis remained active for 4 days at 12" C. Although Johannessen (1978) reported that one copepodid of L. salmonis survived for 30 days it is unlikely that many copepodids are able to survive for this length of time. Voth (1972) reported an average survival time of 4.5 days (range 3 to 8 days) at 20" C. for copepodids of
L. hospitalis. Whitfield et al. (1988) reported a maximum survival time sf 18 days and a 50 96 survival time of 7.5 days at 10" C. for copepodids of the pennellid L. branchialis. As noted by Whitfield et aZ. (1988) copepodid infectivity is likely to decline substantially over this time. - 85 - Adult L. salmonis survived on average 9.5 days after removal from their host at 10" C and 10 760 salinity. In contrast, Ekrger (1970) reported that adult L. salmonis survive for less than 12 hours at 12 % salinity and 1 hour at 4 %O salinity. Hahnenkamp & Fyhn (1985) demonstrated that L. salmonis females were able to maintain a hyperosmotic state at a salinity of 12.4 %, but started to die within 8 hours when exposed to fresh water. The reduced average survival times for adults at low salinities may be due to a higher energy requirement for maintenance of a hyperosmotic state. TABLE Cl. Man time (SD) in hours to first moult of the second nauplius and copepodid stages of Lepeophtheirus salmonis, maintained at three temperatures and ambient salinity (29.0 to 31.0 %o). n = number of cultures that moulted
Development Temperature stage So C lo0 C 15" C
Nauplius I to 52.0 (4.5) 30.5 (2.1) 9.2 (1.8) Nauplius 11 n=5 n = 12 n=5
Nauplius I to 222.3 (4.4) 87.4 (5.0) 44.8 (1.2) Copepodid n=5 n = 10 n=5 TABLE C2. Cumulative development time (CDT), time of first and last appearance, and duration of each Lepeophtheim salmonis developmental stage at 10" C.
Cop.
Chl
Prel Male Female
Pre2 Male Female
Adult male
Adult female
" time to first appearance (direct observation) time to when 50% of the population had attained this stage (dculated from least squares regression of arcsin transformed data) " last copepods obtained on day 52. TABLE C3. Hatching success of egg strings of Lepeophtheirus salmonis, and the mean percentage of total eggs (TE) that produced active nauplii and copepodids at 10" C and salinities of 10 to 30 960. Values are the mean and range (in parentheses); sample size = 10, except for 25 %O where the sample size = 7; nd = no development.
Hatching Active Active Salinity Success Nauplii Copepodids
10
15
20
25
30 (static)
30 (flowing) FIG. C1. Mean development time of eggs of Lepeophrheirus sulmonis at various temperatures and ambient salinity . Hours From Start of Experiment
Temperature Regression Mean Egg Development * C Equation Time hours (days) FIG. C2. Development sequence of a cohort of Lepeophtheirus salmonis at
10" C and ambient salinity (Cop, copepodid; Ch 1, first chalimus, Ch2, second chalimus; Ch3, third chalimus; Ch4, fourth chalimus; Prel , first preadult; Pre2, second preadult, A, adult). 1 Pre2 (male)
Time (days) FIG. C3. Percentage moult increments of total length for Lepeophrheirus salmonis raised at 10" C (Nl, first nauplius; N2, second nauplius; other abbreviations as in Fig. C2). male female
Molt FIG. C4. Percentage moul t increments of cephalothoi-ax width for
Leophtheim salmonis raised at 10" C (Nl, first nauplius; N2, second nauplius;
other abbreviations as in Fig. C2). male
, . i..,...... , ...... , ,., :.:.: zs;sz.;.."".'.'.".:.:.:. ::::.:.:.:.x.:.:::. female
Molt FIG. C5. Distribution of copepodids and chalimus larvae of
Lepeophtheirus salmonis on bady surfaces of Atlantic salmon smolts from laboratory growth experiments, 60 50 n E 40 )r a 30 3 g 20 i2 10 0 om-3''- Q) -
h om a 0 Fins Position on Body FIG. C6. Mean (+ SI)) survival time of newly moulted copepodids of
Lepeophtheincs salmonis at various temperatures and salinities. Values above error bars indicate which salinities have statistically significant differences in survival
(Scheffb's test; p 0.05). Salinity %O CHAPTER FOUR
The morphology and ultrastructure of the alimentary tract of
Lepeuphtheinss salmonis (Copepoda: Caligidae) .
Introduction
The structure of the alimentary tract of parasitic copepods has been investigated in numerous species using light microscopy (Table Dl). These studies include the closely related species Lepeophtheirus dissirnulatus in which Lewis
(1961) identified two cell types in the midgut. These cell types include non-vacuolated columnar cells (type A cells) that are absorptive in function and vacuolated cells (type B cells) that are secretory in function.
At the ultrastructural level, the alimentary tract of only a few species of parasitic copepods has been investigated. Three cell types (referred to as type A,
B, and C cells) have been described from the midgut of the closely related species, Caligus minimus (cf. Poquet, 1980). Type A cells, which are absorptive, have long microvilli, abundant endoplasmic reticulum, and lipid deposits. Type B cells, which are secretory, have short microvilli, and large vacuoles that in mature cells occupy most of the cell volume. Type C cells, which are excretory, have intermediate length microvilli, abundant vesicles, and concretions with a calcareous appearance. - 96 - Three cell types (referred to as type A, B, and C cells,) have been
described from the midgut of Lmmeenicus spratae (cf. El Gharbi, 1984). Type A
cells which are absorptive, have a few short microvilli and few organelles. Type B
cells which are both absorptive and secretory, have long microvilli, large vacuoles, and abundant rough endoplasmic reticulum, Type C cells (similar to the type B cells of Poquet (1980), and the vacuolar cells of Yoshikoshi and K6
(1991a)) are separated into two types (C1 and C2) based on differences in their cytoplasm and the presence of a cell cycle in type C2. Both cell types are digestive and type C2 cells, in addition are secretory.
Two cell types (referred to as vacuolar and non-vacuolar cells) have been described from the midgut of the parasitic copepods Conchyliunrs quintus,
Osrrincola koe, Panaietis yrutugmii, Neoergasilus japonicus, and Lemaea fyprilzacea (cf. Yoshikoshi and K6, 199la). Vacuolar cells (similar to the type B cells of Poquet (19801, and the type C cells of El Gharbi (1984)), which are secretory, have sizeable endocytotic and secretory vacuoles. Non-vacuolar cells
(similar to the type A cells of Poquet (1980)), which are absorptive, have long microvilli, abundant rough endoplasmic reticulum, and lipid deposits.
With exception of a brief mention of the midgut structure of the chalimus stage by Jones er al. (1990) the structure of the alimentary tract of L. salmonis is unknown. The objective of this study was to describe the structure of the alimentary tract of the adult L. salmonis studied based on light, scanning, and - 97 - transmission electron microscopy observations, and to compare it with that of other free-living crustaceans and parasitic copepods. TABLE Dl. Summary of species of parasitic copepsds in which the gross morphology and/or the structure of the alimentary tract has been described based on light microscopy observations.
Species Reference
Suborder Poecillostomatoida
Family Ergasilidae
Ergmilus minor Halisck (1939)
Ergmilus sieboldi Einszpom (1 965)
Suborder Cydopoida
Family Mytilicolidae
Mytilicola intestinalis Durfort (1975)
Suborder Siphonostomatoida
Family Caligidae
Lepeophtheirus dissimufatus Lewis (1961)
Lepeophtheirus pectoralis Scott (1901), Boxshall (1986,1990)
Family Lernaeopodidae
Lernaeopoda scyllicola Gray (1929)
Pseudocharopinus dentatus Rigby and Tunnel1 (1971)
Family Nanaspidae
Allantogymus species Changeux (1960)
Family Pennellidae
Lernaeenicus sayori Honma and Ho (1988)
Lernaeolophus aceratus Honma and Ho (1988) Table D 1. continued
Perodema cylindn'cum Monterosso (1930) El Saby (1933)
Lernaeocera brachialis Schuurmans Stekkoven and Punt (1937) Capart (1948)
Family Pseudocycnidae
John and Nair (1975).
Family Saccopsidae
Saccopsis steemtrupi Bresciani and Lutzen (1 96 1) (= Melinnacktes steemtnrpi)
Family Ineertae sedis
Gonophysema gullmarensis Brescaini and Lutzen (1 960) Materiais and Methods
Adult male and female L. salmonis were collected from chinook
(Oncorhynchus kisutch) and Atlantic salmon (Salmo saZar) held in net pens at the
Pacific Biological Station, Nanaimo, British Columbia.
For light microscopy, specimens were fixed in 10% buffered formalin or
Davidson's solution, dehydrated through to 100% alcohol, and embedded in JB4
plastic resin. Sections were cut to a thickness of 1 to 2 pm and stained with Lee's
stain (methylene blue and basic fuschin).
For scanning electron microscopy, specimens were initially fixed in 3%
glutaraldehyde in Mallonic's buffer at 4" C for at least 24 hours. They were then
prepared following the procedure of Flegenhauer (1987) using Mallonic's solution
as the bufier and acetone as the transitional fluid. Internal structures were revealed by embedding the dehydrated specimens in wax, following the procedure of Oshel
(1985), and sectioning the specimens in the desired plane on a microtome.
Sectioned specimens were de-embedded by reversing the embedding process back to absolute alcohol, transferred to acetone, and critically point dried using CO,.
Specimens were observed in a Autoscan scanning electron microscope operating at an accelerating voltage of 20 Kv.
For transmission electron microscopy, specimens were initially fixed in 4% glutaraldehyde in Mallonic's buffer with 4%NaCl at room temperature for 2 hours then transferred to 4O C crvernight. This was followed by rinsing with several changes of fresh buffer and post-fixation in 2% osmium tetroxide for 2 to - 101 - 3 hours at room temperature. After washing and dehydration through an acetone series, the specimens were infiltrated and embedded in either Spurr's or Epon resin. Semi-thin sections were cut on glass knifes and stained with methylene blue.
Thin sections were cut on a diamond knife, mounted on grids, and stained with uranyl acetate and lead citrate (Reynolds, 1963). Sections were examined using a
Phillips 300 TlEM operating at 80kV. Description of the alimentary tract
The alimentary tract of adult L. salmonis consists of four parts: mouth cone, oesophagus, midgut, and hindgut. Eaeh part is described in detail below.
Mouth Cone
In males and females the structure of the mouth cone is the same. The walls of the mouth cone are formed by the partial fusion of the non-muscular labium and the highly muscularized labrum between which the shafts of the mandibles pass (Fig. D4). The tip of the labium bears the marginal membrane, labial fold, strigil, and the mandible blades, resting in the labial groove (Figs.
B119, D4 and D5). The labrum bears the marginal membrane on its tip and the buccal styletes on its inner surface. Additional descriptions of the mouth cone and associated mouthparts of L. salmonis are given in Kabata (1974b) and in Chapter
Oesophagus
The structure of the oesophagus and its associated tissues is the same in both males and females. A cuticle-lined oesophagus passes from the buccal cavity and joins to the midventral surface of the midgut near the level of the maxiilipeds
(Fig Dl). Two pairs of buccal levator muscles are associated with the oesophagus throughout most of its length (Figs. Dl and D2). These muscles insert anteriorly on the buccal apodemes and posteriorly on the postmaxillulary apodemes. The position of the postmaxillulary apodemes is maintained by broad muscle bands - 103 - which pass both anteriorly and posteriorly to insert on both the dorsal and ventral body walls. These muscles and the levators enable the perpendicular movement of the mouth cone (see also Kabata, 1974b; Boxshall, 1986; 1990). Contraction of the buccal depressor muscles which run from the anterior margin of the buccal apodeme and insert on the body wall near the base of the labrum allow the mouth cone to be folded back against the body wall. Anterior to the nerve ring (formed by the cerebrum, circumoesophageal commissures and suboesophageal ganglion) one dorsal and two ventral nerve cords are associated with the oesophagus (Fig.
D2). The oesophagus and its associated tissues are maintained in position by connective tissue and the nerve ring through which they pass.
In transverse section the lumen of the oesophagus is highly folded (Fig.
D2). The thickness of the oesophageal cuticle increases as it nears the midgut, where a valve like structure is formed (Fig. D3). The cuticle consists of two layers, a thin (5.2 to 6.6 nm) epicuticle and a thicker fibrillar procuticle (Figs. ]US and D9). The outer layer of the procuticle (p2) has an electron-dense outer region which ranges in thickness from approximately 100 to 200 nm and an electron-light inner layer which ranges in thickness from about 1.0 to 1.5 pm, Along the anterior portion of the oesophagus light bodies (ranging up to approximately 200 nm in diameter) occur throughout the $ layer, but are most common in the dark layer adjacent to the lumen (Fig. D9). In the posterior portion of the oesophagus the number of light bodies declines and none is present near the junction with the midgut (Fig. D10). The inner layer of the procuticle @') consists of a region of - 104 - electron-dense granules that commonly extends between the epithelial cells (Fig.
D8). As the oesophagus nears the midgut the thickness of the p' layer increases
markedly.
The lateral cell membranes of the epithelial cells are indistinct. The apical
and basal cell membranes are irregular in shape but not highly folded (Fig D8 and
DlO). The nuclei are elongate, approximately 6 to 7 pm in length and up to 4 pm
in diameter, with a predominant nucleolus positioned usually towards one end.
Mitochondria and irregularly shaped endoplasmic vesicles are abundant within the
cytoplasm. Some rough endoplasmic reticulum is also present. The oesophageal
epithelium is enclosed by a thin basal lamina, a layer of striated circular muscle, and discrete bands of longitudinal muscle. A nerve cord lies between these
muscles and the oesophageal epithelium on the dorsal. aspect (Fig. D2).
Mi& ur
The midgut forms the major portion of the alimentary tract of L. salmonis.
It extends anteriorly past the insertion of the oesophagus to form a small anterior
midgut caecum and two small anterior midgut diverticula (Figs. Dl and Dl 1). In the female the anterior midgut caecum is supported by connective tissue that inserts on the dorsal body wall and ovaries and the diverticula are supported by connective tissue which inserts on the ventral surface of the ovaries. In both males and females the midgut is roughly triangular in shape up to the fifth thoracic segment. it is supported from its apex and base by connective tissue that inserts on the dorsal and ventral body walls. Additional support for the midgut in females is - 105 - provided by connective tissue which inserts on the ventral and lateral surfaces of the ovaries. In both females and males the midgut is laterally compressed in the fifth thoracic segment and supported by connective tissue that inserts on dorsoventral muscle bands and the ventral and dorsal body walls. In the genital complex the midgut is triangular in cross-section in the female and ovoid in cross- section in the male. In both females and males the midgut is ovoid in cross-section throughout most of the abdomen with exception of the region near the junction with the hindgut where it becomes laterally compressed. Throughout the genital segment and abdomen the midgut is supported by connective tissue that inserts on the dorsal, ventral, and lateral body walls.
Well-developed circular muscles connected by finer occasionally branching longitudinal muscles form a mesh-like structure over the external surface of the midgut and are responsible for gut peristalsis (Fig. D6). Short projections of the longitudinal muscles and the connective tissue that supports the midgut insert directly on the basal lamina (Fig. D6). The basal lamina is prom and ranges in thickness from approximately 1 to 4 pm. The pores, which are generally less than
1.0 pm in diameter, can be seen on the external surface of the gut wall (Fig. D6).
At the light microscopy level the midgut lacks distinct zones and appears to have the same cell types present throughout its length. Epithelial cell division and growth result in the epithelial cells being forced apically to form the bulbous topography of the midgut lumen (Figs. D7, Dl 1 to B14). Although these ingrowths may be in excess of 60 pm in height, all cells retain their connection - 106 - with the basal lamina, resulting in regions sf densely packed cell membranes at the center of the ingrowths. Cells of the midgut appear to have distinct cell cycles which leads to their eventual degeneration and release sf cell contents into the midgut lumen. Immature cells are found towards the base of the ingrowths and mature cells at the apex. Dense microvilli that can exceed 5 pm in length cover the internal surfaces of the midgut (Fig. D7).
Five cell types were distinguished in the midgut, and have been designated as type A through E. These designations do not necessarily correspond to the designations of rnidgut cell types given by other authors.
Cells of the midgut epithelium appear to arise from embryonic cells, type
E cells (Figs. D12, Dl5 and D16). These cells occur in groups near the bnse of the ingrowths and do not appear to contact either the basal lamina or the lumen of the rnidgut. They are undifferentiated, containing a few mitochondria, some ribosomes, and a small amount of rough endoplasmic reticulum (Fig. D16).
Type A cells were observed only in the most anterior regions of the midgut
(Figs. Dl8 to D21, D24). These cells are of various shapes and sizes depending on their position with respect to ingrowths of the gut lumen. They have long microvilli, length 4 to 5 pm, ovoid to reniform nuclei with predominant nucleoli, and distinct regions of microtubules surrounding the nuclei. Large irregularly shaped vacuoles, which increase in size as the cells mature, form along the outer surfaces of the microtubule region. Mitochondria, lysosome-like dense bodies, electron-dense vacuoles, phagosomes, and irregularly shaped light vesicles are - 107 - abundant at the cell apex (Fig. D20). These irregularly shaped vesicles appear to coalesce to form the larger vacuoles which surround the nuclei (Fig. D21).
Mitochondria and some endoplasmic reticulum occur in the basal portions of this cell type. The basal membranes of type A cells are nc, highly folded.
Type B cells (vacuolar cells) occur in low numbers throughout the midgut
(Figs. D 13, D18, D22 and D23). These cells are atso of various shapes and sizes, depending upon their position with respect to the midgut wall ingrowths. They are characterized by a distinct sub-apical complex, and the presence of large, irregularly shaped, electron dense vacuoles, which in mature cells occupy mast of their volume. Type B cells have short microvilli, length 0.5 to 0.8 pm, and appear to exhibit extensive pinocytotic activity that results in the formation of large numbers of small electron-translucent pinocytotic vesicles (Fig. D22). These vesicles appear to fuse to form larger, electron-light vacuoles. VacuoIes which contain material with a similar electron-density to that of the dense vacuoles, as well as material with a similar electron density to that of the light vacuoles, commonly occur in the vicinity of the light vacuoles (Fig. D22). Scattered mitochondria and numerous Golgi bodies are present among the dense vacuoles.
Abundant rough endoplasmic reticulum, mitochondria, and the nuclei occur towards the base of these cells (Fig. D23). The cisternae of the endopfasmic reticulum are commonly distended suggesting the storage of cell products. - lo8 - Although extrusion was not observed during this investigation, portions of
older type B cells were present within the lumen of the gut at both the light and
electron microscopy level.
Type C cells are the predominant cell type in the midgut (Figs. D17, D25
to D34). The morphology of this cell type differs between copepods which are feeding on mucus and those which are feeding on blood. In both mucus- and blood-fading copepods the apical cell membranes of type C cells extend into long microvilli, length 4.0 to 5 .O pm, which contain distinct filaments. Mitochondria, rough endoplasmic reticulum, and small vesicles are abundant at their apices.
Their nuclei are ovoid to reniform, with predominant nucleoli. Each nucleus is surrounded by a region of dense microtubules that increases in volume as the cells mature.
Type C cells of mucus-feeding copepods have short single cisternae of rough endoplasmic reticulum, unevenly shaped vacuoles with a light heterogenous content, and distinct intercellular spaces (Figs. D25 to 027). In contrast, type C cells of blood-feeding copepods have large amounts of rough endoplasmic reticulum that is arranged in both parallel arrays and whorls associated with the nuclei (Figs. B30 and D31, D34), numerous Golgi bodies which give rise to electron-dense bodies common in the subapical regions of early stages of these cells (Fig. D33), and lipid deposits that in mature cells may occupy most of the cell volume (Figs. D31 and D32). Rough endoplasmic reticulum was commonly - 109 - seen to be in contact with the lipid deposits. In these instances small deposits of electron-dense material were visible within the cisternae (Fig. D32).
Mature type C cells undergo a process of cellular degeneration in which the apical cell membrane breaks down and the cellular contents are released into the lumen of the midgut. Portions of these cells were commonly observed in the midgut lumen (Fig. D29).
Type D cells (dark cells) were observed infrequently in sections of the anterior midgut (Figs. Dl8 and D24). These cells have long microvilli, length 4.0 to 6.0 pm and electron-dense cytoplasm. Numerous mitochondria and short cisternae of endoplasmic reticulum occur at their cell apices (Fig. D24). Their nuclei are ovoid to reniform and surrounded by regions of microtubules. The middle to basal regions of these cells contain large amounts of endoplasmic reticulum, numerous Golgi bodies, and mitochondria. The basal cell membranes of type D cells appear to be more highly folded tbm those of other cell types.
Hindgut
The midgut opens into a relatively short, cuticle-lined hindgut which terminates at the anus on the dorsal surface. Numerous muscles responsible for the dilation of the hindgut connect the hindgut to the dorsal, ventral, and lateral surfaces of the anal somite (see also Boxshall, 1990).
The cuticle of the hindgut increases in thickness and changes in structure as it nears the anus. The cuticle of the anterior region of the hindgut is thin, generally < 1.0 pm in thickness, highly folded, and consisting of a single fibrillar - 110 - layer (Figs. D35 and D36). The cuticle of the midregion of the hindgut consists of
two layers, a thin < 5.0 nm epicuticle and a thicker fibrillar procuticle which
ranges in thickness up to approximately 2.0 pm (Fig. D37). Light bodies (ranging
up to approximately 80 nm in diameter) which appear similar to those seen in the
oesophagus and electrondense regions which appear to be composed of
microtubu2es are present within the procuticle. In the vicinity of the anus the
procuticle has a similar structure to that of the external cuticle.
Two epithelial cell types occur in the anterior hindgut (Figs. D35 and
D38). These include: 1) electron-transparent cells which contain scattered mitochondria, short cisternae of endoplasrnic reticulum, and irregularly shaped clear vacuoles, and 2) electron-dense cells which contain abundant mitochondria, short cisternae of rough endoplasmic reticulum, scattered Golgi bodies, and electron-dense minerai concretions.
One epithelial cell type occurs in the middle to posterior regions of the hindgut (Fig. D37). These cells have short apical subcuticular microvilli and marked infotdings of their basal plasma membranes. Mitochondria are abundant in both the apical and basal regions of these cells. Short cisternae of rough endoplasmic reticulum are also present throughout these cells. Throughout its length the hindgut epithelium is enclosed by a thin bad lamina and bands of well- developed circular muscles which have interconnecting longitudinal fibers. Discussion
The morphology i-f the mouth cone and the mode of feeding has been previously described for adult L. s.zlmonis (cf. Kabata, 1974b; 1979) and adults ~f the closely allied species L. dissimulaius, Lepeophrheirus pectoralis and Ca1igu.s curtus (cf. Lewis, 1961; Parker er al., 1968; Ebxshall, 1986; 1990).
The position and the gross morphology of the oesophagus and its associated musculature described here for L. sdmonis is the same as that described for L. pectoralis (cf. Boxshall, 1986; 1990). The circular and longitudinal muscles that surround the oesophagus enable the movezxnt of food by peristalsis. The nerve cord which runs between these muscles and the oesophageal epithelium is most likely responsible for their control.
The cuticle lining the oesophagus of L. salmonis is fibrillar, with numerous light bodies present in the p2 layer adjacent to the oesophagus lumen. This cuticle lacks the distinct multilayered epicuticle and a laminate procuticle of the external cuticle seen in L. salmonis and the closely related species Caligus suvulu (cf.
Kannupandi, 1976). The homologies of layers between the oesophagus and external cuticle used in this thesis are uncertain. The ultrastructure of the oesophageal cuticle has received little attention in other species of copepods. From the micrograph given by Sullivan and Bisalputra (1 980) the oesophageal cuticle of
Trigriopus californicus appears to consist of two layers, an electron-dense layer adjacent to the oesophagus lumen and a thicker less electron-dense layer beneath it. Briggs (1977) reports that the cuticle lining the foregut of Purunthessius - 112 - anemoniae consists of three layers (epicutick, exocuticle, and endocuticle) and differs from the external cutkle in that it is much thinner and has a negative reaction for calcium.
The presence of light bodies in the p2 layer of the anterior oesophagus in
L. salmonis suggests that either uptake from, or passage of materials into, the lumen of the oesophagus occurs across the cuticle. Similar structures called dense bodies have been reported in the external cuticle of Gonophysemu gullmarensis, which is a strongly transformed endoparasite of ascidians (Bresciani, 1986). The function of dense bodies is unknown. Studies involving histochemical or tracer techniques are required before the function of the light bodies in the oesophagus cuticle of L. salmonis can be determined.
The epithelial cells of the oesophagus are similar to those reported in T. culifornicus (cf. Sullivan and Bisalputra, 1980). The presence of relatively large mitochondria and numerous, irregularly shaped, endoplasmic vesicles supports the view that active secretion or uptake occurs across the oesophagus cuticle.
The external surface of the midgut of L. salmonis is surrounded by reticular muscle fibers, which are responsible for peristalsis. Loizzi (1971) reports similar muscles encasing each hepatopancreas tubule in the crayfishes Orconectes virilis and Procambarus clarkii. In these crayfishes single myofibrils within the circular muscle branch to form the fibrils of the longitudinal fibers. This structure allows the simultaneous contraction of both fibers increasing the efficiency of movement of materials through the hepatopancreas tubules. The ultrastructure of - 113 - the muscle reticulum may be similar in L. salmonis, but further study is required to confirm this.
In L. salmonis the cells of the rnidgut apparently undergo continual replacement, with mature cells being lost by degeneration or holocrine secretion.
At the light microscopic level cell cycles have been described for the midgut cells of the parasitic copegods Peroderma cylindricurn (cf. Monterosso 1930) and
Ergasilus sieboldi (cf. Einszporn , 1965).
In the present study five cell types have been described from the rnidgut of
L. salmonis. Type E cells are believed to be responsible for replacing mature cells that are lost by degeneration or holocrine secretion. The simple organization of type E cells is similar to undifferentiated cells reported from the midgut sf the copepod T. califomicus (cf. Sullivan and Bisalputra, 1980), the barnacles Balanus. balanoides and Balanus hameri (cf. Rainbow and Walker, 1977), and other higher
Crustacea and insects (reviewed in Sulvian and Bisalputrd, 1980). Type E cells appear to be a definite precursor of type A and C cells in L. salmonis. Whether they also serve as a percussor of type B and D cells is unknown.
The type A cells of L. salmonis appear to be involved in absorption and possibly some intracellular digestion. The presence of long microvilli and abundant mitochondria at the cell apex supports the view that these cells are absorptive. The presence of numerous lysosome-like dense bodies suggests that some intracellular digestion may occur, the products of which may accumulate in the vacuoles surrounding the nucleus. These products would be released into the - 114 - midgut lumen upon degeneration of the cell to be taken up by other cell types.
The lack of rough endoplasmic reticulum supports the view that enzyme
produetion and secretion is not a function of this cell type. Cells of this type have
not been previously reported in parasitic copepods, free-living copepods, or other
crustaceans.
The type B cells of L. sadmomis appear to conduct both intracellular
digestion and enzyme synthesis. In these cells intense pinocytotic activity results in
the formation of pinocytotic vesicles which appear not to contain any particles.
This suggests that the absorbed materid has already undergone some digestion
within the lumen of the midgut. These pinocytotic vesicles apparently fuse to form
light vacuoles that are often seen in close association with Golgi bodies in the sub- apical complex. The vacuoles which contain both electron-dense and electron-light
materials may represent an intermediate stage between the light and dark vacuoles. The abundant rough endoplasmic reticulum may synthesize digestive enzymes which accumulate at first in the reticular cisternae and later in the elec tron-den se vacuoles.
Cells with the same structure of microvilli, the same intense pinocytotic activity, and similar electron-dense vacuoles to those seen in the type B cells of L. salmonis, have been reported in C. minimus (referred to as type B cells) (Poquet,
1980), as well as in C. quim, 0.koe, P. yamagutii, N. japonicus, and L. cyprinacea (referred to as vacuolar cells) (Yoshikoshi and K6, 1991a). These cells are reported to have a dual function of both absorption and secretion of digestive - 115 - enzymes. Yoshikoshi and K6 (199 1b) have demonstrated by histochemical
techniques that the electron-dense vacuoles in these later species contain lysosomal
enzymes.
At the ultrastructural level, type B cells of L. salmonis appear to be
analogous to the type C cells of L. spratrae (cf. El Gharbi, 1984), the type B cells
of the crayfish hepatopancreas (Loizzi, 1971), the type B cells of calanoid
copepods (Amaud et of., 1978; 1980), and the ameboid cells of the harpacticoid
copepod P. anemoniae (cf. Briggs, 1977). With the exception of the ameboid cells
of P. anemoniae these cells are reported to carry out intracellular digestion,
enzyme synthesis, and secretion. Briggs (1977) reports the ameboid cells of P.
anemoniae to be excretory with the large electron-dense central vacuoles
containing indigestible material that is voided into the gut lumen and forms the
fecal pellet.
At the light microscope level the type B cells of L. su!monis appear to be
analogous to the type B cells of L. dissimulutus and E. sieboldi, which are reported to be secretory (Lewis, 1961; Einszporn, 1965), and to the vacuolar cells of Allantogymus sp. which are reported to be excretory (Changeaux, 1960).
The type C cells of L. salmonis appear to conduct both absorption, and in
the case of blood-feeding animals, both storage and metabolism of lipids. The long
microvilli which contain intrarnicrovillar filaments and the presence of large
numbers of mitochondria at the cell apex support an absorptive role for this cell - 116 - type. The abundant rough endoplasmic reticulum which contacts the lipid deposits
is most likely responsible for the synthesis of proteins for export.
Cells that are morphologically almost identical to the type C cells of L.
salmonis have ken reported in C. minimus (referred to as type A cells) (Poquet,
1980), as well as in C. quintus, 0. koe, P. yamagutii, N. japonicus, and L.
cyprinacea (referred to as non-vacuolar cells) (Yoshikoshi and K6, 199 la). Similar
features of these cells include their long microvilli, numerous mitochondria at
their apices, abundant rough endoplasmic reticulum, and distinct lipid deposits.
These cells are reported to have a dual function of both absorption and lipid
storage.
At the ultrastructural level, type C cells of L. salmonis appear to be analogous to the type R cells of the crayfish hepatopancreas (Loizzi, 1971), the cell type three of the harpacticoid copepod T. califsrnicus (cf. Sullivan and
Bisalputra, 198Q),and the columnar cells of P. anemniae (cf. Briggs, 1977).
These cells are reported to have a dual function of both absorption and lipids andlor glycogen storage.
The type C cells of L. salmonis are most likely analogous to the absorptive
R-cells of free-living calanoid copepods (Amaud et al., 1978; 1980). They differ from R-cells in that they have lipid deposits and abundant rough endoplasmic reticulum. In calanoid copepods the sites of storage, synthesis, and catabolism of lipids occur in the midgut mesentery or walls of the lipid sac (see Blades-
Eckelbarger, 1991) and not in the cells of the midgut as in L. salmonis. - 117 -
At the light microscope level the type C cells of mucus-feeding L. sttimonis appear analogous to the type A cells of L. dissirnulurus (cf. Lewis, 1961).
Lepeophtheinzs dissimulatus is reported to feed exclusively on mucous.
As evidenced by the long microvilli, abundant mitochondria, abundant endoplasmic reticulum, and the well-developed infoldings of the basal cell membrane the function of type D cells is most likely that of absorption. Similar celis have not knpreviously reported in parasitic copepods. These cells may be analogous to the D-cells of free-living calanoid copepods for which no function has been attributed (Anaud er al., 1978; 1988). They may also be :i~aalogousto the type two cells of T. cu1~~rnicu.swhich are considered to be absorptive
(Sullivar and Bisalputra, 1980).
Hindgut
The gross morphology of the hindgut and its associated musculature is the same as that described for the closely related species, L. pectoralis (cf. Boxshall,
1990). The dilator muscles serve to dilate the lumen of the hindgut allowing the passage of fecal material. These muscles also enable L process referred to as "anal respiration" or "and drinking". In this process water is taken up through the anus and forced by reverse peristalsis into the posterior midgut, where it is believed that gaseous exchange andlor osmoregulation occurs (reviewed in Giinzl , 199 1).
The circular and longitudinal muscles that surround the hindgut most likely enable the expulsion of fecal material and water taken up during anal respiration. - 118 - The hindgut of L. salmonis may function in crsrnoregulation, excretion,
and/or element storage. The relatively thin cuticle lining of the anterior regions ~f
the hindgut, the presence of light bodies and microtubules within the procuticle,
the folding of the basal cell membranes, and the abundance of mitochondria
suggest an osmoregulatsry function. Lepeaphtheirus salrnnnis shows hyperosmotic
regulation when expod to lower salinity sea waters (Hahnenkamp and Fyhn,
1985). The exchange of seawater through the hindgut during anal respiration may
provide the ions necessary for the maintenance of a hyperosmotic state.
With exception of T. caiifornicus (cf. Scllivan and Bisalputra, 19803,
Eudiaptomus gracilis (cf. Musko, 1988), and P. anemom (cf. Briggs, 1977), which lacks a cuticle-lined hindgut, the ultrastructure of the hindgut in copepods is unknown. The hindguts of both T. califomicus and E. gracilis, which are suggested to function in rismoregulation, have several characteristics in common with the hindgut of L. salmonis. These include: a thin cuticle with invaginations penetrating the epithelial cells, abundant mitochondria, and an invaginated basal plasma membranes. Such characteristics are also common in other crustacean and insect cells involved in ssmoregulatisn (reviewed in Giinzl, 1991).
The mineral concretions seen in the cells of the anterior hindgut may be elements inactivated and stored as wastes, or elements stored awaiting future metabolism. Similar mineral concretions to those seen in the anterior hindgut of L. salmonis have been reported in cells of the hindgut epithelium of P, anemoniae
(cf. Briggs 1977) and E. gracilis (cf. Musko, 198&),in cells of the posterior - 119 - midgut of C. minimus (referred to as type C celIs), and in cells of the midgut epithelium of free-living cyclopoid and calanoid copepods (Duforth, 1980; Amaud et al., 1980). These concretions are believed to be either excretory praducts
(Briggs, 1977) or storage sites of elements such as calcium (Poquet, 1980; 1986).
In summary, the alimentary tract of t. salmonis consists of four parts: a mouth cone, a cutic1.e-lined oesophagus or foregut, a narrow tubular midgut, and a short cuticle-lined hindgut. The structure of the cuticle lining both the oesophagus and hindgut differs markedly from that of the external cuticle. The presence of light bodies within the oesophageal cuticle and light bodies and microtubules within the hindgut cuticle supports the view that active uptake or secretion of materials occurs across these cuticles. Abundant mitochondria and endoplasmic vesicles within the cells of both the oesophagus and hindgut, and marked infolding of the bad cell membranes of the hindgut further support this view.
The midgut of L. salmonis lacks distinct cell zones as seen in free-living copepds. Cells of the midgut appear to undergo distinct cell cycles with mature cells being extruded into the midgut lumen. Five cell types are present in the midgut. The most abundant of these cell types are: E cells (embryonic cells) which replace cells lost through degeneration or holocrine secretion, B cells
(vacuolar cells) which produce digestive enzymes, and C cells which function in absorption, lipid storage, and lipid metabolism. FIGS. Dl to D3. Lepeophtheim salmonis, oesophagus and associated
tissues. Fig. D I. Female, mouth cone, oesophagus, and anterior midgut, median
sagittal section. Scale bar = 200 pm. Fig. D2. Male, oesophagus anterior to the nerve ring, transverse section. Scale bar = 80 pm. Fig. D3. Female, oesophagus junction with the midgut, median sagittal section. Scale bar = 50 pm. (amc, anterior rnidgut caecum; bd, buccal depressor muscles; bl, buccal levator muscles; cb, cerebrum; cm, circular muscle; lab, labrum; labm, labrum muscles; lam, labium; mg, midgut; nc, nerve cord; oe, oesophagus; oec, oesophageal cuticle; oev, oesophageal valve; sg, suboesophageal ganglion).
FIGS. D4 to D7. Lepeophbheirus salmonis, mouth cone and surfaces of the midgut. Fig. D4. Female, mouth cone, ventral view. Scale bar = 60 pm. Fig.
D5. Female, mouth cone tip, median sagittal view. Note the blade of the mandible resting in the labial groove. Scale bar = 25 ym. Fig. D6. Female, external surface of the anterior midgut, lateral view. Note the connective tissue supports and gut musculature. Scale bar = 10 pm. Fig. D7. Female, inner surface of the anterior midgut, lateral view. Note the bulbous topography of the midgut wall and the dense microvilli. Scale bar = 5 pm. @c, buccal cavity; cm, circular muscle; ct, connective tissue; lab, labrum; lam, labium; lg, labial groove; Im, longitudinal muscle; md, mandible; mm, marginal membrane; s, strigil).
FIGS. D8 to 010. Female Lepeop?&irus salmonis, oesophagus structure.
Fig. D8. The oesophageaf cuticle and epithelium, just posterior to the nerve ring, transverse section. Note the light bodies distributed throughout the outer procuticle, and the mitachondria and irregdarly shaped endoplasmic vesicles within the cytoplasm. The epicuticle is not shown at this magnification. Mag.
68200 X. Fig. D9. Oesophageal cuticle, higher magnification of Fig D8, transverse section. Note the thin epicuticle and the concentration of light bodies in he outer procuticle. Mag. 75000 X. Fig. D10. OesophageA cuticle and epithelium near the junction with the midgut, median sagittal section. Note the lack of light bodies in the outer procutick and the presence of cell organelles within the lumen of the amphagus. Mag. 10880 X. (ep, epicuticle; ev, end~plasmicvesicle; lb, light bodies; m, mitochondrion; n, nucleus; oel, oesophagus lumen; rer, rough endoplasmic reticulum; p' , inner procu ticle; p2, outer procuticle).
FIGS. Dl 1 to D14. Lepeophtheinrs salmonis, light micrographs of the midgut. Fig. Dl 1. MJe, midgut near the junction with the oesophagus, transverse section. Scale bar = 50 pm. Fig. D12. Female, anterior midgut, transverse seetion of osmium fixed material. Note the cluster of type E cells (arrow) at the base of the gut wall ingrowth. Scale bar = 10 pm. Fig. D13. Female, anterior midgut, transverse section of osmium fured material. Note the large ingrowths of the midgut wall and the presence of several type B cells (arrows). Scale bar = 10 pm. Fig. D14. Female, posterior midgut, transverse section of osmium fixed material. Note the large lipid deposits (mow)within the type C cells. Scale bar
= 10 pm. (amc, anterior midgut caecum; amd, anterior midgut diverticulum; ct, connective tissue; oe, oesophagus; sg, suboesophageal ganglion; t, testis).
FIGS. D 15 to D 17. Female Lepeophtheinrs salmonis, embryonic (type E cet Is) and early type C cells of a mucus-feeding copepod, transverse sections. Fig.
D15. Two type E cells (arrows) at the base of the gut wall ingrowth. Note that these cells do not contact the basal lamina or midgut lumen. Mag. 7910 X. Fig.
D 16, Higher magnification of the apical region of a type E cell. Note the scarcity of organelles when compared to the surrounding type C cells. Mag. 12190 X. Fig.
D17. Fxly type C cell near the base of a gut wall ingrowth. Note the abundant mitochondria, the rough endqlasmic reticulum, and the region of microtubules surrounding the nucleus. Mag. 9130 X. (bl, basal lamina; m, mitochondrion; mt, microtubules; inv, microvilli; n, nucleus; rer, rough endoplasmic reticulum).
FIG. D 18. Lepeophtheirus sulmonis female, anterior midgut of a mucus- feeding copepod, median sagittal section. Note the three different cell types, an early stage type A cell (A), an early stage type B cell (B), and a type D cell (D).
All, cells are attached to a thick basal lamina which has numerous pores. Mag.
7260 X. (bl, basal lamina; lv, light vacuole; mv, microvilli; n, nucleus; rer, rough endoplasmic reticulum).
FIGS. Dl9 to D21. Lepeophttheiw salmonis female, type A cells from the anterior midgut of a mucus-feeding copepod. Fig. D19. Late stage type A cell at the apex of a gut wall ingrowth, transverse section. Note the long microvilli, the variably shaped vacuoles surrounding the nucleus and its associated microtubules, and the abundant mitochondria. Mag. 7850 X. Fig. D20. Apical portion of a late stage type A cell, transverse section. Note the abr**t&ritmitochondria, the lysosome-like dense bodies, the occasional phagaomes, and the numerous vesicles
(dark arrow) that appear to coalesce to form the large vacuoles that surround the nucleus. Mag. 32160 X. Fig, D2 i . Higher magnification of the large vacuoles and region of microtubules that surrounds the nucleus, trans~~ersesection. Mag. 30520
X. (db, dense bodies; m, mitochondrion; mt, microtubules; mv, microvilli; n, nucleus; ph, phagosome)
FIGS. D22 to D24. Lepeophtheinar salmonis female, type B and type D cells from the anterior midgut. Fig. D22. Apical region of a middle to late stage type B cell, transverse section. Note the short microvilli, the intense pinocytotic activity, the small pinocytotic vesicles (thin mow) fusing to form light vacuoles, and the irregularly shaped dense vacuoles which occupy a large portion of the cells volume. Mag. 13500 X. Fig. 023. Basal region of a middle to late stage type B cell, transverse section. Note the scattered mitochondria and Golgi bodies amongst the dense vacuoles and the distended cisternae of endoplasmic reticulum surrounding the nucleus. Mag. 13500 X. Fig. D24. Comparison of the apical region of a type A and type D cell, median sagittal section. Note the dense cytoplasm and higher numbers of organelles within the type D cell. Mag. 32100
X. (dv, dense vacuoles; gb, Golgi body; lv, light vacuoles; m, mitochondrion; mv, microvilli; n, nucleus; rer, rough endoplasmic reticulum).
FIGS. D25 to D27. Lepeophtheiw salmonis female, type C cells from the posterior rnidgut of a mucus-feeding copepod. Fig. D25. Middle to late stage type
C cell, transverse section. Note the irregularly shaped light vacuoles, the presence of electron-light material within intercellular spaces (arrow), and the dense region of microtubules surrounding the nucleus. Mag. 8540 X. Fig. D26. Later stage type C cell at the crest of a gut ingrowths, transverse section. Note the presence of mitochondria and other cellular debris within the midgut lumen (arrow). Mag.
7060 X. Fig. D27. Higher magnification of the apical region of an early to middle stage type C cell, transverse section. Mag. 2 1820 X. (lv, light vacuoles; m, mitochondrion; mt, microtubules; mv, microvilli; n, nucleus; rer, rough endoplasmic reticulum).
FIGS. D28 to D30. Lepeophtheirus salmonis female, anterior midgut of a blood-feeding copepod. Fig. D28. Apical region of the gut wall, transverse section. Note tne cellular debris within the lumen of the midgut (arrow). Mag.
6020 X. Fig. D29. Apical region of a middle stage type C cell. Note the presence of mitochondria, cisternae of rough endoplasmic reticulum, and the nucleus of a type C cell (arrow) in the lumen of the midgut. Mag. 9790 X. Fig. D30.
Degenerating nucleus of a late stage type C cell. Note the lipid deposit within the whorl of rough endoplasmic reticulum. Mag. 15670 X. (Id, lipid deposit; m, mitochondrion; mt, microtubules; mv, microvilli; n, nucleus; rer, rough endoplasmic reticulum).
FIG. D31. Lepeophtheirus salmonis female, middle stage type C cells from the posterior midgut of a blood-feeding copepod, transverse section. Note the large electron-dense lipid deposit, the irregularly shaped electron-light vacuoles, the abundant cisternae of rough endoplasmic reticulum, and the abundant mitochondria at the cell apices. Mag. 7260 X. (db, dense body; Id, lipid droplet; lv, light vacuole; m, mitochondrion; mt, microtubules; mv, microvilli; n, nucleus; rer, rough endoplasmic reticulum).
FIGS. D32 to D34. Lepeophtheirus salmonis female, late stage type C cells from the posterior midgut of a blood-feeding copepod. Fig. D32. Apex of a late stage type C cell, transverse section. Note the large electron-dense lipid deposit with cisternae of rough endoplasmic reticulum contacting its surface, the presence of electron-dense material (arrow) within the cisternae of the endoplasmic reticulum, the abundant mitochondria, and the scattered dense bodies. Mag. 6920
X. Fig. D33. Higher magnification of the subapical region, transverse section.
Note the Golgi body giving rise to small vesicles (arrow) which may be the precursors to the larger dense bodies. Mag. 18300 X. Fig. D34. Whorls of rough endoplasmic reticulum associated with the nucleus, transverse section. Mag. 8840
X.(db, dense bodies; gb, Golgi body; Id, lipid deposit; rn, mitochondrion; mt, microtubules; mv, microvilli; n, nucleus; rer, rough endoplasmic reticulum).
FIGS. D35 to D37. Lepeophtheim salmonis femalc, hindgut. Fig. D35.
Anterior hindgut, median sagittal section. Note the two cell types with different electron densities. Mag. 5850 X. Fig. D36. Apical region of an electron-light cell of the anterior hindgut, median sagittal section. Note the thin cuticle, the numerous mitochondria, and the numerous short cisternae of rough endoplasmic reticulum. Mag. 10110 X. Fig. D37. Apical region of an epithelial cell from the midregion of the hindgut. Note the electron-dense regions of microtubules within the procuticle, the short subcuticular microvilli, the abundant mitochondria, and cisternae of rough endoplasmic reticulum. The epicuticle and light bodies are not visible at this magnification. Mag. 9600 X. (c, cuticle; m, mitochondrion; mt, microtubules; n, nucleus; p, procuticle; rer, rough endoplasmic reticulum; smv, subcuticular microvilli) .
FIG. D38. Lepeophtheincs salmonis female, anterior hindgut, higher magnification of the subapical region of the epithelium, transverse section. Note the numerous mineral concretions which occur most commonly in the electron- dense cells. Mag. 14830 X. (ev, endoplasmic vesicle; gb, Golgi body; m, mitochondrion; mc, mineral concretions; n, nucleus; rer, rough endoplasmic reticulum).
CHAPTER FIVE
Comparative susceptibility, resistance, and histopathology of the response of naive Atlantic, chinook, and coho salmon to experimental infection with Lepeophtheirus salmonis (Copepoda:
Caiigidae).
Introduction
Legeophtheinrs salmonis has been reported from 10 species of salmonids including: Oncorhynchus clarki (= Salmo clarki) (coastal cutthroat trout),
Oncorhynchus gorbuscha (pink salmon), Oncorhynchus ketu (chum salmon),
Oncorhynchus kisutch (coho salmon), Oncorhynchus masou (cherry or masu salmon), Oncorhynchus mykiss ( = Salmo gairdneri) (rainbow or steel head trout),
Oncorhynchus nerka (sockeye salmon), Oncorhynchus tschuwytscha (chinook salmon), Salvelinus fontinalis (brook trout), and Salmo salar (Atlantic salmon)
(Chapter 1).
Differences in the prevalence and abundance of L. scclrnonLs among salmon species have been reported. Nagasawa (1987) reports for wild salmonids that chinook salmon are the most heavily infected species followed in descending order by steelhead, pink salmon, chum salmon, coho salmon, and sockeye salmon. In
British Columbia, sea-farmed Atlantic salmon are generally more heavily infected - 135 - with L. sulmonis than chinook or coho salmon raised at the same site (unpublished
observations). The higher levels of infection on Atlantic salmon are believed to be
due to their slower swimming speed when compared to both chinook and coho
salmon. To date no experimental work has been conducted to determine other
factors that may explain this difference.
This chapter investigates differences in the susceptibility and resistance of
naive Atlantic, chinook, and coho salmon to experimental infection with L. salmonis. Differences in the histopathology of L. salmonis attachment and feeding
sites between these species are described. Materials and Methods
Ovigerous L. salmonis were collected from sea-farmed Atlantic salmon
(Saltno salar) from Departure Bay on the east coast of Vancouver Island, Canada.
The eggs were hatched and the larvae reared to the infectious copepodid stage at 9 to lo0 C following the methods outlined in Chapter 3.
Thirty-six each of naive Atlantic, chinook, and coho salmon which ranged in size from 15.2 to 22.6 centimeters in length were introduced into a 500 liter tank, acclimated for 1 week, and then exposed for 24 hours to approximately 2000 newly molted copepodid larvae. The infection was carried out under conditions of darkness, low water flow, and aeration. A large surface area screen with 180 pm mesh size was used to prevent copepodid loss during the infection. After exposure the screen was removed and the water flow increased. The fish were maintained in flowing seawater with a temperature of 9.3 to 10.2 OC (mean = 9.6 OC) and ambient salinity 29 to 31 %O . Five of each fish species were killed at 1, 3, 5, 10,
15, and 20 days post-infection with an overdose of the anesthetic MS-222 (tricaine methanesulfonate). The fork length and wet weight was determined for each fish.
Both the anesthetic bath and the body surfaces were examined for copepods and the distribution of the copepods on the fish was noted. The number of copepods present was corrected to a standard wet body weight to compensate for differences in size between hosts.
Intensity data was log (X+ 1) transformed and differences in copepod intensity investigated by analysis of variance (ANOVA) procedures. Multiple - 137 - comparisons of copepod intensity for each host species over time, and between host species at each sampling period were made using Scheffes tests (Zar, 1984).
Tissues for examination by light microscopy were fixed in Davidson's solution and dehydrated through to 100% alcohol. Tissues were either wax- embedded, cut to a thickness of 5 pm, and stained with hematoxlylin and wsin, or they were embedded in JB4 plastic resin, cut to a thickness of 1 to 2 pm, and stained with Lee's stain (methylene blue and basic fuschin). Result•˜
Intensity of Infection
The intensity of L. salmonis on naive Atlantic, coho, and chinook salmon over time is presented in Fig. El. The intensity of infection for each host species was significantly different over time (1-way ANOVA; Atlantic salnon (P < 0.0 1); coho salmon (P < 0.001); chinook salmon (P < 0.01)). The results of multiple range tests (Scheffks test; P < 0.05) over time showed both Atlantic and chinook salmon to have significantly fewer copepods at 10 and 20 days post-infection when compared to 1 day post-infection, and coho salmon to have significantly fewer copepods at 15 and 20 days post-infection when compared to 1, 3, 5, and 10 days post-infection.
There was no significant difference in copepod intensity between host species at 1, 3, and 10 days post-infection (Scheffes test; P-CO.05). At 5 days post-infection there were significantly fewer copepods present on coho salmon than on Atlantic salmon, and no significant difference in copepod intensity between Atlantic and chinook salmon (Scheffks test; P < 0.05). At 15 and 20 days post-infection there were significantly fewer copepods present on coho salmon when compared to both Atlantic and chinook salmon, and no significant difference in copepod intensity between Atlantic and chinook salmon (Scheffes test;
P < 0.05). Distribution
At 1 day post-infection approximately equal percentages of the copepods
recovered from coho were from the anesthetic bath and the fins (Fig. E2). The
percentage of copepods found in the anesthetic bath decrease from approximately
38 % to 0% by 15 days post-infection. Of the copepods identified on the body, the
highest percentage was on the fins at each sampling time. The percentage of copepods on the gills decreased from approximately 22% to 0% by 10 days post-infection. A low percentage of copepods was found on the general body surfaces, including surfaces of the buccal cavity and mouth, at both 1 and 3 days post-infection.
At 1 day post-infection the highest percentage of copepods recovered from
Atlantic salmon was from the anesthetic bath (Fig. E3). This percentage had declined to 0% by 15 days post-infection. Of the copepods identified on the body, the highest percentage was on the gills at 1, 3, and 5 days post-infection, and on the fins at 10, 15, and 20 days post-infection. The percentage of copepods on the body was low throughout the experiment increasing slightly at 20 days post-infection with molting to the preadult stage.
At 1 day post-infection the highest percentage of copepods recovered from chinook salmon was from the anesthetic bath (Fig. I%). This percentage had declined to 0% by 15 days post-infection. Of the copepsds identified on the body, the highest percentage was recovered from the fins, then from the gills, and then from the general body surfaces at all sampling times. - 140 - Copepods were recovered from all gill arches of the host species. With exception of a few, all were attached to the distal half of the gill filaments, with the majority attached to the filament tips. Of the copepods recovered from the fins of coho salmon the majority were on the pectoral (33%) and pelvic (31 %) fins
(Fig. E5). Of those recovered from the fins of Atlantic salmon the majority were on the caudal (28%), pelvic (26%), and pectoral (25%) fins. With exception of the adipose fin, copepods were distributed almost equally among the fins of chinook salmon (pectoral (25 %); pelvic, anal, dorsal, and caudal (1 8- 19 %)).
Developmental Stages
The developmental stages of L. salmonis present at 20 days post-infection on both Atlantic and chinook salmon are presented in Fig. E6. Of the 47 copepods recovered from Atlantic salmon 2% were second chalimus larvae, 60% were late third chalimus larvae, 4% were fourth chalimus larvae, 23 % were first preadult males, and 1I % were first preadult females. Of the 25 copepods recovered from chinook salmon 24% were second chalimus larvae and 76% were third chaiimus larvae.
The developmental stages of L. salmonis present on the different body regions of Atlantic salmon at 20 days post-infection are presented in Fig. E7. Of the 11 cope-pods recovered from the gills 100% were late third chalimus larvae.
Of the 21 copepods recovered from the fins 5% were second chalimus larvae,
43 % were late third chalimus larvae, 10% were fourth chalimus larvae, 33 % were attached first pmdult males, and 10% were attached first preadult females. Of the - 141 - 15 copepods recovered from the body 53% were late third chalimus larvae, 27% were unattached first preadult males, 7% were attached preadult females, and 13% were unattached first preadult females.
Histology of the Gills
At 1, 3, and 5 days post-infection, attachment and feeding sites on gills of coho salmon were characterized by partial to complete erosion of the epidermis and the underlying basement membrane, minor hemorrhaging, and acute inflammation (Figs. E8 to Ell). The inflammatory infiltrate consisted primarily of neutrophils, but lymphocytes were also present. In some sections mild epidermal hyperplasia occurred at the tips of the lamellae.
At 1, 3, and 5 days post-infection, attachment and feeding sites on gills of both Atlantic and chinook salmon were characterized by variable amounts of erosion of the epidermis, small amounts of hemorrhage, and mild inflammation of the dermis (Figs. El2 to E15). The inflammatory infiltrate consisted of abundant neutrophils and a few lymphocytes. In a portion of samples collected at 5 days post-infection the cartilaginous central rods of the filaments were exposed (Fig.
E15).
Gross examination of the gills of both Atlantic and chinook salmon at 10,
15, and 20 days post-infection revealed distinct crypting (atrophy and disappearance of the distal portions of the lamellae) and fusion of the secondary gill lamellae in the vicinity of the copepods. At 10 days post-infection, histological examination of the gills of both species revealed variable amounts of epidermal - 142 - erosion, some hemorrhage, mild inflammation of the dermis, some epidermal hyperplasia, and the fusion of secondary gill lamellae (Figs. E16, El 8 and E19).
The extent of these changes as well as the number of primary lamellae affected increased in both species at 15 and 20 days post-infection (Figs. E20 to E23).
From 10 days post-infection onwards there was a proliferation of goblet cells within the hyperplastic epidermis of the chinook salmon gills (Figs. E18, El9 and
E23). No proliferation of goblet cells was evident in the hyperplastic epidermis of the Atlantic salmon gills. The inflammatory infiltrate of both species consisted primarily of neutrophils, but a few lymphocytes were also present. In general the magnitude of the response of the gills to the presence of L. salmonis was greater in chinook than Atlantic salmon. In both species secondary infection of the gill lesions by rod-shaped and filamentous bacteria occurred at 15 and 20 days post-infection.
Histology of the Fins
At 1, 3, and 5 days post-infection, attachment and feeding sites on fins of all three host species were characterized by partial to complete erosion of the epidermis (Figs. E24 to E27). Over this period the severity of the lesions was highly variable both between and within host species, with later lesions not necessarily more severe than earlier lesions. Mild inflammation of the dermis occurred in coho salmon as early as 1 day post-infection (Figs. E24 and E25).
Neutrophils were the predominant cells at these sites of inflammation, but - 143 - lymphocytes were also present, No inflammatory responses were observed in
either the Atlantic or chinook salmon over the same period (Figs. E26 and E27).
At 10, 15, and 20 days post-infection, attachment and feeding sites on fins
of coho salmon where characterized by well-developed epithelial hyperplasias
(Figs. E28 and E30), which in severe cases resulted in complete encapsulation of
the copepods. In cases of partial or complete encapsulation the spaces surrounding
the copepod were filled with tissue debris and a mixed inflammatory infiltrate
(neutrophils, macrophages, and a few lymphocytes) (Figs. E28 and E29). Lesions
at the point of feeding commonly extend through to the dermis exposing the fin
rays. Necrotic tissue, some hemorrhage, and well-developed inflammation of the dermis occurred at these sites, The inflammatory infiltrate consisted of abundant neutrophils, some macrophages, and a few lymphocytes (Figs. E29 and E3 1).
At 10, 15, and 20 days post-infection, attachment and feeding sites on fins of both Atlantic and chinook salmon showed little tissue response to the presence of the copepods (Figs. E32 to E37). At the point of feeding the epidermis was commonly breached and the underlying dermis and fin rays exposed to the external environment. In some sections mild inflammation of the dermis was evident. The inflammatory infiltrate consisted of abundant neutrophils and a few lymphocytes (Fig. E35). In both species, secondary infection of the fin lesions by rod-shaped and filamentous bacteria occurred in some of the samples collected at
15 and 20 days post-infection.
Histology of the Frontal Filament. - 144 - Chalimus larvae and a portion of first preadult males and females are attached to their hosts by frontal filaments. The frontal filament consists of an elongate stem and a basal plate (Fig. E38). In section the stem appears to consist of two regions. The outer region has similar staining characteristics to that of the body cuticle and appears to be continuous with it. The inner region appears fibrous and has a duct-like structure (axial duct) running along its length (see also
Chapter 2). The basal plate stains darker than both layers of the stem. Basal plates are most commonly attached to the cartilaginous central rods of the primary gill lamellae (Figs. E16, El7 and E20) or to the fin rays (Figs. E32 and E38). Less commonly, basal plates are attached to the basement membranes of the gills and fins.
Within the anterior cephalothorax of late copepodid and chalimus larvae, materials with similar staining characteristics to those of the frontal filaments were commonly observed (Figs. E27, E34, E36). Fully formed frontal filaments (stem and basal plates) were found in the anterior cephalothorax of two chalimus larvae that were in the process of molting (Figs. E39 and E40). These filaments had similar staining characteristics to attached filaments, but differed in the structure of their stems. In both instances the inner regior, of the stems appeared to consist of distinct fibrous bands interspersed with a small amounts of living tissue (Fig.
E4O). The invaginations of the anterior cephalothorax which surrounded the new filaments were lined with newly formed external cuticle, which appeared to be - 145 - continuous with the stems (Fig. E40). Old filaments were attached to and are apparently lost with the molted exoskeletons. - 146 - Discussion
Significant reductions in the intensity of L. salmonis occurred on all three
host species over time. These reductions may be caused by active host rejection and/or natural mortality of the copepis independent of any host response. Host rejection of the cyclopoid copepods Lernaea qpriraacea and Lernaea pslymclrpha has been previously reported in both naive and previously exposed fish (Shields and Goode, 1978; Shariff and Roberts, 1989; Woo and Shariff, 1990). Rejection of these copepods is believed to be due in part to cellular responses and/or possible physical removal by the fish rubbing their bodies against the tank.
There was no significant difference in the intensity of L. sal~misbetween host species early in the experiment. This suggests that all three species are equally susceptible to infection with L. salmonis. Coho salmon appear to be the most resistant spies, having significantly fewer copepods then brrrh chincmk or
Atlantic salmon at 15 and 20 days post-infection. Coho salmon have been shown to be more resistant than both Atlantic and chinook salman when both experimentally and naturally exposed to glochidia of the freshwater mussel
M~rgan'tiferamagaritifera (Myers and Millemann, 1977; Karna and Millemann,
1978).
The intensity data suggest that both chinook and Atlantic salmon have a similar resistance to L. salmonis infection. However, at 20 days post-infection there was a marked difference in the age structure of L. sulrncmis between the chinook and Atlantic salmon, which suggests that copepods develop at a slower - 147 - rate on chinook salmon. By the time that an equal age distribution to that seen on
the Atlantic salmon is attained on the chinook salmon the intensity of E. salmonis
might be significantly lower. Mortality of L. salmonis or loss from their hosts
may be substantial duri~gmolting events. Reduced copepod development rates on chinook salmon are in themselves an indicator of greater host resistance to L. salmonis infection.
This author knows of no reports of different development rates for a parasite on different animal host species. Differences in the development rates of
L. salmonis between chinook and Atlantic salmon may be caused by nutritional factors and/or non-specific host defence mechanisms. Fish may produce humoral factors such as growth inhibitors, enzyme inhibitors, and/or substances that interfere with the feeding activities of L. salmonis.
Host effects on biology of ectoparasites have been previously documented, but the mechanisms are poorly understood. Shariff (1981) suggests that a change in the distribution of Lerneae piscinae growing on big head carp, Aristitchthys noblis, from the general body to the cornea is a reaction to the development of an immune response within the body of the host. The cornea being an avascular site having a lower level of immune response than the body. Paperna ;ind Zwerner
(1982) report for Ergmilus labracis growing on stripped bass, Morone saatilis, that a well-developed tissue response leads ts the interruption of parasite egg sac production and an apparent increase in the rate of detachment of the copepods.
Woo and Shariff (1990) report for Lemea cypriwea growing on kissing - 148 - gourami, Helostoma temmincki, that a higher proportion of egg sacs are lost from copepods growing on previously exposed fish than naive fish. Furthermore, eggs from copepods growing on previously exposed hosts either fail to develop or produce copepodids that have a low infectivity when compared to copepodids hatched from eggs of copepods growing on naive hosts. Observatiorts made during the present study revealed that L. salmonis on Atlantic salmon carries approximately twice as many eggs as those on chinook salmon raised at the same site. In addition, higher numbers of eggs are carried by L. salmonis growing on mature versus immature Atlantic salmon (unpublished observation). Gelnar (1987) reported a higher growth rate of the monogenean Gyroductylus gobienssi on gudgeon, Cobio gobio, when the fish were exposed to conditions of low dissolved oxygen and starvation.
The presence of earlier developmental stages on the gills of Atlantic salmon when compared the fins and body surfaces further supports the hypothesis that non-specific humoral factors may be affecting the development rate of L. salmonis. The gills, as they are highly vascularized in comparison to the fins and body, should have a more pronounced immune response. Blood was found in the guts of all copepods feeding on the gills, whereas blood was found less commonly in the guts of copepods feeding on the fins and general body surfaces. Differences in the development rate of &, salmonis on different body regions of Atlantic salmon may explain the large range of variability in development rates reported between individual copepods in Chapter 3. - 149 - One criticism of this study's experimental design is that the free moving
preadat: stage may have migrated from the chinook to the Atlantic salmon cwsing
the observed difference in the age class distribution. I would argue that this is
unlikely as I would expect a proportion of the preadults that matured on the
chinook to remain attached by their frontal filament. Approximately 63% of the
first preadults found on Atlantic salmon had retained their frontal filaments.
Retention of the frontal filament by the first preadult stage is commonly reported
in many species of caligid copepods from a wide variety of hosts (Chapter 2).
Early in the experiment high percentages of the copepodites on all host
species became detached and were found swimming in the anesthetic bath. This
suggests that settlement may be reversible up to the first chalimus molt and that
the copepodites may change position on the host. This ability to change position
may explain the wide variation in the severity of lesions caused by the copepodid
stage in this study. Migration after settlement from the general body surfaces to
the fins has been reported for the copepodites of Lemaeenicacs sprattue
(Anstensrud and Schram, 1988).
In this experiment copepods were recovered from the gills of all three host
species. The presence of 6. salmonis on the gills of laboratory-infected Atlantic
salmon has been previously reported (Bron et al. 1991). Tljese mthors suggest
that copps$s may settle on the gills of tank-maintained fish due to slower current
through the bucd cavity when compared to wild or pen-reared fish. Although L. salmonis has not been reported on the gills of pen-reared salmonids, both - 150 - copepodids and chalimus stages have been found on the gilIs of mature wild sockeye salmon in British Columbia (unpublished observation).
Copepcxls were eliminated from the gills of coho salmon by 10 days post- infection, possibly by the well-developed inflammatory responses. Glochidia of the freshwater mussel M. margaritifera have been reported to be sloughed from the gills of coho salmon by 4.5 days post-infection at 12O C by well-developed epidermal hyperplasias (Fustish and Milemann , 1978).
The distribution of L. salmonis on Atlantic salmon at 10 days post-infection is similar to that reported by Bron et a1.(1991) for early chalimus stages on experimentaily infected Atlantic salmon. In their study 69% of the copepods were recovered from the fins, 21 % were recovered from the gills, and 18 % were recovered from the body. These authors suggested that distribution is principally a question of local current speed and the ability of the copepodite to hold on in any given arca. The results of the present study indicatzs that other factors such as differences between tissues in their response to L. salmonis are important in determining the distribution on the body.
The extent of tissue damage and the magnitude of the host response was highly variable on all three host species over the period of 1 to 5 days post-infection. Over this period the majority of L. salmonis were present as free- moving copepodid larvae. ,n;Iai.emat of the apepodids on the host would explain this high variability. Copepodids may change positions to locate a suitable site for frontal filament attachment, and/or to avoid host tissue reactions. - 151 - Epidermal erosion, acute inflammation (characterized by a mixed inflammatory infiltrate), mild hyperplasia, and minor hemorrhaging occurred at attachment and feeding sites on coho salmon gills. Intense reactions of coho gill tissue to the presence of parasitic organisms has been previously documented.
Coho gills exposed to the freshwater mussel M. rnargan't@era showed progressive epidermd hhyperplasia, fusion of the secondary gill Iamellae, and infiltration of eosinophilic cells over a periocE of 5 days post-infection (Fustisch and Millemann,
1978). Spores of Lorna salmonae in the vasculature of the primary lamellae of seawater reared coho salmon caused a severe inflammatory response, consisting of a mixed inflammatory infiltrate (Kent et al., 1989).
Epidermal erosion, mild inflammation, and a small amount of hemorrhage occurred at attachment and fding sites on the gills of both chinook and Atlantic salmon over the period of f to 5 days post-infection. Later samples from both species showed progressive development of epidermal hyperplasia, fusion of the secondary gill lamellae, and inflammation of the dermis. In chinook salmon there was a proliferation of mucous cells within the hyperplastic epithelium. The gill tissue responses of both chinook and Atlantic salmon are similar to those reported for other host species infected with other species of parasitic copepods. Previously re~rtedresponses include: hjperplasia resdting in the loss of lamellar structure
(fiabata, 1970; Kabata and Cousens, 1977; Papma and Zwerner, 1982), proliferation of mucous cells within the hyperplastic epidermis (Paperna and
Zwemer, 1982), hypertrophy of the epidermal cells (Kabata and Cousens, 1977), - 152 - and extensive infiltration of macrophages, lymphocytes and eosinophils into heavily infected gills (Paperna and Zwerner, 1982).
The inflammatory response of the gills of chinook salmon seen in this study is similar to the chronic inflammatory response described for the gills of mature chinook salmon infested with the fungus Democystidiwn sy. (Pauley,
1967). This author reports that the inflammatory response was characterized by infiltration of lymphocytes, monocytes, macrophages, and granulocytes.
Differences between coho and chinook salmon with respect to the intensity of their gill tissue reactions to parasitic infection have been previously reported.
Gills of chinook salmon exposed to M. margaritifera showed only mild hyperplastic and inflammatory responses, when compared to gills of coho salmon, which showed well-developed hyperplastic and inflammatory responses as well as fusion of the secondary gill lamellae (Fustisch and Millemann, 1978).
From 10 days post-infection through to the end of the experiment, attachment and feeding sites on the fins of coho salmon were characterized by extensive epiihelial hyperplasias and well-developed inflammatory responses. Over the same period, attachment and feeding sites on the fins of chinook and Atlantic salmon were characterized by extensive epithelial erosion and mild inflammatory responses. The well-developed tissue responses of coho salmon may be responsible for their greater resistance to L. salmonis.
The magnitude of the tissue response reported for fins of Atlantic salmon in the present study is similar to that reported for the general body surfaces of - 153 - Atlantic salmon infmted with L. sttlmrzis chalirnus larvae (Jones et a)., 1990).
These authors report erosion of the epidermis in the vicinity of the mouth cone, the lack of a dermal reaction, and a normal or mildly hyperplastic epithelium in the vicinity of the frontal filament. Boxshall (1977) reports on the histopathology of lesions on the fins of naturally infected flounder, Plati~hthys~flesus,caused by the closely related coppod species Lepeophtheirus pectoralis. Damage to the fins was usually confined to the epidermis and little response to the presence of the copepod was seen unless the dermis was breached. In cases where the dermis was breached, an inflammatory response, comprised sf fibroplasia and ceMar infiltration, occurred and resulted in the formation of a dense fibrous granulation tissue.
In this study the rejection of L. salmonis by naive hosts is most likely due to a non-specific immune response. However, we cannot rule out the possibility that some specific immunity may have developed towards the end of this experiment and contributed to the rejection of L. salmonis. Grayson et al. (1991) report that naturally infected Atlantic salmon mount a low-level specific antibody response to antigens asswiated with the gut epidermis sf L. salmonis.
The mechanisms by which the non-specific immune system rejects L. salmonis remain to be determined. Both normal and activated macrophages of rainbow trout have been shown to have larvicidal activity against the eye fluke
Diplustonum spathacem, but the mechanisms of this activity are unknown (Whyte et al. 1989). - 154 - In this study, neutrophils are the predominant cells at sites of inflammation
in all three host species. Macrophages were common at sites of inflammation of
the fins of coho salmon from 10 days post-infection onwards. Both neutrophils and
macrophages are the predominant cell types reported at sites of inflammation of a wide variety of both naive and previously exposed fish hosts infected with parasitic copepods (Joy and Jones, 1973; Ebxshall, 1977; Shields and Goode,
11978; Shariff, 1981; Paperna and Zwerner, 1982; Shariff and Roberts, 1989). In
this, lymphocytes were present at sites of inflammation only in very low numbers.
This suggests that cell-mediated immunity does not play a major role in the dimination of L. salmonis from these hosts.
The structure of the attached frontal filament is identical to that reported by
Bron et al. (1991) far L. salmonis. The fully formed frontal filaments sen in the anterior cephalothorax of premolt chalimus larvae have not been previously reported. Although fully formed frontal filaments have been reported in the copepodid stage of a wide variety of parasitic copepods (Wilson, 191 1; Gurney,
1934; Hwa, 1965; Kabata, 1972; Kabata, 1976), no frontal filament was reported in newly molted copepodids of L. salmonis (Chapter 2). It is possible that fully formed frontal filaments are only present in late copepodid larvae immediate]jl preceding the molt. This investigation as well as others may have missed reporting the presence of the frontal filament in the copepodid stage by describing copepodids at too early of a stage in their development. - 155 - The current belief is that L. salmonis attaches to its host using a glue-like
secretion. This glue-like secretion is thought to be injected beneath the epidermis
where it spreads out laterally dong the basement membrane to form the "basal
plate" of the frontal filament (see Bron et al. 1991). It is also reported that L. salmonis remains attached by its original frontal filament throughout its development (see Jones et (12. , 1990).
In the present study, "reservoirs of filament materialt' were commonly seen in both the copegodid and chalirnus stages of L. salmonis. These "reservoirs of filament material" have been previously reported in both the copepodid and chdimus stages of L. salmonis (Bron et al., 1991). It is the belief of this author that these "reservoirs of filament material" are in fact new frontal filaments in early stages of development. As suggested for the copepodid stage, fully formed frontal filaments may be present In chalimus larvae only immediately preceding the molt. The cuticle-lined pocket seen surrounding the filament in this study suggests that both the filament and the anterior region cephalothorax are pulled out prior to hardening of the new cuticle. Material which functions as a glue may be secreted via the axial duct and serve to attach the basal plate to the host.
In summary, naive coho, chinook, and Atlantic salmon appear to be equally susceptible to experimental infection with L. salmonis. Coho salmon appear to be the most resistant species with ail copepods lost from the gills by 20 days post-infection and only a few remaining on the fins at 20 days post-infection.
Rejection of L. salmonis on all three host species is most likely due to - 1156 - non-specific host responses. In coho salmon these responses include well- developed epithelial hyperplasias and inflammatory responses, The response of chinook salmon appears to be intermediate in magnitude between that of the coho and Atlantic salmon. Little response to the presence of L. salmonis was observed in Atlantic salmon. The development rate of L. salmonis appears to be higher on
Atlantic salmon when compared to chinook salmon. The development rate of L. salmonis on both chinook and Atlantic salmon appears to be mediated by some form of host response. The presence of fully formed frontal filaments in premolt chalimus larvae indicates that further investigation of the mechanism and timing of filament production is warranted. FIG. El. Mean (+SE) intensity of Lepeophtheirus salmonis on naive
Atlantic, coho, and chinook salmon at various times post-infection. Salmon were maintained at 9.3 to 10.2 OC and ambient salinity (29 to 31 YZ). 1 3 5 10 I5 20 DAYS PQSf -INFECTION FIG. E2. Distribution of Lepeophtheim salmonis on naive coho salmon.
Fish were maintained at 9.3 to 10.2 OC and ambient salinity (29 to 31 %I). Bath
Body
1 3 5 10 15 20 BAYS POST-INFECTION FIG. E3. Distribution of Lepeophrheirus salmonis on naive Atlantic salmon. Fish were maintained at 9.3 to 10.2 "C and ambient salinity
(29 to 3 3. %o). 1 3 5 10 15 20 DAYS POST-INFECTION FIG. E4. Distribution of lepeophtheiw salmonis on naive chinook salmon. Fish were maintained at 9.3 to 10.2 OC and ambient salinity
(29 to 31 %o). Bath
Body
Gills
1 3 5 10 15 20 DAYS POST-INFECTION FIG. E5. Distribution of Lepeophtheirz salmonis on the fms of naive
Atlantic, chinook, and coho salmon. Percentages based on total copepods collected on each host species during the study. V ATLA CHIN COHO HOST SPECIES FIG. E6. Developmental stages of Lepeophtheirms salmonis present on
naive Atlantic and chinook salmon at 20 days post-infection. Percentages based on
total copepods collected on each host species. Fish were maintained at 9.3 to 10.2
"C adambient salinity (29 to 31 %o). (Ch2, second chdimus; Ch3, third chalimus; Ch4, fourth chalimus; Prel (f) first preadult female; Prel (m), first preadult male) 60
40
20
0 Atlantic Chinook HOST SPECIES FIG. E7. Developmental stages of Lepeophtheim salmonis present on different body regions of naive Atlantic salmon at 20 days post-infection.
Percentages based on total copepods collected. Fish were maintained at 9.3 t~
10.2 "C and ambient salinity (29 to 3 1 %o). (Ch2, second chalimus; Ch3, third chalimus; Ch4, fourth chalimus; Prel (f) first preadult female; Prel (m), first preadult male) Gills Fins Body BODY LOCATION FIGS. E8 to El 1. Lepeophtheim salmonis copepodids on the gills of naive coho salmon. Fig. E8. Copepodid on a gill of a coho salmon, 1 day post-infection. Note the hemorrhage and mild inflammatory response. Scale bar =
200 pm. Fig. E9. The inflammatory response of a gill of a coho salmon to L. salmonis, 1 day post-infection. Note the presence of a mixed inflammatory infiltrate comprised mostly of neutrophils. Scale bar = 30 pm. Fig. E10.
Copepodid on a gill of a coho salmon, 5 days post-infection. Note the tip of the second antenna and the mild inflammatory response. Scale bar = 30 pm. Fig.
El 1. Copepodid on a gill of a coho salmon, 5 days post-infection. Note the erosion of the epidermis and the inflammatory response that is comprised mostly of neutrophils. Scale bar = 30 pm. (c, copepodid; cr, central rod; i, mixed inflammatory infiltrate; mc, mouth cone; sa, second antenna).
FIGS. El2 to E15. Lepeophtheim salmo~iscopepodids on the gills of naive Atlantic and chinook salmon. Fig. E12. Coppodid on a gill of an Atlantic salmon, 1 day post-infection. Note the erosion of the epidermis but otherwise little tissue response. Scale bar = 30 pm. Fig. E13. Copepodid on a gill of an Atlantic salmon, 5 days post-infection. Note the tip of the second antenna and the mild inflammatory response. Scale bar = 30 pm. Fig. E14. Copepodid on a gill of a chinook salmon, 5 days post-infection. Note the erosion of the gill tissue, the limited tissue response, and the tip of the second antenna. Scale bar = 30 pm.
Fig. E15. Copepodid on a gill of a chinook salmon, 5 days post-infection. Note the erosion of the epidermis, exposure of the central rod, and limited tissue response. Scale bar = 30 pm. (c, copepodid; cr, central rod; i, mixed inflammatory infiltrate; mc, mouth cone; sa, second antenna).
FIGS. El6 to E19. Lepeophbheim salmonis chalimus larvae on the gills of
naive Atlantic and chinook salmon. Fig. El6. Chalimus larva on a gill of an
Atlantic salmon, 10 days post-infection. Note the epidermal hyperplasia, the inflammatory response, and that the basal plate of the frontal filament is attached to the central rod. Scale bar = 50 pm. Fig. E17. The tissue response of a gill of an Atlantic salmon to the frontal filament of L. salmonis, 20 days post-infection.
Note the limited tissue response to the presence of the frontal filament, and that the basal plate of the frontal filament is attached to both the basement membrane and central rod. Scale bar = 30 pm. Fig. El 8. Chalimus larva on a gill of a chinook salmon, 10 days post-in fection. Note the epidermal hyperplasia, the increased number of goblet cells within the epidermis, and the well-developed inflammatory response. Scale bar = 50 prn. Fig. E19. The tissue response of a gill of a chinook salmon to L. salmonis, 10 days post-infection. Scale bar = 30 pm. (bm, basement membrane; ch, chalimus larva; cr, central rod; ff, frontal filament; g, goblet cell; i, mixed inflammaiory infiltrate).
FIGS. E20 to E23. Lepeophtheirus salmonis chalimus larvae on the gills of naive Atlantic and chinook salmon. Fig. E20. Chalimus larva on a gill of an
Atlantic salmon, 20 days post-infection. Note the extensive hemorrhage, the well- developed inflammatory response, and that the basal plate of the frontal filament is attached to the central rod. Scale bar = 200 pm. Fig. E21. The tissue response of a gill of an Atlantic salmon to L. salmonis, 20 days post-infection. Note the fusion of the secondary lamellae and the presence of some inflammatory cells.
Scale bar = 30 pm. Fig. E22. Feeding site of a chalimus larva on a gill of a chinook salmon, 15 days post-infection. Scale bar = 30 pm. Fig. E23. Chalimus larva on a gill of a chinook salmon, 20 days post-infection. N~tethe well- developed epidermal hyperplasia, the increased number of goblet cells within the epidermis, and the well-developed inflammatory response. Scale bar = 200 pm.
(ch, chalimus law; cr, central rod; ff, frond filament; g, goblet cell; i, mixed inflammatory infiltrate; mc. mouth cr 7e).
FIGS. E24 to E27. Lepegphtheim salmonis copepodids on the fins of naive coho, Atlantic, and chinook salmon. Fig. E24. Copepodid on a fin of a coho salmon, 1 day post-infection. Note the erosion of the epidermis and mild inflammation of the dermis (arrow). Scale bar = 30 pm. Fig. E25. Copepodid on a fin of a coho salmon, 5 days post-infection. Note the tip of the second antenna, erosion of the epidermis, and mild inflammation of the dermis (mow). Scale bar
= 30 gm. Fig. E26. Copepodid on a fin of a chinook salmon, 1 day post-infection. Note the mild erosion of the epidermis in the vicinity of the tip of the mouth cone. Scale bar = 30 pm. Fig. E27. Copepodid on a fin of an Atlantic salmon, 5 days post-infection. Note the severe erosion of the epidermis, the exposed fin ray, the lack of an inflammatory response, and the presence of a frontal filament within the anterior ceyhalothorax. Scale bar = 50 pm. (c, copepodid; ff, frontal filament; fr, fin ray; mc, mouth cone; sa, second antenna).
FIGS. E28 to E3 1. Lepeophtheirus sulmonis chalimus larvae on the fins of
coho salmon. Fig. E28. Chalimus larva on a fin of a coho salmon, 1Q days
post-infection. Note the well-developed epidermal hyperplasia that almost
completely encapsulates the copepod, the well-developed inflammatory response,
and the small amount of hemorrhage associated with the lesion. Scale bar = 200
pm. Fig. E29. The inflammatory response of a fin of a coho salmon to L.
sulmonis, 10 days post-infection. Note the presence of both neutrophils and
rnacrophages. Scale bar = 25 pm. Fig. E30. Chalimus larva on a fin of a coho
sal men, 15 days post-infection . Note the well-developed epidermal hyperplasia and
the well-developed inflammatory response within the dermis. Scale bar = 200
pm. Fig. E31. The inflammatory response of a fin of a coho salmon to a frontal
filament of L. salmonis, 15 days post-infection. Note the hyperplastic epidermis,
and the presence of a mixed inflammatory infiltrate consisting of both neutrophils and macrophages. Scale bar = 30 pm. (ch, chdimus larva; ff, frontal filament;
fr, fin ray; he, hyperplastic epidermis; i, mixed inflammatory infiltrate; mc,
mouth cone).
FIGS. E32 to E35. &epeoph&eincr salmonis chalimus larvae on the fins of naive Atlantic and chinook salmon. Fig. E32. Chalimus larva on a fin of an
Atlantic salmon, 10 days post-infection. Note the erosion of the epidermis and the lack of an inflammatory response. The basal plate of the frontal filament is resting directly on the fin ray. Scale bar = 50 pm. Fig. E33. Higher magnification of a lesion caused by L. salmonis on a fin of an Atlantic salmon, 15 days post-infection. Note that the epidermis is eroded to the basement membrane, and that there is little inflammatory response. Scale bar = 30 pm. Fig. E34. Chalimus larvae on a fin of a chinook salmon, 10 days post-infection. Note the reduced thickness of the epidermis beneath the copepod and the lack of an inflammatory response. Scale bar = 200 pm. Fig. E35. Chalimus larva on a fin of a chinook salmon, 18 days post-infection. Higher magnificaCon of the lesion shown in Fig.
E34. Lesion near the posterior of the chalimus larva. Note that the epidermis has been breached and that there is a mild inflammatory response. Scale bar = 30 pm. (bm, basement membrane; ch, chalimus larva; d, dermis; e, epidermis; ff, frontal filament; fr, fin ray; i, mixed inflammatory infiltrate; mc, mouth cone).
FIGS. E36 and E37. Lepeophtheirus salmonis chalimus larvae on fins of chinook salmon. Fig. E36. Chalimus larva on a fin of a chinook salmon, 20 days post-iizfection. Note the severe erosion of the epidermis anterim to the copepod resulting in exposure of the fin rays and the lack of an inflammatory response.
Scale bar = 200 pm. Fig. E37. Higher magnification of a lesion caused by L. salmonis on a fin of a chinook salmon, 20 days post-infection. Nate that the epidermis is eroded to the basement membrane, and that there is little inflammatory response. Scale bar = 50 pm. (ch, chalimus larva; d, dermis; e, epdermis; fr, fin ray; ff, front. filament; mc, mouth cone).
FIGS. E38 to E40. kpeophtheirus salmonis frontal filament structure. Fig.
E38. Frontal filament of Lepeophtheim salmonis chalimus larva, 10 days post- infection. Note the fibrous stem and the had plate attached to the fin ray. Scale bar = 30 pm. Fig. E39. Premolt chalimus larva, 10 days pst-infection. Note the presence of a fully formed frontal filament contained within a cuticle-lined pocket in the anterior cephalothorax. The old frontal filament is outside of the plane of this section. Scale bar = 30 prn. Fig. 'E40. Premolt chalimus larva, 10 days pst- infection. Note the presence of a fully formed frontal filament contained within a cuticle-lined pocket in the anterior cephalothorax, and the presence of nuclei
(arrow) within the stem of the newly formed filament. Scale bar = 50 pm. (bp, basal plate; ff, frontal filament; fr, fin ray; mc, mouth cone; nc, new cuticle; oc, old cuticle; s, stem).
Effects of cortisol implants on the susceptibility, resistance, and
tissue responses of naive coho salmon to experimental infection
with Lepeophtheims salnonk (Copepoda: Caligidae).
Introduction
In Chapter 5 naive coho salmon (Oncorhynchus kisutck) were shown to be more resistant than naive chinook (Oncorhynchw tschmuytscha) or naive Atlantic salmon (Salmo sttlar) to experimental infection with Lepeophtheirus salmonis.
Histological examination of infected gills from coho salmon revealed a well developed inflammatory response as early as 1 day post-infection. Copepods were eliminated from the gills of coho salmon from 10 days post-infection onwards.
Histological examination of infected fins from coho salmon revealed variable epidermal erosion and inflammation of the dermis up to 5 days post-infection. At
10, 15, and 20 days post-infection the lesions were characterized by well. developed epidermal hyperplasias and intense inflammatory responses.
In the initial phases of the infection, reduction in the intensity of L. safmonis on coho salmon is thought to be caused by nonspecific immune responses, including epidermal hyperplasia, soluble or cellular factors of the - 174 - inflammatory response, and/or possibly serum enzymes or other proteins. In later stages of the infection specific humoral or cellular factors may be important.
Experimental administration of corticostmids to fish has been shown to inhibit inflammatory responses, inhibit phagocytosis, suppress both Rumoral and cellular immune responses, and retard the wound healing process (Pickering,
1987; Kent and Hedrick, 1987; Roubal and Bullock, 1988; Sad, 1988; Naxrul
Islam and Woo, 1991). Cortisol-treated fish are recognized as highly susceptible to a wide variety of diseases (reviewed in Pickering, 1987; Kent and Hedrick,
1987; Naxrul Islam and Woo, 1991).
This chapter investigates the effects of cortisol administration on the susceptibility and resistance of naive coho salmon to experimental infection with
L. salmonis. This study was conducted to confirm whether nonspecific host defenses acting alone or in combination with specific host defenses are responsible for the decline in intensity of L. salmonis on naive coho salmon. The effects of cortisol administration on the hist~pathologyof L. salmonis lesions on the gills and fins are described. - 175 - Materials and Methods
Coconut oil was pasteurized at 7W,cooled to 3W, and mixed with
hydrocortisol (Sigma No. H-4001) to yield a final concentration of 127 mg
hydrocortisol ml-' oil. Forty-five naive coho were anesthetized with MS-222
(tricaine methanesuifonate), adipose fin-clipped, and injected i.p. with 0.2 ml of
cortisol solution per fish. This resulted in an implant of approximately 0.5 mg
hydrocortisol g-' of fish based on an average fish weight of 51.0 grams. Forty-five
naive coho of the same size were selected as controls. These fish were
anesthetized but not injected .
The cortisol-implanted and control fish were introduced into separate 5OQ
liter fiberglass tanks and allowed to acclimate for 1 week. The cortisol-implanted
fish were then transferred into the tank containing the control fish and both groups
of fish were exposed for 24 hours to approximately 4000 newly molted copepodid
larvae. The methods for obtaining the copepodids and for carrying out the
infections have been outlined in Chapters 3 and 5. Prior to copepodid exposure and at 10 and 20 days post-infection, plasma samples were collected from 5 fish
from each of the control and cortisol-implanted groups for cortisol analysis. At sampling, the fish were anesthetized with a high dose of MS-222 (tricaine
methanesulfonate), had their caudal fins severed, and their blood w~scollated in
5 rnl heparinized tubes for plasma cortisol analysis.
Fish were maintained in flowing sea water with a temperature of 10.7 to
12.6T (mean = 1lS•‹C) and ambient salinity (29 to 31 %). All fish were fed a - 176 - commercial dry pellet feed at 1% body weight per day. At 1, 3, 5, 10, 15, and 20 days post-infection six of each of the cortisol-implanted and control fish were rapidly killed with MS-222. The fork length and wet weight was determined for each fish. Both the anesthetic bath and the body surfaces were examined for copepods and the distribution of the copepods on the fish was noted. The number of copepods present was corrected to a standad wet body weight to compensate for differences in size between hosts.
Intensity data was log (X+ i) transformed and differences in copepod intensity investigated by analysis of variance (ANOVA) procedures. Comparisons of copepod intensity for each treatment over time were made using Scheffbs tests
(Zar, 1984). Comparisons of copepod intensity between treatments at each sampling period were made using t-tests.
Cortisol Analysis
Blood samples were spun at 6500 rpm for 5 min, and the resulting plasma was transferred to sterile cryotubes. Plasma samples werc stored frozen at -70•‹C and await analysis. Cortisol values will be determined using an enzyme immunoassay (EIA) technique.
Light Micrascopy
Tissues for exmination by light microscopy were fixed in Davidson's solution and dehydrated through to 100% alcohol. Tissues were either wax- embedded, cut to a thickness of 5 pm, and stained with hematoxlylin and eosin, or - 177 - they were embedded in JB4 plastic resin, cut to a thickness of 1 to 2 pm, and stained with Lee's stain (methylene blue and basic fuschin). Results
Intensity of Infection
The intensitj. of L. salmonis on the cortisol-implanted and control groups over time is presented in Fig. F1. The intensities of iniection for both the cortisol- implanted and control groups were significantly different over time ANOVA; cortisol implanted (P < 0.001); control (P < 0.001)). The results of multiple range tests (Scheffes test; P < 0.05) over time showed the control group to have significantly fewer copepods at 20 days post-infection when compared to 1, 3, 5, and 10 days post-infection, and the cortisol-implanted group to have significantly fewer parasites at 20 days post-infection when compared to 1, 3, 5, 10, and 15 days post-infection (Fig. Fl). At each sampling time, significantly fewer copepods were present on the control group than on the cortisol-implanted group (T-test; P < 0.05). Distribution With the exception of 20 days post-infection the highest percentage of copepods recovered from the control group was from the fins (Fig. F2). The percentage of copepods recovered from the anesthetic bath decreased from approximately 21 % to 0% by 10 days post-infection. Of the copepods identified on the body, approximately 30% were collected from the gills at 1 and 3 days pst-infection. This percentage had declined to 0% by 10 days post-infection. Small numbers of copepods were recovered from the body surfaces at 1, 3, and 5 - 179 - days post-infection. At 15 md 20 days post-infection the percentages on the body surfaces increased due to molting io the preadult stage. With the exception of 20 days post-infection the highest percentages of copepods recovered from the cortisol-implanted group were from the gills and fins (Fig. F3). Small percentages of copepods were recovered from the anesthetic bath up to 5 days post-infection, and from the body surfaces from 1 to 10 days post-infection. At 15 and 20 days post-infection the percentages on the body surfaces increased due to molting to the preadult stage. Mortulity cknd Cross Morphology of the Lesions Sixteen percent of the cortisol-implanted fish died between 9 and 17 days post-infection. These fish showed severe erosion of the snout and fins caused by heavy infections of filamentous bacteria (seawater "myxobacteriosis"). From 5 days post-infection onwards dl of the cortisol-implanted fish were lethargic and showed varying amounts of erosion of the snout and/or fins. No mortality occurred in the control group and no body andlor En erosion was evident in the control fish sampled. Grossly, the gills of both the control and cortisol-implanted groups at 2, 3, and 5 days post-infection showed variable amounts of erosion and clubbing of the filament tips. From 10 to 20 days post-infection the gills of the cortisol-implanted group showed progressive erosion of the filament tips, clubbing, and fusion of the secondary Iarnellae. - 180 - Gross examination of 45 lesions on the fins of the control group over the period of 10 to 20 days post-infection revealed areas of proliferating host tissue immediately adjacent to or surrounding the copepods. Gross examination of 45 lesions on the fins of the cortisol-implanted group over the same period revealed no tissue response to the presence of the copepods. Gill Histology At 1 and 5 days post-infection, attachment and feeding sites on the gills of control coho salmon were characterized by variable amounts of erosion of the epidermis and dermis, hemorrhage, and inflammation of the dermis (Figs. F4 to F7). In some cases minor epidermal hyperplasia resulted in fusion of the secondary lamellae near the tips of the primary larnellae. In severe cases the central rods of the primary lamellae were exposed. The inflammatory infiltrate consisted of abundant neutrophils and a few lymphocytes (Figs. F5 and F?). No secondary infection of the gills was observed over this period. At 1, 5, and 10 days post-infection, attachment and feeding sites on the gills of cortisol-hnplanted coho salmon were characterized by variable amounts of erosion of the epidermis and dermis, hemorrhage, and mild inflammatory responses (Figs. F8 and F9). In most instances the inflammatory response was limited to the presence of scattered inflammatory cells (mostly neutrophils) within the dermis. In some sections a small amount of epidermal hyperplasia and fusion of the secondary lamellae was evident. At 1 and 5 days post-infection, both the intensities of the inflammatory responses and the extent of epidermal hyperplasias - 181 - appeared to be less than those seen in the control group. Examination of attachment and feeding sites at 15 and 20 days post-infection revealed increased numbers of primary Iamellae affected, well developed inflammatory responses, and increased levels of epidermal hyperplasia and fusion of the seccsfidary gill lamellae (Figs. F 10 to F13). Hemorrhage in the tissues at the site of parasite attachment was a common feature of the lesions over this period. The inflammatory infiltrate consisted primarily of neutrophils, but lymphocytes were also present (Fig. F13). Secondary infection of the gill lesions by filamentous bacteria occurred in some sections (Fig. F9). Fin Histology Over the period of 1 to 5 days post-infection, attachment and feeding sites on fins of both the control and cortisol-implanted groups were characterized by variable amounts of epidermal erosion and necrosis (Figs. F14 and F15). Over this period the severity of the lesions was highly variable within treatment groups with later lesions not necessarily more severe than earlier lesions. Mild inflammation of the dermis occurred in the control group as early as 1 day post-infection (Fig. F14). Neutrophils were the predominant cells at these sites of inflammation. No inflammation was observed in the cortisol-implanted group over this period. At 10, 15, and 20 days post infection, attachment and feeding sites on the fins of the control group were characterized by well developed epithelial hyperplasias (Figs. F16 and F 18), which in severe cases resulted in the complete - 182 - encapsulation of the copepod. In cases of encapsulation the spaces surrounding the copepod were filled with a mixed inflammatory infiltrate (neutrophils, macrophages, and a few lymphocytes), and necrotic tissue. Intense inflammation of the dermis occurred in all control samples over this period. The inflammatory infiltrate consisted of abundant neutrophils and rnacrophages and a few lymphocytes (Figs. F 17 and F19). With exception of two samples that showed mild epidermal hyperplasia, hyperplasia was not seen in the cortisol-implanted group over the period of 10 to 20 days post-infection (Figs. F20 and F22). Mild inflammation of the dermis occurred in all samples collected at 15 and 20 days post-infection. The inflammatory infiltrate consisted of neutrophils, some macrophages, and some lymphocytes (Fig. F22). - 183 - Diussion The level of hydrocortisol implanted in this investigation is higher than the lwei of 0.01 mg cortisol g-' body weight which has been Cenionstratd to elevate blood cortisol levels and cause chronic immunosuppression in coho salmon (Maule et ul., 1987). Levels of 0.5 mg cortisol g-' body weight and less have also been demonstrated to cause chronic immunosuppression of both rainbow and brown trout (Pickering and Duston, 1983; Pickering and Pottinger, 1985; Kent and Hedrick, 1987; Woo et al., 1987). That the level of hydrocortisol implanted was sufficient to cause chronic immunosuppression is shown by the high prevalence of snout and fin erosion caused by secondary bacterial infections, as well as 16% mortality in the cortisol-implanted group. The intensity of L. salmonis was significantly higher on the cortisol- implanted group when compared to the control group on each day sampled. Thus treatment with cortisol obviously predisposed naive coho salmon to infection with L. salmonis. This is the first report of a corticosteroid treatment affectkg the susceptibility and/or resistance of fish to infection by parasitic copepods. Administration of corticosteroids to fish has earlier been demonstrated to enhance the establishment and increase the intensity of parasitic infections and to make resistant hosts stisceptible to parasitic infection. Robertson et al. (1963) reported that rainbow trout given intraperitoneal implants of cortisol and cholesterol developed heavy infections of the ciliate parasite Ichthyophthin'us rnultijilis. Implantation of cortisol in rainbow trout resulted in increased numbers of the PKX - 184 - myxosporean present, and enhanced the ability of the myxosporean to reach the sporogonic stage (Kent and Hedrick, 1987). Cortisol-implanted rainbow trout also had a significantly higher prasitaemia, higher mortality, and lower antibody titres than control fish when challenged with the haemoflagellate Cryptobia sulrnosi~r'ca (Woo et al., 1987). Immunized juvenile mirror carp (Qprinus curpio) that were injected with the corticostereid triamcinolone acetonide suffered 100 % mortality compared to 0% in the immunized control groups after exposure to Ichthyophthirius multi$lis (Hoasgh ton and hrlatrkws, 1986). In the present experiment, implantation of hydrocortisol reduced the inflammatory response of both the gills and fins of naive coho salmon to L. salmonis. Administration of corticosteriods have been previously reported to suppress both chemically and surgically-induced inflammation in rainbow trout (Weinreb, 1958), to suppress the inflammatory response of interstitial kidney tissue of rainbow trout to the PKX myxosporean (Kent and Hedrick, 1987), and to cause significant reductions in the extent of inflammatory cell infiltration (mostly neutrophils and macropheges) in the peritoneum of plaice (MacAurther et ul., 1984). Fletcher (1986) demonstrated in vitro that cortisol at concentrations normally found in stressed plaice significantly reduced the migration of ncutrophils derived from the peritoneum. Epidermal hyperplasia was a consistent feature of L. sulmonis lesions on the fins of the control group over the period of 10 to 20 days post-infection. With exception of two mild cases, epidermal hyperplasia was not observed in the - 185 - cortisol-implanted group over the same period. Suppression of epidermal hyperpiasia by cortisol treatment has not been previously reported. Although hyperplasia is a common response of the fish epidermis when exposed to a wide variety of agents such as chemical pollutants, hormonal stimuli, and bacteria or viruses (see Roberts, 1978), the mechanisms by which it is controlled are not understood. Cortisol may limit epidermal hyperpiasia by affecting the mitotic activity of the fish epidermis. This author knows of no data on the effects of corticostemlds on mitotic activity in the fish epidermis. In the amphibian Ram pipiens an inverse relationship between epidermal mitotic rate and the level of plasma corticosteriods has been reported (Garcia-Arce and Mizell, 1972). Hydrocortisone is also known to inhibit mammalian epidermal healing by retarding mitotic activity (Pickering, 1987), Epidermal hyperplasia is commonly reported in association with sites of chronic inflammation in fish. This association suggests that components of the nonspecific and/or specific humoral immune system may be important in the initiation and/or maintenance of hyperplasia. If this is the case cortisol may limit epidermal hyperplasia through its suppression of the nonspecific and specific humoral systems. The percentage of copepods on the gills of the cortisol-implanted group remained approximately constant throughout the experiment. The percentage of copepods on the fins of the cortisol-implanted group declined and the percentage on the body increased by 21) days post-infection with the molting to the preadult - 186 - stage. These results suggest that L. salmonis developed to the preadult stage faster on the fins when compared to the gills. In Chapter 5, f reported a slower development rate of L. salmonis on the gills of Atlantic salmon when compared to the fins. The level of cortisol implanted in this study may have elevated serum cortisol levels beyond the normal physiological range for coho salmon. Therefore, the results cannot be taken as proof that stress or diseaseinduced elevations of serum cortisol would lead to increased susceptibility and decrease resistance of naive coho to infestation with L. salmonis. Further study is required to determine if similar results can be obtained with serum cortisol levels that mimic those seen in stressed or diseased coho salmon. In summary, naive coho salmon implanted with hydrocortisol had higher intensities of L. salmonis than the control group throughout the experiment, suggesting an increased susceptibility and decreased resistance to infection. Non- specific host defence mechanisms, including the magnitude of the inflammatory response and the development of epidermal hyperplasia, were suppressed in the cortisol-implanted group. btkh the inflammatory response and epidermal hyperplasia appear to be responsible for the low susceptibility and high resistance of naive coho salmon to infestation with L. salmonis. Cortisol may suppress epidermal hyperplasia by reducing the mitotic activity of the epidermis, or through its suppression of the nonspecific andlor specific humoral immune systems. FIG. Fl. Mean (+\-SE) intensity of Lepeophtheim salmonis on naive cortisol-implanted and naive control coho salmon at various times post-infection. Fish were maintained at 10.7 to 12.6 "C and ambient salinity (29 to 31 %o). Values above error bars indicated which days do not have significant differences in L. salrnot~isintensity (Scheffbs test; P < 0.05). FIG. F2. Distribution of Lepeophtheiw salmonis on naive control coho salmon. Fish were maintained at 10.7 to 12.6 "C and ambient salinity (29 to 31 %o), Bath Body Fins G~lls I 3 5 10 15 20 DAYS POST-INFECTION FIG. F3. Distribution of Lepegnhtheinrs salmonis on naive cortisol- implrnted coho salmon. Fish were maintained at 10.7 to 12.6 "C and ambient salir'ity (29 to 31 960). 0Bath Body DAYS POST-INFECTION FIGS. F4 to F7. Lepeophtheirus salmnis copepodids on the gills of naive control coho salmon. Fig. F4. Copepodid on a gill of a control coho salmon, 1 day post-infection. Note the mild epidermal hyperplasia and inflammation of the dermis, Scale bar = 200 pm. Fig. F5. The inflammatory response of a gill of a control coho salmon to L. salmonis, 1 day post-infection. Note the presence of a mixed inflammatory infiltrate comprised mostly of neutrophils. Scale bar = 30 pm. Fig. F6. Copepodid on a gill of a control coho salmon, 5 days post-infection. Note the mi!d erosion of the, epidermis and mild inflammatory response. Scale bar = 200 pm. Fig. F7. The inflammatory response of a gill of a control coho salmon to L. salmnis, 5 days post-infection. Note the presznce of a mixed inflammatory infiltrate camprised mostly of neutrophils. Scale bar = 30 pm. (c, copepodid; cr, central rod; i, mixed inflammatory infiltrate; mc, mouth cone). FIGS. F8 to F11. Lepeophtheim salnzonis copepodids and chalirnw larvae on the gills of naive cortisol-implanted coho salmon. Fig. F8. Copepodid on a gill of a cortisol-implanted coho salmon, 5 days post-infection. Note the severe erosion of the epidermis resulting in exposure of the central rod of the primary lamellae. Scale bar = 200 gm. Fig. F9. Feeding site of L. salmonis on a gill of a cortisol-implanted coho salmon, 5 days post-infection. Note the exposed central rod of the primary lamella, and the presence of a secondary bacterial infection. Scale bar = 30 pm. Fig. F10. Chalimus larva on a gill of a cortisol-implanted coho salmon, 15 days post-infection. Note the hyperplastic epidermis causing fusion of the secondary lamellae, the extensive hemorrhage, and the well developed inflammatory response. Scale bar = 200 pm. Fig. F11. Attachment site of L. salmonis on a gill of a cortisol-implanted coho salmon, 15 days post-infection. Note the frontal filament attached to the central rod of the primary lamella, the hemorrhage, and the inflammation. Scale bar = 30 pm. (c, copepodid; ch, chalimus larva; cr. central rod; fb, filamentous bacteria; ff, frontal filament; h, hemorrhage; i, mixed inflammatory infiltrate). FIGS. F 12 and F13. Lepeophtheinrs salmonis chalimus larvae on the gills of naive cortisol-impIanted coho salmon. Fig. F12. Chalimus larva on a gill of a cortisol-implanted coho salmon, 20 days post-infection. Note the hyperplastic epidermis causing fusion of the secondary lamellae, and the well developed inflammatory response. Scale bar = 200 pm. Fig. F13. Feeding site of L. salmonis on a gill of a cortisol-implanted coho salmon, 20 days post-infection. Nok the well developed inflammatory response. Scale bar = 30 pm. (ch, chalimus larva; cr, central rod; i, mixed inflammatory infiltrate). FIGS. F14 to F17. Lepeophtheiw salmonis copepodids on the fins of naive control and naive cortisol-implanted coho salmon. Fig. F14. Copepodid on a fin of a control coho salmon, 1 day post-infection. Note the erosion of the epidermis and the inEarnmatory response within the dermis. Scale bar = 50 pm. Fig. F15. Copepodid on a fin of a cortisol-implanted coho salmon, 1 day post-infection. Note the erosion of the epidermis, the tip of the second antenna penetrating the basemerst membrane (arrow), and the lack of an inflammatory response. Scale bar = 59 pm. Fig. F16. Chalimus larva on a fin of a control coho salmon, 10 days post-infection. Note the hyperplastic epidermis which completely surrmnds the coppod and the well developed inflammatory response. Scale bar = 200 pm. Fig. F 17. Edge of a lesion caused by L. salmonis fixding on a fin of a control coho salmon, 10 days post-infection. Note the well developed inflammatory response with the infiltrate consisting primarily of neutrophils and macrophages. Sdebar = 30 pm. (bm, basement membrane; ch, chalimus larva; fr, fin ray; he, hyperplastic epidermis; i, mixed inflammatory infiltrate; mc, mouth cone; sa, second antenna). FIGS. F18 to F2 1. Lepeophtheirus salmnis preadults and chalimus larvae attached on the fins of naive control and naive cortisol-implanted coho salmon. Fig. F18. Attached preadult on a fin of a control coho salmon, 20 days post-infection. Note the well developed epidermal hyperplasia and inflammation of the dermis. Scale bar = 200 pm. Fig. F19. Higher magnification of the inflammatory infiltrate of a control coho salmon, 20 days post-infection. Note the presence of abundant neutrophils, macrophages, and a few lymphocytes. Scale bar = 25 pm. Fig. F20. Chalimus larva on a fin of a cortisol-implanted coho salmon, li) days post-infection. Note the eroded epidermis, the exposed basement membrane. and lack the of any epidermal hyperplasia andlor inflammation. Scale bar = 200 pm. Fig. F21. Attachment of the frontal filament to a fin of a cortisol- implanted coho salmon, 10 days post-infection. Note the lack of an inflammatory response. Scale bar = 30 pm. (bp, basal plate; fr, fin ray; he, hyperplastic epidermis; i, mixed inflammatory infiltrate; mc, mouth cone; s, stem). Fig. F22. Feeding site of a chalimus larva of Lepeophtheiw sdmonis on a fin of a naive cortisol-implanted coho salmon, 20 days post-infection. Note the erosion of the epidermis, the exposed fin rays, the mild inflammatory respnse, and the secondary bacterial infection. Scale bar = 30 pm. (fb, filamentous bacteria; fr, fin ray). GENERAL DISCUSSION Many people involved in salmonid culture and/or salmonid health find it difficult to distinguish between species of sea lice. The key provided in chapter 1 will enable its users to easily distinguish between all species of sea lice found on trout and salmon at both the preadult and adult stages. Due to marked differences between sea lice species in their biology and behavior, correct identification of the species responsible for a disease outbreak is necessary before decisions regarding their treatment can be made. For example, preadult and adult Lepeophtheirus salmonis rarely become free-swimming and transfer between salmonid hosts (Bruno and Stone, 1990; unpublished observations). Therefore, once the preadults and adults are removed they are unlikely to reinfect the hosts. In contrast, preadults and adults of the Caligus species and Lepeophtheirus cuneifer commonly become free-swimming and transfer from salmon to salmon, and from non- salmonids to salmon (Bruno and Stone, 1990; present study). In British Columbia, large numbers of adult Caligus clemensi have been reported to appear on sea- farmed salmonids following the appearance of herring l., the vicinity of the farm site (unpublished observation). When the preadult and adult stages of these species are removed from their hosts they can easily transfer back andlor infect other salmon stocks held rn tits vicinity. - 197 - The descriptions of the developmental stages of L. salmonis provided in chapter 2 will greatly aid other researchers wishing to conduct research into the population biology of this species. The ability to distinguish L. salmonis from co- occurring C. clemensi at all developmental stages except the nauplius, makes possible future investigations of the population dynamics of these species in British Columbia waters. An important contribution to sea lice research would be a description of the developmental stages of Caligus elongatus. As noted in chapter 1, C. elongatus is the most important species of sea lice in Atlantic Canada, and second only in importance to L. salmonis in Scotland and Norway. The developmental stages of both Caligus cum, and L. cuneifer also require description. However, as these species are relatively rare on salmonids, work on their descriptions is not a high priority. The most important stage in the life cycle of L. salmonis and other species of sea lice is the infectious copepodid stage. In spite of its importance little is known about the behavior and host-finding capability of this stage. In chapter 2 numerous sensory structures of the copepodid stage are identified. However, at present the importance of these structures with respect to dispersal and host- location by L, salmonis can only be speculated upon. Data on the responses sf L. salmonis md other species of sea lice to physical stimuli such as fight (both spectral and intensity sensitivity), shadows, and water movement would greatly increase our understanding of dispersal and - 198 - host-location in these species. Past studies on the behavior of L. salmonis and other species of parasitic copepods (eg. Johannessen, 1978; Poulin et al., 1990) have relied on simple methodologies, which are recognized to produce mostly laboratory artifacts (see Forward, 1981). To obtain meaningful information that which can be extrapolated to behaviors in the field, future studies on the behavior L. salmonis and other species of sea lice need to adopt the equipment and methodologies which are being currently used to study the behavior of free-living zooplankton (eg . Forward, 1981). Data obtained experimentally on development, growth, and survival are provided in chapter 3. These data form an important baseline in our understanding of the epizootiology of L. salmonis on sea-farmed Atlantic salmon. Additional studies on the development rate of L. salmonis to the adult stage should be conducted at other temperatures than used in the present study. Such information would be useful for determining the frequency of sea lice treatments at aquaculture sites. Similar studies to those conducted here are required for the other species of sea lice, especially C. clemensi and C. elongatus. In chapter 5 I report that the development rate of L. salinonis differs on different host species and between different regions of the body of individual host species. Investigations are required to determine the magnitude of these differences as well to identify host defence mechanisms andlor nutritional factors which may be responsible for these differences. Future studies on the development - 199 - rate of L. salmonis and other spies of sea lice need to consider these findings when determining their experimental design. Naturally infected Atlantic salmon show serum responses to only five antigens of L. salmonis in reduced samples and one major antigen in unrduced samples (Grayson et al. 1991). These antigens have been shown by immunohistochemical studies to be associated exclusively with the apices of the midgut epithelium of adult L. salmonis. Based on the results in chapter 4, the cells most likely responsible for eliciting these antigens are mature type-C cells. These cells function in absorption, lipid storage, and lipid metabolism. Lepeophtheirus salmonis shows no preference when given a choice between naive coho, chinook, and Atlantic salmon (chapter 5). Differences between these species in their resistance to L. salmonis infection is related to the intensity of their non-specific and possibly their specific immune responses to the copepod. Naturally infected Atlantic salmon have been demonstrated to mount a low level of antibody response to L. salmonis (Grayson et al., 1991). Research is required to determine if either chinook or coho salmon develop specific antibodies to L. salmonis. Teleost fish respond to conditions of stress by elevation of the level of serum corticosteroid hormones (see Pickering, 1987). Increased levels of these hormones result in immunosuppression and anti-inflammatory effects, In chapter 6 chronic immunosuppression of coho salmon by the implantation of hydrocortisol resdted in an increase in the susceptibility and decrease in the resistance to - 200 - infection with L. salmonis. These data suggest that any factor that causes elevated serum cortisol levels can modiQ the differences in susceptibility and resistance seen between and within salmon hosts challenged with L. salmonis, as well as increase the severity of the disease problem. Numerous ecological and physiological factors, acting alone or in conjunction with each other can stress fish. Examples of these factors include: sudden temperature changes, low dissolved oxygen, plankton blooms, effects of crowding, physical handling, concurrent disease conditions, and sexual maturation. 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