Group Title: new nematode parasite of mole crickets
Title: A new nematode parasite of mole crickets
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Permanent Link: http://ufdc.ufl.edu/UF00099468/00001
 Material Information
Title: A new nematode parasite of mole crickets its taxonomy, biology and potential for biological control
Physical Description: xi, 154 leaves : ill. ; 28 cm.
Language: English
Creator: Nguyen, Khuong Ba, 1941-
Publication Date: 1988
Copyright Date: 1988
 Subjects
Subject: Mole crickets -- Biological control   ( lcsh )
Nematodes   ( lcsh )
Beneficial insects   ( lcsh )
Entomology and Nematology thesis Ph. D
Dissertations, Academic -- Entomology and Nematology -- UF
Genre: bibliography   ( marcgt )
non-fiction   ( marcgt )
 Notes
Thesis: Thesis (Ph. D.)--University of Florida, 1988.
Bibliography: Includes bibliographical references (leaves 149-153).
Statement of Responsibility: by Khuong Ba Nguyen.
General Note: Typescript.
General Note: Vita.
 Record Information
Bibliographic ID: UF00099468
Volume ID: VID00001
Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: All rights reserved by the source institution and holding location.
Resource Identifier: alephbibnum - 001469561
oclc - 20749139
notis - AGY1254

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A NEW NEMATODE PARASITE OF MOLE CRICKETS:
ITS TAXONOMY, BIOLOGY AND POTENTIAL FOR BIOLOGICAL CONTROL

















By

KHUONG BA NGUYEN


A TDSSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


1988















ACKNOWLEDGEMENTS


The author is deeply grateful to the chairman of his

committee, Dr. G. C. Smart, Jr., who has given considerable

time and immeasurable assistance during the term of this

study. His suggestions made during the writing of this

dissertation are greatly appreciated.

Gratitude is expressed also to Drs. R. P. Esser, S. R.

Farrah, J. L. Nation, and A. C. Tarjan for their invaluable

assistance while serving as members of his supervisory

committee.

Thanks are given also to Dr. T. J. Walker for

information related to mole crickets, and to Dr. H. L.

Cromroy for identifying the mite Rhizoqlyphus sp.

Finally, the author is very grateful to his wife and

children for their encouragement and understanding which

were essential for the completion of this work.

















TABLE OF CONTENTS


ACKNOWLEDGEMENTS......................................

LIST OF TABLES ......................................

LIST OF FIGURES .....................................

ABSTRACT ............................................

CHAPTERS


1 INTRODUCTION ..............................

2 STEINERNEMA SCAPTERISCI N. SP.
(STEINERNEMATIDAE : NEMATODA) FROM URUGUAY,
SOUTH AMERICA.................................

Materials and Methods .....................
Results and Discussion ....................

3 MODE OF ENTRY OF STEINERNEMA SCAPTERISCI N.
SP. INTO MOLE CRICKETS ......................

Literature Review .........................
Materials and Methods .....................
Results and Discussion ....................

4 LIFE CYCLE OF STEINERNEMA SCAPTERISCI N. SP..

Materials and Methods .....................
Results and Discussion ....................

5 CULTURE OF STEINERNEMA SCAPTERISCI N. SP.
IN VIVO .......................................

Literature Review .........................
Material and Methods ......................
Results and Discussion ....................

6 CULTURE OF STEINERNEMA SCAPTERISCI N. SP.
IN VITRO. .....................................

Literature Review .........................

iii


Page

ii

v

vii

x



1



3

3
6


33

33
34
39

42

42
44


59

60
60
66


71

71









Materials and Methods ..................... 76
Results and Discussion .................... 82
7 VERTICAL MIGRATION OF STEINERNEMA
SCAPTERISCI N. SP.IN SOIL.................... 89

Literature Review ............. ... .......... 89
Materials and Methods ..................... 90
Results and Discussion .................... 95

8 SURVIVAL OF STEINERNEMA SCAPTERISCI N. SP.... 103

Materials and Methods ..................... 103
Results and Discussion .................... 105

9 STEINERNEMA SCAPTERISCI N. SP. AS A BIOLOGICAL
CONTROL AGENT OF MOLE CRICKETS .............. 110

Materials and Methods ..................... 111
Results and Discussion .................... 115

10 FACTORS INFLUENCING THE CONTROL OF MOLE
CRICKETS BY STEINERNEMA SCAPTERISCI N. SP.... 124

Materials and Methods ..................... 124
Results and Discussion .................... 127

11 NEMATODE PARASITES AND ASSOCIATES OF MOLE
CRICKETS ...................................... 135

Literature Review ........................... 135
Materials and Methods ..................... 136
Results and Discussion .................... 137
Conclusions ................................. 144

12 SUMMARY AND CONCLUSION ...................... 145

LITERATURE CITED ...................................... 149

BIOGRAPHICAL SKETCH .................................... 154















LIST OF TABLES


Table Page

1 Measurements (in um) of first and second
generation females of Steinernema scapterisci n.
sp. (n=10)....................................... 7

2 Measurements (in um) of first and second
generation males of Steinernema scapterisci n.
sp. (n=0)........................................ 8

3 Measurements (in um) of the third stage juvenile
of Steinernema scapterisci n. sp. (n=20)......... 9

4 Effectiveness of different species of Steinernema
and strains of S. carpocapsae in killing four
species of lepidopterous insects, two days after
inoculation...................................... 30

5 Influence of temperature on the life cycle of
Steinernema scapterisci......................... 47

6 Number and percentage of first generation
females and males of Steinernema scapterisci
which developed at 15 C......................... 52

7 Number and percentage of first generation
females and males of Steinernema scapterisci
which developed at 24 C.......................... 53

8 Number and percentage of first generation
females and males of Steinernema scapterisci
which developed at 30 C......................... 54

9 Numbers of mole crickets that died after 2 and 4
days when using 8,000 Steinenema scapterisci
strain H3 and 8,000 strain MC in petri dishes
containing mole crickets....................... 70

10 Cultural and biological characteristics of the
bacterium associated with Steinernema
scapterisci...................................... 84









11 Number of third-stage infective juvenile
nematodes harvested from each culture medium..... 86

12 Number of mole crickets killed by nematodes
produced by in vitro culture..................... 86

13 Comparison of nematodes produced in vitro vs.
in vivo for kill of mole cricket nymphs........... 88

14 Distribution of third-stage juvenile nematodes in
2-cm layers of soil when 5,000 and 10,000 third-
stage juveniles were applied to the soil surface. 97

15 The number of infective-stage juvenile
nematodes recovered from each 2-cm layer of soil
5 days after they were released at point 0...... 100

16 Number of mole crickets that died when buried at
each depth listed below when 200,000
infective-stage juvenile nematodes/square
meter were released on the soil surface........... 102

17 Numbers and percentages of infective-stage
juvenile Steinernema scapterisci recovered weekly
after releasing 2,000 (Experiment 1) and 2,500
(Experiment 2) nematodes in containers of
sterilized soil buried in the field.............. 107

18 Number and percentage of third-stage Steinernema
scapterisci recovered 6 to 14 weeks after
releasing 2,000 infective-stage juveniles
in containers of unsterilized soil and burying
them in the field............................... 109


19 Effect of the nematode Steinernema scapterisci
on different species of mole crickets............ 116


20 Percentage of the mole crickets, Scapteriscus
vicinus and S. acletus, killed by Steinernema
scapterisci..................................... 117


21 Effect of Steinernema scapterisci on earth
worms......................... .. ............... 120

22 Percentage of different insects killed by Steinernema
scapterisci..................................... 122















LIST OF FIGURES


Figure Page

1 Steinernema scapterisci n. sp.. A) Face view
showing unevenly distributed papillae. B) Female
of the second generation, entire body. C) Double-
flapped epiptygma. D) Variation in tails of the
first generation females. E) Variation in tails of
the second generation female. F) Anterior part of
the first generation female showing large
cheilorhabdion and excretory duct with doughnut-
shaped structure................................. 11

2 Steinernema scapterisci n. sp.. A) Anterior part of the
nematode showing the mouth, lips and excretory pore
B) Head with 6 labial papillae. C) Face view showing
unevenly distributed labial papillae; the mouth
circular at the opening, becomes subtriangular. Note
the white material covering the papillae. D) Face
view showing the mouth and labial papillae........ 14

3 Steinernema scapterisci n. sp.. A) Posterior region of
the third-stage infective juvenile showing the
curvature tail. B) Live nematode showing doughnut-
shaped structure. C) Head of the nematode showing thick
cheilorhabdions. D) Spicules and gubernaculum of male.
E) Fixed nematode showing doughnut-shaped
structure......................................... 16

4 Steinernema scapterisci n. sp. A) First generation
male tail with mucron; spicules with angular
head and ribs, gubernaculum with anterior portion bent
upward. B) Variation in tail shape of the
first generation male. C) Tail of the second
generation male showing elongate spicule head.
D) Variation in tail shape of the second
generation male. E) Entire body of the first
generation male ................................... 20

5 Steinernema scapterisci n. sp.. A) Spicule blade
showing the thin posterior part with a small aperture.
B) The spicule shaft showing the angular head, and a
sheath around the shaft. C) Cross section of the
spicule showing 2 lumens in the spicule blade. D)
Gubernaculum showing the long anterior part which









bends upward...................................... 22


6 A, B) Steinernema carpocapsae : strain Breton.
A) Gubernaculum showing short anterior part.
B) Spicule showing the short shaft without a sheath
and with rounded head (compare with Fig. 5B. C, D)
Steinernema scapterisci n. sp.. C) Double-flapped
epiptygma. D) The body of the third-stage infective
juvenile showing annulation and lateral field with 6
incisures........................................ 24

7 A) Steinernema scapterisci n. sp.. Posterior part of
the spicule, thin, with small aperture.
B, C, D) Posterior part of the spicule S. glaseri, S.
carpocapsae, and S. bibionis, respectively, showing
the difference in shape, thickness and size of
the aperture..................................... 26

8 Steinernema scapterisci n. sp.. A) Male tail of the
first generation showed 10 pairs of genital papillae.
B) Flexure of the ovary reflexedd in the body). C, D)
Head and tail of third-stage infective juvenile.
E) Ovary with spermatheca reflexedd in the body).
F) Cross section of spicule blade showed two lumens
(1) and 2 internal ribs (r) .... .................. 28

9 Inoculating a mole cricket through the mouth with
third-stage Steinernema scapterisci n. sp.......... 37

10 Third-stage Steinernema scapterisci n. sp. in the
tracheal tube of a mole cricket. The part of the
tracheal tube above the nematode was broken by the
nematodes from the main tube (nematodes inside)...... 37

11 Diagram of the life cycle of Steinernema
scapterisci n. sp................................. 49

12 The relationship between the percentage of males
and the total number of nematodes in each house
cricket at 24 C................................... 57

13 The relationship between the number of males
and the total number of nematodes in each house
cricket ............................................ 58

14 In vivo culture of the nematode Steinernema scapterisci
n. sp. Top: Live mole crickets placed in an inoculation
chamber with third juvenile nematodes. Bottom: Dead
mole crickets arranged in an incubation chamber..... 62

15 Top: Nematodes which developed within th bodies of mole

viii








crickets have migrated to the exterior. Bottom: Third-
stage juveniles have migrated from the mole crickets
into the water in the incubation chamber......... 63

16 Culture flasks inoculated with bacteria and
nematodes receive humidified and sterilized air
filtered through a bacteriological filter......... 81

17 PVC pipe used in migration test. Two-cm wide PVC pipe
rings were taped together and filled with soil. A
perforated petri dish containing 2 mole crickets was
attached to the bottom............................ 92

18 Encapsulation of the nematode by blood cells of the
mole cricket. Top: Blood cells attached together as
a band along the body of the nematode. Bottom:
Blood cells attached on one end of the nematode... 128

19 Encapsulation of the nematode by blood cells of the
mole cricket. Top: Head and tail of nematode
encapsulated. Note that thread-like structures
connect the cells and capsule. Bottom: Entire nematode
in capsule (flattened by a cover slip on a glass
slide) ............................................ 129

20 SEM photographs of nematodes encapsulated by blood
cells of a mole cricket........................... 130

21 Anterior part of the mite Rhizoqlvphus sp. eating
a nematode. ...................................... 133

22 The nematode Binema sp. found in the mole crickets,
Scapteriscus abreviatus, and Neocurtilla. Top: Entire
body of a female. Bottom: Middle part of a female
showing vulva and eggs............................ 139

23 The nematode Cameronia sp. found in the mole crickets
Scapteriscus acletus, S. vicinus, Neocurtilla
hexadactylla Left: anterior part. Right: egg in the
body. Bottom: male tail............................ 140

24 The nematode Talpicola sp. found in the mole cricket
Scapteriscus acletus. Top: anterior part. Bottom:
Posterior part showing eggs in the body............ 141

25 The nematode Pulchrocephala sp. found in the mole
cricket Neocurtilla hexadactvlla.Top: Anterior part.
Bottom: Posterior part of a female, showing eggs in a
chain ............................................. 143
















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

A NEW NEMATODE PARASITE OF MOLE CRICKETS:
ITS TAXONOMY, BIOLOGY AND POTENTIAL FOR BIOLOGICAL CONTROL

By

KHUONG BA NGUYEN

December, 1988

Chairman: G. C. Smart, Jr.
Major Department: Entomology and Nematology

A new steinernematid nematode parasite of mole crickets

was collected from Uruguay, South America. The nematode

does not fit any nominal species of the genus Steinernema

and is herein described as a new species.

The life cycle and sex ratio of the nematode is

influenced by temperature. At 10 to 15 C the life cycle is

not completed, at 20 C the cycle takes 12 days to complete,

at 24 C, 10 days and at 30 C, 8 days. At 15-24 C the number

of females in the population is greater than the number of

males, but at 30 C the reverse occurs.

The new species does not reproduce well, if at all, in

larvae of the wax moth, Galleria mellonella, which is a

universal host for all other species. Its host range

appears to be much narrower than that of other species, with

a penchant for mole crickets. When released in the field in









North Florida, the nematode became established, has spread

out from the original release sites, and continues to kill

mole crickets after 3 1/2 years. When released on the soil

surface, the nematode moved down and killed mole crickets

placed 10 cm below the soil surface. The nematode survived

in the soil for 10 weeks and retained its ability to kill

mole crickets. These attributes make it a very good

candidate for the biological control of mole crickets

imported accidentally to Florida.

In a survey of nematodes associated with mole crickets,

the following genera were found: Binema, Cameronia,

Cruznema, Diploqaster, Mesorhabditis, Pulchrocephala,

Steinernema, and Talpicola.














CHAPTER 1
INTRODUCTION


According to Walker (1984), the most important pests of

turf and pasture grasses in Florida are mole crickets in the

genus Scapteriscus, which cause damage also to crops such as

vegetables, ornamentals, tobacco, etc. The cost due to mole

cricket activity, including damage and cost of control, to

Floridians is about $45 million annually (Hudson and Short,

1988).

Management of the mole cricket problem in Florida began

in 1940 when tons of calcium arsenate bait were used to

control mole crickets on truck crops in central Florida. The

introduction of synthetic insecticides, especially DDT and

Chlordane, in 1945, temporarily solved the mole cricket

problem. When these insecticides were banned, however, and

when bahiagrass was used increasingly as pasture grasses,

the mole cricket problem became more serious (Walker, 1986).

In response to the concerns of cattlemen, farmers, turf

grass managers, and home owners, a research project on mole

crickets was begun at the University of Florida in 1978. The

objectives of this project were to eliminate or reduce the

damage caused by mole crickets in Florida, and at the same

time to study the fundamental biology of mole crickets. One











of the major directions of the project was to search for

natural enemies of mole crickets.

When searching for natural enemies of an organism, in

principle one should begin the search in areas indigenous to

the organism. The non-native mole crickets in Florida,

those in the genus Scapteriscus, most likely were introduced

to the southeastern United States from South America (Walker

and Nickle, 1981) where they are of little economic

importance. Evidently, there are factors in South America

which keep population densities below economic threshold

levels. A search for natural enemies of mole crickets in

that area showed that a steinernematid nematode may be the

limiting factor. This nematode was found in Brazil and

Uruguay and different isolates from Uruguay were hand-

carried to the United States by Dr. G. C. Smart, Jr. and

myself in 1985. We have worked with this nematode since.

The major objective of the research reported herein

was to study the biology and ecology of the nematode to

determine its effectiveness as a biological control agent of

mole crickets in Florida. A secondary objective was to

determine if nematodes are associated with mole crickets in

Florida.
















CHAPTER 2
STEINERNEMA SCAPTERISCI N. SP.
(STEINERNEMATIDAE : NEMATODA) FROM URUGUAY, SOUTH AMERICA


While searching for natural enemies of mole crickets in

South America, a steinernematid nematode infecting mole

crickets was found in Brazil and Uruguay. The nematode was

thought to be Steinernema carpocapsae (=Neoaplectana

carpocapsae), and several isolates from Uruguay were brought

to the quarantine laboratory in Gainesville, Florida by Dr.

G. C. Smart, Jr. and me in 1985. Its morphology and biology

showed that it is different from that species and it is

herein described as Steinernema scapterisci n. sp. and named

after its host, Scapteriscus, a genus containing mole

crickets.


Materials and Methods


Nematodes collected in Uruguay were inoculated into

mole crickets which were hand-carried to Florida. In

Florida, populations of the nematode were increased in the

mole crickets, Scapteriscus vicinus and S. acletus, and

later in the house cricket, Acheta domestic. These

nematodes, or their progeny were used for all studies.











Morphology

The first generation adults were collected from

infected mole crickets 2-3 days after the crickets died,

the second generation adults were collected 5-7 days after

the crickets died and the third-stage infective juveniles

were collected 7-15 days after the crickets died. The

nematodes were killed in warm water at 40 C, and mounted in

water on glass slides with coverglass supports. Many live

nematodes or nematodes stained with acid fushin were

observed to confirm the presence and/or nature of some

anatomic structures.

Scanning Electron Microscopy (SEM)

Nematodes prepared for SEM were placed live in

lactophenol at 43 C for 30 minutes, transferred to a

desiccator for two days, removed,rinsed with water, and then

prepared by the method of Stone and Green (1971). Specimens

were examined in a Hitachi S450 SEM.

To prepare spicules and gubernacula for SEM, male

nematodes of the first generation were placed in a petri

dish containing water, killed by low heat and stored at room

temperature. After 2-3 days when the bodies had softened

due to decay, they were transferred to clean water, and,

with two small needles, the rear portion of each nematode

was torn open, the spicules and gubernaculum dissected out

and washed free of debris by sloshing them about in water.

Then the spicules and gubernaculum were picked up with a











needle and placed on a previously-prepared SEM stub close to

a hair used as a marker.

Cross Hybridization

These studies were conducted using two different

techniques. In one technique, a drop of blood (hemolymph)

from a mole cricket was placed in a 35 x 10 mm sterile petri

dish, and one third-stage juvenile of S. scapterisci and

one of S. carpocapsae strain Breton added. The dish was

placed in a plastic bag containing a paper towel saturated

with water. The plastic bag was closed, tied and stored in

the dark. The treatment was replicated 25 times.

In the second technique, two drops of blood were prepared as

above, and 10 third-stage juveniles of S. scapterisci were

placed in one drop and 10 third-stage juveniles of S.

carpocapsae strain Breton were placed in the other drop.

They then were handled as above. The treatment was

replicated 10 times for each nematode. The nematodes were

observed daily and when the sexes could be distinguished but

before they became adults, all males in the dishes of S.

scapterisci were removed and placed in a separate drop of

blood. Similarly, the males of S. carpocapsae strain Breton

were removed and placed in a separate drop of blood. Then

the males of S. scapterisci were transferred to the drop of

blood containing females of S. carpocapsae strain Breton,

and males of S. carpocapsae strain Breton were transferred

to the drop of blood containing females of S. scapterisci.











The nematodes were observed frequently to see if they mated

and produced offspring. Nematodes of each species were

retained in drops of blood in two dishes as controls.

Biology

Four insects, fall army worm (Spodoptera fruqiperda),

velvet bean caterpillar (Anticarsia gemmatalis), granulate

cut-worm (Feltia subterrania), and wax-moth larva (Galleria

mellonella) were used to compare the rate of kill by S.

scapterisci to that of some other species and strains of

Steinernema.

Two pieces of Whatman No. 2 filter paper were placed in

a 100 x 15 mm petri dish and 8,000 third-stage juveniles in

2 ml water, and 10 insects were added. Controls were

prepared similarly but without nematodes. Treatments were

replicated 4 times. After 2 days the number of dead insects

was determined.


Results


Measurements: Measurements for first and second generation

females are presented in Table 1, those for first and second

generation males in Table 2 and those for third stage

juveniles in Table 3.

Description

Females, first generation (Fig. ID,1F)

The body cuticle is smooth, the lateral fields and

phasmids were not observed. The head is rounded, continuous












Table 1: Measurements (in um) of first and second
generation females of Steinernema scapterisci n.
sp. (n=10).


Body length

Greatest width

Stoma length

Stoma width

EP

NR

ES

Tail length

Anal body width

Percentage vulva

EP:ES 0.


First generation

Mean (SD) Range

4162 (540) 3531-515

179 ( 13) 159-203

7.5 ( 1) 6-9

10 ( 3) 9-12

89 ( 5) 78-94

174 ( 13) 153-194

242 ( 17) 219-269

46 ( 8) 34-59

58 ( 9) 41-72

53 ( 2) 50-54

.37 (0.03) 0.32-0.4


Second generation

Mean SD)\ Ranne


56





















1


2209

123

6.7

8.9

78

169

241

58

47

52

0.32


(223)

( 14)

(1.4)

(0.9)

(6.8)

( 12)

( 15)

( 4)

(2.8)

( 2)

(0.3)


1841-2530

94-141

5-9

8-11

66-88

147-184

222-266

48-64

43-52

52-60

0.28-0.36


EP = Distance from anterior end to excretory pore

NR = distance from anterior end to nerve ring

ES = distance from anterior end to end of esophagus


Mean (SD% RanICI












Table 2: Measurements (in um) of first and second
generation males of Steinernema scapterisci n.
sp. (n=10).


First generation Second generation

Character Mean (SD) Range Mean (SD) Range


Body length 1728 (358)

Greatest width 156 ( 49)

Stoma length 4.4 ( 1)

Stoma width 6.1 ( 1)

EP 71 (11)

NR 136 ( 11)

ES 187 ( 21)

Testis flexure 374 ( 52)

Anal body width 33 ( 5)

Tail length 25 ( 3)

Spicule length 83 ( 5)

Spicule width 13 ( 4)

Gubernac. length 65 ( 5)

Gubernac. width 8 (0.5)

EP:EF 0.36 (0.02)

Mucro length 4.3 ( 0.6)


1319-2271

97-231

3-5

5-8

63-98

120-152

164-216

306-447

31-45

21-30

72-92

13-14

59-75

8-9

0.32-0.39

3.1-4.7


1147 (95) 1031-1342

73 ( 8) 62-84

4.3 ( 1) 3-6

6.0 (1.2) 5-8

68 (7 ) 50-75

121 (10) 103-131

168 (13) 138-181

205 (19) 176-234

33 ( 4) 28-41

25 ( 3) 22-30

78 ( 3) 75-83

12 ( 1) 11-14

54 ( 3) 47-59

6 (0.7) 5-8

0.40 (0.06) 0.29-0.52

3.9 (0.6) 3.1-4.6


EP = Distance from anterior

EN = Distance from anterior

ES = Distance from anterior


end to excretory pore

end to nerve ring

end to end of esophagus










9

Table 3: Measurements (in um) of the third-stage juvenile
of Steinernema scapterisci n. sp. (n=20).


Character


Body length

Greatest width

EP

NR

ES

Tail length

EP:ES

EP:Tail length


EP = Distance from

NR = Distance from

ES = Distance from

esophagus


Mean

572

24

39

97

127

54

0.31

0.73


3

0.03

0.06


Range

517 609

18 30

36 48

83 106

113 134

48 60

0.27 0.40

0.60 0.80


anterior end to excretory pore

anterior end to nerve ring

anterior end to the end of



























Fig. 1. Steinernema scapterisci n. sp.. A) Face view
showing unevenly distributed papillae. B) Female
of the second generation, entire body. C) Double-
flapped epiptygma. D) Variation in tail of the
first generation females. E) Variation in tails of
the second generation female. F) Anterior part of
the first generation female showing large
cheilorhabdion and excretory duct with doughnut-
shaped structure.
















20pm


CEF 50Mm

D 100pm










12
with the body, and bears both labial and cephalic papillae.

There are six lips which are united at the base with each

terminating in a labial papilla. The six labial papillae

(Fig. 1A, 2A, 2B) are not evenly distributed when viewed en

face, while the 2 subventral and 2 subdorsal papillae are

located as expected, the 2 lateral papillae are located

lateroventrally making the ventral and lateral papillae

closer together than are the lateral and dorsal papillae

(Fig. 2A, C). The apex of each papilla is usually covered

with a thin layer of whitish material, (electron lucent)

(Fig. 2A, C). Cephalic papillae are present, appear to be 4

in number, but they are not distinct; therefore, the exact

number has not been determined. The amphids were not

observed. The stoma is very shallow, circular anteriorly,

then becomes subtriangular. The cheilorhabdions are

strongly sclerotized, unusually thickened, appearing as a

circular or hexagonal ring en face (Fig. IF, 2C, 3C). The

prorhabdions just posterior to the cheilorhabdions, also are

well-sclerotized. Posterior to this ring, no other

sclerotized structure was observed. The esophagus is

typical of the Steinernematidae, i.e. muscular throughout

with the procorpus followed by a slightly swollen,

nonvalvate metacorpus, isthmus, and basal bulb with a small,

but quite visible, valve. The nerve ring is distinct,

located in the region of the isthmus. The esophago-

intestinal valve is long and prominent (Fig. 1F). The



























Fig. 2. Steinernema scapterisci n. sp.. A) Anterior part of
the nematode showing the mouth, lips and excretory
pore. B) Head with 6 labial papillae. C) Face view
showing unevenly distributed labial papillae; the
mouth circular at the opening, becomes
subtriangular. Note the white material covering the
papillae. D) Face view showing the mouth and labial
papillae.















rC ;
~. ~.. ~1

~c~
~u'~;f~,: -'c~
~t~~3 ~.~
r~ ~c

1 C
rf ~





























Fig. 3. Steinernema scapterisci n. sp.. A) Posterior region
of the third-stage infective juvenile showing the
curvature. B) Live nematode showing doughnut-
shaped structure. C) Head of the nematode showing
thick cheilorhabdions. D) Spicules and gubernaculum
of male. E) Fixed nematode showing doughnut-shaped
structure.











































































I -


16

- -


$











excretory pore is located anterior to the middle of the

metacorpus, usually in the procorpus region. The excretory

duct is unusually prominent: it forms a small loop midway

between the excretory pore and the base of the esophagus,

then turns to the right side of the esophagus, or sometimes

extends to the anterior part of the intestine then returns

on the ventral side of the intestine at its junction with

the esophagus; here it coils upon itself 2 or more times

forming a doughnut-shaped structure (Fig. IF) complete with

a hole at the center. A uninucleate gland is located

posteriorly to the doughnut-shaped structure but a junction

of the excretory duct with the gland has not been observed.

This doughnut-shaped structure has been seen in almost every

first generation female and is visible even with a

dissecting microscope. The gonads are didelphic, opposed;

the ovaries are reflexed (Fig. 8B, E). The vulva appears as

a transverse slit with a prominent double-flapped epiptygma

(Fig. IC, 6C) The vagina is sclerotized; it's length is

about 1/3 of the body width at the vulva, and it leads to

paired uteri. The body width of the nematode anterior to the

vulva is always greater than that posterior to the vulva.

The tail is somewhat variable in shape, but usually has a

post-anal swelling ventrally and a mucron (Fig. lD); the

length of the tail is less than the width of the body at the

anus. The pigmy form referred to for other species

(Poinar, 1979) was not observed.









18
Female, second generation (Fig. IB, IE)

The second generation female is similar to the female

of the first generation, but differs in that the second

generation female is much smaller, the valve in the basal

bulb of the esophagus is more prominent, the doughnut-shaped

structure is not as prominent, the tail, which tapers to a

point bearing a mucron, is longer than the body is wide at

the anus (Fig. 1E).

Male, first generation (Fig. 4A, B, E)

The first generation male (Fig. 4E) is much smaller

than the first generation female, but anatomically the two

are similar anteriorly. The body is usually plump. The

nerve ring is located in the isthmus region but its exact

position is variable. The excretory duct does not form the

doughnut-shaped structure which occurs in females. The

posterior part of the nematode is curved ventrally. The

nematode body is spiral in shape when killed by minimal

heat. There is one gonad with a reflexed testis. The

spicules are paired, uniformly curved, dark brown in color

with the head large and somewhat angular (Fig. 4A, 4B, 3D,

5B). The angle formed by the shaft and blade of the

spicules averages 110 degrees (range 100-120). The shaft of

the spicules is long when compared to that of other species

and appears to be encased in a sheath (Fig. 5B, 6B). The

blade tapers smoothly to the end with the posterior portion

thinner than that in other species of Steinernema


























Fig. 4. Steinernema scapterisci n. sp. A) First generation
male tail with mucron; spicules with angular
head and ribs, gubernaculum with anterior portion
bent upward. B) Variation in tail shape of the
first generation male. C) Tail of the second
generation male showing elongate spicule head.
D) Variation in tail shape of the second
generation male. E) Entire body of the first
generation male.























































BD 50 pm




























Fig. 5. A) Steinernema scapterisci n. sp.. Spicule blade
showing the thin posterior part with a small
aperture. B) The spicule shaft showing the angular
head, and a sheath around the shaft. C) Cross
section of the spicule showing 2 lumens in the
spicule blade. D) Gubernaculum showing the long
anterior part which bends upward.










Ti*.


p F



























Fig. 6. A, B): Steinernema carpocapsae strain Breton.
A) Spicule showing the short shaft without a sheath
and with rounded head (compare with Fig. 5). B)
Gubernaculum showing short anterior part. C, D):
Steinernema scapterisci sp. C) Double-flapped
epiptygma. D) The body of the third-stage infective
juvenile showing annulation and lateral field with 6
incisures.







tf.




























Fig. 7. A) Steinernema scapterisci n. sp.. Posterior part
of the spicule, thin, with small aperture.
B, C, D) Posterior part of the spicule of S.
qlaseri, S. carpocapsae, and S. bibionis,
respectively, showing the difference in shape,
thickness and size of the aperture.












26







ll











(Fig. 4A, 4B, 7). In the cross section the blade of the

spicule contains two lumens (Fig. 5C), but only one aperture

was seen on the ventral side close to the tip (Fig. 7A).

This aperture is smaller than that on the spicule in other

species (Fig. 7). Under the compound microscope, each

spicule shows two internal ribs (Fig. 4A, 4B). The point

where the two ribs terminate proximally is variable. These

ribs are strengthening thickenings of the upper and lower

walls between the two lumens in the blade (Fig. 8F). The

gubernaculum is boat-shaped, with a thin, long and ventrally

curved anterior part. Compared to S. carpocapsae strain

Breton, the anterior part of the gubernaculum of S.

scapterisci is much longer (Fig. 5D, 6A). Its posterior end

is bifurcate (Fig. 5D). The spicules glide along the

gubernaculum in two grooves separated by a ridge (Fig. 5D).

The cloaca is on a raised area and bears an anterior flap,

seen easily when the spicules are protracted or retracted.

Ten pairs and one single genital papillae were observed

(Fig. 8A) with pairs 1 and 6 difficult to see. The single

papilla is located ventrally and between pairs 4 and 5;

pairs 1-9 are located ventro-laterally and pair 10

subdorsally. The tail bears a mucron, and the posterior

region is always curved ventrally (Fig. 4A, B, E).

Male, second generation

The second generation male is similar to that of the

first generation but is smaller, especially in width, and































E
S 500pm

A 50Pr
B 120pm F
CD 20pr







Fig. 8. Steinernema scapterisci n. sp.. A) Male tail of the
first generation showed 10 pairs of genital
papillae. B) Flexure of the ovary reflexedd in the
body). C, D) Head and tail of third stage infective
juvenile. E) Ovary with spermatheca reflexedd in the
body). F) Cross section of spicule blade showed two
lumens (1) and 2 internal ribs (r).









29

the spicules have an elongate head (Fig. 4C, 4D).

Juveniles, third stage (Fig. 8C, 8D)

Measurements are given in Table 3. The third stage

juvenile, when newly formed, is always enclosed in the

cuticle of the second-stage juvenile as a sheath. However,

the sheath is lost rather easily, even in storage, and thus

may not always be present. The body is thin. The lip

region is not offset; the oral aperture (mouth) was not

observed; the esophagus is degenerate and thus not seen

clearly, but its basal bulb is elongate and has a valve.

The lateral fields have 6 incisures (Fig. 6D). The tail

tapers gradually dorsally but abruptly ventrally (Fig. 8D).

Cross hybridization

In cross hybridization experiments, males and females

never mated and thus no offspring were present. In the

controls, males and females mated and offspring were present

after 10 days.

Biology

All species of Steinernema, except S. scapterisci, and

all strains of S. carpocapsae, killed from 20 100% of the

insects tested. S. scapterisci killed no more than 10 %

(Table 4). The large difference in the percentage of wax

moth larvae killed by all nematodes except S. scapterisci is

significant since the wax moth larva has been used as a

universal host for in vivo culture of species and strains of

Steinernema. This difference indicates that wax moth larvae












Table 4: Effectiveness of different species of Steinernema
and strains of S. carpocapsae in killing four
species of lepidopterous insects, two days after
inoculation.


Percentage of insects killed*

Nematode FAW VBC GCW WML

S. qlaseri 100 90 50 100

S. bibionis 100 90 55 100

S. carpocapsae

Breton 100 100 -- 100

Italian 100 100 -- 100

Mexican 100 100 80 100

Agriotos 100 100 20 100

All 100 100 -- 100

S. scapterisci 8 3 10 9

Control 0 0 0 0


* Average of four trials

FAW = fall army worm; VBC
GCW = granulate cut worm;


= velvet bean caterpillar;
WML = wax moth larva.









31

can be used as test insects to differentiate between S.

scapterisci and all other species and strains of

Steinernema.

Diagnosis

Steinernema scapterisci n. sp. can be distinguished

from other species of Steinernema as follows: from S.

qlaseri, by the presence of a mucron on the tail of the male

of S. scapterisci, and by the shorter third-stage juvenile

of S. scapterisci (517-609 um) than of S. glaseri (860-1500

um); from S. bibionis and S. intermedia by the shorter

third stage juvenile (700-1000 um for S. bibionis and 608-

800 um for S. intermedia); from S. carpocapsae by the ratio

of head to excretory pore divided by tail length, this ratio

is 0.73 (0.60-0.80) in S. scapterisci compared to 0.60

(0.54-0.66) in S. carpocapsae; and by the shape of the tail

of the third stage juvenile; when relaxed, the tail of S.

scapterisci usually curves ventrally forming an angle about

110 degree with the body. The ratio of the head to excretory

pore/head to end of esophagus is 0.38 compared to 0.90 in S.

glaseri, 0.59 in S. bibionis, 0.72 in S. intermedia, 0.43 in

the Czechoslovakian and DD-136 strains of S. carpocapsae.

S. scapterisci n. sp. also can be separated from all

other species by the following characters: The presence of

thick cheilorhabdions (about 4.8 um thick by 5.8 um long in

lateral view), the doughnut-shaped structure in the

excretory canal, and a prominent double-flapped epiptygma in









32
the 1st generation female. Spicules of the male are brown,

pointed, and taper smoothly to the end; end of the blade

narrow; shaft long and bearing a sheath; gubernaculum with

long and upward-bent anterior part.

S. scapterisci n. sp. cannot be cultured on wax moth

larvae (Galleria mellonella), but sometimes a few wax moth

larvae will be killed by the nematode. When this occurs,

the bodies of the wax moth larvae turn black while those

killed by other species of Steinernema turn whitish or

yellowish but never black. Also, other species of

Steinernema develop very well in wax moth larvae. Finally,

this nematode can be distinguished from other species by

bioassay on 3 insects: fall army worm, velvet bean

caterpillar, and wax moth larvae. In two days, other species

of Steinernema will kill 100% of the test insects, but S.

scapterisci will kill at most only a small percentage of

them (Table 4).
















CHAPTER 3
MODE OF ENTRY OF STEINERNEMA SCAPTERISCI N. SP
INTO MOLE CRICKETS


Animal parasitic nematodes, in general, enter their

hosts in one of three ways: (1) by direct penetration

through the cuticle, (2) through natural openings or wounds

or (3) by vectors. While the first two apply to insect-

parasitic nematodes, transmission by vectors has not been

reported. Most of the insect-parasitic nematodes in the

orders Tylenchida and Mermithida are reported to enter the

hosts by direct penetration, while those in the order

Rhabditida enter the hosts through natural openings or

sometimes by direct penetration through the cuticle. Most

of the information on the rhabditid genus Steinernema has

been collected over a long period of time on the species S.

carpocapsae. Since S. scapterisci is described herein as a

new species, the following experiments were conducted to

determine its mode of entry into its primary hosts, mole

crickets in the genus Scapteriscus.


Literature Review


Bronskill (1962), and Welch and Bronskill (1962)

reported that the infective stage of S. carpocaosae, strain










34
DD-136, entered the body of mosquitoes passively through the

mouth. Weiser (1966) reported that S. carpocapsae entered

the body of some insects through the spiracles and the

tracheal system. Poinar and Himsworth (1967) observed

steinernematids in the crop and mid gut of wax moth larvae

(Galleria mellonella) and concluded that the nematode

entered through the mouth of the insect. Friggiani and

Poinar (1976) showed that S. carpocapsae entered the body

of some adult insects in the order Lepidoptera through the

spiracles.


Materials and Methods


The mole cricket, Scapteriscus acletus, was used in the

following experiments.

Experiment 1

The purpose of this experiment was to determine whether

S. scapterisci entered mole crickets through either the

mouth or anus. Three hours after 5 mole crickets were

exposed to 8,000 infective-stage nematodes in a petri dish,

the mole crickets were killed, the digestive system

dissected out and cut into 3 parts. The first part included

the mouth through the crop; the second part began just

posterior to the crop and continued to the junction of the

Malpighian tubules with the gut; the third part was from

this junction to the anus. Each part was dissected in a

separate petri dish and examined for nematodes.











Experiment 2

The purpose of this experiment was to determine whether

the nematode can kill mole crickets and develop in their

bodies when the crickets were inoculated through the mouth

or through the anus. A small-diameter hypodermic needle

attached to a 1 ml syringe was used as the inoculation

vehicle (Fig. 9). The angled tip of the needle was removed,

the end filed smooth and the syringe mounted on a stand to

facilitate the inoculation process. Twenty mole crickets

were deprived of food and water over night, and then ten of

them were inoculated through the mouth and 10 through the

anus with about 80 third-stage juveniles in 1/10 ml of

water. After they died, the cadavers were incubated, each

separately, to determine if the nematodes reproduced in

them.

Experiment 3

The purpose of this experiment was to determine if the

nematode can enter the mole cricket through the spiracles.

Several hours after the mole crickets were inoculated with

nematodes, as in Experiment 1, the tracheal system (as much

as possible) was dissected out carefully from about 20 mole

crickets and observed under a compound microscope.

Experiment 4

The purpose of this experiment was to determine if the

nematode would kill mole crickets and develop in their

bodies when inoculated through the spiracles. The first and





















Fig. 9. Inoculating a mole cricket through the mouth
with third-stage Steinernema scapterisci.






















Fig. 10. Third-stage nematodes Steinernema scapterisci in
tracheal tube of mole cricket. The part of the
tracheal tube above the nematode was obvious to be
broken by the nematodes from the main tube
(nematodes inside).




































O
O


. o oo


C~b~Y "
~. 0~
~











second thoracic spiracles of mole crickets are large enough

to be used as inoculation ports when using a very small

needle. Thus the same needle and syringe mounted on a stand

that was used in Experiment 2 was used here. Fifteen mole

crickets were inoculated through the first thoracic spiracle

and 4 were inoculated through the second thoracic spiracle

as follows: About 50 nematodes were collected in a droplet

of water at the end of the hypodermic needle. Then, while

viewing through a stereomicroscope, the mole cricket was

squeezed slightly to expel the air in the tracheal system,

and the droplet of water containing the nematodes was

applied to the spiracle and the pressure on the body of the

mole cricket released. The water was sucked through the

spiracle and into the tracheae.

Three days after the mole crickets died, their bodies

were dissected to see if developing nematodes were present.

Experiment 5

The purpose of this experiment was to see if the

nematodes would enter the tracheal system when inoculated

through the spiracle. Ten mole crickets were inoculated

through the first and second thoracic spiracles as in

Experiment 4; all of the crickets were dissected 30 minutes

later and the tracheal system examined for nematodes.










39

Results and Discussion


Experiment 1

When the three parts of the digestive system were

dissected and examined for nematodes, with an average of

five digestive systems examined, 66 nematodes were found in

the anterior part, none in the middle part, and one in the

posterior part.

Since by far the greatest number of nematodes were

found in the anterior part of the digestive system, it seems

that the majority of nematodes enter mole crickets through

the mouth. This agrees with the mode of entry reported by

Poinar and Himsworth (1967) for wax moth larvae. Since one

nematode was found in the posterior part of the digestive

system it is possible that some of the nematodes enter

through the anus.

Experiment 2

Nine of the ten mole crickets inoculated through the

mouth produced large numbers of nematodes one week after the

mole crickets died while none of those inoculated through

the anus produced nematodes after the crickets died. This

suggests that nematodes that enter mole crickets through the

mouth penetrate the gut wall and entered the thoracic cavity

where they reproduce. Any that enter through the anus may

die in the rectum or they may penetrate the gut wall and

entered the abdominal cavity. Previous experiments have











shown that the nematode does not develop in the abdominal

cavity.

Experiment 3

Nematodes were found several times in the tracheal

tubes of mole crickets, mostly in the thoracic area, and

especially in the pronotal region. A few of the nematodes

were observed to break through the wall of the tracheal

tubes (Fig. 10).

Experiment 4

Three of the 4 mole crickets inoculated with nematodes

through the second thoracic spiracle, and 12 of the 15

inoculated through the first thoracic spiracle were found

with developing nematodes in the head and thorax. Since in

vivo culture (See Chapter 5 on culture) showed that most of

the nematodes were produced in the pronotal region, and

since the first thoracic spiracle is located underneath the

pronotum, the first thoracic spiracle may be a very

important entry route for the nematode.

Experiment 5

Nematodes were found in the tracheal system of all mole

crickets inoculated. The nematodes found in very small tubes

with a diameter barely greater than that of the nematode,

thrashed about vigorously, but did not break through the

tracheal tubes during the period of observation.









41

Experiments 3, 4 and 5 demonstrated that the nematode

can kill mole crickets by entering the spiracles, breaking

through the tracheal wall and invading the body cavity.

Thus the spiracles may represent a very important port of

entry for the nematode.
















CHAPTER 4
LIFE CYCLE OF STEINERNEMA SCAPTERISCI N. SP.


Information about the life cycle of a parasitic

nematode is always helpful, but especially so when one wants

to use the nematode as a biological control agent. Such

information is essential to know what life stage to apply in

the field and often gives some indication of the best method

of application. It is useful also in conducting research to

increase the effectiveness of the nematode against the

target pest or group of pests and in developing or improving

efficient and low-cost mass-rearing techniques. The life

cycle of the nematodes Steinernema bibionis, S. carpocapsae,

and S. glaseri were studied by Bovien, 1937, Wouts, 1980 and

Poinar, 1979. In this chapter, I report the results of my

studies on the life cycle of S. scapterisci.


Materials And Methods


Mole crickets were not available for these studies so

house crickets, which react similar to mole crickets in

pathogenicity tests, were used in all experiments. The

house crickets were purchased from a local bait and tackle

shop.








43

Influence of Temperature on the Life Cycle

Fifty house crickets were anesthetized with carbon

dioxide, placed in each of 10 petri dishes (100 x 15 mm) and

exposed to about 8,000 third-stage juvenile nematodes. Two

of the petri dishes were placed in incubation chambers at

10, 15, 20, 24 and 30 C. Starting two days later, up to 3

crickets were dissected daily and the life cycle stage of

the nematode determined.

Intermediate Cycle

At 24 C, male and female nematodes were detected in the

bodies of house crickets. In one experiment with 3

replicates, a small piece of tissue from the thorax (1/2 of

the pronotum) of the cricket was placed on a small piece of

filter paper (1 cm2) saturated with water in a 60 x 15 mm

petri dish. Two gravid nematode females were transferred to

each piece of tissue. Four drops of water were placed in

the petri dish surrounding the filter paper to insure

adequate moisture in the dish. Covers were applied and the

dishes placed in the dark and examined daily. The

experiment was repeated twice. In a second similar

experiment larger pieces of insect tissue (the entire

pronotum) were used.

The reason for using a small piece of tissue was to

determine whether the nematodes, when provided with a

limited food supply, would cease development at the third

stage, engulf and store a pellet of bacteria in the foregut









44
and become third-stage infective juveniles. The reason for

the larger piece of tissue was to determine whether the

nematode, when provided with a sufficient food supply, would

continue to develop to adults instead of becoming third-

stage infective juveniles.

Sex Ratio

Ten to twelve house crickets were placed in a petri

dish and exposed to about 8,000 third-stage juveniles. The

dishes then were placed in an incubator at either 15, 24 or

30 C. After 3 days, all crickets were dissected and the

number of male and female nematodes in each cricket were

counted.


Results and Discussion


Influence of Temperature on the Life Cycle

At 10 C: The third-stage infective juveniles never reached

the adult stage. A few began to develop to the 4th stage

but had poorly-developed, shortened, plump bodies, and all

died within 5 days.

At 15 C: The third-stage infective juveniles developed into

males and females of the first generation after 10 days;

they were in the thoracic cavity. The second generation

juveniles appeared about 15 days after inoculation. Most,

but not all, moved to the abdominal cavity and embedded

themselves in the fat tissue lining the abdominal wall.

These nematodes moved very slowly, many appearing immobile.









45
Until the 18th day, the second generation juveniles had not

increased in size and they became completely immobile. No

third-stage infective juveniles were produced.

At 20 C: First generation males and females were observed

after 7 days, but many of them died, and those that were

alive were not very active. The second generation adults

appeared after 8-9 days; after 10-11 days, some infective-

stage juveniles were observed with increasing numbers

appearing a day later.

At 24 C: First generation adults appeared in less then 3

days. Most of them were in the anterior part of the cricket

cadaver After 4 days, most of the first generation adults

had decayed but a few females containing second generation

juveniles remained. After emerging from the body of the

first generation females, the first-stage juveniles of the

second generation migrated to all parts of the cricket

bodies. During the 5th and 6th days, the second generation

juveniles grew rapidly, and after 7 days, infective third-

stage juveniles, preinfective second-stage juveniles and

second generation adults were seen inside and outside the

cricket bodies. After 8-9 days preinfective second-stage

juveniles and infective third-stage juveniles appeared

increasingly abundant. In the preinfective second-stage

juvenile the esophagus and stoma are not well-developed and

the chamber containing a pellet of bacteria is forming.

When it molts to the third-stage infective juvenile, the











retained as a sheath. At day 10, infective third- stage

juveniles were present throughout the inside and outside of

the cadavers.

At 30 C: First generation adults were observed in 2 days,

and second generation adults in 5 days. Third stage

infective juveniles first appeared at 6 days, the number

increased at 8 days and large numbers appearing at 10 days.

Intermediate Cycle: On the isolated small piece of tissue,

some third stage infective juveniles were observed after 6

days, and many of them appeared after 7 days. None

continued to develop to second generation adults. In the

dish with a larger piece of tissue, some of the juveniles

stopped development and became third stage infective

juveniles but others continued to grow and became adults.

These observations show that third-stage infective

juveniles may be produced by the first generation adults or

by the second generation adults. In this study, those that

were produced by the first generation adults appeared in 6-7

days. With the small piece of tissue, apparently an

insufficient supply of food prevented the juveniles from

becoming second generation adults, so they ceased

development and became third-stage infective juveniles

presumably as a survival mechanism.










47

Table 5: Influence of temperature on the life cycle of
Steinernema scapterisci.


Life cycle stage 10 C 15 C 20 C 24 C 30 C


1st generation adults

2nd generation adults

Intermediate life cycle

Regular life cycle


No. of days

S 10

S NC

S NC


NC 12 10 8


NC = Not completed


appearance


to 1st

7

8


10 6-7









48
Even when adequate food was available, some of the

juveniles wandered away from the food supply and became

third-stage infective juveniles from the first generation

adults, but others developed to second generation adults and

then produced third-stage infective juveniles in 9-10 days.

The regular cycle took 9-10 days at 24 C.

Whether or not S. scapterisci completed the life cycle,

and the length of time required for that cycle when it was

completed, was influenced by temperature. Temperatures

colder than 20 C are not suitable for development (Table 5).

Life Cycle of S. scapterisci n. sp.

The life cycle of S. scapterisci is schematically

presented in Fig. 11 and described as follows: The third-

stage infective juveniles invade the host through the mouth

or spiracles; some may enter through the anus but they fail

to develop (on mole crickets). Immediately after entering

the body cavity of the host, the juveniles accumulate in the

thorax and head and start to develop. The nematode enlarges

in width, the stoma opens and its walls thicken, the

esophagus becomes prominent, the bacterial chamber enlarges

and moves posteriorly releasing the bacteria into the

intestinal lumen, and then into the blood stream or tissues

of the host. This process takes less than 24 hours after

the nematode enters the host. The nematode grows rapidly

and feeds on bacterial cells which develop in the host and

becomes the fourth stage. First the body width increases


































5 days J









6 days


Fig. 11. Diagram of the life cycle of Steinernema
scapterisci n. sp..











much faster than does the body length until its width

reaches almost the width of the adult; the excretory duct

becomes larger and complicated; then the nematode increases

in length until the adult size is reached. During this time

the reproductive system is also formed. It takes 60 to 72

hours to reach the first generation adult stage at 24 C and

just 48 hours at 30 C, but up to 240 hours at 15 C and 168

hours at 20 C. The female becomes about 7 times longer and

7 times wider than the third stage infective juvenile. The

sex ratio is dependent upon temperature and density of

nematodes in the host. At 15-24 C the number of females is

greater but at 30 C the number of males is greater. Males

and females mate and the females initially lay eggs, but

later the eggs are retained and hatch in the female's body.

The nematodes reproduce bisexually. About 24 hours after all

eggs hatch in the body of the female, the second generation

first-stage juveniles break out of the female body and move

into the body cavity of the insect. At that point, an

individual may develop by one of two cycles depending upon

the availability of nutrients and space. If nutrients are

insufficient and/or space is limited (overcrowding), a

number of the second generation second stage juveniles

become third-stage infective juveniles in about 6-7 days

after the nematodes enter the body of the host. The

remaining second generation second-stage juveniles continue

to develop to become second generation females and males











which are much smaller than females and males of the first

generation. These nematodes appear at about 7 days, almost

the same length of time required to become infective-stage

juveniles in the intermediate cycle. These males and

females mate and the females lay some eggs which hatch as

first stage juveniles. A number of eggs are not laid,

however, but hatch in the female's body. These first-stage

juveniles break out of the female's body after she dies to

enter the body cavity of the host. The juveniles develop to

the second stage and then to the third stage infective

juveniles in 9-10 days after inoculation. They emerge from

the host in large numbers and enter the environment to seek

out a new host.

Sex Ratio: The results of the sex ratio experiments are

presented in Tables 6, 7 and 8.

The greatest average number of 1st generation females

and males, 257, (Table 6) were produced in the house

crickets at 15 C. This number was approximately 2 and 4

times as great as the numbers produced at 24 and 30 C

(Tables 7 and 8). This was surprising because the life

cycle was not completed at 15 C (Table 5).

One reason for this seemingly unusual situation could

be that at the lower temperature, a greater number of third

stage infective juveniles entered the house crickets because

of reduced activity of the crickets which had been living at

ambient temperatures. Conversely, the activity of the












Table 6: Number and percentage of first generation
females and males of Steinernema scapterisci
which developed at 15 C.


No. % No. %

Rep Females Females Males Males Total

1 171 60 114 40 285

2 204 50 202 50 406

3 231 59 159 41 390

4 102 48 109 52 211

5 113 55 93 45 206

6 126 55 104 45 230

7 61 56 47 44 108

8 136 53 123 47 259

9 182 59 125 41 307

10 66 40 100 60 166

Means 139 54a 118 46b 257



Numbers with different letters in the same row are
significantly different at the 5% level according to
Duncan's multiple range test.












Table 7: Number and percentage of first generation
females and males of Steinernema scapterisci
which developed at 24 C.



No. % No %

Rep Females Females Males Males Tot


1

2

3

4

5

6

7

8

9

10

11

12

Means


227

80

60

179

77

37

29

83

71

66

232

96

104


al


Numbers with different letters in the same row are
significantly different at 5% level according to
Duncan's multiple range test.










54

Table 8: Number and percentage of first generation
females and males of Steinernema scapterisci
which developed at 30 C.


Rep

1

2

3

4

5

6

7

8

9

10

11

12

Means


No

Females

10

6

23

7

47

65

63

23

40

44

23

41

32


%

Females

56

55

53

44

52

49

51

51

47

47

72

27

47a


No.

Males

8

5

20

9

43

69

60

22

46

50

9

102

36


%

Males

44

45

47

56

48

51

49

49

53

53

28

73

53a


Total

18

11

43

16

90

134

123

45

86

94

32

143

68


Numbers with the same letters
significantly different at 5%
multiple range test.


in the same row are not
level according Duncan's









55
nematodes would have increased at the warmer temperature

because those used for these experiments had been stored in

a refrigerator at 6-10 C.

If this is the case, this information may be useful for

in vivo culture for producing greater populations of third-

stage infective juveniles. First, infective-stage juveniles

stored at 6-10 C and the host crickets kept at ambient

temperatures would be placed in an inoculation chamber at 15

C. After two days when the house crickets were dead, they

would be transferred to an incubation chamber at 24 C.

This technique should allow total populations of 1st

generation adult females to develop at the cooler

temperature and then more third-stage infective juveniles to

develop at the warmer temperature.

When the experimental temperature was increased to 24 C

and to 30 C, the number of first generation adults which

developed in house crickets was reduced. This might be

explained by the higher temperature causing increased

activity of the house crickets which might have caused fewer

infective-stage juveniles to enter the house crickets. It

is possible also, that at the higher temperatures,

especially at 30 C, water in the incubation chamber

evaporated more quickly and after some length of time, the

nematodes may be inactivated because of the lack of

humidity.











At 15 C: The first generation female:male ratio was 54:46.

The nematodes did not develop beyond this stage.

At 24 C: the first generation female:male ratio was 60:40.

When the numbers of adult females and males in a house

cricket were less than or equal to 83 the average

female:male ratio was 69:31 (calculated from Table 7). In

some individual crickets, the female:male ratio was as high

as 76:24 (one cricket), and 70:30 (3 crickets). Regression

analysis suggested that the hypothesis, percentage of males

is a function of the total number of nematodes in house

crickets, is highly significant, P = 0.0003 (Fig. 12). This

implies that nutrients influence the sex ratio.

At 30 C: the first generation female:male ratio was 47:53.

The linear relationship between the number of males and the

total number of nematodes in house crickets was highly

significant, P = 0.0001 (Fig. 13).











48




32
LU
.J

216
o1


P=0.0003


80 160
S + FEMALES/


240
CRICKET


Fig. 12. The relationship between the percentage of males
and the total number of nematodes in each house
cricket at 24 C.


Y= 27.6 +
R2=0. 6


0.08X


0
MALE











1201 Y


Y =-5.31 + 0.6 X
R2 =093
. 80 P 0.0001



Y:-11.4 -1
w3 R2= 0.98
I P= 0.00(

640
z








0 40 80 120 180 2(
No. MALES+ FEMALES/CRICKET




Fig. 13. The relationship between the number
and the total number of nematodes in
cricket.


- 0.5X

)1


of males
each house















CHAPTER 5

CULTURE OF STEINERNEMA SCAPTERISCI N. SP. IN VIVO



Maintaining a continuous supply of nematodes for

research purposes is one of the most important

considerations in studying the effects of nematodes on mole

crickets. This need led to the development of different

methods for producing quantities of the nematodes sufficient

for research purposes. Depending upon the number of

nematodes needed, and other considerations, they were

cultured either in vivo or in vitro. This chapter covers

only in vivo culture.

In vivo culture in the wax moth larva, Galleria

mellonella, has been used by researchers for many years.

Since S. scapterisci reproduces very poorly or not at all in

wax moth larvae, mole crickets have been used as the in vivo

host to maintain populations and strains of the nematode, to

insure that the nematode retains the ability to kill mole

crickets, and to produce numbers sufficient for small-scale

experiments. The purpose of this chapter is to present the

methods used to culture S. scapterisci in mole crickets and

to evaluate other techniques in order to improve production.










60

Literature Review


The culture of nematodes in vivo was first carried out

by Dutky and Hough (1955) when they grew what is now the DD-

136 strain of Steinernema carpocapsae in larvae of the

greater wax moth, Galleria mellonella. Dutky et al. (1964)

reported in detail their culture techniques in which wax

moth larvae were inoculated with infective stage juveniles

of the nematode in petri dishes, then, when the wax moth

larvae died, incubated the cadavers at 24 C. The nematodes

which emerged from the cadavers were trapped in a tray

containing a solution of 1 part formaldehyde and 1,000 parts

water (1/1,000). Each wax moth larva produced up to 200,000

infective-stage juveniles. The juveniles harvested were

stored in a flask at 7.1 C in oxygenated 1/1,000

formaldehyde.


Materials and Methods


In Vivo Culture in Mole Crickets

Two pieces of filter paper, Whatman No. 2, are placed

in the bottom of a petri dish (15 x 100 mm) and 5,000 -

20,000 infective-stage juveniles in water added. Mole

crickets are anesthetized by carbon dioxide, and then placed

in the dish (Fig. 14). The number of mole crickets per dish

varies depending upon their availability and the purpose of

the inoculation. The lids are applied and the petri dishes










61
placed in the dark at room temperature (25 C). After 2-3

days, the dead mole crickets are washed with water to remove

any phoretic nematodes and placed on a filter paper in

incubation chambers (Fig. 14). The incubation chambers

consist of a large petri dish (20 x 150 mm) containing an

inverted lid of a small petri dish (15 x 60 mm) on which a

piece of filter paper (90 mm diameter) is placed. Water is

added to the large petri dish to reach and wet the filter

paper. The mole cricket cadavers are arranged on the filter

paper with the heads directed outward. The dishes are

incubated at 24 C. After 5-7 days, third generation third

stage infective juveniles migrate from the cadavers to the

filter paper and into the water (Fig. 15). The nematodes

are harvested by collecting them on a fine mesh sieve,

openings 20-24 um, washing them onto a filter (dust mask)

and allowing the active nematodes to migrate through the

filter into clean water. The nematodes are used immediately

or stored under aeration from an aquarium pump at 6-12 C for

several weeks. This method of production takes 7-10 days to

produce infective stage juveniles, and yields 20,000 -

80,000 per mole cricket.

Experiment 1

The purpose of this experiment was to determine whether

rates of inoculum greater than 20,000 infective stage

juveniles would yield greater numbers of infective stage

























































Fig. 14. In vivo culture of the nematode Steinernema
scapterisci n. sp. Top: Live mole crickets placed
in an inoculation chamber with third-stage juvenile
nematodes. Bottom: Dead mole cricket arranged in an
incubation chamber.
















-7 7W1


-a r


A .-g


Fig. 15. Top: Nematodes which developed within the bodies
of mole crickets have migrated to the exterior.
Bottom: Third-stage juveniles have migrated from
the mole crickets into the water in the incubation
chamber.









64

juveniles than would 20,000 or less. Inoculum levels of 5,

10, 20, 40, 60, 80, 100 and 120 thousand infective-stage

juveniles were released into each of 8 petri dishes prepared

as reported above. Five anesthetized mole crickets were

placed in each dish and stored in the dark at room

temperature (25 C). After the mole crickets died, they were

washed and placed in incubation chambers. The dishes were

examined daily.

Experiment 2

The purpose of this experiment was to determine whether

the infective-stage juveniles reproduced equally well in all

parts of the mole cricket. Ten mole crickets were

inoculated with nematodes as above. Two days after the mole

crickets died, their bodies were separated into head,

thorax, and abdomen, and each part dissected and examined

for developing nematodes.

In Vivo Culture in House Crickets

Since mole crickets are not available during the winter

months, another good host was sought. An alternate host, to

be effective, must produce a large number of infective stage

juveniles and the nematodes obtained must have similar

virulence to the target host as those produced in mole

crickets. Other experiments (Chapter 9) showed that S.

scapterisci killed a high percentage only of mole crickets

and house crickets, Acheta domestic. House crickets were

used in the experiments reported below to determine whether










65
they would be a satisfactory host in which to produce the

nematode.

Experiment 3

Six house crickets were placed in each of two 15 x 100

mm petri dishes prepared as reported above. After 4 days,

11 of the 12 house crickets died. The cadavers were washed

by shaking them vigorously in a container of water, and then

incubated as in the rearing method for mole crickets. Third

stage juveniles which emerged from the house cricket

cadavers were referred to as the HI population. The H1

population was cycled through house crickets again to yield

the H2 population and the H2 population in turn yielded the

H3 population. The number of infective stage juveniles

produced per house cricket were counted.

Experiment 4

In order to compare the effectiveness of nematodes

produced through three cycles in house crickets, population

H3, with nematodes produced in mole crickets, population MC,

the following experiment was carried out.

Ten petri dishes were prepared as in Experiment 1;

8,000 MC infective-stage juveniles were released in each of

5 dishes, and 8,000 H3 infective-stage juveniles were

released in each of the other 5 dishes. Mole crickets were

anesthetized and 5 placed in each dish. The lids were

applied to the dishes and taped to the bottoms. All dishes









66

were kept in the dark at room temperature (25 C). The

number of dead crickets was reported after 2 and 4 days.


Results and Discussion


In Vivo Culture in Mole Crickets

Experiment 1

At inoculum rates of 5,000-20,000 nematodes per dish of

5 mole crickets, yields of infective-stage juveniles ranged

from 20,000-80,000. At inoculum rates of 40,000-120,000

nematodes per dish of 5 mole crickets, a large number of the

first generation females, due to overcrowding apparently,

emerged from the mole cricket cadavers 4 days after

inoculation. The majority of these females moved down into

the water in the incubation chamber and many of them died

without reproducing. The bodies decayed very quickly and

their decay, plus the decay of the mole cricket cadavers,

gave off a strong odor of ammonia from the incubation

chambers. These decaying bodies rendered the water toxic to

the infective stage-juveniles when they moved into it.

Other females which contained active juveniles in their

bodies, moved down into the water. These females produced

uninfective second stage juveniles which died very quickly

in storage.

When the situation described above occurs during

rearing, the mole cricket cadavers should be rinsed with

water and all of the nematodes and water in the dish must be









67

discarded and clean water added. This process must be

repeated until no, or very few, adult nematodes move into

the water.

Experience showed that only those females on the filter

paper of the incubation chamber, or on or in the cadavers of

the mole crickets produced infective-stage juveniles.

Considering the above problems encountered when 40,000

or more infective stage juveniles per 5 mole crickets were

used as inoculum, it is obvious that those rates were too

high for optimum rearing. One may want to use such high

rates, however, when the first generation females are needed

to obtain juveniles for axenic culture.

Experiment 2

When the head, thorax and abdominal sections of the

mole cricket were examined, developing nematodes were found

mainly in the head and the thorax, especially in the

pronotum; very few were found in the abdomen. These results

indicate that the abdominal cavity is not very suitable for

development of the nematode.

Producing the nematodes on mole crickets has some

disadvantages. Mole crickets, being soil-dwelling insects,

carry many different microorganisms on and in the body;

these develop quickly after the mole crickets die. Also,

the decaying mole cricket cadavers, the dead bacteria and

decaying nematode bodies in the dishes cause a high death

rate of the juveniles. The nematodes harvested usually were









68
contaminated with other bacteriophagous nematodes such as

diplogasterids and rhabditids, sometimes in very high

numbers. These disadvantages can be ameliorated, but not

eliminated, by forcing the mole crickets to swim in a

container of water for 2 minutes before inoculating them.

Immediately after the inoculated mole crickets die, their

wings must be removed, and the mole crickets washed by

shaking them in a container of warm water (40 C) for 10

seconds. Then the posterior 2/3 of the abdomen must be

removed to reduce the amount of body fluids in the

incubation chamber (the abdomen does not support the

development of nematodes as mentioned above). The nematodes

then may be placed in the incubation chamber.

In Vivo Culture Using House Crickets

Experiment 3

It took the same length of time, 7-10 days, to produce

infective-stage juveniles in house crickets as in mole

crickets. Each house cricket can produce from 20,000-

60,000 infective-stage juveniles. Dissection of cadavers of

house crickets showed that the nematodes were present in all

parts of their bodies. Because of this distribution,

reproduction was not much less than that in mole crickets

even though the house cricket is considerably smaller.

Experiment 4

The nematode, H3, produced from three cycles through

house crickets, killed 22 of 25 mole crickets in 2 days and










69
24 of 25 in 4 days. The nematode from mole crickets, MC,

killed 23 of 25 mole crickets in 2 days and 25 of 25 in 4

days (Table 9). These results indicate that nematodes

cycled through house crickets 3 times do not lose their

pathogenicity, but kill mole crickets as readily as do those

produced in mole crickets.

Both house crickets and mole crickets have been used

for in vivo culture of S. scapterisci at the Nematology

Laboratory, University of Florida. The use of house

crickets has the advantage that they are readily available

throughout the year, but the disadvantage that somewhat

fewer infective stage juveniles are produced per house

cricket than per mole cricket. The house cricket also may

serve as a good test insect in lieu of the mole cricket

because of its availability, and because it is killed by S.

scapterisci as readily as is the mole cricket. It may

serve, also, as an excellent host to test the LD50 and LC50

of the nematode.









70

Table 9: Numbers of mole crickets that died after 2 and 4
days when using 8,000 Steinernema scapterisci
strains H3 and 8,000 strain MC Steinernema
scapterisci in petri dishes containing mole
crickets.


No. mole crickets killed/total

Rep 2 days 4 days



H3 MC H3 MC

1 5/5 5/5 5/5 5/5

2 5/5 5/5 5/5 5/5

3 4/5 5/5 5/5 5/5

4 4/5 3/5 5/5 5/5

5 4/5 5/5 4/5 5/5

Total 22/25 23/25 24/25 25/25


H3 = Nematodes
3 cycles.
MC = Nematodes


produced

produced


from house crickets after

from mole crickets.















CHAPTER 6
CULTURE OF STEINERNEMA SCAPTERISCI N. SP. IN VITRO


In vivo culture of entomogenous nematodes is

satisfactory for maintaining strains or species, and for

producing moderate numbers for small-scale experiments, but

when large numbers are needed, as for field experiments, in

vitro culture must be used. In vitro culture has been used

since 1932 with several different methods reported. In this

chapter are presented the different methods used to produce

Steinernema scapterisci.


Literature Review


Glaser (1932) was the first to culture an entomogenous

nematode (Neoaplectana qlaseri) in vitro. He used veal

infusion agar, dextrose and baker's yeast. According to his

report, yeast furnished some nutritional elements found in

the insect host. While his method produced a moderate

number of nematodes, it did not yield quantities sufficient

for field release. McCoy and Glaser (1936) reported an

improved technique by using fermented potato mash with a

mixture of two thirds Irish potato and one third sweet

potato. They believed that sweet potato enhanced the growth

of yeast. The medium was mixed with a pure culture of yeast











grown on sweet potato gruel. Then a 2-cm layer of the

mixture was spread on a tray. After 20-40 hours of

fermentation, the nematodes were added on top. This method

produced about 4 million nematodes per 2,000 cm2 of surface.

A further improvement was described by McCoy and Girth

(1938) who used ground, extracted veal pulp. This method

consisted of grinding fresh veal through a food chopper,

then infusing it with water. After 2 days the infusion was

poured onto a flannel cloth, drained, and the pulp squeezed

as dry as possible. After adding water and a preservative

(0.06% formaldehyde and 0.05% of the sodium derivative of

methyl-hydroxybenzoate) the medium was spread in a thin

layer and inoculated with nematodes. The authors said that

this diet produced larger and more robust nematodes and the

yield was 9,000-12,000 nematodes per cm2 of culture area.

All of the above methods can be used to produce

nematodes but they are either too expensive or do not

produce sufficient quantities for field release.

The symbiotic relationship between the nematode and the

associated bacterium was not understood when the above

culture methods were developed. However, when N.

carpocapsae was described in 1955, the symbiotic

relationship was known and that knowledge led to a

considerable improvement of in vitro culture. The

associated bacterium serves as a food source for the

nematode but apparently does not supply all the nutrients









73

needed (Poinar, 1979). After conducting several different

experiments, Poinar stated that he had been able to obtain

only up to two and a half generations of the nematode on any

kind of bacteriological agar even when the nematodes were

transferred to a new plate seeded with the bacterium. But,

when a medium containing additional nutrients was used,

continuous culture could be achieved. The medium on which

the bacterium is cultured apparently supplies other

nutrients for the nematodes. The first medium used for

nematode production contained commercial dog food as a base

(House et al., 1965). Hara et al. (1981) improved

production of the nematode by growing it monoxenically on a

dog food agar medium. Dutky et al. (1967a, 1967b), proved

that sterols are necessary for growth and development of the

nematode. That information, as well as the knowledge that

the bacterium, Xenorhabdus nematophilus, has a primary and a

secondary form (Akhurst, 1980, see also, section on

"Associated bacterium" below), was very important in

increasing the in vitro production of the nematode. A major

break-through in mass production of the nematode was

accomplished by Bedding (1981) who provided a large surface

area by using a sterilized, polyether-polyurethane sponge

thinly coated with a homogenate of 70% pig's kidney, 10%

beef fat and 20% water. Before adding infective-stage

nematodes, the primary form of the bacterium was added to

the sterilized medium. Bedding (1984) improved his mass-









74

production technique by coating the sponge with a homogenate

of chicken offal and by using autoclavable plastic bags as

containers.

The cost of producing the nematode using the method of

House et al. (1965) was estimated to be $1.00 per million

infective-stage nematodes, that of Hara et al. (1981), 28

cents per million, and that of Bedding (1981), 2 cents per

million.

Associated bacterium:

The bacterium associated with steinernematids was

described by Poinar and Thomas (1965) as Achromobacter

nematophilus. The bacterium was isolated from Steinernema

carpocapsae strain DD-136. Hendrie et al. (1974) rejected

the genus Achromobacter and transferred all but one species

in the genus to other genera. Since A. nematophilus could

not be accommodated by other genera, Thomas and Poinar

created a new genus for it, Xenorhabdus nematophilus.

Another bacterium resembling X. nematophilus, X.

luminescens, was described by Poinar and Thomas (1979) from

Heterorhabditis bacteriophora. The authors placed the genus

Xenorhabdus in the family Enterobiaceae, but pointed out

that it differs from the currently accepted genera in the

family by its large cell size, its immunological properties,

its intimate association with entomogenous nematodes, and


its pathogenesis to insects.











Akhurst (1980) demonstrated that the bacterium was

morphologically and functionally dimorphic having a primary

and a secondary form. The primary form, which is carried

into a new host by the infective-stage nematode, apparently

remains stable in the host (Akhurst, 1980), but the primary

form may revert to the secondary form in in vitro culture.

These two forms can be distinguished by biochemical tests

and by color and size of the colonies. Either form is

equally pathogenic when inoculated into larvae of the

greater wax moth, Galleria mellonella, but the primary form

is much more effective than the secondary form for

production of nematodes. Akhurst (1980) showed that yield

of infective stage juveniles obtained from axenic nematodes

plus the primary form of the bacterium was seven times

greater than that from axenic nematodes plus the secondary

form. He noted that the reason the bacterium changes to the

secondary form is not known, but that no bacteriophage, or

plasmid is involved in mediating the change.

A wide range of microorganisms was found to be

inhibited by the primary form of Xenorhabdus spp. but not by

the secondary form. This inhibition suggests that

Xenorhabdus spp. produce antibiotic substances.

Akhurst (1983) studied the taxonomy of the bacteria

associated with various species of the nematode and, based

on the guanine and cystosine content of DNA, the color of

the colonies, and the production of acid from carbohydrates








76

of the bacterium, suggested the following subspecies: X.

nematophilus subspecies nematophilus for bacteria symbiotic

with S. carpocapsae; X. nematophilus subspecies bovienii

for bacteria symbiotic with S. bibionis; and X.

nematophilus subspecies poinarii for bacteria symbiotic with

S. glaseri.


Materials and Methods


Isolation of Bacteria

The bacteria associated with Steinernema scapterisci

were isolated by one of three methods.

Method 1: This method was a modification of the hanging drop

technique suggested by Poinar (1966). A drop of hemolymph

taken aseptically from a mole cricket was placed in a 35 x

10 mm petri dish. Five infective-stage juvenile nematodes,

sterilized previously by placing them for 2 hours in 0.1%

merthiolate and washing them 3 times in sterilized deionized

water, were transferred into the drop of hemolymph. The

petri dish was placed in a plastic bag containing a folded

facial tissue saturated with water, the bag was tied and

kept in the dark at 25 C. After 24 hours, the bacteria had

developed sufficiently and were transferred to nutrient agar

or tergitol-7 agar with triphenyltetrazolium chloride and

bromothymol blue (T-7 agar) (Poinar and Thomas, 1965) in

order to identify the colonies as those of the associated


bacterium.











Method 2: This was a modification of Akhurst's method

(1980). Infective-stage juveniles were surface-sterilized

by immersion in 0.1% merthiolate for 2 hours, washed 3 times

in sterile water, then about 50 of them in water were

pipetted into a 0.25 cc tissue grinder (Akhurst used yeast

and salt broth instead of water) and macerated. The

macerate was spread on nutrient agar or T-7 agar. The

bacterial colonies appeared after 36-48 hours at 25 C.

Method 3: In this method, bacteria were collected from the

hemolymph of mole crickets as follows: Mole crickets were

inoculated with a suspension of infective-stage juveniles in

an inoculation chamber as explained for in vivo culture.

After 24 hours, a mole cricket that slowed in movement and

had trembling legs (a sure sign of infection) was selected.

Its abdomen was squeezed gently to press out any fecal

matter in the rectum to prevent the release of such matter

into culture media when the hemolymph was collected. The

entire mole cricket was washed in tap water followed by a 2-

3 minute dip of its rear end into 30% hydrogen peroxide.

Then the tip of the cercus was cut off with sterilized

scissors. The hemolymph which oozed from the circus was

collected in a dish containing T-7 agar and spread over the

surface with a loop. After 24 hours bacterial colonies

developed in the culture medium and were transferred to new

T-7 agar plates.











Preparation of Bacteria for Culturing the Nematode

To produce sufficient quantities of the bacteria on

which to grow the nematode in vitro, a bacterial colony was

transferred aseptically to a flask containing brain-heart

infusion broth which had been autoclaved previously. The

flask was attached to a shaker arm and shaken continuously.

After 48 hours, the bacteria were ready to inoculate onto

the solid culture medium.

Experiment 1

To determine the cultural and fermentation

characteristics of the bacterium, tests on various culture

media and test substances (Table 10) were carried out.

Bacterial colonies 24 hour old were inoculated onto

previously prepared media in petri dishes. Different

substances, prepared as instructed by their producers were

placed in separate test tubes with bromcresol purple as a pH

indicator when needed. A drop of 24-hour-old bacteria

produced in liquid medium was placed in one of two tubes of

the same substance, with the other tube serving as a

control. The results were obtained after 24 hours.

Experiment 2

The purpose of this experiment was to determine the

suitability of eight culture media for rearing the nematode.

The 8 culture media listed below were prepared as indicated

for each medium. Where agar was used, the agar was melted

in warm water. Where crickets and animal parts were used,











they were macerated in a blender in the amount of water

indicated for each medium. Where sponge was used, the

sponge was chopped into small pieces. All media were placed

in 500 ml flasks, each plugged with a two-hole stopper

fitted with a glass tube in each hole and a short piece of

autoclavable hose attached to each glass tube. The end of

the two hoses were covered with aluminum foil and all flasks

were autoclaved for 30 minutes at 121 C and 15 psi pressure.

Medium 1: 3 g nutrient agar, 120 ml H20, 1 ml corn

oil, and 10 g sponge. The combination was mixed well in a

500 ml Erlenmeyer flask.

Medium 2: Similar to Medium 1 but without sponge.

Also, after autoclaving and before the agar became firm,

the flask was rotated to distribute agar on the flask wall.

The purpose of this maneuver was to increase the rearing

surface.

Medium 3: 3 g nutrient agar, 120 ml H20, 2 mole

crickets.

Medium 4: 3 g nutrient agar, 1 g brain-heart infusion,

120 ml H20, and 1 ml corn oil.

Medium 5: 20 g pork kidney, 50 ml H20, 1 ml corn oil,

and 10 g sponge.

Medium 6: 20 g pork kidney, 5 g pork brain, 50 ml H20,

and 10 g sponge.

Medium 7: 20 g pork liver, 1 ml corn oil, 50 ml H20,

and 10 g sponge.










80
Medium 8: 20 g pork liver, 5 g pork brain, 50 ml H20,

and 10 g sponge.

One day after the media were prepared and autoclaved,

they were inoculated with bacteria by adding 5 ml of

bacteria-brain-heart infusion broth. Two days later, about

20,000 surface-sterilized infective-stage juveniles were

added to each flask. At that time a bacteriological filter

was fitted to each hose emanating from the stopper in each

flask. One hose served as an inlet for air from an aquarium

pump and the other as an outlet. The air entering the flask

was passed through water to increase its relative humidity

and thus reduce drying of the medium (Fig. 16). The system

was maintained at 25 C. After 14 days the nematodes were

harvested by flooding the flasks with water and emptying the

contents onto a filter in a pan (modified Baermann funnel).

Nematodes that moved out of the media, through the filter

and into the water in the pan were collected after 24 hours.

Experiment 3

This experiment was conducted in order to determine

whether nematodes produced in vitro would kill mole

crickets. Five mole crickets were exposed in a petri dish

to 8,000 infective-stage juvenile nematodes cultured in

vitro. The experiment was replicated four times. After 3

days, the number of dead crickets was recorded.

























N I L. -
mmmm m r
mmm, II


Fig. 16. Culture flasks inoculated with bacteria and
nematodes receive humidified and sterilized air
filtered through a bacteriological filter.


mm











Experiment 4

This experiment was conducted to determine whether

there was any difference in the kill rate of mole cricket

nymphs by nematodes produced in vitro or in vivo. The mole

cricket nymphs used in this experiment had a pronotum length

of 4-9 mm. There were three treatments: (1) the control

without nematodes, (2) infective stage juveniles produced in

vivo and (3) infective stage juveniles produced in vitro.

The inoculation technique was similar to that for in vivo

culture, but the number of nematodes used was 20,000. The

results were determined after 4 days.


Results and Discussion


Isolation of Bacterium

All three methods used to isolate the bacterium are

satisfactory. However, Methods 1 and 3, which involve

isolating the bacterium from mole crickets, were less

satisfactory than Method 2, which involved isolating the

bacterium from the nematode. The reasons are that in

isolating the bacterium from mole crickets, which are

associated with soil, the chance of contamination is high,

and the methods are more complicated. Thus, Method 2 is

simpler and results in less contamination than Method 1 and

3.










83

Description of the Bacterial Colony

After two days on T-7 agar plates, small round colonies

1/2-1 mm in diameter were formed. Initially the colonies

were grey in color, but gradually became darker until after

2 days a small reddish-brown spot was formed at the center

of the colonies. The area surrounding the reddish spot was

clear to slightly blue. The spot in the center of the

colonies increased in size and changed color gradually to

become dark purple. The outer margin of the colonies were

undulated, some more so than others.

Experiment 1

Results of cultural and biological studies are

summarized in Table 10. This bacterium developed on all

media tested. The fermentation test showed that the

bacterium is fermentative in its metabolism of the tested

substances. The negative result in Voges-Proskauer test

suggested that in the fermentation of glucose, the bacterium

cannot produce the neutral end product, acetylmethylcarbinol

(acetoin). The positive results of amino acid tests show

that the bacterium can decarboxylate these acids to form

amines. This bacterium is not capable of utilizing citrate

as the sole source of carbon for its metabolism.










84

Table 10: Cultural and biological characteristics of the
bacterium associated with Steinernema
scapterisci.


Mat-ri al.


PRar- i n


Culture Media


Brain-heart infusion agar
MacConkey agar
Nutrient agar
TSA blood agar base
Tryptic soy agar

Test Substances

Adonitol
Arabinose
Arginine
BCP + glucose
BCP + lactose
BCP + manitol
BCP + rhamnose
BCP + sucrose
BCP + trehalose
Bile esculin
Citrate
Hippurate broth
Inulin broth
Indole
Lysine
Nitrate
OF glucose
ONPG broth
Ornithine
Growth-Peptone + .6% NaC1
Simmon slant
Sorbose
TSB broth
Urease
Voges-Proskauer


ICP = Bromcresol purple
= reaction
= no reaction
'= weak
= not determined


Acid

+
+
+
+
+w
+
+w
+
+
+

+
+w

+
+
+
+
+

+
+
+
+


"""""'Reaction











Experiment 2

By 6 days, nematodes developed rapidly on Culture Media

2, 4, 5, and 7 and were present on the flask walls away from

the media. On day 8, the nematode populations were greater

on Medium 4 than on the other media. On day 12, nematodes

covered the flask walls of Media 4 and 7. On a few other

media, a few nematodes were seen on the flask walls.

Nematodes were harvested from all flasks 15 days after

inoculation. Medium 7, which contained pork liver, oil,

sponge and water, produced the greatest number of nematodes

of all media tested (Table 11). While Media 3 and 4 also

supported fair growth and reproduction of the nematode, the

juveniles were extremely difficult to extract from the agar

in the medium to the extent that the numbers reported in

Table 11 undoubtedly are low.

Experiment 3

Nematodes from in vitro culture killed 100% of the mole

crickets tested while 4 died in the controls (Table 12).

The results show that nematodes produced in vitro are just

as effective in killing mole crickets as are those produced


in vivo.










86

Table 11: Number of third-stage infective juvenile
nematodes harvested from each culture medium.


Medium

1

2

3

4

5

6

7

8


um


Agar, oil, sponge

Agar, oil

Agar, mole crickets

Agar, oil, brain-heart

Kidney, oil, sponge

Kidney, brain, sponge

Liver, oil, sponge

Liver, brain, sponge


Table 12: Number of mole crickets killed
produced by in vitro culture.


o. nematodes

500,000

5,000

1,500,000

3,500,000

1,500,000

1,700,000

40,000,000

1,000,000


by nematodes


No. mole crickets that died

Replicate Treatment Control

1 5/5 0/5

2 5/5 1/5

3 5/5 1/5

4 5/5 2/5

Total 20/20 4/20


Nc











Experiment 4

Fifteen of 25 mole crickets were killed by nematodes

produced by in vitro culture compared to 14 of 25 killed by

those produced by in vivo culture (Table 13). Thus nematodes

produced by in vitro culture were just as effective in

killing mole crickets as were those produced by in vivo

culture.

Comparing the data in Table 12 with that in Table

13, the rate of kill of adult mole crickets was higher than

that of nymphs. This indicates that the nematode does not

kill nymphs as effectively as it kills adults. Preliminary

tests showed that nymphs with a pronotal length equal to or

less than 2 mm were not killed by the nematodes. One of the

reasons for this may be that the natural openings used by

the nematodes to enter the host are smaller in nymphs than

in adults.










88


Table 13: Comparison of nematodes produced in vitro vs.
in vivo for kill of mole cricket nymphs.


No. mole cricket nymphs that died



Replicate In vivo In vitro Control

1 3/5 4/5 1/5

2 2/5 3/5 0/5

3 2/5 3/5 1/5

4 3/5 3/5 1/5

5 4/5 2/5 1/5

Total 14/25 15/25 4/25
















CHAPTER 7
VERTICAL MIGRATION OF STEINERNEMA SCAPTERISCI N.SP. IN SOIL


Both laboratory tests and field tests (Chapter 9) have

shown that Steinernema scapterisci is very effective in

controlling mole crickets in Florida. Some preliminary

experiments in the laboratory indicated that the nematode

moved very little from the site of application. Since

knowledge about movement of the nematode through the soil is

very important in determining when and how to apply the

nematode in the field, additional experiments were

conducted. This chapter reports the results of those

experiments.


Literature Review


Moyle and Kaya (1981) found that when they released

Steinernema carpocapsae in sandy soil, at 15 cm below the

soil surface, 77% of the nematodes were recovered above the

point of release after 48 hours. When they placed them on

the soil surface, 90% of them remaining within 1 cm of the

surface. When they placed nematodes at depths of 2.5 and 5

cm below the soil surface, almost all of them remained at

the release site. Georgis and Poinar (1983) concluded that

an increase in the percentage of clay and silt decreased the

89




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