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Heterorhabditis spp. and Steinernema (= Neoaplectana) spp.: Temperature, and aspects of behavior and infectivity
EXPERIMENTALPARASITOLOGY
62, 169-180(1986)
Heterorhabditis
spp. and Steinernema (= Neoaplectana) spp.: Temperature, and Aspects of Behavior and Infectivity A. S. MOLYNEUX~
Division of Entomology, Commonwealth Scientific and Industrial Research Organization (CSIRO), Stowell Avenue, Hobart, Tasmania 7000, Australia (Accepted for publication 20 November 1985) MOLYNEUX, A. S. 1986. Heterorhabditis spp. and Steinernema (= Neoaplectana) spp.: Temperature, and aspects of behavior and infectivity. Experimental Parasitology 62, 169180. The infectivity of several species/strains of nematodes of the genera Heterorhabditis and Steinernema for postfeeding, 3rd instar larvae of the sheep blowfly, Lucilia cuprina, was tested in sand at various temperatures. Of the two genera, the steinernematids were more active at lower temperatures and parasitized L. cuprina over a greater temperature range. The temperature range of infectivity for L. cuprina differed between nematodes of the same genus and between strains of the same species. Parasitization of L. cuprina and Galleria mellonella occurred in a temperature range that was greater than that permitting nematode development and reproduction. Different strains of the same species were found to have different temperature limits for development and reproduction. Developmental rate was different for each nematode species tested with the heterorhabditids taking longer to complete their life cycle than did the steinernematids. o 1986 Academic PRSS, IK. INDEX DESCRIPTORS: Nematodes, parasitic; Heterorhabditis bacteriophora; Heterorhabditis heliothidis; Heterorhabditis, undescribed spp.; Steinernema (= Neoaplectana) bibionis; Steinernema feltiae (= carpocapsae); Steinernema glaseri; Steinernema kraussei; Steinernema, undescribed spp.; Undescribed steinemematid; Insects; Galleria mellonella; Lucilia cuprina; Bacteria; Xenorhabdus spp.; Dosage/mortality; Parasitization; LD,,; LD,; Temperature; Development; Developmental threshold temperature; Reproduction; Soil.
Nematodes of the genera Heterorhabditis and Steinernema (syn. Neoaplectana) (Wouts et al. 1982) are obligate parasites of insects (Poinar 1979). They have a nonfeeding, free living, infective juvenile, and a mutualistic association with specific bacteria (Xenorhabdus spp.) (Poinar 1979; Akhurst 1983). Infective juvenile nematodes are found usually in the soil where they are attracted to insects from a distance (Bedding and Akhurst 1975). On encountering an insect, they enter via its natural openings (Poinar 1979) and Heterorhabditis spp. also enter via intersegmental mem* Present address: Department of Entomology, Waite Agricultural Research Institute, University of Adelaide, Glen Osmond, South Australia 5064, Australia.
branes (Bedding and Molyneux 1982). After penetration, the nematodes invade the hemocoel where they void their bacteria and secrete an inhibitor of the antibacterial enzymes of the insect host (Gdtz et al. 1981). The bacteria kill the insect thereby providing nutrients for nematode development and reproduction while inhibiting the growth of other bacteria (Akhurst 1982). Within about 2 weeks, the newly formed infective juvenile nematodes leave the cadaver and are ready to parasitize other soil-dwelling insect hosts. Since there is considerable inter- and intraspecific variation in the infectivity of several species/strains of Heterorhabditis and Steinernema for insects (Molyneux et al. 1983; Bedding et al. 1983), an understanding of how environmental factors affect nematode behavior in soil is necessary
169 0014-4894/86$3.00 Copyright 0 1986 by Academic Press, Inc. All rights of reproduction in any form reserved.
170
A.S.MOLYNEUX
in choosing the most appropriate species to give optimal reduction of an insect pest. Previaus studies on the influence of abiotic factors on nematode behavior have largely involved either Steinernema glaseri (Jackson 1973; Georgis and Poinar 1983a) or Steinernema feltiae (syn. Neoaplectana carpocapsae) (Schmiege 1963; Kaya 1977; Burman and Pye 1980; Georgis and Poinar 1983b). However, Molyneux and Bedding (1985) and Molyneux (in press) recently found considerable differences in nematode behavior between heterorhabditids and steinernematids within various soil textures, moisture levels, and temperatures.
The present study was undertaken to compare the behavior and infectivity in soil of several spe&z&ains ofnematodes parasitic in insects and to compare their subsequent development and reproduction at different temperatures. MATERIALSANDMETHODS The Heterorhabditis and Steinernema spp. nematodes were initially obtained from various sources (Table I) and were cultured in vivo using larvae of the wax moth Galleria mellonella (L.) at 23 C to ensure a uniform age of 4 -C 3 days. Larvae of Lucilia cuprina (Wied.) and G. mellonella were obtained from laboratory cultures. Post feeding L. cuprina larvae were used in infectivity experiments because large numbers
TABLE I Sources of Nematodes Nematode Heterorhabditis spp.
Strain DI Q380 V16 T310
H. heliothidis
NC NZ T321
H. bacteriophora Steinernema spp.
Wl Wll
S. bibionis
T33.5
N60 S. glaseri
KG
S. feltiae
Agriotos
5’. kraussei
-
Undescribed steinernematid
Ql
a Isolated by the method of Bedding and Akhurst (1975)
Source Soil”, Darwin, Northern Territory, Australia Soil”, Yeppoon, Queensland, Australia ex Graphognathus leucoloma, Geelong, Victoria, Australia Soil”, Sandy Bay, Tasmania, Australia W. Wouts, DSIR, New Zealand W. Wouts, DSIR, New Zealand Soil”, Dysart, Tasmania, Australia W. Wouts, DSIR, New Zealand Soil”, Dwellingup, Western Australia, Australia Soil”, Dwellingup, Western Australia, Australia ex Otiorhynchus sulcatus, Nicholl’s Rivulet, Cygnet, Tasmania, Australia Soila, (sheep campsite), Canberra, A.C.T., Australia H. K. Kaya, University of California, Davis, CA, USA G. 0. Poinar, University of California, Berkeley, CA, USA Z. Mratek, CSAV, Budejovice, Czechoslovakia Soil=, Mirani, Queensland, Australia
Heterorhabditis, Steinernema SPP.: ASPECTS OF BEHAVIOR of larvae of uniform age could be easily obtained from laboratory cultures and at this stage of their development they,,busrow into soil. Clean active suspensions of infective juveniles were obtained using the methods of Whitehead and Hemming (1965). Nematode suspensions were diluted to lo3 infective juveniles/ml and 0.1 ml aliquots were transferred on 1.0 cm2 pieces of filter paper to 1.0% purified agar in Petri dishes and incubated at the required temperatures in controlled temperature rooms and incubators. Nematodes were classified as inactive if no movement occurred, as barely active if they moved part of their body, and active if able to move along the agar surface after mechanical stimulation. Postfeeding, 3rd instar L. cuprina larvae were exposed individually to nematode parasitization within plastic screwcap specimen jars (diameter 4.2 cm, height 6.0 cm) tilled to within 1.0 cm of the top with approximately 80 g of clean fine sand, moisture content 7% (0.03 bars). Infective juvenile nematodes were introduced in 1.O ml of water into a centrally located hole (diameter 0.5 cm, depth 2.0 cm), covered with sand, and left for 16 hr to allow equilibration at the required temperature. Dosages of 1 to 10 nematodes were individually counted whereas dosages greater than 10 were estimated by dilution counts. A single blowfly larva was placed on the sand surface of each jar, and the lid secured after ensuring that the larva had started to burrow. The range of temperature allowing parasitization was established using a single dosage of 320 (102.5)infective juveniles for each of the 16 nematode species/ strains with 20 replications for each nematode/temperature combination. In a separate experiment, the parasitization of L. cuprina larvae by 12 nematode species/strains was analyzed quantitatively for three temperatures: 18, 23, and 28 C. For each nematode/temperature combination, there were eight dosages of infective juveniles: lOa, lOi, 101.5,102, 102.5,103, 103.5,and 104.Each combination of nematode/temperature/dosage was tested on two occasions, with each combination consisting of 20 specimen jars. After lo-14 days at temperatures of 18 C or more and 28 days at temperatures below 18 C, the jars were emptied and the sand sieved in water. The larvae or puparia found were examined microscopically in insect Ringer’s solution for nematode parasitization and reproduction. In both experiments, it was not feasible to examine simultaneously the 16 and 12 species/strains of nematodes. The species/strains were divided into four groups of four or three groups of four; when each group was tested, Heterorhabditis sp. Dl was included as a standard and there were 20 nematode-free controls. LD,, and LD,, values for each nematode/insect combination were computed using the probit analysis
171
of Finney (1971). The coincidence of the two probit lines obtained for each nematode species/strain was tested and comparisons between the selected nematode/temperature combinations were made as pairwise tests of coincident lines using the general linear model concept in the Genstat statistical package (Alvey et al. 1977). Because nematode reproduction is so variable in L. cuprina larvae (Molyneux 1984), final instar G. mellonella larvae were exposed individually to nematode infection within specimen jars filled with moist sand (moisture content 7% = 0.03 bars). Five Heterorhabditis sp. or 10 Steinevnema sp. infective juveniles were added to each specimen jar with 20 replications for each nematode/temperature combination. Jars were then left for 16 hr (S. gluseri) or 48 hr (S. feltiae, Heterorhabditis sp. Dl, H. heliothidis T327) at 23 C to allow parasitization. Cadavers of G. mellonelln were then placed in glass vials (diameter 2.5 cm, height 7.5 cm) containing 5 ml of 0.1% Formalin. Formalin was added to reduce the risk of foreign microorganisms dominating the nematodes’ bacterial symbiont at the lower temperatures. Each cadaver was supported above the water level by a piece of pleated filter paper (diameter 9.0 cm., height 4.5 cm) inserted into and placed at right angles to the water as described by Kaya (1977). The tops of the glass vials were covered with parafilm and placed at the required constant temperatures. The total number of infective juveniles that emerged per cadaver was counted and cadavers which did not produce infective juveniles were dissected and examined microscopically in insect Ringer’s solution. The approximate rates of development of four nematode species were determined by measuring the time taken between parasitization of the insect host and emergence of the newly formed infective juveniles .
RESULTS
Very few Heterorhabditis spp. were mobile at temperatures below 9 C whereas at least half of the Steinernema spp. were active at temperatures below 7 C (Table II). Each nematode species/strain parasitized some Lucilia cuprina larvae with the steinernematids favoring the lower end of the temperature range (Fig. 1). Maximal parasitization of L. cuprina larvae by Heterorhabditis spp. occurred within a narrower range of temperatures than that associated with Steinernema spp. The results of dosage parasitization experiments are summarized in Figs. 2-4 and Table III. There was no significant differ-
172
A. S. MOLYNEUX
TABLE II Immobilization of Heterorhabditis and Steinernema Infective Juvenile Nematodes at Low Temperatures Temperature (C) at which nematode is Nematode Heterorhabditis sp. Heterorhabditis sp. Heterorhabditis sp. H. bacteriophora H. heliothidis H. heliothidis H. heliothidis Heterorhabditis sp. Steinernema sp. Undescribed steinernematid Steinernema sp. S. glaseri S. feltiae S. bibionis S. kraussei S. bibionis
Strain
Active
Barely active
Inactive
Q380 Dl V16 NZ T327 NC T310 WI
16 12 12 12 12 10 10 10 14
1.5 11 10-11 10-11 9-10 7-8 8-9 8-9 11-12
14 10 9 9 8 6 7 7 10
Ql
14 10 10 8 4 4 3
11-12 8-9 8-9 6-7 3 3 l-2
10 7 I 5 2 2 0
WI1 KG Agriotos T335 N60
ence between the results obtained for any nematode/temperature/dosage combination repeated at different times. LD,, and LD, values were not calculated for S. kraussei tested at 28 C because parasitization did not reach 50% at any dosage. There were considerable differences between the 12 nematode species/strains, and more importantly, between strains belonging to the same species. S. bibionis T335 was significantly better (P < 0.001) than S. bibionis N60 at each temperature tested, and H. heliothidis T327 was significantly better than H. heliothidis NC and NZ at 18 (P < 0.001) and 23 C (P < 0.001). However, at 28 C, there was no significant difference between the three strains of H. heliothidis. At 28 C, Heterorhabditis sp. Dl was more effective (P < 0.001) than any other nematode, and H. heliothidis T327 was the most effective nematode below 28 C (P < 0.05). At 18 C, Heterorhabditis sp. V16, H. heliothidis NC and T327, S. kraussei, and both strains of S. bibionis displayed optimum infectivity, whereas H. heliothidis NZ was more effective at 23 C.
Heterorhabditis spp. Dl and Q380 were most effective at 28 C. In contrast, there was no significant difference in the infectivity of H. bacteriophora, S. glaseri, and S. feltiae for L. cuprina between 18 and 28 C. The steinernematids produced infective juveniles in the majority of L. cuprina cadavers resulting from exposure to dosages of 10’ to lo3 nematodes at 18 and 23 C but no reproduction occurred at 28 C. At 28 C, the majority of Heterorhabditis spp. reproduced effectively in all cadavers resulting from exposure to dosages of loo to 102.5infective juveniles with occasional reproduction at dosages of 103. However, H. heliothidis strain T327 did not reproduce at any dosage at 28 C. At 23 C, all Heterorhabditis spp. tested reproduced in cadavers at dosage rates of loo to 102.5 infective juveniles. However, at 18 C, Heterorhabditis spp. Dl and Q380 did not reproduce at any dosage in L. cuprina cadavers and H. bacteriophora reproduced only occasionally. The remaining heterorhabditids reproduced at 18 C in all cadavers
Heterorhabditis, Steinernema SPP.: ASPECTS OF BEHAVIOR
TEMPERATURE C
FIG. 1. Parasitization of postfeeding, 3rd instar larvae of Lucilia cuprina exposed individually to dosages of 320 heterorhabditid.(A, B) or steinernematid (C, D) infective juvenile nematodes in 80 g sand (0.03 bars) at various temperatures. O-O, Heterorhabditis sp. Dl; X---X, Heterorhabditis sp. Q380; A- --A, Heterorhabditis sp. V16; *----Q, Heterorhabditis sp. T310; 0- - -0, H. heliothidis T327; n - - 4, H. heliothidis NC; A-A, H. heliothidis NZ; m---M, H. bacteriophora; f~- - -6, Steinernema sp. Wl; X-X, Steinernema sp. Wll; A- - -A, S. bibionis N60; O---O, 5’. bibionis T33.5; A-A, S. feltiae Agriotos; U-0, S. glaseri KG; l - - -0, S. kraussei; q - - -0, steinernematid species Ql.
resulting from dosages of loo to 102.5infective juveniles except H. heliothidis T327 which reproduced only in cadavers resulting from dosages of loo to lo2 infective juveniles. At dosage rates above 102.s,no production of Heterorhabditis spp. infec-
173
tive juveniles occurred at 18, 23, and 28 C. At those dosages, the characteristic pigmentation normally produced by the bacterial symbiont was absent and cadavers were invariably fetid. Although the temperature range supporting nematode reproduction inside L. cuprina and Galleria mellonella was similar, the steinernematids reproduced at higher temperatures inside larvae of G. mellonella. Of the four nematode species studied in detail, parasitization occurred over a greater range of temperatures than the temperature range allowing nematode reproduction, and substantial differences in the production of infective juveniles in G. mellonella occurred between the different nematode species. Heterorhabditis sp. Dl produced infective juveniles over the narrowest temperature range (20-32 C), whereas S. glaseri produced infective juveniles across a temperature range of 12 to 32 C. Heterorhabditis sp. Dl did not reproduce at 18 C and only developed partially to the L4 stage within 9 days after parasitization of the host. At temperatures below 10 C, H. heliothidis T327, S. glaseri, and S. feltiae did not reproduce and no live nematodes were recovered at the end of 14 days. At 10 C, S. glaseri and S. feltiae molted to the L4 stage and H. heliothidis T327 developed into first generation adults, but died after 4 weeks without further development. At 12 C, a few infective juveniles emerged from approximately 10% of cadavers infected with S. glaseri or H. heliothidis T327 after 8 weeks and after 10 weeks from cadavers infected with S. feltiae. At 28 and 32 C, only first generation adults were produced in cadavers infected with H. heliothidis and S. feltiae, respectively. At 35 C, S. glaseri developed to first generation adults but failed to reproduce while Heterorhabditis sp. Dl did not develop. The time taken between parasitization of the insect and emergence of the newly formed infective juveniles from the cadaver (measured as rate of development) was dif-
174
A. S. MOLYNEUX 380
1%
Y = 1.436.x
+ 0.176
-.-
23c
Y = 1.472.x
+ 1.148
--o--
28C
Y = 0.839X
+ 2.739
..
.o.....
99 :i .‘/ :ix .’ /
0
0
7
..‘/ .’ / :’ ,
90 6
70 5
50 30
4 10 3
:
: :’ .’
2
Zl z a
B
18C
Y = L.242.7 + 2.299
2%
Y = i.062.x
+ 2.508
-.--n--
28C
Y ,= 0.958X
+ 2.813
. . . ..o.....
16
,
/
, /
18C
Y = 0.840X
+ 3.572
23c
Y = 0.801X
+ 3.267
-.--mm-
28C
Y = 0.806X
+ 3.082
. . . ..o...
2 CA0 99 7
90 6
70 50 30
10
1 10"
lo=
lo3
10'
loo
DOSAGE (No.
10'
lo2
ld"
10'
NEMATODES)
FIG. 2. Dosage/parasitization lines for postfeeding, 3rd instar larvae of Lucilia cup&a exposed to infective juveniles of Heterovhabditis sp. Dl (A), Heterorhabdifis sp. Q380 (B), H. bacteriophora (C), and Heterovhabditis sp. V16 (D) in sand (0.03 bars) at 18, 23, and 28 C.
Heterorhabditis, Steinernema SPP. : ASPECTS OF BEHAVIOR :
175
1%
Y = 1.344x
+ 2.772
-.-
18C
Y = 0.947x
+ 2.613
23C
Y = ,.*59x
+ 2.560
--a--
2x
Y = 1.206.x
+ 2.470
--o--
28C
Y = 0.920x
+ 2.948
. . . ..o.....
28C
Y = 1.14*x
+ 2.416
.....o.....
12
-.-
99 7
so 6 70 5
50 30
4 10 E 2 4 F: s LI
3 1 327
-.-
18C
Y = ,.340x
+ 3.561
23C
Y = 1.188X
+ 3.458
--m--
2%
Y = 0.927x
+ 2.854
. . . ..o.....
23C
Y = 0.799x
--•--
+ 2.421
if o,n SE 7
9c 6 7C 5
5c 3(
4 l( _’
C
_’_’
3 I 10'
10=
103
lo4 DOSAGE
(No.
loo
10:
10=
10"
lo4
NEMATODES )
FIG. 3. Dosageiparasitization lines for postfeeding, 3rd instar larvae of Lucilia cuprina exposed to infective juveniles of Heterorhabditis heliothidis NC (A), H. heliothidis NZ (B), H. heliothidis T327 (C), and Steinernema kruussei (D) in sand (0.03 bars) at 18, 23, and 28 C.
2 z -2
176
A. S. MOLYNEUX MO
1%
Y -
23c
Y - 0.870X + 2.802
1.291.x
+ 2.424
*WC
Y = 0.438X
-.-
r335
---o-. . . ..o.....
+ 3.082
18C
Y = 1.757x
+ 2.7cm
-.-
2x
Y = 1.3761(
+ 2.785
--•.--
28C
Y = 1.237): + 2.291
. . ..a.....
99 7
so 6 70 50
5
30 4
10 5 2 2 s 2
3
l
1%
Y = 1.167X + 3.067
23C
Y = 1.300x
+ 2.898
28C
Y = 1.,19x+
2.824
-.--n--
18C
Y = 0.852X
23C
Y = 0.918X + 2.942
. . . ..a....
28C
Y = 0.953x
-.-
+ 2.809
--a-.o..
+ 2.687
2 D 99
co 7
so 6
70 50
5
30 4
10
3
1 lo9
I
lOI
I
lo2
I
I
lo3
I
lo4 loo 10' DOSAGE (No. NEMATODES)
lo2
lo3
lo4
FIG. 4. Dosage/parasitization lines for postfeeding, 3rd instar larvae of Lucike cuprina exposed to infective juveniles of Steinevnema bibionis N60 (A), S. bibionis T33S (B), S. glasevi KG (C), and 5’. feltiae Agriotos (D) in sand (0.03 bars) at 18, 23, and 28 C.
2 1 & PI
177
Heterorhabditis, Steinernema SPP.: ASPECTS OF BEHAVIOR TABLE III Effect
of Temperature
Nematode
on Infectivity
Strain
Heterorhabditis sp. Heterorhabditis sp. Heterorhabditis sp. H. bacteriophora H. heliothidis H. heliothidis H. heliothidis Steinernema glaseri S. bibionis S. bibionis S. feltiae S. kraussei
Dl Q380 V16
NC NZ T327 KG T335 N60 Agriotos
of
Heterorhabditis
spp. and
18 c
571 4,080 50 149 45 331 12 45 20 99 372 213
402-827 2,760-6,860 30-78 106-208 31-63 223-497 7-17 30-65 14-28 70-136 243-580 149-303
Steinernema
spp. for
23 C
76 385 145 222 87 125 20 41 41 336 174 1,710
LDSO, 95% limits 52- 107 272-562 92-227 153-321 64-116 88-175 13-29 29-57 30-54 221-519 121-249 1,040-3,160
28 C
32 222 239 192 170 183 206 88 155 23,900 267
25-39 157-318 167-342 145-254 123-233 138-240 148-284 63-120 129-186 10,600-75,000 199-366
b
LD,,
Heterorhabditis sp. Heterorhabditis sp. Heterorhabditis sp. H. bacteriophora H. heliothidis H. heliothidis H. heliothidis Steinernema glaseri S. bibionis S. bibionis S. feltiae S. kraussei a Individual b Mortality
DI Q380 V16
NC NZ T327 KG T335 N60 Agriotos
7,820 46,700 1,670 1,610 408 7,480 107 569 109 972 11,900 2,950
4,450-17,100 22,300-143,000 912-3,810 1,020-2,970 268-711 3,960-18,000 69-189 359- 1,050 76- 179 643-1,670 5,750-33,000 1,770-5,980
941 8,370 5,790 3,580 905 1,440 239 400 347 10,000 4,320 68,900
95% limits 590- 1,750 4,510-19,300 2,880-15,300 2,080-7,460 597- 1,580 906-2,690 152-434 270-669 238-564 4,980-26,700 2,500-8,840 25,500-31,400
Lucilia cuprina”
609 5,920 9,300 4,190 4,190 2,420 4,970 1,230 1,690 2.01 x 10’ 5,890
446-881 3,270-13,100 5,220-19,600 2,700-7,250 2,570-7,800 1,610-4,060 3,030-9,300 815-2,070 1,260-2,400 3.2 x 106-3.0 x lo* 3,440-11,820
b
larvae in 80 g sand, moisture content 7% CO.03 bars). did not reach 50% at any dosage.
ferent for each nematode species tested at movement of the majority of steinernevarious temperatures (Fig. 5). S. glaseri matid infective juveniles may reflect the completed its life cycle in the fastest time colder climates of their European and and H. heliothidis had generally the slow- North American origins (Table II). Alest rate of development although the devel- though both strains of Steinernema biopmental rate of S. feltiae was the slow- bionis were isolated from soils or insects est of the four species at 12 C (Fig. 5). For taken from temperate areas of Australia each nematode species tested, second gen- (Table I), they were still mobile at 4 C. Aceration infective juveniles emerged from cording to Bedding (personal communicacadavers of G. mellonella. The relationship tion), it is likely that S. bibionis has been between developmental rate and tempera- introduced into Australia, most probably ture appeared to be linear, and the calcu- from Europe. In contrast, the inability of lated theoretical threshold temperatures for the heterorhabditids Dl and Q380 to move development gave a close approximation to at 10 and 14 C, respectively, suggests that the observed lower temperature limits for they are native to the warm, humid tropical development. areas. The variability in infectivity for Lucilia DISCUSSION cuprina larvae is, in part, related to the Important differences between the hete- length of time the insect host remdins susrorhabditids and steinernematids were ob- ceptible to nematode parasitization at difserved in the bioassays at various tempera- ferent temperatures (Molyneux 1984). Untures. The lower temperature limits for like lepidopteran pupae (Kaya and Hara
178
A. S. MOLYNEUX 0.3 -
l
Y=O.O07X-0.093,r=0.989,
13.S"
oY=O.O04X-O.O37,r-0.989,
8.3'
*Y=O.O07X-0.071,r=0.996,
9.9O
q Y=O.O13X-0.153,r=0.979,
11.8
0.2 -
01 0
I
I
I
5
10
15
I
I
I
20 25 M TEMPERATURE C
35
40
I
I
FIG. 5. Average rate of development for completion of life cycle of 0 -0, Heterorhabditis sp. Dl; 0 -0, H. heliothidis T327; k -*, Steinernema feltiae Agriotos; q -------a, S. glaseri KG in final instar Galleria mellonella cadavers at various temperatures. r, Correlation coefficient; a, calculated developmental threshold temperature C
1981; Moyle and Kaya 1981), L. cuprina larvae become almost completely invulnerable to parasitization soon after pupariation (Molyneux 1984). The greater infectivity of some nematodes (e.g., Heterorhabditis sp. Dl) at 28 C than at 18 C suggests that the increased activity of nematodes at the higher temperatures more than offsets any increase in the rate of pupariation of L. cuprina. The relationship between nematode activity and insect susceptibility is not peculiar to L. cuprina alone. Many of these insects that pupate in soil are probably similarly influenced, and other insects not pupating may be less active and thus become vulnerable to nematode parasitization. With all nematodes tested in this study, the range of temperature allowing parasitization was greater than the temperature range supporting nematode reproduction. This is of little benefit to the nematode at
the upper end of the temperature range because thermal damage soon resulted (unpublished data). However, at lower temperatures, some nematode species will remain in the hosts’ hemocoel as infective juveniles (unpublished data). There they can remain without killing the host until temperatures rise allowing bacterial growth and nematode reproduction. In the meantime, depending on the insect species, the insect may migrate and thus disseminate the nematode. The failure of Steinernema spp. to reproduce in L. cuprina at 28 C (Molyneux et al. 1983) like the failure of both genera when applied in high dosages at lower temperatures is apparently due to the contamination of the cadaver by foreign microorganisms (Molyneux 1984). The introduction of a high inoculum of gut bacteria, particularly by Steinernema spp. penetrating Lucilia cuprina larvae via the anus, can result in failure of the nematode’s bacterial symbiont to dominate within the host hemocoel. Unlike Steinernema spp., Heterorhabditis spp. infective juveniles can penetrate the intersegmental membranes of their insect hosts thereby avoiding the gut bacteria and reducing the risk of contamination. Furthermore, the outer cuticle is shed just before or during penetration, enabling Heterorhabditis species to enter with a surface that is largely bacteria free (Bedding and Molyneux 1982). Reproduction of steinernematids at temperatures of 28 C or more in Galleria mellonella larvae is probably the result of fewer foreign microorganisms being introduced due to nematode entry via the spiracles in addition to the anus and mouth. In agreement with previous observations (Jackson 1962; Kaya 1977; Poinar 1979), the most favorable temperature for the growth and reproduction of S. feltiae and S. glaseri lies between 23 and 28 C although the temperature limits for reproduction of S. feltiae Agriotos exceed those reported by Kaya (1977) and Pye and
Heterorhabditis, Steinernema
SPP.: ASPECTS
OF BEHAVIOR
179
otically associated with the insect pathogenic nemaBurman (1978) for the DD136 and Agriotos todes Neoaplectana and Heterorhabditis. Journal strain of Steinernema feltiae, respectively. of General Microbiology 121, 303-309. In contrast, the most favorable tempera- AKHURST, R. J. 1982. Antibiotic activity of Xenotures for growth and reproduction of Hetrhabdus spp., bacteria symbiotically associated erorhabditis sp. Dl and H. heliothidis with insect pathogenic nematodes of the families Heterorhabditidae and Steinernematidae. Journal T327 fell outside this range (30 and 20 C, of General Microbiology 128, 3061-3065. respectively) reflecting once again the cliAKHURST, R. J. 1983. Neoaplectana species: Speckmatic zones of their original locality. The ficity of association with bacteria of the genus Xedecline in nematode reproduction at high norhabdus. Experimental Parasitology 55, temperatures may be the result of competi258-263. tion with its own bacterial symbiont which ALVEY, N. G., et al. 1977. Regression and generalised linear models. In “GENSTAT: A General Statismay explain why Jackson (1962) reported tical program,” Chap. 7. Rothamsted Experimental minimal reproduction of S. glaseri at 35 C Station, England. in axenic liquid media. In addition, thermal BEDDING, R. A., AND AKHURST, R. J. 1975. A simple damage may result in a lack of viable technique for the detection of insect parasitic rhabditid nematodes in soil. Nematologica 21, 109- 110. sperm or ova or a change in the mating beBEDDING, R. A., AND MOLYNEUX, A. S. 1982. Penehavior of the nematode (Kaya 1977). At tration of insect cuticle by infective juveniles of Helow temperatures, the lack of nematode reterorhabditis spp. (Heterorhabditidae: Nematoda). production is probably a direct effect of Nematologica 28, 354-359. BEDDING, R. A., MOLYNEUX, A. S., AND AKHURST, temperature on nematode maturation. AkR. J. 1983. Heterorhabditis spp., Neoaplectana hurst (1980) reported that the primary form spp., and Steinernema kraussei: Interspecific and of Xenorhabdus spp. is essential for opintraspecific differences in infectivity for insects. timum nematode reproduction. The relaExperimental Parasitology 55, 249-257. tively few infective juveniles of SteinerBURMAN, M., AND PYE, A. E. 1980. Neoaplectana carpocapsae: Movements of nematode populations nema glaseri produced per cadaver at difon a thermal gradient. Experimental Parasitology ferent temperatures may have been due to 49, 258-265. the absence of the primary form of its bacFINNEY, D. J. 1971. “Probit Analysis,” 3rd ed. Camterial symbiont in the KG strain. bridge University Press, London. The results from these studies indicate GEORGIS,R., AND POINAR, G. 0. 1983a. Effect of soil texture on the distribution and infectivity of Neothat when selecting nematodes as possible aplectana glaseri (Nematoda: Steinernematidae). control agents for a particular pest insect Journal of Nematology 15, 329-332. nematode dosage rates, species of nemaGEORGIS,R., AND POINAR, G. 0. 1983b. Effect of soil tode, and insect type will need to be contexture on the distribution and infectivity of Neosidered in relation to temperature for their aplectana carpocapsae (Nematoda: Steinernematidae). Journal of Nematology 15, 308-3 11. permanent establishment in the field. ACKNOWLEDGMENTS
I thank Dr. R. A. Bedding for encouragement and guidance throughout this study; J. G. Moss, V. S. Patel, M. A. Stanfield, and W. H. Edwards for valuable technical assistance; Dr. D. A. Ratkowsky and R. Lowry for statistical advice; and Professor H. R. Wallace, Department of Plant Pathology, Waite Agricultural Research Institute, University of Adelaide, for constructive criticism of the manuscript. REFERENCES AKHURST, R. J. 1980. Morphological and functional dimorphism in Xenorhabdus spp., bacteria symbi-
G~~Tz,P., BOMAN, A., AND BOMAN, H. G. 1981. Interactions between insect immunity and an insectpathogenic nematode with symbiotic bacteria. Proceedings of the Royal Society of London Series B 212, 333-350.
JACKSON, G. J. 1962. The parasitic nematode, Neoaplectana glaseri in axenic culture, II. Initial results with defined media. Experimental Parasitology 12, 25-32. JACKSON, G. J. 1973. The ageing of Neoaplecfana glaseri. Proceedings of the Helminthological Society of Washington 40, 74-76. KAYA, H. K. 1977. Development of the DD-136 strain of Neoaplectana carpocapsae at constant temperatures. Journal of Nematology 9, 346-349.
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MOLYNEUX, A. S. 1984. The influence of temperature on the infectivity of heterorhabditid and steinernematid nematodes for larvae of the sheep blowfly, Lucilia cuprina. In “Proceedings of the Fourth Australian Applied Entomological Research Conference” (P. T. Bailey and D. W. Swincer, eds.), pp. 344-351. Govt. Printer, Adelaide. MOLYNEUX, A. S. Survival of infective juveniles of Heterorhabditis spp., and Steinernema spp. (Nematoda: Rhabditida) at various temperatures and their subsequent infectivity for insects. Revue de Nematologie, in press. MOLYNEUX, A. S., AND BEDDING, R. A. 1985. Influence of soil texture and moisture on the infectivity of Heterorhabditis sp. Dl and Steinernema glaseri for larvae of the sheep blowfly, Lucilia cuprina. Nematologica 30, 358-365. MOLYNEUX, A. S., BEDDING, R. A., AND AKHURST, R. J. 1983. Susceptibility of larvae of the sheep blowfly Lucilia cuprina to various Heterorhabditis spp., Neoaplectana spp., and an undescribed steinernematid (Nematoda). Journal of Invertebrate Pathology 42, l-7.
MOYLE, P. L., AND KAYA, H. K. 1981. Susceptibility of pupae of two cocoon-forming lepidopterous species to the entomogenous nematode Neoaplectuna carpocapsae (Rhabditida: Steinernematidae) Journal of Nematology 13, 419-421. POINAR, G. 0. 1979. “Nematodes for Biological Control of Insects.” CRC Press, Boca Raton, FL. PYE, A. E., AND BURMAN, M. 1978. Neoaplectana carpocapsae: Infection and reproduction in large pine weevil larvae, Hylobius abietis. Experimental Parasitology 46, 1- 11. SCHMIEGE, D. C. 1963. The feasibility of using a neoaplectanid nematode for control of some forest insect pests. Journal of Economic Entomology 56, 427-431. WHITEHEAD, A. G., AND HEMMING, J. R. 1965. A comparison of some quantitative methods of extracting small vermiform nematodes from soil. Annals of Applied Biology 55, 25-38. WOUTS,W. M., MR&?EK,, Z., GERDIN, S., AND BEDDING, R. A. 1982. Neoaplectana Steiner, 1929 a junior synonym of Steinernema Travassos, 1927 (Nematoda: Rhabditida). Systematic Parasitology 3, 147-154.
62, 169-180(1986)
Heterorhabditis
spp. and Steinernema (= Neoaplectana) spp.: Temperature, and Aspects of Behavior and Infectivity A. S. MOLYNEUX~
Division of Entomology, Commonwealth Scientific and Industrial Research Organization (CSIRO), Stowell Avenue, Hobart, Tasmania 7000, Australia (Accepted for publication 20 November 1985) MOLYNEUX, A. S. 1986. Heterorhabditis spp. and Steinernema (= Neoaplectana) spp.: Temperature, and aspects of behavior and infectivity. Experimental Parasitology 62, 169180. The infectivity of several species/strains of nematodes of the genera Heterorhabditis and Steinernema for postfeeding, 3rd instar larvae of the sheep blowfly, Lucilia cuprina, was tested in sand at various temperatures. Of the two genera, the steinernematids were more active at lower temperatures and parasitized L. cuprina over a greater temperature range. The temperature range of infectivity for L. cuprina differed between nematodes of the same genus and between strains of the same species. Parasitization of L. cuprina and Galleria mellonella occurred in a temperature range that was greater than that permitting nematode development and reproduction. Different strains of the same species were found to have different temperature limits for development and reproduction. Developmental rate was different for each nematode species tested with the heterorhabditids taking longer to complete their life cycle than did the steinernematids. o 1986 Academic PRSS, IK. INDEX DESCRIPTORS: Nematodes, parasitic; Heterorhabditis bacteriophora; Heterorhabditis heliothidis; Heterorhabditis, undescribed spp.; Steinernema (= Neoaplectana) bibionis; Steinernema feltiae (= carpocapsae); Steinernema glaseri; Steinernema kraussei; Steinernema, undescribed spp.; Undescribed steinemematid; Insects; Galleria mellonella; Lucilia cuprina; Bacteria; Xenorhabdus spp.; Dosage/mortality; Parasitization; LD,,; LD,; Temperature; Development; Developmental threshold temperature; Reproduction; Soil.
Nematodes of the genera Heterorhabditis and Steinernema (syn. Neoaplectana) (Wouts et al. 1982) are obligate parasites of insects (Poinar 1979). They have a nonfeeding, free living, infective juvenile, and a mutualistic association with specific bacteria (Xenorhabdus spp.) (Poinar 1979; Akhurst 1983). Infective juvenile nematodes are found usually in the soil where they are attracted to insects from a distance (Bedding and Akhurst 1975). On encountering an insect, they enter via its natural openings (Poinar 1979) and Heterorhabditis spp. also enter via intersegmental mem* Present address: Department of Entomology, Waite Agricultural Research Institute, University of Adelaide, Glen Osmond, South Australia 5064, Australia.
branes (Bedding and Molyneux 1982). After penetration, the nematodes invade the hemocoel where they void their bacteria and secrete an inhibitor of the antibacterial enzymes of the insect host (Gdtz et al. 1981). The bacteria kill the insect thereby providing nutrients for nematode development and reproduction while inhibiting the growth of other bacteria (Akhurst 1982). Within about 2 weeks, the newly formed infective juvenile nematodes leave the cadaver and are ready to parasitize other soil-dwelling insect hosts. Since there is considerable inter- and intraspecific variation in the infectivity of several species/strains of Heterorhabditis and Steinernema for insects (Molyneux et al. 1983; Bedding et al. 1983), an understanding of how environmental factors affect nematode behavior in soil is necessary
169 0014-4894/86$3.00 Copyright 0 1986 by Academic Press, Inc. All rights of reproduction in any form reserved.
170
A.S.MOLYNEUX
in choosing the most appropriate species to give optimal reduction of an insect pest. Previaus studies on the influence of abiotic factors on nematode behavior have largely involved either Steinernema glaseri (Jackson 1973; Georgis and Poinar 1983a) or Steinernema feltiae (syn. Neoaplectana carpocapsae) (Schmiege 1963; Kaya 1977; Burman and Pye 1980; Georgis and Poinar 1983b). However, Molyneux and Bedding (1985) and Molyneux (in press) recently found considerable differences in nematode behavior between heterorhabditids and steinernematids within various soil textures, moisture levels, and temperatures.
The present study was undertaken to compare the behavior and infectivity in soil of several spe&z&ains ofnematodes parasitic in insects and to compare their subsequent development and reproduction at different temperatures. MATERIALSANDMETHODS The Heterorhabditis and Steinernema spp. nematodes were initially obtained from various sources (Table I) and were cultured in vivo using larvae of the wax moth Galleria mellonella (L.) at 23 C to ensure a uniform age of 4 -C 3 days. Larvae of Lucilia cuprina (Wied.) and G. mellonella were obtained from laboratory cultures. Post feeding L. cuprina larvae were used in infectivity experiments because large numbers
TABLE I Sources of Nematodes Nematode Heterorhabditis spp.
Strain DI Q380 V16 T310
H. heliothidis
NC NZ T321
H. bacteriophora Steinernema spp.
Wl Wll
S. bibionis
T33.5
N60 S. glaseri
KG
S. feltiae
Agriotos
5’. kraussei
-
Undescribed steinernematid
Ql
a Isolated by the method of Bedding and Akhurst (1975)
Source Soil”, Darwin, Northern Territory, Australia Soil”, Yeppoon, Queensland, Australia ex Graphognathus leucoloma, Geelong, Victoria, Australia Soil”, Sandy Bay, Tasmania, Australia W. Wouts, DSIR, New Zealand W. Wouts, DSIR, New Zealand Soil”, Dysart, Tasmania, Australia W. Wouts, DSIR, New Zealand Soil”, Dwellingup, Western Australia, Australia Soil”, Dwellingup, Western Australia, Australia ex Otiorhynchus sulcatus, Nicholl’s Rivulet, Cygnet, Tasmania, Australia Soila, (sheep campsite), Canberra, A.C.T., Australia H. K. Kaya, University of California, Davis, CA, USA G. 0. Poinar, University of California, Berkeley, CA, USA Z. Mratek, CSAV, Budejovice, Czechoslovakia Soil=, Mirani, Queensland, Australia
Heterorhabditis, Steinernema SPP.: ASPECTS OF BEHAVIOR of larvae of uniform age could be easily obtained from laboratory cultures and at this stage of their development they,,busrow into soil. Clean active suspensions of infective juveniles were obtained using the methods of Whitehead and Hemming (1965). Nematode suspensions were diluted to lo3 infective juveniles/ml and 0.1 ml aliquots were transferred on 1.0 cm2 pieces of filter paper to 1.0% purified agar in Petri dishes and incubated at the required temperatures in controlled temperature rooms and incubators. Nematodes were classified as inactive if no movement occurred, as barely active if they moved part of their body, and active if able to move along the agar surface after mechanical stimulation. Postfeeding, 3rd instar L. cuprina larvae were exposed individually to nematode parasitization within plastic screwcap specimen jars (diameter 4.2 cm, height 6.0 cm) tilled to within 1.0 cm of the top with approximately 80 g of clean fine sand, moisture content 7% (0.03 bars). Infective juvenile nematodes were introduced in 1.O ml of water into a centrally located hole (diameter 0.5 cm, depth 2.0 cm), covered with sand, and left for 16 hr to allow equilibration at the required temperature. Dosages of 1 to 10 nematodes were individually counted whereas dosages greater than 10 were estimated by dilution counts. A single blowfly larva was placed on the sand surface of each jar, and the lid secured after ensuring that the larva had started to burrow. The range of temperature allowing parasitization was established using a single dosage of 320 (102.5)infective juveniles for each of the 16 nematode species/ strains with 20 replications for each nematode/temperature combination. In a separate experiment, the parasitization of L. cuprina larvae by 12 nematode species/strains was analyzed quantitatively for three temperatures: 18, 23, and 28 C. For each nematode/temperature combination, there were eight dosages of infective juveniles: lOa, lOi, 101.5,102, 102.5,103, 103.5,and 104.Each combination of nematode/temperature/dosage was tested on two occasions, with each combination consisting of 20 specimen jars. After lo-14 days at temperatures of 18 C or more and 28 days at temperatures below 18 C, the jars were emptied and the sand sieved in water. The larvae or puparia found were examined microscopically in insect Ringer’s solution for nematode parasitization and reproduction. In both experiments, it was not feasible to examine simultaneously the 16 and 12 species/strains of nematodes. The species/strains were divided into four groups of four or three groups of four; when each group was tested, Heterorhabditis sp. Dl was included as a standard and there were 20 nematode-free controls. LD,, and LD,, values for each nematode/insect combination were computed using the probit analysis
171
of Finney (1971). The coincidence of the two probit lines obtained for each nematode species/strain was tested and comparisons between the selected nematode/temperature combinations were made as pairwise tests of coincident lines using the general linear model concept in the Genstat statistical package (Alvey et al. 1977). Because nematode reproduction is so variable in L. cuprina larvae (Molyneux 1984), final instar G. mellonella larvae were exposed individually to nematode infection within specimen jars filled with moist sand (moisture content 7% = 0.03 bars). Five Heterorhabditis sp. or 10 Steinevnema sp. infective juveniles were added to each specimen jar with 20 replications for each nematode/temperature combination. Jars were then left for 16 hr (S. gluseri) or 48 hr (S. feltiae, Heterorhabditis sp. Dl, H. heliothidis T327) at 23 C to allow parasitization. Cadavers of G. mellonelln were then placed in glass vials (diameter 2.5 cm, height 7.5 cm) containing 5 ml of 0.1% Formalin. Formalin was added to reduce the risk of foreign microorganisms dominating the nematodes’ bacterial symbiont at the lower temperatures. Each cadaver was supported above the water level by a piece of pleated filter paper (diameter 9.0 cm., height 4.5 cm) inserted into and placed at right angles to the water as described by Kaya (1977). The tops of the glass vials were covered with parafilm and placed at the required constant temperatures. The total number of infective juveniles that emerged per cadaver was counted and cadavers which did not produce infective juveniles were dissected and examined microscopically in insect Ringer’s solution. The approximate rates of development of four nematode species were determined by measuring the time taken between parasitization of the insect host and emergence of the newly formed infective juveniles .
RESULTS
Very few Heterorhabditis spp. were mobile at temperatures below 9 C whereas at least half of the Steinernema spp. were active at temperatures below 7 C (Table II). Each nematode species/strain parasitized some Lucilia cuprina larvae with the steinernematids favoring the lower end of the temperature range (Fig. 1). Maximal parasitization of L. cuprina larvae by Heterorhabditis spp. occurred within a narrower range of temperatures than that associated with Steinernema spp. The results of dosage parasitization experiments are summarized in Figs. 2-4 and Table III. There was no significant differ-
172
A. S. MOLYNEUX
TABLE II Immobilization of Heterorhabditis and Steinernema Infective Juvenile Nematodes at Low Temperatures Temperature (C) at which nematode is Nematode Heterorhabditis sp. Heterorhabditis sp. Heterorhabditis sp. H. bacteriophora H. heliothidis H. heliothidis H. heliothidis Heterorhabditis sp. Steinernema sp. Undescribed steinernematid Steinernema sp. S. glaseri S. feltiae S. bibionis S. kraussei S. bibionis
Strain
Active
Barely active
Inactive
Q380 Dl V16 NZ T327 NC T310 WI
16 12 12 12 12 10 10 10 14
1.5 11 10-11 10-11 9-10 7-8 8-9 8-9 11-12
14 10 9 9 8 6 7 7 10
Ql
14 10 10 8 4 4 3
11-12 8-9 8-9 6-7 3 3 l-2
10 7 I 5 2 2 0
WI1 KG Agriotos T335 N60
ence between the results obtained for any nematode/temperature/dosage combination repeated at different times. LD,, and LD, values were not calculated for S. kraussei tested at 28 C because parasitization did not reach 50% at any dosage. There were considerable differences between the 12 nematode species/strains, and more importantly, between strains belonging to the same species. S. bibionis T335 was significantly better (P < 0.001) than S. bibionis N60 at each temperature tested, and H. heliothidis T327 was significantly better than H. heliothidis NC and NZ at 18 (P < 0.001) and 23 C (P < 0.001). However, at 28 C, there was no significant difference between the three strains of H. heliothidis. At 28 C, Heterorhabditis sp. Dl was more effective (P < 0.001) than any other nematode, and H. heliothidis T327 was the most effective nematode below 28 C (P < 0.05). At 18 C, Heterorhabditis sp. V16, H. heliothidis NC and T327, S. kraussei, and both strains of S. bibionis displayed optimum infectivity, whereas H. heliothidis NZ was more effective at 23 C.
Heterorhabditis spp. Dl and Q380 were most effective at 28 C. In contrast, there was no significant difference in the infectivity of H. bacteriophora, S. glaseri, and S. feltiae for L. cuprina between 18 and 28 C. The steinernematids produced infective juveniles in the majority of L. cuprina cadavers resulting from exposure to dosages of 10’ to lo3 nematodes at 18 and 23 C but no reproduction occurred at 28 C. At 28 C, the majority of Heterorhabditis spp. reproduced effectively in all cadavers resulting from exposure to dosages of loo to 102.5infective juveniles with occasional reproduction at dosages of 103. However, H. heliothidis strain T327 did not reproduce at any dosage at 28 C. At 23 C, all Heterorhabditis spp. tested reproduced in cadavers at dosage rates of loo to 102.5 infective juveniles. However, at 18 C, Heterorhabditis spp. Dl and Q380 did not reproduce at any dosage in L. cuprina cadavers and H. bacteriophora reproduced only occasionally. The remaining heterorhabditids reproduced at 18 C in all cadavers
Heterorhabditis, Steinernema SPP.: ASPECTS OF BEHAVIOR
TEMPERATURE C
FIG. 1. Parasitization of postfeeding, 3rd instar larvae of Lucilia cuprina exposed individually to dosages of 320 heterorhabditid.(A, B) or steinernematid (C, D) infective juvenile nematodes in 80 g sand (0.03 bars) at various temperatures. O-O, Heterorhabditis sp. Dl; X---X, Heterorhabditis sp. Q380; A- --A, Heterorhabditis sp. V16; *----Q, Heterorhabditis sp. T310; 0- - -0, H. heliothidis T327; n - - 4, H. heliothidis NC; A-A, H. heliothidis NZ; m---M, H. bacteriophora; f~- - -6, Steinernema sp. Wl; X-X, Steinernema sp. Wll; A- - -A, S. bibionis N60; O---O, 5’. bibionis T33.5; A-A, S. feltiae Agriotos; U-0, S. glaseri KG; l - - -0, S. kraussei; q - - -0, steinernematid species Ql.
resulting from dosages of loo to 102.5infective juveniles except H. heliothidis T327 which reproduced only in cadavers resulting from dosages of loo to lo2 infective juveniles. At dosage rates above 102.s,no production of Heterorhabditis spp. infec-
173
tive juveniles occurred at 18, 23, and 28 C. At those dosages, the characteristic pigmentation normally produced by the bacterial symbiont was absent and cadavers were invariably fetid. Although the temperature range supporting nematode reproduction inside L. cuprina and Galleria mellonella was similar, the steinernematids reproduced at higher temperatures inside larvae of G. mellonella. Of the four nematode species studied in detail, parasitization occurred over a greater range of temperatures than the temperature range allowing nematode reproduction, and substantial differences in the production of infective juveniles in G. mellonella occurred between the different nematode species. Heterorhabditis sp. Dl produced infective juveniles over the narrowest temperature range (20-32 C), whereas S. glaseri produced infective juveniles across a temperature range of 12 to 32 C. Heterorhabditis sp. Dl did not reproduce at 18 C and only developed partially to the L4 stage within 9 days after parasitization of the host. At temperatures below 10 C, H. heliothidis T327, S. glaseri, and S. feltiae did not reproduce and no live nematodes were recovered at the end of 14 days. At 10 C, S. glaseri and S. feltiae molted to the L4 stage and H. heliothidis T327 developed into first generation adults, but died after 4 weeks without further development. At 12 C, a few infective juveniles emerged from approximately 10% of cadavers infected with S. glaseri or H. heliothidis T327 after 8 weeks and after 10 weeks from cadavers infected with S. feltiae. At 28 and 32 C, only first generation adults were produced in cadavers infected with H. heliothidis and S. feltiae, respectively. At 35 C, S. glaseri developed to first generation adults but failed to reproduce while Heterorhabditis sp. Dl did not develop. The time taken between parasitization of the insect and emergence of the newly formed infective juveniles from the cadaver (measured as rate of development) was dif-
174
A. S. MOLYNEUX 380
1%
Y = 1.436.x
+ 0.176
-.-
23c
Y = 1.472.x
+ 1.148
--o--
28C
Y = 0.839X
+ 2.739
..
.o.....
99 :i .‘/ :ix .’ /
0
0
7
..‘/ .’ / :’ ,
90 6
70 5
50 30
4 10 3
:
: :’ .’
2
Zl z a
B
18C
Y = L.242.7 + 2.299
2%
Y = i.062.x
+ 2.508
-.--n--
28C
Y ,= 0.958X
+ 2.813
. . . ..o.....
16
,
/
, /
18C
Y = 0.840X
+ 3.572
23c
Y = 0.801X
+ 3.267
-.--mm-
28C
Y = 0.806X
+ 3.082
. . . ..o...
2 CA0 99 7
90 6
70 50 30
10
1 10"
lo=
lo3
10'
loo
DOSAGE (No.
10'
lo2
ld"
10'
NEMATODES)
FIG. 2. Dosage/parasitization lines for postfeeding, 3rd instar larvae of Lucilia cup&a exposed to infective juveniles of Heterovhabditis sp. Dl (A), Heterorhabdifis sp. Q380 (B), H. bacteriophora (C), and Heterovhabditis sp. V16 (D) in sand (0.03 bars) at 18, 23, and 28 C.
Heterorhabditis, Steinernema SPP. : ASPECTS OF BEHAVIOR :
175
1%
Y = 1.344x
+ 2.772
-.-
18C
Y = 0.947x
+ 2.613
23C
Y = ,.*59x
+ 2.560
--a--
2x
Y = 1.206.x
+ 2.470
--o--
28C
Y = 0.920x
+ 2.948
. . . ..o.....
28C
Y = 1.14*x
+ 2.416
.....o.....
12
-.-
99 7
so 6 70 5
50 30
4 10 E 2 4 F: s LI
3 1 327
-.-
18C
Y = ,.340x
+ 3.561
23C
Y = 1.188X
+ 3.458
--m--
2%
Y = 0.927x
+ 2.854
. . . ..o.....
23C
Y = 0.799x
--•--
+ 2.421
if o,n SE 7
9c 6 7C 5
5c 3(
4 l( _’
C
_’_’
3 I 10'
10=
103
lo4 DOSAGE
(No.
loo
10:
10=
10"
lo4
NEMATODES )
FIG. 3. Dosageiparasitization lines for postfeeding, 3rd instar larvae of Lucilia cuprina exposed to infective juveniles of Heterorhabditis heliothidis NC (A), H. heliothidis NZ (B), H. heliothidis T327 (C), and Steinernema kruussei (D) in sand (0.03 bars) at 18, 23, and 28 C.
2 z -2
176
A. S. MOLYNEUX MO
1%
Y -
23c
Y - 0.870X + 2.802
1.291.x
+ 2.424
*WC
Y = 0.438X
-.-
r335
---o-. . . ..o.....
+ 3.082
18C
Y = 1.757x
+ 2.7cm
-.-
2x
Y = 1.3761(
+ 2.785
--•.--
28C
Y = 1.237): + 2.291
. . ..a.....
99 7
so 6 70 50
5
30 4
10 5 2 2 s 2
3
l
1%
Y = 1.167X + 3.067
23C
Y = 1.300x
+ 2.898
28C
Y = 1.,19x+
2.824
-.--n--
18C
Y = 0.852X
23C
Y = 0.918X + 2.942
. . . ..a....
28C
Y = 0.953x
-.-
+ 2.809
--a-.o..
+ 2.687
2 D 99
co 7
so 6
70 50
5
30 4
10
3
1 lo9
I
lOI
I
lo2
I
I
lo3
I
lo4 loo 10' DOSAGE (No. NEMATODES)
lo2
lo3
lo4
FIG. 4. Dosage/parasitization lines for postfeeding, 3rd instar larvae of Lucike cuprina exposed to infective juveniles of Steinevnema bibionis N60 (A), S. bibionis T33S (B), S. glasevi KG (C), and 5’. feltiae Agriotos (D) in sand (0.03 bars) at 18, 23, and 28 C.
2 1 & PI
177
Heterorhabditis, Steinernema SPP.: ASPECTS OF BEHAVIOR TABLE III Effect
of Temperature
Nematode
on Infectivity
Strain
Heterorhabditis sp. Heterorhabditis sp. Heterorhabditis sp. H. bacteriophora H. heliothidis H. heliothidis H. heliothidis Steinernema glaseri S. bibionis S. bibionis S. feltiae S. kraussei
Dl Q380 V16
NC NZ T327 KG T335 N60 Agriotos
of
Heterorhabditis
spp. and
18 c
571 4,080 50 149 45 331 12 45 20 99 372 213
402-827 2,760-6,860 30-78 106-208 31-63 223-497 7-17 30-65 14-28 70-136 243-580 149-303
Steinernema
spp. for
23 C
76 385 145 222 87 125 20 41 41 336 174 1,710
LDSO, 95% limits 52- 107 272-562 92-227 153-321 64-116 88-175 13-29 29-57 30-54 221-519 121-249 1,040-3,160
28 C
32 222 239 192 170 183 206 88 155 23,900 267
25-39 157-318 167-342 145-254 123-233 138-240 148-284 63-120 129-186 10,600-75,000 199-366
b
LD,,
Heterorhabditis sp. Heterorhabditis sp. Heterorhabditis sp. H. bacteriophora H. heliothidis H. heliothidis H. heliothidis Steinernema glaseri S. bibionis S. bibionis S. feltiae S. kraussei a Individual b Mortality
DI Q380 V16
NC NZ T327 KG T335 N60 Agriotos
7,820 46,700 1,670 1,610 408 7,480 107 569 109 972 11,900 2,950
4,450-17,100 22,300-143,000 912-3,810 1,020-2,970 268-711 3,960-18,000 69-189 359- 1,050 76- 179 643-1,670 5,750-33,000 1,770-5,980
941 8,370 5,790 3,580 905 1,440 239 400 347 10,000 4,320 68,900
95% limits 590- 1,750 4,510-19,300 2,880-15,300 2,080-7,460 597- 1,580 906-2,690 152-434 270-669 238-564 4,980-26,700 2,500-8,840 25,500-31,400
Lucilia cuprina”
609 5,920 9,300 4,190 4,190 2,420 4,970 1,230 1,690 2.01 x 10’ 5,890
446-881 3,270-13,100 5,220-19,600 2,700-7,250 2,570-7,800 1,610-4,060 3,030-9,300 815-2,070 1,260-2,400 3.2 x 106-3.0 x lo* 3,440-11,820
b
larvae in 80 g sand, moisture content 7% CO.03 bars). did not reach 50% at any dosage.
ferent for each nematode species tested at movement of the majority of steinernevarious temperatures (Fig. 5). S. glaseri matid infective juveniles may reflect the completed its life cycle in the fastest time colder climates of their European and and H. heliothidis had generally the slow- North American origins (Table II). Alest rate of development although the devel- though both strains of Steinernema biopmental rate of S. feltiae was the slow- bionis were isolated from soils or insects est of the four species at 12 C (Fig. 5). For taken from temperate areas of Australia each nematode species tested, second gen- (Table I), they were still mobile at 4 C. Aceration infective juveniles emerged from cording to Bedding (personal communicacadavers of G. mellonella. The relationship tion), it is likely that S. bibionis has been between developmental rate and tempera- introduced into Australia, most probably ture appeared to be linear, and the calcu- from Europe. In contrast, the inability of lated theoretical threshold temperatures for the heterorhabditids Dl and Q380 to move development gave a close approximation to at 10 and 14 C, respectively, suggests that the observed lower temperature limits for they are native to the warm, humid tropical development. areas. The variability in infectivity for Lucilia DISCUSSION cuprina larvae is, in part, related to the Important differences between the hete- length of time the insect host remdins susrorhabditids and steinernematids were ob- ceptible to nematode parasitization at difserved in the bioassays at various tempera- ferent temperatures (Molyneux 1984). Untures. The lower temperature limits for like lepidopteran pupae (Kaya and Hara
178
A. S. MOLYNEUX 0.3 -
l
Y=O.O07X-0.093,r=0.989,
13.S"
oY=O.O04X-O.O37,r-0.989,
8.3'
*Y=O.O07X-0.071,r=0.996,
9.9O
q Y=O.O13X-0.153,r=0.979,
11.8
0.2 -
01 0
I
I
I
5
10
15
I
I
I
20 25 M TEMPERATURE C
35
40
I
I
FIG. 5. Average rate of development for completion of life cycle of 0 -0, Heterorhabditis sp. Dl; 0 -0, H. heliothidis T327; k -*, Steinernema feltiae Agriotos; q -------a, S. glaseri KG in final instar Galleria mellonella cadavers at various temperatures. r, Correlation coefficient; a, calculated developmental threshold temperature C
1981; Moyle and Kaya 1981), L. cuprina larvae become almost completely invulnerable to parasitization soon after pupariation (Molyneux 1984). The greater infectivity of some nematodes (e.g., Heterorhabditis sp. Dl) at 28 C than at 18 C suggests that the increased activity of nematodes at the higher temperatures more than offsets any increase in the rate of pupariation of L. cuprina. The relationship between nematode activity and insect susceptibility is not peculiar to L. cuprina alone. Many of these insects that pupate in soil are probably similarly influenced, and other insects not pupating may be less active and thus become vulnerable to nematode parasitization. With all nematodes tested in this study, the range of temperature allowing parasitization was greater than the temperature range supporting nematode reproduction. This is of little benefit to the nematode at
the upper end of the temperature range because thermal damage soon resulted (unpublished data). However, at lower temperatures, some nematode species will remain in the hosts’ hemocoel as infective juveniles (unpublished data). There they can remain without killing the host until temperatures rise allowing bacterial growth and nematode reproduction. In the meantime, depending on the insect species, the insect may migrate and thus disseminate the nematode. The failure of Steinernema spp. to reproduce in L. cuprina at 28 C (Molyneux et al. 1983) like the failure of both genera when applied in high dosages at lower temperatures is apparently due to the contamination of the cadaver by foreign microorganisms (Molyneux 1984). The introduction of a high inoculum of gut bacteria, particularly by Steinernema spp. penetrating Lucilia cuprina larvae via the anus, can result in failure of the nematode’s bacterial symbiont to dominate within the host hemocoel. Unlike Steinernema spp., Heterorhabditis spp. infective juveniles can penetrate the intersegmental membranes of their insect hosts thereby avoiding the gut bacteria and reducing the risk of contamination. Furthermore, the outer cuticle is shed just before or during penetration, enabling Heterorhabditis species to enter with a surface that is largely bacteria free (Bedding and Molyneux 1982). Reproduction of steinernematids at temperatures of 28 C or more in Galleria mellonella larvae is probably the result of fewer foreign microorganisms being introduced due to nematode entry via the spiracles in addition to the anus and mouth. In agreement with previous observations (Jackson 1962; Kaya 1977; Poinar 1979), the most favorable temperature for the growth and reproduction of S. feltiae and S. glaseri lies between 23 and 28 C although the temperature limits for reproduction of S. feltiae Agriotos exceed those reported by Kaya (1977) and Pye and
Heterorhabditis, Steinernema
SPP.: ASPECTS
OF BEHAVIOR
179
otically associated with the insect pathogenic nemaBurman (1978) for the DD136 and Agriotos todes Neoaplectana and Heterorhabditis. Journal strain of Steinernema feltiae, respectively. of General Microbiology 121, 303-309. In contrast, the most favorable tempera- AKHURST, R. J. 1982. Antibiotic activity of Xenotures for growth and reproduction of Hetrhabdus spp., bacteria symbiotically associated erorhabditis sp. Dl and H. heliothidis with insect pathogenic nematodes of the families Heterorhabditidae and Steinernematidae. Journal T327 fell outside this range (30 and 20 C, of General Microbiology 128, 3061-3065. respectively) reflecting once again the cliAKHURST, R. J. 1983. Neoaplectana species: Speckmatic zones of their original locality. The ficity of association with bacteria of the genus Xedecline in nematode reproduction at high norhabdus. Experimental Parasitology 55, temperatures may be the result of competi258-263. tion with its own bacterial symbiont which ALVEY, N. G., et al. 1977. Regression and generalised linear models. In “GENSTAT: A General Statismay explain why Jackson (1962) reported tical program,” Chap. 7. Rothamsted Experimental minimal reproduction of S. glaseri at 35 C Station, England. in axenic liquid media. In addition, thermal BEDDING, R. A., AND AKHURST, R. J. 1975. A simple damage may result in a lack of viable technique for the detection of insect parasitic rhabditid nematodes in soil. Nematologica 21, 109- 110. sperm or ova or a change in the mating beBEDDING, R. A., AND MOLYNEUX, A. S. 1982. Penehavior of the nematode (Kaya 1977). At tration of insect cuticle by infective juveniles of Helow temperatures, the lack of nematode reterorhabditis spp. (Heterorhabditidae: Nematoda). production is probably a direct effect of Nematologica 28, 354-359. BEDDING, R. A., MOLYNEUX, A. S., AND AKHURST, temperature on nematode maturation. AkR. J. 1983. Heterorhabditis spp., Neoaplectana hurst (1980) reported that the primary form spp., and Steinernema kraussei: Interspecific and of Xenorhabdus spp. is essential for opintraspecific differences in infectivity for insects. timum nematode reproduction. The relaExperimental Parasitology 55, 249-257. tively few infective juveniles of SteinerBURMAN, M., AND PYE, A. E. 1980. Neoaplectana carpocapsae: Movements of nematode populations nema glaseri produced per cadaver at difon a thermal gradient. Experimental Parasitology ferent temperatures may have been due to 49, 258-265. the absence of the primary form of its bacFINNEY, D. J. 1971. “Probit Analysis,” 3rd ed. Camterial symbiont in the KG strain. bridge University Press, London. The results from these studies indicate GEORGIS,R., AND POINAR, G. 0. 1983a. Effect of soil texture on the distribution and infectivity of Neothat when selecting nematodes as possible aplectana glaseri (Nematoda: Steinernematidae). control agents for a particular pest insect Journal of Nematology 15, 329-332. nematode dosage rates, species of nemaGEORGIS,R., AND POINAR, G. 0. 1983b. Effect of soil tode, and insect type will need to be contexture on the distribution and infectivity of Neosidered in relation to temperature for their aplectana carpocapsae (Nematoda: Steinernematidae). Journal of Nematology 15, 308-3 11. permanent establishment in the field. ACKNOWLEDGMENTS
I thank Dr. R. A. Bedding for encouragement and guidance throughout this study; J. G. Moss, V. S. Patel, M. A. Stanfield, and W. H. Edwards for valuable technical assistance; Dr. D. A. Ratkowsky and R. Lowry for statistical advice; and Professor H. R. Wallace, Department of Plant Pathology, Waite Agricultural Research Institute, University of Adelaide, for constructive criticism of the manuscript. REFERENCES AKHURST, R. J. 1980. Morphological and functional dimorphism in Xenorhabdus spp., bacteria symbi-
G~~Tz,P., BOMAN, A., AND BOMAN, H. G. 1981. Interactions between insect immunity and an insectpathogenic nematode with symbiotic bacteria. Proceedings of the Royal Society of London Series B 212, 333-350.
JACKSON, G. J. 1962. The parasitic nematode, Neoaplectana glaseri in axenic culture, II. Initial results with defined media. Experimental Parasitology 12, 25-32. JACKSON, G. J. 1973. The ageing of Neoaplecfana glaseri. Proceedings of the Helminthological Society of Washington 40, 74-76. KAYA, H. K. 1977. Development of the DD-136 strain of Neoaplectana carpocapsae at constant temperatures. Journal of Nematology 9, 346-349.
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MOLYNEUX, A. S. 1984. The influence of temperature on the infectivity of heterorhabditid and steinernematid nematodes for larvae of the sheep blowfly, Lucilia cuprina. In “Proceedings of the Fourth Australian Applied Entomological Research Conference” (P. T. Bailey and D. W. Swincer, eds.), pp. 344-351. Govt. Printer, Adelaide. MOLYNEUX, A. S. Survival of infective juveniles of Heterorhabditis spp., and Steinernema spp. (Nematoda: Rhabditida) at various temperatures and their subsequent infectivity for insects. Revue de Nematologie, in press. MOLYNEUX, A. S., AND BEDDING, R. A. 1985. Influence of soil texture and moisture on the infectivity of Heterorhabditis sp. Dl and Steinernema glaseri for larvae of the sheep blowfly, Lucilia cuprina. Nematologica 30, 358-365. MOLYNEUX, A. S., BEDDING, R. A., AND AKHURST, R. J. 1983. Susceptibility of larvae of the sheep blowfly Lucilia cuprina to various Heterorhabditis spp., Neoaplectana spp., and an undescribed steinernematid (Nematoda). Journal of Invertebrate Pathology 42, l-7.
MOYLE, P. L., AND KAYA, H. K. 1981. Susceptibility of pupae of two cocoon-forming lepidopterous species to the entomogenous nematode Neoaplectuna carpocapsae (Rhabditida: Steinernematidae) Journal of Nematology 13, 419-421. POINAR, G. 0. 1979. “Nematodes for Biological Control of Insects.” CRC Press, Boca Raton, FL. PYE, A. E., AND BURMAN, M. 1978. Neoaplectana carpocapsae: Infection and reproduction in large pine weevil larvae, Hylobius abietis. Experimental Parasitology 46, 1- 11. SCHMIEGE, D. C. 1963. The feasibility of using a neoaplectanid nematode for control of some forest insect pests. Journal of Economic Entomology 56, 427-431. WHITEHEAD, A. G., AND HEMMING, J. R. 1965. A comparison of some quantitative methods of extracting small vermiform nematodes from soil. Annals of Applied Biology 55, 25-38. WOUTS,W. M., MR&?EK,, Z., GERDIN, S., AND BEDDING, R. A. 1982. Neoaplectana Steiner, 1929 a junior synonym of Steinernema Travassos, 1927 (Nematoda: Rhabditida). Systematic Parasitology 3, 147-154.