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Open Access 04-03-2023

Susceptibility of white grubs from forestry and sugarcane plantations in South Africa to entomopathogenic nematodes

Authors: Agil Katumanyane, Bernard Slippers, Mesfin Wondafrash, Antoinette P. Malan, Brett P. Hurley

Published in: BioControl | Issue 2/2023

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Abstract

The control of white grub (Coleoptera: Scarabaeidae) pests of sugarcane and forest plantations is difficult due to their cryptic nature and resistance to chemicals. This study evaluated the potential use of entomopathogenic nematodes (EPNs) as an alternative control method. Laboratory bioassays were performed with 12 locally isolated EPN species to determine the susceptibility of third instar larvae of the white grubs Schizonycha affinis Boheman, Pegylis sommeri Burmeister, Monochelus sp. and Maladera sp. 4. Concentration trials to determine lethal dosages for three of the white grub species were performed using Heterorhabditis zealandica Poinar MJ2C. Bioassays were performed to determine whether nematodes could develop inside the cadavers of S. affinis and P. sommeri as these had shown the lowest susceptibility to EPNs. The mortality percentage of the white grubs, although varying significantly, was found to be low for most of the EPN species, except H. zealandica. The highest percentage mortality of white grubs was observed at four weeks post inoculation with the rate of mortality being highest in the first week. Schizonycha affinis had the lowest LD50 of 38 Infective juveniles (IJs) per larva after 28 days, compared to Maladera sp. 4, with 284 IJs per larva, and P. sommeri, with 1035 IJs per larva. The dissection of insect cadavers revealed possible limiting factors for low susceptibility, due to the nematodes and their associated symbiotic bacteria’s inability to infect the insect haemocoel, with no EPNs being found inside some of the cadavers.
Notes
Handling Editor: Ralf-Udo Ehlers.

Introduction

White grubs (Coleoptera: Scarabaeidae) are the root-feeding larvae of Scarabid beetles which are sporadic pests of various crops (Ritcher 1966; Jackson and Klein 2006). These larvae feed on plant roots, resulting in significant root damage. The adults tend to feed on plant leaves or to bore into underground stems (Jackson and Klein 2006). In South Africa, white grubs are the main insect pests of sugarcane and important establishment pests in plantation forestry (Echeverri-Molina and Govender 2016a, b; Sivparsad et al. 2018). Govender (2007, 2014) reported that white grubs were responsible for 13% of the damage caused in black wattle plantations in the KwaZulu-Natal (KZN) province, while McArthur and Leslie (2004) reported an average of 23–55% reduction in sugarcane yield in the KZN Midlands North area. Forestry and sugarcane plantations are often in proximity and an overlap of white grub species affecting these two crops has been observed (Sivparsad et al. 2018).
The control of white grubs is difficult mainly because of their soil-dwelling nature, their resistance to chemical insecticides (Grewal et al. 2005) and the nocturnal lifestyle of the adults (Jackson and Klein 2006). While white grub control efforts have been dominated using chemical insecticides, the use of entomopathogenic nematodes (EPNs) offers a relatively safe and efficient control option (Grewal et al. 2005; Koppenhöfer et al. 2020). EPNs (Rhabditida: Steinernematidae and Heterorhabditidae) are known to infect various below- and above-ground insects (Grewal et al. 2005; Lacey and Georgis 2012). The two EPN families are used worldwide in the biological control of various insect pests (Shapiro-llan et al. 2010; Lacey and Georgis 2012). EPNs kill their insect hosts using symbiotic bacteria, which they carry in their digestive system and in specialised bacterial chambers (Nobuyoshi 2002). Death of the insect normally occurs within 48 h after infection (Kaya et al. 1993; Nobuyoshi 2002).
South Africa has a diversity of native EPNs, with a total of 17 Steinernema and seven Heterorhabditis species reported by 2017 (Malan and Ferreira 2017). A recent description of Steinernema bertusi Katumanyane, Tiedt, Malan & Hurley brought the total of Steinernema species described from South Africa to 12, constituting 12% of the described species of this genus in the world (Katumanyane et al. 2020). Despite the diversity of native EPNs in South Africa and their potential as biological control agents for white grubs, so far only one study has been undertaken to evaluate the potential use of EPNs to control white grubs in South Africa. Abate et al. (2019) evaluated the efficacy of both native and non-native EPNs to control the white grub, Heteronychus licas Klug in sugarcane plantations. There is need to evaluate the use of EPNs on more white grub pests in sugarcane and plantation forests in South Africa.
In the current study, we screened locally isolated EPNs for their biocontrol potential against native white grubs from forestry and sugarcane plantations in South Africa. More specifically, we (1) screened twelve locally isolated EPN species for their biocontrol potential against four most dominant white grub species, (2) determined whether (and how) mortality rate of white grub species varied with infected EPN species and over time, (3) tested different concentrations of EPN to determine the lethal doses needed, and (4) examined the potential for different EPNs to infect, kill and reproduce in two of the most resistant white grub species.

Materials and methods

Source of white grubs

Four white grub species, namely Schizonycha affinis Boheman, Pegylis sommeri Burmeister, Maladera sp. 4 and a Monochelus sp. were used in this study. Insect identification was done by means of a LUCID key, previously developed for white grubs in sugarcane plantations in southern Africa:https://​keys.​lucidcentral.​org/​keys/​v3/​sugarcane_​white_​grubs/​sugarcane_​white_​grubs.​html. The white grubs were collected from wattle and sugarcane plantations in the KwaZulu-Natal province of South Africa. Each white grub was placed in a 30 ml plastic vial, filled with moist autoclaved peat moss (Hygrotech sustainable solutions) and provided with a fresh carrot disc to feed. The vials containing white grubs were placed in cooler boxes for transport and later rearing at the Forestry and Agricultural Biotechnology Institute (FABI) Biocontrol Centre at the University of Pretoria, South Africa. Rearing was done at 23 °C and 60–70% RH. The autoclaved peat and carrot discs were replaced weekly. Only third instar larvae were used in the experiments. The grubs were only used three weeks onwards after collection to eliminate those that could have come infected from the field.

Source of EPNs

Twelve EPN species namely Heterorhabditis bacteriophora Poinar strain SF351, Heterorhabditis baujardi Phan, Subbotin, Nguyen & Moens strain BA, Heterorhabditis indica Poinar, Karunakar & David strain SGS, Heterorhabditis noenieputensis Malan, Knoetze & Tiedt strain SF669, Heterorhabditis safricana Malan, Nguyen, de Waal & Tiedt strain SF281, Heterorhabditis zealandica Poinar strain MJ2C (green), H. zealandica strain SF41 (blue), Oscheius myriophila Poinar strain AK30, S. bertusi Katumanyane, Tiedt, Malan & Hurley strain AK27, Steinernema fabii Abate, Malan, Tiedt, Wingfield, Slippers & Hurley strain ML15, S. fabii strain SCH10, Steinernema jeffreyense Malan, Knoetze & Tiedt strain J194, Steinernema sacchari Nthenga, Knoetze, Berry, Tiedt & Malan strain DUK and Steinernema yirgalemense Nguyen, Tesfamariam, Gozel, Gaugler & Adams strain 157-C were used in this study. The nematode isolates were obtained from the nematode collections at the FABI Biocontrol Centre, University of Pretoria and the Department of Conservation Ecology and Entomology at Stellenbosch University, Stellenbosch, South Africa. All nematodes used were reared using the third instar larvae of the greater wax moth, Galleria mellonella L. (Lepidoptera: Pyralidae). Infective juveniles (IJs) were harvested on modified White’s traps and stored in distilled water in horizontally placed culture flasks at 12 °C. The EPNs were shaken periodically for ventilation and used within three weeks after harvesting. A fresh batch of nematodes was used for each repetition of the experiment.

Screening bioassays

Soil bioassays

The methods used by Koppenhöfer and Fuzy (2003), An et al. (2012) and Wu et al. (2014) were modified and used in the present experiment. The EPNs H. bacteriophora, H. zealandica-MJ2C, S. fabii, S. jeffreyense and S. yirgalemense were separately tested for their potential to kill the third instar larvae of P. sommeri, Monochelus sp., Maladera sp. 4. Two additional species, namely S. sacchari and a strain of S. fabii-SCH10, were further tested against the larvae of S. affinis.
For the setup, 30 ml plastic cups were filled with 30 g autoclaved sandy loam soil. A white grub was then placed on top and allowed to dig itself into the soil, and the soil was inoculated with the EPNs. From a nematode suspension, 400 IJs were sprayed onto the surface of the soil in each cup and the soil moisture was adjusted to 18% v/w. A carrot disc was provided for white grub feeding. The vials were then placed in a plastic container lined with moist tissue paper (100% moisture) and stored at 25 °C in a dark room. The experiment included five treatments (EPN species), and each was tested on ten white grubs. This was replicated three times. Each replicate was undertaken on a different test date using a different batch of nematodes. The cumulative percentage mortality for each treatment was recorded over a period of one month, at weekly intervals. For S. affinis, two additional EPN species were tested.

12-well laboratory bioassays

Based on the preliminary results of the soil bioassays, the most resistant white grubs, namely S. affinis and P. sommeri, were subjected to further tests in 12-well bioassay plates. The aim was to investigate any changes that occurred in the susceptibility of the white grub species, when they were directly exposed to the EPNs. The susceptibility of the two white grub species to nine EPNs, namely H. bacteriophora, H. baujardi, H. indica H. noenieputensis, H. safricana, H. zealandica-MJ2C, H. zealandica-SF41, O myriophila and S. fabii-SCH10, was tested. Additionally, the susceptibility of S. affinis to S. bertusi was also tested.
For the experimental setup, a single white grub was placed in a cell of a 12-well bioassay plate lined with a filter paper disc. It was inoculated with 200 IJs suspended in 50 µl distilled water. Ten grubs of each white grub species were used for each of the treatments plus the control. The controls received water only. The plates were placed in a plastic container lined with wet paper towels (at 100% moisture level), closed with the lid, and kept in a growth chamber, adjusted at 25 °C. Mortality reading was taken daily for up to seven days for S. affinis and for up to ten days for P. sommeri, as the mortality for the latter was observed to take longer than for the former. Any dead larvae were removed, rinsed with distilled water and placed in a new 12-well plate lined with moist filter paper. Dead larvae were returned to the growth chamber to allow any EPNs, if present, to develop. All dissections of the cadavers, aimed at confirming the presence of EPNs, were performed on the 10th day post-inoculation (PI). Only those cadavers that were observed to have EPNs present were counted as part of the mortality (equal infection) data. The experiment was replicated three times, on a different test date, for each nematode species, and using a fresh batch of nematodes.

Concentration trial

Based on the preliminary results obtained with both the soil and the 12-well bioassay disk (Figs. 1, 2, 3), H. zealandica MJ2C was selected for the probit test to determine its lethal dosages (LD50) on the third instar larvae of P. sommeri, S. affinis and Maladera sp. 4. Monochelus sp. was not included in the experiment, as its larvae were not available in sufficient numbers. Similar methods to those that were used in the soil bioassay were employed in this experiment. However, the treatments included different concentrations of H. zealandica MJ2C, namely 0, 100, 200, 400, 800 and 1600 IJs per vial. Ten white grubs were used in each treatment (EPN concentration × ten white grub species), with the experiment being replicated three times, and with each replicate being undertaken on a different test date, using a different batch of nematodes. Cumulative percentage mortality was recorded at weekly intervals over a period of one month.

Examining EPN survival and development inside insect cadavers

The ability of EPNs to develop in the insect cadavers of S. affinis and P. sommeri was tested using H. safricana, H. noenieputensis, H. indica, H. zealandica-MJ2C, H. baujardi, S. fabii-SCH10, H. bacteriophora, H. zealandica-SF41, O. myriophila and S. bertusi. The aim was to test the survival and development of the nematodes to gain improved understanding of the mechanisms of resistance employed by the two most resistant grub species, namely S. affinis and P. sommeri, as observed in the soil and 12-well bioassays. Similar methods to those used in the 12-well bioassays were employed in the experiment. Grubs were individually placed in 90 mm diam. Petri dishes and inoculated with 200 IJs of the nematodes. Ten grubs each of S. affinis and P. sommeri were used per EPN treatment. The inoculated insects were incubated at 25 °C, in containers lined with moist tissue paper. Insect mortality was recorded daily for a period of ten days. Irrespective of the day of insect death, all dissections of dead insects were performed on the 10th day PI to be able to observe and record the status of the EPNs developing inside the haemocoel. The presence or absence of the EPNs and the symbiotic bacteria, the development stage of the EPNs, and the general state of the cadaver were recorded. The EPN symbiotic bacteria colonisation was visually observed in the haemolymph for characteristics typical to a successful entomopathogenic symbiotic bacteria (ESB) colonisation which included gumminess of the bacterial body mass for Photorhabdus bacteria, colour changes, absence of a putrid smell. The limitation with visual observation for ESB could lie in the inability to observe these characters in the presence of a supressed ESB colonization.

Statistical analysis

All statistical analyses were performed using the R statistical software, version 4.00 (R Development Core Team 2020). To determine whether mortality rates of white grub species (percentage of white grubs that died during the experiment) varied with infected EPN species and over time, we used generalised linear models (GLM) with a binomial distribution and a logit link function (Zuur et al. 2009), which assessed the main and interaction effects of time and EPN species. In order to better depict the trends in these effects, the variation in mortality among treatments (EPN species) was analysed and represented using bar plot (with error bars representing SE) for each time interval (7, 14, 21 and 28 days). Due to the limited number of observations for each time interval, the Kruskal–Wallis non-parametric tests were performed to compare treatments (EPN species) for each time interval.
To test for the effect of time and different concentrations of EPN on mortality of white grubs, and determine the lethal doses needed, the log-logistic dose–response model (Eq. 1) was fitted with the ‘drm’ function of the package drc (Ritz et al. 2015) in R:
$$M(t)=\frac{1}{1 + exp (b\cdot log(t)-log(\mathrm{LD}_{50}))}$$
(1)
where M(t) is the expected mortality rate at time t (number of weeks); b is proportional to the slope of M, at time t; and LD50 is the concentration dose by which 50% of the white grubs would have died.

Results

Screening bioassays

Schizonycha affinis

The results of the soil bioassays showed that the mortality rate of S. affinis varied significantly among different EPN species (χ2 = 167.48; df = 7; p < 0.001) and over time (χ2 = 29.42; df = 3; p < 0.001). However, we found no interaction effects between EPN species and time (χ2 = 8.73; df = 21; p = 0.991). Schizonycha affinis mortality increased over time for all treatments (Fig. 1a). Its highest mortality rate was constantly observed with H. zealandica MJ2C (Fig. 1a). The maximum percentage mortality of 70.0% ± 1.2%, which was obtained after 21 days, was significantly different from the next closest efficient species, S. jeffreyense, at 43.0% ± 1.2%, and S. yirgalemense, at 30.0% ± 1.2%. Overall, H. bacteriophora, S. fabii (BA), S. sacchari and an isolate of S. fabii (SCH10) showed very low mortality rates of less than 20% after 28 days. For all the tested nematodes, the mortality graph levelled off after 21 days (Fig. 1a). In keeping with the results obtained in the soil bioassays, the highest percentage mortality for the 12-well laboratory bioassays was obtained from H. zealandica MJ2C at 63.0% ± 2.2%, seven days PI. Heterorhabditis zealandica SF41 gave the second highest mean mortality at 46.7% ± 3.3%, followed by Oscheius myriophila Poinar, at 40.0% ± 1.5%. The rest of the tested nematodes resulted in less than 30% mortality, seven days PI (Fig. 1b).

Pegylis sommeri

The results of the soil bioassay showed that mortality percentage varied significantly among the different EPN species investigated (χ2 = 214.42; df = 5; p < 0.001), but not over time (χ2 = 0.66; df = 3; p = 0.883). Furthermore, there was no significant interaction effect between time and EPN species (χ2 = 6.19; df = 15; p = 0.976). These results suggest that the mortality did not increase over time. At 28 days, the mortality rates of P. sommeri were < 5% across all EPNs, except for in the case of H. zealandica MJ2C (Fig. 2a). Heterorhabditis zealandica MJ2C showed the highest percentage mortality, although it was only 30.0% ± 5.8%. Steinernema jeffreyense and S. yirgalemense caused no mortality. In the 12-well bioassay disks, P. sommeri still maintained a very high resistance towards the EPNs tested (Fig. 2b). The highest mortality of 26.7% ± 1.2%, was obtained using H. zealandica SF41.

Maladera sp. 4

Mortality rates of Maladera sp. varied significantly among the different EPN species (χ2 = 90.74; df = 5; p < 0.001), and over time for all EPN species (χ2 = 35.23; df = 3; p < 0.001). However, there was no interaction effects between EPN species and time (χ2 = 2.96; df = 15; p = 0.999). At 28 days PI, H. bacteriophora exhibited the highest percentages of mortality, 93.0% ± 6.7%, followed by H. zealandica MJ2C, with a percentage mortality of 76.7% ± 6.7% (Fig. 3a). Steinernema fabii and S. jeffreyense resulted in 63% ± 15% and 60% ± 10% percentage mortality, respectively, while the lowest mortality rate (40% ± 15%) was recorded for S. yirgalemense (Fig. 3a).

Monochelus sp.

The results of the soil bioassays for Monochelus sp. showed that its mortality also varied significantly among the different EPN species (χ2 = 50.71; df = 5; p < 0.001), and over time for all EPN species (χ2 = 25.55; df = 3; p < 0.001), but there was no significant interaction effect between time and EPN species (χ2 = 4.17; df = 15; p = 0.997). More specifically, we found that mortality rate increased over time but varied considerably among treatments (Fig. 3b). The highest mortality after 28 days of inoculation was recorded from H. bacteriophora (60.0% ± 11.5%) and S. jeffreyense (60.0% ± 15.3%) treated grubs. The average mortality of 50% ± 10%, caused by H. zealandica MJ2C, was closely followed by the average mortality caused by S. yirgalemense, at 47% ± 3%. The lowest level of mortality was caused by S. fabii, which attained a 30% ± 0% mean mortality on day 28 PI.

Concentration trial

Heterorhabditis zealandica MJ2C was used in the trial because it was the most efficient EPN for three of the white grub species, namely S. affinis, P. sommeri and Monochelus sp., in the screening bioassays. According to the results of the study, the LD50 was found to be generally lowest for S. affinis, at 38 IJs per grub, intermediate for Maladera sp. 4, at 284 IJs per grub, and highest for Pegylis sommeri, at 1035 IJs per grub, at 28 days PI (Table 1).
Table 1
Parameter estimates of the log-logistic dose response model showing the LD50 obtained for different white grubs, after inoculation with different concentrations of Heterorhabditis zealandica MJ2C: 0, 100, 200, 400, 800 and 1600 IJs
 
Schizonycha affinis
Pegylis sommeri
Maladera sp. 4
Est
SE
P
Est
SE
P
Est
SE
P
Slope
         
7 days
− 0.65
0.19
 < 0.001
− 0.99
0.26
 < 0.001
− 0.86
0.21
 < 0.001
14 days
− 0.38
0.17
0.029
− 1.21
0.26
 < 0.001
− 1.10
0.21
 < 0.001
21 days
− 0.39
0.18
0.031
− 1.23
0.25
 < 0.001
− 1.04
0.20
 < 0.001
28 days
− 0.53
0.21
0.011
− 1.30
0.25
 < 0.001
− 1.29
0.23
 < 0.001
LD50
         
7 days
762
235.39
0.001
2226
857.17
0.009
1152
329.65
 < 0.001
14 days
238
117.98
0.044
1369
315.44
 < 0.001
497
84.58
 < 0.001
21 days
74
63.47
0.241
1136
231.46
 < 0.001
319
56.91
 < 0.001
28 days
38
34.08
0.269
1035
191.72
 < 0.001
284
43.23
 < 0.001
Estimate slope refers to the parameter b in the log-logistic function; and LD50 is the dosage by which 50% of the white grub population would have died
Est estimate; P probability value testing the significance of the estimates

Examining EPN development inside the insect cadaver

Heterorhabditis zealandica MJ2C showed a high mortality potential for S. affinis, causing the highest number of infections (19 out of the 30 white grubs tested), and was able to complete its life cycle inside the haemocoel of the grubs (Table 2). It was also observed to be the only nematode species able to protect its cadavers from attack by unidentified mite species. The cadavers were observed to remain intact, not smelly, or semi-decomposed, typical of EPN infected cadavers. Oscheius myriophila showed a comparatively high mortality for S. affinis, killing, on average, four out of ten individuals and being able to complete its life cycle within a week (Table 2). However, O. myriophila was unable to protect any of its cadavers, resulting in most of them being putrid and full of mites. Whenever H. indica colonised and killed S. affinis, the EPNs were unable to develop inside the cadavers, with only bacteria being observable at the end of the experiment (Table 2). For H. bacteriophora and H. zealandica SF41, mostly entomopathogenic nematode symbiotic bacterial (ESB) colonisation was observed, with no surviving EPNs appearing to remain in the cadavers. Steinernema bertusi showed a low percentage mortality towards S. affinis, with all the infections recorded being successful but was unable to keep the cadavers clean. The rest of the EPN species tested showed very low mortality rates. For H. safricana, H. noenieputensis and S. fabii SCH10, no ESBs/EPNs were observed in the dead cadavers (Table 2).
Table 2
Dissection observations for Schizonycha affinis
EPN species
Days PI and cumulative number of dead larvae
Notes
1
2
3
4
5, 6, 7, 8, 9, 10
 
Heterorhabditis bacteriophora
0
7
8
9
9
Mostly ESB present but no successful EPNs
H. baujardi
0
0
1
4
5
Three successful EPN cycles. No ESB/EPNs observed for the rest of the dead grubs
H. indica
0
4
5
6
8
All life cycles, except for one, unsuccessful. ESB unable to develop in the haemocoel
H. noenieputensis
0
1
3
4
4
No ESB/EPNs observed in any of the dead grubs
H. safricana
0
0
0
1
1
No ESB/EPNs observed in any of the dead grubs
H. zealandica MJ2C
0
9
16
19
19
All colonisations successful and cadavers kept clean
H. zealandica SF41
0
5
7
12
14
Mostly ESB colonisations and dead EPNs, or no EPNs at all. Only two EPNs completed their life cycle. ESB was unable to keep a cadaver clean. Cadavers semi-decomposed and full of mites
Oscheius myriophila
5
10
11
12
12
Mostly successful colonisations but could not keep cadavers clean. The life cycle was completed within a short period of time, being less than seven days
Steinernema bertusi
0
5
6
6
6
All infections successful. However, ESB unable to keep a cadaver clean. Cadavers semi decomposed and full of mites
S. fabii-SCH10
0
0
1
2
2
No ESB/EPNs observed for any of the dead grubs
Control
0
0
0
0
0
No mortality recorded
The table shows the cumulative number of dead white grubs out of 30 white grubs inoculated per EPN species, the status of the entomopathogenic nematodes (EPNs) and of the symbiotic bacteria (ESB) inside the insect cadaver, as well as the status of the insect cadaver over a period of ten days post-infection, PI. No new mortalities recorded after five days PI. The nematode species are listed alphabetically
Heterorhabditis bacteriophora were found to have killed 20% of the P. sommeri grubs (Table 3) and were observed to have kept the cadavers clean and free of mites. In the cadavers, the red colour of the symbiotic bacteria was observed, although no EPN development of H. bacteriophora was observed. In two of the grubs, the symbiotic bacteria of H. bacteriophora were observed to have colonised only the upper body of P. sommeri. However, the rest of the EPNs used were found mostly to be unable to infect P. sommeri. Heterorhabditis zealandica SF41 was also found to have infected about 20% of the grubs involved (Table 3), but the EPNs were unable to complete their life cycle. Instead, stunted and dead nematodes were observed.
Table 3
Dissection observations for Pegylis sommeri
EPN species
Days PI and cumulative number of dead larvae
Notes
1
2
3
4
5
6
7
8
9
10
Heterorhabditis bacteriophora
0
1
1
4
5
6
6
6
6
6
EPNs mostly able to complete life cycle in cadavers. However, ESB present in some, although adult EPNs dead or absent. ESB able to maintain a clean cadaver
H. baujardi
0
0
0
0
1
1
2
2
2
2
No ESP/EPNs observed in cadavers
H. indica
0
0
0
0
0
0
0
0
0
0
No mortality recorded
H. noenieputensis
0
0
0
1
2
2
4
4
4
4
Mostly ESB present, but dead or stunted adult EPNs observed. No ESB/EPNs present in the cadavers that died after seven day
H. zealandica
0
1
2
2
3
3
3
3
3
3
Life cycle completed in two of the cadavers. ESB was unable to keep a cadaver clean. Cadavers semi-decomposed and full of mites. ESB present in one cadaver, but no EPNs present
MJ2C
          
H. zealandica SF41
0
0
1
5
6
7
8
8
9
9
Mostly ESB present, but dead or stunted adult EPNs observed. ESB unable to protect some of the cadavers
H. safricana
0
0
0
0
0
0
0
0
0
0
No mortality recorded
Oscheius myriophila
0
0
0
0
0
0
0
0
0
0
No mortality recorded
Steinernema bertusi
0
0
0
0
0
0
0
0
0
0
No mortality recorded
S. fabii-SCH10
0
0
0
0
0
0
0
0
0
0
No mortality recorded
Control
0
0
0
0
0
0
0
0
0
0
No mortality recorded
The table shows the number of white grubs that were found dead over a period of ten days post-infection, PI, the status of entomopathogenic nematodes (EPNs) and of symbiotic bacteria (ESB) inside the haemolymph, as well as the status of the insect cadaver. The nematode species are listed alphabetically
Most of the EPN-associated bacteria were generally observed to be unable to protect the P. sommeri and S. affinis cadavers from colonisation by other bacteria, possibly due to interference from the insects’ own gut bacteria. The above could be seen in terms of the colour changes occurring in the cadaver, from colours typical of the symbiotic bacteria concerned to darkened colours only hours after infection, coupled with the emission of a foul odour after a few days had elapsed.

Discussion

The results of the current study showed varying degrees of susceptibility of the white grubs S. affinis, P. sommeri, Maladera sp. 4 and Monochelus sp. to South African EPNs. Generally, Maladera sp. 4 was found to be the most susceptible, while P. sommeri was found to be the least susceptible. Schizonycha affinis and Monochelus sp. were found to be moderately susceptible. Other than P. sommeri, all the white grubs tested were found to be moderately susceptible to H. zealandica MJ2C. From the probit analysis, the LD50 values obtained using H. zealandica MJ2C were found to be relatively high for P. sommeri (1035 IJs per larva), followed by the values obtained for Maladera sp. 4 (284 IJs per larva) and, lastly, the values obtained for S. affinis (38 IJs per larva), at 28 days PI.
The varying degrees of susceptibility of white grubs for the different EPN species or isolates in the current study aligns well with what has already been shown for white grub susceptibility to EPNs elsewhere (Grewal et al. 2005; An et al. 2012; Abate et al. 2019). Consensus exists that the efficacy of EPNs against white grubs can be difficult to predict, due to the control efficacy recorded from previous experiments being inconsistent (Grewal et al. 2005). Such inconsistency can be explained by the presence of various biotic differences, including differences in the attractiveness of hosts, EPN species, their dispersal rates and ability to penetrate the grubs involved through their cuticle/gut wall (Georgis and Gaugler 1991; Koppenhöfer et al. 2007). In addition, environmental factors, like moisture, temperature and the physical properties of soil (Kaya et al. 1993), can also be responsible for the inconsistencies. In our experiments, only H. zealandica MJ2C was able to provide consistency in terms of mortality potential across the white grub species tested. Even then, the percentage mortality varied with repetitions and H. zealandica MJ2C proved unable to kill off the third instar larvae of P. sommeri effectively.
The obtained LD50 values were found to gradually decrease over time, in alignment with the increasing rate of mortality of the white grubs over time for all the EPN species apart from P. sommeri, for which the mortality, in the vial experiments, remained constant after seven days PI. At 28 days PI, a LD50 of 1035 IJs per larva was obtained for P. sommeri, of 284 IJs per larva for Maladera sp. 4 and of 38 IJs per larva for S. affinis. Although the obtained LD50 is high, it is nevertheless comparable to what has previously been obtained in other studies conducted on white grubs, further emphasising the high resistance of some white grubs to EPNs. For example, Sankaranarayanan et al. (2019), in using 24-well plates to test different strains of H. indica on the third instars of the white grub, Holotrichia serrata F. (Coleoptera: Scarabaeidae), obtained mean lethal dosage in the range 2015 IJs per grub to 7359 IJs per grub. However, Pokhrel et al. (2018) obtained rather lower LD50 after exposing second instar larvae of Chiloloba acuta (Coleoptera: Scarabaeidae) to S. abbasi and S. siamkayai in 40 g of silt loam soil. The resulting LD50 values were 44.9 IJs ml−1 and 50 98.1 IJs ml−1, respectively, 14 days PI. These LD50 values were very low compared to the values obtained during the probit experiment 14 days PI, namely 238, 497 and 1369 IJs per larva for S. affinis, Maladera sp. 4 and P. sommeri, respectively. However, the low LD50 of Pokhrel et al. (2018) was obtained with second instar larvae, which are generally known to be less resistant than are third instars (Alvandi et al. 2017).
Observing the EPN development occurring in the haemocoel of S. affinis and P. sommeri revealed several factors that might cause the low susceptibility of the two white grubs to EPNs. Such factors might include the inability of the EPNs to penetrate the insect’s cuticle and the gut cavity, as the haemocoel of some of the grubs dissected was found to contain no EPNs. Additionally, the inability of the EPN symbiotic bacteria to multiply and protect the EPNs and the cadaver concerned might have hindered the spreading of otherwise successful infection.
Stokwe and Malan (2017) observed that ESB were unable to multiply on the haemolymph of the woolly apple aphid, Eriosoma lanigerum, and that the nematodes involved were unable to effect successful infection. For white grubs, possible interference with symbiotic bacteria multiplication can include antagonistic effects brought about through the action of the white grubs’ gut bacteria (Skowronek et al. 2020). For example, H. zealandica MJ2C was observed to be able to protect its cadaver from the inroads of mites, with the cadavers not developing a foul smell. The fact that H. zealandica SF41 did not yield similar results suggests that H. zealandica MJ2C’s unique characteristic may be attributed to its different symbiotic bacteria. Booyzen (2022) reported the existence of three different symbiotic bacteria associated with South African H. zealandica, turning early infected Galleria mellonella larvae ‘blue’, ‘green’ and ‘red’. The H. zealandica MJ2C (green) and isolate SF41 (blue) used in the current study are associated with P. thracensis and P. heterorhabditis subsp. Heterorhabditis, respectively. Of key importance is the fact that the H. zealandica (MJ2C) associated with P. thracensis was found to be the most pathogenic EPN, which can probably be ascribed to the nature of the associated bacteria. The observation of the red colour in the H. bacteriophora infected cadaver and yet no EPNs in the dissected cadavers indicate that the bacteria of H. bacteriophora is able to multiply in the haemolymph of the white grubs.
Previously, Skowronek et al. (2020) found that bacteria from the midgut of the common white grub, Melolantha melolantha L. larvae, exhibited antagonistic activity against ESB. The possibility that the symbiotic bacteria of H. zealandica MJ2C can counteract the white grubs’ antagonistic bacteria requires further investigation. Similar to the observations made in the current study are the observations made in the study conducted by Karagoz et al. (2007). These authors observed that when the white grubs investigated were killed by EPNs the associated Sancassania sp. (Acari: Acaridae) mites moulted to the adult stage and began feeding on the host tissues and/or microbes associated with the cadavers, as well as on the IJs. Such a finding could serve to emphasise still further H. zealandica MJ2C’s use of its symbiotic bacteria to protect its cadaver.
The results of the current study confirm the results obtained in previous studies, showing that white grubs have varying degrees of susceptibility to EPNs. Variations in susceptibility cut across different EPN species and strains of the same EPN species. The implication of the occurrence of such a pattern for the biological control of white grubs, through the effective utilisation of EPNs, is that EPN screening is required for each white grub species. Furthermore, different strains of the same EPN species might need to be tested for their utilisation in different geographical regions, as inconsistency in efficacy has been shown in such regard.
For South African white grub species, the current study shows that high mortality can be achieved using H. zealandica MJ2C. However, H. zealandica MJ2C showed limited capacity to control P. sommeri. Further investigation is therefore required to test for the resistance mechanisms of P. sommeri. Most of the EPNs tested were found to lack the ability to develop within the haemocoel of the two most resistant grubs investigated, namely P. sommeri and S. affinis. This finding implies that such EPNs, even when they prove capable of penetrating the white grubs involved, tend to lack the biological compatibility that they require to kill the white grubs. Such incompatibility might be present because of the immunological defences possessed by the white grubs. Future studies should focus on identifying the specific mechanisms used by the white grubs to defend themselves against the EPNs investigated, while further screening studies can be conducted to identify the comparatively efficient EPN species.

Acknowledgements

The authors thank the members of the Tree Protection Cooperative Programme (TPCP) and the National Research Foundation of South Africa (TP14062571871 and ITR150119112367) for their financial support made available during the study. We acknowledge the effort of Des Conlong, Tom Webster and Janet Edmonds at the South African Sugarcane Research Institute (SASRI) for their assistance with the white grub collection. Additionally, we thank Benice Sivparsad from the Institute for Commercial Forestry Research (ICFR), for assistance with the white grub monitoring data and for providing some of the grubs that were used in the experiments.
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Literature
go back to reference Abate BA, Slippers B, Wingfield MJ, Conlong DE, Burger DA, Hurley BP (2019) Virulence and survival of native entomopathogenic nematodes for the management of white grubs in South Africa. Biol Control 137:104043CrossRef Abate BA, Slippers B, Wingfield MJ, Conlong DE, Burger DA, Hurley BP (2019) Virulence and survival of native entomopathogenic nematodes for the management of white grubs in South Africa. Biol Control 137:104043CrossRef
go back to reference Alvandi J, Karimi J, Ghadamyari M, Sharifi M, Asoodeh A (2017) Physiological defence of the white grub, Polyphylla adspersa Motschulsky (Col., Scarabaeidae) against entomopathogenic nematodes. J Asia Pac Entomol 20:878–885CrossRef Alvandi J, Karimi J, Ghadamyari M, Sharifi M, Asoodeh A (2017) Physiological defence of the white grub, Polyphylla adspersa Motschulsky (Col., Scarabaeidae) against entomopathogenic nematodes. J Asia Pac Entomol 20:878–885CrossRef
go back to reference An R, Voss M, Jagdale GB, Grewal PS (2012) Differences in immune defence evasion of selected inbred lines of Heterorhabditis bacteriophora in two white grub species. Insects 3:378–389CrossRefPubMedPubMedCentral An R, Voss M, Jagdale GB, Grewal PS (2012) Differences in immune defence evasion of selected inbred lines of Heterorhabditis bacteriophora in two white grub species. Insects 3:378–389CrossRefPubMedPubMedCentral
go back to reference Booysen E, Malan AP, Dicks LTM (2022) Colour of Heterorhabditis zealandica-infected-Galleria mellonella dependent on the Photorhabdus symbiont, with two new nematode-symbiotic associations reported. J Invertebr Pathol 189:107729CrossRefPubMed Booysen E, Malan AP, Dicks LTM (2022) Colour of Heterorhabditis zealandica-infected-Galleria mellonella dependent on the Photorhabdus symbiont, with two new nematode-symbiotic associations reported. J Invertebr Pathol 189:107729CrossRefPubMed
go back to reference Echeverri-Molina D, Govender P (2016a) Community structure and morphospecies composition of whitegrubs (Coleoptera: Scarabaeidae) attacking plantation Acacia mearnsii seedlings in KwaZulu-Natal, South Africa. Afr Entomol 24:170–179CrossRef Echeverri-Molina D, Govender P (2016a) Community structure and morphospecies composition of whitegrubs (Coleoptera: Scarabaeidae) attacking plantation Acacia mearnsii seedlings in KwaZulu-Natal, South Africa. Afr Entomol 24:170–179CrossRef
go back to reference Echeverri-Molina D, Govender P (2016b) Pest status of whitegrub species (Coleoptera: Scarabaeidae) attacking commercially grown Acacia mearnsii seedlings and its management implications in KwaZulu-Natal plantations, South Africa. African Entomol 24:382–392CrossRef Echeverri-Molina D, Govender P (2016b) Pest status of whitegrub species (Coleoptera: Scarabaeidae) attacking commercially grown Acacia mearnsii seedlings and its management implications in KwaZulu-Natal plantations, South Africa. African Entomol 24:382–392CrossRef
go back to reference Georgis R, Gaugler R (1991) Predictability in biological control using entomopathogenic nematodes. J Econ Entomol 84:713–720CrossRef Georgis R, Gaugler R (1991) Predictability in biological control using entomopathogenic nematodes. J Econ Entomol 84:713–720CrossRef
go back to reference Govender P (2007) Status of seedling establishment pests of Acacia mearnsii De Wild. (Mimosaceae) in South Africa. S Afr J Sci 103:141–147 Govender P (2007) Status of seedling establishment pests of Acacia mearnsii De Wild. (Mimosaceae) in South Africa. S Afr J Sci 103:141–147
go back to reference Govender P (2014) Effects of plantation residue management on the community structure of wattle regeneration invertebrate pests in South Africa. South for 76:229–236CrossRef Govender P (2014) Effects of plantation residue management on the community structure of wattle regeneration invertebrate pests in South Africa. South for 76:229–236CrossRef
go back to reference Grewal PS, Koppenhöfer AM, Choo HY (2005) Lawn, turfgrass and pasture applications. In: Grewal PS, Ehlers RU, Shapiro-Ilan DI (eds) Nematodes as biocontrol agents. CABI Publishing, Wallingford, pp 115–146CrossRef Grewal PS, Koppenhöfer AM, Choo HY (2005) Lawn, turfgrass and pasture applications. In: Grewal PS, Ehlers RU, Shapiro-Ilan DI (eds) Nematodes as biocontrol agents. CABI Publishing, Wallingford, pp 115–146CrossRef
go back to reference Jackson TA, Klein MG (2006) Scarabs as pests: a continuing problem. Coleopt Bull 60:102–119CrossRef Jackson TA, Klein MG (2006) Scarabs as pests: a continuing problem. Coleopt Bull 60:102–119CrossRef
go back to reference Karagoz M, Gulcu B, Cakmak I, Kaya HK, Hazir S (2007) Predation of entomopathogenic nematodes by Sancassania sp. (Acari: Acaridae). Exp Appl Acarol 43:85–95CrossRefPubMed Karagoz M, Gulcu B, Cakmak I, Kaya HK, Hazir S (2007) Predation of entomopathogenic nematodes by Sancassania sp. (Acari: Acaridae). Exp Appl Acarol 43:85–95CrossRefPubMed
go back to reference Katumanyane A, Malan AP, Tiedt LR, Hurley BP (2020) Steinernema bertusi n. sp. (Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa. Nematology 22:343–360CrossRef Katumanyane A, Malan AP, Tiedt LR, Hurley BP (2020) Steinernema bertusi n. sp. (Rhabditida: Steinernematidae), a new entomopathogenic nematode from South Africa. Nematology 22:343–360CrossRef
go back to reference Kaya KH, Bedding AR, Akhurst JR (1993) An overview of insect-parasitic and entomopathogenic nematodes. In: Bedding AR, Akhurst JR, Kaya KH (eds) Nematodes and the biological control of insect pests. CSIRO Publications, Melbourne, pp 1–15 Kaya KH, Bedding AR, Akhurst JR (1993) An overview of insect-parasitic and entomopathogenic nematodes. In: Bedding AR, Akhurst JR, Kaya KH (eds) Nematodes and the biological control of insect pests. CSIRO Publications, Melbourne, pp 1–15
go back to reference Koppenhöfer AM, Fuzy EM (2003) Ecological characterization of Steinernema scarabaei, a scarab-adapted entomopathogenic nematode from New Jersey. J Invertebr Pathol 83:139–148CrossRefPubMed Koppenhöfer AM, Fuzy EM (2003) Ecological characterization of Steinernema scarabaei, a scarab-adapted entomopathogenic nematode from New Jersey. J Invertebr Pathol 83:139–148CrossRefPubMed
go back to reference Koppenhöfer AM, Grewal PS, Fuzy EM (2007) Differences in penetration routes and establishment rates of four entomopathogenic nematode species into four white grub species. J Invertebr Pathol 94:184–195CrossRefPubMed Koppenhöfer AM, Grewal PS, Fuzy EM (2007) Differences in penetration routes and establishment rates of four entomopathogenic nematode species into four white grub species. J Invertebr Pathol 94:184–195CrossRefPubMed
go back to reference Koppenhöfer AM, Shapiro-Ilan DI, Hiltpold I (2020) Entomopathogenic nematodes in sustainable food production. Front Sustain Food Syst 4:125CrossRef Koppenhöfer AM, Shapiro-Ilan DI, Hiltpold I (2020) Entomopathogenic nematodes in sustainable food production. Front Sustain Food Syst 4:125CrossRef
go back to reference Lacey LA, Georgis R (2012) Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. J Nematol 44:218–225PubMedPubMedCentral Lacey LA, Georgis R (2012) Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. J Nematol 44:218–225PubMedPubMedCentral
go back to reference Malan AP, Ferreira T (2017) Entomopathogenic nematodes. In: Fourie H, Spaull VW, Jones RK, Daneel MS, De Waele D (eds) Nematology in South Africa: a view from the 21st century. Springer International, Berlin, pp 459–480CrossRef Malan AP, Ferreira T (2017) Entomopathogenic nematodes. In: Fourie H, Spaull VW, Jones RK, Daneel MS, De Waele D (eds) Nematology in South Africa: a view from the 21st century. Springer International, Berlin, pp 459–480CrossRef
go back to reference McArthur DG, Leslie G (2004) Preliminary observations on the impact of white grubs on the sugarcane yields in the Midlands north region of the South African sugar industry. Proc S Afr Sugar Technol Assoc 75:283–286 McArthur DG, Leslie G (2004) Preliminary observations on the impact of white grubs on the sugarcane yields in the Midlands north region of the South African sugar industry. Proc S Afr Sugar Technol Assoc 75:283–286
go back to reference Nobuyoshi I (2002) Behaviour of entomopathogenic nematodes. In: Dl L (ed) The biology of nematodes. Taylor & Francis, London, pp 511–520 Nobuyoshi I (2002) Behaviour of entomopathogenic nematodes. In: Dl L (ed) The biology of nematodes. Taylor & Francis, London, pp 511–520
go back to reference Pokhrel M, Thapa RB, Gharty-Chhetry YD, Sporleder M (2018) Efficacy of two entomopathogenic nematodes strains Steinernema siamkayai and S. abbasi against the 3rd instar larvae of Chiloloba acuta. J Agric Environ 17:73–81CrossRef Pokhrel M, Thapa RB, Gharty-Chhetry YD, Sporleder M (2018) Efficacy of two entomopathogenic nematodes strains Steinernema siamkayai and S. abbasi against the 3rd instar larvae of Chiloloba acuta. J Agric Environ 17:73–81CrossRef
go back to reference Ritcher PO (1966) White grubs and their allies. A study of North American Scarabaeoid larvae. Monograph series no. 4. Oregon State University Press, Corvallis Ritcher PO (1966) White grubs and their allies. A study of North American Scarabaeoid larvae. Monograph series no. 4. Oregon State University Press, Corvallis
go back to reference Sankaranarayanan C, Singaravelu B, Rajeshkumar M (2019) Entomopathogenic nematodes (EPN): diversity in Indian tropical sugarcane ecosystem and its biocontrol potential against white grub Holotrichia serrata F. on sugarcane. Sugar Tech 21:371–382CrossRef Sankaranarayanan C, Singaravelu B, Rajeshkumar M (2019) Entomopathogenic nematodes (EPN): diversity in Indian tropical sugarcane ecosystem and its biocontrol potential against white grub Holotrichia serrata F. on sugarcane. Sugar Tech 21:371–382CrossRef
go back to reference Sivparsad BJ, Germishuizen I, Conlong DE, Webster T, Morris AR (2018) Abundance and diversity of white grubs in KwaZulu-Natal: interim results of the 2017–2018 monitoring programme. ICFR Technical Note 06–2018. Institute for commercial forestry research, Pietermaritzburg, South Africa. pp 1–12 Sivparsad BJ, Germishuizen I, Conlong DE, Webster T, Morris AR (2018) Abundance and diversity of white grubs in KwaZulu-Natal: interim results of the 2017–2018 monitoring programme. ICFR Technical Note 06–2018. Institute for commercial forestry research, Pietermaritzburg, South Africa. pp 1–12
go back to reference Skowronek M, Sajnaga E, Pleszczyńska M, Kazimierczak W, Lis M, Wiater A (2020) Bacteria from the midgut of common cockchafer (Melolontha melolontha l.) larvae exhibiting antagonistic activity against bacterial symbionts of entomopathogenic nematodes: isolation and molecular identification. Int J Mol Sci 21:580CrossRefPubMedPubMedCentral Skowronek M, Sajnaga E, Pleszczyńska M, Kazimierczak W, Lis M, Wiater A (2020) Bacteria from the midgut of common cockchafer (Melolontha melolontha l.) larvae exhibiting antagonistic activity against bacterial symbionts of entomopathogenic nematodes: isolation and molecular identification. Int J Mol Sci 21:580CrossRefPubMedPubMedCentral
go back to reference Stokwe NF, Malan AP (2017) Laboratory bioassays to determine susceptibility of woolly apple aphid, Eriosoma lanigerum (Hausmann) (Hemiptera: Aphididae), to entomopathogenic nematodes. Afr Entomol 25:123–138CrossRef Stokwe NF, Malan AP (2017) Laboratory bioassays to determine susceptibility of woolly apple aphid, Eriosoma lanigerum (Hausmann) (Hemiptera: Aphididae), to entomopathogenic nematodes. Afr Entomol 25:123–138CrossRef
go back to reference Wu S, Youngman RR, Kok TL, Laub AC, Pfeiffer DG (2014) Interaction between entomopathogenic nematodes and entomopathogenic fungi applied to third instar southern masked chafer white grubs, Cyclocephala lurida (Coleoptera: Scarabaeidae), under laboratory and greenhouse conditions. Biol Control 76:65–73CrossRef Wu S, Youngman RR, Kok TL, Laub AC, Pfeiffer DG (2014) Interaction between entomopathogenic nematodes and entomopathogenic fungi applied to third instar southern masked chafer white grubs, Cyclocephala lurida (Coleoptera: Scarabaeidae), under laboratory and greenhouse conditions. Biol Control 76:65–73CrossRef
go back to reference Zuur AF, Ieno EN, Walker NJ, Saveliev AA, Smith GM (2009). GLM and GAM for absence–presence and proportional data. In: Zuur AF et al. (eds) Mixed effects models and extensions in ecology with R. Statistics for biology and health. Springer, New York, pp 245–259 Zuur AF, Ieno EN, Walker NJ, Saveliev AA, Smith GM (2009). GLM and GAM for absence–presence and proportional data. In: Zuur AF et al. (eds) Mixed effects models and extensions in ecology with R. Statistics for biology and health. Springer, New York, pp 245–259
Metadata
Title
Susceptibility of white grubs from forestry and sugarcane plantations in South Africa to entomopathogenic nematodes
Authors
Agil Katumanyane
Bernard Slippers
Mesfin Wondafrash
Antoinette P. Malan
Brett P. Hurley
Publication date
04-03-2023
Publisher
Springer Netherlands
Published in
BioControl / Issue 2/2023
Print ISSN: 1386-6141
Electronic ISSN: 1573-8248
DOI
https://doi.org/10.1007/s10526-023-10185-7

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