Developmental characteristics and life cycle of the lawn cutworm, Spodoptera depravata (Lepidoptera: Noctuidae)

Abstract

We investigated the developmental characteristics and life cycle of the lawn cutworm, Spodoptera depravata (Lepidoptera: Noctuidae), which is one of the most important pests causing economic damage in grass production. For larval culture, we provided the zoysiagrass at $25^{\circ}C$ and $60{\pm}5%$ humidity. The durations of the developmental stages were as follows: $4.11{\pm}0.19$ days for eggs, $25.17{\pm}3.02$ for larvae, $8.80{\pm}0.28$ for pupae, and $7.57{\pm}0.95$ for adults. We grew the larvae to the 7th instar stage, unlike previous studies, in which it was assumed that the 6th instar was the final age. There was a significant positive correlation between the body length and head capsule width of each instar larvae. In terms of morphology, the eggs changed from light green immediately following oviposition to black as they developed, and the grass-fed larvae changed from light yellow immediately after hatching to green as development continued. We observed a pattern of black spots at regular intervals on the dorsal sides of the abdomens of the final instar larvae. Furthermore, we detected two notable designs on the dorsal side of the front of the head. The pupal colors changed from light brown and green immediately after pupation, to dark brown as the pupal cuticle hardened. The wingspans of the adults were similar in both sexes. However, the forewings of the males had obvious outer lines and eyespots with dark gray-brown backgrounds, whereas the corresponding features on the female forewings were less obvious. The oviposition preperiod was 2.11 days, the oviposition period was 4.2 days, the average fecundity per female was approximately 341 eggs, and the hatching rate was approximately 76.1%.

Introduction

Grasses, which are groundcover plants, contribute to the quality of the urban environment by preventing soil erosion caused by water or wind, increasing the organic matter content of the soil, purifying the air, and controlling air temperature (Choo and Lee, 2017). They are also used in the construction of natural landscapes, sports facilities, and the improvement of residential environments (Choo et al., 2000; Bae et al., 2013). The demand for grasses is increasing rapidly owing to the diversification of grass consumption and an increase in incomes (Youn et al., 2006; Bae et al., 2013).Grasses are a major forest product in Korea, and 2,148.9 ha is devoted to their production. Jangseong-gun, which is located in north Jeollanam-do, is the principle production region; it accounts for 49.6% of Jeollanam-do production and 73.7% of the domestic production of the Republic of Korea (Korea Forest Service, 2017). The pests that affect grasses mostly comprise soil-dwelling insects, particularly beetles and moths (Choo et al., 1998; Yang, 2013; Lee et al., 2014). Of these, Spodoptera depravata and Aceria zoysiae are more numerous and cause greater damage than any other insect pests (Lee et al., 2014).

Studies of the physiological and ecological aspects of grasses were conducted in 1968 (Cho and Kim, 1968; Yu and Youm, 1968), but since then, research has been limited compared to that carried out on other agricultural crops, although grasses are one of the main forest products (Hyun et al., 2012; Choo and Lee, 2017). Of the papers published on grasses in the Republic of Korea up to 2016, a substantial number have focused on the ecological effects of grass pests (Choo et al., 1998; Yang, 2013; Lee et al., 2014), but there have been no investigations into their developmental characteristics or life cycles.

Spodoptera depravata (Lepidoptera: Noctuidae) is one of the major lepidopteran pests, and attacks the linear leaves of Gramineae crops (Iwano, 1987; Kang et al., 2004). It is distributed throughout northeastern Asian countries such as the Republic of Korea, Japan, and China (Mochida and Okada, 1974; Kang et al., 2004). However, the year-round prevalence differs regionally: three occurrences per year at Kanto, Japan (Iwano, 1987), four to five at Tianjin, China (Guo et al., 1993), and five at Shanghai, China (Qian et al., 2003). In the Republic of Korea, S. depravataoccurs four times per year during early May to November, and is most prevalent between early August and late September (Lee, 2013). During this period, it was captured 2.8 times more often than other species of moths such as Parapediasia teterrellus and Pseudaletia separata (Lepidoptera: Noctuidae) (Lee, 2013).

It is thought that S. depravata females produce three to five egg masses—with 50–150 eggs per mass—during their lifespan, but no actual experimental data are available (Lee, 2013). It has also been reported that the species feeds actively from the third instar, resulting in rapid damage to grassland within a few days (Lee, 2013). In cases of severe infestation, the symptoms of early-stage larval feeding include grass leaves with a whitish appearance, because young instar larvae gnaw the surfaces of the leaves, resulting in linear pale white strip damage that excludes the veins. However, third-instar larvae feed on whole leaves including the veins, and feeding damage increases rapidly as the larvae approach the final instar (e.g., the 6th instar), resulting in yellowish-looking grasses and brown patches on the ground (Kang et al., 2004).

In contrast to the Republic of Korea, several ecological studies on S. depravata have been conducted in Japan and China. These include investigations of: the flight activity of three Spodopteraspp.—including S. depravata—measured by flight actograph (Saito, 2000); the relevant biological characteristics and control methods (Qian et al., 2003; Zhou et al., 2008); life tables (Huang and Xu, 2004); survivorship and reproductive capacity at room temperature (Huang et al., 2004); temperature-dependent survival; the survival of pupae at lower temperatures; and survival under various soil moisture conditions (Huang et al., 2005).

Ecological studies, particularly on the developmental period and the characteristics of each stage, are of fundamental importance to an understanding of the patterns of occurrence in the field, the patterns of damage, and the control strategies required for a given pest species. Furthermore, studies on the life cycle could be the basis for the development of effective agricultural pesticides, artificial diets, and mass indoor breeding systems. Therefore, the present study comprised an investigation of the life cycle of S. depravata and its developmental characteristics at each stage.

Materials and methods

DNA extraction

To identify S. depravata, we extracted genomic DNA from one or two legs of a specimen reared in the laboratory using a Wizard™ Genomic DNA Purification Kit (Promega, Wisconsin, USA), according to the manufacturer’s instructions.

Primers, polymerase chain reaction (PCR), and sequencing

To amplify the DNA barcoding region of the COI gene, we adapted a pair of primers from those described by Folmer et al. (1994): LCO1490 (5′-GGTCAACAAATCATAAAGATATTGG-3′), and HCO2198 (5′-TAAACTTCAGGGTGACCAAAAAATAC-3′). The PCR was conducted using AccuPower® PCR PreMix (Bioneer, Daejeon, Republic of Korea) under the following conditions: initial denaturation at 94°C for 7 min; 35 cycles of amplification (94°C for 1 min, 50°C for 1 min, and 72°C for 1 min); and a final extension at 72°C for 7 min. We then purified the PCR product using an AccuPower® PCR Purification kit (Bioneer, Daejeon, Republic of Korea). Electrophoresis was performed to confirm successful DNA amplification in 0.5× TAE buffer (a mixture of Tris base, acetic acid, and ethylenediaminetetraacetic acid) on 1% agarose gel. DNA sequencing was conducted using an ABI PRISM® BigDye® Terminator ver. 3.1 Cycle Sequencing kit with an ABI 3100 Genetic Analyzer (PE Applied Biosystems, Foster City, CA, USA). All products were sequenced from both strands. The sequences of both strands from each individual were aligned to obtain the finalized sequence using the ClustalW2 program (Larkin et al., 2007; http://www.ebi.ac.uk/Tools/msa/clustalw2). The sequences of the five specimens had one haplotype, and the species was identified using the Basic Local Alignment Search Tool (BLAST) and the Barcode of Life Data System (BOLD).

Specimens

To investigate the life cycle of S. depravata, we collected egg masses stuck to grass leaves, larvae at the 2nd to 6th instars hiding in the thatch layer between grasses and soil, and pupae located under the soil at a depth of 1 cm at the grass plantation in Jangseong-gun, Jeollanam-do, Republic of Korea between 18 and 25 May 2017 (Fig. 1). We also collected adults using a pheromone trap (Fig. 1C) containing an S. depravata-specific sex pheromone lure (Green Agro Tech Co., Ltd., Gyeongsan, Republic of Korea). The egg masses deposited on the wall of the trap by the collected adults were removed using thin paper (Fig. 1C). The collected egg masses, larvae, pupae, and adults were reared in a growth chamber at the insectarium of the Insect Molecular Phylogeny and Ecology Laboratory of Chonnam National University at 25°C in 60 ± 5% relative humidity, with a 16:8 light:dark cycle.

Host plant

Zoysia japonica Steud (Poales: Poaceae) grass was cultivated from May to November 2017 at a greenhouse at Chonnam National University (28°C, 32 ± 15%). The grass was cultivated without any use of agrochemicals and was on sale in Jangseong-gun, Jeollanam-do, where we collected the S. depravata specimens.

The grasses were transplanted to a plastic basket (48 × 38 × 7.5 cm) perforated for draining after covering with more than 2 cm of soil, and the plastic basket was placed in a 93 × 47.5 × 47.5 cm BugDorm-4S4590DH specimen handling cage (MegaView Science Co., Ltd., Taichung, Taiwan) to prevent the invasion of other pests and diseases. The grass was watered every 1–3 days depending on the dryness of the soil. A Yellow Sticky Trap (FarmHannong Co., Ltd., Seoul, Republic of Korea) was installed inside the specimen handling cage to remove small flies emerging from the soil. Grass leaves that had reached 8–10 cm were provided for the larvae by cutting the rhizome and stolon after washing the roots 3–4 times in water to remove the soil, which potentially harbors pests and diseases.

Insect rearing

To investigate the life cycle of the moth, we primarily used egg masses rather than other stages to ensure we had enough individuals. After hatching, all the 1st–3rd instar larvae from the same egg mass were reared together in an SP310102 insect breeding dish [diameter (Ø) 10 cm, height 4 cm; SPL Life Science, Pocheon, Republic of Korea], whereas ten larvae from each of the 4th–final instars were separately reared in an insect breeding dish to prevent cannibalization. Once enough individual larvae had been obtained the life cycle was regularized.

We allowed the adults to mate in a sex ratio of 1:1 to obtain egg masses in the mating space, which was formed into a triangular pyramid (approximately 17 cm per side; Fig. 2A) made from butter paper (38 cm × 25 cm) to provide a space for mating activity. The pyramid was provided with an SP10050 petri dish (Ø 5 cm, height 1.5 cm; SPL Life Science, Pocheon, Republic of Korea) supplied with 10% sugar water as adult feed. The portion of the butter paper on which the egg masses had been laid (Fig. 2B) was removed together with the egg masses, and placed on an insect breeding dish with a large moistened filter paper (No.131 Qualitative; Ø 9 cm; Advantec MFS, Inc., Dublin, CA, USA). The breeding dish was covered with a lid without meshes to maintain the moisture level inside the breeding dish.

Approximately 50 hatched larvae were kept until the 3rd instar on a petri dish, and fed with a small moistened filter paper (No.131 Qualitative; Ø 5.5 cm; Advantec MFS, Inc., Dublin, CA, USA), which was placed on the bottom of the dish (Fig. 4C). During this period, the larvae were provided with two or three 3–4 cm-long grass leaves without roots. The lid without a mesh was used to keep the moisture in the petri dish, preventing the leaves from drying out (Fig. 2C). Basically, new grass was provided each day, but the amount varied depending on the level of consumption. The petri dish and small filter paper were replaced every 2 days.

From the 4th instar, each individual larva was reared separately in a specimen bottle (Ø 4.6 cm, height 7.2 cm; SPL Life Science, Pocheon, Republic of Korea). A moistened filter paper was placed on the bottom of the specimen bottle, and grass including roots and leaves was provided after insertion into a black plastic mesh (Ø 4.5 cm, thickness of line 1 mm, spaces between lines 7 mm) to allow the grass to stand straight and for the larvae to hold the grasses while eating (Fig. 2D). Once or twice in a day, three to four units of 4–8 cm-long grass were provided depending on the level of consumption. As the larvae approached the final instar, six to eight units of grass were provided. The specimen bottle was equipped with a screw-type grooved plastic lid so that the bottle and the lid were in close contact, and the moisture of the internal filter paper was maintained, thereby ensuring the freshness of the grass. The specimen bottle and small filter paper were replaced every 5–6 days.

When the larvae reached the prepupa stage, they stopped eating and constructed a cocoon from feces, grass roots, and leaves in the space between the plastic mesh and the small filter paper. We then stopped supplying grass and maintained the internal humidity with moistened filter paper until pupation was complete. Immediately after pupation, the pupal cuticle is very weak. Therefore, 1 day after pupation we transferred the pupae to new specimen bottles, depending on their sex, and kept them there until they emerged.

After emergence, we provided the adults in the specimen bottles with cotton soaked in a 10% sugar solution (Fig. 2E), and allowed them to mate in a sex ratio of 1:1 in the mating space described above. If the male to female ratio was not 1:1, they were left in the specimen bottle until an adult of the opposite sex emerged. To determine the duration of egg development, we transferred each pair of adults to a new mating space. The newly obtained egg masses were counted to determine the number of 1st instar larvae using the same method described for egg mass rearing. When the 1st egg mass was observed, the female adult was transferred to a new mating space.

Life cycle of S. depravat

We investigated the developmental period of each immature stage—i.e., eggs, larvae, and pupae—using the oviposited egg masses. Using a total of ten egg masses, which were obtained from five adult pairs, the period of egg development was checked every 6 h until the second day of observation, based on a preliminary experiment in which the eggs took 3–4 days to hatch. From the third day, we observed the eggs at 1-h intervals until hatching was complete. Newly hatched larvae were counted and removed from the insect breeding dish to avoid redundant counting. Larvae hatched in the same period (within 1 h) were used for the observation of the larval period at 12-h intervals, whereas pupae and adults were checked for pupal and adult periods at 24-h intervals. The egg development period was defined from oviposition to hatching, each larval period and larval instar was defined by the presence or absence of the exuvium, and the prepupal period was defined from the start of cocoon formation by the last instar larvae to the completion of pupation. The pupal period was defined from the completion of pupation to emergence

For the close examination of egg and larval development, and to confirm the pupal sexes, we used a personal computer with an IMTcam3 camera (IMT i-Solution Inc., Daejeon, Republic of Korea) connected to an SZ51 Olympus microscope (OLYMPUS, Tokyo, Japan). Using the IMT iSolution Lite System (IMT i-Solution Inc., Daejeon, Republic of Korea) installed on the computer, we measured the individual body length and head width of each larval stage after taking the respective pictures. After sclerotization of the pupae had been confirmed, we measured the body length 24 h after pupation using Digimatic calipers (Mitutoyo, Kawasaki, Kanagawa, Japan) to distinguish the sexes. The pre-period of oviposition was defined as the time between emergence and the observation of the first egg mass; the period of oviposition was defined as the time between the observation of the first and last egg masses; and the adult period was defined as the time between emergence and death. We made observations at 12-h intervals to determine the proportion of hatching egg masses. The number of larvae hatching from the egg masses was difficult to count at the egg stage, so the number of hatched eggs and the number of dead eggs that turned black after a certain period were used to calculate the hatching ratio. For a close examination of the adult stage, dried male and female specimens were prepared, and their wingspans and body lengths were measured using calipers.

Fig. 1. Collection of Spodoptera depravata. A, collection site in Samseo-myeon, Jangseong-gun, Jeollanam-do (35°13′13′′ N, 126°38′25′′ E); B, eggs spawned on the grasses; C, adults collected in the pheromone trap (inset: isolated egg mass from the trap); D, larvae collected from the grasses; and E, pupae collected from the ground.

Fig. 2. Equipment used for the investigation of the life cycle of Spodoptera depravata. A, mating and oviposition cage constructed from butter paper; B, spawned egg mass on butter paper or petri dish; C, petri dish for larval culture at instar stages 1–3 (inset: petri dish covered with lid); D, top-to-bottom view of specimen bottle for larval culture at instar stages 4–7 (inset: side view of the specimen bottle with lid); and E, an adult emerged from a pupa in the specimen bottle.

Statistical analys

All the obtained data—i.e., the developmental periods of each immature stage, fecundities, hatching ratios, pre-periods of oviposition, periods of oviposition, body lengths, head widths of larvae, and adult wingspans—were input into Microsoft Excel (Microsoft Office Excel 2016; Microsoft Corp., Seattle, WA, USA). The body lengths, head widths, and developmental periods at each larval stage were analyzed using one-way analysis of variance (ANOVA), and means were compared using Student’s t-test at a significance level of 0.05. Correlation analysis was used to determine the relationship between body length and head width. The significance of the correlation coefficient (r) was tested using the t-test at (n – 2) degrees of freedom. Statistical analysis was performed on a computer using JMP statistical software (ver. 13.2.1; SAS Institute Inc., Cary, NC, USA).

Results and discussion

Species identification using molecular biology techniques

To confirm that S. depravata was the species collected in the pheromone traps, we sequenced the 658 bp of the COI gene—which is used as a DNA barcoding sequence—from five adult specimens. The sequences of the five individuals had 99–100% homology with S. depravata according to BLAST and BOLD (Fig. 3).

Fig. 3. Identification of Spodoptera depravata using molecular biology techniques. A barcoding sequence [658 bp of the cytochrome oxidase I (COI) gene] of five adult specimens collected using pheromone traps; and B, summary of the results of Blast and Bold searches for the S. depravata DNA barcoding sequence.

Morphological characteristics of the developmental stages

Eggs

oviposited as a mass changed from yellow green to black over time, and the larva within the eggshells rolled their bodies until just before hatching as long as the eggshell was present (Fig. 4). The eggs were 0.53 ± 0.01 mm in diameter.

Fig. 4. Development of Spodoptera depravata egg mass. A, twenty-two hours after oviposition; B, eighty-eight hours after oviposition (five hours before hatching); and C, an egg shell that has been almost completely consumed by a hatching larva. Note that only the bottom portion of the egg shell remains, and that the hatching larva is still rolling up its body.

Larva

Each larval stage is presented in Fig. 5. Immediately after hatching, the larvae had orange heads and light yellow bodies (Fig. 5A). As the larvae ate the grass, their bodies became green, and the last instar larvae were brown (Fig. 5G). Starting with the 3rd instar, the larvae developed an irregular series of white dots and lines, which were visible in both the dorsal and lateral views of the abdominal segments; and white lines (dorsal and subdorsal) appeared from the thorax to the terminal abdominal segment (Fig. 5C, 5D). From the 5th instar, two black lines were visible parallel to the adfrontal suture of the head, which resembled the Chinese character eight (八), and this line darkened and sharpened as the larvae approached the final instar (Figs. 5 and 6; Zhou et al., 2008). The adfrontal suture on the head was outlined in white with an inverted “Y” (Fig. 6F). The line of the epicranial notch is long in species of the genus Spodoptera (Passoa, 1991). The final instar larvae had black markings at regular intervals on the abdominal segment (Fig. 5G). The markings take the form of thick lines, unlike the triangles or semicircles observed in some species of Spodoptera (Passoa, 1991). There are pairs of black markings on each abdominal segment, and the markings on the 1st abdominal segment are much smaller than those on the 4th segment (Fig. 5G, Passoa, 1991). However, in the case of S. eridania, the black markings on the 1st segment are much larger than those on the 4th segment (Passoa, 1991). The 1st to 3rd instar larvae consumed the mesophyll excluding the veins of the grass, whereas subsequent instar larvae fed on the whole grass leaves, and the quantity consumed by the 7th instar larvae increased rapidly.

Fig. 5. Instars of Spodoptera depravata larvae. A, 1st instar; B, 2nd instar; C, 3rd instar; D, 4th instar; E, 5th instar; F, 6th instar; and G, 7th instar. The thick arrows in Fig. 7C indicate the white dorsal lines on the thorax and abdomen (C). The thin arrows in Fig. 7C indicate the irregular spots scattered mainly on the abdomen (C). The thick arrows in Fig. 7D indicate the irregular series of short, white, wavy lines (D). The thick arrow in Fig. 7G indicates the black markings at the first abdominal segment (G), and the thick arrow in Fig. 7G indicates the black markings at the fourth abdominal segment (G).

Fig. 6. Head capsules of larval instars of Spodoptera depravata. A, 1st instar; B, 2nd instar; C, 3rd instar; D, 4th instar; E, 5th instar; F, 6th instar; and G, 7th instar. The thick arrows in Fig. 8F indicate the two black lines in the adfrontal suture, and the thin white arrow in Fig. 8F indicates the inverted “Y” in the adfrontal region(F).

The average larval body lengths increased from a minimum of 2.77 mm to a maximum of 23.06 mm (Fig. 5; Table 1). The lengths of the larval instars were as follows: 1st instar, 2.77 ± 0.02 mm; 2nd instar, 3.86 ± 0.27 mm; 3rd instar, 5.55 ± 0.48 mm; 4th instar, 7.94 ± 1.14 mm; 5th instar, 11.62 ± 1.06 mm; 6th instar, 16.30 ± 0.74 mm; and 7th instar, 23.06 ± 1.07 mm. Each larval instar had a distinct body length, and there were statistically significant differences between the body lengths of the instars (Table 1; P < 0.05). There was a tatistically positive correlation (r = 0.995) between the lengths and widths of each larval instar (Fig. 7; P < 0.05 ).

Table 1. Body length and head width of Spodoptera depravata larvae (mm).

Values are the mean ± S.D. of three replicates for each stage of S. depravata development.

Means within rows followed by superscripted letters are significantly different at P < 0.05 (Student’s t-test).

Fig. 7. Correlation between the head width and body length of Spodoptera depravata during larval growth. There was a positive correlation between the head width and the body length.

Pupae

The pupae were obtect (Fig. 8), in common with many other members of the order Lepidoptera (Mosher, 1969). Immediately after pupation, the potential forewing region was clearly visible at the abdominal segment, with overall green and light brown pupal colors (Fig. 8A). After 5 days of pupation, the pupae were sclerotized and brown. The larval abdominal segment disappeared in the pupae, and the pupal eyes changed to black (Fig. 8B). The pupae darkened immediately prior to emergence, and the wings became visible through the pupal case (Fig. 8C). Similar changes to the pupal color and wing patterns immediately prior to emergence have been reported in other members of the order Lepidoptera (Ślisińska et al., 2006; Park, 2008; Zheng et al., 2011). Pupal gender was distinguished by the shape of a notch at abdominal segments 8 or 9. All the pupae had two bumps separated by a narrow groove that resembled an Arabic numeral eight (8) on abdominal segment 9, whereas only the female pupae had a longitudinal notch at the genital opening on abdominal segment 8 (Fig. 8D, 8E). The bodies of the female pupae were longer (13.00 ± 0.34 mm) than those of the male pupae (12.33 ± 0.12 mm) (Table 2).

Fig. 8. Spodoptera depravata pupae. A, Initial stage (1 day after pupation); B, middle stage (5 days after pupation); C, end stage (immediately before emergence); D, genital opening of male pupa; and E, genital opening of female pupa

Adults

The adult males had bipectinate antennae with abundant fringes, and forewings with a gray–brown background and a black–brown outer line, which appeared to be disconnected at each vein, and had a dark brown wedge-shaped spot located at the outside of the outer line (Fig. 9A). There was a near-white orbicular stigma at the center of the forewing, and a dark brown reniform stigma between the orbicular stigma and the outer line of the forewing (Fig. 9A). The costal region and outer margin of the hindwing had a light gray–brown background, whereas the other regions were white and had numerous fringes (Fig. 9A, 9B). The adult females had simple filiform antennae, and their overall color and the patterns on their wings were lighter than those of males (Fig. 9C, 9D). The females had forewings with light gray–brown backgrounds and brown outer lines, and hindwings with gray–white costal regions and outer margins; the other regions of their hindwings were white (Fig. 9C, 9D). The adult males were 11.83 ± 0.31 mm long and the females were 10.36 ± 0.07 mm long. The wing spans of the adult males were 25.37 ± 0.62 mm, and those of the females were 25.28 ± 0.04 mm. Therefore, the females had relatively short bodies compared to the males, whereas males and females had wings of similar width (Table 2).

Fig. 9. Spodoptera depravata adults. A, dorsal view of male; B, ventral view of male; C, dorsal view of female; and D, ventral view of female.

Table 2. Body lengths of Spodoptera depravata pupae and adults, and wingspans of adults (mm).

Values are the mean ± S.D. of three replicates.

Life cycle of Spodoptera depravata

The average developmental periods of the immature stages were: eggs, 4.11 ± 0.19 days; larvae, 25.17 ± 3.02 days; and pupae, 8.80 ± 0.28 days, and the average lifespan of an adult was 7.57 ± 0.95 days (Table 3).

Table 3. Developmental period of immature stages of Spodoptera depravata.

Values are the mean ± S.D. of three replicates for each stage of S. depravata development.

 Means followed by superscripted letters are significantly different at P < 0.05 (Student’s t-test). Figures within parentheses indicate the total number of individuals examined.

The time taken for S. depravata eggs to develop has been estimated at 3–4 days (Zhou et al., 2008), so is slightly longer in this study. This duration was similar to that of other species of Noctuidae, as follows: 3.00 ± 0.00 days for S. exigua (Azidah and Sofian-Azirun, 2006); 4.0 ± 0.03 days for S. litura (Fand et al., 2015); and 4.51 ± 0.51 days for Pseudoips fanaga (Kwon and Park, 2011).

With regard to the larval developmental period Huang and Xu (2004), however, reported it to be 18.66 ± 1.33 days for the 1st generation (at an average culture temperature of 27.96°C) and 20.09 ± 2.10 days for the 2nd generation (at an average culture temperature of 26.43°C). Qian et al. (2003) reported 18.9 days for the 3rd generation under the natural temperature condition (26°C during July to early August 2000-2002). Therefore, our results were 2–10 days longer than those reported in previous studies, but were closest to those of Huang et al. (2004), i.e., a larval development period of 23.05 days at an average temperature of 26°C.

For the developmental period of the pupae our result was similar to that reported by Huang et al. (2004), i.e., 8.21 ± 2.37 days, whereas the authors of other studies reported 7.27 ± 0.93 days for the 1st generation, 10.30 ± 0.99 days for the 2nd generation (Huang and Xu, 2004), and approximately 5.7 days without a specific generation (Qian et al., 2003). Therefore, the developmental period was longest in our study.

Adults lifespan in our study was longer than the results reported from other studies, i.e., 5.94 ± 1.92 days for the 1st generation (Huang and Xu, 2004), 5.94 ± 1.92 days for the 2nd generation (Huang and Xu, 2004), 3.24 ± 1.12 days (Huang et al., 2004), and approximately 5.2 days without a specific generation (Qian et al., 2003).

Development period of each larval stage

We determined the development periods of the larval stages to be 2.76 ± 0.30 days, 2.79 ± 0.30 days, 3.17 ± 0.29 days, 3.42 ± 0.58 days, 3.54 ± 0.62 days, 3.7 ± 0.64 days, and 4.81 ± 0.72 days for instars 1–7, respectively, and the prepupal period to be 2.1 ± 0.20 days (Table 3). As the larval stages progressed, the development periods also increased from 2.76 ± 0.30 days for the 1st instar to 4.81 ± 0.72 days for the 7th instar (Table 3). The larval 7th instar period differed significantly from the other larval instar periods, whereas there were no statistically significant differences between the others periods. However, Huang and Xu (2004) reported larval periods ranging from 2.08 ± 0.46 days for the 3rd instar to 4.90 ± 1.90 days for the 6th instar in the 2nd generation. In other studies, the minimum larval periods were observed to be the 4th instar (approximately 2.1 days; Qian et al., 2003) and the 2nd instar (2.91 ± 1.37 days; Huang et al., 2004). Differences in the culture environment, the season during which the investigation was conducted, the rearing temperature and humidity, and the feed used probably contributed to these discrepancies. However, in all previous studies the final larval instar (denoted the 6th instar) was the longest, and its duration was similar to that of our final larval instar, which we concluded to be 7th.

Development of 7th instar of S. depravata

In the present study, the final larval instar of S. depravatawas confirmed to be the 7th, whereas in all previous studies of S. depravata, the 6th was reported to be the final instar (Qian et al., 2003; Huang and Xu, 2004; Huang et al., 2004; Zhou et al., 2008). The available report-type materials also indicated the 6th as the final larval instar, without data presentation (Lee, 2013). However, our life cycle data clearly indicated that the S. depravata larvae continued to develop to a 7th instar, i.e., they molted six times (Fig. 6).

To clarify the existence of the 7th instar, we measured the head width of each instar by the observation of the presence or absence of exuviae. There were obvious and statistically significant differences between the head widths of the larval instars (P < 0.05; Table 1; Fig. 7): 0.42 ± 0.07 mm for the 1st instar, 0.53 ± 0.04 mm for the 2nd instar, 0.78 ± 0.03 mm for the 3rd instar, 1.09 ± 0.07 mm for the 4th instar, 1.55 ± 0.10 mm for the 5th instar, 2.22 ± 0.03 mm for the 6th instar, and 2.82 ± 0.01 mm for the 7th instar (Table 1). There were also statistically significant differences between the body lengths of the larval instars (P < 0.05; Table 1). However, except for Huang and Xu (2004)—who reported larval development to the 7th instar in some individuals, although they did not publish their data—the authors of most life cycle studies of S. depravata have reported that the larvae develop to the 6th instar only. One possible reason for this discrepancy is the influence of the host plants. In the case of the congeneric species S. exigua, the final larval instar is reported to differ depending on the host plant provided for consumption: the final instar is the 6th when the larvae are fed on cabbages [Brassica oleracea var. capitata (Brassicales: Brassicaceae)] or long beans [Vigna unguiculata (Fabales: Fabaceae)]; but the final instar is the 8th when the larvae are fed on shallots [Allium cepa (Asparagales: Amaryllidaceae)] or lady’s fingers [Abelmoschus esculentus (Malvales: Malvaceae)] (Azidah and Sofian-Azirun, 2006). A developmental study on S. exigua also indicated differing periods for the larval instars depending on the host plants provided to the larvae. For example, the 8th instar larvae took 27.04 ± 0.86 days and 25.14 ± 2.24 days to develop when shallots and lady’s fingers were provided, respectively, even though both host plants enabled the S. exigualarvae to continue to the 8th instar (Azidah and Sofian-Azirun, 2006). Similarly, the 6th instar larvae took 16.71 ± 0.33 days and 12.5 ± 0.46 days to develop when cabbage and long beans were provided, respectively, even though both host plants enabled the larvae to continue to the 6th instar (Azidah and Sofian-Azirun, 2006). Considering these results, we assume that the differences in the developmental periods of the final instars resulted from differences in the grass varieties: we used Zoysia grass, which is a turfgrass (Z. japonica) with a medium leaf blade width in the present study, whereas tall fescue or ray grasses were used in previous studies (Huang and Xu, 2004; Huang et al., 2004; Huang et al., 2005). Studies on the effects of the host plants on the life cycle are needed for further accurate comparisons.

Table 4. Oviposition and fecundity of Spodoptera depravata adults.

Values are the mean ± S.D. of three replicates for each measurement. ^† Total numbers examined.

Oviposition and hatching rates

We investigated the period of oviposition, the number of eggs deposited, and the hatching rate. The oviposition pre-period lasted 2.11 ± 0.69 days, the oviposition period lasted 4.20 ± 0.98 days, and the fecundity per female was 341.57 ± 180.08 (Table 4). The oviposition pre-period was similar to the pre-periods reported by Huang and Xu (2004), i.e., 2.69 ± 1.52 days (1st generation) and 2.33 ± 1.02 days (2nd generation). However, the oviposition period was longer and the fecundity was greater in our study than in the study by Huang and Xu (2004), i.e., 2.62 ± 1.63 days and 276.28 ± 225.22, respectively, for the 1st generation, and 2.33 ± 1.02 days and 228.23 ± 175.42, respectively, for the 2nd generation. Once more, we assume that the discrepancies were due to differences between the host plants, the types, the host plant culture conditions, etc. However, the hatching ratio in the present study—i.e., 76.17% on average—was similar to the hatching ratios reported by Huang and Xu (2004)—i.e., 84.38% and 90.32% for the 1st and 2nd generations, respectively.

Conclusions

We investigated the duration and characteristics of each stage of development in the life cycle of S. depravata, which occurs in the Republic of Korea. The developmental period of total stage, consisting egg, larvae, pupae, and adults was 45.67 ± 2.71 days. In contrast to previous studies conducted in other countries, we found that the larvae continued to develop to the 7th instar, which lasted longer than the other larval instars. The pupal period was similar to that reported by other authors, whereas adult development (8.8 days) took longer than previously reported (5.7–10.3 days). The body length and head width of the larvae differed significantly between instars (P < 0.05). Host plant-dependent life cycle studies are required for a better understanding of S. depravata ecology. We hope that the results of the present study will be used to develop an artificial diet for mass breeding and subsequent studies to produce eco-friendly control agents.