Greenhouse Evaluation of Conventional and Biorational Insecticides for Managing the Invasive Thrips parvispinus (Karny) (Thysanoptera: Thripidae)

Abstract

Thrips parvispinus (Karny) is an invasive pest of vegetable and ornamentals in the United States. To support ornamental growers to control T. parvispinus infestations, we tested seven conventional (spinosad, chlorfenapyr, sulfoxaflor–spinetoram, pyridalyl, tolfenpyrad, abamectin, and cyclaniliprole–flonicamid) and two biorational insecticides (mineral oil and sesame oil) under greenhouse conditions on mandevilla (Mandevilla splendens) and gardenia (Gardenia jasminoides), primary T. parvispinus ornamental hosts. Two insecticide applications were performed: a curative, treating an existing infestation, and a prophylactic, treating a plant prior to the thrips release. In the curative application, ten larvae and ten adults were released two weeks prior to treatment. Three leaves from the upper, middle, and lower canopy were collected 24 h, 7-, and 14-days post-treatment to assess thrips mortality. In the prophylactic application, plants were first sprayed with insecticides, and thrips were introduced 24 h later, but followed the same sampling method. In mandevilla, chlorfenapyr, abamectin, and spinosad caused the highest thrips mortality in both application types. Among horticultural oils, mineral oil and sesame oil increased mortality in prophylactic applications only. In gardenia, neither curative nor prophylactic applications of these products led to significant thrips mortality, and the possible reasons and recommendations for best thrips management are presented.

1. Introduction

Thrips parvispinus (Karny) (Thysanoptera: Thripidae) is a pest of vegetable and ornamentals and an invasive species originally from southeast Asia [1,2,3]. This pest is highly polyphagous, infesting over 43 plants from 19 families [4]. Over the last two decades, its geographic distribution and host range have expanded dramatically. Nowadays, T. parvispinus can be found on five continents (Africa, Asia, Oceania, Europe, and North America) [3,4]. Its spread has been largely driven by the increased international trade of plants. In countries such as India [3] and Indonesia [2], the pest has caused significant yield losses of chili pepper. Likewise, it has been associated with yield losses of ornamentals, like greenhouse-grown gardenias [5], and crops, such as green beans, potatoes, strawberries, brinjal [6], and papaya [7].

In 2020, the pest was found infesting ornamental plants in Florida, raising concerns about its impact on both ornamental and vegetable crops in the United States [8]. Thrips parvispinus has rapidly spread within the country, reaching states such as Colorado, Georgia, North and South Carolina, Pennsylvania, and Ohio [9]. Considering that Florida contributes 29% of the total value of potted flowering plants in the country [10], T. parvispinus represents a threat to the ornamental industry in Florida. Recently, heavy infestations of T. parvispinus have been documented in ornamental plants, such as gardenia and mandevilla [11], impacting the cultivation and distribution of these plants, particularly in light of the significant economic value associated with nursery, greenhouse, floriculture, and related agricultural commodities [12].

Initially, due to the severity of the threat posed by T. parvispinus, the Florida Department of Agriculture and Consumer Services-Division of Plant Industry (FDACS-DPI) implemented strict quarantine measures in the state. Nurseries in which T. parvispinus was found were placed under “Stop Sale and Hold Order” and were prohibited from moving plant materials until no thrips were present on the plants. During Florida’s 2023 spring shipping season, the most critical time of the year for the ornamental plant industry, over 60 nurseries faced quarantine due to infestations of T. parvispinus. Recently, the FDACS-DPI deregulated T. parvispinus and only nurseries with large, active infestations will be placed under quarantine (Revynthi pers. Com. with FDACS-DPI).

Thrips parvispinus mainly infests leaves and flowers as adults and nymphs, cause severe leaf scarring, flower drop, and upward leaf curling [13], significantly impacting plant aesthetics and marketability in an industry where perfection is non-negotiable. Given that the ornamental plant industry is extremely demanding, quarantine situations like this mandate swift action and an urgent need for effective monitoring and rapid pest control strategies to protect both growers and their livelihoods. To date, insecticide applications stand out as the most reliable method for the rapid suppression of invasive pests.

Thrips parvispinus completes its development from egg to adult in approximately 12 to 15 days, depending on the temperature, allowing for up to 20 generations per year [6]. After egg-hatching (typically around 5 days), the species progresses through two larval instars: the first lasting 1–2 days and the second 3–5 days. This is followed by a short pre-pupal phase (~1 day) and a pupal stage of about 2 days before adult emergence [14]. This rapid generation time raises significant concerns about the potential development of insecticide resistance. Considering the thrips’ ability to develop insecticide resistance [15] and the low-level introduction of new formulated insecticides for thrips control in the last decades, the proper application of these products is essential. Ideally, prioritizing insecticides with minimal non-target effects and rotation programs will prevent the emergence of resistant T. parvispinus populations. Horticultural oils offer safe and effective alternatives, as long as direct contact with beneficial organisms is carefully avoided [16]. Horticultural oils are petroleum- or plant-based products used to manage soft-bodied pests, like thrips, mites, aphids, and whiteflies [17]. Horticultural oils may help manage T. parvispinus by repelling adults, deterring oviposition and feeding or directly killing thrips, either by blocking their spiracles and causing asphyxiation or by disrupting metabolic processes [17]. These multiple modes of action make oils valuable tools for control while reducing the risk of resistance development [15,18].

To support ornamental growers facing all these challenges, we recently tested 32 products (21 conventional insecticides, 10 horticultural oils, and 1 insecticidal soap) for their potential to control T. parvispinus infestations [19]. We only tested products that are registered for ornamental plants, and we included horticultural oils and an insecticidal soap to provide growers with options of products with minimal non-target effects for integration into rotation schemes. All experiments were conducted under laboratory conditions using a Spray Potter Tower. Here, we present a follow-up study in which we tested the nine most efficacious products under greenhouse conditions on mandevilla (Mandevilla splendens (Hook.f.) Woodson) (Magnoliopsida: Apocynaceae) and gardenia (Gardenia jasminoides Ellis) (Magnoliopsida: Rubiaceae), primary T. parvispinus ornamental hosts. The products were spinosad, chlorfenapyr, sulfoxaflor–spinetoram, pyridalyl, tolfenpyrad, abamectin, cyclaniliprole–flonicamid, mineral oil, and sesame oil. The selected products represent different IRAC insecticide groups [20] to encourage rotation practices, reduce the risk of thrips developing resistance, and provide growers with a broader range of options for effective and sustainable pest management. We conducted toxicity experiments using two approaches: (i) curative application, in which plants already infested with thrips were treated and (ii) prophylactic application where plants were sprayed before infesting them with thrips. By evaluating both curative and prophylactic applications, we identified how these products perform under realistic scenarios, offering practical solutions for growers to integrate into their pest management strategies.

2. Materials and Methods

2.1. Host Plants

Mandevilla plants var. ‘Scarlet’ and ‘White’ (M. splendens) and gardenia plants (G. jasminoides) var. ‘Aimee’ were obtained from local nurseries in Homestead, Florida (mandevilla: 25°33′35″ N 80°27′08″ W, gardenia: 25°32′25″ N 80°25′35″ W). The gardenia plants were received as liners and grown to maturity before use. Both host plants were approximately one year old at the time of this study. The two mandevilla varieties were used simultaneously in the experiments. Plants were free of visible pests and were transferred to a quarantine greenhouse at 27 ± 2 °C, 60 ± 10% RH, and 12:12 h (L:D), where the plants received water twice a week. Bean plants (Phaseolus vulgaris L. var. ‘Roman’) (Goya Foods^®, Jersey City, NJ, USA) and pepper plants (Capsicum annuum var. ‘Pepper Mini Bell Red Organic’) (Harris Seeds^®, Rochester, NY, USA) were cultivated from seeds and used as host plants to sustain the T. parvispinus colony. Plants were watered three times a week. Bean seeds were sown weekly in 140 mL plastic pots. Pepper seeds were sown in 4 L pots. In both cases, pots were filled with ProMix soil (ProMix BX Mycorhizae, Denver, CO, USA). The plants were grown in a climate-controlled room at 25 ± 2 °C, RH 50%, and a 12:12 h (L:D) photoperiod. All experiments were carried out using gardenia and mandevilla plants, but each plant species was tested at different times in the greenhouse.

2.2. Thrips Parvispinus Rearing and Egg Cohort

A laboratory colony of T. parvispinus was established at the containment facility at the Tropical Research and Educational Center (TREC) in Homestead, Florida (25.50° N, 80.49° W). The permit was issued by FDACS-DPI (#2022-105). The founding individuals were obtained from mandevilla plants submitted to the TREC Plant Diagnostic Clinic, and species identity was verified by FDCAS-DPI. Since its establishment, the colony has been maintained inside a mesh cage (W47.5 × D47.5 × H93.0 cm; mesh diameter = 160 µm aperture; BugDorm-4M4590, BugDorm, Taichung, Taiwan) under controlled conditions (27 ± 1 °C, RH 70%, 12:12 h L:D) in a growth chamber (Panasonic Versatile Environmental Test Chamber MLR-352H). To maintain the T. parvispinus colony, four newly developed (approximately two weeks old) bean plants were added to the rearing cage three times per week. The colony’s diet was supplemented with Typha spp. pollen (Biobest^®, Westerlo, Belgium), which was applied uniformly using a soft brush. Every two weeks, old and dried plant materials were removed from the colony, and all waste was autoclaved prior to disposal to maintain sanitation and prevent contamination. The colony has been maintained under laboratory conditions since October 2022 without any exposure to pesticides.

Before starting the experiments, larval cohorts were generated by transferring adult males and females from the stock colony into large Petri dishes (135 mm diameter). These dishes contained a layer of moist cotton wool (Fisherbrand^®, Pittsburgh, PA, USA) topped with a bean leaf placed with the abaxial surface facing upward. Adults were introduced into the dishes using a manual aspirator, and the dishes were sealed with modified lids fitted with fine mesh (80 × 80, 350 µm aperture) to allow ventilation. The adults were allowed to mate and oviposit for 24 h, after which they were removed. The Petri dishes containing eggs were then incubated in a growth chamber set (27 ± 1 °C, 70% RH, and 12:12 h L:D) to promote development. For experiments, six-day-old second-instar larvae (L2) and adult females of an unknown age were used.

2.3. Prophylactic Spray Applications

Pesticide solutions were made following the highest recommended label rates for thrips control (where applicable) (

Table 1. List of conventional and biorational insecticides tested against Thrips parvispinus in greenhouse experiments.

	Active Ingredient(s)	Trade Name	EPA Registration	Insecticide Group	Rate *	Rate in 1L Solution	Site **
Conventional Insecticides	Spinosad	Conserve SC	62719-291	5	1.20 mL/ha	0.78 mL	G, N, L
	Tolfenpyrad	Hachi-Hachi SC	71711-31-67690	21A	323.1 mL/ha	2.11 mL	G, N, S, L
	Chlorfenapyr	Piston TR	91234-19	13	119.7 mL/ha	0.78 mL	G
	Cyclaniliprole–Flonicamid	Pradia	71512-33-59807	28–29	209.4 mL/ha	1.37 mL	G, N, S
	Abamectin	Timectin 0.15 EC	84229-1	6	95.7 mL/ha	0.63 mL	S, G, N
	Sulfoxaflor–Spinetoram	Xxpire	62719-676	4C-5	31.5 g/ha	206 mg	G, N
	Pyridalyl	Overture 35 WP	59639-125	Unclassified	91.8 g/ha	599 mg	G
Biorational Insecticides	Sesame oil	Bee Safe 3-in-1	FIFRA 25 (b) exempt	Unclassified	35.9 mL/ha	23.02 mL	S, G, N, L
	Mineral oil	Ultra-fine	86330-11	Unclassified	3%	30 mL	G, N, L, I

* Application rates are calculated according to the product amounts recommended on the label for treatment of one hectare (ha). ** Application site for each product (G: greenhouse, S: shadehouse, N: nursery, L: landscape, and I: interior).

). Each solution (1000 mL) was then loaded into 1.7 L handheld sprayers (HDX™, Mansfield, OH, USA) for application. Mandevilla and gardenia plants were placed in individual mesh cages (W 24.5 × D 24.5 × H 63.0 cm, mesh diameter 160 µm; BD4F2260 Bugdorm^®) (Bugdorm^® China). Each cage was labeled with the corresponding treatment, replicate, and randomly distributed within the greenhouse. One plant was placed in each cage, with each plant representing a replicate. A total of six replicates were conducted per treatment. The plants were sprayed until runoff (before thrips release) and allowed to dry for 24 h. After this drying period, the plants were artificially infested with ten second-instar larvae from a synchronized population (from the cohorts) and ten female adults from the stock colony. The larvae were transferred using a fine brush (Cotman™, Winsor & Newton, London, UK) onto 1.4 cm diameter bean leaf discs, cut with a cork borer (Fisherbrand^®, Pittsburgh, PA, USA). These discs were placed on 24-well cell culture plates (Falcon^®, Fisher Scientific, Pittsburgh, PA, USA), which were filled with water to prevent the larvae from escaping. The adults were collected using an aspirator attached to a 1000 µL pipette tip (Fisherbrand^®, Pittsburgh, PA, USA) sealed with parafilm (American National CanTM, Greenwich, CT, USA) to contain them.

The thrips were released on the plants by placing one leaf disc with ten larvae and one pipette tip containing ten female adults on each plant. We added adults and larvae simultaneously for infestation purposes, ensuring an asynchronous population. This approach allowed for the presence of multiple thrips stages on the plants, enabling a more comprehensive evaluation of product efficacy across the entire life cycle. The plants were watered three times per week throughout the experiment. Thrips mortality evaluations were performed 24 h, 7 days, and 14 days after thrips release. Samples were taken from three sections of the plant canopy: bottom, middle, and upper parts. One leaf from each section was cut using a Micro-Tip Blade Pruning Shears (Fiskars^®, Espoo, Finland) and placed into plastic bags (Ziploc^®, San Diego, CA, USA). These samples were then examined in the laboratory using a Leica Stereozoom S9E stereomicroscope (Heerbrugg, Switzerland), where all developmental stages were recorded, including both live and dead thrips. Each product was evaluated in two separate experimental blocks, with six replicates per block (N = 12). Water was included as the untreated control. For logistical reasons, mandevilla and gardenias were tested separately, in the same greenhouse, but at different times.

2.4. Curative Spray Applications

The experimental design for the curative applications followed the same procedure as the prophylactic evaluating the same pesticides (Table 1), with the only distinction being the application method, where thrips (second-instar larvae—L2 and adults) were released on the plants and allowed to establish for two weeks before applying the treatments. Consistently with the prophylactic applications, each product was evaluated in two separate experimental blocks, with six replicates per block (N = 12). A water treatment was included as the control. Mortality evaluations were performed 24 h, 7 days, and 14 days post-treatment, following the same procedure used in the prophylactic applications. Curative and prophylactic spray applications were performed simultaneously in the same greenhouse.

2.5. Statistical Analyses

All statistical analyses were conducted in R version 4.1.3 [21,22]. Thrips mortality (the proportion of dead thrips) observed on each evaluation day was analyzed via generalized linear mixed-effects models (GLMMs) using the glmmTMB package [23]. Mortality was presented as a single cumulative value, rather than separated by developmental stage. The response variable was modeled as binomially distributed [24,25] with the logit link function. The model included the following fixed effects: ‘treatments’ (9 products and water control) and ‘application type’ (curative and prophylactic). All interactions among these factors were included as well. The model also included a random effect, labeled ‘block’, and corresponding to the repetition of the entire experiment. A GLMM with a negative binomial error distribution was also used to analyze differences in the mean number of thrips observed on the two mandevilla varieties (‘Scarlet’ and ‘White’), including ‘treatments’, ‘application type’, and their interaction as fixed effects and ‘block’ as the random effect. The dataset used in this case included only plants that were treated with water as the control.

After fitting the models, F-tests were performed followed by the estimation of marginal mean proportion of thrips killed [26] for each application method. Each treatment’s efficacy was compared with that of the control, with tests of these odds ratios conducted separately for each method of application, using Dunnett’s p-value adjustment. For thrips mortality, the odds ratio corresponds to the relative chance of a thrips surviving following exposure to some treatment compared with one exposed only to the control. For an additional explanation of the interpretation of odds ratios in such contexts, see [19].

The data on the mean number of thrips observed on each part of the canopy were fit to GLMMs implemented with the glmmTMB package with a negative binomial error distribution. The models included the fixed effects of ‘thrips stage’ (L1, L2, and adults), ‘plant section’ (top, middle, and bottom), and ‘application type’ (curative and prophylactic) and random effects of the set of treatments evaluated independently twice in time, included as ‘block’ in the model. The dataset used in this case included only plants that were treated with water as the control. Statistics for mandevilla and gardenia were calculated separately.

3. Results

The two mandevilla varieties (‘Scarlet’ and ‘White’) did not differ significantly in thrips mortality (GLMM: Wald χ² = 0.01; d.f. = 1; p = 0.90); therefore, mortality data were analyzed together for both varieties. Curative and prophylactic applications of conventional and biorational insecticides on mandevilla plants caused significant thrips mortality (GLMM: Wald χ² = 95.0; d.f. = 8; p ≤ 0.001, Figure 1). The conventional insecticides chlorfenapyr (curative: p ≤ 0.001; prophylactic: p ≤ 0.001), abamectin (curative: p = 0.006; prophylactic: p ≤ 0.001), and spinosad (curative: p ≤ 0.001; prophylactic: p = 0.004) caused the highest thrips mortality and were effective in both application methods. However, while chlorfenapyr caused 80 to 100% mortality, spinosad caused 10 to 50% mortality, and abamectin caused 20 to 50% mortality, suggesting that the first insecticide was more lethal than the other two. Among the biorational insecticides, mineral oil (curative: p = 0.015; prophylactic: p ≤ 0.001) and sesame oil (curative: p = 0.33; prophylactic: p = 0.007) caused significant mortality in prophylactic applications only; however, mortality was below 40%. In addition, in mandevilla, most thrips were found in the top part of the canopy (GLMM: Wald χ² = 25.7; d.f. = 2; p ≤ 0.001; Figure 2A). The average number of first- (L1) and second-instar (L2) larvae was significantly higher at the top compared to the middle (p ≤ 0.001) and bottom (p ≤ 0.001) plant sections while the location of the adults on the plant canopy was random (p = 0.1).

In gardenia plants, neither curative nor prophylactic applications of conventional and biorational insecticides resulted in significant thrips mortality (GLMM: Wald χ² = 3.2; d.f. = 8; p = 0.88, Figure 3). The number of thrips found in the top, middle, or bottom section of the gardenia’s canopy was not significantly different (GLMM: Wald χ² = 0.00; d.f. = 2; p = 1.0; Figure 2B). However, the average number of L2 was significantly higher at the top compared to the middle (p = 0.03) and bottom (p = 0.04) plant sections, while no differences were found for the location of L1 (p = 0.1) and adults (p = 0.1) on the plant canopy.

4. Discussion

Here, we evaluated the potential of nine conventional and biorational insecticides to control an existing T. parvispinus infestation (curative application) or to prevent the establishment of the pest on the plant (prophylactic application) under greenhouse conditions. The aim of the prophylactic application was to evaluate whether the tested products, in addition to causing immediate mortality through direct contact, also possess residual activity capable of affecting thrips over time. In mandevilla (Figure 1), most products that caused high mortality upon direct exposure also showed residual effects lasting for one generation of thrips (approximately 14 days). Exceptions included sulfoxaflor–spinetoram, mineral oil, and sesame oil, which exhibited residual toxicity despite lacking significant direct-contact effects. Three chemical insecticides, spinosad (IRAC insecticide group = 5), chlorfenapyr (13), and abamectin (6), stood out as the most efficacious active ingredients against T. parvispinus infesting mandevilla in the greenhouse. We also observed that larvae were primarily located in the upper part of the canopy, particularly on mandevilla, while adults were distributed more randomly across the plant (Figure 2). These findings corroborate our previous assays conducted under laboratory conditions using a Potter Spray Tower in which spinosad and chlorfenapyr also excelled in all trials exhibiting direct and residual toxicity [19]. In gardenia (Figure 3), however, the same active ingredients did not cause thrips mortality to the same extent.

Their differential efficacy in mandevilla and gardenia suggests that host plant characteristics may have played an important role in thrips survival and development. At least two hypotheses can explain why the insecticides were effective on mandevilla but not on gardenia. First, we observed that the reduced thrips mortality on gardenia plants was associated with lower thrips infestation levels compared to mandevilla. Since we artificially infested both plant species with the same number of thrips from a colony well-adapted on beans, the thrips population did not establish as well on gardenia as it did on mandevilla. This may be due to the fact that the original specimens were collected from mandevilla, potentially influencing their adaptability to this host plant. Thrips parvispinus may exhibit a stronger preference for mandevilla or may have struggled to adapt to gardenia leading to reduced population growth, similar to what was observed for Frankliniella occidentalis Pergande (Thysanoptera: Thripidae) [27,28]. As a result, the lower thrips density on gardenia plants may have limited their contact with insecticidal residues, reducing treatment efficacy. Second, variations in plant morphology, such as leaf structure, trichome density, wax composition, or plant secondary metabolites, can induce physiological changes in pests or interfere with their level of tolerance to xenobiotics from their environment [29,30]. Plant morphology may even interfere with insecticide adherence, penetration, and retention [31]. Thus, mandevilla plants may have morphological and/or chemical traits favoring thrips establishment or leaf surfaces that facilitate better pesticide deposition and absorption, whereas gardenia leaves could possess traits that disturb its establishment or pesticide retention. Although these hypotheses remain to be tested, our findings highlight the importance of considering host plant traits and plant–insect interactions when developing IPM strategies. They also suggest that pesticide efficacy should be evaluated across multiple host plants.

Many biological invaders, including T. parvispinus, might experience Allee effects that hinder their establishment in new environments [32]. According to the Allee effect, population growth or individual fitness tends to increase as the population density increases, particularly at low densities, potentially resulting in a critical minimum population size below which populations cannot sustain themselves [33,34]. It is possible that the establishment of T. parvispinus on gardenia plants may be hindered by an Allee effect, where at low population densities, thrips may face challenges to overcome the plant’s defenses, leading to poor establishment. This suggests that introducing a higher number of thrips may be necessary to initiate successful colonization on gardenia plants. Such density-dependent processes have been observed in other species, where low-density populations face challenges in establishment and growth [35,36,37]. This hypothesis warrants further investigation, but exploring this effect could provide a valuable insight into how to optimize control measures in IPM programs targeting T. parvispinus.

Chlorfenapyr is known for providing thrips control and other listed pests for several days, offering long residual activity. It is labeled in Florida and is an excellent product to be included by ornamental growers in IPM programs under greenhouse conditions. For instance, F. occidentalis and Frankliniella intonsa Trybom (Thysanoptera: Thripidae) are highly susceptible to chlorfenapyr, with mortality reaching 100% within 12 h of exposure, whether via direct spray, contact with treated surfaces, or ingestion [38]. It is an example of a pro-insecticide, whose active form is generated inside the body of the target pest. Because its mode of action is different from others that are conventionally used to control thrips, the chances of developing resistance, although noted against the two-spotted spider mite [39], is less probable.

Spinosad is a widely used insecticide among greenhouse producers in the United States, primarily for managing F. occidentalis. Initially, it was introduced in 1998, and provided excellent thrips control [40,41]. However, its efficacy has declined in recent years, likely due to resistance development [42,43]. It is a naturally derived insecticide produced by the Actinomycete bacterium Saccharopolyspora spinosa that functions through its active metabolites, spinosyns A and D [44]. Spinosad provides both contact and residual properties, delivering rapid insect mortality within one to three days and maintaining effectiveness for up to two weeks after application [45]. Its mode of action involves stimulating the insect nervous system, leading to paralysis and death, similar to neonicotinoid-based insecticides (imidacloprid, thiamethoxam, acetamiprid, and dinotefuran) and the macrocyclic lactone insecticide/miticide (abamectin). Additionally, spinosad exhibits translaminar movement within leaf tissue, enhancing its effectiveness against pests like thrips that feed on leaf undersides and are challenging to target with contact insecticides alone. Although spinosad is highly effective, studies have reported harmful effects on natural enemies following both direct (topical) application and exposure to residues [46]. To reduce the risks of thrips resistance, we recommend rotating spinosad and chlorfenapyr with other conventional insecticides, such as abamectin and sulfoxaflor–spinetoram, and oils, such as mineral oil and sesame oil, in programs for T. parvispinus control. Furthermore, spinosad application should be carefully scheduled to ensure compatibility with biological control agents implemented in nurseries, greenhouses, and other crop systems [47].

Abamectin is a widely used insecticide and miticide in greenhouse and field crop production, particularly for managing thrips, mites, and other piercing–sucking pests. It belongs to the macrocyclic lactone class and is derived from the soil bacterium Streptomyces avermitilis [48]. Abamectin primarily acts as a neurotoxin, binding to glutamate-gated chloride channels in the insect nervous system, leading to paralysis and death. It has contact and residual activity, with visible effects occurring within hours and mortality typically observed within two-to-four days [49]. Additionally, it also has translaminar movement, penetrating leaf tissue and providing coverage against thrips. Despite its efficacy, abamectin has limitations. It degrades rapidly under sunlight, reducing its persistence in field applications and can be toxic to beneficials. Abamectin has been found to be highly toxic to nymphs and adults of predatory mites, such as Phytoseiulus persimilis Athias-Henriot [50,51] and Neoseiulus californicus McGregor (Mesostigmata: Phytoseiidae) [52], although the impact on N. californicus appears to be less severe compared to P. persimilis. Moreover, repeated use has led to resistance development in multiple pest species, including thrips and spider mites [53,54,55]. As suggested above, abamectin should be incorporated into rotation programs and compatibility between chemical and biological products should be considered when designing IPM strategies in nurseries, greenhouses, and various cropping systems [47].

The management of thrips using insecticides is very challenging due to their cryptic feeding habits, high mobility, rapid reproduction, short life cycle, and increasing resistance to insecticides [56,57,58]. Poorly timed applications and insufficient spray coverage, especially when using contact insecticides, are common issues that can compromise their overall efficacy. Although horticultural oils have not yet been assigned an IRAC group, their mode of action is primarily mechanical and physical disruption. Mineral oils, for example, function by disrupting respiration and mobility [59] in an array of insect pests [60]. Along with their varied modes of action, these oils are biodegradable, breaking down naturally without leaving residues in the environment. In our study, mineral and sesame oils proved to be good alternatives for rotation schemes corroborating our previous results obtained under laboratory conditions with these products against T. parvispinus [19]. Note that horticultural oils also need careful application since their effects can lead to phytotoxicity when administered at high concentrations [61] or when applied at high temperatures [62]. Nevertheless, resistance development would be prevented using a proper insecticide rotation scheme that encompasses a mix of traditional insecticides with biorational products that carry different modes of action [18,63].

In our study, we tested nine conventional insecticides representing seven IRAC groups, but this serves only as an initial step toward controlling the spread of this invasive pest. Further studies are essential to develop a more comprehensive IPM program and to evaluate specific rotation strategies. In this respect, we recently also addressed the urgent issue of thrips management by presenting an alternative to mitigate the presence of T. parvispinus from propagative material [61]. Our findings demonstrate that dip treatments with biorational and microbial insecticides, including mineral oil and Beauveria bassiana-based products, can effectively dislodge and kill T. parvispinus larvae from infested cuttings. In addition, no adverse effects were observed on mandevilla and gardenia cuttings treated with these products.

According to the label, chlorfenapyr is approved for use only in greenhouses. Therefore, nurseries can apply it in the early production stages of young ornamental plants before moving them outdoors. Drench or foliar applications in holding areas ensure a thorough coverage and early suppression of thrips populations, reducing infestations before plants are exposed to outdoor conditions. During the growing stage of mandevilla, gardenia, or any other ornamental plant outdoors, the use of the abamectin, spinosad, and sulfoxaflor–spinetoram insecticides, along with horticultural oils and biological control agents compatible with these pesticides is recommended. This strategy minimizes chemical reliance and reduces the risk of T. parvispinus resistance while maintaining long-term thrips management.

5. Conclusions

Our study demonstrates that three chemical insecticides (spinosad, chlorfenapyr, and abamectin) were the most effective in controlling established thrips populations on mandevilla under greenhouse conditions, showing both contact and residual activity. However, their efficacy was notably different on gardenia suggesting that host plant traits influenced our results. The observed differences highlight the importance of evaluating pest management tools across multiple host plants and incorporating plant–insect interactions into IPM strategies. Effective T. parvispinus management requires a multifaceted approach that incorporates biorational insecticides, insect growth regulators, effective resistance management practices, and precision agriculture to minimize the use of insecticides. Biological control agents along with chemical practices still need to be better investigated but could help in enhancing T. parvispinus suppression. In addition, knowledge of the biology and population dynamics of this pest will be crucial for designing an effective management strategy. In summary, the effective management of T. parvispinus requires integration of chemical, biological, and cultural control strategies. We must continue uncovering key interactions among these strategies to optimize their combined effectiveness against this pest.