Study on the Vectoring Potential of Halyomorpha halys for Pantoea stewartii subsp. stewartii, the Pathogen Causing Stewart’s Disease in Maize

Abstract

Pantoea stewartii subsp. stewartii (Pss) is a Gram-negative bacterium first documented in North America, and is the causal agent of Stewart’s disease in maize (Zea mays), especially in sweet corn. First identified in North America, it is primarily spread by insect vectors like the corn flea beetle (Chaetocnema Pulicaria) in the United States. However, Pss has since spread globally—reaching parts of Africa, Asia, the Americas, and Europe—mainly through the international seed trade. Although this trade is limited, it has still facilitated the pathogen’s global movement, as evidenced by numerous phytosanitary interceptions. Recent studies in Italy, as indicated in the EFSA journal, reported that potential alternative vectors were identified, including Phyllotreta spp. and the invasive Asian brown marmorated stink bug (Halyomorpha halys); the latter tested positive in PCR screenings, raising concerns due to its broad host range and global distribution. This information has prompted studies to verify the ability of Halyomorpha halys to vector Pss to assess the risk and prevent the further spread of Pss in Europe. In this study, we explored the potential transmission of Pss by the brown marmorated stink bugs in maize plants, following its feeding on Pss-inoculated maize, as well as the presence of Pss within the insect’s body.

1. Introduction

Pantoea stewartii subsp. stewartii (Pss) is a Gram-negative bacterium. The main host of this bacterium is Zea mays (maize), especially sweet corn, but dent, flint, flour, and popcorn cultivars can also be infected. Pss causes a maize disease known as Stewart’s wilt, responsible for serious maize crop losses throughout the world [1]. Pss has been introduced to other parts of the world through maize seeds: the bacterium has now spread to Africa, North, Central and South America, Asia, and Ukraine. In the EU, it has been reported as transient in Slovenia and in Italy, where it has a restricted distribution and has now been eradicated (Pantoea stewartii subsp. stewartii). The bacterium is regulated under the Commission Implementing Regulation (EU) 2019/2072 of 28 November 2019 (Annex II) as a harmful organism, prohibiting its introduction and spread in the EU territory on seeds of Z. mays. Other potential host plants include various species of the family Poaceae, such as weeds, rice (Oryza sativa), oat (Avena sativa), and common wheat (Triticum aestivum), as well as jackfruit (Artocarpus heterophyllus), the ornamental Dracaena sanderiana, and the palm Bactris gasipaes [2].

In the USA, Pss spread largely relies on insect vectors, primarily the corn flea beetle Chaetocnema pulicaria Melsheimer (Coleoptera, Chrysomelidae), which is currently not present in the EU. In the north-central and eastern regions of the United States, the economic impact of this disease is minimal due to the presence of resistant cultivars and the use of systemic seed-applied insecticides for pest control. It is worth noting that susceptible maize varieties are severely affected and may be destroyed at the seedling stage [3].

To prevent the diffusion of Stewart’s wilt, the diagnosis of Pss is an effort supported by EU countries, as highlighted by the project financed by the EU (Valitest, grant agreement No. 773,139), and the activity of the European EURL-BAC project and Italian project Proteggo financed by the Italian Ministry of Agriculture, Food Sovereignty and Forests (MASAF). Various detection methods have been described in the literature, utilizing isolation, serology, or molecular biology techniques. Additionally, a diagnostic procedure for Pss was published in PM 7/60 (2) EPPO [4].

In the framework of EUROPHYT (European Union Notification System for Plant Health Interceptions), the Italian Plant Protection Organization carried out the surveillance of the territory for possible vectors associated with the bacterial pathogen in areas where symptoms of Stewart’s wilt were found [5,6]. In Italy, within the framework of Pss studies, during entomological surveys in maize fields, some insects were caught feeding on plants, and some of them were found positive under PCR screening: one crysomelid beetle belonged to the genus Phyllotreta (this genus is similar to the main vector in the country of origin C. pulicaria), as well as some specimens of the brown marmorated stink bug Halyomorpha halys (Stål) (Heteroptera, Pentatomidae) [5].

Following the detection of Pss in Halyomorpha halys (H. halys) in the framework of the PROTEGGO 1.3 project financed by the MASAF, a laboratory assay was conducted in 2021 to assess the stink bug’s potential as a vector of the bacterium. Originally from East Asia, H. halys has become an invasive pest in many parts of the world, including North America, Europe, the Caucasus region, Western Central Asia, and Chile. This polyphagous species feeds on a wide variety of crops, often causing severe damage even at early developmental stages. Injuries caused by H. halys result in rot deformities, and unmarketable produce, leading to considerable economic losses [7]. In addition to its direct damage, H. halys is also known to act as a vector for plant-associated microorganisms. For instance, it has been shown to transmit Eremotecium coryli Kurtzman (Saccharomycetaceae) to fruits and vegetables [8,9].

2. Materials and Methods

2.1. Bacterial Strain

The lyophilized strain of Pantoea stewartii subsp. stewartii (Pss) CREA-DC 1775 [6] was revived and cultured in NAG. The bacteria strain was cultured on nutrient agar 0.25% d-glucose (NAG) [4] for 48 h at 27–28 °C. Bacterial suspensions were prepared in phosphate buffer (PB 50 mM, pH = 7) and cell density was measured using a spectrophotometer (DS-11 Fx+, Spectrophotometer-Fluorometer Denovix Inc., Wilmington, DE, USA) at OD₆₆₀ = 0.05, corresponding to approximately 10⁸ colony-forming units (CFU)/mL [6].

2.2. Halyomorpha Halys

Adult specimens of Haliomorpha halys were collected in the field during the 2021 maize growing season and subsequently maintained under controlled laboratory rearing conditions (26 °C, 16:8 L:D, 65%RH). Insects were reared in mesh cages and provided with a mixed diet consisting of fruits, vegetables and seeds to ensure adequate nutrition and physiological maintenance prior to experimentation. A sub-sample of 25% of the batch of adults of the population used in the tests were analyzed and found to be negative for Pss. Following starvation [10,11,12], the insects were allowed to feed on maize plants under controlled conditions. The experimental procedures were conducted according to a stepwise protocol, detailed in the following sections.

2.3. Inoculation of Plant

Sweet maize seedlings, 8 to 14 days old (1–2 leaf stage), were artificially inoculated with Pss strain IPV-BO 2766 at a concentration of 10⁷–10⁸ cells/mL prepared in phosphate buffer. The 14-day-old maize plants (1–2 leaf stage) were stem-inoculated using an insulin needle, and grown in a quarantine glasshouse at 22–28 °C [4]. Negative control plants were inoculated with sterile distilled water. The disease symptoms appeared after 7 days. Plants were kept under observation for 30 days.

2.4. Halyomorpha halys, Zea mays and Pantoea stewartii subsp. stewartii Greenhouse Experiments

Approximately 100 female and 50 male Halyomorpha halys adults were used in this experiment. Adult specimens were reared in the Quarantine greenhouse in insect cages (BugDorm BD4E4545). The experimental design included three plant treatments: (i) non-inoculated maize plants; (ii) maize plants artificially inoculated with Pss; (iii) maize plants inoculated with distilled water as control. Each plant was placed in a rearing cage together with five adults of Halyomorpha halys specimens; in total, thirty maize plants were set with H. halys, unless otherwise specified. The experiment was divided into three steps, as follows:

• Step I. All the insects were fed for ten days in rearing cages containing healthy maize plants of 4–5 leaves supplemented with apple slices as an additional food source (a total of 30 maize plants were employed). After 1 week of feeding, 6 randomly selected insects (2 males and 4 females) were collected for molecular analysis. DNA was extracted separately from body and head parts to verify the absence of Pss. Before starting step II, adults, both females and males, were starved for 5 days with only water supply to avoid potential false negatives due to less or no feeding on the host [10,11,12];
• Step II. In this step, the remaining insects were divided into two treatment groups and exposed for one week to either maize plants artificially inoculated with Pss or to control plants treated with sterile distilled water. A total of 5 males and 25 females were placed in cages with healthy maize plants as negative control, while 40 males and 80 females were housed in cages with Pss-inoculated maize plants. In total, 5 adults were placed with one maize plant, and a total of 6 not inoculated and 24 inoculated maize plants were employed. After one week of feeding, 26 males and 34 females were randomly selected from insects exposed to inoculated maize plants. These individuals were collected and subjected to molecular analysis to assess the potential presence of Pss in both their body and head;
• Step III. The remaining 58 insects previously fed on Pss-inoculated plants, along with 30 insects previously fed on healthy control plants, were individually transferred to new insect-rearing cages. Each cage contained one adult insect and one healthy maize plant at the 4–5 leaf development stage (a total of 88 maize plants were employed). The insects were allowed to feed on the healthy plants for 30 days. At the end of this period, all insects were collected and analyzed to assess the presence of Pss.

The plants used in step III to feed the insects were observed for the possible development of symptoms of bacterial wilt disease over the next 21 days. At the end of the observation period, the plants were harvested and PCR-tested for the presence of Pss using qPCR according to Tambong et al. [13].

2.5. DNA Extraction

Plant DNA was extracted from leaves tissue with the DNeasy Plant Mini kit (Qiagen, Venlo, The Netherlands). The DNA concentration was evaluated by Qubit (dsDNA HS Assay kit, Invitrogen, Waltham, MA, USA) and the DNA was stored at ≤−15 °C until analysis. The bacterial DNA was extracted with the Gentra Puregene Yeast/Bact Kit (Qiagen, Venlo, The Netherlands).

Insects were stored in 95% ethanol until DNA extraction. Each specimen was dissected with scalpel and forceps in its storage solution to process the head and body separately for each sample. This procedure is essential for identifying where Pss may be retained within the insect [14]. Both anatomical parts were ground using seven Ø 2.0 mm glass beads in a 2.0 mL tube and Precellys 24 (Precellys) bead beater. DNA was extracted from the homogenized tissues using the DNeasy Blood & Tissue QIAcube Kit (QIAGEN, Redwood City, CA, USA) according to the manufacturer’s recommendations. The final elution step was performed in 50 µL of AE buffer provided in the kit.

2.6. Real-Time qPCR Detection from Plant and Insect

To confirm the presence of Pss in plant tissue, the diagnosis of Pss was performed according to Tambong et al. [13] and Pal et al. [15]. Pss re-isolation from symptomatic tissues was performed. Pss-like colonies were identified by qPCR using a TaqMan SYBR green [13,15]. The presence of Pss in the insect tissue was confirmed [13] according to the EPPO protocol [4] employing the Sso Advanced Universal Probes Supermix (Biorad, Hercules, CA, USA). All the real-time PCR reactions were carried out using CFX96 Real-Time System, BioRad. For all the real-time PCR, the standard deviation of the cycle threshold (Ct) values is calculated from the arithmetic mean of every sample that was amplified in two technical replicates. In each real-time PCR, positive and negative amplification controls in technical duplicate were added. For Pss DNA quantification, a standard quantification procedure was performed based on the real-time PCR of [13].

3. Results

3.1. Transmission Experiments

In step I, out of the 100 females and 50 males tested, 2 males and 4 females were selected for Pss analysis. The head and body were separated, and DNA was independently extracted from each tissue, as described in the materials and methods section. The DNA was analyzed by real-time PCR [13] and no signal relative to the presence of Pss was observed during the amplification. After a 5-day starvation period [10,11,12] and prior to step II, 5 male and 25 female insects (see

Table 1. Ct and CFU/mL for Pss found in H. halys, both males and females, in the different phases of the experiment.

	Insect Part	Step I		Step II		Step III
		Male	Female	Male	Female	Male	Female
H. halys	Head/Body	7	29	26	34	14	46
H. halys positive to PSS	Head	0	0	24	29	5	37
	Body	0	0	24	32	8	37
H. halys negative to PSS	Head	7	29	2	5	9	12
	Body	7	29	2	2	6	12
Ct range 2	Head	NA 1	NA	30–35	25–36	30–35	31–36
	Body	NA	NA	20–35	20–36	29–35	28–36
CFU/mL range 2	Head	0	0	104–102	105–102	104–102	104–102
	Body	0	0	107–102	107–102	104–102	104–102

¹ NA = No amplification signal. ² Minimum and maximum values obtained in the analyzed samples.

) were maintained in step I conditions for the entire duration of the experiment. These insects were housed in cages with healthy plants and served as negative controls. At the end of step III, they were collected for DNA extraction and analyzed using real-time PCR. No amplification signals were detected, indicating the absence of Pss. Simultaneously, in step II, 34 females and 26 males were selected from the 40 males and 80 females that were fed for 1 week in cages containing maize plants inoculated with Pss (see Figure S1 in the Supplementary File). DNA was extracted separately from the head and the body tissues, and analyzed by real-time PCR (Table 1) [13]. Figure 1 shows the histogramfit (histfit) plot of the threshold cycles (Ct) values obtained from the real-time PCR analysis of DNA extracted from insects after step II. The histfit plots a histogram of values in data using the number of bins equal to the square root of the number of elements in data, and fits a normal density function.

Figure 1 indicates that both male and female insects tested positive for Pss, with Ct values ranging from 20 to 35 (Table 1). The Ct values from male head tissues showed a narrower range (30 ≤ Ct ≤ 35) compared to those from females (25 ≤ Ct ≤ 36) (Figure 1a,c; Table 1). These values correspond to Pss concentrations of 10⁴–10² CFU/mL in males and 10⁵–10² CFU/mL in females (Table 1). CFU/mL concentrations were calculated using a calibration curve generated from real-time PCR, following the method described by Tambong et al. [13] (see Figure S2 in the Supplementary File). According to this curve, samples with Ct > 36 were considered negative and assigned a Ct value of 0.

The analysis of DNA extracted from the insect bodies revealed similar Ct values for both females (Figure 1b) and males (Figure 1d), ranging from 20 to 35, corresponding to a Pss concentration of 10⁷–10² CFU/mL (Table 1). After step II, 86% of female heads and 94% of female bodies were tested positive for Pss, while 87% of both male heads and bodies were positive. The scatter plots in Figure 2a,b show the Pearson’s correlation between Ct values from male and female heads and bodies, while Figure 2c,d compare Ct values between heads and bodies across sexes. In all cases, the p values were greater than 0.05, indicating that the correlations were not statistically significant. These results suggest that the distribution of the bacterium within the head and body is random.

In step III, 46 females and 14 males (Table 1) were transferred into a cage containing healthy maize plants and left to feed for 30 days. At the end of this period, all insects were analyzed by real-time PCR to detect the possible presence of Pss. Figure 3 shows Ct values obtained from the DNA extracted from these insects. Specifically, Figure 3a,b show the histfit plot of the Ct values for female heads and bodies, while Figure 3c,d display the corresponding histograms for male heads and bodies.

The results shown in Figure 3 indicate that, after step III, 73.9% of female insects (combined head and body data) tested positive for Pss. In males, 57% of heads and 43% of bodies were positive. Ct values for both sexes ranged from 30 to 40, corresponding to Pss concentrations between 10⁴ and 10² CFU/mL (Table 1). However, a higher proportion of male insects tested negative for Pss (Table 1). Ct values from both females’ (Figure 3a,b) and males’ (Figure 3c,d) head and body samples showed similar distributions. Figure 4 presents the results of the Pearson’s correlation analysis of Ct values, between heads and bodies within each sex (Figure 4a,b) and between sexes for both tissues (Figure 4c,d). In all cases, the p-values were greater than 0.05, indicating no statistically significant correlation. As previously observed in step II, this supports the conclusion that Pss is randomly distributed between the heads and the bodies of both male and female insects.

3.2. Plants

All the plants that were artificially inoculated were tested positive using molecular methods and pathogen isolations. All the plants from step III tested negative using both molecular methods [13,15] and isolation procedures.

4. Discussion

Stewart’s wilt on maize, caused by the Gram-negative bacterium Pantoea stewartii subsp. stewartii (Pss), has been reported as endemic in the mid-Atlantic USA states, the Ohio River Valley and the southern portion of the Corn Belt [16]. Stewart’s wilt is reported to have declined in prevalence in the USA due to the use of resistant varieties and the widespread use of neonicotinoid seed treatment. Neonicotinoids reduced the population abundance of the vector of Pss, the corn flea beetle Chaetocnema pulicaria, in which the bacterium overwinters [17,18]. In the Americas, C. pulicaria is the main vector and the main overwintering site of Pss. However, Stewart’s wilt is also transmitted by other chrysomelid beetles, the toothed flea beetle Chaetocnema denticulata Illiger and by the spotted cucumber beetle Diabrotica undecimpunctata howardi Barber [5], and by the larvae of the seed corn maggot Delia platura Meigen (Diptera, Anthomyidae), the wheat wireworm Agriotes mancus Say (Coleoptera, Elateridae) and May beetles Phyllophaga spp. (Coleoptera, Scarabaeidae). The transmission of bacteria among plants via insect vectors occurs during the insect feeding stage. Each contaminated insect vector can infect several healthy plants after feeding on an infected plant [2]. Insect screening in some of the Italian maize fields showing symptoms of Stewart’s wilt yielded positive tests for Pss in one case involving Phyllotreta sp. and in a few specimens of H. halys. In this paper, the potential role of H. halys in vectoring Pss is shown for the first time, based on the positive detection via a molecular test of the bacterium in insect body parts. Although the detection rate varied, Pss was found in both the heads and in the rest the bodies of male and female insects. These results suggest that H. halys can acquire the bacterium from the artificially inoculated plants under laboratory conditions. However, analyses failed to demonstrate successful transmission to healthy plants. Although insects tested positive in step III (Figure 3), all the plants analyzed remained negative. These findings indicate that the insect can ingest the bacterium during feeding, but it is not able to transmit it into plants. Moreover, the increase in the number of Pss-negative insects in step III may point to mechanical contamination or the elimination of the bacterium during digestion, suggesting H. halys is unable to effectively inoculate Pss into plant tissues while feeding.

Further studies are needed to clarify the exact mechanisms behind the behavior of H. halys, and to determine whether certain environmental or physiological conditions could enable transmission.

Considering the wide spread of H. halys in Italian corn fields and the growing issues related to Pss on maize, its epidemiological role remains an open question and requires further investigation. Similar outputs were obtained when assessing the transmission ability of H. halys of the yeast E. coryli on hazelnuts: although adults tested positive for the yeast in their mouthparts, the experiments failed to prove successful transmission [19].

5. Conclusions

This study provides the first experimental assessment of the brown marmorated stink bug (Halyomorpha halys) as a potential vector of Pantoea stewartii subsp. stewartii (Pss), the causative agent of Stewart’s wilt in maize. While molecular analyses confirmed that H. halys can acquire and retain Pss in both its head and body after feeding on infected plants, no evidence was found of successful pathogen transmission to healthy maize plants. These findings suggest that H. halys may act as a passive carrier but not an effective vector capable of spreading the disease under the tested conditions. The possibility of mechanical contamination or digestive elimination further complicates the interpretation of its epidemiological role. Given the widespread presence of H. halys and the economic significance of Pss, further investigations are warranted to clarify the conditions under which transmission might occur, and to better understand the potential risk this insect poses to maize health in Europe.