Kiwifruit (Actinidia chinensis) is one of the most popular fruits worldwide owing to its rich source of vitamin C, balanced nutritional components of minerals, and low calories which contribute to their health effects (Li et al., 2017; Stonehouse et al., 2013). However, when exporting kiwifruits, there are problems including postharvest rots, which cause significant losses during storage, distribution, and the shelf-life period (Manning et al., 2016). In general, the commencement of domestic kiwifruit harvesting occurs around mid-October and extends until April, thanks to the utilization of low-temperature storage (Choi et al., 2019). During the storage of kiwifruit, various fungal pathogens deteriorate the quality of fruits and cause postharvest fruit rots (Kim and Koh, 2018; Shin et al., 2021).

Postharvest fruit rots of kiwifruit have been related to more than seven fungi (Beraha and O’Brien, 1979; Hawthorne et al., 1982; Pennycook, 1985). Among them, Botryosphaeria spp. and Diaporthe spp. were found to be the primary cause of the kiwifruit rots (Du et al., 2021). Botryosphaeria dothidea is usually isolated from the postharvest fruit rots of kiwifruit, which was frequently reported in New Zealand, South Korea, and China. In addition, B. dothidea is a fungal pathogen capable of invading the kiwifruit via undamaged pericarp through natural openings (Li et al., 2017; Marsberg et al., 2017). Among the major pathogens that cause kiwifruit rots, Diaporthe spp. (anamorph: Phomopsis spp.) is a significant pathogen. Diaporthe eres has been reported in association with postharvest fruit rot in hardy kiwifruit in China, causing severe disease and thus significant economic losses in Northeast China (Liu et al., 2021). In recent years, D. eres is reported to cause the rotting of kiwifruit (A. deliciosa) in South Korea (Gi et al., 2022).

Several diagnostic methods have been developed to solve the problem of kiwifruit rots. Polymerase chain reaction (PCR) is required to verify the PCR amplicons increasing the overall cost and time expenditure (Wambua et al., 2017). Recent research has shown the availability of isothermal nucleic acid amplification, such as loop-mediated isothermal amplification (LAMP), which has also been applied for detection of kiwifruit rots pathogens (Qian et al., 2018; Wang et al., 2021). It requires four to six primers to increase specificity as well as isothermal devices, such as heat blocks for incubating at 60-65°C (Lu et al., 2021). Compared with conventional PCR and LAMP methods, the recombinase polymerase amplification (RPA) assay has been proven to be specific, sensitive, rapid, and cost-effective (Lobato and O’Sullivan, 2018). In this study, we developed a rapid and highly sensitive RPA assay for detection of B. dothidea and D. eres, respectively, to ensure effective control of fruit rots throughout the stages of cultivation, storage, and distribution. We compared the specificity and sensitivity of the RPA assay to those of the conventional PCR. In addition, we evaluated the effectiveness of the RPA assay by detecting B. dothidea and D. eres in kiwifruits showing rotting symptoms.

In 2019, fungal isolates were obtained from kiwifruits (cv. Hayward) grown in orchards located in Boseong County, South Korea. Kiwifruits with typical postharvest rot symptoms were treated with 70% ethanol for 60 s to disinfect the surface of kiwifruits, then rinsed for 60 s in sterile distilled water, and air-dried for 1 h. Skins were peeled and four pieces of flesh (1 cm²) were separated from the margin of symptomatic tissues and cultured on potato dextrose agar media (PDA) and grown for 6 days at 25°C. After incubation, a mycelial plug was transferred to fresh PDA and incubated at 25°C for 7 days. To obtain pure cultures of B. dothidea and D. eres, sporulation was achieved in a modified Barley media (Kim and Park, 1998). After 10 days, pycnidia were produced on the Barley media. Barley seeds with pycnidia were immersed in sterilized water for 10 min and mycelial fragments were removed by filtering through sterile Miracloth (Merck, Rahway, NJ, USA).

Genomic DNA (gDNA) was extracted from PDA culture of B. dothidea and D. eres, using the i-genomic BYE DNA extraction mini kit (iNtRON, Seongnam, Korea). For molecular identification of fungal isolates, internal transcribed spacer and rDNA region were amplified using the universal primer pair, ITS4 (5′-TCC TCC GCT TAT TGA TTG ATT GC-3′) and ITS5 (5′-GGA AGT AAA AGT CGT AAC AAG G-3′). Conventional PCR was conducted using 1 μl of fungal gDNA, 1 μl of each primer (20 μM of ITS4 and ITS5), and 7 μl of sterile distilled water to 10 μl of 2× TOPsimple DyeMIX (aliquot)-nTaq pre-mixture (Enzynomics, Daejeon, Korea). The reaction conditions were an initial preheated phase at 95°C for 2 min, followed by 32 cycles of DNA denaturation at 95°C for 30 s, a process of primer annealing at 57°C for 60 s, extension at 72°C for 1 min, and the final extension at 72°C for 5 min. PCR products were detected by electrophoresis using a 1.5% agarose gel. Amplified products were purified using a DNA purification kit (Nucleogen, Siheung, Korea) and were sent to Macrogen Inc. (Daejeon, Korea) for sequencing reactions.

RPA primers for detection of B. dothidea and D. eres were designed according to the TwistDx RPA User’s Guide (TwistDX, Ltd., Cambridge, UK). For B. dothidea, sequences of β-tubulin (TUB) genes (KU565871.1, KC864747.2, KJ801793.1, MG564762.1, MG564761.1, MG564760.1, MG564759.1, MG564758.1, and MG564757.1) were downloaded from the NCBI GenBank. For D. eres, sequences of histone H3 (HIS) gene (KJ420842.1, KJ420871.1, KP714488.1, KP714489.1, MN224562.1, MH121476.1, OL436035.1, OL436036.1, OL436037.1, and OL436038.1) were downloaded from the NCBI GenBank. RPA primers were designed based on highly conserved regions that were identified by alignment of the collected sequences, using BioEdit (v7.2.5) and Mega version 11 programs (Table 1). RPA reactions were performed in a total volume of 50 μl using a TwistAmp basic RPA kit (TwistDX, Ltd.) following the manufacturer’s instructions. The reaction mixture consisted of 2.4 μl of each RPA primer (10 μM), 29.5 μl of 1× rehydration buffer, 2 μl of gDNA, 11.2 μl sterile distilled water, and 2.5 μl of magnesium acetate (280 nM). The mixture was incubated at 39°C for 20 min. After the RPA reaction, 10 μl of amplified products were confirmed by loading on a 1.5% agarose gel.

A total of six primer sets were specifically designed for RPA analyses, targeting TUB in B. dothidea and HIS in D. eres. For TUB in B. dothidea, the primer sets included five primers, TUB_F1, TUB_F2, TUB_R1, TUB_R2, and TUB_R3, and for HIS in D. eres, the primer sets comprised of five primers, HIS_F1, HIS_F2, HIS_F3, HIS_R1, and HIS_R2 (Table 1, Supplementary Fig. 1). To assess the specificity of these primer sets, we used gDNA extracted from the mycelia of Botryosphaeria dothidea, Diaporthe eres, Botrytis cinerea, Alternaria alternata, and Pestaloptiopsis sp. These fungal isolates were obtained from infected kiwifruits in orchards in Boseong County, South Korea. Both the RPA and conventional PCR analyses demonstrated remarkable specificity, successfully amplifying the target genes, TUB in B. dothidea and HIS in D. eres (Fig. 1, Supplementary Fig. 2). Out of the six primer sets evaluated, primer set B. d_1 for TUB in B. dothidea and the primer set D_1 for HIS in D. eres displayed superior amplification of gDNA from B. dothidea and D. eres, respectively, while exhibiting minimal background amplification for gDNA derived from other fungal pathogens. Based on these findings, these primer sets were chosen for further optimization of the RPA analysis.

Next, the chosen primer sets were evaluated to ascertain the ideal temperature and incubation duration for RPA analysis. Three distinct temperatures, namely 25°C, 35°C, and 39°C, were tested using 10 ng of gDNA, and the RPA reactions were terminated after a 20 min incubation period. Notably, well-defined bands were observed for TUB in B. dothidea at both 35°C and 39°C, while for HIS in D. eres, well-formed bands were specifically observed at 39°C (Fig. 2A). In order to identify the ideal incubation time, RPA analyses were performed at a temperature of 39°C. The reactions were halted at different time points: 1 min, 5 min, 10 min, and 20 min after incubation. As a result, distinct and well-defined bands became visible as early as 5 min for both TUB in B. dothidea and HIS in D. eres. However, the most favorable incubation time, resulting in the best-defined bands, was observed after 20 min (Fig. 2B). Taken together, the RPA assays were optimized to achieve the best results by maintaining an incubation temperature of 39°C for a minimum of 5 min.

To assess the sensitivity of the RPA assay, a series of 10-fold dilutions was prepared for both B. dothidea and D. eres gDNA. The dilution series covered a range from 12.6 ng/μl to 1.26 pg/μl for B. dothidea gDNA and from 8.4 ng/μl to 0.84 pg/μl for D. eres gDNA. Well-defined bands were observed for all the reactions added with 2 μl of the dilution series of gDNA, and there was no amplification without input gDNA (Fig. 3). In comparison to the conventional PCR results, the RPA assay exhibited superior sensitivity. Specifically, for TUB in B. dothidea, the RPA assay demonstrated a sensitivity that was at least 1,000 times greater (Fig. 3A), while for HIS in D. eres, it was 100 times more sensitive (Fig. 3B).

In order to evaluate the effectiveness of the RPA assay, healthy kiwifruits were inoculated with varying concentrations of conidia of B. dothidea and D. eres and subjected to both RPA and conventional PCR analyses. Conidia of B. dothidea and D. eres were filtered and adjusted to concentrations ranging from 1 × 10¹ conidia/ml to 1 × 10⁵ conidia/ml. Healthy kiwifruits, obtained from a grocery store, were wounded and inoculated with 20 μl of the spore suspensions on the fruit surface. The inoculated fruits were then placed in a chamber and incubated at 25°C. Following a 10 days incubation period, the peel of the kiwifruits at the site of inoculation was carefully removed and subjected to gDNA extraction. Subsequently, a 2 μl volume of extracted gDNA was used for RPA and conventional PCR analyses. Genomic DNA extracted from kiwifruit samples that had been wounded, but not inoculated with spores was used as a negative control (Fig. 4). The RPA assays were successful in detection of B. dothidea and D. eres in kiwifruit showing typical rot symptoms, as well as in asymptomatic kiwifruits that had been inoculated with a very small number of conidia.

Kiwifruit production suffers greatly from postharvest fruit rot that leads to significant economic losses for kiwifruit farms throughout the harvesting, storage, distribution, and marketing processes (Hawthorne et al., 1982). In South Korea, there has been a lack of research concerning fruit rots in kiwifruit and effective methods for identifying infected fruits. During storage at low temperatures after harvesting, it is crucial to exercise great caution in order to prevent the spread of postharvest fruit rot diseases to healthy kiwifruits. However, it is very challenging to distinguish infected kiwifruit due to long latent periods. Therefore, it becomes necessary to develop molecular diagnostic techniques to detect the pathogens responsible for kiwifruit rots. The causal agents of kiwifruit rots, namely B. dothidea and D. eres quickly destroy ripening kiwifruits, emphasizing the need for a simple, rapid, sensitive, and accurate method like the developed RPA method to detect these pathogens. In the current study, the RPA assay demonstrated superior speed and specificity in detecting B. dothidea and D. eres in infected kiwifruits compared to the conventional PCR assay. The assay exhibited rapid performance, taking only 5 min, and could be conducted at a relatively low temperature of 39°C. As a result, it holds great promise as a diagnostic method for identifying infected kiwifruits during storage and transportation, enables effective controls of kiwifruit rot disease, and contributes to extending the shelf-life period of kiwifruits.