Introduction

Hazelnut (Corylus avellana) is an important nut tree with significant economic importance worldwide. Türkiye maintained its position as the leading global hazelnut producer in 2022, contributing to 60% of the world’s production with a total of 765 thousand tons, according to FAO statistics (FAOSTAT 2022). Predominantly cultivated across the Black Sea region of Türkiye, hazelnuts serve as a primary income source for farmers. With increasing global demand and economic significance, the hazelnut industry is expanding, highlighting the importance of orchard renewal and the need for true-to-type cultivars to establish new orchards in the region. Hazelnuts are mainly propagated by suckers (Erdoğan 2018; İslam 2018). However, this conventional method for vegetative propagation is slow and labor-intensive. Micropropagation is the suitable technique for producing a large number of clonal plants quickly (Nas and Read 2004a, b).

Microbial contamination presents a notable obstacle to the effective initiation of hazelnut micropropagation (Hand et al. 2016). Therefore, it is crucial to reduce or eliminate contamination for a successful outcome. The control of fungal and bacterial contamination is a complex and time-consuming process influenced by various factors, including the explant source, type, age, size, and season of explant collection. In hazelnut micropropagation, various protocols have been developed to address microbial contamination during the initiation phase. Among these protocols, the source of explants plays a crucial role in the success of initiation. Initiating culture with shoot nodes obtained from plants cultivated in a controlled, healthy environment has shown lower contamination rates compared to those sourced from field-grown plants (Messeguer and Mele 1983; Yu and Reed 1995; Bacchetta et al. 2008). Many researchers have adopted a similar approach, potting donor plants from the field, growing them in greenhouses or growth chambers, and then initiating culture (Hand et al. 2016; Silvestri et al. 2020; Yahyaoui et al. 2021). Maintenance of field-grown branches in a forcing solution within a controlled environment prior to initiating culture has also been documented to decrease contamination (Diaz-Sala et al. 1990; Damiano et al. 2005; Nas 2004).

As woody plants age, their morphological capacities decline (Rodriguez et al. 1988; Hand 2013). Hazelnut shoot micropropagation from juvenile tissues typically exhibits higher in vitro response rates compared to those from adult tissues (Messeguer and Mele 1987; Al Kai et al. 1984; Anderson 1984; Perez et al. 1987; Rodriguez et al. 1988). Furthermore, studies suggest that juvenile plants tend to harbor fewer microbial contaminants than mature plants (Anderson 1984; Messeguer and Mele 1987). Hand et al. (2016) demonstrated that the viability and contamination levels of shoot explants are influenced by nodal position, recommending the utilization of the first three nodes below the apex of plants. These findings highlight the importance of employing healthy and juvenile plant sources to minimize microbial contamination and enhance in vitro regeneration during the initiation phase.

Sampling season has a significant influence on contamination levels and in vitro regeneration. Numerous studies have documented variations in microbial contamination rates when nodal explants are collected at different times of the year and used for shoot culture (Messeguer and Mele 1983; Bassil et al. 1991; Yu and Reed 1995; Gao et al. 2008). Similarly, many researchers have investigated the culture responses of plants collected in different seasons to identify the optimal period for initiating successful cultures, revealing significant variations (Messeguer and Mele 1983; Yu and Reed 1995; Bacchetta et al. 2005; Gao et al. 2008; Yahyaoui et al. 2021).

Surface sterilization is a widely explored method for reducing microbial contamination during culture establishment. However, excessive sterilization can lead to the accumulation of phenolic compounds in the culture medium, which may have toxic effects, potentially hindering successful micropropagation in woody plants (Bassil et al. 1991). Hence, it is crucial to identify the appropriate chemicals, doses, and exposure times for effective sterilization. In hazelnut micropropagation, commonly used chemical disinfectants for surface sterilization include ethanol, Tween 20, and sodium hypochlorite. These sterilization agents have been extensively studied and documented in various research works (Messeguer and Mele 1983; Bassil et al. 1991; Yu and Reed 1995; Bacchetta et al. 2008; Kaplan et al. 2020; Elçiçek 2020).

Thermotherapy is a well-documented technique in plant biotechnology aimed at controlling or eliminating pathogens, particularly viruses, bacteria, and fungi, from infected plant materials. By exposing plants to elevated temperatures, usually ranging from 35 °C to 42 °C, this method disrupts the replication and dissemination of viruses, resulting in their inactivation or elimination (Barba et al. 2015; Laimer and Barba 2011; Panattoni et al. 2013; Faccioli 2001; Faccioli and Marani 1998). This approach can be applied to both in vivo and in vitro plants. Combining heat treatment with meristem culture shows promise for producing plants free from viral infections (Skiada et al. 2009; Lozoya-Saldana and Dawson 1982; Kaya 2021).

Türkiye has a rich hazelnut biodiversity owing to its favorable geographical location. The national collection of hazelnut genetic resources located in Giresun currently holds 459 C.avellana genotypes. In order to efficiently propagate and conserve genotypes for breeding and biotechnology research purposes several in vitro culture establishment studies were conducted. However, initial micropropagation attempts encountered challenges, particularly with axillary nodes sourced from mature field-grown trees, which exhibited a high level of contamination and failed to establish in vitro. Consequently, various approaches were employed to reduce contamination and enhance the success of culture initiation. This paper investigates the different strategies utilized to reduce microbial contamination during the establishment phase of culture.

The study aimed to devise efficient techniques for initiating plant cultures, encompassing several key strategies. Firstly, it sought to assess the most favorable seasons for collecting plant material to initiate cultures. Secondly, the study aimed to establish a sanitation program for donor plants to ensure the procurement of healthy explants suitable for culture initiation. Additionally, it aimed to employ thermotherapy on potted donor plants as a means of pathogen eradication. Lastly, the research involved exploring different disinfectants to enhance the surface sterilization process for the explants. These methodologies collectively aimed to refine and optimize the procedures involved in initiating plant cultures, ensuring the health and viability of the resulting cultures.

Materials and methods

Plant material

The study utilized Turkish cultivars of C. avellana; ‘Tombul’, ‘Çakıldak’, and ‘Foşa’. Plant materials were sourced from both mature field-grown trees and 1-year-old plants propagated by suckers. In all experiments, upper shoots measuring 30–40 cm in length were collected and transferred to the laboratory for surface sterilization. Three shoots were harvested from each tree. Before surface sterilization, the shoot apex was removed due to their vulnerability to sterilization (Bacchetta et al. 2008; Hand et al. 2016).

Initial surface sterilization

In experiments 1 and 2 the following surface sterilization method was employed. First, the leaves were removed from the shoots, which were then washed with running tap water. Subsequently, the shoots were treated with antibacterial liquid soap (Activex) and rinsed with tap water. Shoots were excised into 2–3 cm-long single nodal segments, and the first three nodes below the apex were used as plant material. The explants were disinfected with 70% ethanol for 10 s and 2% NaOCl for 20 min. Following this, the explants were rinsed three times in distilled sterile water for 10 s each time. Finally, the explants were placed into 16 × 100-mm culture tubes containing 5 ml of culture medium.

Experiment 1: season of explant collection

To investigate the impact of different explant collection seasons on clean shoots and bud-break, explants were obtained from 30-year-old ‘Tombul’, ‘Çakıldak’, and ‘Foşa’ mother plants in the field collection. This was carried out across five distinct periods spanning the years 2021–2022. No fungicide was applied to the trees throughout the duration of the study. Explants were collected during five distinct periods: November and December, January and February, May and June, July and August, and September and October. For each designated period, the culture initiation process commenced with the collection of 30 samples per interval. Sampling occurred once during the 2nd, 4th, and 6th weeks within an 8-week timeframe, totaling three sampling occasions over the span of 2 months. Consequently, the total sample size (n) amounted to 90 samples across all periods. This sampling protocol ensured systematic monitoring and assessment of cultures at multiple time points throughout the designated period. Precipitation and temperature data for the specified periods (2021–2022) were collected and recorded for the experiment.

Experiment 2: sanitation and thermotherapy programs

In November 2021, young ‘Tombul’, ‘Çakıldak’, and ‘Foşa’ suckers were selected and transplanted into 5-liter pots filled with a peat-perlite mixture in a 2:1 ratio. The pots were placed in the greenhouse with natural light, subjected to regular fungicide treatments (Aldeid®, active ingredient: 200 g/l Azoxystrobin + 125 g/l Difenoconazole) at 2-week intervals, and fertilized weekly with a 20-20-20 N-P-K plus micronutrients fertilizer (Grow-Up) at a concentration of 1000 ppm (Hand et al. 2016).

Sanitation program

In July 2022, new upper shoots from approximately 1-year-old plants, cultivated in a controlled environment within the greenhouse, were trimmed to a length of 30–40 cm. The preparation of explants and surface sterilization procedures were conducted according to the instructions provided in the ‘Initial surface sterilization’ section.

Thermotherapy

In June 2022, the potted plants were transferred to a growth chamber with a temperature of 25°C, a 16-hour photoperiod, and 55% relative humidity. The plants were kept there for one week. Afterward, the temperature in the growth chamber was increased to 37°C, and the plants were subjected to thermotherapy for 30 days, following the method described by Hu et al. (2015). Throughout the experiment, the plants were regularly watered, and fallen leaves were collected and removed. Once the thermotherapy ended, dried-out and dead plants were removed from the chamber, while the surviving plants were maintained at a temperature of 25°C and 55% humidity for 60 days. The plants were regularly fertilized with 20-20-20 N-P-K plus micro elements (Grow-Up), applied at a concentration of 1000 ppm on a weekly basis until the collection of explants. At the end of the 60-day period, newly developed shoots were brought to the laboratory for surface sterilization. The explants were prepared and subjected to surface sterilization following the guidelines outlined in the ‘Initial surface sterilization’ section.

Experiment 3: Surface sterilization

In January 2022, shoots were collected directly from 30-year-old ‘Tombul’ mother plants in the HRI collection orchard. The shoots underwent a series of steps for surface sterilization: first, the leaves and apex were removed, and the shoots were washed under running water. Afterward, the shoots were cut to a length of 2–3 cm, with each cut including a single axillary node. In the next step, various concentrations of disinfectants, including ethanol (70%), NaOCl (1%, 2%, 3%), H2O2 (3%, 6%, 9%), HgCl2 (0.1%, 0.5%, 1%), and AgNO3 (1%), were utilized for explant sterilization. The explants were exposed to these disinfectants for different periods of time, as specified in Table 1. After all the treatments, the explants were rinsed three times with distilled sterile water.

Table 1 Surface sterilization treatments with varying concentration and exposure durations used to assess the visually clean shoots, necrosis and viability percentages for mature field-grown ‘Tombul’ plants
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Culture and conditions

Following the surface sterilization procedure outlined in all experiments, the explants were placed into 16 × 100-mm culture tubes containing 5 ml culture medium. Murashige and Skoog (Murashige and Skoog 1962) medium mineral salts were supplemented with 30 g L− 1 sucrose, 5 mg L− 1 6-benzylaminopurine (BAP), 0.01 mg L− 1 indole-3-butyric acid (IBA), and solidified with 7 g L− 1 plant agar. The pH of the medium was adjusted to 5.7, followed by autoclaving at 120 °C for 20 min. The explants were cultured at 23 ± 2 °C under 16 h photoperiod using cool white fluorescent lamps (40 µmol m− 2 s− 1 light intensity).

Data analysis

Microbial contamination, necrosis, and bud break numbers were assessed through visual inspection and recorded on the 30th day of the culture period. Viability was defined by the presence of a green appearance in the explants, with no observable contamination or necrosis. The experiments were performed using a completely randomized design (CRD) and were independently replicated three times, with 30 explants per treatment (n = 90). Arcsine transformation was applied to the data before analysis. Data analysis was conducted using the SPSS 27 program. The normality of the data was assessed using the Shapiro-Wilk test, which yielded significant results (p ≤ 0.05) in all experiments. Means were analyzed with two-way ANOVA and differentiated using Tukey’s Honest Significant Difference test. The significance level was set at p < 0.05 for experiments 1 and 2, and p < 0.01 for experiment 3.

Results

Effects of explant collection period

The timing of explant collection significantly influenced visually clean shoot levels during culture initiation (p < 0.001) (Tables 2 and 3) (Fig. 1). The average clean shoots for all cultivars reached its highest point at 60% in May-June and was slightly lower at 53.7% in July-August. Conversely, the lowest average clean shoots were observed in November-December and January-February, at 8.9% and 12.6%, respectively. There were no significant differences found in the visually clean shoot percentage among cultivars (p = 0.472) (Table 3).

Table 2 Visually clean shoots and bud-break percentages of three mature field-grown C.avellana cultivars at different sampling periods
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Table 3 Analysis of variance and interaction between cultivars and sampling periods on visually clean shoots (%) and bud-break (%) of axillary nodes
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Fig. 1
figure 1

Average visually clean shoots and bud-break percentages in three cultivars’ cultures initiated with field-grown plants during five different periods, and mean precipitation of the study area (2022–2023, data source: Turkish State Meteorological Service). Significant differences in Tukey’s post hoc test at p ≤ 0.05 were indicated by different letters among the periods (Table 2) (n = 90)

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Bud sprouting varied significantly by different collection periods (p < 0.001) (Tables 2 and 3) (Fig. 1). Explants collected in July-August demonstrated the highest response across all cultivars, with a bud-break of 70%. In contrast, dormant explants collected in November-December and September-October exhibited poor success, with only 13.3% and 22.2% of buds sprouting, respectively. Furthermore, bud sprouting varied significantly among three cultivars (p = 0.047) (Table 3).

Effects of explant source; sanitation and thermotherapy programs

There was a significant increase in visually clean shoots when 1-year-old potted suckers were subjected to a sanitation program in the greenhouse and thermotherapy in the growth chamber compared to field-grown mature plants (p < 0.001) (Table 4). The average visually clean shoots were 97.4% for thermotherapy-treated plants, 75.6% for greenhouse plants, and 53.7% for field-grown plants (Fig. 2). However, clean shoots did not vary among the cultivars (p = 0.168) (Table 5).

Table 4 Visually clean shoots and bud-break percentages in cultures from mature field-grown plants, 1-year-old greenhouse-grown plants, and 1-year-old thermotherapy-treated plants
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Table 5 Analysis of variance and interaction between cultivar and treatments on visually clean shoots (%) and bud-break (%) of axillary nodes
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Bud sprouting varied significantly among different explant sources (p < 0.001). The success rate of culturing was high for buds collected from mature field-grown plants (73.3%) and young greenhouse-grown plants (73.3%). However, thermotherapy reduced bud sprouting rates across all three varieties, with an average sprouting rate of 47.4%. Significant necrosis was observed in buds subjected to thermotherapy. There was no notable variance in bud-break among the cultivars (p = 0.168) (Table 5).

Effects of surface sterilization

The impacts of different concentrations of, and exposure times to, NaOCl, HgCl2, AgNO3, and H2O2, both individually and in combination, on visually clean shoots, necrosis, and viability were investigated (Table 1). Higher concentrations and specific combinations of these sterilizing agents were more effective in obtaining visually clean and viable shoots. All NaOCl and H2O2 concentrations alone were ineffective at reducing contamination. Even at their highest doses, these treatments resulted in only 24.4% and 30% visually clean shoots, respectively. In contrast, HgCl2 and AgNO3 treatments alone showed significant improvement in obtaining uncontaminated shoots. HgCl2 resulted in 36.7–72.3% visually clean shoots, while AgNO3 yielded 54.4–81.1% visually clean shoots. However, these treatments had adverse effects, increasing necrosis. HgCl2 and AgNO3 caused the highest necrosis, 38.9% and 56.7%, respectively, among all treatments. Pre-treatment with ethanol and NaOCl, followed by HgCl2, AgNO3, and H2O2, generally improved contamination reduction and viability. Notably, certain treatment combinations achieved the highest visually clean and viable shoots. Overall, T19, T23, and T25 provided the best results, with viability ranging between 66.7% and 72.2%.

Fig. 2
figure 2

The effect of thermotherapy on visual microbial contamination (a) Plants propagated by suckers in November 2021 and maintained in the greenhouse for 6 months. (b) Plants regularly fertilized for 60 days after the thermotherapy (c) Shoots used to initiate in vitro cultures (d) Three weeks after culturing nodal explants from the thermotherapy treatment (e) Five weeks after culturing nodal explants from the thermotherapy treatment

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Discussion

Hazelnuts, like many other woody plants, face various challenges that can limit the success of culture initiation. These challenges include microbial contamination and difficulties in regenerating new plants from mature materials (Bassil et al. 1991; Diaz-Sala et al. 1990; Messeguer and Mele 1987; Hand et al. 2016). Establishing a successful micropropagation program requires optimizing all processes and beginning with clean cultures free from microbial contamination. While researchers have explored numerous methods to reduce microbial contamination, there is no single best method. Previous studies on hazelnuts have primarily focused on the explant source, the culturing period, and surface sterilization. Our study introduces the application of thermotherapy to mother plants as a novel approach to reduce microbial contamination during hazelnut in vitro culture establishment. This method was directly compared with conventional approaches to assess its effectiveness.

The timing of explant collection is closely associated with clean shoot rates during culture establishment (Messeguer and Mele 1983; Bassil et al. 1991; Yu and Reed 1995; Gao et al. 2008). However, existing studies on this subject have not reached a definitive consensus. In our study, the highest average visually clean shoot rates occurred in the May-June and July-August periods, at 60% and 53.7%, respectively, while the lowest rates were observed in November-December and January-February, at 12.6% and 8.9%, respectively (Table 2). Damiano et al. (2005) noted that clean cultures initiated with axillary buds from May to September varied between 5 and 60%, which aligns with our findings for the May-June period but shows a broader range. Similarly, Gao et al. (2008) reported lower clean shoot rates in late March (10.71%) and mid-April (58.04%) compared to mid-May (91.57), indicating a seasonal variation that partially matches our observation of higher clean shoots in late spring and summer. Contamination rates are influenced by weather conditions (Hand 2013). We observed a negative correlation between clean culture levels and monthly precipitation. During our study periods, the highest precipitation occurred in the autumn and winter months, correlating with the lowest clean shoots, while the lowest rainfall was recorded in the summer months, correlating with the highest clean shoots (Fig. 1). This observation (Fig. 1) confirmed that of Yu and Reed (1995) through their culture initiation using nodal explants from field-grown suckers. They noted that, of cultures initiated between March and September, the highest contamination was in June and was attributed to rain.

The timing of culture initiation impacts not only the contamination rate but also the in vitro response of the culture. We observed that the highest bud burst rate occurred in July-August (70%), gradually declining in September-October (22.2%), November-December (13.3%), before increasing again in winter months (52.2%) (Table 3) (Fig. 1). Prior research has also suggested that the success of culturing buds is low during the dormancy period but increases during active growth periods (Pincelli-Souza et al. 2018; Yu and Reed 1995). Yu and Reed (1995) reported that hazelnuts enter the dormancy period starting from September, resulting in low culture success for samples cultured after this month, decreasing from 43.33 to 3% from July to September. Yahyaoui et al. (2021) found that sampling in July (33%) and September (49%) produced more regenerated shoots than December and January. They achieved the lowest shoot proliferation sampling in December (4%). Our findings partially align with their data, confirming that summer months are more conducive to successful culture establishment. Previous studies on hazelnuts have shown that the early part of the growing season has a greater potential for in vitro response (Yu and Reed 1995; Bacchetta et al. 2005, 2008). Bacchetta et al. (2008) initiated in vitro cultures in two different physiological phases of the plants (spring and winter) and found higher success rates when collecting buds in spring. This could be attributed to the active growth phase of hazelnut trees during this season, leading to a higher success rate of shoot nodes collected during this period. In our current study, explants taken in May-June browned and became necrotic after surface sterilization, resulting in a large loss of culture. Similarly, Bacchetta et al. (2008) and Yahyaoui et al. (2021) reported that actively growing explants in spring were particularly sensitive to surface disinfectants, leading to tissue necrosis. Messeguer and Mele (1983) also showed that in spring, explants did not grow when they were less than 1.5 mm in diameter, so fewer viable shoots were formed during this period.

Considering the explant source, the results are in line with those reported by previous studies in hazelnut. Growing the donor plant in a controlled greenhouse generally yields explants with lower microbial loads than those grown in the field (Messeguer and Mele 1983; Yu and Reed 1995; Bacchetta et al. 2008). In our study, the suckers grown in the greenhouse were regularly treated with fertilizers and fungicides, and achieved 21.9% more clean shoots compared to material collected directly from the field (Table 4). Similar trends have been reported in earlier studies. Bacchetta et al. (2008) showed that growing plants in pots reduces contamination by about 20–30% compared to material collected directly from the field. Yu and Reed (1995) achieved success with shoots grafted onto rootstocks and grown in greenhouses. Similarly, Perez et al. (1987) found that sampling vegetative shoots from 12-month-old greenhouse-grown plants resulted in 70% clean explants. In addition to the environmental conditions surrounding the mother plant, the age of the plant from which the explant is sourced significantly influences contamination levels and culture success. Typically, cleaner cultures with higher rates of bud bursting success are obtained from explants taken from younger plants (Bassil et al. 1991; Diaz-Sala et al., 1990; Messeguer and Mele 1987). For this reason, recent studies have opted for juvenile plants or suckers of adult trees grown in greenhouse conditions as an explant source when initiating culture (Jyoti 2013; Hand et al. 2016; Elçicek 2020).

Thermotherapy has proven to be an efficient method to produce virus free plants (Kunkel 1936; Deogratias et al. 1989; Koubouris et al. 2007). In thermotherapy-based strategies, in vitro hazelnut plants infected with apple mosaic virus (ApMV) have been subjected to heat treatments (Elçicek 2020; Kaya 2021). There are very few studies using thermotherapy for reducing fungal and bacterial contaminants in plants. Torres et al. (2019) showed that applying humid heat to bamboo shoots for 5 and 10 min at 50 °C significantly reduced fungal contamination in culture. Linck et al. (2019) applied thermotherapy to raspberry and blackberry plants infected with phytoplasma and showed that all plantlets regenerated in tissue culture from these plants were negative for the bacterium. Likewise, Klimenko et al. (2020) managed to eliminate phytoplasma by applying thermotherapy to grape plants in vitro. Thermotherapy in hazelnut was used for the first time in this study as a contamination reducing agent in micropropagation. This technique includes thermotherapy of not sterile potted plants, followed by axillary node culture using newly emerged shoots. While this method effectively reduced contamination, the rate of bud bursting remained relatively low compared to explants obtained from greenhouse and field. Previous studies have noted that buds subjected to heat treatment may experience necrosis (Torres et al. 2019; Díaz-Barrita et al. 2008). In this study, the decrease in bud bursting percentage may be due to the damage caused by thermotherapy to plant buds. Despite this, cultures established with thermotherapy-treated plant explants achieved 97.4% clean shoots (Table 4). This is 43.7% more than mature field-grown trees and 21.8% more than plants maintained in the greenhouse and regularly treated with fungicides (Table 4). By treating mother plants with thermotherapy, we aimed to improve the initial cleanliness of cultures, potentially leading to higher success rates in micropropagation. Our findings offer an innovative solution to a persistent problem in hazelnut micropropagation, providing a new strategy that could complement existing methods.

Surface sterilization is critical, particularly when initiating culture with field-grown explants. The chemical agents employed in this process significantly impacted contamination, browning, phenolic compounds in culture medium, necrosis formation, viability, in vitro response of cultures, and ultimately regeneration. Consequently, the type, dose, and application duration of these disinfectants play a crucial role in the initial stages of tissue culture. Ethanol is a common disinfectant that is generally applied briefly before NaOCl at concentrations of 70–95% to increase the effectiveness of surface sterilization. While ethanol treatment may reduce contamination, it is important to note that it could potentially elevate the production of phenolic compounds and lead to browning in hazelnut buds, as reported by Bassil et al. (1991). Elçiçek (2020) applied 0.5-2% NaOCl for 10 min to nodal explants taken from the greenhouse and found that the viability rate in the 10-minute treatment of 20% NaOCl was 90.5%, 96% and 87% in Foşa Çakıldak and Tombul hazelnut varieties, respectively. However, in our study, application of 1–3% NaOCl for 20 min alone was not effective in reducing contamination in field-grown explants.

HgCl2 is typically utilized in the concentration range of 0.05–1% for sterilization in tissue culture (Hesami et al. 2019; Gammoudi et al. 2022; Dagne et al. 2023). In Corylus colurna micropropagation, Kosenko et al. (2008) utilized a 0.1% HgCl2 solution for 10 min to sterilize nodal explants. Similarly, Yahyaoui et al. (2021) employed a 0.1% HgCl2 solution along with ethanol, NaOCl and H2O2 for surface sterilization of Sicilian hazelnuts. In this study, the application of 0.1 HgCl2 for 10 min was not effective. The use of 0.5-1% HgCl2 solution for 10 min significantly decreased contamination but led to a high rate of necrosis. HgCl2 is highly effective in surface sterilization (Hesami et al. 2019). However, even in small amounts, HgCl2 can be toxic to plants and animals (Vaidya and Mehendale 2014; Ge et al. 2022), making it unsuitable for tissue culture (Da Silva et al. 2016).

AgNO3 is recognized for its antifungal and antibacterial properties (Mihaljević et al., 2013; Dagne et al. 2023). While AgNO3 has been utilized in surface sterilization for woody plants, its use in hazelnuts has not been documented previously. In our study, compared to NaOCl applications, both HgCl2 and AgNO3 were effective in reducing contamination, albeit at the cost of increased tissue browning and necrosis. However, doses of 0.5% and 1% of HgCl2 and application durations of 20 and 30 min for 1% AgNO3 resulted in adverse effects, indicating that these applications are not suitable for hazelnut node surface sterilization.

Considering all surface sterilization applications in this study, it was found that chemical agents provided more effective results when applied in combination. After the application of 70% ethanol for 30 s and 2% NaOCl for 20 min, the subsequent application of 0.1% HgCl2, 1% AgNO3, or 3% H2O2 yielded the highest results. Similarly, Yahyaoui et al. (2021) used a combination of 0.1% HgCl2 for 2 min, 3% H2O2 for 3 min, and 70% ethanol for 10 min in the sterilization of nodal explants of plants grown in the greenhouse. When choosing the appropriate surface sterilization method, it is crucial to prioritize practices that not only minimize contamination but also maintain high explant viability. Successful surface sterilization should effectively kill microbial contaminants without damaging plant cells and tissues.

Despite various efforts to maintain the mother plant in a healthy environment, choose the best time for culture initiation, and utilize effective surface sterilization agents, significant losses can still occur during the initiation and especially subculturing stages of in vitro culture due to endophytic microorganisms (Reed et al. 1998; Hand et al. 2016). Endophytic fungi and bacteria are not eradicated after surface sterilization and negatively impact the growth of in vitro plants. Internal bacteria were identified in C.avellana shoot culture using by surface sterilization followed by liquid contaminant detection medium (Reed et al. 1998; Hand et al. 2016). After applying thermotherapy to the mother plants, employing a similar approach as utilized by Hand et al. (2016) may aid in identifying contaminant-free cultures, consequently enhancing culture viability and success.

Conclusion

In conclusion, this study highlights the difficulties in establishing contamination-free hazelnut cultures from mature field-grown plants. The timing of explant collection plays a critical role, with July to August identified as the optimal period for culture initiation. Surface sterilization using a combination of NaOCl, HgCl2, and AgNO3 showed promise in reducing contamination, yet the potential toxicity of chemicals raises concerns. Alternatively, maintaining a sanitation program for mother plants proved to be more effective. Thermotherapy of mother plants followed by in vitro culture establishment yielded the highest visually clean shoots. Overall, these findings underscore the significance of timing, appropriate surface sterilization techniques, and the selection of explant sources to minimize contamination and improve success rates in hazelnut culture initiation.