Acacia mangium × A. auriculiformis micropropagation in a non-sterile environment

ABSTRACT

Autoclaving eliminates microbial contamination during micropropagation, but the process is complex, time-consuming and expensive. Chemical sterilisation also effectively disinfects culture media and is relatively simple and cost-effective. Prior studies have focused on the effects of chemical sterilisation on bud induction, but the effects of sterilant on proliferation and rooting are unknown. We investigated the effect of sterilant on Acacia mangium × A. auriculiformis bud induction, rooting and subculture rooting. The optimal bud induction medium comprised 1/8 Murashige and Skoog medium $+$ 7 g l⁻¹ agar $+$ 0.2 g l⁻¹ chlorothalonil $+$ 0.5 mg l⁻¹ 6-benzylaminopurine. The maximum bud induction rate (99.54%) with zero contamination was achieved using the third to fifth stem segments collected in October and treated with 0.8 g l⁻¹ carbendazim for 3 min. The maximum rooting rate (97.62%) was attained using a rooting medium consisting of 7 g l⁻¹ agar $+$ 0.2 g l⁻¹ chlorothalonil $+$ 1.5 mg l⁻¹ indolebutyric acid $+$ 0.5 mg l⁻¹ naphthaleneacetic acid. Proliferation ratio and subculture duration were positively correlated. The maximum proliferation rate (3.58%) was realised in the fourth subculture rooting. Chlorothalonil can effectively replace autoclaving of A. mangium × A. auriculiformis bud induction and rooting media. The present study provides insights for improving the rapid propagation method of Acacia sp. and a new direction for the development of micropropagation technology.

KEYWORDS:

• Acacia mangium × A. auriculiformis
• chemical sterilisation
• chlorothalonil
• micropropagation
• non-sterile

Introduction

The hybrid of Acacia mangium Willd. and A. auriculiformis A. Cunn. ex Benth (Samba et al. 2007) is a promising tree cultivar with superior growth characteristics, adaptability, fibre quality, pulping properties and insect and wind resistance compared with its parental lines (Galiana et al. 1998; Diouf et al. 2006; Yong et al. 2011; Monteuuis et al. 2013; Muhammad et al. 2018). However, asexual propagation is the only method by which the superior traits can be perpetuated (Kim et al. 2008; Sunarti and Insiana Putri 2009).

Micropropagation is a widely used method of asexual propagation (Yang et al. 2006; Dutra et al. 2009). It is characterised by a rapid micro-multiplication rate and high reproduction coefficients and conserves the traits of the variety cultured (Debergh and Maene 1981; Singh 2015). However, complex and time-consuming operating procedures and strict, meticulous technical requirements hinder the wide application of this technique (Loyola-Vargas et al. 2008). Micropropagation is expensive due to the equipment required, energy consumption, labour and other factors (Peiris et al. 2012). Moreover, sterilisation by autoclaving is time-consuming and increases micropropagation production costs (Macek et al. 1995; Zhai et al. 1996), but reducing culture medium contamination is one of the main challenges of plant propagation. It is important to sterilise the growth medium as a potential contamination source and eliminate microorganisms that hinder in vitro plant development and growth.

Chemical sterilisation using substances that eliminate fungi and bacteria from the culture medium represents a promising alternative to autoclave-based sterilisation. Chlorine dioxide (ClO₂) (Yanagawa et al. 1995; Cardoso 2009; Cardoso and Inthurn 2018), sodium hypochlorite (NaClO) (Yanagawa et al. 1995; Teixeira et al. 2006; Siekierzyńska and Litwińczuk 2018), peracetic acid (Cardoso 2009), and hydrogen peroxide (H₂O₂) (Snow 1986; Curvetto et al. 2006) have been evaluated as candidate chemical sterilants. Chlorothalonil is a protective fungicide that inhibits and kills bacteria, and it is also used as a plant-growth regulator (Van Scoy and Tjeerdema 2014). Chlorothalonil is widely applied because it is readily available and relatively inexpensive (Kishore and Pande 2005; Kiefer et al. 2020). We previously reported that chlorothalonil can be used as a chemical sterilant for A. auriculiformis bud induction (Huang et al. 2019). Compared with other chemical disinfectants, chlorothalonil is simple to apply and ensures low contamination and high germination rates (Huang et al. 2019). However, the sterilisation effect of chlorothalonil on other tree species and in the processes of proliferation and rooting has not been studied.

To address these issues, we used A. mangium × A. auriculiformis as the test material in this study and performed bud-induction and rooting tests in a completely open non-sterile environment. No autoclaving was required to sterilise the culture medium or containers. The work was not performed in a sterile environment. We aimed to find a high-efficiency, anti-polluting and low-cost sterilisation method and, to a certain extent, to reduce the difficulty of the experiments so that plants in the laboratory could grow normally in a non-sterile environment. For this reason, the present study investigated the impact of adding a sterilisation agent to non-sterile culture media at various stages of tree explant growth. We evaluated the feasibility of adding chlorothalonil to A. mangium × A. auriculiformis explant culture medium and assessed the impact of this sterilant on explant bud induction, rooting and subculture rooting. Thereafter, the best medium for bud induction and rooting of A. mangium × A. auriculiformis in a non-sterile environment was selected. This work provides technical support for a new sterilisation method and the rapid propagation of A. mangium × A. auriculiformis in the future and for industrial development.

Materials and methods

Materials

In 2014, elite A. mangium and A. auriculiformis trees were selected from plantations in Shadui Town, Jiangmen City, Guangdong province, China, and subjected to controlled pollination. Mature seeds were harvested and used to establish a hedge garden at 50 cm × 50 cm intervals. The hedge garden plants, used as ‘maternal plants’, were periodically fertilised, pruned and treated with fungicides to ensure healthy growth. Semi-lignified annual shoots with axillary buds were used as explants and were randomly selected in different maternal plants. Intellectual property rights were secured for study materials.

Explant pre-treatment

All experiments were conducted in 2016. Semi-lignified annual shoots with axillary buds were excised from the hedge garden, washed in tap water with a soft brush, soaked in water with washing powder for 30 min, and rinsed in tap water for 10 min. The stems were trimmed to uninodes with axillary buds until their lengths were in the range of 2.0–3.0 cm. They were then rinsed under running water for 1 h.

Sterilant screening

Carbendazim and chlorothalonil at 0.2, 0.4, 0.6 and 0.8 g l⁻¹ were used to determine the optimal bacteriostatic agent and concentration (Huang et al. 2019). After pre-treatment, 24 explants per treatment were inoculated in triplicate. Explant survival, contamination and browning rates were recorded after 30 days of culturing.

Explant disinfection

According to the method described by Huang et al. (2019), the collected stem segments were selected from the hedge garden in 2016 and used as explants. The latter consisted of upper stem segments with axillary buds from the first to second segments and leaves trimmed to approximately one-quarter of a whole phyllode and middle explants from the third to fifth stem segments containing axillary buds and with or without leaves.

All pre-treated explants were disinfected according to the protocol shown in Table 1. The sterilised explants were then inoculated into media containing optimal sterilant concentrations. In total, 21 explants were inoculated per treatment and repeated in triplicate. After 30 days of incubation, growth parameters (survival, contamination and browning rates) of each explant were recorded. The optimal explant types and disinfection strategies were selected.

Table 1. Explant type and disinfection scheme for Acacia mangium × A. auriculiformis bud induction

Explant type	Disinfection strategy
Middle stem without phyllode	0.8 g l^−1 carbendazim for 3 min
	75% (v/v) alcohol for 10 s
	75% (v/v) alcohol for 15 s
Middle stem with phyllode	0.8 g l^−1 carbendazim for 3 min
	75% (v/v) alcohol for 10 s
	75% (v/v) alcohol for 15 s
Upper stem with phyllode	0.8 g l^−1 carbendazim for 1 min
	75% (v/v) alcohol for 2 s
	75% (v/v) alcohol for 5 s

Bud induction medium

To obtain the optimal bud induction medium for A. mangium × A. auriculiformis in a non-sterile environment, we used modified Murashige and Skoog (MS) medium (1/2 macronutrient salts), 1/2 MS, 1/4 MS, 1/8 MS and blank controls (with only water and agar added to the medium), and these were tested with 0, 0.5 or 1.0 mg l⁻¹ 6-benzylaminopurine (6-BA) in each treated medium. The optimal explants were selected for pre-treatment and sterilised with 0.8 g l⁻¹ carbendazim for 3 min. The explants were then inoculated into the various bud induction media containing 0.2 g l⁻¹ chlorothalonil. Each treatment consisted of 24 explants and was repeated in triplicate. The survival, contamination, browning and budding rates were calculated after culturing for 30 days.

Explant collection times

Shoots with axillary buds were collected as explants in January, April, July and October 2016. The optimal explants were selected for pre-treatment and the ideal sanitisation strategy was used to disinfect them. The optimal induction medium was used to promote buds under routine laboratory conditions without strict bacterial control. A total of 72 explants from each of the foregoing months was selected for bud induction. The numbers of induced buds was recorded after 30 days of culturing and the optimal explant collection month was selected.

Rooting medium

To screen the best A. mangium × A. auriculiformis rooting medium, well-developed induced buds were inoculated into rooting medium containing 0.2 g l⁻¹ chlorothalonil and various concentrations of indolebutyric acid (IBA) and naphthaleneacetic acid (NAA) that were conducive to rooting (Table 2). In total, 24 induced buds were selected per treatment and all treatments were repeated in triplicate. Bud rooting was observed after 15, 30 and 45 days and the rooting rates were determined after 30 days of culturing.

Table 2. Indolebutyric acid (IBA) and naphthaleneacetic acid (NAA) concentrations used in rooting medium

Treatment no.	IBA (mg l^−1)	NAA (mg l^−1)
1	0
2	0.75
3	1.0	0.5
4	1.25
5	1.5
6		0
7		0.25
8	1.0	0.5
9		0.75
10		1

Proliferation by subculture rooting

Well-developed buds that had turned into plantlets after culture in the first-generation rooting medium were selected and inoculated into the optimal rooting medium containing 0.2 g l⁻¹ chlorothalonil. After 45 days of rooting, the seedlings were cut into 2.5-cm-long segments containing one or more axillary buds for subculture rooting. The numbers of inoculated seedlings per generation and seedlings obtained after rooting subculture were recorded. Each subculture was inoculated with 16 plantlets and there were three replicates per treatment.

Culture medium preparation

After adding all required nutritional components, the culture medium pH was adjusted to 5.8, and then 7.0 g l⁻¹ agar was added to the medium. The culture medium was heated to boiling in a microwave oven to dissolve the agar; when the temperature of the medium reached 50°C, 0.2 g l⁻¹ chlorothalonil was added to the medium, which was then stirred well. The medium was aliquoted and dispensed into culture vessels and cooled until the aliquots solidified for use in further experiments. All operations of culture medium preparation were carried out in a non-sterile environment on a laboratory bench, and the cultures were maintained under a white fluorescent light with a photon flux of 30 μmol m⁻² s⁻¹.

Data analysis

Analysis of variance was conducted using SPSS v. 19.0 for Windows (IBM Corp., Armonk, NY, USA). Significant differences (P < 0.05) between treatment means were determined using Duncan’s multiple range test. The following parameters were recorded:

Survival rate = number of surviving explants/total number of inoculated explants × 100%

Browning rate = number of browning explants/total number of inoculated explants × 100%

Contamination rate = number of contaminated explants/total number of inoculated explants × 100%

Bud induction rate = number of explants with bud induction/total number of surviving explants × 100%

Rooting rate = number of seedlings with roots/total number of surviving seedlings × 100%

Proliferation ratio = number of proliferating plantlets/total number of inoculated explants.

Results

Sterilant screening

The culture conditions and various sterilants added to the medium are listed in Table 3. The survival rates of A. mangium × A. auriculiformis explants differed significantly depending on the type of sterilant added to the medium (Table 3). Contamination levels were significantly (P < 0.05) higher in the media containing carbendazim than in those containing chlorothalonil (Table 3). Browning rates significantly (P < 0.05) increased but survival rates significantly (P < 0.05) decreased with increasing chlorothalonil concentration, and the browning mainly started from the bottom of the stem segment (Figure 1). In contrast, the contamination rates did not significantly (P > 0.05) differ among chlorothalonil concentrations (Table 3). The highest survival rate (80.55%) was determined for 0.2 g l⁻¹ chlorothalonil (Table 3).

Figure 1. Effects of chlorothalonil concentration on Acacia mangium × A. auriculiformis explants

Scale bars (top right-hand corner) = 1 cm.

Table 3. Effects of sterilants on Acacia mangium × A. auriculiformis explants

Sterilant	Concentration (g l^−1)	Survival rate (%)	Contamination rate (%)	Browning rate (%)
Chlorothalonil	0.2	80.6^a	7.0^b	12.5^b
	0.4	68.1^a	6.9^b	25.0^b
	0.6	63.9^a	9.7^b	26.4^b
	0.8	16.7^b	20.8^b	62.5^a
Carbendazim	0.2	18.6^b	56.9^a	25.0^b
	0.4	8.3^b	62.5^a	29.2^b
	0.6	12.5^b	65.3^a	22.2^b
	0.8	19.4^b	59.7^a	20.8^b
Control	0.0	31.9^b	54.2^a	13.9^b

Superscript letters indicate significant difference according to Duncan’s multiple range test at 5% probability. Data are means ± standard error.

Explant disinfection

There were significant differences in browning, contamination and survival rates among the various A. mangium × A. auriculiformis explant disinfection strategies (Table 4). For all disinfectant types and concentrations, browning rates were significantly higher and survival rates were significantly lower for the upper stems with phyllode and the middle stems without phyllode than those for the middle stems with phyllode. Hence, the middle stems with leaves were suitable for use as explants for A. mangium × A. auriculiformis bud induction (Figure 2a). The browning, contamination and survival rates did not significantly differ (P > 0.05) among the disinfection strategies. When the middle stems with phyllode were used as explants, the survival rate was 96.83% after treatment with 0.8 g l⁻¹ carbendazim for 3 min.

Figure 2. Bud induction of Acacia mangium × A. auriculiformis. (a) Bud induction of middle explants with leaves. (b) Induced buds after 30 days of culture

Scale bars (top right-hand corner in each slide) = 1 cm.

Table 4. Effects of explant type and disinfection strategy on Acacia mangium × A. auriculiformis bud induction

Explant type	Disinfection strategy	Survival rate (%)	Contamination rate (%)	Browning rate (%)
Middle stem without phyllode	0.8 g l^−1 carbendazim for 3 min	28.6^bc	9.5^ab	61.9^b
	75% (v/v) alcohol for 10 s	47.6^b	0.0^b	52.4^bc
	75% (v/v) alcohol for 15 s	38.1^b	0.0^b	61.9^b
Middle stem with phyllode	0.8 g l^−1 carbendazim for 3 min	96.8^a	0.0^b	3.2^d
	75% (v/v) alcohol for 10 s	76.2^a	14.3^ab	9.5^d
	75% (v/v) alcohol for 15 s	90.5^a	0.0^b	9.5^d
Upper stem with phyllode	0.8 g l^−1 carbendazim for 1 min	38.1^b	23.8^a	38.1^c
	75% (v/v) alcohol for 2 s	6.4^c	4.8^ab	88.9^a
	75% (v/v) alcohol for 5s	4.8^c	9.5^ab	85.7^a

Superscript letters indicate significant difference according to Duncan’s multiple range test at 5% probability. Data are means ± standard error.

Bud induction medium

The contamination rate decreased to <10% after adding chlorothalonil to the bud induction medium (Table 5). After adding different concentrations of MS basic medium to the induction medium, the browning rate decreased significantly with the decrease of MS concentration. No explant contamination was observed when 1/8 MS was used; survival rate was >90%, and the bud induction rate was higher than that for all other media. When 0–1 mg l⁻¹ 6-BA was used, the budding rate increased initially and decreased thereafter. The budding rate was highest in the presence of 0.5 mg l⁻¹ 6-BA (Table 5). Therefore, the optimal A. mangium × A. auriculiformis bud induction medium comprised 1/8 MS +0.5 mg l⁻¹ 6-BA and the maximum budding rate therein was 92.55%. The induced buds were about 2–3 cm and grew vigorously (Figure 2b).

Table 5. Effects of medium type and 6-benzylaminopurine (6-BA) concentration on Acacia mangium × A. auriculiformis bud induction

Treatment medium type	6-BA (mg l^−1)	Survival rate (%)	Contamination rate (%)	Browning rate (%)	Budding rate (%)
1/2 MS	0	11.1^cd	8.3^a	80.6^a	0.0^c
	0.5	56.9^b	1.4^a	41.7^b	16.7^c
	1	26.4^c	1.4^a	72.2^a	6.7^c
1/4 MS	0	55.6^b	0.0^a	44.4^b	2.1^c
	0.5	68.1^b	2.8^a	29.2^bc	45.5^b
	1	56.9^b	1.4^a	41.7^b	2.2^c
1/8 MS	0	90.2^a	0.0^a	9.7^cd	92.1^a
	0.5	94.5^a	0.0^a	5.6^d	92.6^a
	1	90.3^a	0.0^a	9.7^cd	89.4^a
Modified MS	0	5.6^cd	4.2^a	90.3^a	0.0^c
	0.5	23.6^cd	4.2^a	72.2^a	4.8^c
	1	1.4^d	4.2^a	94.4^a	0.0^c
Control	0	11.1^cd	8.3^a	80.6^a	0.0^d
	0.5	5.6^cd	4.2^a	90.3^a	3.0^d
	1	55.6^b	0.0^a	44.4^b	2.1^d

Superscript letters indicate significant difference according to Duncan’s multiple range test at 5% probability. Data are means ± standard error. MS = Murashige and Skoog.

Explant collection time

The bud induction rates differed significantly (P < 0.05) among the explants collected in different months (Figure 3). The bud induction rates were lowest in January (66.21%) and highest in October (99.54%). The bud induction rates of the explants collected in April and July were slightly lower than those collected in October but were nonetheless > 95%; this showed that the growth status of A. mangium × A. auriculiformis seedlings would be negatively affected in low-temperature conditions. Thus, April to October was the optimum period to collect explants for bud induction.

Figure 3. Bud induction rates in Acacia mangium × A. auriculiformis explants collected in different months

Different letters indicate significant differences at P < 0.05 according to Duncan’s multiple range test.

Rooting medium

Various concentrations of IBA and NAA had significant (P < 0.05) effects on rooting (Table 6). For treatments 1–5, the rooting rate increased with IBA concentration when the NAA concentration was fixed at 0.5 mg l⁻¹. For treatments 6–10, increasing the NAA concentration increased and then decreased the rooting rate when the IBA concentration was fixed at 1 mg l⁻¹. The maximum rooting rate was 97.62% in the presence of 1.5 mg l⁻¹ IBA and 0.5 mg l⁻¹ NAA (Table 6). Therefore, high IBA concentrations improved rooting potential, and the optimal NAA concentration was in the range of 0.5–1.0 mg l⁻¹. Roots began to emerge after 12 days of culturing (Figure 4a A1), and growth was vigorous (Figure 4a A2). After 20 days of culture, the growth of rooted seedlings became stable (Figure 4b B1 and B2).

Figure 4. Acacia mangium × A. auriculiformis rooting culture. Rooted seedlings cultivated for (a) 12 days, (b) 20 days and (c) 45 days

Scale bars (top right-hand corner in each slide) = 1 cm.

Table 6. Effects of different indolebutyric acid (IBA) and naphthaleneacetic acid (NAA) concentrations on Acacia mangium × A. auriculiformis rooting

Treatment no.	IBA (mg l^−1)	NAA (mg l^−1)	Rooting rate
1	0		31.68^ef
2	0.75		19.28^f
3	1	0.5	39.04^de
4	1.25		64.56^bc
5	1.5		97.62^a
6		0	18.38^f
7		0.25	26.28^ef
8	1	0.5	65.60^bc
9		0.75	77.78^b
10		1	52.54^cd

Superscript letters indicate significant differences at P < 0.05 according to Duncan’s multiple range test.

Proliferation by subculture rooting

After 45 days of rooting culture, seedling roots had grown to approximately 5 cm in length (Figure 4c C1 and C2). The seedlings were then cut into 2–3 sections for subculture rooting. The proliferation ratio increased significantly with subculture duration. The maximum proliferation ratio (3.58) was attained by the fourth subculture (Figure 5).

Figure 5. Proliferation rates in Acacia mangium × A. auriculiformis rooting subcultures

Different letters indicate significant differences at P < 0.05 according to Duncan’s multiple range test.

Discussion

Sterilant screening

Micropropagation is an important technique for plant cloning, with the sterile environment required for this method mainly relying on the decontamination of culture vessels and media by autoclaving. In recent years, relatively simple and low-cost chemical sterilisation methods have also been used in the in vitro culture of various plants. For example, during the in vitro rooting of pineapple shoots (Oliveira et al. 2015), the proliferation and rooting of sugarcane shoot tips (Tiwari et al. 2012), the callus induction process of Cochlospermum regium (Schrank) Pilg (Gavilan et al. 2018), and the in vitro establishment of Eucalyptus (Molinari et al. 2021) and Sequoia sempervirens L. (Ribeiro et al. 2011), autoclaving of media and glass vessels can be replaced by chemical sterilisation using NaClO. During in vitro establishment, the stages of root elongation of gerbera (Cardoso and Teixeira da Silva 2012) and elongation and rooting of Anthurium andraeanum Linden ex André (Cardoso 2009) and potato seedlings (Duan et al. 2019) can be chemically sterilised using ClO₂. During the induction of adventitious buds and regenerated plants from lily scales, chemical sterilisation with H₂O₂ can achieve better plant growth performance than using autoclave sterilisation (Curvetto et al. 2006). Finally, in vitro culture of Gossypium hirsutum L. using sterilisation via n-hexane in combination with H₂O₂ yielded the lowest contamination and highest germination rates, with the germinated seedlings healthy, fresh and rooted robustly in the medium (Bakhsh et al. 2016). Together, such results indicate that selection of the appropriate chemical bacteriostatic agents for different cultures can be an effective means for sterilising the medium during in vitro propagation of plants.

In the present study, we compared the sterilisation effects of chlorothalonil and carbendazim. Chlorothalonil mainly disrupts the metabolism of pathogenic bacteria to inhibit their vitality and prevents the infection and spread of pathogenic bacteria. Carbendazim inhibits spindle formation during the cell division of pathogenic bacteria, thereby inhibiting the growth of hyphae. Our results demonstrated that carbendazim did not have an obvious sterilisation effect on the culture medium because the contamination rate exceeded 50%; moreover, the survival rate was very low after adding carbendazim at different concentrations. These findings indicate that Acacia seedlings are sensitive to carbendazim, with the use of carbendazim having a toxic effect thereon. Similarly, Brondani et al. (2013) reported negative effects of sterilant agents on in vitro cultures at higher concentrations, while emphasising the positive effects at lower concentrations. Thus, sterilant agents added to a culture medium can damage tissues or exert beneficial effects, depending on the concentration, indicating the need to establish the optimal concentrations for each explant type. In the present study, the low concentration of chlorothalonil controlled the contamination rate to below 10% and obtained a high survival rate of plants. Chemical sterilisation during rooting and sub-proliferation was also suitable, and no toxic effects were observed on the growth of plants. Therefore, we speculate that when a high concentration of chlorothalonil disrupts the metabolism of pathogenic bacteria, it also inhibits the growth of Acacia seedlings. Additionally, excessive accumulation of chlorothalonil inhibits the activity of indoleacetic acid oxidase, resulting in the destruction of the endogenous plant auxin indoleacetic acid, affecting plant growth (Díaz Rodríguez et al. 2019). Thus, lower concentrations of sterilant agents can significantly promote plant growth.

Explant disinfection

In vitro explant establishment varies with type, size, sampling site (Murashige 1974) and disinfection method (Niedz and Bausher 2002; Iliev 2008; Borges et al. 2012). The explant source significantly influences contamination rates and axillary bud growth. Here, phyllode and without-phyllode mid-stem segments and stem tips without phyllode served as explants. The semi-woody mid-stem segment was the most suitable explant (Shi et al. 2015).

Under natural conditions, both the interior and exterior of an explant harbour bacteria and mould that must be completely eliminated during in vitro reproduction (Niedz and Bausher 2002; Chen and Yeh 2007; Yong et al. 2011). Explant pre-treatment effectively lowers contamination rates. Different types of explants require different disinfection methods. Selecting the appropriate disinfectant type, concentration and disinfection time are all crucial for successful explant pre-treatment and disinfection. In the present study, explants soaked in 0.8 g l⁻¹carbendazim for 3 min were free of contamination and their survival rate was 96.83%. Thus, the disinfection program must be customised according to the explant type, location and size as well as the sterilant type and disinfection time.

Bud induction medium

Studies have shown that, during the in vitro induction of sugarcane sprouts, 0.1% NaClO can be used to sterilise the medium, with comparable effects on survival rate, bud number and bud viability to those on the cultures grown on autoclaved nutrient medium (Tiwari et al. 2012). Gerbera shoot tips grown in medium sterilised with ClO₂ showed similar or better development to that resulting from autoclaved medium; sterilisation of the medium with 0.0025% ClO₂ during the in vitro establishment phase of shoot tips resulted in better plant development; and the percentages of calluses formed or regenerated shoots were similar in both treatments (Cardoso and Teixeira da Silva 2012). These results suggest that the use of appropriate bacteriostatic agents can both effectively control the contamination of the medium and facilitate the growth of the cultured buds. In our prior study of A. mangium × A. auriculiformis bud induction, in which the medium was sterilised by autoclaving, we observed a contamination rate of 3.33% and an induction rate of 97.33% (Wang et al. 2016). In the present study, 0.2 g l⁻¹ chlorothalonil was used to sterilise the medium, yielding a contamination rate of zero (Table 5). Moreover, in the explants collected in November that were inoculated in 1/8 MS medium supplemented with 0.5 mg l⁻¹ 6-BA, the induction rate was 99.54% (Figure 3). Thus, chemical sterilisation of the medium yielded better results than autoclaving for A. mangium × A. auriculiformis bud induction.

The success of the in vitro bud induction process depends on various factors; for example, the concentration of the medium can affect the in vitro culture. The results of this study showed that bud induction in A. mangium × A. auriculiformis was significantly affected by MS salt concentration, resulting in the highest browning rate, indicating that growing A. mangium × A. auriculiformis explants were highly sensitive to a medium with high salt concentrations. Inhibition of shoots in the presence of high concentrations of salt may be due to osmotic and toxic effects. Studies have shown that an excessive concentration of inorganic salts in the medium leads to a large amount of phenolic substances in the explants, which promotes the oxidation reaction and induces browning. Moreover, the osmotic stress induced by the salt-rich medium may affect the metabolism of plant tissues, which stimulates ready oxidation and leads to plant growth and the release of toxic compounds (Rodriguez et al. 2014). However, we found that the browning rate of the control group without adding salt was also high, which is inconsistent with the conclusion discussed above. Thus, the specific reasons underlying the significant decrease in browning rate observed with decreased medium concentration to the culture medium requires further exploration.

Overall, in this study, we found that adding 0.5 mg l⁻¹ 6-BA to the low MS medium concentration (1/8) provided a favourable environment for the survival of Acacia seedlings, with no contamination occurring in the medium after adding 0.2 g l⁻¹ chlorothalonil (Table 5). Notably, explants collected in October that were selected and cultured in the optimal medium exhibited an induction rate of 99.54%.

Rooting medium

Several studies have addressed the selection and optimisation of rooting medium type, sucrose concentration and phytohormone type and concentration (Vahdati et al. 2004; Akila et al. 2011; Dewir et al. 2015). A few reports examined autotrophic micropropagation technology (Kubota 2001; Valero-Aracama et al. 2001; Ding et al. 2010) and demonstrated that sucrose must be included in the culture medium used for traditional micropropagation. However, sucrose should not be added to the culture media used in photoautotrophic micropropagation systems. Instead, the explants must assimilate atmospheric carbon dioxide and undergo photosynthesis. In the present study, sucrose was omitted from the culture medium because the latter was non-sterile. No carbon substitute was added to the medium. Forced to grow in this non-sterile environment, the seedlings better adapted to the external environment during the subsequent domestication process. The addition of appropriate concentrations of plant-growth regulators ensured optimal explant rooting rates. The phytohormone concentration markedly affected rooting. The high rooting rate in the presence of chlorothalonil showed that it did not negatively impact plant growth regulators. Future research should endeavour to determine the optimal phytohormone concentration that promotes culture growth and to identify and evaluate sucrose substitutes that serve as carbon sources and inhibit bacterial growth in the culture media.

Chemical disinfectants significantly lower contamination rates in plant media. However, few studies have focused on the effects of sterilants on rooting. In the present study, 0.2 g l⁻¹ chlorothalonil was added to the culture medium as a sterilant; compared with autoclaved media, the rooting rate (99.43%) (Shi et al. 2015) was not significantly different from the media supplemented with chemical sterilants (97.62%). Hence, chemical sterilisation can effectively replace autoclaving because it lowers the contamination rate without inhibiting rooting. However, the effects of chemical sterilisation on culture growth were not apparent based on the rooting rate analysis alone. Other growth performance parameters after transplantation must also be evaluated.

Proliferation by subculture rooting

Several studies reported the successful proliferation of explants grown on chemically sterilised culture media (Zhao et al. 2009; Huang and Mo 2012). NaClO and sugar were added to banana medium and the sterilant proved to be highly phytotoxic (Huang and Mo 2012). Most cluster buds presented with severe browning in the initial stages of inoculation and the bananas did not thrive. H198 was used as a sterilant in konjac micropropagation but caused most explants to turn brown during proliferation (Zhao et al. 2009). In the present study, 0.2 g l⁻¹ chlorothalonil was added to the proliferation culture medium and had no apparent negative effect on differentiation or proliferation. Neither browning nor contamination was detected, indicating that chlorothalene can be used as a sterilant for the non-sterile reproduction of A. mangium × A. auriculiformis.

In the present study, seedling height was approximately ≥5 cm after 45 days of rooting culture. The seedlings were cut into 2–3 segments and successfully proliferated via the rooting subculture. The proliferation rate was 3.58% at the fourth subculture; in contrast, it was 2.52% for the traditional micropropagation (proliferation was activated by inducing cluster buds). Hence, the culture method used in the present study was superior to the traditional micropropagation method in terms of the proliferation rate. The explants developed roots relatively more easily after the first rooting subculture. The number of days required for rooting gradually decreased with increasing subculture frequency. The average time required for single-subculture rooting was approximately 40 days. The rooting culture environment resembled the nursery culture. Therefore, the seedlings could be directly transplanted without acclimatisation. When sterilised by autoclaving, the proliferation time of A. mangium × A. auriculiformis explants is 40 days, the rooting time is 15 days, and the acclimatisation period is 5–10 days (Shi et al. 2015). Hence, the chemical sterilisation methodology used in the present study was superior to the autoclaving method because the former can save seedling acclimatisation time while improving seedling utilisation rates.

Author contributions

LH conceived and designed the experiments. HW performed the experiments. YL, LH and HW analysed the data. YL and LH wrote the paper. YL and LH revised the paper. All authors read and approved the final manuscript.

Disclosure statement

No potential conflict of interest was reported by the author(s).

Additional information

Funding

This work was supported by a grant from the Fundamental Research Funds of CAF [No. CAFYBB2017MB009]. The funders had no role in study design, data analysis or manuscript preparation.