Processing math: 100%
ARTICLE   Open Access    

Characterization and identification of rhizobacteria associated with Liberica and Robusta coffee rhizosphere

More Information
  • Coffee is a viable agricultural commodity that makes a positive impact to the Philippine economy. However, with an increasing trend in domestic consumption, the local coffee production has declined. Chemical fertilization has been considered by many farmers to improve coffee production and yield but it causes a serious threat to public health and the environment. Biofertilizer using rhizobacteria has beneficial effects to improve the growth and yield of many crops, which is cost effective and safer than synthetic fertilizers. This study characterized the indigenous and beneficial rhizobacteria obtained from the Liberica and Robusta coffee rhizosphere, in terms of phosphate solubilization, biocontrol activities, and tolerance to abiotic stresses. Six rhizobacterial isolates were molecularly identified and belonged to genera Bacillus, Burkholderia, and Pantoea. These rhizobacteria solubilized inorganic phosphate with solubilization index ranging from 2.5 to 3.5 mm. For biocontrol activities, Bacillus sanguinis showed activity in terms of HCN and multiple hydrolytic enzymes production. Also, Burkholderia sp. demonstrated amylase, protease, and pectinase activities. Moreover, all isolates were found to be relatively tolerant to a wide range of pH and concentrations of salt and heavy metals. The performance of these rhizobacterial isolates in terms of phosphate solubilization, biocontrol activities, and tolerance to stresses is promising and shown to have potential in coffee cultivation in the Philippines.
  • Greenhouse cultivation is a widespread method of cultivating high-quality plants to improve survival and adaptability. However, with the impacts of global climate change, abiotic stresses, such as higher temperature or stronger light intensity due to higher solar radiation, are becoming a great threat to plants especially in tropical, subtropical and temperate regions, consequently retarding plant growth or causing plant death under greenhouse cultivation[14]. To date, shade practice has been a common and effective method that alleviates or prevents the damage on plant cultivation under greenhouse conditions[2, 57]. Aied et al.[2] reported that eggplant grown under greenhouse conditions with 30%–70% shading, performed better than that grown in the open field.

    Light is an important factor that affects plant growth[7, 8]. Shade practice reduces light intensity under greenhouse cultivation, and excessive or slight shading causes unfavorable light intensity and further affects plant growth, hence, it is necessary to determine the optimal shading levels for plant growth. In practice, shading levels in greenhouse cultivation depend on plant species and genotypes. Slight shade (25%–50% of full sunlight) was suitable for Azolla growth[9]. Two dwarf Lisianthus species showed the best growth under 20% shade condition in a study performed by Rezai et al.[10].

    Plants respond to shading conditions through morphological changes and physiological behaviors. With regard to morphology, suitable shading conditions benefited the morphological development of Anthurium andreanum[7] and increased the plant height, internode length, leaf number, and leaf area[11]. Meanwhile, a well-developed root system is formed to increase nutrient absorption capacity[12,13]. In contrast, many plants under light-limiting or excessive light conditions showed undesirable morphological traits and reduced plant quality[11]. Regarding physiological behaviors, the biosynthesis and degradation of chlorophyll is actively regulated by light[14,15], and suitable shading levels upregulate chlorophyll content[8, 11]. In addition, relative water content (RWC) and soluble sugar content are also related to shading levels[11,16]. Moreover, the massive production of reactive oxygen species (ROS) owing to abiotic stress leads to accelerated activity of antioxidant enzymes, such as superoxide dismutase (SOD), catalase (CAT), and peroxidase (POD) in plant cells to minimize the negative impacts[11]. In all, various physiological activities of plants are in order when plants are cultivated under greenhouse cultivation with suitable shading conditions.

    Cyclocarya paliurus (Batal) Iljinskaja (C. paliurus), belonging to the family Juglandaceae, is a native, medicinal and ornamental tree distributed in the highlands of southern China[17]. In recent years, the demand for its leaf production has increased with the exploration and exploitation of its medical value because the leaf contains many bioactive compounds, such as triterpenoids, polysaccharides, and phenolic compounds etc., which benefit human health[1820]. The harvesting of its leaf was mainly from natural forests of Cyclocarya species, resulting in serious damage to the natural forests. Moreover, the shape of Cyclocarya species can be used as ornamental trees in urban parks due to its graceful shape and copper-like fruits. Thus, it is necessary to develop artificial cultivation of Cyclocarya species.

    Seedlings are the only way to artificially cultivate Cyclocarya species, and seedling cultivation has an important effect on the quality of artificial cultivation. Previous reports on seedling cultivation of C. paliurus only focused on improving leaf bioactive content using light and fertilization treatments under laboratory conditions[2124]. For example, Deng et al.[21] reported that leaf production per seedling increased under intermediate shade and fertilization treatments, and Yang et al.[23] investigated the effect of differential light quality and intensity on the growth and water-soluble polysaccharides of C. paliurus. However, in the actual process of forestry breeding, seedling is often cultivated in the open field. Considering the difference in experimental controllability under laboratory conditions, changeable environmental factors in the open field directly or indirectly lead to the uneven growth and low quality of seedling. This reduced the survival and adaptability of seedlings after transplantation. Therefore, it is necessary and urgent to cultivate high-quality seedlings of Cyclocarya species.

    Plastic greenhouse cultivation has been used to cultivate seedling growth of Cyclocarya species[25]. Feng et al.[25] reported that 50%–60% soil water capacity in plastic greenhouses significantly improved seedling quality. However, during the process of seedling cultivation, seedling growth was inhibited or seedlings died which was found to be caused by higher temperatures, especially in summer. To avoid or alleviate this problem, we hypothesized that shade could improve the seedling quality of C. paliurus under plastic greenhouse conditions. Therefore, the objective of this study was to determine the optimal shading level for C. paliurus seedlings from June to September and obtain high-quality seedlings based on their growth performances and physiological behaviors under different shading levels.

    Seeds were collected from the natural forest of Jinggangshan (Jiangxi province, China, named JX), and the collected seeds were treated using the method of Fang et al.[17]. After stratification treatment for five months, the germinated seedlings were first transplanted into cultivation bags (10.8 cm in diameter and 14.5 cm in height) filled with a medium (a mixture of nutritional soil: perlite (10:1, g/g)) and then cultured in an incubation room at 15 ± 2 °C (day) and 10 ± 2 °C (night), with natural light environment (about 15 ± 1 h photoperiod). Each bag contained one plant.

    Seedlings with even growth (height, marked H1) were selected and exposed to different shading levels in a plastic greenhouse under natural condition. Shade nets were set at 1.8 m height from the ground with different layers of shade net: no layer of shade net (marked SH0, as control, 60%–70% of full sunlight), one layer of shade net (marked SH1, 30%–40% of full sunlight) and two layers of shade net (marked SH2, 10%–20% of full sunlight); and plastic sheeting were not fitted around the plastic greenhouse. Soil water capacity was maintained at 50%–60%, and the lost water was supplemented at approximately 6:00 AM every day, and the position of the seedling was adjusted every 5 d. The experiment was established in a randomized complete block design, and each shading treatment contained three replicates and 15 seedlings per replicate.

    After 100 d of shading treatment (from June to September), the surviving seedlings were observed and investigated for growth analysis. Three seedlings were randomly selected and separated into leaves, shoots, and roots to determine and analyze their water content (WC) and mineral content, and the fourth compound leaf from the top to bottom with the same orientation were obtained from other seedlings and then immediately frozen in liquid nitrogen to determine and analyze their total soluble sugar (TSS) content, antioxidant enzyme activity, chlorophyll content and malondialdehyde (MDA) content.

    Seedling height (the final height, marked H2), ground diameter (DH), and leaf area (LA) of the seedlings under each shading treatment were measured. After washing, three parts of seedling were weighed (marked FW), and then heated at 105 °C for 15 min and dried to a constant weight (marked DW) at 85 °C. Finally, the growth and biomass of seedlings were assessed and the growth index was calculated according to the method described by Feng et al.[25].

    Based on the measurement of FW and DW, WC was calculated with the formula 1[26]:

    WC(%)=FWDWFW×100 (1)

    According to the method described by Feng et al.[25], dried samples were digested and extraction was obtained using the electric-heating digestion method by the Block Digestion System (AIM600, Clontarf Qld 4019 Australia), then the mineral content was measured using Inductively Coupled Plasma - Optical Emission Spectrometer (ICP-OES, optima 7000DV, PerkinElmer Co., USA), and finally the mineral content (potassium (K), calcium (Ca), magnesium (Mg), sodium (Na)) and translocation factor (TF) were calculated according to Eqns 2 & 3:

    C(mg/g)=Cm×VDW (2)
    TF=Cshoot+CleafCroot (3)

    Cm is the mineral content measured by ICP-OES, mg/L; V is the total volume of extraction, L; DW is the dried weight of the sample, g; Cshoot, Cleaf, and Croot are the mineral content in the shoots, leaves and roots, respectively.

    Total soluble sugar (TSS) was extracted and measured according to the method described by Li[27]. First, fresh leaves were ground and added to 10 mL distilled water, and then heated at 100 °C for 30 min, finally filtered to a volume of 25 mL. Next, 0.5 mL extraction was added to a mixture of 1.5 mL of distilled water, 0.5 mL of Anthrone and ethyl acetate mixture (Anthrone:ethyl acetate = 1:50 g/mL) and 5 mL of H2SO4, and then kept at 100 °C for 1 min. Finally, the absorbance was measured at 630 nm using a UV-visible spectrophotometer (TU-1900, Beijing Purkinje General Instrument Co., Ltd, China). TSS content was calculated using Eqn 4:

    C(%)=Cs×VW×0.5×1000000×100 (4)

    Cs was obtained from the standard curve in which sugar was used as the standard curve, mg/L; V is the total volume of extraction, mL; W indicates the fresh weight of the sample, g.

    Chlorophyll content was determined using the method explained by Li[27]. Briefly, fresh leaves were ground and added to 10 mL 80% acetone, and then filtered to a volume of 25 mL. Then, the absorbance was measured using a UV-visible spectrophotometer (TU-1900, Beijing Purkinje General Instrument Co., Ltd, China) at 470 nm (A470), 663 nm (A663) and 645 nm (A645). Finally, chlorophyll content and carotenoid content were calculated using Eqns 5, 6, 7 & 8 respectively.

    Ca=13.95A6656.88A649 (5)
    Cb=24.96A6497.32A665 (6)
    Cx.c=1000A4702.05Ca114.8Cb245 (7)
    Chlorophyllcontent(mg/g)=(Ca+Cb)×VW (8)

    V is the total volume of extraction, mL; W indicates the fresh weight of the sample, g.

    MDA content was extracted from fresh leaves using an improved thiobarbituric acid-malondialdehyde (TBA-MDA) assay method[27], and then measured at 450 nm, 532 nm and 600 nm by a UV-visible spectrophotometer (TU-1900, Beijing Purkinje General Instrument Co., Ltd, China), respectively. Finally, MDA content was calculated using Eqns 9 & 10, respectively:

    C(umoLL)=6.45(A532A600)0.56A450 (9)
    MDAcontent(umoL/g)=C×VW (10)

    V is the total volume of extraction, L; W indicates the weight of the fresh sample, g.

    Fresh leaves were ground and added to 15 mL 0.05 mol/L phosphate buffer (PBS, pH 7.8), then kept at 4 °C for 15 min, finally centrifuged at 4 °C, 11,000 ×g for 10 min (TGL-16M, Hunan Xiangyi Laboratory Instrument Development Co., Ltd, China). Enzyme extraction (labeled E) was obtained and antioxidant enzyme activity was immediately determined.

    SOD activity was determined using the method explained by Li[27] with slight modifications. Reaction mixture including 0.1 mL enzyme extraction (labeled E1), 3.0 mL 0.05 mol/L PBS (pH 7.8), 0.6 mL 130 mol/L methionine, 0.6 mL 0.1 mmol/L ethylene diamine tetraacetic acid disodium, 0.4 mL distilled water, 0.6 mL 0.02 mmol/L riboflavin and 0.6 mL 0.75 mmol/L nitro-blue tetrazolium was reacted at 4,000 lx for 20 min, but the control was kept in the dark. After that, the absorbance was measured at 560 nm by a UV-visible spectrophotometer (TU-1900, Beijing Purkinje General Instrument Co., Ltd, China).

    POD activity was measured according to the method described by Li[27] with slight modification. Reaction mixture containing 2.9 mL 0.05 mol/L PBS (pH 7.8), 1 mL 2% H2O2 (v/v), 1 mL 50 mmol/L guaiacol and 0.1 mL enzyme extraction (labeled E2) was heated at 34 °C for 3 min. Then, the absorbance was immediately measured at 470 nm, 6 times with an interval of 1 min by a UV-visible spectrophotometer (TU-1900, Beijing Purkinje General Instrument Co., Ltd, China).

    Polyphenol oxidase (PPO) activity was determined using the method described by Li[27] with slight modification. Reaction mixture containing 3.5 mL 0.05 mol/L PBS (pH 7.8), 1 mL 0.1 mol/L catechol and 0.5 mL enzyme extraction (labeled E3) was heated at 37 °C for 10 min. Then, 2 mL 20% trichloroacetic acid (w/v) was quickly added to the reaction mixture and centrifuged at 4 °C, 11,000 ×g for 10 min (TGL-16M, Hunan Xiangyi Laboratory Instrument Development Co., Ltd, China). Finally, the absorbance was measured immediately at 420 nm, 6 times with an interval of 1 min by a UV-visible spectrophotometer (TU-1900, Beijing Purkinje General Instrument Co., Ltd, China).

    CAT activity was measured using the method described by Li[27] with slight modification. Reaction mixture was prepared using 2 mL 0.05 mol/L PBS (pH 7.8), 0.5 mL 0.18% H2O2 (v/v), 1 mL distilled water and 0.5 mL enzyme extraction (labeled E4). The absorbance was immediately measured at 240 nm, 6 times with an interval of 1 min by a UV-visible spectrophotometer (TU-1900, Beijing Purkinje General Instrument Co., Ltd, China).

    Antioxidant enzyme activity was calculated with the following equations:

    SODactivity(U/gFW)=(AckAe)×Ve0.5×Ack×W×Ve1 (11)
    PODactivity(U/gmin)=D470×Ve0.01×W×Ve2×T (12)
    PPOactivity(U/gmin)=D420×Ve(0.01×W×Ve3×T) (13)
    CATactivity(U/gmin)=D240×Ve0.1×W×Ve4×T (14)

    Ack and Ae are the absorbance of the control and samples at 560 nm, respectively; D470 is the change of absorbance measured at 470 nm; D420 is the change of absorbance measured at 420 nm; D240 is the change of absorbance measured at 240 nm; Ve is the total volume of E, mL; Ve is the total volume of E, mL; Ve1 is the volume of E1, mL; Ve2 is the volume of E2, mL; Ve3 is the volume of E3, mL; Ve4 is the volume of E4; T is the reaction time, min; W is the weight of fresh leaf, g.

    One-way analysis of variance was conducted to evaluate the effect of shading treatment on growth and physiological characteristics, followed by Tukey's Highly Significant Differences (HSD), and the standard error of differences between means was calculated with p set to 0.05. All statistical analyses were performed using SPSS Statistics 18 version 16.0 for Windows (SPSS Inc., Chicago, IL, USA). Correlation analysis was performed to identify the relationship between the growth index of seedlings and physiological index using the software package Origin 9.1 (Northampton, MA01060, USA). Network analysis was performed using Gephi (version 0.9.2, WebAtlas, France) to analyze the correlation between various physiological index.

    In the treatment without shade net (SH0, as a control), the survival rate of seedlings was 84.44%, and seedling growth was significantly inhibited. Specifically, seedlings had the lowest height and growth biomass. Leaves were seriously burnt, but RSR was the highest (Fig. 1, Table 1). Under shading conditions, the survival rate of seedlings increased significantly, but there was no difference under SH1 and under SH2. Seedling growth and biomass significantly increased with increasing layers of shade net. In particular, seedlings under SH2 showed favorable growth with the largest leaf area, highest plant height, and highest plant weight, but RSR decreased with increasing layers of shade net (Table 1, Fig. 1).

    Figure 1.  Growth of Cyclocarya paliurus seedlings under different shading levels. (a), (c) Seedling growth in plastic greenhouse under three shading levels; (b), (d) seedling growth in plastic greenhouse under SH1 and SH0, respectively; SH0 indicates seedling growth with no shade net (about 60%−70% of full sunlight); SH1 indicates seedling growth with one layer of shade net (about 30%−40% of full sunlight); SH2 indicates seedling growth with two layers of shade net (about 10%−20% of full sunlight). Seedlings were treated in plastic greenhouses with different shading levels for 100 d (from June to September), and plastic sheeting was not fitted around the plastic greenhouse and the shade net was set at 1.8 m height from the ground.
    Table 1.  Variations in seedling growth of Cyclocarya paliurus with different shading treatments
    Treatment Survival rate
    (%)
    The fourth leaf area
    (cm2)
    DH
    (mm)
    The growth rate of stem
    (cm/d)
    DM:FM
    (g/plant)
    RSR
    SH0 84.44 ± 3.85b 3.46 ± 0.42c 1.48 ± 0.24c 0.02 ± 0c 4.04 ± 0.72b 0.70 ± 0.19a
    SH1 95.56 ± 3.85a 9.19 ± 1.045b 2.76 ± 0.42b 0.08 ± 0.02b 6.34 ± 2.79b 0.58 ± 0.13ab
    SH2 100 ± 0a 33.19 ± 3.09a 5.29 ± 0.76a 0.31 ± 0.03a 18.84 ± 3.72a 0.41 ± 0.08b
    SH0 means seedling growth with no shade net; SH1 means seedling grwoth with one layer of shade net; SH2 means seedling growth with two layers of shade net. Different letters (a,b,--) indicate significant differences between shading treatments by Tukey's Highly Significant Differences at p set to 0.05. DH represents ground diameter; DM:FM represents dry mass: fresh mass; RSR represents the ratio of underground weight to aboveground weight.
     | Show Table
    DownLoad: CSV

    WC of seedlings under SH1 and SH2 was slightly higher than that under SH0, but no differences exist between SH0, SH1, and SH2 (Fig. 2a). However, WC varied in the three parts (root, shoot, and leaf) under different shading levels. Specifically, WC in the root increased, but WC in the shoot decreased with increasing layers of shade net, and no difference was observed between SH0, SH1, and SH2. WC in the leaf under SH1 and SH2 significantly increased by 12.54% and 12.78%, respectively, in comparison with that under SH0 (Fig. 2a).

    Figure 2.  (a) Water content and (b) total soluble sugar content in Cyclocarya paliurus seedlings under different shading levels. SH0 indicates seedling growth with no shade net; SH1 indicates seedling growth with one layer of shade net; SH2 indicates seedling growth with two layers of shade net. * means significant differences (p < 0.05 according to Tukey's HSD) between SH0, SH1 and SH2.

    Leaves under SH2 showed the highest TSS content, TSS content under SH0 and SH1 decreased by 13.93% and 14.67%, respectively, but there was no difference between three shading levels (Fig. 2b).

    Chlorophyll (Chl) content of leaves under SH2 was 2.52 times and 1.67 times higher than that under SH0 and SH1, respectively (Fig. 3a). Chl a content was much higher than Chl b content and carotenoid content, but both of them showed a similar trend (Fig. 3b, c, d).

    Figure 3.  Chlorophyll content and carotenoid content in Cyclocarya paliurus leaves under different shading levels. (a) Chlorophyll content; (b) chlorophyll a content; (c) chlorophyll b content; and (d) carotenoid content. SH0 indicates seedling growth with no shade net; SH1 indicates seedling growth with one layer of shade net; SH2 indicates seedling growth with two layers of shade net. * means significant differences (p < 0.05 according to Tukey's HSD) between SH0, SH1 and SH2.

    Under the three shading levels, MDA content in leaf under SH2 was the lowest, and its content under SH0 and SH1 significantly increased by 59.15% and 60.25%, respectively (Fig. 4). But there was no difference between under SH0 and SH1.

    Figure 4.  MDA content in Cyclocarya paliurus leaves under different shading levels. SH0 indicates seedling growth with no shade net; SH1 indicates seedling growth with one layer of shade net; SH2 indicates seedling growth with two layers of shade net. * means significant differences (p < 0.05 according to Tukey's HSD) between SH0, SH1 and SH2.

    Four kinds of mineral (K, Ca, Mg, and Na) were detected in seedlings, and their contents were in the order of Ca>K>Mg>Na (Fig. 5a). The translation factor of the four minerals decreased with the increasing layers of shade net, and there was a significant difference between under SH0, SH1, and SH2 (Fig. 5b).

    Figure 5.  Mineral contents in Cyclocarya paliurus seedling under different shading levels. (a) mineral contents in seedlings; (b) translocation factor; (c) K content; (d) Ca content; (e) Mg content; (f) Na content. SH0 indicates seedling growth with no shade net; SH1 indicates seedling growth with one layer of shade net; SH2 indicates seedling growth with two layers of shade net. * means significant differences (p < 0.05 according to Tukey's HSD) between SH0, SH1 and SH2.

    K content of seedlings was significantly decreased with the increasing layers of shade net (Fig. 5a), K content under SH0 and SH1 was rich in the leaf (Fig. 5c). K content in the root under SH1 was higher than that under SH0 and SH2, but no change was observed between under SH0, SH1 and SH2. K content in the shoot and leaf decreased with increasing layers of shade net (Fig. 5c).

    Ca content of seedlings under SH1 significantly decreased, but increased significantly under SH2 in comparison with that under SH0 (Fig. 5a). Ca content in the leaf was higher than that in the root and shoot (Fig. 5d). Ca content in the root and shoot had a similar trend with that in seedlings. Leaves showed a decreasing Ca content with increasing layers of shade net.

    Seedling under SH1 had the highest Mg content (Fig. 5a), Mg content was also rich in leaf. Mg content in the root slightly increased, but Mg content in the shoot and leaf significantly decreased with increasing layers of shade net (Fig. 5e).

    Na content of seedlings reached significant difference under SH2 (Fig. 5a), Na content in the root was higher than that in the leaf and in shoot. Na content in the root and shoot significantly increased, but decreased in the leaf with increasing layers of shade net (Fig. 5f).

    Four kinds of antioxidant enzymes (SOD, PPO, POD, and CAT) were detected in the leaf and their activities were in the following order: SOD>PPO>POD>CAT (Fig. 6). However, their activities varied with different shading levels. Leaf under SH0 had the highest SOD activity, and SOD activity decreased significantly with increasing layers of shade net. The change of POD and CAT activities were similar to that of SOD activity. However, PPO activity increased significantly with increasing layers of shade net, and its activity under SH2 was 25.67 and 23.03 times higher than that under SH0 and SH1, respectively.

    Figure 6.  Variations in antioxidant enzyme activities in Cyclocarya paliurus leaves under different shading levels. (a) CAT activity; (b) PPO activity; (c) POD activity; and (d) SOD activity. SH0 indicates seedling growth with no shade net; SH1 indicates seedling growth with one layer of shade net; SH2 indicates seedling growth with two layers of shade net. * means significant differences (p < 0.05 according to Tukey's HSD) between SH0, SH1 and SH2.

    The growth index of seedlings had positive or negative relationships with various physiological indices under three shading levels (Fig. 7a). And various relationships exist between them. For example, seedling height and leaf area had a negative relation with growth biomass (DM:FM, RSR); PPO activity had a significant positive relation with TSS and Chl content. Antioxidant enzyme activity (SOD, POD, CAT) had a positive relation with mineral content (K, Ca, Mg, Na); but physiological index had a negative relation with antioxidant enzyme activity and mineral content (Fig. 7b).

    Figure 7.  (a) Correlation analysis and (b) network analysis between different studied factors. LA, DH, RH, DF, RSR indicates leaf area, ground diameter, the growth rate of shoot, the ratio of dry mass to fresh mass, the ratio of underground weight to aboveground weight, respectively; WC, TSS, Chl, PPO indicate water content, total soluble sugar content, Chlorophyll content, polyphenol oxidase activity;* represents significant differences (p < 0.05 according to Tukey's HSD) between SH0, SH1, and SH2. antioxidant enzyme; mineral content; physiological index; MDA content. Blue line means a positive relation between each other; red line means a negative relation between each other.

    Greenhouse cultivation is an effective method of cultivating high-quality seedlings. However, higher temperatures or stronger light intensity impairs the successful growth of many plants under greenhouse cultivation, especially plastic greenhouse cultivation. The morphological characteristics of many seedlings were the increase of leaf thickness, the reduction of leaf area and plant height, and the serious sunburn of leaves[2,3,14]. In accordance with previous morphological changes, our findings also showed that plastic greenhouse cultivation with no shade net (SH0) inhibited plant height and leaf area and caused serious leaf sunburn (Table 1, Fig. 1). Our finding of MDA content under SH0 further inferred that seedling growth and metabolism were seriously inhibited or damaged even at the cellular level (Figs 26), resulting in higher activity of antioxidant enzymes and higher content of minerals to alleviate this damage according to network analysis (Fig. 7). This result suggested that plastic greenhouse cultivation without shading condition was unsuitable to cultivate seedlings of C. paliurus.

    Shading practices effectively alleviate or prevent damages in plants under greenhouse cultivation[2,58,14]. Our findings further confirmed that suitable shading conditions favor the growth of seedlings. For one reason, growth index and biomass serve as a direct indicator of plant response to environmental conditions[2830]. Our findings showed that both shading levels (SH1 and SH2) benefited seedling growth, especially two layers of shade net (SH2), where growth index and biomass of seedlings were much better than that under SH0 (Fig. 1, Table 1). Similar findings were reported by Lee et al.[31], who showed that 30% shade promoted plant growth. However, Liu et al.[22] reported that full light intensity conditions resulted in the highest growth and total biomass production of one-year-old C. paliurus plants in an open field, probably because cultivation conditions were different from our study. For another reason, chlorophyll takes part in light utilization and promotes plant growth by improving light interceptions and absorptions and accelerating photosynthetic metabolism[3234]. In corroboration with previous findings[2,35], our finding further revealed that the chlorophyll content in leaves had a significant positive relationship with plant height and TSS content or PPO activity (Fig. 7). However, the significant decrease of chlorophyll content under SH0 and SH1 may be caused by the serious damage to the cellular membrane, leading to the inhibition of chlorophyll synthesis, the promotion of chlorophyll degradation, or the prevention of the conversion of Chl a to Chl b to a large extent. This was further illustrated by the negative relation between Chl content and MDA content (Fig. 7b). This was in corroboration with the results for rose[8], Azolla microphylla[11] and lettuce[34, 36]. Finally, the change of MDA content under different shading levels further inferred that shading condition was suitable for seedling cultivation by alleviating the damage of adverse conditions. This was shown from the network analysis between MDA and other physiological index (Fig. 7b).

    Sugar is a substrate that participates in many biosynthetic processes and energy production in plants, including the regulation of lateral root emergence and development and biosynthesis and degradation of auxin via an interaction with phytochrome[37]. In addition, sugar is also a substrate for sensing and signaling plant systems under biotic or abiotic stress[38]. Sugar can protect olive trees during drought conditions[39,40] and improve C. paliurus seedlings during drought conditions[25]. In the present study, total sugar content in the leaves under three shading levels had no difference (Fig. 2), but TSS content positively correlated with morphological traits (plant height, leaf area) and other physiological components (Chl content, PPO activity), and negatively correlated with RSR (Fig. 7), suggesting that sugar acts as a major integrator for osmotic response to abiotic stress and influences root development by promoting auxin degradation[25,33,37,38,41].

    Minerals are indispensable nutrients for the growth and development of plants, and as non-enzymatic antioxidant defense systems, minerals respond to adverse conditions by regulating many physiological activities[42,43]. In our study, the content of four minerals varied in the three parts and had different relationships with physiological index (Figs 5 & 7b). K is essentially for stomatal function, cell expansion, osmoregulation, and cellular or whole-plant homeostasis[4446]. Mg is the central atom of chlorophyll and enzyme activator for photosynthesis. High K and Mg contents were observed in the leaf under SH0 and SH1, and they had a positive relationship with antioxidant enzymes, but had a negative relationship with Chl content, TSS content or WC (Figs 5 & 7b). This result can be attributed to the transfer of high content of K and Mg from roots to leaves to protect seedlings from high solar radiation. Ca is important for preserving membrane integrity, signaling osmoregulation, and influencing K/Na selectivity[47,48]. This was further inferred from the change of Ca content under different shading levels (Fig. 5). Na improves plant water balance and water use efficiency by modifying stomatal control and contributes to the maintenance of cell turgor and expansion[49]. A decreasing tendency of Na transfer ability from root to stem or leaf was observed with increasing shading levels in our study (Fig. 5).

    Enzymatic defense mechanisms (superoxide dismutase (SOD), catalase (CAT), and peroxidase (POD)) have been developed to reduce the negative effects of reactive oxygen species (ROS), which affect plants at the cellular level under stress[50]. In accordance with Feng et al.[25], who reported that C. paliurus seedlings had increased antioxidant enzyme activities under different soil water capacities, higher activities of SOD, CAT and POD were observed in leaf under SH0 in comparison with that under SH2, highlighting the effectiveness of antioxidant enzyme systems in protecting cellular apparatuses under unsuitable conditions. Similar result was also reported in rose[8]. Moreover, SOD activity was much higher than CAT activity and POD activity, and they had positive correlation with MDA and mineral content (Figs 6 & 7b). This indicates that SOD is the main antioxidant enzyme in the enzymatic defense mechanism, and three antioxidant enzymes (SOD, CAT, and POD) participated in alleviating excessive ROS at the cellular level via a cascade mechanism[50] and coordinated non-enzymatic mechanisms (K, Ca, Mg, and Na) to maintain the healthy growth of plants. However, our findings showed that PPO activity under SH0 and SH1 had a lower level compared to that under SH2 and positively correlated with growth index except for RSR (Figs 4 & 7b), inferring that PPO may have another role in seedling growth; however, further experimental investigation is needed to confirm this inference.

    Cyclocarya paliurus seedlings were cultivated in a plastic greenhouse under three shading levels. The results showed that (1) shading conditions with two layers of shade net (SH2) improved the survival rate and growth of Cyclocarya paliurus seedlings, and seedlings significantly increased plant height, leaf area, and biomass accumulation; (2) in comparison with that under SH0 and SH1, chlorophyll content increased and MDA content reduced significantly under SH2, indicating that suitable shading conditions was beneficial to seedling growth and normal metabolism; and (3) the physiological responses of seedlings varied with shading levels and showed various correlation with seedling growth, indicating that mineral and antioxidant enzymes could coordinate to alleviate or protect seedling from damage under SH0 and SH1, and SOD was the main enzymatic mechanism.

    This work was supported by Natural Science Foundation of Fujian province of China (No. 2022J011107), Talent project of Quanzhou city of China (2021C043R) and The college students innovations special project of Fujian province (S202110399065; S202210399049).

  • The authors declare that they have no conflict of interest.

  • [1]

    Bae JH, Park JH, Im SS, Song DK. 2014. Coffee and health. Integrative Medicine Research 3(4):189−91

    doi: 10.1016/j.imr.2014.08.002

    CrossRef   Google Scholar

    [2]

    Krishnan S. 2017. Sustainable Coffee Production. In Oxford Research Encyclopedia of Environmental Science. pp. 1−29. https://doi.org/10.1093/acrefore/9780199389414.013.224

    [3]

    Department of Agriculture - High Value Crops Development Program. 2022. Philippine Coffee Industry Roadmap 2021-2025. Department of Agriculture - Bureau of Agricultural Research through UPLB Foundation, Inc. in collaboration with the Philippine Council for Agriculture and Fisheries. 150 pp. www.pcaf.da.gov.ph/wp-content/uploads/2022/06/Philippine-Coffee-Industry-Roadmap-2021-2025.pdf.

    [4]

    Hasibuan AM, Ferry Y, Wulandari S. 2022. Factors affecting farmers' decision to use organic fertilizers on Robusta Coffee Plantation: A case study in Tanggamus, Lampung. IOP Conference Series: Earth and Environmental Science 974(1):012105

    doi: 10.1088/1755-1315/974/1/012105

    CrossRef   Google Scholar

    [5]

    Bhardwaj D, Ansari MW, Sahoo RK, Tuteja N. 2014. Biofertilizers function as key player in sustainable agriculture by improving soil fertility, plant tolerance and crop productivity. Microbial Cell Factories 13:66

    doi: 10.1186/1475-2859-13-66

    CrossRef   Google Scholar

    [6]

    Beneduzi A, Ambrosini A, Passaglia LMP. 2012. Plant growth-promoting rhizobacteria (PGPR): Their potential as antagonists and biocontrol agents. Genetics and Molecular Biology 35:1044−51

    doi: 10.1590/s1415-47572012000600020

    CrossRef   Google Scholar

    [7]

    Goswami D, Parmar S, Vaghela H, Dhandhukia P, Thakker JN. 2015. Describing Paenibacillus mucilaginosus strain N3 as an efficient plant growth promoting rhizobacteria (PGPR). Cogent Food & Agriculture 1(1):1000714

    doi: 10.1080/23311932.2014.1000714

    CrossRef   Google Scholar

    [8]

    Gupta K, Dubey NK, Singh SP, Kheni JK, Gupta S, et al. 2021. Plant Growth-Promoting Rhizobacteria (PGPR): Current and Future Prospects for Crop Improvement. Environmental and Microbial Biotechnology. In Current Trends in Microbial Biotechnology for Sustainable Agriculture, eds. Yadav AN, Singh J, Singh C, Yadav N. Singapore: Springer. pp. 203–26. https://doi.org/10.1007/978-981-15-6949-4_9

    [9]

    Majeed A, Abbasi MK, Hameed S, Imran A, Rahim N. 2015. Isolation and characterization of plant growth-promoting rhizobacteria from wheat rhizosphere and their effect on plant growth promotion. Frontiers in Microbiology 6:198

    doi: 10.3389/fmicb.2015.00198

    CrossRef   Google Scholar

    [10]

    Cortes AD, Opulencia RB, Aggangan NS. 2020. Characterization of plant growth promoting diazotrophic bacteria isolated from Cacao (Theobroma cacao L.) rhizosphere treated with bamboo biochar and arbuscular mycorrhizal fungi. Philippine Journal of Science 149(4):1063−70

    doi: 10.56899/149.04.05

    CrossRef   Google Scholar

    [11]

    Philippot L, Raaijmakers JM, Lemanceau P, Van der Putten WH. 2013. Going back to the roots: the microbial ecology of the rhizosphere. Nature Reviews Microbiology 11(11):789−99

    doi: 10.1038/nrmicro3109

    CrossRef   Google Scholar

    [12]

    Hassen AI, Khambani LS, Swanevelder ZH, Mtsweni NP, Bopape FL, et. al. 2021. Elucidating key plant growth-promoting (PGPR) traits in Burkholderia sp. Nafp2/4-1b (=SARCC-3049) using gnotobiotic assays and whole-genome-sequence analysis. Letters in Applied Microbiology 73(5):658−71

    doi: 10.1111/lam.13556

    CrossRef   Google Scholar

    [13]

    Fulthorpe R, Martin AR, Isaac ME. 2020. Root endophytes of coffee (Coffea arabica): variation across climatic gradients and relationships with functional traits. Phytobiomes Journal 4:27−39

    doi: 10.1094/PBIOMES-04-19-0021-R

    CrossRef   Google Scholar

    [14]

    Backer R, Rokem JS, Ilangumaran G, Lamont J, Praslickova D, et al. 2018. Plant growth-promoting rhizobacteria: context, mechanisms of action, and roadmap to commercialization of biostimulants for sustainable agriculture. Frontiers in Plant Science 9:1473

    doi: 10.3389/fpls.2018.01473

    CrossRef   Google Scholar

    [15]

    Ahemad M, Kibret M. 2013. Recent trends in microbial biosorption of heavy metals: a review. Biochemistry & Molecular Biology 1:19−26

    doi: 10.12966/bmb.06.02.2013

    CrossRef   Google Scholar

    [16]

    Walpola BC, Yoon MH. 2012. Prospectus of phosphate solubilizing microorganisms and phosphorus availability in agricultural soils: A review. African Journal of Microbiology Research 6(37):6600−5

    doi: 10.5897/AJMR12.889

    CrossRef   Google Scholar

    [17]

    Liu M, Liu X, Cheng BS, Ma XL, Lyu XT, et. al. 2016. Selection and evaluation of phosphate-solubilizing bacteria from grapevine rhizospheres for use as biofertilizers. Spanish Journal of Agricultural Research 14(4):e1106

    doi: 10.5424/sjar/2016144-9714

    CrossRef   Google Scholar

    [18]

    Kunwar VS, Chimouriya S, Lamichhane J, Gauchan DP. 2018. Isolation and characterization of phosphate solubilizing bacteria from rhizosphere of coffee plant and evaluating their effects on growth and development of coffee seedlings. BioTechnology: An Indian Journal 14(5):173

    Google Scholar

    [19]

    Sembiring M, Sabrina T, Mukhlis M. 2020. Phosphate solubilizing microbes and coffee skin compost to increase Robusta coffee plant growth in Andisol of Mount Sinabung area. Bulgarian Journal of Agricultural Science 26(4):766−71

    Google Scholar

    [20]

    Kundan R, Pant G, Jadon N, Agrawal PK. 2015. Plant growth promoting rhizobacteria: Mechanism and current prospective. Journal of Fertilizers & Pesticides 6:155

    doi: 10.4172/2471-2728.1000155

    CrossRef   Google Scholar

    [21]

    Abawari R, Tuji F, Yadete D. 2021. Multi traits of phosphate solublizing bacterial and fungal isolates and evaluation of their potential as biofertilizer agent for coffee production. International Journal of Applied Agricultural Sciences 7(1):1−15

    doi: 10.11648/j.ijaas.20210701.11

    CrossRef   Google Scholar

    [22]

    Mardanova AM, Fanisovna Hadieva G, Tafkilevich Lutfullin M, Valer'evna Khilyas I, Farvazovna Minnullina L, et. al. 2017. Bacillus subtilis strains with antifungal activity against the phytophatogenic fungi. Agricultural Sciences 8(1):1−20

    doi: 10.4236/as.2017.81001

    CrossRef   Google Scholar

    [23]

    Tsegaye Z, Gizaw B, Tefera G, Feleke A, Chaniyalew S, et. al. 2019. Isolation and biochemical characterization of Plant Growth Promoting (PGP) bacteria colonizing the rhizosphere of Tef crop during the seedling stage. Journal of Plant Science and Phytopathology 3(1):13−27

    doi: 10.29328/journal.jpsp.1001027

    CrossRef   Google Scholar

    [24]

    Kenneth OC, Nwadibe EC, Kalu AU, Unah UV. 2019. Plant Growth Promoting Rhizobacteria (PGPR): A Novel Agent for Sustainable Food Production. American Journal of Agricultural and Biological Sciences 14(1):35−54

    doi: 10.3844/ajabssp.2019.35.54

    CrossRef   Google Scholar

    [25]

    Subiramani S, Ramalingam S, Muthu T, Nile SH, Venkidasamy B. 2020. Development of abiotic stress tolerance in crops by Plant Growth-Promoting Rhizobacteria (PGPR). In Phyto-microbiome in Stress Regulation. Environmental and Microbial Biotechnology eds. Kumar M, Kumar V, Prasad R. Springer, Singapore. pp. 125−45. https://doi.org/10.1007/978-981-15-2576-6_8

    [26]

    Vocciante M, Grifoni M, Fusini D, Petruzzelli G, Franchi E. 2022. The role of plant growth-promoting rhizobacteria (PGPR) in mitigating plant's environmental stresses. Applied Sciences 12(3):1231

    doi: 10.3390/app12031231

    CrossRef   Google Scholar

    [27]

    Ma Y, Rajkumar M, Zhang C, Freitas H. 2016. Beneficial role of bacterial endophytes in heavy metal phytoremediation. Journal of Environmental Management 174:14−25

    doi: 10.1016/j.jenvman.2016.02.047

    CrossRef   Google Scholar

    [28]

    Kumar A, Kumari M, Swarupa P, Shireen S. 2019. Characterization of pH dependent growth response of agriculturally important microbes for development of plant growth promoting bacterial consortium. Journal of Pure and Applied Microbiology 13(2):1053−61

    doi: 10.22207/JPAM.13.2.43

    CrossRef   Google Scholar

    [29]

    Muleta D, Assefa F, Börjesson E, Granhall U. 2013. Phosphate-solubilising rhizobacteria associated with Coffea arabica L. in natural coffee forests of southwestern Ethiopia. Journal of the Saudi Society of Agricultural Sciences 12(1):73−84

    doi: 10.1016/j.jssas.2012.07.002

    CrossRef   Google Scholar

    [30]

    Pino AFS, Espinosa ZYD, Cabrera EVR. 2023. Characterization of the rhizosphere bacterial microbiome and coffee bean fermentation in the Castillo-Tambo and Bourbon varieties in the Popayán-Colombia Plateau. BMC Plant Biology 23(1):217

    doi: 10.1186/s12870-023-04182-2

    CrossRef   Google Scholar

    [31]

    Walterson AM, Stavrinides J. 2015. Pantoea: insights into a highly versatile and diverse genus within the Enterobacteriaceae. FEMS Microbiology Reviews 39(6):968−84

    doi: 10.1093/femsre/fuv027

    CrossRef   Google Scholar

    [32]

    Urgiles-Gómez N, Avila-Salem ME, Loján P, Encalada M, Hurtado L, et. al. 2021. Plant growth-promoting microorganisms in coffee production: From isolation to field application. Agronomy 11(8):1531

    doi: 10.3390/agronomy11081531

    CrossRef   Google Scholar

    [33]

    Knežević MM, Stajković-Srbinović OS, Assel M, Milić MD, Mihajlovski KR, et al. 2021. The ability of a new strain of Bacillus pseudomycoides to improve the germination of alfalfa seeds in the presence of fungal infection or chromium. Rhizosphere 18:100353

    doi: 10.1016/j.rhisph.2021.100353

    CrossRef   Google Scholar

    [34]

    Chukwuma OB, Rafatullah M, Kapoor RT, Tajarudin HA, Ismail N, et. al. 2023. Isolation and characterization of lignocellulolytic bacteria from municipal solid waste landfill for identification of potential hydrolytic enzyme. Fermentation 9(3):298

    doi: 10.3390/fermentation9030298

    CrossRef   Google Scholar

    [35]

    Sherpa MT, Sharma L, Bag N, Das S. 2021. Isolation, characterization, and evaluation of native rhizobacterial consortia developed from the rhizosphere of rice grown in Organic State Sikkim, India, and their effect on plant growth. Frontiers in Microbiology 12:713660

    doi: 10.3389/fmicb.2021.713660

    CrossRef   Google Scholar

  • Cite this article

    Navarro GVD, Quirong DD, Maghanoy GA, Cortes AD. 2023. Characterization and identification of rhizobacteria associated with Liberica and Robusta coffee rhizosphere. Technology in Horticulture 3:24 doi: 10.48130/TIH-2023-0024
    Navarro GVD, Quirong DD, Maghanoy GA, Cortes AD. 2023. Characterization and identification of rhizobacteria associated with Liberica and Robusta coffee rhizosphere. Technology in Horticulture 3:24 doi: 10.48130/TIH-2023-0024

Figures(2)  /  Tables(3)

Article Metrics

Article views(3139) PDF downloads(522)

ARTICLE   Open Access    

Characterization and identification of rhizobacteria associated with Liberica and Robusta coffee rhizosphere

Technology in Horticulture  3 Article number: 24  (2023)  |  Cite this article

Abstract: Coffee is a viable agricultural commodity that makes a positive impact to the Philippine economy. However, with an increasing trend in domestic consumption, the local coffee production has declined. Chemical fertilization has been considered by many farmers to improve coffee production and yield but it causes a serious threat to public health and the environment. Biofertilizer using rhizobacteria has beneficial effects to improve the growth and yield of many crops, which is cost effective and safer than synthetic fertilizers. This study characterized the indigenous and beneficial rhizobacteria obtained from the Liberica and Robusta coffee rhizosphere, in terms of phosphate solubilization, biocontrol activities, and tolerance to abiotic stresses. Six rhizobacterial isolates were molecularly identified and belonged to genera Bacillus, Burkholderia, and Pantoea. These rhizobacteria solubilized inorganic phosphate with solubilization index ranging from 2.5 to 3.5 mm. For biocontrol activities, Bacillus sanguinis showed activity in terms of HCN and multiple hydrolytic enzymes production. Also, Burkholderia sp. demonstrated amylase, protease, and pectinase activities. Moreover, all isolates were found to be relatively tolerant to a wide range of pH and concentrations of salt and heavy metals. The performance of these rhizobacterial isolates in terms of phosphate solubilization, biocontrol activities, and tolerance to stresses is promising and shown to have potential in coffee cultivation in the Philippines.

    • Coffee is among the most widely consumed pharmacologically active beverage world wide and its consumption has become part of everyday life[1]. Many coffee-producing countries in the coffee belt are benefiting from this agricultural commodity because their locations support the ideal growth and production of coffee, serving as a major source of income[2]. In particular, the Philippines is an ideal place to grow quality coffee of different types. However, despite the 2.1% increase in consumption, the local coffee production has been decreasing by 3.5% per year for the last 10 years[3].

      Coffee cultivation is usually associated with agri-chemical inputs, such as fertilizers and pesticides, to increase soil fertility and crop growth. However, overuse of chemical fertilizers causes soil nutrient imbalance and environmental contamination[4]. Soil acidification due to unabsorbed chemical fertilizers may lead to plant toxicity, resulting in growth deterioration and low yield of crops. In addition, chemical fertilizers are expensive and may be a burden for smallholder coffee farmers. Sustainable strategies in agricultural farming are being introduced to combat such threats, such as the exploitation of beneficial microbes as a biofertilizer[5]. Rhizospheric microorganisms are being characterized to explore their potential role in food safety and sustainable crop production.

      Rhizobacteria with plant growth promoting attributes and biocontrol activities are ubiquitous and highly abundant in the plant rhizosphere, which primarily colonize the roots and promote plant growth. They can act as biocontrol agent and their effects can occur via local antagonism to soil-borne pathogens or by induction of plant systemic resistance against pathogens[6]. Plant growth promotion can be attributed to their ability to produce phytohormones (e.g., indole acetic acid, cytokinins, and gibberellins), fix atmospheric nitrogen, solubilize phosphate, and mineralize organic substances[79].

      This study focused on the isolation, characterization, and identification of rhizobacteria with phosphate solubilization ability from the coffee rhizosphere and we evaluated their biocontrol activities and tolerance to different abiotic stresses. Their potential use as a biofertilizer will provide new insights in coffee cultivation and production in the Philippines.

    • Soil samples were randomly collected at 35-cm soil depth along the roots of Coffea liberica (Liberica) and Coffea canephora (Robusta) trees at the Coffee Genebank of National Coffee Research, Development and Extension Center (NCRDEC) in Cavite State University. A total of three 100 g of rhizospheric soil sub-samples from each coffee tree were pooled to make one composite sample per tree. A portion of the collected samples were immediately processed for microbiological analysis and the rest were air-dried at room temperature for soil physicochemical analysis.

    • Dried soil samples were sent to the Agricultural System Institute, University of the Philippines Los Baños, for the analysis of pH, moisture content, and organic matter content. The pH of the soil was found to be at 5.2 and 5.7, moisture content was 41.6% and 42.3%, and organic matter content was about 2.88% and 3.28% in the Liberica and Robusta coffee rhizosphere, respectively.

    • We selectively isolated rhizobacteria with phosphate solubilization activity on tricalcium phosphate medium. Briefly, a total of 10 g of fresh composite soil samples were mixed in 90 mL sterile distilled water and was serially diluted to the 106 dilution. One mL from each dilution was spread-plated on Pikovskaya's agar medium, containing 0.5 g·L−1 yeast extract, 10.0 g·L−1 dextrose, 5.0 g·L−1 tricalcium phosphate, 0.5 g·L−1 ammonium sulphate, 0.2 g·L−1 potassium chloride, 0.1 g·L−1 magnesium sulphate, 0.0001 g·L−1 manganese sulphate, 0.0001 g·L−1 ferrous sulphate, and 15.0 g·L−1 agar. Agar plates were incubated at 30 °C for 5 d and the clear zone around colonies were observed as indication of phosphate solubilization. The colonies with a clear zone were purified on Nutrient Agar (NA) plates. The colony characteristics of the isolates, such as elevation, margin, shape, and color were recorded. In addition, the purity of the cultures was verified using Gram stain reaction, where bacterial shapes, arrangement, and Gram reaction were also recorded. Pure cultures were then stored at 4 °C for further analysis.

      The phosphate solubilization index of rhizobacteria was measured, following the spot-plating method[10]. Initial numbers of cells were adjusted to 0.5 McFarland standard that is approximately 1.5 × 108 CFU·mL−1, this was used in all experimental assays of the study. A total of 4 μL 24-h bacterial suspension grown in Nutrient Broth (NB) was spot inoculated on Pikovskaya's agar plates containing tricalcium phosphate and incubated at 30 °C for 5 d. The colony and clear zone diameters (mm) were measured, then the solubilization index was calculated by dividing the sum of colony diameter and clear zone over the colony diameter.

    • All isolates were grown in a 5-mL glycine-supplemented NB medium. Initially, a Whatman filter paper was saturated with picric acid solution, containing 2% sodium carbonate and 0.5% picric acid, and was immediately placed at the inner top of the screw cap tubes. Tubes were sealed with parafilm and incubated with shaking for 4 d at room temperature. A color change from yellow to reddish brown in the filter paper indicate a positive HCN production.

    • The bacterial inoculum was spot inoculated, in triplicates, on starch agar medium (containing 0.5 g·L−1 peptone, 3 g·L−1 beef extract, 0.5 g·L−1 NaCl, 1% starch powder, and 12 g·L−1 agar) and incubated for 48 h at 30 °C. The plates were then flooded with iodine solution for 1 min and then drained off. The appearance of a clear zone around colonies indicate a positive for amylase activity.

    • The protease activity of the isolates was screened using milk agar medium (containing 5 g·L−1 peptone, 3 g·L−1 yeast extract, 100 mL·L−1 UHT non-fat milk and 12 g·L−1 agar). A total of 4 μL fresh inoculum was spot inoculated on the medium, in triplicate, and incubated for 48 h at 30 °C. A clear zone around the colonies indicates a positive protease activity.

    • Pectinase activity of isolates was screened using the pectin agar medium (containing 0.5 g·L−1 peptone, 0.3 g·L−1 beef extract, 0.5 g·L−1 NaCl, 4 g·L−1 citrus pectin, and 12 g·L−1 agar). Spot inoculation was similarly used but incubation was extended to 96 h. A 50 mM potassium iodide-iodine solution was then flooded on the surface of the agar plates. A positive result was indicated by the appearance of a clear zone around colonies.

    • The ability of the isolates to tolerate abiotic stresses, such as acidity, salinity, and heavy metal contents (i.e., lead, copper, and manganese) was screened. Briefly, the isolates were subjected to varying pH level, namely 4, 5, 7, 9 and 11. Uniform initial number of cells were grown in NB for 24 h with shaking and bacterial suspension and spot inoculated on NA plates. Also, NA medium was supplemented with varying sodium chloride concentrations (i.e., 1%, 3%, 5%, 7%, 9%, and 11%) and was used to screen for salt tolerance of the isolates. Lastly, the ability of the isolates to tolerate heavy metals such as lead, copper, and manganese was screened using NA medium supplemented with varying concentrations (i.e., 100, 150, 200, 400, 800, and 1,600 ppm) of lead acetate (Pb(C2H3O2)2), copper sulphate (CuSO4), and manganese sulphate (MnSO4), respectively. Growth was observed after 48 h at room temperature, indicating resistance/tolerance to varying levels of pH, salt, and heavy metals.

    • A total of six isolates were subjected to DNA extraction using Vivantis GF-1 Bacterial DNA Extraction Kit, following the manufacturer's protocol. The quality of the isolated genomic DNA was verified in 0.8% agarose gel (dissolved in 0.5X TAE buffer), following gel electrophoresis (Mupid One) and gel documentation (Vilber Lourmat). The purity and concentration of DNA were quantified using a NanoDrop 2000c UV-Vis Spectrophotometer (Thermo Scientific™).

      Genomic DNA samples were subjected to polymerase chain reaction (PCR) amplification using universal primers to target the 16S rRNA gene, which are 27F (5'-AGAGTTTGATCCTGGCTCAG-3') and 1492R (5'-GGTTACCTTGTTACGACTT-3'). PCR was performed in a 50-μL reaction containing 1X Taq Master Mix (Vivantis), 2 mM MgCl2, 0.2 μM each of 27F and 1492R primers, and 100 ng of DNA template. A final volume of 50 μL was adjusted with molecular grade water. PCR reactions were performed using MiniAmp™ Plus Thermal Cycler (Applied Biosystems™) with the following conditions: initial denaturation step at 95 °C for 5 min, followed by 30 cycles of 94 °C for 30 s, annealing at 55 °C for 30 s, and 72 °C for 1 min, with a final extension step of 72 °C for 10 min. The PCR products were ran using 1.2% agarose gel (stained with GelRed® nucleic acid dye) under 100 V for 35 min and verified under UV transilluminator.

      The PCR products were sent to Apical Scientific Sdn Bhd (Selangor, Malaysia) for Sanger sequencing. The electropherogram of the sequences was inspected and verified using Sequence Analyzer of MEGAX software. Homologous partial 16S rRNA gene sequences with > 97% similarity were mined in the GenBank database using BLAST algorithm. In MEGAX, ClustalW was used to perform multiple sequence alignment of the unknown and homologous nucleotide sequences. Phylogenetic trees were constructed using Neighbor joining method with 1,000 replicated bootstrap values. The identity of the rhizobacterial isolates was verified based on the clustering of target sequence with the closest annotated sequence (type strain).

    • A total of six rhizobacteria with phosphate solubilization activity (Fig. 1a) were recovered and were subjected to morphological characterization. Two isolates were Gram positive and four were Gram negative, by which all are rod-shaped cells. Colony shapes vary from irregular to circular; elevation is either flat, raised, convex, and crateriform; margin varies from being entire, serrate, undulate, and curled; and the color is yellow, cream, or white (Table 1). The solubilization index (SI) ranges 2.5 to 3.5 mm. Isolate PCL 2.1 and PCR 1.1 obtained the highest phosphate SI of 3.5 mm, whereas isolate PCL 1.3 obtained the lowest (2.5 mm) (Table 2).

      Figure 1. 

      Representatives of rhizobacteria (a) capable of phosphate solubilization as indicated by the clear zone around the colony (arrow) and (b) not capable of phosphate solubilization grown on Pikovskaya's agar medium for 5 d at 30 °C.

      Table 1.  Morphological characteristics of rhizobacteria from the coffee rhizosphere.

      Rhizosphere sourceIsolateColony shapeColony elevationColony marginColony colorGram reactionCell shape
      LibericaPCL 1.2IrregularRaisedEntireWhiteNegativeRods
      LibericaPCL 2.1CircularFlatSerrateWhitePositiveRods
      LibericaPCL 2.3IrregularConvexEntireWhiteNegativeRods
      RobustaPCR 1.1CircularRaisedEntireYellowNegativeRods
      RobustaPCR 1.3IrregularCrateriformUndulateCreamPositiveRods
      RobustaPCR 1.5CircularFlatCurledWhiteNegativeRods

      Table 2.  Six rhizobacterial isolates showing phosphate solubilization, biocontrol activities, and abiotic stress tolerance.

      IsolatePhosphate
      solubilization
      index (mm)
      Biocontrol activityAbiotic stress tolerance
      HCN productionAmylaseProtease PectinaseNaCl (%)pHMnSO4 (ppm)CuSO4 (ppm)Pb(C2H3O2)2 (ppm)
      PCL 1.23.1 ± 0.574 – 111,600400800
      PCL 2.13.5 ± 0.5+74 – 111,600400800
      PCL 2.33.1 ± 0.534 – 111,600400800
      PCR 1.13.5 ± 0.5+34 – 111,600800800
      PCR 1.32.5 ± 0.5+++54 – 11800400800
      PCR 1.52.8 ± 0.5+++34 – 111,600400800
      HCN, hydrogen cyanide; NaCl, sodium chloride; MnSO4, manganese sulphate; CuSO4, copper sulphate; Pb(C2H3O2)2, lead acetate.
    • Among the six rhizobacterial isolates, only PCR 1.3 produced HCN. For the hydrolytic enzyme production, PCL 2.1 produced amylase, PCR 1.1 produced protease, PCR 1.3 produced amylase and protease, and PCR 1.5 produced amylase, protease, and pectinase (Table 2).

    • For pH tolerance, all isolates were tolerant to a wide range of pH (4 to 11). For salt tolerance, PCL 1.2 and PCL 2.1 were tolerant to 7% NaCl, while the lowest tolerance was observed at 3% NaCl. For heavy metal tolerance, all rhizobacteria tolerated 1600 ppm of MnSO4, except PCR 1.3 that only tolerated 800 ppm. PCR 1.1 was able to tolerate up to 800 ppm of CuSO4, while the rest were only tolerant to 400 ppm. Meanwhile, all of them were tolerant to 800 ppm of Pb(C2H3O2)2 (Table 2).

    • Based on the 16S rRNA gene analysis of six promising rhizobacterial isolates, PCL 1.2, PCL 2.1, PCL 2.3, PCR 1.1, PCR 1.3, and PCR 1.5 were identified as Pantoea rwandensis (97.44%), Bacillus pseudomycoides (97.23%), Pantoea sp. (93.62%), Bukholderia cepacia (97.17%), Bacillus sanguinis (99.51%), and Bukholderia sp. (86.66%), respectively (Table 3). Based on the phylogenetic tree analysis, each of the unknown sequence clustered to its closest neighbor (type strain) with bootstrap values (Fig. 2).

      Table 3.  Molecular identity of rhizobacteria isolates from coffee rhizosphere.

      IsolateIdentityClosest neighbor (type strain)Similarity (%)Accession no.
      PCL 1.2Pantoea rwandensisPantoea rwandensis strain LMG 2627597.44NR_118121.1
      PCL 2.1Bacillus pseudomycoidesBacillus pseudomycoides97.23NR_114422.1
      PCL 2.3Pantoea sp.Pantoea agglomerans strain JCM123693.62NR_111998.1
      PCR 1.1Burkholderia cepaciaBurkholderia cepacia ATCC 25416 strain LMG 122297.17NR_114491.1
      PCR 1.3Bacillus sanguinisBacillus sanguinis strain BML-BC00499.51NR_175555.1
      PCR 1.5Burkholderia sp.Burkholderia pseudomallei strain ATCC 2334386.66NR_043553.1

      Figure 2. 

      Neighbor joining tree of rhizobacteria isolates and their closely related species generated using BLAST. Bootstrap values are based on 1000 replications analyzed using MEGAX.

    • Rhizosphere is a nutrient-rich region consisting of a wide variety of microorganisms living in the small area of soil that surrounds and is associated with plant roots. The organisms in the rhizosphere microbiome can have a significant influence on the development, nutrition, and health of plants[11]. Rod-shaped bacteria are commonly found in the rhizosphere and known to have plant growth-promoting attributes, including those species that belong to the genera Bacillus[6], Burkholderia[12], and Pantoea[13], which were isolated and characterized in our study.

      The microbes in the rhizosphere play key roles in nutrient acquisition and assimilation, improved soil texture, secreting, and modulating extracellular molecules such as hormones, secondary metabolites, antibiotics, and various signal compounds, all leading to enhancement of plant growth[14]. Our study initially screened phosphate solubilizing bacteria, because their activity is essential to address plant phosphate requirements of various plants. These bacteria may provide the plants with available phosphorus from sources that would otherwise be scarce due to a wide range of mechanisms. They can also contribute to the natural biogeochemical cycle of nutrients in the rhizosphere[10,15]. Besides, inoculation of phosphate-solubilizing bacteria species has reportedly improved phosphorus absorption and grain production of Triticum aestivum[16] and it significantly facilitated the growth of Vitis vinifera under greenhouse conditions[17]. Besides, the indigenous bacteria with phosphate solubilization activity obtained from coffee rhizosphere were able to stimulate the growth of the Arabica coffee seedlings under nursery conditions[18]. In addition, phosphate solubilizing microbes generally improved the growth and nutrient uptake of Robusta coffee grown in phosphorus deficient soil[19]. The ability of the rhizobacterial isolates to solubilize inorganic phosphate indicates their potential role to improve the growth of crops. It is recommended to evaluate the amount of phosphorus that these rhizobacteria will contribute to the coffee plants once inoculated.

    • In this study, Bacillus sanguinis demonstrated HCN production, suggesting its potential role as a biological agent against plant pathogens. HCN is mostly synthesized by Bacillus species, which is believed to disrupt several cellular processes such as the electron transport chain, correct functioning of enzymes and even impedes the action of cytochrome oxidase of the target pathogens[20]. HCN production of phosphate-solubilizing Bacillus isolates was also observed as a biochemical trait of potential biofertilizer agent for coffee production[21].

      On the other hand, Bacillus sanguinis and Burkholderia sp. had promising potential as a biocontrol agent due to its capability to produce multiple lytic enzymes, which is an essential strategy for fungal inhibition. Production of lytic enzymes of rhizobacteria is considered as a plant disease inhibitor since these enzymes are involved in cell wall degradation of plant pathogens present in the soil environment[22,23]. Lytic enzymes can be used as biocontrol agents to inhibit fungal pathogens that cause diseases in crops[24].

    • Abiotic stresses such as salinity, acidity or alkalinity, and heavy metal contamination may have a detrimental effect on agricultural production by affecting plant growth, nutritional imbalance, and physiological and metabolic changes. Rhizobacteria may help in plant growth promotion and alleviation of the stress-induced changes in the host plant[25]. Rhizobacteria that tolerate high concentrations of NaCl can help plants thrive in saline soil through their mechanisms such as osmolytes regulation, nitrogen fixation, solubilization of phosphate, as well as formation of auxin, siderophore and exopolysaccharides[26]. Meanwhile, plants are vulnerable to heavy metal stress exposure, which may came from industrial and other environmental pollutants. Rhizobacteria are essential for the reduction of toxic heavy metals in plants, they can contribute to the improvement of heavy metal tolerance and plant growth through improved phytoremediation and metal accumulation inside the plant[27]. The pH is also considered as a limiting condition for plants but stress tolerant rhizobacteria considerably enhance the seed germination of crops in acidic or alkaline soils[28]. The tolerance of isolates at low pH is expected since the pH of the soil used in the present study is acidic. The ability of the rhizobacteria to withstand a wide range of abiotic stresses indicates their potential to promote plant growth and protect plants from the detrimental effects of abiotic stresses in the soil.

    • The bacterial isolates obtained from the Liberica rhizosphere are Pantoea rwandensis, Bacillus pseudomycoides, and Pantoea sp. that exhibited tolerance to high salt concentration and wide range of pH and heavy metal concentrations, but only B. pseudomycoides exhibited amylase activity. On the other hand, the Robusta rhizobacterial isolates identified in our study were Burkholderia cepacia, Bacillus sanguinis and Burkholderia sp. that showed promising performance, specifically the B. sanguinis that has the ability to produce multiple lytic enzymes and HCN. The result of this study indicates that both Liberica and Robusta rhizosphere contained beneficial rhizobacteria that establish mutual symbiotic relationship with the host plant. Previously, phosphate solubilizing species under the genera Bacillus and Burkholderia and the species of endophytic Pantoea were reported to be abundant in the rhizosphere of forest-grown Coffea arabica[12,29]. These rhizobacterial isolates demonstrated a great potential to be utilized as a biofertilizer to improve the growth and yield of coffee.

      The bacterial genus Pantoea was found to be abundant in the rhizosphere microbiome and bean fermentation of coffee[30]. This genus comprises many versatile endophytic species that possess a variety of biosynthetic and biodegradative capabilities. They are found to be useful for biocontrol of plant pathogens, bioremediation of contaminated environments, biosensors, and a source of therapeutic drugs[31]. In addition, Bacillus species was discovered to be a phosphate-solubilizer and enhanced the growth of coffee seedlings and the availability of phosphorus in the soil[32]. Specifically, B. pseudomycoides was found to be a good choice in phytoremediation and an active agent in biofertilizers or biofungicide[33]. Meanwhile, B. sanguinis was similarly found to produce multiple lytic enzymes such as amylase, cellulase, protease, and xylanase[34], indicating its important role against plant pathogens. Lastly, Burkholderia species in general can establish symbiotic relationships with terrestrial plants, functioning as active rhizosphere components, endophytic plant colonizers, or microsymbionts in legume root nodules[35].

    • Our work successfully screened indigenous and beneficial rhizobacteria associated with Robusta and Liberica coffee rhizosphere. They belong to the genera Bacillus, Burkholderia, and Pantoea, which were reported to have plant growth-promoting attributes, biocontrol activities, and tolerance to abiotic stresses. These bacteria can be utilized as an alternative to chemical fertilization and be used as a potential biofertilizer for coffee cultivation in the Philippines. The performance of these isolates as a biofertilizer may vary, but their effectivity can only be verified upon application to crops.

    • All the authors confirm equal contributions for the following: study conception and design, data collection, analysis and interpretation of results, and manuscript preparation.

    • All data generated or analyzed during this study are included in this published article.

      • The study was funded by the Research Center of Cavite State University, Philippines through the Small Scale CvSU Research Grant. The authors are grateful to CvSU-Research Center for allowing them to conduct experiments in the Bacteriology and Genetics Laboratories of the Interdisciplinary Research Building.

      • The authors declare that they have no conflict of interest.

      • Copyright: © 2023 by the author(s). Published by Maximum Academic Press, Fayetteville, GA. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (2)  Table (3) References (35)
  • About this article
    Cite this article
    Navarro GVD, Quirong DD, Maghanoy GA, Cortes AD. 2023. Characterization and identification of rhizobacteria associated with Liberica and Robusta coffee rhizosphere. Technology in Horticulture 3:24 doi: 10.48130/TIH-2023-0024
    Navarro GVD, Quirong DD, Maghanoy GA, Cortes AD. 2023. Characterization and identification of rhizobacteria associated with Liberica and Robusta coffee rhizosphere. Technology in Horticulture 3:24 doi: 10.48130/TIH-2023-0024

Catalog

  • About this article

/

DownLoad:  Full-Size Img  PowerPoint
Return
Return