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Article

Proper Biochar Increases Maize Fine Roots and Yield via Altering Rhizosphere Bacterial Communities under Plastic Film Mulching

by
Yanghui Sui
*,
Yanbo Wang
*,
Wanxin Xiao
,
Cheng Chang
,
Shuping Zhang
and
Haiyan Zhao
Corn Research Institute, National Engineering Laboratory of Maize, Liaoning Academy of Agricultural Sciences, Shenyang 110161, China
*
Authors to whom correspondence should be addressed.
Agronomy 2023, 13(1), 60; https://doi.org/10.3390/agronomy13010060
Submission received: 7 November 2022 / Revised: 8 December 2022 / Accepted: 21 December 2022 / Published: 24 December 2022
(This article belongs to the Special Issue Crop Yield Formation and Fertilization Management)
Browse Figures
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Abstract

:
Biochar amendment is considered a sustainable agricultural strategy to improve crop yields. However, information on grain yield, fine roots and in relation to rhizosphere microbial communities in maize under plastic film mulching is very limited. Herein, biochar applied every 2 years (8.4 t ha−1, B1) and biochar applied every 5 years (21 t ha−1, B2) combined with nitrogen (225 kg ha−1), or nitrogen alone, were tested in a field experiment. The results showed that a biochar–fertilizer application significantly decreased the root length at the V9 stage, but biochar applied every 5 years significantly maintained the root length at the R6 stage. Biochar–fertilizer application increased grain yield under the B1 treatment while slightly decreasing under the B2 treatment. The rhizosphere of maize was preferentially colonized by Proteobacteria, Firmicutes, Sphingomonas, and Bradyrhizobium. Dominant phyla including Proteobacteria were enriched in bulk soils, while Bacteroidetes and Firmicutes were depleted in rhizosphere and bulk soils under the biochar–fertilizer application. Changes in root morphology and soil properties were responsible for bacterial community structure in response to different biochar applications. Thus, we concluded that the differential responses of maize yield and root attributes might be related to the specific biochar dose-specific effects on soil microbiome diversity.

1. Introduction

Biochar is a carbon-rich and porous solid material produced by the pyrolysis of biomass in oxygen-limited conditions [1,2]. It is characterized by a large surface area, high porosity, and high sorption ability [3,4], which may help to mitigate the impact of climate change through enhancing soil’s carbon sequestration capacity [5,6]. In addition, biochar has positive effects on soil physicochemical properties, such as improving soil water-holding capacity and enhancing nutrient retention and cation exchange capacity; hence, biochar amendment can improve soil quality and crop yield [7,8,9]. In recent decades, increasing attention has been paid to the mechanisms underpinning crop growth promotion by biochar [8], especially how crop plants may adapt to soil amendment with biochar.
Roots are the bridge between plants and soil, and are the first point of contact with biochar particles. The root system is highly sensitive to prevailing soil conditions such as soil structure and nutrient status. Moreover, biochar amendment can affect root–soil interactions by facilitating mineral nitrogen retention in the rhizosphere of barley [10]. However, previous studies on root responses to biochar amendment are generally limited to root morphological and biomass measurements [10]. Therefore, the mechanisms controlling biochar–root interactions remain poorly understood [11].
Changes in root morphology and physiology are direct responses of plants to biochar amendment. Fine roots, which play a major role in plant uptake of soil nutrients and water during growth, may extend into the internal pores of biochar [12,13,14]. The lifespan of fine roots is of significance to plant growth, crop productivity, plant–environment interactions, and soil carbon and nutrient cycles in farmland [15,16,17,18]. Indeed, Lambers et al. (1987) [19] estimated that ~50% of photosynthetic diurnal products in crop plants are used to support fine root growth and maintenance. However, information on the responses of fine roots following biochar amendment is scarce.
The rhizosphere is the direct region of nutrient transformation and absorption in the root–soil system. It represents a dynamic zone for diverse biochemical interactions between plant roots and soil microorganisms [20,21], and microorganisms play a vital role in regulating nutrient transformation and absorption. Through its effects on soil structure [22] and nutrient cycles [23], biochar is believed to provide a preferential habitat for potentially beneficial soil microorganisms [24], which in turn may promote plant growth [25]. In addition, biochar–root interactions can indirectly affect soil biogeochemistry, such as nutrient availability and water-holding capacity [26], thereby altering soil microbial diversity and community structure in the rhizosphere [27,28].
The soil fertility and structure have a direct impact on root growth. The biochar treatments can change the soil fertility (e.g., TN and MBN) by increased soil enzymes activities related to nitrogen metabolism [29]. According to Yang et al., biochar altered the behavior of several nitrifying soil bacteria to elevated concentrations of TN and AN in the soil [30]. Biochar could increase wheat growth traits, which were linked to improving the main environmental factors (e.g., soil NH4+-N, NO3−-N, TN, urease and catalase activities) with a significant change in the rhizobacteria community [31,32]. The effects of biochar on soil microbial abundance and community composition may differ between soil compartments [14] and crop species [27,33]. Sarma et al. [34] showed that biochar amendment had a positive influence on the total microbial biomass in a wheat field. By contrast, in acidic soils, Noyce et al. (2015) [35] found that 5 t ha−1 biochar amendment had neither beneficial nor toxic effects on soil microbial communities in a northern hardwood forest. Root exudates of various crops have distinct effects on the microbial community in rhizosphere soil [36,37]. In this process, biochar may simultaneously play direct and indirect roles due to its particular properties [38]. Interactions between plant roots and specific microbial taxa can induce plant systemic resistance and promote plant nutrient uptake [39,40]. In fact, some microbial species accelerate the decomposition of soil organic matter, providing their host plants with mineral nutrients [41]. However, the effects of biochar were investigated mainly in terms of changes in soil physicochemical properties, crop yield, and biomass production, while changes in the rhizosphere microbial communities of maize were not explored.
In the present study, a field experiment was conducted to explore interactions between fine roots and rhizosphere bacteria in response to biochar and nitrogen fertilizer application in maize. We hypothesized that biochar–fertilizer application may facilitate the development and delay the senescence of fine roots by promoting the aggregation of specific bacterial taxa toward the rhizosphere of maize.

2. Materials and Methods

2.1. Study Area

The field experiment was conducted in Jianping County, northwest Liaoning Province, China (126°45′ E, 45°44′ N). The study area has a north temperate continental monsoon climate, with mean annual temperature and precipitation of 7.6 °C and 614.7 mm, respectively. The mean duration of annual sunshine is 2850 h, and the frost-free period lasts 120 days. Most hot and rainy days occur in the same season, from June to August. Continuous cultivation of maize is the main cropping system in this area. The soil at the experimental site was classified as Haplustalfs by the USDA ‘Soil Taxonomy’ system. The soil texture was sandy loam [42]. At the beginning of the experiment, the basic properties of the sampled soil (0–20 cm depth) are shown in Table 1. The test cornstalk derived biochar was produced by Liaoning Jinhefu Agricultural Technology Co., Ltd. (Anshan, China). Biochar preparation process was using the methods of Gao et al. (2019) [43]. The biochar had the following properties: total carbon content = 366.0 g∙kg−1, total nitrogen content = 12.2 g∙kg−1, pH = 7.6 (1:2.5, biochar/water, w/v).

2.2. Experimental Design

A complete randomized block design with three treatments and three replications was employed (n = 9) in the field trial conducted during 2017–2018. Before the experiment, the land was cultivated each year with spring maize. Our experiment used film-mulching of big ridge double line (Figure 1). The plastic film used for film mulching was black polyethylene film with a thickness of 0.008 mm. The treatments were as follows: (1) conventional nitrogen fertilization treatment (CK, control, based on local practices, 225 kg ha−1 nitrogen fertilizer); (2) application of 8.4 t ha−1 biochar (first year addition but second year without biochar) plus 225 kg ha−1 nitrogen fertilizer (B1); (3) application of 21 t ha−1 biochar (first year addition but second until fifth year without biochar) plus 225 kg ha−1 nitrogen fertilizer (B2).
We experimented with conventional tillage before sowing in both years. Biochar was applied 5 days before sowing, while all fertilizers (N, P, and K) were applied in furrows 1 day before sowing. Biochar was uniformly distributed over the furrow and fully mixed with the 0–20 cm soil layer using a cultivator. All treatments including the control received equal amounts of mineral fertilizers in the form of triple super phosphate and potassium chloride (120 kg P ha−1 and 90 kg K ha−1, respectively). Maize (Zea mays L.) from a common maize hybrid variety in China (Zhengdan 958) was planted on 4 May 2017 and 2018, and harvested on 8 October 2017 and 28 September 2018. The planting density was 6.75 plants m−2. The field management followed local cultivation practices.

2.3. Sampling and Measurements

2.3.1. Root Analysis

Root sampling was performed at V9 and R6 maize growth stages. In each plot, soil monoliths were taken in three cores (20 cm wide, 60 cm long, and 60 cm depth) for root distribution analysis. Root morphology was measured by Epson Expression 12000XL flatbed scanner (Epson, Long Beach, CA, USA) [44]. Large root samples were divided to avoid overlapping during scanning. Images were analyzed using WinRhizo Pro V.2007d (Regent Instruments, Inc., Quebec, QC, Canada) to measure the total root length (RL), total root surface area (RSA), total root volume (RV), and average root diameter. Root diameters were measured and categorized referring to Brennan et al. (2014) [45]. Six plants per treatment were measured to assess the root traits.

2.3.2. Soil Physicochemical Analysis

Soil cores (5.1 cm diameter, 20 cm long) were collected from the field following harvest. In each plot, five cores were collected and mixed thoroughly to form one composite sample. Composite samples were sealed in plastic bags and maintained on ice in an insulated box. After being transported to the laboratory, root fragments and other debris were removed manually. Soil samples were air-dried, ground, and passed through a 2 mm sieve. Total carbon and nitrogen were analyzed using an Elementar Variomax CNS Analyzer (Elementar). For bulk density analysis, soil samples were collected from field plots using a cylinder with a volume of 100 cm3. Soil porosity was calculated as follows: soil porosity = (1 − bulk density/soil specific gravity) × 100 [46].

2.3.3. Yield Measurements

When 95% of the maize crop had dried in the field after reaching physiological maturity, we collected plants from a 12 m2 area (5 m × 4 rows) in the center of each plot to determine the population-level grain yield under the condition of 14.0% grain water content.

2.3.4. Bacterial Community Analysis

Soil that was tightly attached to the fine roots was collected as rhizosphere soil, while the remaining soil served as bulk soil. Eighteen fresh samples (nine rhizosphere soils and nine bulk soils) were sieved through a 2 mm sieve to remove roots and stones. Sieved samples were divided into two subsamples, one of which was used for DNA extraction. Total genomic DNA was extracted from 0.5 g of soil samples using a PowerSoil DNA Isolation Kit (Mo Bio Laboratories, Inc., Carlsbad, CA, USA) following the manufacturer’s instructions. DNA concentration and purity were monitored on 1% agarose gels. The extracted DNA was diluted to 1 ng/µL with sterile water and used as template for PCR amplification.
Fragments of the bacterial 16S rRNA gene were amplified by PCR using primer pair 16S-341F/16S-785R targeting the V3–V4 hypervariable region (forward 5′-CCTACGGGNGGCWGCAG-3′, reverse 5′-GACTACHVGGGTATCTAATCC-3′) [47]. Sequencing libraries were constructed using an NEB Next Ultra DNA Library Prep Kit for Illumina (NEB, Hitchin, UK) following the manufacturer’s recommendations, and index codes were added. Library quality was assessed on a Qubit 2.0 Fluorimeter (Thermo Fisher Scientific, Waltham, MA, USA) and an Agilent Bioanalyzer 2100 system (Agilent Technologies, Santa Clara, CA, USA). Sequencing was performed on an Illumina MiSeq platform (Illumina, San Diego, CA, USA) and 250/300 bp paired-end reads were generated. Sequence analysis was carried out using the QIIME software package [48]. After quality control and chimeric filtering, sequences were clustered into operational taxonomic units (OTUs) at a threshold of 97% similarity.

2.4. Statistical Analysis

One-way analysis of variance (ANOVA) was conducted using SPSS v18.0 (IBM Corp., Armonk, NY, USA) to test the effects of different treatments on root morphological characteristics, soil properties, and bacterial alpha-diversity indices (e.g., observed OTUs, Shannon, Simpson, Chao1, ACE). Duncan’s multiple range tests (p < 0.05) were performed to test for significant differences between treatments. Differences in the relative abundance of individual taxa between two groups were determined using Metastats [49]. For beta-diversity analysis, OTU counts at 97% sequence identity were normalized using the variance-stabilizing transformation function implemented in DESeq2 [50]. Unconstrained principal coordinate analysis (PCoA) was performed with the PCoA function from the ape package [51]. Taxonomy dendrogram displaying the log2 fold change in relative abundance with respect to soil treatments. The relationships between ten dominant bacterial phyla, root characteristics, and soil variables were measured with Pearson correlations. RDA was used to identify the relationship between the bacterial phyla and environmental variables from rhizosphere soil by Canoco 5.0.

3. Results

3.1. Changes in Soil Properties

During the two years of the experiment, the B1 treatment resulted in the highest mean soil total nitrogen content after harvest (Table 2). The total nitrogen content of the B1 treatment was ~75% and ~34% higher than that of CK in 2017 and 2018, respectively (p < 0.05). In addition, both B1 and B2 treatments significantly decreased the carbon-to-nitrogen (C/N) ratio, by 32–34% compared with controls in the first year (p < 0.05), while no evident differences were observed in the second year. Soil bulk density and porosity did not differ significantly between treatments.

3.2. Responses of Maize Root and Yield

To explore the effects of biochar–fertilizer application on root growth and senescence, we analyzed root morphological characteristics during the vegetative (V9) and reproductive (R6) growth stages of maize. At the V9 stage, the mean diameter, surface area, and volume all increased with an increasing biochar application rate, albeit not significantly (p > 0.05; Figure 2). For example, the root volume of the B2 treatment was 54.8% greater than that of CK. By contrast, the total root length responded negatively to biochar–fertilizer treatment. The total root length was lowest in the B2 treatment, which decreased by 5.8% and 13.2% compared with those of CK and B1, respectively (p < 0.05). At the R6 stage, the mean root diameter, surface area, and volume did not respond significantly to biochar–fertilizer treatment, while total root length increased significantly in the B2 treatment compared with CK (p < 0.05). Comparison of the two growth stages revealed that all root values were decreased at the R6 stage relative to the V9 stage. The smallest change in total root length was observed for the B2 treatment (27.9%), while the largest change was observed for CK (41.7%). Specifically, the total root length decreased from 7800.1 cm (V9) to 5623.6 cm (R6) in the B2 treatment, and from 8983.3 cm (V9) to 5233.2 cm (R6) in CK.
Classifying the root diameters into different classes revealed that fine roots with a diameter < 0.5 mm were predominant in all treatments (>60%), suggesting that fine roots contributed most to the total root length of maize. At the V9 stage, the largest percentage of fine roots was observed in the B1 and CK treatments, while the percentage of coarser roots (main and lateral roots, diameter > 0.5 mm) increased with increasing the biochar application rate. At the R6 stage, the root diameter distribution was consistent across all treatments (Figure 3). In terms of root length, maize roots growing in the biochar-amended soils (especially B2) appeared to have fewer fine root patterns than those in CK at the V9 stage (p < 0.05). At the R6 stage, the root length contributed by lateral roots with a diameter ranging from 0.5 to 1 mm substantially increased from 899.8 cm for CK to 1133.7 cm for B2. These results suggest that the B2 treatment decreased the senescence level of fine roots by 15.3% compared with CK (Figure 4a). The root length of main roots > 1 mm in diameter did not differ significantly between treatments during the two growth stages. Maize grain yield in B1 was 12.50 t ha−1 on average across the treatment (Figure 4b), which was higher than CK (11.28 t ha−1) and B2 (10.91 t ha−1) (p > 0.05; p < 0.05).

3.3. Differences in Bacterial Diversity and Community Composition

To investigate the changes in bacterial alpha-diversity, we profiled the communities assembled in the rhizosphere and bulk soils via high-throughput sequencing. Subsampling of bacterial 16S rRNA gene sequence reads resulted in 35,449–44,250 sequences within each sample. Good’s coverage estimates exceeded 99% for all samples, indicating that the sequencing depth was sufficient to capture the bacterial diversity present within the soils (Table 3). A total of 347,975 high-quality sequences were clustered into 1094 OTUs with an average length of 450.40 bp. Irrespective of soil compartment, the alpha-diversity indices did not significantly differ between the treatments. However, higher species richness was observed in bacterial communities from rhizosphere soils compared with bulk soils, according to the observed OTUs, Chao1, and ACE indices. Soil compartment was not significantly associated with the differences in bacterial diversity based on the Shannon and Simpson indices.
To evaluate the differences in bacterial beta-diversity across 18 different soil samples, we conducted PCoA based on the Bray–Curtis distance matrix. Beta-diversity analysis revealed distinct differences in rhizosphere soil samples compared with bulk soil samples during the V9 stage. The PCoA plot displays a clustering pattern of samples depending on soil compartment (Figure 5). Regarding the biochar–fertilizer treatment, B1 is separated from the other two groups along the second principal coordinate (PCo2). The first (PCo1) and second (PCo2) principal coordinates cumulatively explain 54.32% of the total variation in bacterial community composition. B1 is clearly distinguished from B2, reflecting the pronounced effects of the biochar application rate on the bacterial community composition.

3.4. Predominant Bacterial Phyla

The taxonomic distributions of the bacterial communities were evaluated at different levels of classification (Figure 6). The relative abundance of dominant bacterial phyla (>1% of total sequences) was similar among all soil samples. Proteobacteria (47.2%), Bacteroidetes (13.4%), Acidobacteria (9.8%), Firmicutes (7.6%), Planctomycetes (6.3%), Actinobacteria (6.1%), Verrucomicrobia (3.0%), Gemmatimonadetes (2.2%), Nitrospirae (1.6%), and Chloroflexi (1.2%) were the most abundant phyla in both the rhizosphere and bulk soil samples (Figure 6). These phyla accounted for >97% of the total bacterial community in each soil compartment. Particularly, Proteobacteria was dominant across all treatments, with a relative abundance varying from 42.3% to 54.2% in rhizosphere soils and from 41.7% to 48.3% in bulk soils. The B2 treatment significantly enriched Proteobacteria in bulk soils, by 3.3% compared with CK, whereas Bacteroidetes and Firmicutes were depleted by 1.1% and 10.3% (or 2.2% and 4.5%) in bulk soils (or rhizosphere soils), respectively. The community of the B1 treatment contained a significantly higher relative abundance of Verrucomicrobia (p < 0.05) in bulk soil. In rhizosphere soils, biochar affected the abundance of some dominant phyla, but not significantly compared with the CK.
A comparison of different phyla (Figure 6) showed that some of the major taxa in the rhizosphere and bulk soils exhibited consistent responses to biochar–fertilizer treatment in both cases. For example, in biochar-amended soils, Proteobacteria and Verrucomicrobia were enriched, while Firmicutes and Actinobacteria were depleted compared with those in controls. Whether biochar promoted specific bacteria in the rhizosphere while decreasing in bulk soil remained unknown. Thus, we next explored whether any particular bacteria aggregate in the rhizosphere of maize following biochar and nitrogen fertilizer application. To address this question, we calculated the percentage of rhizosphere-responsive OTUs for which the relative abundance shifted in the same direction (i.e., enrichment or depletion in the rhizosphere) within the major taxa (Figure 7). At the phylum level, Chlamydiae showed the greatest enrichment toward the rhizosphere. By contrast, Bacteroidetes, Planctomycetes, Acidobacteria, and Firmicutes were all depleted in the rhizosphere.

3.5. Root Morphology and Soil Properties Shaped the Rhizosphere Bacterial Community

The relative abundance of the 10 dominant rhizosphere bacterial phyla exhibited strong relationships with root characteristics and soil properties (Table 4). For example, Firmicutes abundance was negatively correlated with soil porosity and positively correlated with the C/N ratio and bulk density (p < 0.05 or 0.01). Actinobacteria abundance was negatively correlated with the root surface area (p < 0.01). Verrucomicrobia abundance was negatively correlated with the soil C/N ratio and positively correlated with the soil total nitrogen content (p < 0.05). Chloroflexi abundance was negatively correlated with the root length and soil bulk density, and positively correlated with soil porosity (p < 0.05).
To further determine the relative contribution of environmental variables on shaping the rhizosphere bacterial community, the TN was identified as a variable that significantly contributed to the soil bacterial community. The three root variables correlated with soil bacterial community, including root length, root surface area, and different root diameter classes. RDA analysis showed that these factors together explained the 93.96% variation of the soil bacterial community composition (Figure 8).

4. Discussion

4.1. Biochar Alters the Maize Fine Root Morphology

The root system (the major plant–soil interface) plays an essential role in the uptake of nutrients and water required for crop productivity [52]. In particular, fine roots constitute a key part of the root system for maintaining normal crop physiology [53,54,55]. Plant roots are associated with a diverse range of microorganisms that are highly responsive to the soil environment [39]. Previous studies showed a clear shift in the root-associated microbial community structure of sweet pepper plants grown in biochar-amended soil, characterized by a substantial induction of several chitin- and aromatic compound-degrading genera [39]. This prompted us to hypothesize that physical and chemical factors (e.g., biochar-associated organic compounds) may collectively contribute to microbial community shifts, and altered microbial communities may be at least partially responsible for plant growth promotion and resistance induction. Characterization of fine root morphology, together with rhizosphere bacterial community structure, is therefore a useful approach to explore microbially mediated rhizosphere processes of crops following biochar–fertilizer application.
We found that at the V9 stage, the total root length of maize treated with different rates of biochar plus nitrogen fertilizer was decreased considerably, while the corresponding root surface area and volume were slightly increased compared with controls (Figure 2). This result indicates that the total number of maize roots increased in biochar-amended soils during the vegetative growth stage. However, at the R6 stage, the roots of maize treated with 21 t ha−1 biochar plus nitrogen fertilizer became longer and slightly thinner than those of controls, suggesting a delay in root senescence. These results are consistent with previous observations of Sun et al. [56] showing that 2 t ha−1 year−1 biochar amendment increased the total root length of maize and decreased the root diameter at the three-leaf stage (V3). In addition, several studies indicated that biochar–fertilizer application could increase the root biomass and fine root length of maize [57], and the total root length of lettuce [58]. Consistently, we also found that the root system of maize comprised more lateral roots (diameter 0.5–1 mm) when growing in soil amended with 21 t ha−1 biochar plus nitrogen fertilizer, relative to controls (Figure 3). This observation suggests that the biochar–fertilizer application delayed the senescence of fine roots in maize at the mature stage by facilitating plant acquisition of soil nutrients.
Several studies have suggested that biochar has a positive effect on root length because it directly expands the crop rhizosphere to facilitate nutrient and water uptake [10,59]. There is also evidence indicating that this positive effect on root growth involves indirect pathways; biochar acts as a nutrient source enhancing nutrient availability [10], and as a porous matrix providing a preferential habitat for potentially beneficial soil microorganisms [36]. Indeed, we observed that the biochar–fertilizer application altered soil total nitrogen content, bulk density, and porosity (Table 2), while species richness in rhizosphere bacterial communities was also altered in the maize field (Table 3). The possible mechanisms are discussed in the following section.

4.2. Biochar Shapes Maize Rhizosphere Bacterial Communities

Soil bacteria can adapt to a changing environment of cropland via motility, chemotaxis, quorum sensing [60], and/or specific substrate utilization [61]. Previous reports revealed that biochar amendment alters soil physicochemical properties, and such environmental changes subsequently shift bacterial community structure [11,62]. As expected, we found that biochar–fertilizer application increased soil total nitrogen content and porosity, while decreasing soil bulk density (Table 2). These changes in the soil environment are beneficial for the growth of specific bacteria [11,63]. This relationship suggests that biochar improved soil bacterial diversity by altering the physicochemical conditions [64].
Biochar has variable effects (positive, negative, and null) on rhizosphere soil microbial communities [65,66,67], depending on the raw material, preparation technology, and application rate [68]. Herein, we revealed that the relative abundance of several rhizosphere-associated phyla (e.g., Actinobacteria) was much higher in the maize rhizosphere than in bulk soils, regardless of the biochar application rate (Figure 6).
At the phylum level, some studies have indicated that biochar amendment reduces the abundance of Proteobacteria, Acidobacteria, Bacteroidetes, and Firmicutes in soil [39,69]. In contrast, others have shown that biochar amendment increases the abundance of Nitrospiraceae, Gemmatimonadetes, Chloroflexi, and Firmicutes in rhizosphere soil [63]. In the present study, biochar resulted in an enrichment of Proteobacteria and a depletion of Firmicutes in the maize rhizosphere compared with controls (Figure 6). Xu et al. [64] and Mueller et al. [70] similarly observed that the relative abundance of Proteobacteria increased in metal-contaminated soils after amendment with wine lees-derived biochar. It is well known that Proteobacteria plays a vital role in the metabolism of chemically diverse carbon compounds [71], possibly due to an increase in carbon sources in biochar-amended soil [72].
Our results suggest that biochar-induced changes in the maize root morphology and rhizosphere bacterial community structure may indirectly affect root exudation. Sun et al. [56] found that biochar amendment strongly increased the number of organic acids in root exudates and thereby promoted root elongation in Northeast China. The root system releases carbon exudates, which in turn alter the microbial community structure and carbon pool turnover in soil [11,73]. However, interactions between fine roots and rhizosphere microorganisms are often ignored in root research, and this hinders our ability to determine the role of biochar in soil for crop growth. In the present study, biochar extended the fine root lifespan and altered the community structure of bacterial responders in the rhizosphere of maize. We also observed a significant negative correlation between Actinobacteria abundance and root surface area (Table 4). This result indicates that biochar–fertilizer application directly influenced root morphology and indirectly affected rhizosphere bacterial community structure by mediating root exudation.

5. Conclusions

This study revealed that biochar amendment had greater effects on root properties, grain yield, and microbiome diversity than nitrogen addition alone. Maize plants subjected to biochar amendment tended to develop roots (diameter < 1 mm) longer, indicating that biochar input delayed root senescence from the V9 to the R6 stage. In addition, biochar influenced the relative abundance of dominant bacterial taxa involved in soil carbon and nitrogen cycling, including Proteobacteria, Firmicutes, Sphingomonas, and Bradyrhizobium. For the top 10 dominant bacterial phyla, the abundance of specific taxa was closely related to soil properties and root characters. Our results reveal a clear relationship between root morphological characteristics and the rhizosphere bacterial community structure of maize. Root morphology plays a distinct role in shaping the structure of rhizosphere bacterial communities under different biochar management strategies, and distinct microorganisms in turn regulate crop productivity.

Author Contributions

Conceptualization, Y.W. and Y.S.; methodology, Y.S. and H.Z.; software, Y.S.; formal analysis, Y.S.; investigation, W.X., C.C. and S.Z.; data curation, Y.S. and S.Z.; writing—original draft preparation, Y.S.; writing—review and editing, Y.S., Y.W., W.X., C.C., S.Z. and H.Z.; funding acquisition, Y.S. and Y.W. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Natural Science Foundation of Liaoning Province, China, grant number 2020-MS-048; the National Natural Sciences Foundation of China, grant number 32101832; the National Modern Maize Industry Technology System, China, grant number CARS-02-52; and the China Postdoctoral Science Foundation, grant number 2017M611255.

Data Availability Statement

Raw reads have been deposited in the Short Read Archive of NCBI under project no. PRJNA908286.

Acknowledgments

The authors particularly thank Wenfu Chen and Jiping Gao from Liaoning Biochar Engineering & Technology Research Center for technical assistance. The authors are grateful to the anonymous reviewers for their constructive comments on the manuscript.

Conflicts of Interest

The authors declare no conflict of interest.

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Figure 1. Schematic illustration of the field layout showing plastic film mulching of big ridge double line.
Figure 1. Schematic illustration of the field layout showing plastic film mulching of big ridge double line.
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Figure 2. Root morphological characteristics of maize altered by biochar treatments. CK, conventional nitrogen fertilization; B1, application of 8.4 t ha−1 biochar plus 225 kg ha−1 nitrogen fertilizer; and B2, application of 21 t ha−1 biochar plus 225 kg ha−1 nitrogen fertilizer. (a) Root diameter; (b) Root length; (c) Root surface area; (d) Root volume. Data are mean ± SE (n = 3). Different letters above the error bars indicate significant differences between treatment groups at p < 0.05.
Figure 2. Root morphological characteristics of maize altered by biochar treatments. CK, conventional nitrogen fertilization; B1, application of 8.4 t ha−1 biochar plus 225 kg ha−1 nitrogen fertilizer; and B2, application of 21 t ha−1 biochar plus 225 kg ha−1 nitrogen fertilizer. (a) Root diameter; (b) Root length; (c) Root surface area; (d) Root volume. Data are mean ± SE (n = 3). Different letters above the error bars indicate significant differences between treatment groups at p < 0.05.
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Figure 3. Percentage of maize root lengths in different diameter classes during two growth stages of maize under biochar treatments. Main root axes, diameter ˂ 1.0 mm; first-order lateral roots, diameter 0.5–1.0 mm; and second-order fine roots, diameter < 0.5 mm. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption. Data are mean ± SE (n = 3).
Figure 3. Percentage of maize root lengths in different diameter classes during two growth stages of maize under biochar treatments. Main root axes, diameter ˂ 1.0 mm; first-order lateral roots, diameter 0.5–1.0 mm; and second-order fine roots, diameter < 0.5 mm. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption. Data are mean ± SE (n = 3).
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Figure 4. Root lengths of different diameter classes (a) and maize grain yield (b) under biochar treatments. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption. Data are mean ± SE (n = 3). Different letters above the error bars indicate significant differences between treatment groups at p < 0.05.
Figure 4. Root lengths of different diameter classes (a) and maize grain yield (b) under biochar treatments. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption. Data are mean ± SE (n = 3). Different letters above the error bars indicate significant differences between treatment groups at p < 0.05.
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Figure 5. Principal coordinate analysis (PCoA) plot based on Bray–Curtis’s distance matrices showing the differences in bacterial community composition between rhizosphere and bulk soils of maize under biochar application (V9 stage). Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption.
Figure 5. Principal coordinate analysis (PCoA) plot based on Bray–Curtis’s distance matrices showing the differences in bacterial community composition between rhizosphere and bulk soils of maize under biochar application (V9 stage). Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption.
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Figure 6. Relative abundance of different phyla in rhizosphere and bulk soils of maize following biochar application. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption.
Figure 6. Relative abundance of different phyla in rhizosphere and bulk soils of maize following biochar application. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption.
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Figure 7. Taxonomic dendrogram displaying the log2 fold change in relative abundance with respect to soil treatments. Only the OTUs belonging to the most represented higher taxa (innermost circle) are shown. The three outermost rings indicate the number of soil treatments in which the relative abundance of an OTU was significantly higher (brown) or lower (green) in the rhizosphere communities under CK, B1, and B2 (p < 0.05). The nodes in the cladogram indicate the phylum (Phy.), class (Cla.), order (Ord.), and family (Fam.) to which each OTU belongs. The color of the node represents the response consistency (measured as the percentage of OTUs enriched or depleted in the rhizosphere) within the subtree rooted at that node: consistent enrichment toward the rhizosphere is marked in red color, while consistent depletion is marked in blue color. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption.
Figure 7. Taxonomic dendrogram displaying the log2 fold change in relative abundance with respect to soil treatments. Only the OTUs belonging to the most represented higher taxa (innermost circle) are shown. The three outermost rings indicate the number of soil treatments in which the relative abundance of an OTU was significantly higher (brown) or lower (green) in the rhizosphere communities under CK, B1, and B2 (p < 0.05). The nodes in the cladogram indicate the phylum (Phy.), class (Cla.), order (Ord.), and family (Fam.) to which each OTU belongs. The color of the node represents the response consistency (measured as the percentage of OTUs enriched or depleted in the rhizosphere) within the subtree rooted at that node: consistent enrichment toward the rhizosphere is marked in red color, while consistent depletion is marked in blue color. Treatment abbreviations (CK, B1, and B2) are defined in Figure 2 caption.
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Figure 8. Redundancy analysis (RDA) of bacterial community composition (phylum level) and environmental variables from rhizosphere soil of maize fields without and with biochar. Abbreviations: D: root diameter.
Figure 8. Redundancy analysis (RDA) of bacterial community composition (phylum level) and environmental variables from rhizosphere soil of maize fields without and with biochar. Abbreviations: D: root diameter.
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Table
Table 1. Chemical properties of the experimental soil at the beginning of the experiment.
Table 1. Chemical properties of the experimental soil at the beginning of the experiment.
Organic MatterTotal NAvailable PAvailable KpH
(g kg−1)(mg kg−1)
Sandy loam11.190.8632.3569.006.95
Table
Table 2. Soil properties of topsoil (0–20 cm) in maize field following biochar and nitrogen fertilizer application.
Table 2. Soil properties of topsoil (0–20 cm) in maize field following biochar and nitrogen fertilizer application.
YearTreatmentTN (mg g−1)C/NBD (g cm−3)SP (%)
2017CK0.61 ± 0.06a18.57 ± 1.34a1.63 ± 0.02a38.55 ± 0.88a
B11.07 ± 0.26a12.67 ± 3.24b1.54 ± 0.07a41.96 ± 2.66a
B20.89 ± 0.16a12.21 ± 0.62b1.48 ± 0.05a44.01 ± 2.05a
2018CK0.61 ± 0.12a14.86 ± 1.78a1.34 ± 0.07a49.50 ± 2.56a
B10.82 ± 0.11a14.06 ± 1.18a1.25 ± 0.08a52.83 ± 2.90a
B20.80 ± 0.05a13.55 ± 0.54a1.22 ± 0.09a53.99 ± 3.49a
Source of variation
Treatment*****
Yearns**ns**
Treatment × Yearnsnsnsns
Abbreviations: TN: total content of nitrogen; C/N: the ratio of TC and TN; BD: bulk density; SP: soil porosity; * p < 0.05; ** p < 0.01.
Table
Table 3. Effects of biochar and nitrogen fertilizer application on species richness and diversity of bacterial communities in the rhizosphere and bulk soils of maize.
Table 3. Effects of biochar and nitrogen fertilizer application on species richness and diversity of bacterial communities in the rhizosphere and bulk soils of maize.
Soil CompartmentTreatmentObserved OTUsShannonSimpsonChao1ACEGood’s Coverage
RhizosphereCK877 ± 275.89 ± 0.160.0060 ± 0.0023921 ± 19905 ± 230.9973 ± 0.0000
B1868 ± 135.87 ± 0.070.0058 ± 0.0008908 ± 11929 ± 60.9975 ± 0.0005
B2903 ± 35.94 ± 0.020.0053 ± 0.0001940 ± 5896 ± 120.9975 ± 0.0000
BulkCK797 ± 885.63 ± 0.320.0089 ± 0.0045891 ± 51859 ± 610.9961 ± 0.0010
B1847 ± 215.87 ± 0.040.0056 ± 0.0006907 ± 28879 ± 300.9968 ± 0.0013
B2841 ± 315.90 ± 0.090.0052 ± 0.0006887 ± 32889 ± 280.9970 ± 0.0006
Biochar
B0837 ns5.76 ns0.0075 ns906 ns0.9967 ns882 ns
B1858 ns5.87 ns0.0057 ns907 ns0.9972 ns904 ns
B2872 ns5.92 ns0.0053 ns913 ns0.9972 ns893 ns
Soil compartment
Rhizosphere883 a5.90 ns0.0057 ns923 a0.9974 a910 a
Bulk829 b5.80 ns0.0066 ns895 b0.9966 b876 b
Significance
Biocharnsnsnsnsnsns
Soil compartment*nsnsnsns*
Biochar × Compartmentnsnsnsnsnsns
OTU, Operational taxonomic units. Values are means ± standard deviation. Values in the same column followed by different lowercase letters indicate a significant difference. * p ≤ 0.05; ns, not significant.
Table
Table 4. Correlations between dominant bacterial phyla, root characteristics, and soil variables.
Table 4. Correlations between dominant bacterial phyla, root characteristics, and soil variables.
Bacterial PhylumRDRLRSRVD < 0.50.5 < D < 1D > 1TNC/NBDSP
Proteobacteria0.43−0.370.280.40−0.39−0.170.380.09−0.33−0.360.36
Bacteroidetes−0.300.540.39−0.100.430.520.10.60−0.42−0.270.27
Acidobacteria0.020.090.240.090.020.180.230.38−0.59−0.530.53
Firmicutes−0.440.29−0.45−0.450.360.05−0.51−0.600.80 **0.69 *−0.69 *
Planctomycetes0.08−0.110.140.06−0.06−0.320.20.57−0.66−0.620.62
Actinobacteria−0.17−0.12−0.70 *−0.330.03−0.30−0.63−0.290.190.24−0.24
Verrucomicrobia0.46−0.280.460.48−0.370.110.470.70 *−0.81 **−0.630.63
Gemmatimonadetes−0.260.08−0.54−0.380.090.11−0.340.13−0.36−0.380.38
Nitrospirae−0.150.450.630.070.270.580.410.03−0.030.15−0.15
Chloroflexi0.50−0.70 *−0.270.30−0.62−0.450.020.08−0.51−0.69 *0.69 *
Abbreviations: RD: root diameter; RL: root length; RS: root surface area; RV: root volume; BD: bulk density; SP: soil porosity; * p ≤ 0.05; ** p ≤ 0.01.
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Sui, Y.; Wang, Y.; Xiao, W.; Chang, C.; Zhang, S.; Zhao, H. Proper Biochar Increases Maize Fine Roots and Yield via Altering Rhizosphere Bacterial Communities under Plastic Film Mulching. Agronomy 2023, 13, 60. https://doi.org/10.3390/agronomy13010060

AMA Style

Sui Y, Wang Y, Xiao W, Chang C, Zhang S, Zhao H. Proper Biochar Increases Maize Fine Roots and Yield via Altering Rhizosphere Bacterial Communities under Plastic Film Mulching. Agronomy. 2023; 13(1):60. https://doi.org/10.3390/agronomy13010060

Chicago/Turabian Style

Sui, Yanghui, Yanbo Wang, Wanxin Xiao, Cheng Chang, Shuping Zhang, and Haiyan Zhao. 2023. "Proper Biochar Increases Maize Fine Roots and Yield via Altering Rhizosphere Bacterial Communities under Plastic Film Mulching" Agronomy 13, no. 1: 60. https://doi.org/10.3390/agronomy13010060

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