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Volume 4 Issue 4
Oct.  2022
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Article Contents
Abstract
Introduction
Results
Discussion
Conclusion
Materials and methods
Acknowledgements
Author contributions
Compliance with ethical standards
References
Zhe Wang, Yong Chi, Tao Li, Wenya Song, Yunfeng Wang, Tong Wu, Gongaote Zhang, Yujie Liu, Honggang Ma, Weibo Song, Khaled A. S. Al-Rasheid, Alan Warren, Borong Lu. 2022: Biodiversity of freshwater ciliates (Protista, Ciliophora) in the Lake Weishan Wetland, China: the state of the art. Marine Life Science & Technology, 4(4): 429-451. DOI: 10.1007/s42995-022-00154-x
Citation: Zhe Wang, Yong Chi, Tao Li, Wenya Song, Yunfeng Wang, Tong Wu, Gongaote Zhang, Yujie Liu, Honggang Ma, Weibo Song, Khaled A. S. Al-Rasheid, Alan Warren, Borong Lu. 2022: Biodiversity of freshwater ciliates (Protista, Ciliophora) in the Lake Weishan Wetland, China: the state of the art. Marine Life Science & Technology, 4(4): 429-451. DOI: 10.1007/s42995-022-00154-x
shu

Biodiversity of freshwater ciliates (Protista, Ciliophora) in the Lake Weishan Wetland, China: the state of the art

  • 1.

    Institute of Evolution and Marine Biodiversity, Ocean University of China, Qingdao, 266003, China

  • 2.

    Marine College, Shandong University, Weihai, 264209, China

  • 3.

    Laoshan Laboratory, Qingdao, 266237, China

  • 4.

    Zoology Department, College of Science, King Saud University, Riyadh, 11451, Saudi Arabia

  • 5.

    Department of Life Sciences, Natural History Museum, London, SW7 5BD, UK

  • Corresponding author:

    Borong Lu boronglu@126.com

  • Zhe Wang, Yong Chi, Tao Li, Wenya Song, Yunfeng Wang, Tong Wu and Gongaote Zhang contributed equally to this work.

  • Received Date: 2022-04-14
  • Accepted Date: 2022-10-19
  • Published online: 2022-11-27
    Special topic: Ciliatology.
    Springer Nature or its licensor (e.g. a society or other partner) holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.
    Edited by Jiamei Li.
    Graphical Abstract
    • Abstract
  • Ciliates are core components of the structure of and function of aquatic microbial food webs. They play an essential role in the energy flow and material circulation within aquatic ecosystems. However, studies on the taxonomy and biodiversity of freshwater ciliates, especially those in wetlands in China are limited. To address this issue, a project to investigate the freshwater ciliates of the Lake Weishan Wetland, Shandong Province, commenced in 2019. Here, we summarize our findings to date on the diversity of ciliates. A total of 187 ciliate species have been found, 94 of which are identified to species-level, 87 to genus-level, and six to family-level. These species show a high morphological diversity and represent five classes, i.e., Heterotrichea, Litostomatea, Prostomatea, Oligohymenophorea, and Spirotrichea. The largest number of species documented are oligohymenophoreans. A comprehensive database of these ciliates, including morphological data, gene sequences, microscope slide specimens and a DNA bank, has been established. In the present study, we provide an annotated checklist of retrieved ciliates as well as information on the sequences of published species. Most of these species are recorded in China for the first time and more than 20% are tentatively identified as new to science. Additionally, an investigation of environmental DNA revealed that the ciliate species diversity in Lake Weishan Wetland is higher than previously supposed.
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  • Chitin is a natural mucopolysaccharide that consists of 2-acetamino-2-deoxygen-d-glucose and 2-amino-2-deoxygen-d-glucose linked by a β (1 → 4) glycosidic bond (Kumar 2000; Shahidi et al. 1999). The amount of chitin in nature is large, as it is found in crabs and shrimps, fungus, and the cell walls of lower plants (Rinaudo 2006). Chitin also has distinctive biological characteristics; it is non-toxic, antibacterial, biocompatible, and biodegradable (Hirano 1999; Jayakumar et al. 2010; Kurita 2006). Moreover, chitin can show low immunogenicity as well as it has film forming properties and can form fibers (Zheng et al. 2002).

    Functional groups, such as acetyl groups, hydroxyl groups, and carbonyl groups, in the main chain structure of the chitin molecule make it easy to combine with itself or other hydroxyl compounds by hydrogen bonds (Rinaudo 2006; Tamura et al. 2006, 2011). As a result, chitin is difficult to dissolve in most acidic or alkaline solutions and organic solvents, limiting its usage (Tamura et al. 2011). To be usable, chitin needs to be chemically modified, such as through deacetylation and esterification, before application in the medical, food, agricultural, biotechnical, and environmental fields, resulting in an increased interest in chitin nanofibers (Ifuku et al. 2009; Ifuku and Saimoto 2012; Jayakumar et al. 2010; Wang et al. 2018a, b). Chitin nanofibers are generally referred to as a linear fiber or whisker with a nanometer diameter and a high aspect ratio (Ifuku and Saimoto 2012). They have been prepared by a variety of methods, including mechanical treatment, dissolution and regeneration, TEMPO oxidation, and electrostatic spinning (Jia et al. 2002; Kato et al. 2004; Muzzarelli et al. 1999; Zhong et al. 2010). The nanofibers prepared by chitin not only have the features of chitin itself as mentioned above but also are characterized by a high specific surface area and stability in an aqueous solution (Ifuku and Saimoto 2012). The nanofibers are easier to process further and utilize in the food and biomedical industries, e.g., chitin/chitosan packaging film and cell attachment (Jayakumar et al. 2010; Mushi et al. 2014).

    Enzyme immobilization, the restriction of enzymes on solid materials, is a requisite for the application of enzymes as biocatalysts (Han et al. 2016; Sheldon 2007; Wang and Caruso 2005). Enzyme immobilization can prevent instability and unrepeatability of free enzymes because the support of chitin nanofibers facilitates recovery from reaction mixtures (Wang et al. 2009). Nanofibers have been applied to immobilization, with immobilization methods including covalent binding, physical adsorption, and polymer embedding (Iyer and Ananthanarayan 2008; Sheldon 2007; Zhang et al. 2009). In enzyme immobilization, chitin nanofibers are a kind of inert immobilized carrier, making the enzyme reusable, easy to separate, and more stable (Iyer and Ananthanarayan 2008; Khoshnevisan et al. 2011; Mateo et al. 2007). Magnetic nanoparticles, used in enzyme immobilization, can be rapidly separated and immobilized enzymes can be recovered from the reaction system using an external magnet (Deng et al. 2009; Xiao et al. 2017). The use of magnetic supports can also reduce operation costs and prevent problems that can limit recovery (Khoshnevisan et al. 2011; Pan et al. 2009).

    We previously developed a chitin-nanofiber-based support for enzyme immobilization (Huang et al. 2018). Chitin-based supports for enzyme immobilization with high biocompatibility, stability, and recyclability have been previously studied (Huang et al. 2018). However, the relationship and optimization between oxidized time of immobilization support and enzyme efficiency have not been studied. In the present study, chitin nanofibers were prepared using the TEMPO/NaBr/NaClO system with various oxidation times and then used for chymotrypsin immobilization. The characteristics of the TEMPO-oxidized chitin nanofibers were measured by X-ray diffraction (XRD), transmission electron microscopy (TEM), dynamic light scattering (DLS), Fourier transform infrared spectrometry (FT-IR), ultraviolet–visible (UV/VIS) spectrophotometry, and optical microscopy. The relationship between different nanofibers and their carboxylate content, crystallinity, and light transmittance was analyzed. Finally, the effects of oxidation time on the chymotrypsin immobilization were evaluated.

    Characterization of chitin nanofibers

    X-ray diffraction patterns of oxidized chitin

    The diffraction pattern of the original chitin was the crystalline form of the α-chitin. The XRD patterns are shown in Supplementary Fig. S1 and the crystallinity in Table 1. The peaks at 9.3°, 12.8°, 19.2°, 21.3°, 23.2°, and 26.3° correspond to the chitin crystal planes of (020), (021), (110), (120), (130), and (013), respectively (Zhang et al. 2005). The crystallinity of the starting chitin was more than 85%. When the oxidation time was less than 60 min, the small peaks at 12.8° or 23.2° disappeared; however, the (020) and (110) peaks of the oxidized chitin were stable. Furthermore, the crystallinity increased slightly with a short oxidation time (less than 60 min), indicating the disordered regions of chitin may have been destroyed (Jiang et al. 2018). The XRD curves were obviously altered when the oxidation time was between 60 and 480 min. The crystallinity decreased significantly by 10.57% and the intensities of the (020) and (110) peaks were also reduced. These results indicated chitin chains underwent depolymerization via TEMPO oxidation (Kato et al. 2004; Muzzarelli et al. 1999). The water-insoluble CNFs obtained were below 1% when the oxidation time of chitin was 480 min, making it impossible to analyze the characteristics of CNFs.

    Table  1.  Crystallinity of TEMPO-oxidation chitin with various oxidation times
    Oxidization time/min ICR ± RSD/%
    0 89.70 ± 0.64
    20 89.91 ± 0.77
    40 90.16 ± 0.94
    60 87.02 ± 1.12
    120 84.37 ± 2.15
    240 83.30 ± 1.20
    360 80.89 ± 1.29
    480 79.13 ± 1.43
     | Show Table
    DownLoad: CSV
    Transmission electron microscopy images of chitin nanofibers

    The water-insoluble CNFs were obtained by centrifugation and the depolymerized part was removed. The TEM images are shown in Fig. 1 and size distribution of CNFs is shown in Fig. 2. The prepared nanofibers were needle-shaped or whisker-shaped (Fig. 1). The width of the CNFs increased by about 1.4 nm with an increased oxidization time, as shown in Fig. 2. The increased width was related to the inter-whisker linkages formed by electrostatic interactions between carboxyl and amino groups (Fan et al. 2007). However, the average length of the CNFs decreased from 450.38 ± 122.11 nm (at an oxidation time of 20 min) to 241.70 ± 74.61 nm (at an oxidation time of 360 min). These results were consistent with the patterns of dynamic light scattering (Supplementary Table S1 and Fig. S2). The proportion of CNFs with lengths in the range of 200–500 nm was about 68% when the TEMPO-mediated time was 360 min, compared with 32% at 20 min and 49% at 60 min, indicating TEMPO-oxidation could shorten the length of nanofibers with ultrasonic treatment. Meanwhile, the CNFs with lengths of more than 5000 nm with a short oxidation time were about 5% because blocks of chitin fibers existed in the form of aggregation or bundles in the CNFs dispersion.

    Figure  1.  Transmission electron microscopy (TEM) images of chitin nanofibers with various oxidization times (magnification: a–c, × 200, 000; d–f × 100, 000)
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    Figure  2.  Size distribution of chitin nanofibers calculated by TEM images
    DownLoad: Full-Size Img PowerPoint
    Infrared spectroscopy characterization of chitin nanofibers

    The FT-IR spectra of the TEMPO-oxidized chitins are shown in Fig. 3. The absorption peaks at 3431 and 3262 cm−1 are attributed to the O–H and N–H stretching vibrations, respectively. The absorption peaks at 1654 and 1622 cm−1 correspond to the amide I band. These bands are associated with the typical features of α-chitin (Wang et al. 2018a, b). The sodium carboxylate groups of CNFs were converted to protonated carboxyl groups under acidic condition. An absorption band appeared at approximately 1740 cm−1, indicating the hydroxyl groups had been successfully oxidized. The C–O stretching bands at approximately 1030 cm−1 were the internal standard band. As shown in Supplementary Fig. S3, the absorption ratios A1740/A1030 corresponded to the carboxylate contents of the CNFs. The degree of acetylation was plotted as the ratio of A1560/A1030, where the absorbance of the amide II bands appeared at 1560 cm−1 (Shigemasa et al. 1996). The carboxylate contents increased when the oxidation time was shorter than 60 min; the maximum appeared at 60 min. The contents decreased slightly with longer oxidation times. However, the degree of acetylation of CNFs treated with various times was mostly constant, indicating that almost no deacetylation occurred during the TEMPO/NaClO/NaBr oxidation.

    Figure  3.  FT-IR spectra of the chitin nanofibers prepared with various oxidized times
    DownLoad: Full-Size Img PowerPoint

    The transmittance of chitin nanofibers

    The transmittance of chitin nanofibers was evaluated by full wavelength scanning using an ultraviolet–visible spectrophotometer (Fig. 4). While the oxidation time was less than 40 min, the dispersive properties of CNFs were poor and the light transmittance was under 10%. When the oxidation time was 20 min, the transmittance was less than 2% (i.e., the highest transmittance was 1.15% at a wavelength of 800 nm), and the chitin crystal bundles made the dispersion muddy (Fig. 5). As seen in the optical microscopy images (Fig. 6), the CNFs oxidized for 20 min were flake shaped. Results showed there was undissolved chitin or chitin derivatives in the dispersion, making the dispersion non-transparent. The light transmittance was also continuously increased until the oxidation time was 360 min. Meanwhile, the dispersion tended to be transparent. There was no macroscopic flake chitin derivative in the dispersion treated for more than 120 min, suggesting CNFs subjected to ultrasonic treatment tended to be individual rather than aggregated.

    Figure  4.  Light transmittance of chitin nanofibers' dispersions treated with various oxidized times
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    Figure  5.  Photographs of CNFs dispersions prepared with various oxidized times
    DownLoad: Full-Size Img PowerPoint
    Figure  6.  Optical micrographs of chitin nanofibers treated with various oxidation times (magnification: a–c × 100; d–f × 400)
    DownLoad: Full-Size Img PowerPoint

    Content of carboxylate groups

    The carboxylate contents of CNFs are shown in Fig. 7. The carboxylate content increased during TEMPO-treatment of up to 60 min. The content of carboxylate groups was 0.52 mmol/g CNFs, corresponding well to the tendency with the A1740/A1030 ratio (Supplementary Fig. S3). The carboxylate groups slightly decreased with oxidation times beyond 60 min. The results showed depolymerization and excessive oxidation can lead to an increase in water-soluble chitin with large consumption of NaOH and NaClO, with the content of carboxyl groups declining in the water-insoluble part.

    Figure  7.  The content of carboxylate group about different chitin nanofibers' dispersions
    DownLoad: Full-Size Img PowerPoint

    Immobilization enzymes

    CNFs can be applied to enzyme immobilization because carboxyl groups of CNFs have covalent bindings with amino groups. The effects of oxidation time on immobilization are presented in Fig. 8. The enzyme loading increased with increased addition of enzymes when the CNFs were a type of immobilization carrier. While the additional amount of CTs was 500 or 2000 mg/g carrier, the highest loading immobilization and relative enzyme activity were at the oxidation time of 60 min. The highest enzyme loading was 307.17 ± 4.08 mg/g carrier with the addition of CTs at 500 mg/g carrier. When the additional amount was 2000 mg/g carrier, the maximum was 726.82 ± 12.05 mg/g carrier. When the oxidation time was below 60 min, the enzyme loading and relative activity increased with the increase of carboxyl group content. When the oxidation time was beyond 60 min, the CTs immobilization capacity decreased. While the oxidation time was 360 min, the catalytic activity of immobilized CTs only retained about 90% of the maximum relative enzyme because of the decreased carboxyl group content.

    Figure  8.  The immobilization of NH2-Fe3O4 modified chitin nanofibers (CNFs) and chymotrypsins (a 500 mg/g carrier; b 2000 mg/g carrier)
    DownLoad: Full-Size Img PowerPoint

    Chitin has high crystallinity because of the hydrogen bond, and it is insoluble in water or organic solvents, leading to limited usage of chitin. Improving its solubility is a key factor for the industrial application of chitin. TEMPO is a type of organic radical that is widely used as a catalyst for oxidation. The C6 hydroxyl groups on the carbon chain can be converted into carboxylate groups using the TEMPO/NaBr/NaClO system (Muzzarelli et al. 1999). Hence, TEMPO oxidation can effectively improve the solubility and dispersion of chitin. When the oxidation time was less than 60 min, TEMPO could not completely oxidize chitin. Therefore, TEMPO-oxidized chitin was present only on the surface of the chitin bundles, with most chitin residues remaining unreacted in the product (Fan et al. 2007). As a result, the dispersity of the TEMPO-meditated chitin nanofibers with short oxidation time was low and bundles of the fibers and chitin flakes remained. Conversely, long-time TEMPO-oxidation can cause depolymerization of chitin, with the degrees of polymerization of products generally lower than 50 (Muzzarelli et al. 1999). The resulting crystal form of chitin was deformed and the crystallinity decreased. Additionally, solubility and the dispersity of the CNFs increased because of the TEMPO-causing depolymerization.

    CNFs have great potential as a support for enzyme immobilization in various biotechnological applications (Wang et al. 2018a, b). Through the activation of the carboxylate groups using EDC/NHS, the CNFs can immobilize the NH2-Fe3O4 and CTs by covalent bonding. The degree of oxidation of chitin has a significant effect on the immobilization efficiency. As shown in Fig. 8, the enzyme activity and loading capacity of the immobilized CTs increased with the carboxylate group content of the chitin nanofibers. These results occurred because the enzymes were immobilized onto the chitin nanofibers via interaction with the carboxylate groups. Moreover, more carboxylate groups could conjugate more enzymes, resulting in increased enzyme activity and loading capacity of the immobilized enzymes.

    In this study, TEMPO-oxidized chitin nanofibers treated with various oxidation time were prepared and characteristics analyzed. Chitin nanofibers were applied to enzyme immobilization using chymotrypsin as a model enzyme. It was demonstrated that enzyme efficiency could be improved by controlling TEMPO-oxidation time by using chitin nanofibers as support. The CNFs treated with TEMPO for 60 min retained the highest activity of immobilized enzymes and enzyme loading, corresponding to the maximum carboxylate content (0.52 mmol/g CNFs). While the additional amount of CTs was 500 or 2000 mg/g carrier, the highest loading amount of CTs was 307.17 ± 4.08 or 726.82 ± 12.05 mg/g carrier, respectively. The immobilized enzymes showed good correlation to the TEMPO oxidation time. Therefore, the optimal enzyme immobilization achieved with various oxidation times can reduce production costs when used in industrial reactions.

    Materials

    2, 2, 6, 6-Tetramethylpiperidine-1-oxyl (TEMPO) was purchased from Adamas Reagent Ltd. Chitin was purchased from Yuanye Bio-Technology Co. (Shanghai, China). Sodium hypochlorite (NaClO), N-hydroxysuccinimide (NHS) and chymotrypsins (CTs) were purchased from Macklin Biochemical Technology Co. (Shanghai, China). 2-morpholinoethanesulfonic acid (MES) and 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) were purchased from Aladdin Co. (Shanghai, China). Other materials were purchased from Sinopharm Group Co. Ltd. (Shanghai, China). All chemical reagents were of analytical grade.

    Preparation of chitin nanofibers

    The method to prepare chitin nanofibers (CNFs) was similar to that used by Fan et al. (2007) and Huang et al. (2018). Chitin powder (5.00 g, degree of acetylation was 96%) was suspended in water (417.5 ml) containing 1 mmol/L TEMPO (0.08 g) and 10 mmol/L sodium bromide (0.51 g). The oxidation of chitin was started by adding NaClO solution (4%, 82.50 ml, equal to 10 mmol/g chitin) into the slurry. The pH of the chitin slurry was maintained at 10 through 1 mol/L NaOH with magnetic stirring at room temperature. After different reaction times (20–480 min), the oxidization reaction ended by adding ethanol until the pH did not change. The pH of the mixture was adjusted to 7.0 with 0.5 mol/L HCl or 0.5 mol/L NaOH. The slurry was centrifuged at 9500 rpm for 15 min. The precipitate was washed with deionized water through repeated centrifugation and reduplicative suspension. After the precipitate was mixed with deionized water and a consistent concentration (0.1%) remained, the CNFs were treated with ultrasonic disruption (360 W, 30 min, Scientz IID). The CNFs were freeze-dried for further use.

    Characterization of chitin nanofibers

    The original chitin and the CNFs of different TEMPO-oxidized times were converted to powder and measured by XRD. The range of diffraction angle (2θ) was from 5° to 60° in an X-ray diffraction (Bruker D2 PHASE, German) that used Co Kα at 30 kV and 10 mA. The crystallinity (ICR, %) was calculated as ICR = (I020 − Iam)/I020 × 100, where I020 is the maximum intensity at about 9.6° and Iam is the intensity of amorphous diffraction at 16° (Zhang et al. 2005). The average size and size distribution of the chitin nanofibers were obtained by dynamic light scattering (DLS, Malvern Zetasizer Nano ZS 90) before 1‰ (m/v) CNF suspensions were filtered through 5 μm mixed cellulose ester (MCE) filtration. The images of CNFs were detected through transmission electron microscopy (MIC-JEM 1200EX) at an accelerating voltage of 120 kV after the dried CNFs were dispersed into 0.5% (m/v) dispersion. CNFs with free carboxylate groups were prepared by immersing them in 0.01 mol/L HCl, repeatedly washing them with deionized water, then drying them at 50 ℃ using a vacuum dryer for FT-IR measurement (Thermo Fisher-Nicolet 6700).

    The dispersion properties were characterized by optical transmittance. The CNFs were dispersed in water suspensions of 1% (m/v). The transmittance of each sample was measured through a UV–Vis spectrophotometer (Shimadzu UV-2550) with the wavelength ranging from 400 to 800 nm (Saito and Isogai 2004). The CNFs were then observed through optical microscopy (BM-1000).

    Determination of carboxylate groups content

    The carboxylate content of the CNFs was determined by the electrical conductivity titration method (Fan et al. 2007). Water (40 ml) was added to the dried CNFs (0.20 g), the mixture stirred for 2 h to prepare a well-dispersed slurry, then 50 mmol/l HCl was added to the mixture to set the pH at 2.5. Next, 50 mmol/L NaOH solution was added at a rate of 0.1 ml/min up to pH 11 using a pH–Stat titration system. The carboxylate content and consumption of NaOH curves were obtained to reflect the content of both carboxylate groups.

    Enzyme immobilization

    The synthesis of NH2-Fe3O4

    Preparation of the amine-functionalized magnetic nanoparticles (NH2-Fe3O4) was done using the same procedure as Mohapatra et al. (2007). FeCl2·4H2O (1.25 g) and FeCl3·6H2O (3.40 g) were dissolved in 100 ml deionized water under nitrogen protection. Ammonium hydroxide (6.00 ml) was added and stirred for 30 min. The system was then magnetically separated and precipitate was washed with deionized water and ethyl alcohol. The magnetic materials (Fe3O4) were vacuum dried for further use. Fe3O4 (1.00 g) was dispersed through a 50% (v/v) ethyl alcohol solution (100 ml) and the pH was adjusted to 4.0 by 0.1 mol/L HCl at 70 ℃. Aminopropyltriethoxysilane (APTES, 3.00 ml) was added and stirred for 12 h under nitrogen atmosphere; then NH2-Fe3O4 was washed with deionized water and ethyl alcohol.

    Immobilization of chymotrypsins and chitin nanofibers

    Enzyme immobilization was performed using the method of Huang et al. (2018). CTs were used for immobilization. CNFs were activated through the EDC/NHS system to covalently immobilize the enzymes onto the surface of the supports. CNFs [1% (m/v)] treated with different oxidation times were dispersed in an MES buffer (100 mmol/L pH 6) containing EDC (10 mg/ml) and NHS (50 mg/ml). The mixture was then shaken for 3 h. The activated CNFs were separated via centrifugation at 13, 000 r/min for 10 min. The water-insoluble part was washed with deionized water and centrifuged repeatedly.

    In previous work, the CTs were immobilized onto CNFs without magnetic nanoparticles; however, the immobilized CTs were aggregated and enzyme activities not detected. Therefore, the CNFs (1 mg) were covalently bonded with NH2-Fe3O4 (0.5 mg) for quick separation. The suspension was shaken for 1 h at 20 ℃. The magnetic chitin nanofibers were separated by external magnets. The citric acid–sodium citrate buffer solution (50 mmol/L pH 5) containing the CTs (500 or 2000 mg/g CNFs) was added to the CNFs. The resulting mixture was incubated for 2 h at 20 ℃ under shaking. The immobilized CTs were separated by external magnets and washed with a citric acid–sodium citrate buffer solution (50 mmol/L, pH 5). The concentration of CTs without immobilization was determined by the Bradford method. The activities of the immobilized enzymes were analyzed using the Lowry–Folin method (Bradford 1976; Kawamura et al. 1981).

    This work was supported by the National Key Research and Development Program of China (no. 2019YFD090 1902), National Natural Science Foundation of China (31922072), China Agriculture Research System (CARS-48), Taishan Scholar Project of Shandong Province (tsqn201812020).

    W-CH and XM conceived and designed the study. RC and WW performed the experiments. RC and W-CH wrote the paper. W-CH and XM reviewed and edited the manuscript. All authors read and approved the manuscript.

    Conflict of interest

    The authors have no conflicts of interest to declare.

    Animal and human rights statement

    This article does not contain any studies with human participants or animals performed by any of the authors.

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