A fluorescence nanoparticles-based method for visualizing and quantifying root cap mucilage

Translated Abstract

The root cap mucilage, a gel-like substance secreted by border cells or border-like cells at the root cap, plays a crucial role in plant root adaptation to soil environments. Despite its importance, methods for analyzing root cap mucilage, especially visualization techniques, remain limited. This study introduces a simple method using fluorescent silica nanoparticles (Quartz Dot) to visualize and quantify the area of secreted mucilage with confocal laser scanning microscopy. The nanoparticles used in this study are embedded with fluorescent probes and large enough to be excluded from the apoplastic mucilage matrix, thereby outlining the boundary of the mucilage region surrounding the root cap. Combined with fluorescent probes for cell wall staining, this approach allows the quantification of the minute mucilage areas surrounding the Arabidopsis root cap. The method was validated by comparing the mucilage region in wild-type Arabidopsis and the smb brn1 brn2 mutant, revealing a significant reduction in mucilage areas in the mutant. Additionally, we investigated the effects of several abiotic stresses, including toxic ions and nutrient starvation, on the area of Arabidopsis root cap mucilage, and further identified soil stresses that promote expansion of the root cap mucilage region, underscoring the sensitivity and precision of our method. Collectively, this study established a simple and robust technique for visualizing and quantifying the root cap mucilage region. The demonstrated utility of this method indicates its potential for broader applications in understanding the physiological and molecular mechanisms underlying root cap functionalization and its role in adaptation to soil environments.

INTRODUCTION

The tips of plant roots are commonly covered by a dome-shaped tissue known as a root cap. The root cap plays a critical role in constructing an interface between the root and the soil environment through its specialized functional properties (Ganesh et al. 2022). As a part of these functions, the outermost cells of the root cap are continuously released into the soil. These cells, called border cells (or border-like cells in Brassicaceae species), secrete gelatinous substances known as root cap mucilage (Driouich et al. 2021). This cell release occurs in a sustained manner to ensure consistent coating of the root tip with the mucilage. Root cap mucilage, along with released root cap cells, has been reported to have various functions in root adaptation to soil environments. For example, the root cap mucilage protects biotic stress by forming a protective shield against pathogenic microbes and nematodes (Zhao et al. 2000, Driouich et al. 2013, Castilleux et al. 2018). The root cap mucilage also preserves root integrity against abiotic stresses, including drought (Aslam et al. 2022), aluminum (Ryan et al. 1995, Miyasaka and Hawes 2001, Watanabe et al. 2008, Cai et al. 2013, Nagayama et al. 2019), salt (Edmond Ghanem et al. 2010), and heavy metals (Morel et al. 1986, Huskey et al. 2018). Therefore, the regulation of mucilage formation and secretion at the root cap plays a critical role in plant adaptation.

The root cap mucilage is mainly composed of pectic polysaccharides and proteoglycans (Chaboud and Rougier 1984, Bacic et al. 1986, Amicucci et al. 2019, Nazari 2021). It also contains non-polysaccharide molecules, such as extracellular DNA and antimicrobial proteins and peptides (Wen et al. 2009, Ma et al. 2010, Driouich et al. 2013, Weiller et al. 2017, Pozzo et al. 2018). Recent studies have uncovered molecular mechanisms underlying the root mucilage formation, including polysaccharide biosynthesis, modification (Donohoe et al. 2013, van de Meene et al. 2017, Amicucci et al. 2019), and secretion mediated by intracellular trafficking (Gunning and Steer 1996, Wang et al. 2017, Wang and Kang 2018, Liu et al. 2024). Furthermore, recent studies in Arabidopsis thaliana (hereafter Arabidopsis) revealed mechanisms that link mucilage secretion and root cap cell separation (Durand et al. 2009, Maeda et al. 2019, Liu et al. 2024), highlighting the tight regulation of mucilage formation by the developmental program involving root cap morphogenesis. In addition, in various plant species other than Arabidopsis, environmental stimuli, such as biotic and abiotic stresses, also alter the secretion of root cap mucilage (Hawes et al. 2000, Miyasaka and Hawes 2001, Wen et al. 2007, Huskery et al. 2018). These findings suggest that the formation and secretion of the root cap mucilage are regulated through the integration of developmental programs and environmental signals.

To further dissect the mechanisms underlying the formation and secretion of the root cap mucilage, advanced imaging techniques are required for simultaneously capturing the dynamics of apoplastic mucilage in relation to morphogenesis and gene expression in living root cap cells. In particular, fluorescence imaging using confocal laser scanning microscopy is expected to serve as a powerful tool for quantifying the elaborate dynamics of root cap mucilage in Arabidopsis. The conventional method of visualizing the outline of root cap mucilage on the basis of the non-permeability of the colloidal particles of India ink, is incompatible with fluorescence imaging (Maeda et al. 2019). To achieve a breakthrough, we here established a quantitative imaging method using fluorescently labelled silica nanoparticles for Arabidopsis roots.

MATERIALS AND METHODS

Plant materials and growth conditions

Arabidopsis thaliana (L.) Heynh. accession Col-0 was used as the wild type. The sombrero bearskin1 bearskin2 (smb brn1 brn2) triple mutant was reported previously (Bennet et al. 2010, Kamiya et al. 2016, Willemsen et al. 2008). Arabidopsis seeds were allowed to germinate on plates containing 0.5× Arabidopsis nutrient solution (Haughn and Somerville 1986) supplemented with 1% (w/v) sucrose and 1% (w/v) agar and adjusted to pH 5.8 (hereafter called standard medium). Plants were grown at 23°C with 16 h light/8 h dark cycles. For abiotic stress treatment, three days-after-germination plants were transferred to various stress media prepared by modifying the aforementioned standard medium as described below, and root tips were observed 48 h after the transfer. For aluminum stress, AlCl₃ was added to the standard medium at the final concentration of 50 μM, and pH was adjusted to 4.3. For salt stress, NaCl was added to the standard medium at the final concentration of 100 mM. For phosphate-starved stress, K₃PO₄ concentration (1.25 mM in the standard medium) was reduced to 10 µM. For sulfate-starved stress, MgSO₄ concentration (1.0 mM in the standard medium) was reduced to 100 µM.

Imaging and image quantification

For confocal microscopy, dissected root tips were mounted with a solution containing 1.0 µg/mL Calcofluor White (Fluorescent Brightener 28, MP Biomedicals, LLC, Irvine, CA, USA) and 1 mg/mL Quartz Dot QD-GO (300 nm particle, Ex 514 nm/Em 519–540 nm, Furukawa Electric Advanced Engineering, Chiba, Japan), or a mixture of 50 µg/mL propidium iodide and 1 mg/mL Quartz Dot QD-VG (300 nm particle, Ex 405 nm/Em 438–506 nm, Furukawa Electric Advanced Engineering). Confocal microscopy observation was performed using Leica Stellaris 5 equipped with five lasers (50-mW diode laser at 405 ± 3 nm, 40-mW diode laser at 448 ± 3 nm, 20-mW solid state lasers at 488 ± 1 nm, 514 ± 2 nm, and 561 ± 1 nm). For Figure 1A, all lasers were used at 2.5% of full laser power and emission wavelength for each excitation was mentioned in the figure. All images were taken using a 60x water-immersion objective lens unless noted otherwise. Microscope images were edited and analyzed using ImageJ. Quantification of the root cap mucilage area was conducted as follows (Figure 1B–D). First, the central longitudinal axis of the root cap was defined by connecting the distal tip of the root cap and the center of the quiescent center (QC). A horizontal line was delineated perpendicular to the central axis and at a distance of 200 μm from the root cap tip. Quantification was performed in the area distal to the horizontal line. Outline of the root cap and the boundary between the regions with and without the fluorescent silica nanoparticles were traced using polygons and used to calculate the area devoid of the fluorescent particle to quantify the area of root cap mucilage. For time-lapse imaging, fluorescent images were captured using a DsRed filter and a Leica K8 CMOS camera.

RESULTS AND DISCUSSION

The root cap mucilage, with its gel-like properties, prevents the penetration of large particles. Taking advantage of this property, India ink, whose colloidal particles are approximately 100 nm or larger in diameter, is widely used to trace the edge of the root cap mucilage region surrounding the root tip (Madsen et al. 1992, Taban et al. 2005). Using a similar approach, we employed a commercially available fluorescent silica nanoparticle Quartz Dot (QD) as an alternative for fluorescent-based observation. These silica nanoparticles are comparable in size to India ink colloids and are embedded with a fluorescent probe, whereby allowing visualization by confocal laser scanning microscopy. We tested two types of QDs: QD-VG and QD-GO, which incorporate the fluorescent dyes Fluolid-N-green and Rhodamine 6G, respectively. QD-VG, excitable by a 405-nm and 448-nm lasers, is compatible with propidium iodide staining of cell walls and with red-fluorescent protein such as RFP and its derivatives (Figure 1A). In contrast, QD-GO is efficiently excitable by a 514-nm laser and is compatible with Calcofluor White staining of cell walls (Figure 1A). Also, since QD-GO could be excited by a 560-nm laser, it allows for simultaneous observation with green-fluorescent proteins, such as GFP and its derivatives. To assess whether the QDs could be used to simultaneously visualize the root cap mucilage region and the root cap cell walls as intended, we applied a solution containing QD-VG and propidium iodide to the root tip of Arabidopsis primary roots (Figure 1B, C). As expected, we could observe regions around the root tips where QD-VG particles were excluded, resembling the pattern observed with India ink. By tracing the regions where the nanoparticles were excluded, we quantified the area of apoplastic mucilage surrounding the root cap (Figure 1D). In addition, we performed time-lapse imaging to visualize fluorescent silica nanoparticles suspended around the root. The nanoparticles remained in suspension but were excluded from a distinct microregion covering the root cap surface (Supplementary Movie 1). This result further supports the presence of a distinct apoplastic layer surrounding the root cap, which corresponds to root cap mucilage.

Figure 1 Fluorescent nanoparticle-based methods for visualizing the root cap mucilage of Arabidopsis. (A) Fluorescence profiles of two types of fluorescent silica nanoparticles, Quartz Dot (QD): QD-VG and QD-GO, with various excitation and emission filter sets. QD particles are visualized as colored dots. Scale bars = 10 μm. (B–D) Visualization and quantification of root cap mucilage region using QDs. Confocal images of an Arabidopsis root tip stained with propidium iodide (B), QD-VG particles surrounding the root tip (C), and their merged image with a magnification (D). The central axis was defined by connecting the root tip and the QC. A horizontal line was then drawn 200 μm from the tip to establish the quantification area (D). Tracing of the region devoid of QDs for the quantification of root tip mucilage area is shown (D). Scale bars = 100 μm.

In Arabidopsis, two paralogous NAC-type transcription factors, BEARSKIN1 (BRN1) and BRN2, play a key role in the root cap mucilage formation in a genetically redundant manner (Maeda et al. 2019, Liu et al. 2024). It has been reported that brn1 brn2 double mutants show a reduction in root cap mucilage. SOMBRERO (SMB) is another NAC-type transcription factor that controls root cap cell differentiation and cell removal in a partially redundant manner with BRN1 and BRN2 (Bennett et al. 2010, Fendrych et al. 2014, Kamiya et al. 2016). To further evaluate the utility of this QD-based method, we here compared the mucilage areas between the wild type and the smb brn1 brn2 mutant. Silica nanoparticles were distributed closer proximity to the root cap cells of smb brn1 brn2 than those of the wild type (Figure 2A, B). Impenetrable areas of the fluorescent silica nanoparticles were significantly smaller in smb brn1 brn2 triple mutant, as previously reported for brn1 brn2 double mutants (Maeda et al. 2019). These data demonstrate that fluorescent silica nanoparticles, like India ink, are unable to penetrate the gelled root cap mucilage region, highlighting the reliability of the nanoparticles in visualizing the root cap mucilage with confocal microscopy.

Figure 2 Root cap mucilage regions of wild-type (Col, A) and smb brn1 brn2 triple mutant roots (B). Magenta dots represent the fluorescence of QD-GO particles. Cyan color indicates the root tip cells stained with Calcofluor White. Scale bars = 50 μm. (C) Quantification of the mucilage area for each genotype. The mucilage area in smb brn1 brn2 was significantly smaller than that of the wild type (p < 0.01, t-test; wild type, n = 4; smb brn1 brn2, n = 5).

In various plant species, the root cap mucilage has been reported to expand in response to the stresses derived from soil environments (Edmond Ghanem et al. 2010, Cai et al. 2011, Cai et al. 2013, Huskey et al. 2018). Aluminum (Al) is one of the major abiotic stresses in soil environments. Ionized aluminum (Al³⁺) causes cellular damage to roots, resulting in stunted root growth. In addition, Al³⁺ readily binds with phosphate to form insoluble aluminum phosphate complexes, reducing the available amount of phosphate for plant growth. Previous studies have reported that root caps cells promote mucilage production in response to Al in some plant species, including rice (Oryza sativa), snap bean (Phaseolus vulgaris), and soybean (Glycine max) (Miyasaka and Hawes 2001, Cai et al. 2011, Cai et al. 2013, Nagayama et al. 2019). While Al response has been extensively analyzed in Arabidopsis, such as its perception and related cell signaling mechanisms (Rounds and Larsen 2008, Sawaki et al. 2009, Liu et al. 2016, Zhou et al. 2023, Cao et al. 2024, Ding et al. 2024), there are no studies linking root cap mucilage to the aluminum stress in Arabidopsis. Therefore, using the fluorescent silica nanoparticles, we next investigated how root cap mucilage in Arabidopsis responds to excessive Al. We found that the root cap mucilage area was significantly larger under Al stress than under control conditions. (Figure 3A, B). This result suggests that, similar to other plant species, the Arabidopsis root cap promotes mucilage expansion upon excessive Al. By contrast, such mucilage expansion was not observed under phosphate or sulfate starvation (Figure 3D, E), suggesting that the expansion of the mucilage region under the Al stress is due to the Al toxicity rather than Al-mediated phosphate sequestration. We also found that salt stress induces the expansion of the root cap mucilage region in Arabidopsis (Figure 3C). These observations suggest that the Arabidopsis root cap, like those of other reported plant species, increases the root cap mucilage region in response to varying levels of soil minerals such as Al and sodium.

Figure 3 Quantification of root cap mucilage regions of wild-type Arabidopsis roots under various abiotic stress conditions. (A–E) Three day-after-germination plants grown on the standard media were transferred to a fresh standard media (control, A), aluminum stress media (B), salt stress media (C), phosphate-starved media (D), and sulfate-starved media (E). Root tips were observed 48 h after the treatments. Magenta dots represent the fluorescence of QD-GO particles. Cyan color indicates the root tip cells stained with Calcofluor White. White dashed lines indicate the boundary of the root cap mucilage region. Scale bars = 50 μm. (F) Quantification of the mucilage areas surrounding the root tips under designated abiotic stress condition. Statistical analysis was performed using Tukey’s HSD test (aluminum stress, n = 5; other conditions, n = 4).

PERSPECTIVE

Here, we developed a simple method for visualizing and quantifying the root cap mucilage area using the commercially available fluorescently labeled silica nanoparticles, Quartz Dot. By utilizing two types of silica nanoparticles with distinct fluorescence properties, our method allows simultaneous observation with cell wall stains and possibly with fluorescent proteins. We demonstrated its capability to quantitatively analyze changes in the root cap mucilage region in response to various environmental stimuli in Arabidopsis, a species known for its minute mucilage region at the root tip. The data presented here highlight the sensitivity and precision of our method. Further integration of molecular biology tools, such as transcriptome analysis and genome editing, will significantly improve our understanding of physiological functions of root cap mucilage and molecular mechanisms underlying its formation. Given the association between the root cap mucilage formation and developmental processes involving root cap cell separation, our method will facilitate integrative studies combining developmental biology and environmental stress responses. Such studies could provide a comprehensive framework to uncover the multifaceted roles of root cap mucilage in plant root adaptation and development.

Acknowledgments

This work was supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan and Japan Society for the Promotion of Science to H.S. (23KJ1580), R.S. (24K09142), K.N. (19H05671 and 19H03248), and S.M. (20H03277 and 21H05152); the Fukushima Institute for Research, Education and Innovation to R.S. (JPFR24040101); the Japan Science and Technology Agency to S.M. (PRESTO [JPMJPR20D6]); MEXT Promotion of Distinctive Joint Usage/Research Center Support Program (JPMXP0723833155) to S.M.; Takeda Science Foundation to S.M.

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