Wen
Liu†
ab,
Liuqin
Huang†
ab,
Rachel
Komorek
c,
Pubudu P.
Handakumbura
a,
Yadong
Zhou
a,
Dehong
Hu
a,
Mark H.
Engelhard
a,
Hongchen
Jiang
b,
Xiao-Ying
Yu
c,
Christer
Jansson
a and
Zihua
Zhu
*a
aEnvironmental Molecular Science Laboratory, Pacific Northwest National Laboratory, Richland, WA 99354, USA. E-mail: Zihua.zhu@pnnl.gov
bState Key Laboratory of Biogeology and Environmental Geology, China University of Geosciences, Wuhan, 430074, China
cEnergy and Environment Directorate, Pacific Northwest National Laboratory, Richland, WA 99354, USA
First published on 7th November 2019
The rhizosphere is arguably the most complex microbial habitat on Earth, comprising an integrated network of plant roots, soil and a highly diverse microbial community (the rhizosphere microbiome). Understanding, predicting and controlling plant-microbe interactions in the rhizosphere will allow us to harness the plant microbiome as a means to increase or restore plant ecosystem productivity, improve plant responses to a wide range of environmental perturbations, and mitigate the effects of climate change by designing ecosystems for long-term soil carbon storage. To this end, it is imperative to develop new molecular approaches with high spatial resolution to capture interactions at the plant-microbe, microbe-microbe, and plant-plant interfaces. In this work, we designed an imaging sample holder that allows integrated surface imaging tools to map the same locations of a plant root-microbe interface with submicron lateral resolutions, providing novel in vivo analysis of root-microbe interactions. Specifically, confocal fluorescence microscopy, time-of-flight secondary ion mass spectrometry (ToF-SIMS), X-ray photoelectron spectroscopy (XPS), and scanning electron microscopy (SEM) were used for the first time for the correlative imaging of the Brachypodium distachyon root and its interaction with Pseudomonas SW25, a typical plant growth-promoting soil bacterium. Imaging data suggest that the root surface is inhomogeneous and that the interaction between Pseudomonas and Brachypodium roots was confined to only a few spots along the sampled root segments and that the bacterial attachment spots were enriched in Na- and S-related and high-mass organic species. We conclude that the attachment of the Pseudomonas cells to the root surface is outcompeted by strong root-soil mineral interactions but facilitated by the formation of extracellular polymeric substances (EPS).
Brachypodium distachyon (Brachypodium) is a widely distributed annual monocot grass that has been proposed as a model organism for grasses, including bioenergy grasses, due to its suitable traits such as small genome, short lifetime, simple growth conditions and amenability to genetic modification.14–17 It has been widely used for many fundamental studies including plant-microbe interactions.18–21Pseudomonas sp. are commonly found in the rhizosphere and many isolates (e.g., Pseudomonas SW25 used in this study22 and P. fluorescens23) have been widely studied as model PGPB. However, the mechanisms of Pseudomonas on promoting plant growth are still under debate and may include pathogen suppression, P release, N fixation and hormonal regulation.23–25 Even less is known about the fundamental principles that control the interactions between plant roots and the Pseudomonas bacteria.
The rhizosphere is a highly heterogeneous system, mainly composed of roots, mineral particles, organic matters and various microbes.1 Most commonly used analysis tools in this field such as total carbon analysis,26 FT-ICR-MS, NMR,27 and genomics28 are bulk analysis techniques, i.e., samples need to be extracted from the system using solvents. Although such approaches have provided key information for developing various models to explain mass transfer in the rhizosphere, they are associated with at least two intrinsic drawbacks. First, these approaches lack spatial information to describe the distribution of microbes or organic matter, e.g., along a root segment. Second, although some organic or bioorganic molecules are soluble, many molecular species may not be soluble and/or be firmly attached to mineral or root surfaces. Scanning electron microscopy (SEM)29 and transmission electron microscopy (TEM)30 have been introduced in the field, providing morphological information with good spatial resolution, albeit with mostly elemental and no or little molecular information. Traditionally, fluorescence microscopy has been widely used in this field, and recently nanoscale secondary ion mass spectrometry (NanoSIMS) has been employed to map the distribution of organic matter on mineral surfaces.31 These techniques can provide high spatial-resolution (down to tens of a nm) chemical maps, although, generally, only selected species with specially labelled fluorescent or isotope tags can be tracked.
Time-of-flight secondary ion mass spectrometry (ToF-SIMS) is a powerful surface analysis tool with several unique advantages.32 First, it can provide elemental, isotopic and molecular information simultaneously. Second, its information depth is very shallow (normally 1–3 nm), so surface-specific information can be collected. In addition, it has excellent sensitivity (ppm level) and very good spatial resolution (sub-micron).33 Therefore, it is a very useful tool in studies of the rhizosphere. Clearly, each technique has its own strength and weaknesses, and a single technique approach normally can provide only limited information. Therefore, a multi-technique approach is highly desirable for comprehensive interrogation of a complex system such as the rhizosphere.
In this work, confocal fluorescence microscopy, ToF-SIMS, X-ray photoelectron spectroscopy (XPS) and SEM were collectively used for the first time to image the Brachypodium root and root-microbe interactions.
Two root segments (about 12–15 mm each) close to the bacterial inoculation areas were selected and excised from each of the I1 and I2 plants. Similarly, two roots from the corresponding areas were selected and cut from each of the U1 and U3 plants. The roots were gently rinsed with deionized water to remove loosely attached soil particles and then immobilized onto a special sample holder (Fig. 1a) that was developed as part of this study. In brief, eight stainless steel pins were used to immobilize four molybdenum (Mo) masks on a ∼5.0 cm diameter aluminum (Al) sample holder (thickness ∼ 6.0 mm). Four Mo masks (12.7 mm diameter and 0.10 mm thick each) were used to press root samples to make them flat. Sample flatness was critical to ensure high quality data from surface analysis tools such as ToF-SIMS and XPS. On each Mo mask, three 5.0 × 1.5 mm2 windows were open for imaging analysis. Such a design allowed us to easily determine accurate locations for multi-imaging analysis. For example, as shown in Fig. 1b, two root samples were immobilized under a Mo mask, and six selected locations, including locations (1) and (2), could be easily located and imaged using different imaging tools.
The initial analysis of fresh root segments was performed by fluorescence microscopy using an upright confocal fluorescence microscope (Zeiss LSM 710) under ambient conditions. The segments were excited using an Ar ion laser with a 488 nm wavelength. The objective was 40× NA 0.75. The fluorescence detection had a wavelength range of 498 nm to 550 nm for GFP. A 3D Z stack was acquired for every sample. The locations of fluorescence images were recorded for subsequent ToF-SIMS, XPS and SEM imaging analysis.
Fluorescence microscopy was followed by ToF-SIMS imaging using a TOF-SIMS5 instrument (IONTOF GmbH, Műnster, Germany). Before SIMS analysis, the sample holder was put into the introduction chamber of the SIMS instrument, and the root samples were dried under vacuum. After drying, the sample holder was introduced into the analysis chamber for SIMS imaging analysis. A 25 keV Bi3+ beam was used as the analysis beam to collect SIMS spectra and images. The Bi3+ beam was focused to be ∼0.5 μm diameter and scanned over 200 × 200 μm2 to 500 × 500 μm2 areas. The current of the Bi3+ beam was about 0.36 pA with 10 kHz pulse frequency, and the data collection time was 600 s per imaging testing. The total ion dose was under the static limit so only surface information (<2 nm) was collected. While collecting data, a low energy electron flood gun (10 eV, ∼1.0 μA current) was used to compensate for surface charging. The pressure in the analysis chamber was about 2 × 10−8 mbar. A positive ion imaging testing and a negative ion imaging testing were performed on each selected location (totally 24 locations, 6 on each Mo mask, as shown in Fig. 1a and b). Due to the complexity of ToF-SIMS spectral data, principal component analysis (PCA)34,35 was used to extract effective information, following procedures described in our previous work.36–38 Because only SIMS signals on the root surface were of interest for PCA analysis, data reconstruction was required. SIMS imaging capability showed its importance here. During data reconstruction, only root surface areas were selected based on SIMS images (e.g., Fig. 1d). Thus, SIMS signals on root surfaces were shown in reconstructed mass spectra for further PCA analysis, and interference signals from the substrate could be removed.
XPS measurements were conducted on the same samples after ToF-SIMS analysis. A Physical Electronics Quantera Scanning X-ray Microprobe was used. This system used a focused monochromatic Al Kα X-ray (1486.7 eV) source for excitation and a spherical section analyzer. The instrument had a 32-element multichannel detection system. The X-ray beam was incident normal to the sample and the photoelectron detector was at 45° off-normal. High energy resolution spectra were collected on root surfaces using a pass-energy of 69.0 eV with a step size of 0.125 eV. For the Ag 3d5/2 line, these conditions produced a FWHM of 0.92 eV ± 0.05 eV. The binding energy (BE) scale was calibrated using the Cu 2p3/2 feature at 932.62 ± 0.05 eV and the Au 4f7/2 feature at 83.96 ± 0.05 eV. The sample experienced variable degrees of charging. Low energy electrons at ∼1 eV, 19 μA and low energy Ar+ ions were used to minimize this charging.
SEM imaging was performed after the XPS measurements. A Hitachi TM-1000 scanning electron microscope (Chiyoda, Tokyo, Japan) was used with an accelerating voltage of 15.0 kV. Before SEM imaging, about 2 nm Au was coated on top of the sample to reduce charging. The imaging areas were ranging from 100 × 75 μm2 to 500 × 375 μm2.
The fluorescence microscopy results prompted the question of how the surface properties of the attachment spots differed from the rest of the root. Here, ToF-SIMS spectra (Fig. 2) can provide valuable chemical information, including elemental, isotopic and molecular information, to elucidate the difference. However, ToF-SIMS spectra are complex, and each spectrum may be composed of hundreds of ion signals. Thus, statistical analysis was used to distinguish features that differentiate the regions with and without Pseudomonas.
Fig. 2 Representative ToF-SIMS spectra of I1, I2, U1 and U3 samples. (a) Negative ion spectra and (b) positive ion spectra. |
Principal Component Analysis (PCA) has been widely used in ToF-SIMS data analysis for over a decade. Fig. 3 shows the PCA analysis results of negative ion spectra collected from the four sets of Brachypodium root samples. The PC1 scores plot (Fig. 3a) revealed that only two spots were clearly separated. It should be noted that after PCA analysis, each spectrum has its own PC1 score value, that is to say, each data spot in Fig. 3a corresponds to a spectrum. Thus, based on the PC1 score values, these two separated data spots are found to be corresponding to the spectra from the locations (1) and (2) of the I2 sample (Fig. 1b). The two separated data spots are close to each other, and the major difference between them and the remaining data spots is the decrease in PC1 scores. The PC1 loadings plot (Fig. 3b) shows that the positive loadings of PC1 are mainly CN and POx-related species, as well as low-mass organic species, while the negative PC1 loadings are mainly Cl and organic SOx species, as well as high-mass organic species. The loadings plot data suggest that the surface of attachment spots for the Pseudomonas cells had relatively more Cl, organic SOx and high-mass organic species, but less CN, POx-related species and low-mass organic species. Moreover, the PC1 scores of the remaining data spots are close together, suggesting that all Pseudomonas-free root surfaces exhibited a similar micro-chemical environment.
Fig. 3 PCA analysis results of negative ion spectra of I1, I2, U1 and U3 samples. (a) PC1 scores plot and (b) PC1 loadings plot. It should be noted that the data spots (1) and (2) in (a) correspond to the two locations (1) and (2) in Fig. 1b, where Pseudomonas attachment was observed. |
Fig. 4 shows the PCA analysis results of positive ion spectra collected from the four sets of Brachypodium root samples. The PCA scores plot (Fig. 4a) is qualitatively consistent with the negative ion results. The PC1 scores of locations (1) and (2) are well separated from the remaining data spots. The loadings plot (Fig. 4b) shows that the major positive loadings of PC1 are Na-related species and high-mass organic species, and the negative loadings are K+, NH4+ and low-mass organic N species. The data suggest that the root surface of the Pseudomonas attachment spots had relatively more Na-related species and high-mass organic species, but less K+, NH4+ and low-mass organic N species.
Fig. 4 PCA analysis results of positive ion spectra of I1, I2, U1 and U3 samples. (a) PC1 scores plot and (b) PC1 loadings plot. It should be noted that the data spots (1) and (2) in (a) are corresponding to the two locations (1) and (2) in Fig. 1b, where Pseudomonas was observed. |
Importantly, PCA analysis results from negative ion spectra and positive ion spectra are consistent with each other. First, PC1 scores could separate Pseudomonas attachment spots from Pseudomonas-free root segments. Second, both positive ion results and negative ion results show that small N-contained organic species are enriched on the Pseudomonas-free root surface, while high-mass organic species are enriched on the surface of Pseudomonas attachment spots.
Fig. 5 shows XPS data from two representative locations on the roots of the I2 and U1 plants. More N, K and Si were observed on the Pseudomonas-free root surface, while more S was observed on Pseudomonas attachment spots. These results are well consistent with the results from ToF-SIMS/PCA analysis. For example, both XPS S spectra and PCA analysis results of ToF-SIMS negative ion spectra show more –SOx groups on the surface of Pseudomonas attachment spots. Such a consistency is very reasonable, because both techniques are surface sensitive, sharing similar information depth (a few nanometers) during analysis. It should be noted that XPS can provide chemical state information, which cannot be obtained from fluorescence, SIMS and SEM analysis. A notable observation in S 2p spectra (Fig. 5b) is two chemical states of S (S6+ and S0/S2−) on the Pseudomonas-free root surface, but only one dominant chemical state of S (S6+) on the Pseudomonas attachment spots. One possible explanation for such an observation is that there was some (∼1 mM) MgSO4 in the growth medium for Pseudomonas, so that some SO42− might stay in the extracellular polymeric substances of the Pseudomonas biofilm.
Fig. 5 High energy resolution XPS data showing the chemical difference between the Pseudomonas-free root surface (U1 plant) and the root surface of Pseudomonas attachment spots (I2 plant, location (1) in Fig. 2b). |
Fig. 6 displays the SEM images of roots with and without Pseudomonas attachment. The root surface with Pseudomonas attachment looks smoother, almost free of soil particles. In contrast, many small soil particles were observed on the Pseudomonas-free root surface. This situation is in agreement with the ToF-SIMS and XPS data that there are more Si and K (from soil mineral particles) on the Pseudomonas-free root surface.
Fig. 6 SEM images of root surfaces obtained after ToF-SIMS measurements. (a) Root surface from the I2 plant at location (1) with Pseudomonas attachment shown in Fig. 1b. (b) Pseudomonas-free root surface on a root segment from the U1 plant. |
The above SEM observation is very interesting. From the literature, biofilms are usually implicated in aggregation processes because soil particles stick to them.39 If so, a possible explanation for the above observation is that Pseudomonas could only attach on soil particle-free areas. However, SEM images show that all root surfaces from U1 and U3 are with some considerable amount of soil particles, indicating that soil particle-free root surfaces are rare before Pseudomonas treatment. If so, another possibility is that Pseudomonas attachment might reduce direct interactions between root and soil particles.
It is notable that Pseudomonas attachment was observed only on two locations along the four root segments from two Pseudomonas-inoculated Brachypodium plants (I1 and I2), i.e., locations (1) and (2) of the I2 plant. One possible explanation for such an observation is that the Pseudomonas treatment process was not uniform. However, this is doubtful since a large amount of Pseudomonas bacterial solution was added to the soil around the I1 and I2 plants, and root segments for imaging were selected close to the inoculation site. A more plausible explanation is that direct interactions between Pseudomonas cells and Brachypodium roots is weak and outcompeted by root-mineral interactions (Fig. 7a). Such a weak interaction between Pseudomonas cells and Brachypodium roots has been confirmed by a separate research study in our lab.40 In that work, the Brachypodium roots grew in liquid media with glass beads (not in soil), in which Pseudomonas seemed to have difficulty in attaching to the free root surface in the media, but could aggregate on the glass bead-root interface.40 Therefore, we tend to believe that Pseudomonas may produce extracellular polymeric substances to form a biofilm on a small amount of root surfaces and that the biofilm can physically separate roots and soil particles (Fig. 7b).
Footnote |
† These authors equal contribution. |
This journal is © The Royal Society of Chemistry 2020 |