This study was co-funded by the Austrian Science Fund (FWF): P30085-N28 (Thomas Prohaska) and the Austrian Science Fund (FWF) and the Federal State of Lower Austria: P27571-BBL (Jakob Santner).

The authors have nothing to disclose.

The solute imaging protocol presented here is a versatile method to visualize and quantify 2D nutrient and contaminant fluxes in soil-plant environments. It is unique in its capability to generate sub-mm scale multi-element images of labile solute species at the soil-root interface, exceeding the achievable spatial resolution of alternative methods for measuring solute gradients in the rhizosphere substantially4. The targeted in situ sampling approach of DGT, in combination with a highly sensitive chemical analysis method such as LA-ICP-MS, facilitates the detailed investigation of solute flux dynamics around individual plant roots grown in soil or similar substrates. Due to the sink-based sampling process, the obtained images reflect the lability of the visualized solutes, and therefore are an estimation of their plant-availability10. Although the method-inherent measurement of solute fluxes bears considerable advantages like the interpretability as plant-available nutrient fractions, flux measurements are much less straight-forward to understand than porewater concentration measurements. The standard DGT sampling geometry in bulk soil applications (specifically the 0.8 mm-thick diffusion gels used in that setup) allows for comparing the actual porewater concentration, csoln, and a time-averaged porewater concentration estimate by a bulk DGT measurement, cDGT, and for the interpretation of these parameters regarding the resupply dynamics of a solute species. However, such comparison cannot be done based on imaging DGT application with very thin diffusion layers, as the derived cDGT values are unrealistically small34. DGT imaging results are therefore not always simple and quick to interpret and are often not directly comparable to more conventional porewater concentration measurements.
When applying the method, a few critical steps need to be carefully considered, mainly related to filling and watering the rhizotron growth containers. During filling the soil into the rhizotron, it is very important to avoid compacting the soil too much, as the plant roots cannot penetrate strongly compacted soil and root growth will be inhibited. We have observed roots avoiding strongly compacted soil and growing along the inner edges of the rhizotron growth container, where the soil is usually less compacted. In this case, individual roots located in the center of the rhizotrons, where DGT gels can be applied conveniently, may not develop at all, effectively inhibiting successful gel application. In our laboratory, experience showed that dry soil bulk densities of 1.0-1.4 g cm-3 allow unimpeded root development. Moreover, excessive soil compaction is also a potential source of artefacts regarding the solubility of redox-sensitive elements and biogeochemically associated species. As the total pore volume is reduced and the pore diameter distribution is shifted towards lower diameters in highly compacted soil, less air-filled larger-diameter pore volume is available, which may lead to reductive conditions locally. Consequently, MnIII/IV- and FeIII-oxides may be reduced, leading to increased Mn2+ and Fe2+ fluxes. The dissolution of Fe-oxides, which are important sorption sites e.g., for phosphate and micronutrients, may liberate sorbed and/or co-precipitated species and thereby cause artificially elevated fluxes of the biogeochemically associated species. A similar issue may arise if the growth containers are watered too much. Evaporation via the small soil surface area at the top of the growth container is low and the soil may remain water-saturated for up to several weeks after planting, which may also cause redox artefacts.
Another important consideration is the chemical functionality of the fabricated HR-DGT binding gel. By following the protocol, thin gels with a homogeneous distribution of binding phases are obtained. If the gels have areas of inhomogeneous material distribution (e.g., holes in the gel or aggregates of binding phases) these areas need to be removed or, if too extensive, the gel fabrication protocol needs to be repeated. If prepared correctly, the gel must be able to bind the target solute species that diffuse into the gel immediately and quantitatively27, which is determined by the analyte-specific gel binding capacity. While exceeding the gel capacity is less problematic in uncontaminated soils, it should be considered in metal-contaminated soils and saline soil environments. Saturation of the gel binding phases will not only impair quantitative solute sampling, but also result in lateral diffusion of solutes between binding phases in the gel, leading to an indefinite localization of small-scale solute flux features. Thus, if very high quantities of labile nutrient/contaminant species are expected in the target soil environment, preliminary tests should be performed. For estimating expected DGT loadings, bulk soil DGT piston sampling followed by gel elution and wet-chemical analysis can be applied15,49. If necessary, DGT deployment times may be adjusted to reduce the gel contact time and thus avoid gel saturation above capacity thresholds. Conversely, preliminary tests can also be helpful to identify required gel contact times and/or LA-ICP-MS sensitivities if very low solute loadings are expected, which may be important for mapping trace element solutes at natural soil background levels15. Besides, correct DGT gel functioning should be verified before its experimental application via controlled loading of gels in the preparation of DGT LA-ICP-MS calibration standards. The gel standard provides a matrix-matched reference gel analyte loading that can be used to assess if the sample gel loading determined by LA-ICP-MS is within the expected range. If unable to obtain a signal which is different from the gas and method blank background noise, the operator must ensure that laboratory procedures for trace element analysis were implemented and all protocol steps were performed correctly. Sometimes, the DGT gel is accidentally flipped after solute sampling with the soil-exposed, loaded side facing towards the glass plate rather than the laser beam, resulting in low signal intensities and erroneously flipped features in the final solute flux images.
During the LA-ICP-MS analysis, a large quantity of data is generated, which takes considerable time to evaluate. In our lab, we use in-house data evaluation scripts tailored for our target data output format using standard spreadsheet software. After semi-automated sorting and calibration, image plotting is conducted using open source, open access image analysis tools (ImageJ, Fiji50). This approach allows full control over data sorting, evaluation, and presentation, which is essential because the collected data correspond to rectangular, and not quadratic pixels, which needs to be properly displayed in the generated solute maps. Moreover, during data processing, any pixel interpolation should be carefully avoided. Pixel interpolation leads to smoothed gradients in the chemical images, resulting in softened, often circular element distribution features and is therefore an undesirable alteration of the original data. Pixel interpolation is a standard procedure in re-scaling and re-formatting operations in many image processing software products but can be deselected usually.
In conclusion, the described method is a significant advancement for understanding nutrient and contaminant dynamics in natural soil-rhizosphere-plant systems. In addition to DGT-only applications, the method can be combined with other, diffusion-based imaging techniques like planar optodes3,33,42,43,48,51 and zymography20,21,22,23,24, and may be developed further for including additional elements and soil parameters.

Nutrient acquisition by crop plants is a key factor in determining crop productivity. The processes governing efficient uptake of nutrients by crops have been studied intensely, especially the mechanisms controlling nutrient availability to and nutrient internalization by plant roots at the soil-root interface, the rhizosphere, are recognized for their role in crop nutrient acquisition. Important processes for plant nutrient uptake include: nutrient transport towards the root; dynamic sorption equilibria between species dissolved in the soil porewater and species bound to solid soil surfaces; microbial competition for nutrients; microbial mineralization of nutrients that are contained in soil organic matter; and nutrient internalization into the root symplasm. The uptake of inorganic trace metal(oid) contaminants is largely controlled by the same mechanisms.
Depending on nutrient and contaminant availability, plant demand and diffusivity in soil, differential nutrient patterns in the rhizosphere can be observed. For strongly sorbing elements with comparatively high internalization rates (e.g., P, Fe, Mn, Zn, As, Cd, Pb), depletion of the labile (i.e., reversibly adsorbed) element fraction compared to the bulk soil is found, with depletion zone widths often being &#8804;1 mm, while for more mobile nutrients such as NO3-, depletion zones can extend up to several centimeters1. Moreover, accumulation of elements such as Al and Cd has been observed when availability exceeds plant uptake rates2,3.
Given the importance of rhizosphere processes in nutrient and contaminant cycling, several techniques for measuring the plant-available element fraction at high spatial resolution have been developed4,5. However, measuring small-scale labile solute distributions has proven to be challenging for several reasons. A major difficulty is to sample very small (low &#181;L range) volumes of soil and/or porewater at defined positions adjacent to living plant roots to resolve the steep nutrient gradients in the rhizosphere. One approach to address this problem is to use micro-suction cups for the extraction of porewater samples6. With this method, A. G&#246;ttlein, A. Heim and E. Matzner7 measured soil porewater nutrient concentrations in the vicinity of Quercus robur L. roots at a spatial resolution of ~1 cm. A difficulty of analyzing &#181;L volumes of soil or soil solution is, that these small sample volumes, in combination with the low concentrations of all but the major nutrient species, require highly sensitive chemical analysis techniques.
An alternative system, capable of resolving nutrient gradients at a resolution down to ~0.5 mm, is to grow a root mat on the surface of a soil block, with a thin hydrophilic membrane layer separating soil from the roots8,9. In this configuration, solutes can pass through the membrane and roots can take up nutrients and contaminants from the soil while root exudates can diffuse into the soil. After the establishment of a dense root layer, the soil block can be sampled and sliced to obtained soil samples for subsequent extraction of element fractions. In this way, one-dimensional nutrient, and contaminant gradients, averaged across a relatively large area (~100 cm2) can be analyzed.
A further challenge is to obtain samples of the labile, plant-available element fraction, since most chemical soil extraction techniques operate very differently compared to the mechanisms by which plants take up nutrients and contaminants. In many soil-extraction protocols, soil is mixed with an extractant solution with the aim to establish a (pseudo-)equilibrium between dissolved and sorbed element fraction. However, plants continuously internalize nutrients and, therefore, often progressively deplete the rhizosphere soil. Although equilibrium extraction protocols have been widely adopted as soil tests as they are easy to implement, the extracted nutrient fraction often does not represent the plant-available nutrient fraction well10,11,12,13. Sink methods which continuously deplete the sampled soil for nutrients have been proposed as advantageous methods and may better resemble the underlying nutrient uptake mechanism by mimicking the root uptake processes10,11,14,15.
In addition to the methods described above, genuine imaging applications, capable of measuring continuous parameter maps with resolutions &#8804;100 &#181;m across fields of view of several cm2 have been developed for specific elements and soil (bio)chemical parameters5. Autoradiography can be used to image the element distribution in the rhizosphere provided that suitable radioisotopes are available16. Planar optodes enable visualization of important soil chemical parameters such as pH and pO217,18,19, and enzyme activity or total protein distributions can be mapped using fluorescent indicator imaging techniques such as soil zymography20,21,22,23 and/or root blotting methods24. While zymography and autoradiography are limited to the measurement of a single parameter at a time, pH and pO2 imaging using planar optodes can be done concurrently. The more traditional root mat techniques provide 1D information only, while micro suction cups provide point measurements or low resolution 2D information, however both approaches allow for multi-element analysis. More recently, P. D. Ilhardt, et al.25 presented a novel approach using laser induced breakdown spectroscopy (LIBS) to map 2D total multi-element distributions at a resolution of ~100 &#181;m in soil-root core samples where the natural element distribution was preserved by careful sample preparation.
The only technique capable of targeted 2D sampling of multiple nutrient and contaminant solutes at high spatial resolution is the diffusive gradients in thin films (DGT) technique, a sink-based sampling method that immobilizes labile trace metal(loid) species in situ on a binding material embedded in a hydrogel layer26,27. DGT was introduced as a chemical speciation technique for measuring labile solutes in sediments and waters, and was soon adopted for its use in soils28. It enables sub-mm scale multi-element solute imaging, which was initially demonstrated in a river sediment29, and has been developed further for its application in plant rhizospheres30,31,32,33.
For DGT sampling, a gel sheet of a size of approximately 3 cm x 5 cm is applied onto a single plant root that is growing in the surface layer of a soil block, with a hydrophilic membrane separating the gel from the soil. During the contact time, labile nutrients and/or contaminants diffuse towards the gel and are bound immediately by the binding material incorporated in the gel. In this way, a concentration gradient, and thus a continuous net flux towards the gel is established and prevailed during the sampling time. After sampling, the hydrogel can be removed and analyzed using an analytical chemical technique allowing for spatially resolved analysis. A highly specialized and frequently used technique for this purpose is laser ablation inductively coupled plasma mass spectrometry (LA-ICP-MS). In some early studies, micro particle induced X-ray emission (PIXE) was also used29. DGT sampling combined with LA-ICP-MS analysis allows for multi-element chemical imaging at a spatial resolution of ~100 &#181;m. If highly sensitive ICP-MS techniques (e.g., sector field ICP-MS) are employed, exceptionally low limits of detection can be achieved. In a study on the effect of liming on Zn and Cd uptake by maize15, we were able to map labile Cd in the maize rhizosphere in uncontaminated soil with a limit of detection of 38 pg cm-2 of Cd per gel area. DGT, planar optodes, and zymography rely on diffusion of the target element from soil into a gel layer, which can be exploited for combined application of these methods in order to simultaneously, or consecutively, image a large number of parameters relevant for plant nutrient and contaminant uptake. Detailed information on analytical chemical aspects of DGT imaging, on the potential of combining DGT and other imaging methods, and on its applications is comprehensively reviewed in ref.34,35.
In this article we describe how to carry out a solute imaging experiment using the DGT technique on roots of terrestrial plants in a unsaturated soil environment, including plant cultivation, gel fabrication, gel application, gel analysis and image generation. All steps are elaborated in detail, including notes on critical steps and experimental alternatives.

<table>
	<tbody>
		<tr>
			<td>(NH4)2S2O8 (ammonium persulfate; APS)</td>
			<td>VWR</td>
			<td>21300.260</td>
			<td>&#8805;98.0%, analytical reagent</td>
		</tr>
		<tr>
			<td>2-(N-morpholino)-ethanesulfonic acid (MES)</td>
			<td>Sigma-Aldrich</td>
			<td>M8250-100G</td>
			<td>&#8805;99.5%</td>
		</tr>
		<tr>
			<td>Acrylamide solution</td>
			<td>Sigma-Aldrich</td>
			<td>A4058-100ML</td>
			<td>40%, for electrophoresis</td>
		</tr>
		<tr>
			<td>Analyte salts</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Use water soluble analyte salts of analytical grade or higher</td>
		</tr>
		<tr>
			<td>Buechner funnel</td>
			<td>VWR</td>
			<td>511-0065</td>
			<td>13 cm plate diameter</td>
		</tr>
		<tr>
			<td>Chemical equilibrium modelling software</td>
			<td>KTH Sweden</td>
			<td>n/a</td>
			<td>Visual MINTEQ</td>
		</tr>
		<tr>
			<td>Clamp</td>
			<td>Local warehouse</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Desktop publishing software</td>
			<td>Adobe Inc.</td>
			<td>n/a</td>
			<td>InDesign CS6</td>
		</tr>
		<tr>
			<td>DGT cross-linker</td>
			<td>DGT Research Ltd</td>
			<td>n/a</td>
			<td>2%, agarose derivative</td>
		</tr>
		<tr>
			<td>DGT piston sampler</td>
			<td>DGT Research Ltd</td>
			<td>n/a</td>
			<td>2 cm diameter exposure window</td>
		</tr>
		<tr>
			<td>Digital single-lens reflex (DSLR) camera</td>
			<td>Canon Inc.</td>
			<td>n/a</td>
			<td>Canon EOS 1000D</td>
		</tr>
		<tr>
			<td>Dispersion device</td>
			<td>IKA</td>
			<td>3737000</td>
			<td>Ultra-Turrax T10 Basic</td>
		</tr>
		<tr>
			<td>Double-sided adhesive tape</td>
			<td>Tesa</td>
			<td>56171</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sigma-Aldrich</td>
			<td>34923</td>
			<td>Puriss. p.a., absolute, &#8805;99.8%</td>
		</tr>
		<tr>
			<td>Gel blotting paper</td>
			<td>Whatman</td>
			<td>10426981</td>
			<td>Blotting Papers, Grade GB005, 20 &#215; 20 cm, 1.5 mm thickness</td>
		</tr>
		<tr>
			<td>Gel drier</td>
			<td>UniEquip</td>
			<td>n/a</td>
			<td>UNIGELDRYER 3545</td>
		</tr>
		<tr>
			<td>High-pressure microwave system</td>
			<td>Anton Paar</td>
			<td>n/a</td>
			<td>Multiwave 3000</td>
		</tr>
		<tr>
			<td>HNO3</td>
			<td>VWR</td>
			<td>1.00456.2500P</td>
			<td>65%, ISO for analysis</td>
		</tr>
		<tr>
			<td>Horizontal shaker</td>
			<td>GFL</td>
			<td>305</td>
			<td></td>
		</tr>
		<tr>
			<td>HydroMed D4</td>
			<td>AdvanSource Biomaterials Corp.</td>
			<td>n/a</td>
			<td>Ether-based hydrophilic urethane</td>
		</tr>
		<tr>
			<td>ICP-MS software</td>
			<td>Perkin Elmer</td>
			<td>n/a</td>
			<td>Syngistix</td>
		</tr>
		<tr>
			<td>Image analysis software</td>
			<td>National Institutes of Health (NIH)</td>
			<td>n/a</td>
			<td>ImageJ Fiji, freely available at https://fiji.sc/</td>
		</tr>
		<tr>
			<td>Knife-coating device</td>
			<td>BYK</td>
			<td>5561</td>
			<td>Single Bar 6&#8243;, 0.5 mils</td>
		</tr>
		<tr>
			<td>LA software</td>
			<td>Elemental Scientific Lasers</td>
			<td>n/a</td>
			<td>ActiveView</td>
		</tr>
		<tr>
			<td>LA system</td>
			<td>Elemental Scientific Lasers</td>
			<td>n/a</td>
			<td>NWR193</td>
		</tr>
		<tr>
			<td>Laminar flow bench</td>
			<td>Telstar Laboratory Equipment B.V.</td>
			<td>n/a</td>
			<td>Class II biological safety cabinet</td>
		</tr>
		<tr>
			<td>Magnetic stirrer</td>
			<td>IKA</td>
			<td>0003582400</td>
			<td>C-MAG MS 7</td>
		</tr>
		<tr>
			<td>Moisture-retaining film</td>
			<td>Bemis Company, Inc.</td>
			<td>PM999</td>
			<td>Parafilm M, 4&#34; x 250&#39;</td>
		</tr>
		<tr>
			<td>N,N,N&#8217;,N&#8217;-tetramethylethylenediamine (TEMED)</td>
			<td>Sigma-Aldrich</td>
			<td>T9281-50ML</td>
			<td>BioReagent, suitable for electrophoresis, ~99%</td>
		</tr>
		<tr>
			<td>NaNO3</td>
			<td>Sigma-Aldrich</td>
			<td>229938-10G</td>
			<td>99.995% trace metals basis</td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Sigma-Aldrich</td>
			<td>1064980500</td>
			<td>Pellets for analysis</td>
		</tr>
		<tr>
			<td>Overhead shaker</td>
			<td>GFL</td>
			<td>3040</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfluoroalkoxy alkane (PFA) vials</td>
			<td>Savillex</td>
			<td>200-015-20</td>
			<td>15 mL Standard Vial, Rounded Interior</td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>Thermo Scientific</td>
			<td>13-644-928</td>
			<td>Orion 3-Star Benchtop pH Meter</td>
		</tr>
		<tr>
			<td>pH probe</td>
			<td>Thermo Scientific</td>
			<td>8157BNUMD</td>
			<td>Orion ROSS Ultra pH/ATC Triode</td>
		</tr>
		<tr>
			<td>Plastic cutter</td>
			<td>DGT Research Ltd</td>
			<td>n/a</td>
			<td>Use empty cross-linker vials from DGT research Ltd</td>
		</tr>
		<tr>
			<td>Plastic tweezers</td>
			<td>Semadeni</td>
			<td>602</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasticine</td>
			<td>Local stationary shop</td>
			<td>n/a</td>
			<td>non-drying plastic modelling mass based on paraffin wax and bulking agents</td>
		</tr>
		<tr>
			<td>Polycarbonate membrane discs</td>
			<td>Whatman</td>
			<td>110606</td>
			<td>Nuclepore Hydrophilic Membrane, 25 mm diameter, 0.2 &#181;m pore size, 10 &#181;m thickness</td>
		</tr>
		<tr>
			<td>Polycarbonate membrane sheet</td>
			<td>Whatman</td>
			<td>113506</td>
			<td>Nuclepore Hydrophilic Membrane, 8 &#215; 10 in, 0.2 &#181;m pore size, 10 &#181;m thickness</td>
		</tr>
		<tr>
			<td>Polyethersulfone membrane discs</td>
			<td>Pall Corporation</td>
			<td>60172</td>
			<td>Supor 450 Membrane Disc Filters, 25 mm diameter, 0.45 &#181;m pore size, 0.14 mm thickness</td>
		</tr>
		<tr>
			<td>Polyethersulfone membrane sheet</td>
			<td>Pall Corporation</td>
			<td>60179</td>
			<td>Supor 450 Membrane Disc Filters, 293 mm diameter, 0.45 &#181;m pore size, 0.14 mm thickness</td>
		</tr>
		<tr>
			<td>PTFE foil</td>
			<td>Haberkorn</td>
			<td>n/a</td>
			<td>50 &#181;m thickness</td>
		</tr>
		<tr>
			<td>PTFE spacer</td>
			<td>Haberkorn</td>
			<td>n/a</td>
			<td>Variable thicknesses available</td>
		</tr>
		<tr>
			<td>PTFE-coated razor blades</td>
			<td>Personna GEM</td>
			<td>62-0178</td>
			<td>Stainless steel single edge blades (coated)</td>
		</tr>
		<tr>
			<td>PTFE-coated Tygon tubing</td>
			<td>S-prep GmbH</td>
			<td>SP8180</td>
			<td>0.32 cm inner diameter</td>
		</tr>
		<tr>
			<td>Quadrupole ICP-MS</td>
			<td>Perkin Elmer</td>
			<td>N8150044</td>
			<td>NexION 2000B</td>
		</tr>
		<tr>
			<td>Quantitative filter paper, 454</td>
			<td>VWR</td>
			<td>516-0854</td>
			<td>Particle retention 12-15 &#181;m</td>
		</tr>
		<tr>
			<td>Spreadsheet software</td>
			<td>Microsoft Corporation</td>
			<td>n/a</td>
			<td>Microsoft Excel 2016 (v16.0)</td>
		</tr>
		<tr>
			<td>Stainless-steel cutter</td>
			<td>Local locksmithery</td>
			<td>n/a</td>
			<td>2.5 cm diameter</td>
		</tr>
		<tr>
			<td>Suspended particulate reagent-iminodiacetate (SPR-IDA)</td>
			<td>Teledyne CETAC Technologies</td>
			<td>n/a</td>
			<td>10 &#181;m diameter polystyrene beads, 10 % (w/v) bead suspension</td>
		</tr>
		<tr>
			<td>Transistor-transistor logic (TTL) cable</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Consult ICP-MS technician to identify a suitable TTL cable for a specific instrument</td>
		</tr>
		<tr>
			<td>Two-volume cell</td>
			<td>Elemental Scientific Lasers</td>
			<td>n/a</td>
			<td>Two-volume cell 1</td>
		</tr>
		<tr>
			<td>Vinyl electrical tape</td>
			<td>3M</td>
			<td>n/a</td>
			<td>Scotch Super 33+</td>
		</tr>
		<tr>
			<td>Water purification system</td>
			<td>Termo Electron LED GmbH</td>
			<td>n/a</td>
			<td>TKA-GenPure</td>
		</tr>
		<tr>
			<td>ZrOCl2 &#215; 8H2O</td>
			<td>Alfa Aesar</td>
			<td>86108.30</td>
			<td>99.9 %, metals basis</td>
		</tr>
	</tbody>
</table>

two-dimensional visualization and quantification of labile, inorganic plant nutrients and contaminants in soil

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic nutrient (e.g., P, Fe, Mn) and contaminant (e.g., As, Cd, Pb) solute species in the soil adjacent to plant roots (the &#8216;rhizosphere&#8217;) at sub-millimeter (~100 &#181;m) spatial resolution. The method combines sink-based solute sampling by the diffusive gradients in thin films (DGT) technique with spatially resolved chemical analysis by laser ablation inductively coupled plasma mass spectrometry (LA-ICP-MS). The DGT technique is based on thin hydrogels with homogeneously distributed analyte-selective binding phases. The variety of available binding phases allows for the preparation of different DGT gel types following simple gel fabrication procedures. For DGT gel deployment in the rhizosphere, plants are grown in flat, transparent growth containers (rhizotrons), which enable minimal invasive access to a soil-grown root system. After a pre-growth period, DGT gels are applied to selected regions of interest for in situ solute sampling in the rhizosphere. Afterwards, DGT gels are retrieved and prepared for subsequent chemical analysis of the bound solutes using LA-ICP-MS line-scan imaging. Application of internal normalization using 13C and external calibration using matrix-matched gel standards further allows for the quantification of the 2D solute fluxes. This method is unique in its capability to generate quantitative, sub-mm scale 2D images of multi-element solute fluxes in soil-plant environments, exceeding the achievable spatial resolution of other methods for measuring solute gradients in the rhizosphere substantially. We present the application and evaluation of the method for imaging multiple cationic and anionic solute species in the rhizosphere of terrestrial plants and highlight the possibility of combining this method with complementary solute imaging techniques.

To demonstrate the capability and data detail of the DGT imaging method, we compiled the sub-mm, 2D flux distribution of multiple labile nutrient and contaminant solute species in soil adjacent to roots of Fagopyrum esculentum and Salix smithiana (Figure 7). Approximate times for general procedural steps of the protocol are presented in Table 1.
Solute images in Figure 7 were generated in three different studies using either HR-MBG or HR-CBG binding gels. The chemical images show the 2D solute flux distribution at a spatial resolution of 82-120 &#181;m along the x-axis and 300-400 &#181;m along the y-axis, depending on the LA-ICP-MS parameters used. Because no interpolation was applied during image calibration and resizing, single pixels represent measured datapoints. Alignment of the solute images with a photographic image of the ROI reveals that the sub-mm, 2D solute flux distribution of different elements is highly variable according to soil structure and root morphology. This can be attributed to the differential biogeochemical behavior of the elements in the soil-rhizosphere-plant system, and their interaction with the soil matrix and the plant roots.
In Figure 7A labile inorganic Mg, Al, P, Mn and Fe solute fluxes were visualized around a young F. esculentum root grown in carbonate-free soil fertilized with NH4NO3. The sub-mm solute distribution showed zones of decreased Al, P and Fe fluxes alongside older root sections due to root uptake, and highly increased Mg, Al, P, Mn and Fe fluxes at the root apex due to localized P mobilization processes of the F. esculentum root. Note that the root tip is located somewhat behind the soil surface and therefore hardly visible in the photographic image. Figure 7B shows the distribution of labile trace metals, including Mn, Fe, Zn, Cd and Pb around a root of metal-tolerant S. smithiana grown in a soil moderately contaminated with Zn, Cd and Pb. The solute images visualized distinct depletion particularly of Zn, Cd and Pb at the immediate root position, showing that S. smithiana roots act as a localized sink for labile trace metals in contaminated soil. Besides, localized Zn, Cd and Pb flux increases can be observed, indicating accumulation of these trace metals at the immediate soil-root interface.
In addition to multi-elemental solute imaging, the presented method can also be combined with complementary diffusion-based imaging techniques such as planar optodes34. This is demonstrated in Figure 7C, where the distribution of labile trace metals in the rhizosphere of S. smithiana was co-localized with the distribution of pH using a combined, single-layer planar optode-DGT cation binding gel33. The soil substrate was fertilized with (NH4)2SO4, leading to a pH decrease along the root axes by ~1 unit as compared to bulk soil. The pH decreases were co-localized with increased solute fluxes of Mn, Fe, Co, Ni, Cu and Pb, suggesting pH-induced metal solubilization.
Moreover, these example results show some of the potential imaging artefacts that may be obtained. For example, structural soil inhomogeneities, e.g., observed as a horizontal line in the lower third of the ROI image of Figure 7A, can cause soil-gel contact discontinuities resulting in limited diffusion at this location into the binding gel. Conversely, excessive soil compaction in the rhizotron can lead to poor porosity resulting in an artificial shift of the soil redox status towards anoxia. This is illustrated in Figure 7B, where extensive areas of highly elevated Mn and Fe fluxes in the solute images visually matched with a dense layer of soil in the ROI image. This suggests a decreased soil redox potential due to high soil compaction, resulting in reductive dissolution and solubilization of the highly redox-sensitive elements. Careful rhizotron filling and visual inspection of the soil surface directly after filling is therefore recommended.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61661/61661fig07.jpg" /> 
Figure 7: Sub-mm 2D distribution of labile nutrient and contaminant solute species across different soil-root interfaces.&#160;(A) Distribution of anionic P and cationic Mg, Al, Mn, and Fe solutes around a young F. esculentum root. Co-localized sampling of anionic and cationic solutes was achieved using HR-MBG for 24 h at a soil water saturation of ~75% WHC. The Al, P and Mn images are displayed as calibrated fDGT values (pg cm-2 s-1), whereas Mg and Fe images show 13C-normalized intensities. Scale bar represents 1 cm. This figure is adapted from ref.48. (B) Distribution of Mn, Fe, Zn, Cd and Pb around a S. smithiana root grown in soil moderately contaminated with Zn, Cd and Pb. Cationic trace metal solutes were sampled using HR-CBG for 20 h at a soil water saturation of ~80 % WHC. All images are displayed as calibrated fDGT values (pg cm-2 s-1). Scale bar represents 0.5 cm. This figure is adapted from ref.3. (C) Distribution of pH and multiple cationic solutes around S. smithiana roots grown in soil spiked with Cd. Co-localization of pH and solute dynamics was achieved using a modification of the HR-MBG protocol, allowing for simultaneous solute sampling and planar optode imaging33. The Mn, Cu, Zn and Cd images are displayed as calibrated fDGT values (pg cm-2 s-1), whereas Fe, Co, Ni and Pb images show 13C-normalized intensities. Scale bar represents 1 cm. This figure is adapted from ref.33. The presented figures are reproduced from the cited articles3,33,48 licensed under CC BY. <a href="https://www.jove.com/files/ftp_upload/61661/61661fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental File 1.&#160;<a href="https://www.jove.com/files/ftp_upload/61661/JoVE61661_Supporting Information_Updated 2021_.pdf" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil at JoVE.com

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

This protocol presents a workflow for sub-mm 2D visualization of multiple labile inorganic nutrient and contaminant solute species using diffusive gradients in thin films (DGT) combined with mass spectrometry imaging. Solute sampling and high-resolution chemical analysis are described in detail for quantitative mapping of solutes in the rhizosphere of terrestrial plants.

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic ...

two-dimensional-visualization-quantification-labile-inorganic-plant

Research

JoVE Journal

Environment

5.9K Views.  Montanuniversität Leoben. This protocol presents a workflow for sub-mm 2D visualization of multiple labile inorganic nutrient and contaminant solute species using diffusive gradients in thin films (DGT) combined with mass spectrometry imaging. Solute sampling and high-resolution chemical analysis are described in detail for quantitative mapping of solutes in the rhizosphere of terrestrial plants.

Video: Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

Tracing rare stable isotopes from plant material through the ecosystem provides the most sensitive information about ecosystem processes; from CO2 fluxes and soil organic matter formation to small-scale stable-isotope biomarker probing. Coupling multiple stable isotopes such as 13C with 15N, 18O or&#160;2H has the potential to reveal even more information about complex stoichiometric relationships during biogeochemical transformations. Isotope labeled plant material has been used in various studies of litter decomposition and soil organic matter formation1-4. From these and other studies, however, it has become apparent that structural components of plant material behave differently than metabolic components (i.e. leachable low molecular weight compounds) in terms of microbial utilization and long-term carbon storage5-7. The ability to study structural and metabolic components separately provides a powerful new tool for advancing the forefront of ecosystem biogeochemical studies. Here we describe a method for producing 13C and 15N labeled plant material that is either uniformly labeled throughout the plant or differentially labeled in structural and metabolic plant components.
Here, we present the construction and operation of a continuous 13C and 15N labeling chamber that can be modified to meet various research needs. Uniformly labeled plant material is produced by continuous labeling from seedling to harvest, while differential labeling is achieved by removing the growing plants from the chamber weeks prior to harvest. Representative results from growing Andropogon gerardii Kaw demonstrate the system&#39;s ability to efficiently label plant material at the targeted levels. Through this method we have produced plant material with a 4.4 atom%13C and 6.7 atom%15N uniform plant label, or material that is differentially labeled by up to 1.29 atom%13C and 0.56 atom%15N in its metabolic and structural components (hot water extractable and hot water residual components, respectively). Challenges lie in maintaining proper temperature, humidity, CO2 concentration,&#160;and light levels in an airtight 13C-CO2 atmosphere for successful plant production. This chamber description represents a useful research tool to effectively produce uniformly or differentially multi-isotope labeled plant material for use in experiments on ecosystem biogeochemical cycling.

Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

This method explains how to build and operate a continuous 13C and 15N isotope labeling chamber for uniform or differential plant tissue labeling. Representative results from metabolic and structural labeling of Andropogon gerardii are discussed.

Tracing rare stable isotopes from plant material through the ecosystem provides the most sensitive information about ecosystem processes; from CO2 fluxes ...

Experimental Protocol for Manipulating Plant-induced Soil Heterogeneity

Coexistence theory has often treated environmental heterogeneity as being independent of the community composition; however biotic feedbacks such as plant-soil feedbacks (PSF) have large effects on plant performance, and create environmental heterogeneity that depends on the community composition. Understanding the importance of PSF for plant community assembly necessitates understanding of the role of heterogeneity in PSF, in addition to mean PSF effects. Here, we describe a protocol for manipulating plant-induced soil heterogeneity. Two example experiments are presented: (1) a field experiment with a 6-patch grid of soils to measure plant population responses and (2) a greenhouse experiment with 2-patch soils to measure individual plant responses. Soils can be collected from the zone of root influence (soils from the rhizosphere and directly adjacent to the rhizosphere) of plants in the field from conspecific and heterospecific plant species. Replicate collections are used to avoid pseudoreplicating soil samples. These soils are then placed into separate patches for heterogeneous treatments or mixed for a homogenized treatment. Care should be taken to ensure that heterogeneous and homogenized treatments experience the same degree of soil disturbance. Plants can then be placed in these soil treatments to determine the effect of plant-induced soil heterogeneity on plant performance. We demonstrate that plant-induced heterogeneity results in different outcomes than predicted by traditional coexistence models, perhaps because of the dynamic nature of these feedbacks. Theory that incorporates environmental heterogeneity influenced by the assembling community and additional empirical work is needed to determine when heterogeneity intrinsic to the assembling community will result in different assembly outcomes compared with heterogeneity extrinsic to the community composition.

Understanding the role of environmental heterogeneity in species coexistence has typically focused on types of heterogeneity that are extrinsic to the community&#8217;s species composition. We provide novel detailed methods for creating soil heterogeneity treatments using soils subject to plant-soil feedback conditioning, or heterogeneity intrinsic to the community composition.

Coexistence theory has often treated environmental heterogeneity as being independent of the community composition; however biotic feedbacks such as ...

Quantification of Heavy Metals and Other Inorganic Contaminants on the Productivity of Microalgae

Increasing demand for renewable fuels has researchers investigating the feasibility of alternative feedstocks, such as microalgae. Inherent advantages include high potential yield, use of non-arable land and integration with waste streams. The nutrient requirements of a large-scale microalgae production system will require the coupling of cultivation systems with industrial waste resources, such as carbon dioxide from flue gas and nutrients from wastewater. Inorganic contaminants present in these wastes can potentially lead to bioaccumulation in microalgal biomass negatively impact productivity and limiting end use. This study focuses on the experimental evaluation of the impact and the fate of 14 inorganic contaminants (As, Cd, Co, Cr, Cu, Hg, Mn, Ni, Pb, Sb, Se, Sn, V and Zn) on Nannochloropsis salina growth. Microalgae were cultivated in photobioreactors illuminated at 984 &#181;mol m-2 sec-1 and maintained at pH 7 in a growth media polluted with inorganic contaminants at levels expected based on the composition found in commercial coal flue gas systems. Contaminants present in the biomass and the medium at the end of a 7 day growth period were analytically quantified through cold vapor atomic absorption spectrometry for Hg and through inductively coupled plasma mass spectrometry for As, Cd, Co, Cr, Cu, Mn, Ni, Pb, Sb, Se, Sn, V and Zn. Results show N. salina is a sensitive strain to the multi-metal environment with a statistical decrease in biomass yieldwith the introduction of these contaminants. The techniques presented here are adequate for quantifying algal growth and determining the fate of inorganic contaminants.

Integration of microalgal cultivation with industrial flue gas will ultimately introduce heavy metals and other inorganic compounds into the growth media. This study presents a procedure used to determine the end fate and impact of heavy metals and inorganic contaminants on the growth of Nannochloropsis salina grown in photobioreactors.

Increasing demand for renewable fuels has researchers investigating the feasibility of alternative feedstocks, such as microalgae. Inherent advantages ...

Assessment of Labile Organic Carbon in Soil Using Sequential Fumigation Incubation Procedures

Management practices and environmental changes can alter soil nutrient and carbon cycling. Soil labile organic carbon, a readily decomposable C pool, is highly sensitive to disturbance. It is also the primary substrate for soil microorganisms, which is fundamental to nutrient cycling. Due to these attributes, labile organic carbon (LOC) has been identified as an indicator parameter for soil health. Quantifying the turnover rate of LOC also aids in understanding changes in soil nutrient cycling processes. A sequential fumigation incubation method has been developed to estimate soil LOC and potential C turnover rate. The method requires fumigating soil samples and quantifying CO2-C respired during a 10 day incubation period over a series of fumigation-incubation cycles. Labile organic C and potential C turnover rate are then extrapolated from accumulated CO2 with a negative exponential model. Procedures for conducting this method are described.

Labile organic carbon (LOC) and the potential carbon turnover rate are sensitive indicators of changes in soil nutrient cycling processes. Details are provided for a method based on fumigating and incubating soil in a series of cycles and using the CO2 accumulated during the incubation periods to estimate these parameters.

Management practices and environmental changes can alter soil nutrient and carbon cycling. Soil labile organic carbon, a readily decomposable C pool, is ...

A Gusseted Thermogradient Table to Control Soil Temperatures for Evaluating Plant Growth and Monitoring Soil Processes

Thermogradient tables were first developed in the 1950s primarily to test seed germination over a range of temperatures simultaneously without using a series of incubators. A temperature gradient is passively established across the surface of the table between the heated and cooled ends and is lost quickly at distances above the surface. Since temperature is only controlled on the table surface, experiments are restricted to shallow containers, such as Petri dishes, placed on the table. Welding continuous aluminum vertical strips or gussets perpendicular to the surface of a table enables temperature control in depth via convective heat flow. Soil in the channels between gussets was maintained across a gradient of temperatures allowing a greater diversity of experimentation. The gusseted design was evaluated by germinating oat, lettuce, tomato, and melon seeds. Soil temperatures were monitored using individual, battery-powered dataloggers positioned across the table. LED lights installed in the lids or along the sides of the gradient table create a controlled temperature chamber where seedlings can be grown over a range of temperatures. The gusseted design enabled accurate determination of optimum temperatures for fastest germination rate and the highest percentage germination for each species. Germination information from gradient table experiments can help predict seed germination and seedling growth under the adverse soil conditions often encountered during field crop production. Temperature effects on seed germination, seedling growth, and soil ecology can be tested under controlled conditions in a laboratory using a gusseted thermogradient table.

Traditional thermogradient tables create a range of temperatures across the surface. Welding gussets perpendicular to the surface of a thermogradient table will control temperature in depth increasing possible research applications.

Thermogradient tables were first developed in the 1950s primarily to test seed germination over a range of temperatures simultaneously without using a ...

Continuous Instream Monitoring of Nutrients and Sediment in Agricultural Watersheds

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in water resources is a prerequisite for understanding the drivers of the pollutant loads and for making informed water resource management decisions. The commonly used &#34;grab sampling&#34; method provides the concentrations of pollutants at the time of sampling (i.e., a snapshot concentration) and may under- or overpredict the pollutant concentrations and loads. Continuous monitoring of nutrients and sediment has recently received more attention due to advances in computing, sensing technology, and storage devices. This protocol demonstrates the use of sensors, sondes, and instrumentation to continuously monitor in situ nitrate, ammonium, turbidity, pH, conductivity, temperature, and dissolved oxygen (DO) and to calculate the loads from two streams (ditches) in two agricultural watersheds. With the proper calibration, maintenance, and operation of sensors and sondes, good water quality data can be obtained by overcoming challenging conditions such as fouling and debris buildup. The method can also be used in watersheds of various sizes and characterized by agricultural, forested, and/or urban land.

With the advancement of technology and the rise in end-user expectations, the need and use of higher temporal resolution data for pollutant load estimation has increased. This protocol describes a method for continuous in situ water quality monitoring to obtain higher temporal resolution data for informed water resource management decisions.

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in ...

Isolation and Analysis of Microbial Communities in Soil, Rhizosphere, and Roots in Perennial Grass Experiments

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The purpose of this manuscript is to provide detail on how to rapidly sample soil, rhizosphere, and endosphere of replicated field trials and analyze changes that may occur in the microbial communities due to sample type, treatment, and plant genotype. The experiment used to demonstrate these methods consists of replicated field plots containing two, pure, warm-season grasses (Panicum virgatum and Andropogon gerardii) and a low-diversity grass mixture (A. gerardii, Sorghastrum nutans, and Bouteloua curtipendula). Briefly, plants are excavated, a variety of roots are cut and placed in phosphate buffer, and then shaken to collect the rhizosphere. Roots are brought to the laboratory on ice and surface sterilized with bleach and ethanol (EtOH). The rhizosphere is filtered and concentrated by centrifugation. Excavated soil from around the root ball is placed into plastic bags and brought to the lab where a small amount of soil is taken for DNA extractions. DNA is extracted from roots, soil, and rhizosphere and then amplified with primers for the V4 region of the 16S rRNA gene. Amplicons are sequenced, then analyzed with open access bioinformatics tools. These methods allow researchers to test how the microbial community diversity and composition varies due to sample type, treatment, and plant genotype. Using these methods along with statistical models, the representative results demonstrate there are significant differences in microbial communities of roots, rhizosphere, and soil. Methods presented here provide a complete set of steps for how to collect field samples, isolate, extract, quantify, amplify, and sequence DNA, and analyze microbial community diversity and composition in replicated field trials.

Excavation of plant roots from the field as well as processing of samples into endosphere, rhizosphere, and soil are described in detail, including DNA extraction and data analysis methods. This paper is designed to enable other laboratories to use these techniques for the study of soil, endosphere, and rhizosphere microbiomes.

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The ...

An Optimized Rhizobox Protocol to Visualize Root Growth and Responsiveness to Localized Nutrients

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive harvest or expensive equipment. We present a customizable and affordable rhizobox method that allows the non-destructive visualization of root growth over time and is particularly well-suited to studying root plasticity in response to localized resource patches. The method was validated by assessing maize genotypic variation in plasticity responses to patches containing 15N-labeled legume residue. Methods are described to obtain representative developmental measurements over time, measure root length density in resource-containing and control patches, calculate root growth rates, and determine 15N recovery by plant roots and shoots. Advantages, caveats, and potential future applications of the method are also discussed. Although care must be taken to ensure that experimental conditions do not bias root growth data, the rhizobox protocol presented here yields reliable results if carried out with sufficient attention to detail.

Visualizing and measuring root growth in situ is extremely challenging. We present a customizable rhizobox method to track root development and proliferation over time in response to nutrient enrichment. This method is used to analyze maize genotypic differences in root plasticity in response to an organic nitrogen source.

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive ...

High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling in soils, to pathogen competition, plant growth promotion, and cross-kingdom signaling. Furthermore, siderophores are also of commercial interest in bioleaching and bioweathering of metal-bearing minerals and ores. A rapid, cost effective, and robust means of quantitatively assessing siderophore production in complex samples is key to identifying important aspects of the ecological ramifications of siderophore activity, including, novel siderophore producing microbes. The method presented here was developed to assess siderophore activity of in-tact microbiome communities, in environmental samples, such as soil or plant tissues. The samples were homogenized and diluted in a modified M9 medium (without Fe), and enrichment cultures were incubated for 3 days. Siderophore production was assessed in samples at 24, 48, and 72 hours (h) using a novel 96-well microplate CAS (Chrome azurol sulphonate)-Fe agar assay, an adaptation of the traditionally tedious and time-consuming colorimetric method of assessing siderophore activity, performed on individual cultivated microbial isolates. We applied our method to 4 different genotypes/Lines of wheat (Triticum aestivum L.), including Lewjain, Madsen, and PI561725, and PI561727 commonly grown in the inland Pacific Northwest. Siderophore production was clearly impacted by the genotype of wheat, and in the specific types of plant tissues observed. We successfully used our method to rapidly screen for the influence of plant genotype on siderophore production, a key function in terrestrial and aquatic ecosystems. We produced many technical replicates, yielding very reliable statistical differences in soils and within plant tissues. Importantly, the results show the proposed method can be used to rapidly examine siderophore production in complex samples with a high degree of reliability, in a manner that allows communities to be preserved for later work to identify taxa and functional genes.

We present a protocol for rapid screening of environmental samples for siderophore potential contributing to micronutrient bioavailability and turnover in terrestrial systems.

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling ...

Measuring Phosphorus Release in Laboratory Microcosms for Water Quality Assessment

Phosphorus (P) is a critical limiting nutrient in agroecosystems requiring careful management to reduce transport risk to aquatic environments. Routine laboratory measures of P bioavailability are based on chemical extractions performed on dried samples under oxidizing conditions. While useful, these tests are limited with respect to characterizing P release under prolonged water saturation. Labile orthophosphate bound to oxidized iron and other metals can rapidly desorb to solution in reducing environments, increasing P mobilization risk to surface runoff and groundwater. To better quantify P desorption potential and mobility during extended saturation, a laboratory microcosm method was developed based on repeated sampling of porewater and overlying floodwater over time. The method is useful for quantifying P release potential from soils and sediments varying in physicochemical properties and can improve site-specific P mitigation efforts by better characterizing P release risk in hydrologically active areas. Advantages of the method include its ability to simulate in situ dynamics, simplicity, low cost, and flexibility.

Accurate quantification of phosphorus (P) desorption potential in saturated soils and sediments is important for P modeling and transport mitigation efforts. To better account for in situ soil-water redox dynamics and P mobilization under prolonged saturation, a simple approach was developed based on repeated sampling of laboratory microcosms.

Phosphorus (P) is a critical limiting nutrient in agroecosystems requiring careful management to reduce transport risk to aquatic environments. Routine ...