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. 2026 Apr 12;10(4):e70164. doi: 10.1002/pld3.70164

Detection and Quantification of Dysprosium in Plant Tissues

Edmaritz Hernández‐Pagán 1, Kanjana Laosuntisuk 1, Alex T Harris 1, Allison N Haynes 1, David Buitrago 2, Anisa Guedira 1, Cyprian Rajabu 1, Michael W Kudenov 2,✉, Colleen J Doherty 1,✉
PMCID: PMC13070872  PMID: 41982647

ABSTRACT

The growing demand for rare‐earth elements (REEs), particularly dysprosium (Dy), underscores the need for sustainable extraction methods. Recovery of Dy, particularly from geographically distributed waste sources, is challenging. This gap positions phytomining, a technique using plants to accumulate metals, as a promising alternative. However, plant species differ in their ability to accumulate metals in high concentrations, necessitating efficient screening methods. In this study, we developed a high‐throughput fluorescence‐based assay to detect and quantify Dy uptake in plant tissues. The Dy detection method described in the present work exploits Dy's unique spectroscopic properties for sensitive and efficient analysis, enabling the detection of concentrations as low as 0.07 μM, with a detection limit of 0.2 μM in a plant matrix. By incorporating sodium tungstate (Na2WO4) as a fluorescence enhancer, we achieved robust emission intensities at 480 and 580 nm, facilitating Dy quantification in complex plant matrices. Additionally, the use of time‐resolved fluorescence techniques reduces background autofluorescence from plant tissues, enhancing signal specificity. Validation of the fluorescence method with inductively coupled plasma mass spectrometry (ICP‐MS) demonstrated a strong correlation in Dy levels. Greenhouse trials confirmed the method's utility for screening Dy accumulation in living plants and highlighted the potential for rapid stand‐off detection. This fluorescence‐based approach offers a scalable, efficient tool for identifying Dy‐accumulating plants and advances phytomining as a sustainable strategy for REE recovery.

Keywords: agromining, dysprosium detection, phytomining, rare‐earth elements, spectral imaging

Abbreviations

Ce

cerium

Dy

dysprosium

DyCl3

dysprosium chloride

ICP‐AES

ICP atomic emission spectroscopy

ICP‐MS

inductively coupled plasma mass spectrometry

La

lanthanum

Na2WO4

sodium tungstate

Nd

neodymium

REE

rare‐earth element

UV

ultraviolet

UV–Vis–NIR

ultraviolet–visible‐near‐infrared

XRF

X‐ray fluorescence

Y

yttrium

1. Background

Rare‐earth elements (REEs) comprise 17 elements, including scandium (Sc), yttrium (Y), and the lanthanides ranging from lanthanum (La) to lutetium (Lu) (Voncken 2016). The rising use of clean energy technologies has sharply increased the demand for REEs, particularly dysprosium (Dy) (Seo and Morimoto 2014; Xiao et al. 2022; Eheliyagoda et al. 2023). Dy is added to neodymium–iron–boron (NdFeB) permanent magnets to improve high‐temperature resistance (Riaño and Binnemans 2015). These magnets are used in hybrid electric vehicles, wind turbines, computer hard drives, home appliances, and other applications (Seo and Morimoto 2014).

Extracting and processing REEs is challenging despite their relative abundance in the Earth's crust. REEs are usually dispersed and mixed with non‐REE metals, making mining and purification costly and potentially environmentally harmful (Balaram 2019). As of 2025, China holds 49% of global REE reserves (U.S. Department of the Interior U.S. Geological Survey 2025) and, as of 2019, 87% of the world's Dy reserves (Xiao et al. 2022). This dominance, combined with projected increases in demand, has intensified interest in identifying alternative REE sources to support the green energy transition (Dutta et al. 2016; Li et al. 2019).

Concerns over supply risks and environmental impacts have driven efforts to identify new REE sources and reduce the carbon footprint of traditional mining (Balaram 2019; Talan and Huang 2022). Secondary sources such as coal fly ash, acid mine drainage, and recycled magnets are being explored to ensure a sustainable REE supply (Peiravi et al. 2021; Mwewa et al. 2022).

Phytomining—using plants to extract metals from soil (Dinh et al. 2022; van der Ent et al. 2020)—shows promise for sustainable Dy recovery. Some plants can tolerate, uptake, concentrate, and translocate metals to aerial tissues at levels significantly higher than, or even toxic to, other plants in the same environment (Yang et al. 2005; van der Ent and Rylott 2024). Metal accumulation in plants varies considerably from species to species, but some plants can accumulate metal concentrations ranging from 100 to 10,000 mg/kg or higher. Plants that accumulate metals at these high levels tend to be highly specific to a particular element (van der Ent and Rylott 2024). Due to their ability to concentrate metals, such accumulator plants are valuable for extracting valuable elements and cleaning contaminated soils (Rylott and Bruce 2022). For example, nickel phytomining research has successfully identified nickel‐accumulating species such as Pycnandra acuminata and Phyllanthus balgooyi, which are being scaled for commercial nickel recovery (Tognacchini et al. 2020; Mesjasz‐Przybylowicz et al. 2016; Isnard et al. 2020; Chaney 2020).

Efforts are ongoing to identify plant species that may be suitable for REE phytomining. For example, plants in the genus Phytolacca are promising candidates for phytomining and phytoremediation (Dinh et al. 2022; Liu, Liu, et al. 2021; Grosjean et al. 2019). Phytolacca americana can accumulate up to 1040‐mg/kg REEs in the leaf tissues when grown in REE‐contaminated soils (Yuan et al. 2018). Expanding the knowledge of these and other species across various environments is crucial for developing effective REE phytomining methods.

To develop a successful Dy phytomining system for secondary REE sources, it is essential to rapidly identify plants capable of accumulating high Dy levels in their leaf tissues. Conventional methods for measuring Dy in plants rely on inductively coupled plasma‐based (ICP) techniques such as ICP mass spectrometry (ICP‐MS) and ICP atomic emission spectroscopy (ICP‐AES). ICP‐MS can detect elements at concentrations as low as parts per trillion (ppt), equivalent to 0.001 μg/L, corresponding to picomolar concentrations (e.g., 6.15 × 10−6 μM Dy). In contrast, ICP‐AES can detect elements at parts per billion (ppb), equivalent to 1 μg/L and down to nanomolar concentrations (Zawisza et al. 2011). However, these techniques can require large amounts of plant tissue, and sample preparation results in tissue destruction due to a digestion step, often via acid‐ or microwave‐assisted digestion. This requirement for tissue destruction limits single‐plant and time‐course Dy detection and measurement (Huang and Schulte 1985). Since plant species vary in REE uptake, developing alternative, nondestructive screening methods will help identify plants with the highest potential for REE accumulation.

Alternative measurement techniques for REE detection include X‐ray fluorescence (XRF). XRF microscopy detected and mapped La and cerium (Ce) in Dicranopteris linearis growing on REE mine tailings in Ganzhou, Jiangxi Province, China. The results showed that La and Ce were colocalized in the pinnae and pinnules and were enriched in areas of dead tissue and leaf veins (Liu et al. 2020). Similarly, XRF spectroscopy was used to identify Y hyperaccumulating plants from herbarium specimens (Goudard et al. 2024). This screen identified plants previously described as high REE accumulators, including D. linearis and Blechnopsis orientalis. In addition, this study identified 15 new REE‐accumulating species. However, P. americana , a species known to accumulate REEs and to preferentially accumulate heavy REEs, such as Dy, in its leaves (Grosjean et al. 2019; Yuan et al. 2018), was not identified in the herbarium screen. Herbarium collections include plants from various locations. Therefore, the P. americana plants in the herbarium collection may have been collected from areas with low or no REEs in the soil. In addition to screening herbarium samples to identify REE‐accumulating plants, Goudard et al. (2024) also employed XRF microscopy to examine the distribution of five other REEs in addition to Y (Ce, La, Nd, Praseodymium [Pr], and Ytterbium [Yb]) in D. linearis tissue. The results showed that Ce, La, Nd, and Pr were concentrated in the vascular bundles of the pinnules. In contrast, Y was primarily localized in the tissue surrounding the vasculature, suggesting different distribution patterns of individual REEs in plant tissue. These findings highlight XRF's potential for the nondestructive measurement of the concentration and distribution of REEs in plants. However, XRF can be sensitive to the sample matrix, making it challenging to employ across diverse, living biological samples. Additionally, XRF primarily analyzes the upper surface, with reduced effectiveness at greater depths (Guagliardi et al. 2025).

Here, we present an alternative method for screening Dy uptake in plants using deep‐UV‐induced fluorescence. The spectroscopic properties of REEs allow high‐throughput detection from limited plant material. Lanthanide 4f‐4f transitions produce multicolor emission from the ultraviolet–visible‐near‐infrared (UV–Vis–NIR) range, long luminescence decay, and narrow absorption and emission bands (Sato et al. 2016; Runowski et al. 2020). Dy emits at 480 nm (blue) and 575–580 nm (yellow) when excited with UV light (Sruthi et al. 2022; Gitty et al. 2022). Here, we describe a 96‐well plate UV fluorescence assay for phytomining‐relevant Dy concentrations, demonstrate its application to plant tissue, and compare it with traditional ICP‐MS quantification methods. This approach can screen plant tissues for Dy uptake in a high‐throughput 96‐well plate format, and we demonstrate its potential for standoff detection in in‐field measurements.

2. Methods

2.1. Dy Absorbance and Fluorescence Measurement From DyCl3 Solution to Determine Dy Excitation Wavelength

Solutions containing 100 mM (16,250 mg/L) Dy were prepared in water at room temperature using a 96‐well plate and agitated for 2 h and 30 min. Absorbance measurements (BioTek Synergy H1 Multimode Reader, BioTek, USA) were taken from 250 to 700 nm at 10‐nm intervals, and from 250 to 500 nm at 1‐nm intervals. The wavelength of maximum absorbance obtained (350 nm) was used to excite the Dy ions in the solutions, implementing 60‐μs time‐resolved fluorescence and a 300‐μs delay collection time. The sensitizer, Na2WO4, was added to 0‐ and 0.8‐μM Dy solutions for a final concentration of 0.25 M, and excitation wavelengths were tested to determine the highest Dy emission using 60‐μs time‐resolved fluorescence and a 300‐μs delay collection time. Emission was measured at 480 and 580 nm. The instrument's general settings are as follows: The optics are on the top of the plate, the light source is a xenon flash, and the lamp energy is “High.” The “High” mono flash power is 883‐V input voltage with a 20‐ms interval (50 Hz) and about 0.4 J of energy per flash.

2.2. General Settings for Dy Fluorescence Assay in the Plate Reader

Dy emissions were measured using 20‐, 30‐, or 60‐μs time‐resolved fluorescence and a 300‐μs delay collection time. The excitation wavelength used was 275 nm, and the emission was collected from 450 to 700 nm as RFU based on the peak height at the wavelength. The instrument's general settings are as follows: The optics are on the top of the plate, the light source is a xenon flash, and the lamp energy is set to “High,” as described above.

2.3. Dy Fluorescence Measurement From DyCl3 Solution

In the 96‐well plates, 10 μL of DyCl3 solutions were mixed with 100‐μL Na2WO4 for a final concentration of 0.25 M. The concentrations of DyCl3 in this test were 0, 0.07, 0.1, 0.3, 0.5, 1, and 1.5 μM. The concentrations of Dy in μg/L were as follows: 0, 11.375, 16.25, 48.75, 81.25, 162.5, and 243.75, respectively. The plates were agitated at room temperature for 2 h and 30 min before the fluorescence assay. Dy emissions were measured using time‐resolved fluorescence as described above.

2.4. Preparation of Dy‐Spiked Samples

P. acinosa aerial tissue was dried in an oven overnight at 90°C, and 0.31 g of dried tissue was mixed with 31 mL of DyCl3 solution in Digitubes (SCP Science) to get final concentrations of 0, 0.05, 0.2, 0.8, 3.2, and 25.6 μM (0, 8.125, 32.5, 130, 520, and 4160 μg/L) of Dy in the samples. An aliquot was collected to use for fluorescence spectroscopy measurements. The samples were used for Dy emission spectra measurements and ICP‐MS. Dy emissions were measured using time‐resolved fluorescence as described above.

2.5. P. acinosa Growth and Treatment With DyCl3 Solution

P. acinosa seeds were stratified in concentrated sulfuric acid for 5 min with agitation and rinsed with DI water five times. Seeds were then surface sterilized with 50% bleach supplemented with 0.02% (v/v) Triton‐X for 7 min with agitation and rinsed with DI water three times. Afterward, seeds were kept in sterile DI water in the dark at 25°C for 2 days and exposed to 12/12‐h light/dark cycles for 7–10 days for germination. Germinating seeds were transferred to growing medium (OASIS ROOTCUBES, Oasis grower solution, USA) containing 0.1× Hoagland solution (0.5‐mM KNO3, 0.5‐mM Ca(NO3)2·4H2O, 0.6% Sprint138 chelated iron, 0.2‐mM MgSO4·7H2O, 0.1‐mM NH4NO3, 0.2‐mM KH2PO4, 5‐μM H3BO3, 1‐μM MnCl2·4H2O, 0.08‐μM ZnSO4·7H2O, 0.03‐μM CuSO4·5H2O, and 0.01‐μM H3MoO4·H2O). Once the plants had four fully expanded leaves, plants were transferred to 0.25× Hoagland solution (1.25‐mM KNO3, 1.25‐mM Ca(NO3)2·4H2O, 1.5% Sprint138 chelated iron, 0.5‐mM MgSO4·7H2O, 0.25‐mM NH4NO3, 12‐μM H3BO3, 2‐μM MnCl2·4H2O, 0.2‐μM ZnSO4·7H2O, 0.08‐μM CuSO4·5H2O, 0.02‐μM H3MoO4·H2O) supplemented with 0‐, 5‐, 10‐, 15‐, and 20‐mM DyCl3 solution (equivalent to 0, 812.5, 1625, 2437.5, and 3250 mg/L of Dy). KH2PO4 was excluded from the Hoagland solution to prevent Dy precipitation with phosphate (Grosjean et al. 2019). The leaf discs were cut from two areas of one leaf, leaf lamina and midrib, after growing in DyCl3 for 7 days using a 2‐mm diameter cork borer and placed in a 96‐well plate. The discs were then submerged in 100 μL of 0.5‐M Na2WO4, and the plates were agitated at room temperature for 2 h and 30 min before taking a measurement. Dy emissions were measured using time‐resolved fluorescence as described above.

2.6. Dy Screening in Intact Plant Tissue

A commercial deep UV Raman and fluorescence spectrometer by Photon Systems Inc. (DUV Raman/PL 200) was used to measure Dy signal from P. acinosa plants treated with a water solution containing 0, 10, and 50 mM (0, 1625, and 8125 mg/L) of DyCl3 following the fluorescence spectroscopy methods in Banah et al. (2024). The laser had a pulse frequency rate of 40 pulses per second, and 10 pulses with an energy of 5.2 μJ were used with a diameter between 100 and 200 μm. After plant treatment for 48 h, fluorescence emission spectra in the youngest leaves of the plants were measured using a 248‐nm excitation wavelength. The emission at 480 and 575–580 nm wavelengths was used as the Dy signal. For the comparative ICP‐MS measurements used to validate the spectroscopic readings, we pooled tissue from the stems and all the leaves of each plant. This pool of tissues was processed for ICP‐MS measurements.

2.7. Dy Analysis by ICP‐MS

Plants were dried in an oven at 110°C overnight, ground, and mineralized with concentrated nitric acid and 30% hydrogen peroxide for 1 h each in a DigiPrep MS (SCP Science). The sample volume was adjusted to 25 mL, adding 10,000 ppb of indium internal standard solution (SCP Science) for a final concentration of 50 ppb and 2% (v/v) HNO3. Sample precipitates were filtered using a manifold (SCP Science) and 1.0‐μm Digi filters (SCP Science). Dy concentrations were determined using an iCAP RQ quadrupole‐based ICP‐MS (Thermo Scientific). A certified reference (BCR‐670, Lemna minor ) was processed for quality control (Berky et al. 2023).

2.8. Dy Detections With Deep UV LED Light

P. acinosa leaves were incubated in 100‐mM DyCl3, 100‐mM CeCl3, or water for 3 h at room temperature. Then, small holes were generated with a thin needle (0.45 mm × 13 mm), and 3 μL of 2‐M Na2WO4 were added to the pierced area, followed by a 5‐min incubation at room temperature to promote tungstate penetration through the epidermis. Following the Dy and tungstate treatment, the leaves were exposed to LED light (ThermoLabs, M275L4) at 275 nm and generated images using a smartphone camera under dark conditions.

3. Results

3.1. Detection of Dy Using Deep‐UV Fluorescence Spectroscopy

We first scanned the Dy absorbance using 100‐mM DyCl3 in solution to determine the maximum Dy absorbance wavelengths. A 1‐nm resolution scan from 250 to 500 nm identified seven major and five minor peaks (Figure S1A). No additional peaks were detected between the 500‐ and 700‐nm wavelength region at 10‐nm resolution (Figure S1). The maximum absorbance was 350 nm (Figure S1A,B), consistent with the Dy UV–Vis absorption spectrum (Runowski et al. 2020).

Sodium tungstate (Na2WO4) enhances Dy fluorescence via excitation energy transfer (Alberti and Massucci 1966; Xu et al. 2014). We added 0.25‐M Na2WO4 to DyCl3 solutions and evaluated the emission peaks at the excitation wavelengths identified in the absorbance assays. The characteristic Dy emission peaks were at 480 and 580 nm (Figure 1A; Sruthi et al. 2022). We used time‐resolved fluorescence in these assays because endogenous plant molecules cause autofluorescence (Donaldson 2020), which can interfere with fluorescence spectra. Excitation between 255 and 295 nm generated robust Dy emission peaks, while 325 and 450 nm showed minimal emission (Figures 1 and S1C,D). The highest fluorescence occurred with 275‐nm excitation, which was selected for further experiments (Figures 1A and S1C,D).

FIGURE 1.

FIGURE 1

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Dy emission signals and emission of minimum Dy levels detected by fluorescence spectroscopy in an aqueous DyCl3 solution. (A) Dy emission spectrum from 0‐ and 0.8‐μM DyCl3 treated with 0.25‐M Na2WO4, using 255‐, 265‐, 275‐, and 295‐nm excitation wavelengths, 100 gain, and 60‐μs time‐resolved fluorescence. Lines indicate the mean measured value from four to five technical replicates. (B, C) Scatter plot and R‐squared values between the 0‐ and 1‐μM concentrations of DyCl3 with 0.25‐M Na2WO4 and Dy emission at (B) 480 and (C) 580 nm measured with an excitation wavelength of 275 nm, 150 gain, and 20‐μs time‐resolved fluorescence. Data are mean ± SD from four replicates.

A standard curve of DyCl3 concentrations (0 to 1.5 μM) with 0.25‐M Na2WO4 and fluorescence emission at 480 and 580 nm showed a linear relationship (R 2 > 0.99) across the concentrations tested (Figure 1B,C). A significant Dy signal was detected at concentrations as low as 0.05‐μM DyCl3, with a robust and linear response from 0.1 μM in DyCl3 solutions (Figures 1B,C and S1E).

3.2. Mitigation of the Background Effects of the Plant Matrix by Fluorescence Lifetime

We tested whether Dy fluorescence was detectable in plant tissues. Homogenized leaf and stem tissue from Phytolacca acinosa , a species known to preferentially accumulate Dy over low REEs (Grosjean et al. 2019), was mixed with 5‐ and 500‐μM DyCl3 and 0.25‐M Na2WO4. Dy was detectable at both concentrations (Figure 2A), but high autofluorescence was observed in the control plant tissues without Dy, particularly at 480 nm. The autofluorescence from endogenous plant molecules (Donaldson 2020) confounded the Dy signal at 480 nm and made detection at 5‐μM concentration challenging. Implementing time‐resolved fluorescence removed the background signal (Figure 2B). Since Dy has a microsecond‐range fluorescence lifetime (Berezin and Achilefu 2010), delays of 20 to 80 μs after excitation substantially reduced the background signal from the plant matrix while still allowing Dy detection (Figure S2).

FIGURE 2.

FIGURE 2

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Dy screening in plant matrix with time‐resolved fluorescence. (A, B) Dy emission spectra of 0‐, 5‐, and 500‐μM DyCl3 with 0.25‐M Na2WO4 in plant matrix (A) without and (B) with 60‐μs time‐resolved fluorescence at 100 gain. Lines indicate the mean measured value from five technical replicates.

We tested DyCl3 concentrations (0.05–25.6 μM) mixed with homogenized P. acinosa tissue and 0.5‐M Na2WO4 to evaluate if Dy fluorescence is quantitative. We could detect the canonical Dy signal in the plant matrix background at concentrations of 0.05 μM (8.1 μg/L), with a robust signal starting from 0.2 μM (32.5 μg/L) (Figure S3). The relative fluorescence units (RFU) at 480 and 580 nm after 275‐nm excitation responded linearly to the Dy concentration (R 2 > 0.99) (Figure 3A,B). ICP‐MS results from the same sample confirmed these fluorescence measurements, with the fluorescence signal correlating with the ICP‐MS determined concentrations (R 2 > 0.99) (Figure 3C,D). Thus, Dy can be quantitatively detected at concentrations as low as 0.2 μM in a plant tissue background using fluorescence spectroscopy.

FIGURE 3.

FIGURE 3

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Dy detection in plant matrix by fluorescence spectroscopy. (A, B) Scatter plot and R‐squared values of Dy concentrations between 0.05 and 25.6 μM (8.1 to 4160 μg/L) and 0.5‐M Na2WO4 added to the plant matrix and the Dy emission signal at (A) 480 and (B) 580 nm. (C, D) Scatter plot and R‐squared values between Dy concentrations measured by ICP‐MS and the Dy emission signal at (C) 480 and (D) 580 nm. Measurements were taken with an excitation wavelength of 275 nm, 100 gain, and 30‐μs time‐resolved fluorescence. Error bars indicate the standard deviation from four to five technical replicates.

3.3. Screening Plant Leaf Tissue for Dy Uptake Using Fluorescence Spectroscopy

We used time‐resolved fluorescence to detect Dy in the leaves of plants grown in Dy solutions (Figure 4). P. acinosa plants were grown in modified Hoagland's solution supplemented with 0‐, 5‐, 10‐, 15‐, and 20‐mM DyCl3 for 7 days. To evaluate uptake in the above‐ground tissue, we took leaf discs (2 mm) from either the midrib to represent the vascular tissue or from the lamina, avoiding the veins to represent nonvascular tissue. Leaf discs were treated with 0.5‐M Na2WO4 and excited at 275 nm. The emission spectrum (450 to 700 nm) showed Dy signals in leaves treated with 5‐mM DyCl3 and higher but not in control plants (Figure S4). Dy fluorescence at 580 nm increased with the Dy concentrations provided to the plants (Figure 4), indicating the method can screen for Dy translocated from roots to the leaves. Higher levels of Dy were consistently observed in the midrib compared to the lamina tissue.

FIGURE 4.

FIGURE 4

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Dy screening of REE accumulation in plants from leaf discs. (A, B) Dy fluorescence at (A) 480 and (B) 580 nm from the leaf discs of P. acinosa plants grown in 0.25× Hoagland solution (without KH2PO4) supplemented with 0‐, 5‐, 10‐, 15‐, and 20‐mM DyCl3 for 7 days. Two discs, one from leaf lamina and another from midrib, were cut from four different plants and treated with 0.5‐M Na2WO4. Dy fluorescence was detected from the leaf discs with a 275‐nm excitation wavelength, 100 gain, and 60‐μs time‐resolved fluorescence. Data are mean ± standard deviation.

3.4. Evaluation of Dy Levels in Living Plants by Fluorescence Spectroscopy

We designed a method to detect Dy fluorescence directly from intact plant leaves in the field (Figure 5). This portable method does not implement time‐resolved fluorescence, so plant autofluorescence is included, which is a limitation of this implementation. P. acinosa plants were treated with 10‐ and 50‐mM DyCl3 or water (control) for 2 days. Using 248‐nm excitation, Dy emissions at 480 and 575 nm increased with higher Dy treatments (Figure 5A). The plate reader quantified Dy emission at 580 nm, whereas the portable detector measured emission at 575 nm, both capturing the same characteristic Dy emission band (~575–580 nm). Autofluorescence was detected in control plants, especially at 480 nm (Figure S5A), but plants treated with DyCl3 were distinguishable from controls and each other (10 vs. 50 mM) (Figures 5A and S5A).

FIGURE 5.

FIGURE 5

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Nondestructive screening of Dy in plant tissue. (A) Box and whisker plots of Dy emission at 575‐nm wavelength after excitation at 248 nm in the tissue of plants treated with DyCl3. Each box represents a measurement from five biological replicates (individual plants). (B) Point plot of Dy emission at 575‐nm wavelength after 248‐nm excitation and Dy concentrations measured by ICP‐MS represented by a color gradient. Yellow‐hued colors represent lower concentration values as determined by ICP‐MS, and blue represents high concentration values. Each point represents an individual plant.

After the fluorescence measurement, leaves were analyzed by ICP‐MS. At higher Dy concentrations, measurements by both Dy emission and ICP‐MS showed increased variability in Dy detection between replicates (Figures 5B and S5B,C). A positive correlation was found between Dy emission and ICP‐MS measurements (Figures 5B and S5C). Variability in these measurements reflects biological heterogeneity in Dy uptake among independently grown wild plants, as fluorescence measurements were performed on single leaf discs while ICP‐MS analyses were conducted on pooled tissue samples from the same plants.

Variations in the plant autofluorescence also contribute to differences in the background signal. Developing a portable device with delayed fluorescence would reduce this background noise. Although further improvements to portable detection will increase the accuracy, this system offers a rapid first‐pass screening to identify plants with high Dy uptake levels.

3.5. Detection of Dy at a Standoff Distance

The Dy fluorescence signal in the presence of Na2WO4 is bright and visible to the eye. Therefore, since imaging at a distance could increase the high‐throughput screening, we tested the ability to detect the Dy uptake in plants at a standoff distance. We treated P. acinosa leaves with 100‐mM DyCl3 for 3 h and induced tissue penetration of Na2WO4 by piercing the leaves several times with a thin needle (0.45 mm × 13 mm) and adding 3 μL of 2‐M Na2WO4 on the pierced area. We illuminated the leaf with an LED of 275 nm and took a picture with a smartphone in a dark box. Control samples included leaves treated with water (+/− Na2WO4) or Ce (+/− Na2WO4). The Dy fluorescence is already evident at 3 h in plants exposed to this high Dy concentration from the smartphone pictures of the leaf areas treated with Dy and Na2WO4 (Figure 6). There is no discernible fluorescence from the controls with Ce, water, or Dy without Na2WO4. This demonstrates that Dy detection can be visualized at a stand‐off distance, which could facilitate rapid, high‐throughput screening by remote monitoring.

FIGURE 6.

FIGURE 6

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Standoff detection of Dy uptake in plant leaves. Fluorescence from P. acinosa leaves incubated with water (H2O), Ce, and Dy, captured by a smartphone camera. Image panels are as follows: Top panel, − Na2WO4, − LED: an image of punctured leaves before Na2WO4 treatment and 275‐nm LED exposure. Middle panel, − Na2WO4, + LED: an image of punctured leaves, before Na2WO4 treatment exposed to the 275‐nm LED light. Bottom panel, + Na2WO4, + LED: an image of fluorescence from punctured leaves treated with Na2WO4 exposed to 275‐nm LED light. Arrows indicate where Na2WO4 treatments were placed.

4. Discussion

Phytomining is a promising technique for enhancing the recovery of REEs while minimizing the environmental footprint associated with traditional mining methods. The number of species identified as suitable for metal accumulation is limited. For example, Liu et al. identified only 22 plant species as REE hyperaccumulators or potential REE hyperaccumulators (Liu, Yuan, et al. 2021). To advance REE phytomining, it is crucial to identify plants that can absorb unusually high REE concentrations and to understand how environmental factors and microbial interactions influence REE uptake. Strategies to improve plant uptake through genetic modification, microbial treatments, or other environmental factors will further enhance phytomining efforts. Critically, the selection of improved genotypes and environmental parameters that improve REE uptake and specificity is limited by the need for rapid, high‐throughput screening methods. Rapid screening methods to identify REE uptake will accelerate the assessment of natural and engineered genotypes and enhance our understanding of how external factors affect REE uptake by plants.

Fluorescence spectroscopy is a fast method for quantifying Dy levels in plant tissue. We quantified the blue Dy band at 480 nm and the yellow Dy band using 580 nm on the plate reader and 575 nm on the portable device, capturing the same 575–580‐nm emission feature. Using a plate reader for these assays offers the advantage of time‐resolved fluorescence, which overcomes the challenges of the autofluorescence produced by the plant. Dy produces a longer‐lived luminescence than the compounds responsible for plant autofluorescence (Figure 2) (Hagan and Zuchner 2011). This method can differentiate Dy levels in various parts of the leaf, such as the lamina and the midrib, providing insights into the localization and distribution of Dy within different plant organs (Figure 4). This screening can be done with only a small tissue sample provided by a 2‐mm diameter leaf punch. This approach enables rapid screening of Dy emission in the tissue while preserving the rest of the plant's aerial parts.

Our UV fluorescence approach also enables rapid, nondestructive detection of Dy in plant tissue in field settings (Figure 5). This nondestructive sampling can facilitate monitoring the same plant across time, allowing insights into dynamics and how different environmental treatments affect Dy uptake. Nondestructive Dy monitoring can facilitate tracking Dy levels at different growth stages and how Dy is distributed across different tissues as the plant matures. Time‐course monitoring of individual plants can improve the accuracy of studies examining how Dy uptake is affected by environmental changes, such as the addition of microbes or alterations to the soil environment. These findings can aid in optimizing growth conditions to enhance Dy uptake. Additionally, continuous Dy monitoring can help determine the optimal harvesting time to maximize Dy recovery, ensuring plants are collected when Dy concentrations peak.

The present instrument we used for field‐based assays lacks time‐resolved fluorescence. Therefore, the plant's autofluorescence can interfere with the Dy signal. It is crucial to account for autofluorescence when comparing plant species, as different plants have varying background signals that can change in response to environmental changes (Donaldson 2020). Although currently limited by the lack of time‐resolved fluorescence, this field‐based evaluation can be used as an initial screen. Leaf discs collected from the top Dy accumulation candidates identified by this screen can be evaluated using time‐resolved fluorescence in the laboratory. Developing a portable device with time‐resolved capability would enhance sensitivity and accuracy for nondestructive Dy detection in the field and herbarium samples, similar to advancements achieved with XRF (Goudard et al. 2024).

Additionally, the emission from Dy complexed with Na2WO4 is sufficiently bright to be visible to the naked eye (Figure 6), enabling practical applications for remote imaging and high‐throughput, nondestructive screening of large plant populations. Standoff detection can be achieved using a phenotyping device to measure plant Dy uptake from a distance after treating the plants with Na2WO4. This method is beneficial for detecting plants that accumulate high concentrations of Dy, as the visible emission will highlight those plants more effectively. A key limitation is that detecting the Dy emission remotely can be affected by ambient light levels. The images presented here were acquired in a dark enclosure to demonstrate the visualization of Dy fluorescence. Additional imaging optimization will be required to measure the Dy levels in field conditions. This approach would simplify and improve the efficiency of plant screening to identify high Dy‐accumulating plants in research and agricultural settings.

This investigation focused on developing a quick method to screen Dy in plant tissue. The plants grew in high Dy concentrations that were applied hydroponically in the form of DyCl3. This system allowed us to focus on Dy detection and eliminate variability in uptake due to environmental variables. However, this is an artificial delivery of REEs (van der Ent and Rylott 2024). This delivery system demonstrates that the detection correlates with measures by ICP‐MS and can identify plants with uptake from those without. The ability to measure down to 0.2 μM in the plant background indicates that this detection method can screen Dy in plants and potentially identify new Dy hyperaccumulators. Plants grown in environments with high REE levels can accumulate REEs to these levels. For example, in mining areas where REEs are enriched, D. linearis plants had Dy concentrations of 113.2 μM (18.4 μg/g) in the stem and 227.7 μM (37.0 μg/g) in the leaves (Wei et al. 2001). Even in areas with no known REE contamination, plants have been identified with Dy levels above our detection minimum. In Ehime, Japan, D. linearis plants had Dy concentrations in their aerial tissue of ~14 μM (2.4 μg/g), Athyrium yokoscence plants had ~53 μM (8.7 μg/g), and P. americana plants had ~73 μM (12 μg/g) (Ichihashi et al. 1992). However, we have demonstrated the feasibility only in one species, P. acinosa . REE uptake and distribution vary not only across species but also depending on the environment in which the plant is grown (Gmur et al. 2025). It is essential to acknowledge that this sensitivity might differ in woody species and may be affected by environmental factors. Expanding these studies to include additional species would help clarify both the limitations and the wider relevance of this approach.

Fluorescence assays can be expanded to screen for other REEs in plant tissue. REEs exhibit a wide range of emission characteristics across the UV–Vis–NIR spectrum, which makes them detectable using fluorescence spectroscopy. Further investigation is needed to understand how Dy and other REE signals behave in the presence of sodium tungstate, the interaction between emissions of different REEs, and strategies to mitigate such effects. The extremely long fluorescence lifetime of most REEs can be leveraged to eliminate the interference from most other elements, like transition metals; however, interference from other REEs might require alternative strategies. While the fluorescence assay has been demonstrated in P. acinosa , its applicability to other plant species remains unexplored. Expanding the evaluation to additional species will be important for confirming the reproducibility and generalizability of the assay. Although many questions remain, fluorescence spectroscopy offers new opportunities for research into REE uptake by plants and for advancing phytomining techniques.

5. Conclusion

In conclusion, this work demonstrates the development of rapid and nondestructive methods for detecting Dy in plant tissues. The ability to quantitatively assess Dy levels in living plant tissues represents a significant advancement in phytomining. The use of fluorescence spectroscopy offers promising avenues for improving the efficiency and effectiveness of phytomining research. By quickly identifying plants with high REE uptake, these methods can accelerate the screening and selection of potential hyperaccumulators, facilitating further research and optimization of phytomining practices. Evaluating the response of the same plant across time will inform how environmental factors affect Dy uptake. The unique spectral characteristics of other REEs suggest that this method could be adapted to quantify the uptake of several different REEs. Expanding these assays to include other REEs could significantly enhance our understanding of plant–REE interactions and contribute to more sustainable and environmentally friendly mining solutions. In summary, adapting fluorescence spectroscopy for REE detection can enhance phytomining research by enabling the rapid quantification and localization of REEs, paving the way for more sustainable and efficient resource recovery practices.

Author Contributions

M.W.K. and C.J.D. designed the initial project. E.H‐P., A.T.H., and K.L. optimized the detection assays. E.H‐P., A.T.H., A.N.H., C.R., D.B., A.G., and M.W.K. performed the measurements. M.W.K. developed the imaging system used for Figures 5 and 6. E.H‐P., M.W.K., and C.J.D. prepared the initial draft. All authors contributed to the writing and review of the manuscript.

Funding

This work was supported by DARPA Young Investigator Award (D19AP00026), NC State Research and Innovation Seed Funding Program, and the Office of Surface Mining, Reclamation, and Enforcement (S24AC00044‐00).

Ethics Statement

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Dy absorbance spectra and emission signals. (A) Dy absorbance spectra scanned from 250 to 500 nm at 1‐nm intervals for 100‐mM DyCl3. (B) Dy absorbance spectra from 100‐mM DyCl3 in aqueous solution scanned from 250 to 700 nm at 10‐nm intervals. Lines indicate the mean measured value from four to five technical replicates. (C and D) Dy emission from 255‐, 265‐, 275‐, 295‐, 325‐, 350‐, 365‐, 390‐, 425‐, and 450‐nm excitation wavelengths from 0‐ and 0.8‐μM DyCl3 in aqueous solutions with 0.25‐M Na2WO4. Emission was measured at (C) 480 and (D) 580 nm using a 100 gain and 60‐μs time‐resolved fluorescence. Error bars indicate the standard deviation from four to five technical replicates. (E) Dy emission spectra from 0‐ to 1‐μM DyCl3 solution and 0.25‐M Na2WO4 at 275‐nm excitation, 150 gain, and 20‐μs time‐resolved fluorescence. The vertical dashed lines indicate 480‐ and 580‐nm emission wavelengths. The blue‐shaded ribbon indicates the standard deviation from four technical replicates.

PLD3-10-e70164-s003.tiff (3.4MB, tiff)

Figure S2: Dy screening in plant matrix with time‐resolved fluorescence. Time‐resolved fluorescence lengths of 0, 20, 30, 40, 50, 60, and 80 μs were tested to evaluate the detection of Dy in the presence of P. acinosa tissue. A solution of 3.2‐μM Dy and 0.5‐M Na2WO4 was mixed with P. acinosa homogenized leaf and stem tissue. Dy fluorescence was measured with a 275‐nm excitation wavelength and 150 gain. Lines indicate the mean measured value from five technical replicates.

PLD3-10-e70164-s001.tiff (92.2MB, tiff)

Figure S3: Dy emission in plant matrix. Dy emission spectra of 0.05 to 25.6 μM (10 to 5158 μg/L) DyCl3 concentrations with 0.5‐M Na2WO4 in plant matrix. Measurements were taken with an excitation wavelength of 275 nm and 100 gain with 30‐μs time‐resolved fluorescence. The blue‐shaded ribbon indicates the standard deviation from four to five technical replicates.

PLD3-10-e70164-s005.tiff (3.4MB, tiff)

Figure S4: Dy emission spectra of REE uptake in plants from leaf discs. Leaf discs (2 mm) were taken from P. acinosa plants grown in 0.25× modified Hoagland solution supplemented with 0‐, 5‐, 10‐, 15‐, and 20‐mM DyCl3 for 7 days. Two discs, one from leaf lamina and another from midrib, were cut from four different plants and treated with 0.5‐M Na2WO4. Dy fluorescence was measured from the leaf discs with a 275‐nm excitation wavelength and 100 gain with 60‐μs time‐resolved fluorescence. The vertical dashed lines indicate 480‐ and 580‐nm emission wavelengths.

PLD3-10-e70164-s004.pdf (187.5KB, pdf)

Figure S5: Nondestructive Dy detection in plant tissue. (A) Box and whisker plots of Dy emission at 480‐nm wavelength after excitation at 248 nm in the tissue of plants treated with DyCl3. (B) Box and whisker plots of Dy concentrations in plants treated with DyCl3 measured by ICP‐MS. (C) Point plot of Dy emission at 480‐nm wavelength after 248‐nm excitation and Dy concentrations measured by ICP‐MS represented by a color gradient. Yellow‐hued colors represent lower concentration values as determined by ICP‐MS, and blue represents high concentration values. Each box represents a measurement from five biological replicates. Each point represents an individual plant.

PLD3-10-e70164-s002.pdf (83.1KB, pdf)

Acknowledgments

We would like to thank Kendrick Turner and Blake Bextine for their advice on assay development and Elizabeth Rylott and Thomas Makris for their guidance and advice on manuscript preparation.

Contributor Information

Michael W. Kudenov, Email: mwkudeno@ncsu.edu.

Colleen J. Doherty, Email: colleen_doherty@ncsu.edu.

Data Availability Statement

All data generated or analyzed during this study are included in this published article (and its Supporting Information files).

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1: Dy absorbance spectra and emission signals. (A) Dy absorbance spectra scanned from 250 to 500 nm at 1‐nm intervals for 100‐mM DyCl3. (B) Dy absorbance spectra from 100‐mM DyCl3 in aqueous solution scanned from 250 to 700 nm at 10‐nm intervals. Lines indicate the mean measured value from four to five technical replicates. (C and D) Dy emission from 255‐, 265‐, 275‐, 295‐, 325‐, 350‐, 365‐, 390‐, 425‐, and 450‐nm excitation wavelengths from 0‐ and 0.8‐μM DyCl3 in aqueous solutions with 0.25‐M Na2WO4. Emission was measured at (C) 480 and (D) 580 nm using a 100 gain and 60‐μs time‐resolved fluorescence. Error bars indicate the standard deviation from four to five technical replicates. (E) Dy emission spectra from 0‐ to 1‐μM DyCl3 solution and 0.25‐M Na2WO4 at 275‐nm excitation, 150 gain, and 20‐μs time‐resolved fluorescence. The vertical dashed lines indicate 480‐ and 580‐nm emission wavelengths. The blue‐shaded ribbon indicates the standard deviation from four technical replicates.

PLD3-10-e70164-s003.tiff (3.4MB, tiff)

Figure S2: Dy screening in plant matrix with time‐resolved fluorescence. Time‐resolved fluorescence lengths of 0, 20, 30, 40, 50, 60, and 80 μs were tested to evaluate the detection of Dy in the presence of P. acinosa tissue. A solution of 3.2‐μM Dy and 0.5‐M Na2WO4 was mixed with P. acinosa homogenized leaf and stem tissue. Dy fluorescence was measured with a 275‐nm excitation wavelength and 150 gain. Lines indicate the mean measured value from five technical replicates.

PLD3-10-e70164-s001.tiff (92.2MB, tiff)

Figure S3: Dy emission in plant matrix. Dy emission spectra of 0.05 to 25.6 μM (10 to 5158 μg/L) DyCl3 concentrations with 0.5‐M Na2WO4 in plant matrix. Measurements were taken with an excitation wavelength of 275 nm and 100 gain with 30‐μs time‐resolved fluorescence. The blue‐shaded ribbon indicates the standard deviation from four to five technical replicates.

PLD3-10-e70164-s005.tiff (3.4MB, tiff)

Figure S4: Dy emission spectra of REE uptake in plants from leaf discs. Leaf discs (2 mm) were taken from P. acinosa plants grown in 0.25× modified Hoagland solution supplemented with 0‐, 5‐, 10‐, 15‐, and 20‐mM DyCl3 for 7 days. Two discs, one from leaf lamina and another from midrib, were cut from four different plants and treated with 0.5‐M Na2WO4. Dy fluorescence was measured from the leaf discs with a 275‐nm excitation wavelength and 100 gain with 60‐μs time‐resolved fluorescence. The vertical dashed lines indicate 480‐ and 580‐nm emission wavelengths.

PLD3-10-e70164-s004.pdf (187.5KB, pdf)

Figure S5: Nondestructive Dy detection in plant tissue. (A) Box and whisker plots of Dy emission at 480‐nm wavelength after excitation at 248 nm in the tissue of plants treated with DyCl3. (B) Box and whisker plots of Dy concentrations in plants treated with DyCl3 measured by ICP‐MS. (C) Point plot of Dy emission at 480‐nm wavelength after 248‐nm excitation and Dy concentrations measured by ICP‐MS represented by a color gradient. Yellow‐hued colors represent lower concentration values as determined by ICP‐MS, and blue represents high concentration values. Each box represents a measurement from five biological replicates. Each point represents an individual plant.

PLD3-10-e70164-s002.pdf (83.1KB, pdf)

Data Availability Statement

All data generated or analyzed during this study are included in this published article (and its Supporting Information files).

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