Populus tremula x alba (clone INRA 717-1B4) plants were propagated from ~1 cm cuttings of in vitro plantlets and grown in half-strength Murashige-Skoog medium (Caisson Laboratories, UT, USA) supplemented with 0.5 μM indole-3-butyric acid (IBA). Plantlets were grown in culture in Magenta boxes for a minimum of eight weeks until they were approximately 8 cm in height. Plantlets were then transplanted into small seedling pots containing vermiculite, covered, and acclimated for three weeks in a mist-chamber. They were then repotted into vermiculite-filled, one-gallon round pots and moved into the greenhouse and grown for approximately 8 weeks. Plants used for histochemistry were fertilized three times per week with 100 mL of general purpose 20-20-20 Plant Prod^® NPK fertilizer (100 ppm nitrogen, phosphorus and potassium; Plant Products Co. Ltd, Brampton, ON, Canada), and watered with an equal volume of distilled H₂O. Plants used for quantitative CT analysis and the nitrogen experiment were fertilized three times per week with a modified Long Ashton’s solution optimized for poplar growth (1 mM NH₄NO₃, 0.9 mM CaSO · 2H₂O, 0.6 mM KH₂PO₄, 0.5 mM KCl, 0.04 mM K₂PO₄, 0.33 mM MgCl₂ · 7 H₂O, and 0.03 g L-1 standard micronutrient mix (Plant Products Co. Ltd, Brampton, CT ON, Canada), pH 5.6).

For N manipulation experiments, a P. tremula x tremuloides hybrid (clone INRA 353-38) was used. Plants propagated as above were grown in Sunshine Basic Mix #2 (Sungro, Seba Beach, AB, Canada) under three nitrogen fertilization treatments. Three plants were assigned to each nitrogen condition: 0.1 mM, 1 mM or 10 mM NH₄NO₃ considered to be low, medium (normal) and high nitrogen treatments, respectively. The plants were fertilized on alternate days with 100 mL of the desired NH₄NO₃ concentration within modified Long Ashton’s nutrient solution (0.9 mM CaCl₂, 0.6 mM KH₂PO₄, 0.5 mM KCl, 0.04 mM K₂HPO₄, 0.3 mM MgSO₄ x 7H₂O and 0.03 g L^-1 standard micronutrient mix (Plant Products Co. Ltd, Brampton, ON, Canada), pH 5.6). All other concentrations of nutrients were standardized in these solution. Plants were grown in greenhouse conditions for 12 weeks and watered with dH₂O as necessary.

Roots were harvested after eight weeks of growth, divided into segments according to distance from root tip and root color, and then pooled into groups for each plant (n>30). Samples were frozen immediately in liquid nitrogen and then freeze-dried for three days. An aliquot of 8 mg was weighed from each pooled sample. For some samples (mainly root tips) where material was limited, the entire sample was weighed and extracted and the precise weight used in final calculations. Pooled root samples were extracted thrice in 100% MeOH by grinding freeze-dried samples in a PreCellys 24 homogenizer for two minutes using three 2 mm steel beads per sample, sonicating for ten minutes and centrifuging for five minutes at 1500 rpm. This procedure was followed using 1 x 1.5 mL, and 2 x 1 mL of 100% MeOH resulting in 3.5 mL of extract. The extract volume was made up to 5 mL to ensure it was within the linear range for soluble CT quantification. Soluble CTs were measured using the butanol-HCl assay (Porter et al., 1986) as described previously (Gourlay and Constabel, 2019). CTs purified from P. tremula x tremuloides leaf tissue (mean degree of polymerization ~9) was used as a standard, isolated and characterized as described by Preston and Trofymow (2015).

Insoluble CTs were quantified by depolymerizing CTs directly in the previously-extracted pellet. Six mL of 1-butanol:HCl (95:5) and 200 μL Fe reagent (Porter et al., 1986) were added directly to the dried pellet and heated to 95 oC for 40 minutes. 400 μL of MeOH was added to each sample before heating to correct for the volume of soluble extract used in the standard curve. Spectrophotometer readings were converted to CT concentration (μg mL-1) for the total sample using a standard curve prepared using purified Populus CTs (Mellway et al., 2009).

Roots were carefully removed from plants, washed to remove potting soil, and cut into 1 cm sequential segments beginning at the root tip. Samples were immediately fixed at room temperature in modified Karnovsky’s fixative (25% glutaraldehyde and 16%) paraformaldehyde in 0.5 mM sodium phosphate buffer (PBS), pH 7.4). Samples were stored in fixative at 4 °C for up to four weeks before embedding. Samples were then rinsed three times for 30 minutes each with 0.5 mM PBS, pH 7.4, and dehydrated in sequential ethanol rinses (30%, 50%, 70%, 95%, and 100%) over the course of one day. Throughout the following week, plants were infiltrated with increasing concentrations of Technovit^® 7100 Glyco Methycrylate (Electron Microscopy Sciences) in 100% ethanol, before being cured.

Longitudinal sections (5 μm) were cut using a glass blade microtome (Sorval JB-4). Sections were dried onto slides and stained with 0.1% 4-dimethylaminocinnamaldehyde (DMACA) (w/v) in 0.5 M sulphuric acid in 1-butanol according to the method of Gutmann and Feucht (1991). The slides were covered in 1 mL of stain solution and heated on a Thermolyne Type 1000 hot plate (setting 2.5) for two minutes or until the stain began to evaporate and discolor. The slides were removed from the heat before bubbling occurred in the resin, rinsed three times for 30 s in 1- butanol or until all unbound stain was removed, and cover slips were then secured with PermountTM mounting medium. Sections were imaged within 24 hours on a transmitted light microscope (Zeiss [47-30-12-9902], Germany) fitted with a SPOT RTKE diagnostic 7.2 Color Mosaic camera. A deep burgundy stain indicated the presence of CTs, consistent with other reports of DMACA stained CTs in embedded samples hydrolyzed by hot sulphuric acid (Abeynayake et al., 2011; Gutmann, 1993).

Five clonal P. tremula x alba plants were grown for nutrient ion flux analysis. The Long Ashton’s solution was altered from previous experiments to match the measuring solution used during MIFE™ analysis, with overall concentrations of each ion similar to previous studies (Hawkins et al., 2014; 1 mM NH₄NO₃, 0.9 mM CaSO₄ · 2H₂O, 0.6 mM KH₂PO₄, 0.5 mM KCl, 0.04 mM K₂PO₄, 0.3 mM MgCl₂ · 7 H₂O and 0.03 g L^-1 standard micronutrient mix, Plant Products Co. Ltd, Brampton, ON, Canada, pH 5.6). Plants were grown in pairs in a continuous rotation to ensure that they were all the same age when harvested after six weeks of growth. Healthy, individual roots were excised from the plant, and placed in aerated solution containing 500 μM NH₄NO₃ and 500 μM CaSO₄ for 30 minutes prior to measurement to allow the roots to acclimate.

Borosilicate glass capillaries (1.5 mm diameter) were pulled, dried at 200 °C for five hours and silanized with tributylchlorosilane to make sterile, measuring electrodes. Once cooled, electrodes were backfilled with 200 mM NH₄Cl for NH₄ ⁺, 500 mM CaCl₂ 2H₂O for Ca²⁺, and 500 mM KNO₃ 100 mM KCl for NO₃ ^-. Electrode tips were then filled with ion-selective resins: NH₄ ⁺ and Ca²⁺ -selective cocktails (Fluka), and NO₃ ^–selective cocktail containing 0.5% methyltrididecylammoniumnitrate (MTDDA NO₃ ^-), 0.084% methyltriphenylphosphonium (MTPPB) and 99.4% n-phenyloctylether (NPOE) (Plassard et al., 2002). Electrodes were calibrated using a set of standards at pH 5.6. A reference electrode filled with 100 mM KCl in 1% agar and electrolyzed, chloride silver wire was placed in the solution for each measurement to complete the electrical current. Calibration was conducted at the beginning and end of each day to ensure readings were representative and reliable.

The electrodes were mounted in a specialized holder (MMT-5, Narishige, Tokyo, Japan) providing three-dimensional positioning. A maximum distance of 20 μm was recorded between the electrodes and the root surface, and 4 μm between the electrode tips. Roots were secured and placed in the measuring chamber with fresh measuring solution (500 μM NH₄NO₃ and 500 μM CaSO₄). The chamber was moved by a computer-controlled manipulator (PatchMan NP2, Eppendorf AG, Hamburg, Germany). During flux measurements, the MIFE™ computer caused the chamber to gently oscillate over a distance of 40 μm, back and forth from the root surface in a 10 s square-wave cycle. The concentration of each ion was calculated from its electrochemical potential at the two positions and ion net flux calculated from the difference in concentration between the two positions (Shabala et al., 1997).

Flux measurements were taken at four discrete positions along the root in the white zone: at the root tip (<2 mm), and three positions within the maturation zone of the white root zone (0.5 cm, 1.5 cm and 3.5 cm from the root tip). These root distances were chosen as they included the zones where uptake is expected to be highest (Hawkins et al., 2008; 2014).

Data was analyzed and graphed using R v.3.0.3. Once model assumptions and normality were checked, data was statistically analyzed using a two-sample T-test or one-way analysis of variance (ANOVA) with Tukey’s Honestly Significant Difference (HSD) test.

In order to determine the overall distribution of CTs in roots, root systems from greenhouse-grown poplar were carefully removed from pots and soil and root zones separated and dissected. Root systems were white at the actively growing regions, with older sections becoming brown, and developing obvious secondary growth towards the crown ( Figure 1 ). We used both color and root diameter to divide root systems into four zones similar to the classification of Peterson et al. (1999): root tips (meristematic and elongation zone; <0.5 cm), white zone (maturation zone; 1.0-12 cm), brown zone (without periderm; 13-20 cm), and cork zone (with periderm; >20 cm). Any root tips that showed necrosis were excluded.

The sum of soluble and insoluble CT concentration was highest in the root tips and within the white zone, and then decreased with increasing distance from the root tip ( Figure 2 ). In the youngest parts of the roots (< 5 cm from the root tip), CTs accumulated to more than 100 mg/g DW; this is as much as 20-fold greater than the CT concentration in leaves of greenhouse grown plants. Older parts of the root system contained approximately 25-50 mg/g DW. The lowest CT content was found in the oldest zone, which showed secondary growth - the cork zone. Soluble CTs comprised the predominant fraction of CTs in all regions except in the cork zone, where insoluble CTs were predominant ( Figure 2 ).

To further define areas of CT accumulation at the cellular level, embedded tissues from excised root samples from different root zones were stained with 4-dimethylaminocinnamaldehyde (DMACA). In embedded tissue sections, this stain colors CTs reddish brown rather than blue (Abeynayake et al., 2011). In our root sections, CTs were strongly stained in several cell layers of the root cap, but not in the underlying smaller meristematic cells ( Figure 3A ). In the white zone immediately adjacent (~ 1 cm from tip), CTs were detected in the epidermal cell layers just behind the root cap. Staining was also seen sporadically in cortical cells ( Figure 3B ). Some DMACA positive staining was observed in the endodermis, but not within vascular tissue of the stele. In older roots, where secondary xylem expands laterally and begins to disrupt the cortex, only very low levels of staining were observed ( Figure 3C ). Overall, the pattern of DMACA staining was consistent with the chemical analysis: the most intense staining of CTs was seen in the tips and younger sections of the roots, predominantly in epidermal and subepidermal regions around the tip.

At higher magnification, distinct patterns of CT accumulation distribution were visible within different cell types within the white zone. In the outer epidermal layer of the young white root, CT staining was typically observed in large vacuoles filling most of the intracellular space ( Figure 4 ). By contrast, in cortical cells below the epidermal layer, CTs tended to occur in smaller, more concentrated vacuole-like structures. In many cases, these CT-containing structures were arranged around the periphery of the cells, possibly around a central zone or structure, or were found throughout the cell. CTs were not present in all cortical cells but occurred somewhat sporadically, as we had noted at lower magnification ( Figure 3 ). This pattern was observed consistently in multiple sections observed within this region.

MIFE™ technology is a non-invasive method to measure net fluxes of specific nutrient ions at precise root locations (Shabala and Newman, 1997; Shabala et al., 1997). To determine if there is a potential interaction of CTs with N uptake, we looked for correlations of N fluxes with CT accumulation. We measured the net flux of NO₃- and NH₄+ along the longitudinal axis of young poplar roots using a MIFE system. Since the root tip and youngest part of the root is most active in nutrient acquisition (Hawkins et al., 2014), we focused on the white zone and root tip. Ion net fluxes were measured for eight minutes at each of four positions along the root axis to ensure that a reliable average flux measurement was gained for each replicate. Five plants were grown and three roots were measured per plant. The highest NH₄ ⁺ net fluxes were observed at the root tip, which were significantly greater than fluxes at more distal positions ( Figure 5 ). NO₃- uptake was lower than NH₄ ⁺ flux at the root tip and did not vary significantly along the length of the root. We also measured the net fluxes of another cation, Ca²⁺, at the three positions, and these also did not vary with distance from the root tip.

We next tested if CT concentration in the different root zones is influenced by external N supply. Young potted poplar plants were fertilized with a modified Long Ashton nutrient solution that provided 0.1, 1.0 or 10 mM NH₄NO₃ (low, medium, and high N). After the six-week experimental period, the impact of N limitation at the lowest N level was shown by reduced plant growth and a lighter leaf color compared to plants grown at medium and high N availability. Roots were excised and samples taken from the tip, white, and brown zones for CT assays. Under all three N treatments, we observed the same pattern: highest CT concentration at the root tip, which decreased with increasing distance from the tip in the white and brown zones ( Figure 6 ). Within each root zone, the highest CT concentration was measured in roots grown with 0.1mM NH₄NO₃; roots of plants grown under intermediate N levels showed intermediate CT concentration, and the lowest CT concentration was measured for all three root zones in plants supplied with 10 mM NH₄NO₃. In the white zone, CT concentration was two-fold lower at the highest compared to lowest N availability. This confirmed the repressive effect of N on CT metabolism, and demonstrated this effect across all three root zones.