Cellulose biosynthesis in plant cell walls is continually modified to accommodate developmental stages of the plant and to respond to environmental changes. Here we used radioactive isotopes of ¹¹C (t_1/2 20 m), administered to leaves as ¹¹CO₂, and ¹⁸F (t_1/2 110 m), administered to roots as the glucose analogue, 2-deoxy-2-[¹⁸F] fluoro- D-glucose (¹⁸FDG), and developmental leaf aging in Nicotiana tabacum to explore the different contributions of intracellular (¹¹C) sugar versus the exchange of intercellular (¹⁸F) sugar. Leaf cellulose content decreased with leaf age, and was correlated with a decreasing influx of ¹⁸F tracer. However, the incorporation of intracellular ¹¹C sugar into cellulose was unchanged with leaf age, suggesting that the rate limiting step in controlling cellulose biosynthesis was the intercellular exchange of sugar. This hypothesis was tested using known inhibitors of cellulose biosynthesis. Isoxaben, a pre-emergence herbicide and methyl jasmonate, a plant defense hormone, were topically applied to same aged source leaves prior to tracer administration. Treatments exerted opposite effects on sugar transport, as well as opposite effects on ¹¹C and ¹⁸F tracer flux into cellulose. These combined results support the model where cellulose biosynthesis was tightly coupled to intercellular sugar trafficking and not to the state of the metabolic machinery.

Keywords: Intercellular sugar trafficking; cellulose synthesis; developmental aging; carbon-11; ¹⁸FDG;

Plant cell walls are complex macromolecular structures that surround cells and are compositionally rich in polysaccharides, proteins, aromatic compounds and aliphatic compounds. They are highly dynamic compartments, differentiated by cell type and developmental stage, and serve a multitude of important aspects to plant survivability under various environmental conditions [1, 2].

Growing or expanding cells are typically surrounded by a primary wall comprised largely of polysaccharide macromolecules which contribute to the structural integrity of the cell and to cell adhesion. This structure provides an important conduit for solute transfer including growth resources and key signal molecules that help maintain cellular function [3].

In many cells, this wall can be thickened and further strengthened by the addition of a secondary wall containing lignin, or polyaromatic macromolecules that help maintain cellular water conductance. As plant cells continue to grow, becoming more specialized in function and differentiated by the tissue type, the walls that encase them often can possess a vast array of compositional differences that help serve the diverse cellular functions across the plant.

Plants are sessile organisms, and as such have evolved with the capacity to be highly plastic in their responses to growth conditions imposed by a changing environment. This unique feature empowers them with the ability not only to survive harsh conditions, but also to survive predation by herbivores and attack by pathogens. Their ability to respond in this capacity requires that the cell wall structure, and its composition, be continually modified providing physical barriers as a mechanism to ward off attack by predators. For example, strategically situated parenchyma cells will rapidly develop invaginations or reinforcements of their cell wall as a consequence of plant defense [4]. Jasmonic acid (JA) and it’s methyl ester are key hormones that trigger this response by stimulating re-programming of key biochemical pathways providing essential biosynthetic precursors for lignin as a physical barrier [5,6]. In parallel, JAs also mediate reduced biosynthesis of cellulose, the most abundant polysaccharide macromolecular component of cell walls, and likely the most nutritious to feeding herbivores [7, 8].

The highly dynamic nature of cellulose biosynthesis attests to the complexity of the underpinnings that regulate cell wall construction. To this day, that regulation is still not entirely understood, especially in the context of changing cellular functions that coincide with normal plant growth and development [2,3,9,10 and 11]. To properly generate or modify the cell wall, proteins involved in the process, including cellulose synthesis, glucan synthesis and pectin methyl esterase, are transported intracellular via membrane trafficking to the plasma membrane, or the extracellular space [12,13]. It is presumed that such processes must be tightly regulated [14]. Recent studies even go as far as to suggest that the diversification of membrane trafficking is what contributes to cell wall differentiation across all plant cell types [15]. Even so, there remain many open questions concerning the molecular mechanisms that underlie these transport processes. For example, some proteins at the plasma membrane may become endocytosed into the cytoplasm in response to feedback signals associated with extracellular conditions [16].

Unfortunately, very little attention has been given to understanding the dynamics underpinning the exchange of the simple sugar building blocks that are important to cell wall construction and/or modification. Herein, we report on the use of a short-lived radiocarbon isotope, carbon-11 (t_½ 20.4 min), administered to intact leaves in Nicotiana tabacum as 11CO2 to explore the effect of leaf age on the intracellular partitioning of “new carbon” into cell wall cellulose [17]. We also report on the use of the fluorine-18 (t_½ 110 min) radioisotope, administered to plants as 2-[¹⁸F]fluoro-2-deoxy-D-glucose (¹⁸FDG), a radioactive glucose surrogate, to examine the role of intercellular sugar trafficking in cellulose biosynthesis using developmental leaf aging as a model [18]. Over the years, fluorine- 18 has been extremely useful for radio-labeling molecules of interest for use in animal and human research, but in recent years this isotope has found its way into plant science with the radio synthesis of 18F-labeled sucrose analogues to study sugar transport [19, 20]. Additionally, 18FDG has also been shown to actively transport in plant vasculature [21-26] and utilized in secondary metabolism of essential defense compounds [27]. Here, we showed evidence that plants will metabolically assimilate 18FDG into cell wall cellulose and the tracer can be used as a marker for quantifying extracellular glucose flux into cellulose.

Plant Growth: Tobacco plants (Nicotiana tabacum L. cv Samsun) were grown from seeds in commercial potting mix with a slow-release fertilizer (Osmocote; Scotts Company, Marysville, OH, USA) in commercial growth chambers (Conviron, Inc., Winnipeg, Canada) at 24°C with a 16/8 h photoperiod at 350 μmol m^-2 s^-1. At 3 weeks into their growth cycle, seedlings were transferred to aerated hydroponics stations (8 plants per station). The nutrient status in the hydroponics solution was maintained using commercial a Hoagland modified basal salt mixture (PhytoTechnology Laboratories, Shawnee Mission, KS, USA) prepared by dissolving 4.9 g of the salt mix in 3 L of de-ionized water and buffering with 1.66 g MES hydrate. The pH of the solution was adjusted to 6.5 by adding 1N potassium hydroxide solution. Hydroponics solutions were changed on a 5-d cycle. Plants were used for experiments when they had seven fully expanded leaves.

Treatments: All chemicals used in these studies were obtained from Sigma Aldrich (St. Louis, MO, USA) and were used without any further purification. Isoxaben (ISX), N-(3’[1-ethyl-lmethylpropyl]- 5-isoxazolyl)-2,6-dimethoxybenzamide is a preemergence, broad leaf herbicide used primarily on small grains, turf grasses and ornamental plants [28].

This herbicide is extremely active, with IC50 values in the nanomolar range [29]. Isoxaben treatment has been shown to selectively inhibit 14C-glucose incorporation into the acid insoluble cellulosic cell wall fraction [30,31] and so has become a powerful tool enabling the chemical inhibition of cellulose biosynthesis in vivo.

Isoxaben interferes with rosette formation, and by blocking the demonization of CesA polypeptides 130 it inhibits cellulose biosynthesis [32]. Jasmonic acid (JA), including its methyl ester (MeJA), is ubiquitous in all higher plants filling numerous roles in plant growth and development [33] and especially plant defense hormones eliciting rapid reprogramming of plant metabolism when applied topically to tissues [5,34]. Like ISX, MeJA will also inhibit cellulose biosynthesis in vivo. In all tracer flux measurements involving measuring radio labeled cellulose, we applied either a 5 nm solution of ISX or a 5 μM solution of MeJA as topical treatments to upper intact leaf surfaces. Differences in treatment concentrations were based on perceived substrate biological activities. Treatments were applied to a small single layer section of Kim Wipe™ for 1 hr prior to tracer administration (i.e. ¹¹CO₂ to leaves or ¹⁸FDG to roots). In special studies designed to assess the effect of treatment dose on ¹¹C-photoassimilate transport, we administered treatments through the cut petiole of an excised leaf. The amount of treatment taken up by the leaf tissue was quantified by measuring the volume of solution assimilated over time. Treatment solutions were significantly reduced for this experimental protocol because a significant volume of solution was always assimilated by the cut leaf over a 1hr time course.

¹¹CO₂ Production and Leaf Administration: ¹¹CO₂ was produced via the ¹⁴N(p, α)¹¹C nuclear transformation from a 20 ml target filled with high-purity nitrogen gas (400 ml @ STP) using 18MeV protons from the TR-19 (Ebco Industries Ltd, Richmond, BC, Canada) cyclotron at BNL, and captured on a molecular sieve (4Å). The ¹¹CO₂ that was trapped on the molecular sieve was desorbed and quickly released into an air stream at 200 ml/min as a discrete pulse for labeling a leaf affixed within a 5 x 10 cm lighted (350 μmol m^-2 s^-1) leaf cell at 21°C to ensure a steady level of fixation [35, 36]. For mature source leaves, only a portion of the leaf was affixed within the cell. The leaf was positioned in the cell so as to capture part of the outside leaf edge. This configuration allowed efficient air/tracer flow around the upper side of the leaf as well as the underside. The load leaf tissue was exposed to ¹¹CO₂ tracer for 1 minute as a transient pulse in the air stream, and then was chased with clean air for the duration of the exposure. A PIN diode radiation detector (Carroll Ramsey Associates, Inc, Berkeley, CA, USA) affixed to the bottom of the leaf cell enabled continuous measurement of radioactivity levels within the cell during the pulse.

¹⁸FDG Synthesis and Root Administration: Fluorine-18 as fluoride anion (¹⁸F-) was produced using an 18 MeV proton beam on the TR-19 cyclotron (Ebco Industries Ltd, Richmond, BC, Canada). This beam was focused onto an oxygen-18 enriched water target causing the ¹⁸O (p, n) ¹⁸F nuclear reaction to occur. After irradiation, ¹⁸F-fluoride was collected on a QMA sep-pak (Waters Accell Plus QMA) located at the target station, then extracted and transferred under pressure to the radiochemistry laboratory using a dilute solution of acetonitrile and potassium bicarbonate (0.1 N). The contents of the transfer were received into a FDG-Plus™ Automated Radiochemistry Module (Bioscan, Inc., Washington, DC, USA) where the ¹⁸F-fluoride was dried under vacuum with Kryptofix 2.2.2. ¹⁸F-Fluoride was then reacted with mannose trifoliate (1, 3, 4, 6-tetra-O-acetyl-2-O-trifluoromethane sulfonyl-β-D-mannopyrannose). Reaction by nucleo philic displacement with the trifoliate starting material yielded an intermediate radio labeled product in high yield that could be hydrolyzed easily in base and at room temperature to yield 2-deoxy-2- [¹⁸F]fluoro-D-glucose (¹⁸FDG). The reaction mixture was processed through a C18 Sep-Pak cartridge to remove the phase-transfer catalyst, Kryptofix 2.2.2. The final product was formulated in approximately 1 ml of de-ionized water yielding approximately 5 mCi per dose of purified product at end-ofsynthesis. Doses were measured using a CRC-12 Dose Calibrator (Capintec, Inc., Ramsey, NJ, USA).

¹⁸FDG was administered directly to the hydroponics solution of study plants. In staging these studies, individual plants were removed from their larger hydroponics growth chambers and transferred to individual 50 ml beakers filled with 30 ml de-ionized water. Forced air from an aquarium air pump maintained proper solution aeration and provided good mixing of the tracer. Studies leveraging ¹⁸F-fluoride as a water flow tracer were conducted in the same manner as the ¹⁸FDG studies. For ¹⁸F-fluoride uptake studies, the dried tracer was reconstituted in 1 mL of de-ionized.

Leaf Autoradiography: Counting down from the apex, leaf-2 (a nearly fully expanded source leaf) was subjected to topical treatments while still attached to the plant. MeJA or ISX treatments were applied on half of the leaf’s upper surface, as described above. 2.5 hour following ¹⁸F-fluoride or ¹⁸FDG tracer administration to the roots, leaf-2 was excised and imaged for quantifying either ¹⁸F-fluoride or ¹⁸FDG uptake and distribution using autoradiography (Typhoon 7000: GE Healthcare, Piscataway, NJ, USA). Differences in the radioactivity levels between the 190 treated and untreated halves were determined using Image Quant TL 7.0 software (GE Healthcare Bio-Sciences AB, Uppsala, Sweden) where 1 cm diameter circular regions-of interest (ROI) we retraced onto both sides of the leaf’s midrib (placed equidistant from that reference point), and the amount of radioactivity within each ROI was determined from the image intensity. No correction was made for tissue attenuation of radioactivity in the acquired images since it was assumed that each leaf half was identical in nature and thickness to the other.

Autoradiography was also used in ¹¹C tracer studies. In these studies, leaf-2 was detached from the plant to enable treatments to be administered via the cut petiole (see Treatment section for details). Gaseous ¹¹CO₂ tracer was administered to the detached leaf in much the same way as in the intact leaf studies and autoradiography was used to quantify ¹¹C-photoassimilate transport(after ¹¹CO₂ fixation) as a function of treatment type and treatment dose. Imaging was performed 30 min after ¹¹CO₂ fixation to avoid ¹¹C-photoassimilate transport beyond the “fieldof- view” resulting in exudation of tracer out through the petiole cut. Using the same Image Quant software, ROIs were traced around the ¹¹CO₂ fixation site, as well as the remaining leaf tissue including petiole outside that site which contained radioactivity. The amount of activity found outside of the fixation site was correlated to transported ¹¹C-photoassimilate.

Magnetic Resonance Imaging: A single intact leaf imaging experiment was conducted using BNL’s 4-Tesla MRI scanner as a way to verify the utility of the ¹⁸F-fluoride tracer as a water flow agent in plants. Past studies [37, 38 and 39] have attested to the fact that this tracer can be used as a proxy for tracing the dynamics of water transport in plants. Even so, we designed our own test. In a single experiment we compared leaf autoradiography (Figure. 1) of ¹⁸F-fluoride distribution in a ‘masked’ leaf study with that of a proton MRI of the same ‘masked’ leaf. Half of the leaf was masked using an opaque cardboard cover. The other half was illuminated using an LED illuminated fiber optic wand that allowed us to transmit light to the leaf surface while the plant was positioned within the magnet’s bore.

The BNL 4-Tesla MRI scanner was driven by a Varian INOVA console with a shielded whole-boy SONATA-Siemens gradient set (using three K2217 Siemens Cascade Gradient Power Amplifiers with 2000 V and 500 A) to produce gradient pulses with 44 mT/m peak amplitude at 0.25 m sec maximum rise time. A multiplespin- echo series of pulses was applied as a train of 180o rf pulses giving a series of echoes with a single phase encoding gradient [40]. By varying the phase encoding gradient, a spin-echo image can be obtained that can be translated into a planar T2 mapping of tissue water content.

Whole-Plant 18FDG Translocation: The movement of 18FDG from roots up into the shoots and leaves of the plant was monitored using two sodium iodide scintillation detectors (Ortec, Oak Ridge, TN, USA) that were positioned facing perpendicular to the apex area and to leaf-2. Each detector was well-shielded using small tantalum blocks to ensure minimal detector crosstalk.

Detectors were cross-calibrated against a NIST traceable point source of radioactivity prior to each experiment. Each detector was set to report radioactivity levels on 1 min time intervals. The temporal allocation patterns of radiotracer translocation to the apex region and to leaf-2 were based on the amount of tracer assimilated by the plant over the time course of the experiment, and reported as fractional total activities. The amount of tracer assimilated by the plant was determined by measuring the level of radioactivity in the beaker (after the plant was removed at the end of the experiment) and comparing that value to the original dose of administered. All calculations were performed using decay corrected values to account for ¹⁸F radioisotope decay over time.

¹⁸FDG-6-Phosphate Metabolite Analyses: Approximately 100 mg of leaf-2 tissue was extracted 2.5 hr after 18FDG was administered through the roots of the plant. Extraction was carried out in 4x w/v of methanol, briefly vortexes (VWR analog vortex mixer; Sigma-Aldrich Corp. St. Louis, MO, USA) and then sonicated (Branson Bransonic 32; Sigma-Aldrich Corp. St. Louis, MO, USA) in an iced water bath for 10 min. The tubes were centrifuged (Eppendorf Centrifuge 5424) for 2 min at 15,000 rpm and the supernatant was filtered through 0.2 μm Acrodiscs (Gelman Sciences, Ann Arbor, MI, USA). An aliquot of the extract was analyzed by radio-TLC [41] using silica gel– coated plastic sheets (Polygram SIL G/UV254; Macherey-Nagel) and acetonitrile/tetrabutylammonium hydroxide, 9.5 mmol/L, 8:2 (v/v), as eluent. The formation of ¹⁸FDG-6-P was confirmed by co-spotting the TLC plate with 245 nonradioactive ¹⁹FDG-6-P (Sigma-Aldrich, St. Louis, MO USA).

Doses of pure ¹⁸FDG-6-P were prepared using in vitro enzyme chemistry and applied to leaf tissue to test for ¹⁸FDG-6-P mobility. To generate these doses, 5 mCi of ¹⁸FDG was added to 1 ml NaH₂PO₄ buffer solution (70 mmol/L) that contained 10 mg (250– 400 units) of hexokinase type IV from bakers’ yeast (EC 2.7.1.1; Sigma-Aldrich, St. Louis, MO USA), 2 mg (3.6 μmol) of adenosine triphosphate (Sigma-Aldrich, St. Louis, MO USA) and 1 mg (5 μmol) MgCl₂·6H₂O (Sigma-Aldrich, St. Louis, MO USA) and was pH adjusted to 7.4. The solution was stirred at room temperature (30°C) for 60 min and samples taken for radio-TLC analysis of product purity (Figure 2) ¹⁸FDG-6-P transport assays were carried out by applying tracer to an abraded leaf surface and measuring the extent of transport over 1 hr using autoradiography.

One hour after ¹¹CO₂ tracer was administered to either the young apex leaves or the older source (leaves 2, 4 or 6), the respective leaf was excised from the plant at the petiole. The tissue area that was captured in the leaf cell during tracer exposure was then cut away from the remaining leaf tissue, weighed, and then flash frozen in liquid nitrogen. Using a mortar and pestle, the tissue was powdered in liquid nitrogen, and extracted using published procedures [17, 42-45]. In the first step the tissue was refluxed in 6 ml of aq MeOH (50% v/v) for 10 min at 90°C. The extract was separated by pipette, and the remaining tissue was washed twice using 6 ml of demonized water. The washings were combined with the aqueous alcohol fraction prior to counting the 11C. This fraction removed small soluble compounds including radio labeled monosaccharides and disaccharides and amino acids that were subjected to separate radio-HPLC analyses discussed below. The remaining tissue (considered cell-wall components) was subjected to an extraction using 6 ml of dilute 1 N Naoh for 10 min at 95°C. This step separated callose (1,3-β-glucans), as well as pectin’s (branched chain 1,4-β-glucans) from the remaining cell wall polymers. The extract was again separated by pipette, and the remaining tissue washed twice using 6 ml of de-ionized water. The washings were combined with the base portion prior to counting the ¹¹C. The remaining tissue was then subjected to acid digestion using a 3:1 mixture of dilute 1 N nitric acid: acetic acid for 30 min at 100°C enabling hemicelluloses to be solubilized, leaving behind cellulose as the undigested portion. The contents were cooled to ambient temperature and filtered onto 2-cm disks of pre-weighed glass microfiber filters (GF/A: 2.5 cm diameter; What man, Maid stone, UK). The collected tissue was washed 3-times during this filtration step using de-ionized water. Samples were immediately counted using a static NaI gamma radiation detector, decay-corrected back to a common zero time and fraction-corrected to allow correlation to total ¹¹C-activity fixed within the tissue. Once counted, filtered samples were dried under a heat lamp and reweighed to determine 280 the amount of cellulosic biomass.

This same cellulose analysis protocol was used to measure ¹⁸F-cellulose derived from ¹⁸FDG, and in one study application from ¹⁸FDG-6P. Unlike the ¹¹C-tracer studies, leaf tissue samples were harvested for ¹⁸F-cellulose analysis 2.5 hr after ¹⁸FDG tracer was administered to the roots. We tried to target the same region of leaf tissue and mass as in the ¹¹C-tracer studies. A longer time period was used in the ¹⁸F-tracer studies than in the ¹¹C-tracer studies to allow sufficient time for initial uptake and transport of ¹⁸F-tracer to the aerial portions of the plant. At the end of the cellulose assay, the acid indigestible portion was dried and further deconstructed using the phenol–sulfuric acid assay [46]. This method involved refluxing the material in a 1:10 solution of phenol (5%) and concentrated sulfuric acid at 100°C until all of the solid material was digested. Aliquots of this extract were neutralized and analyzed by radio HPLC for recovered ¹⁸FDG.

Measurement of Leaf Water Potential: In a separate set of measurements, leaf water potentials (Ψ) were determined for a set of different aged leaves by using a Scholander pressure bomb (model 3005, Soil moisture Equipment Corp., Santa Barbara, USA).

Statistical Analysis: Data was subjected to the Student t-test for unpaired samples assuming an unequal variance. Statistical significance levels were assigned to the following rating scale (*, P < . 05; **, P < . 01; ***, P < . 001).

Characterization of ¹⁸FDG Uptake and Translocation in Plants: ¹⁸FDG was taken up by intact plant roots and transported to the aerial portions of the plant via the xylem. This process was driven by water flow regulated by the plant’s transpiration stream. On arrival into mature leaves, ¹⁸FDG can be phosphorylated or utilized in cell-wall construction. In either case the 18F-tracer becomes immobilized. Additionally, ¹⁸FDG delivered into mesophyll cells could be converted to ¹⁸F-sucrose or some other mobile metabolite that is reloaded into the phloem and transported across longdistances to distal sink tissues such as young developing leaves and roots. Results in (Figure 1) reflect this dynamic exchange over a 4 hour time period. We observed that mature leaf-2 radioactivity, presented as a ratio of tissue radioactivity divided by the total plant radioactivity, increased during the early stages of ¹⁸FDG incubation, and then decreased during the later time points.

All data presented in Fig. 3 was corrected for radioactive decay so this trend reflected phloem re-loading and long-distance transport of ¹⁸F-tracer. In concert with this behavior, radioactivity in the plant’s apex was seen to pass through an inflection point that was slightly delayed in time from the maximum activity time point in leaf-2. This too is a reflection of phloem loading and longdistance transport of ¹⁸F-tracer from the source leaf to this sink.

Treatments using ISX and MeJA Impact ¹⁸F-Tracer Mobility: Chemical inhibitors for cell-wall cellulose, isoxaben (ISX) and methyl jasmonate (MeJA) were tested for their effects on ¹⁸F-tracer mobility derived from ¹⁸FDG. Treatments were applied to half of leaf-2, and autoradiography was used to quantify effects of treatment on phloem loading of the ¹⁸F-tracer (Figure 4). Data was presented as a ratio of the treated tissue activity divided by the untreated tissue activity (T/U) that was measured across the two halves of same leaves. Control data was derived from application of mock treatments using deionized water. Results from these studies were summarized in (Table 1). This table also contained data on the effect of treatment on leaf water flow where ¹⁸F-fluoride was used as a water tracer. We felt that it was important to access treatment effects on water flow as this dynamic had a strong bearing on xylem unloading of ¹⁸FDG into leaf-2. Furthermore, Table 1 also contained data on the effect of treatment on ¹⁸FDG phosphorylation as this process would impact ¹⁸F-tracer mobility in phloem loading, as well as remove ¹⁸F-tracer from metabolically active sugar pools used in cellulose

biosynthesis. Our application of ¹⁸FDG-6P to leaf tissue showed no evidence of physical transport, nor evidence of chemical assimilation of ¹⁸F-tracer into cellulose (data not shown). Taken together, we were able to correct ¹⁸F-tracer mobility data for changes in leaf water conductance and for changes in ¹⁸FDG phosphorylation imposed by the chemical treatment (see bottom row in Table 1). In summary, ISX treatment had no significant effect on ¹⁸FDG delivery to leaf-2 by the xylem nor did it have a significant effect on ¹⁸FDG phosphorylation in that tissue, but it did significantly reduce ¹⁸F-tracer loading into the phloem, as reflected by the significantly higher T/U value relative to controls, 1.820 ± 0.116 versus 0.949 ± 0.123, respectively (P=0.0009). In contrast, MeJA treatment significantly reduced ¹⁸FDG delivery to leaf-2 by the xylem, it slightly increased

¹⁸FDG phosphorylation and it slightly increased 18F-tracer loading into the phloem as reflected by the lower T/U value relative to controls, 0.546 ± 0.373 versus 0.949 ± 0.123, respectively (P=0.0520).

Similarly, ISX and MeJA treatments were tested for their impact on ¹¹C-photoassimilate transport in leaf-2. In these studies

the leaf was detached, and treatment was applied through the cut petiole (Figure 5) prior to administration of ¹¹CO₂. As before, autoradiography was used to quantify effects of treatment on phloem loading and transport of ¹¹C-photoassimilates within the leaf.

The distance tracer traveled from the boundary of the ¹¹CO₂ leaf cuvette over a fixed period of time was used as the metric of transport. Since treatments were applied via the cut petiole, it was also possible to quantify the dose assimilated by the study leaf during the incubation period and map transport against treatment in a dose dependent manner. Treatments were applied over 107 orders of magnitude ranging from 0.004 pmol • g fresh wt^-1 leaf tissue to 50,000 pmol • g fresh wt^-1.The results in (Figure 6)were plotted on a logarithmic scale to reflect the exponential

dose dependencies of ¹¹C-photoassimilate transport to both treatments. Transport was normalized in this figure relative to control (untreated) studies which was shown as the ‘zero’ dotted line.

Positive deviation from this ‘zero’ line reflected increased ¹¹C-photoassimilate transport, whereas negative deviation from this line reflected decreased transport. Similar to the 18FDG results, ISX treatment significantly decreased ¹¹C-photoassimilate transport in a dose dependent manner while MeJA significantly increased transport.

Leaves at different stages of development (apex, leaf-2, leaf- 4 and leaf-6) were tested to determine whether developmental leaf age influenced the extent of intracellular sugar (¹¹C) and intercellular sugar trafficking (¹⁸F) contributed to cell-wall cellulose biosynthesis. Results from these studies were presented in Table 2 along with cellulose content data that was presented as percent gram fresh weight (% gfw) of leaf tissue. Cellulose content decreased significantly from 1.024 ± 0.048 % gfw to 0.399 ± 0.124 % gfw with increasing leaf age. Leaf water potentials also increased from -0.35 ± 0.03 to -0.22 ± 0.01 MPa across the same range of leaf ages resulting from increased water content in the older leaves.

This phenomenon would inherently lower the measured values of the cellulose content with increasing leaf age when based on fresh tissue mass, but the magnitude of the observed drop in cellulose content (61%) exceeded the change in water potential (37%), so likely the measured decrease in cellulose with increasing leaf age was real. However, ¹¹C-cellulose content remained unchanged with leaf age while ¹⁸F-cellulose decreased significantly from 0.083 ± 0.011 % of tissue radioactivity to 0.010 ± 0.007 % of tissue radioactivity with increasing leaf age.

Having placed the impact of ISX and MeJA treatments on cellular sugar transport in a proper context, we next conducted tests to see if these treatments had any impact on the flux of tracers (¹¹C vs.¹⁸F) into leaf-2 cellulose. The results in Table 3 showed that ISX and MeJA treatments impacted the flux of ¹¹C-tracer into ¹¹C-cellulose, increasing it from 0.282 ± 0.019 % of leaf-2 radioactivity in controls to 0.406 ± 0.051 % with ISX treatment (P = 0.0527), and significantly decreasing it to 0.090 ± 0.026 % with MeJA treatment (P = 0.0003). These same treatments were found to have just the opposite effect on the flux of 18F-tracer (as 18FDG) into ¹⁸F-cellulose where ISX significantly decreased the flux from 0.042 ± 0.003% of leaf-2 radioactivity in controls to 0.021 ± 0.001 in treated tissues (P = 0. 0010), while MeJA significantly increased the flux to 0.055 ± 0.005 in treated tissues (P = 0.0496).

Over the years, most attention has been given to the transport of sucrose as it represents the major mobile form of photo synthetically assimilated carbon in higher plants [47, 48]. Sucrose synthesized in fully mature source leaves is exported via the phloem to supply non-photosynthetic organs with energy and carbon resources. Its long-distance transport requires loading of sucrose into phloem sieve elements from symplastic or apoplastic compartments [49, 50]. This is facilitated by sucrose transporters (SUT), also called sucrose carriers (SUC), which are driven by negative plant potentials. In Nicotiana tabacuum, the sucrose transporter 1 protein (SUT1) responsible for transporting sucrose over long-distances functions as a H⁺/sucrose symporter [51,52].

Since the identification of the first eukaryotic monosaccharide transporter in Chlorellakessleri [53], monosaccharide transporters have also been identified in a variety of higher plants [54-57]. For example, in Arabidopsis thaliana, the AtSTP1 and AtSTP3 monosaccharide transporters display both high and low affinity for D-glucose transport, respectively, suggesting functional differences [58, 59]. Marked differences in the pattern of expression have also been noted across tissues, organs and even cell types. For example, AtSTP1 transcripts are present in leaves, stems, flowers, and roots, while AtSTP2 is expressed only in developing pollen [60]. Analysis of expression patterns also suggests that these transporters are highly regulated and responsive to environmental cues, such as in response to pathogen infection or after wounding.

Additionally, from past studies, the monosaccharide transporter, MST1, was isolated from Nicotiana tabaccum and found to be most strongly expressed in sink tissues including roots, flowers and young leaves [61] where it appears to play a role in unloading of sugars. The protein is homologous to those hexose transport proteins found in Arabidopsis thaliana and Chlorella kessleri and possesses 12 putative membrane-spanning domains which can function as a H⁺/monosaccharide symporter just like SUT1, catalyzing the cellular uptake of hexoses (e.g. D-glucose and D-galactose) or pentoses (e.g. D-xylose). Even so, overwhelming evidence from ¹⁴C-tracer studies and detailed phloem sap analyses has led to the general consensus in the phloem transport field that sucrose is the predominant sugar substrate that is carried in the sieve tubes of higher plants and transported over long distances to distal tissues [62]. More specifically, hexose sugars do not transport over such long distances.

Furthermore, to the best of our knowledge, no one has seen evidence for the presence of such monosaccharide transporters in the companion cells or sieve tube elements that would facilitate such long-distance transport of hexose sugars. Of course, we cannot deny the fact that our tracer data clearly showed evidence of phloem loading and long-distance transport of ¹⁸F-tracer that was derived from ¹⁸FDG after its uptake into source leaf-2. This observation is consistent with past observations indicating phloem loading and long distance transport of ¹⁸F-tracer after application of ¹⁸FDG to sorghum leaf tips [22]. One possible explanation of these observations is that ¹⁸FDG is converted to a ¹⁸F-sucrose analog that can be actively transported [19]. Alternatively, conversion of ¹⁸FDG to its respective sugar alcohol form might provide a mechanism for moving the ¹⁸F-tracer. Polyols are reduced forms of aldoses and ketoses, and can be found in all living forms [63]. In some higher plants these sugar alcohols are, together with sucrose, direct products of photosynthesis and serve similar functions as sucrose such as translocation of carbon skeletons and energy resources between sources and sink organs [64, 65].

Chemical treatments using ISX and MeJA were found to impose interesting systematic differences on the way they affected longdistance transport of ¹⁸F-tracer and ¹¹C-photoassimilate that suggests they may act universally on H⁺-symport mechanisms. Specifically, ISX was seen to inhibit both phloems reloading of ¹⁸F-tracer, as well as long distance transport of ¹¹C-photoassimilate in a dose dependent manner. Classical sugar transport inhibitors like carbonyl cyanide 3-chlorophenylhydrazone (CCCP) behave as protonophores that can shuttle protons across the lipid bilayer of the cell’s plasma membrane, and in doing so, uncouple the membrane proton gradient from sugar transport thus shutting down active transport [66]. However, protonophoric activity is not restricted to weakly acidic compounds like CCCP. There are claims that basic compounds like (Z)-5-Methyl-2-[2-(1-naphthyl) ethenyl]-4-piperidinopyridine exhibit protonophoric activity likely due to the amine group [67]. As a weakly basic benzamide, ISX may behave in a similar capacity resulting in its ability to universally shut down sugar transport. Furthermore, we note that past studies have shown that MeJA can re-instate sugar transport after treatment with a classical sugar transport inhibitor like CCCP presumably by re-coupling proton symport. Our observation that MeJA increased long-distance ¹¹C-photoassimilate transport, as well as increased phloem loading and long distance transport of ¹⁸F-tracer suggests that its ability to act on the membrane proton gradient serves to increase transport activity of sugars in general.

Our present work also showed that intercellular sugar trafficking was an integral part of the regulatory mechanism for cellulose biosynthesis. Here we used developmental leaf aging as a model for exploring changes in cellulose biosynthesis and for measuring changes in the flux of intra and intercellular sugar resources in the process. Our data clearly demonstrated that ¹⁸FDG can be taken up by roots and delivered to all leaf types via xylem transport, and unloaded at the parenchyma symplastxylem interface where the sugar could migrate from cell-tocell through the plasmodesmatal cytosolic sleeve in between the plasma membrane and the endoplasmic reticulum. As cells expand and differentiate, their fate determines the extent to which their cytoplasmic connectivity to other cells is maintained [68]. Some cell types, such as those in the leaf mesophyll remain closely connected with their neighboring cells and this connectivity could extend to the parenchyma symplast-xylem interface. However, cytoplasmic continuity is not a constant and will decline with tissue development as simple plasmodesmata (PD) are lost during leaf sink-source transition [69], and the frequency of even the more complex branched PDs decline with maturation [70]. Taken together, we would expect ¹⁸FDG’s ability to move cell-to-cell via the symplast to systematically decline with aging, consistent with our finding that ¹⁸F-tracer flux into ¹⁸F-cellulose decreased with increasing leaf age. Furthermore, we were surprised to find that there was no change in the flux of ¹¹C into ¹¹C-cellulose from intracellular sugar sources. What this implies is the cellulose synthetic machinery remained active regardless of leaf age, and that the diminished supply of intercellular sugar was what controlled the process. We tested this hypothesis using ISX and MeJA treatments where from our earlier discussion it was concluded that ISX inhibited sugar transport (both in long-distance movement through the phloem, and in cell-to-cell movement within same tissues) while MeJA enhanced it. Consistent with this model, the inhibition of sugar transport by ISX increased the flux of ¹¹C-tracer into ¹¹C-cellulose from intracellular sugar while it decreased the flux of ¹⁸F-tracer into ¹⁸F-cellulose from intercellular sugar. Also consistent with this model, the increase in sugar transport by MeJA decreased the flux of ¹¹C-tracer into ¹¹C-cellulose from intracellular sugar while it increased the flux of ¹⁸F-tracer into 18F-cellulose from intercellular sugar.

To date, there exists an extensive body of information providing insight into the structural architecture of plant cell walls which are comprised of cellulosic micro fibrils embedded in a matrix of no cellulosic polysaccharides that include heteroxylans and 1,3; 1,4)-β-glucans [71]. Even so, progress in understanding the mechanisms for regulating the synthesis of these macromolecular structures has been relatively slow. For example, past work has shown that β-glucan synthesis, responsible for 1, 3-β-glucan (callose) synthesis in cell wall construction, decreases in activity with plant tissue age [72]. In contrast to this, our work suggests that cellulose synthesis remains uniformly active; at least in developing leaf tissues, and most importantly that intercellular sugar trafficking is a critical component to regulating cell-wall cellulose biosynthesis. We expect that the use of sugar transport mutants could help shed additional light on this highly regulated process and this will certainly be the focus of future studies.

Finally, an area of keen interest is to understand the responses of the plant cell wall to a biotic stresses including drought, flooding, and temperature, or to biotic stresses including attack by pathogens and feeding insects. The effects of these stresses on cell wall metabolism are complex, but important to unravel if we are to build adaptation and/or resistance traits in future cropping systems.

This article has been authored by Brookhaven Science Associates, LLC under contract number DE-SC0012704 with the U.S. Department of Energy, Office of Biological and Environmental Research. Additional support was provided by the German Academic Exchange Service (Deutscher Akademischer Austauschdienst, DAAD) Bonn. The United States Government retains and the publisher, by accepting the article for publication, acknowledges that the United States Government retains a nonexclusive, paid-up, irrevocable, world-wide license to publish or reproduce the published form of this manuscript, or allow others to do so, for United States Government purposes. The authors would like to acknowledge B. Rooney for his assistance in acquiring the MR image, D. Alex off and C. Shea for their assistance in preparing 18FDG, and D. Braun for providing feedback during manuscript preparation.

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