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. 2026 Mar 27;74(13):11001–11014. doi: 10.1021/acs.jafc.5c13415

A Plant-Derived Arabinoxylan Platform for Biomolecule Delivery into Plant Cells

Mohammad Rashid ∥, Kari Vinzant ∥, Ashique Al Hoque ‡, Mohiuddin Quadir ‡,§, Mariya Khodakovskaya ∥,*
PMCID: PMC13067356  PMID: 41891846

Abstract

Nanotechnology offers innovative solutions for enhancing plant productivity through biocompatible, nontoxic nanocarriers capable of delivering diverse agrochemicals and biomolecules directly to plants. In this study, we demonstrated the potential of a nanocarrier system based on arabinoxylan (AX), a wheat bran-derived polymer. Two AX-based nanocarriers, fluorescein isothiocyanate-labeled AX (AX–FITC) and positively charged AX (AX+), were synthesized and used to determine possible uptake and delivery of 35S-eGFP-Nos plasmid DNA (pDNA). Plant uptake of AX nanocarriers was assessed by various methods, including passive uptake through incubation, leaf injection, and vacuum infiltration. Among the tested methods, injection of conjugates into tobacco leaves and vacuum infiltration of seedlings proved to be the most efficient for delivering pDNA polyplexes. Furthermore, a comprehensive analysis of the phenotype and transcriptome of model Solanaceae species (tomato) exposed to varying nanoparticle concentrations revealed no signs of phytotoxicity or genotoxicity, reinforcing its biosafety for seed and plant treatments. This work highlights the potential of AX as a sustainable, plant-derived nanocarrier, offering a “green” alternative to synthetic nanomaterials for the delivery of DNA and agrochemicals in plant biotechnology and agriculture.

Keywords: arabinoxylan, nontoxic nanocarriers, plant-based polymers, green chemistry, DNA delivery, nanoparticles

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Introduction

Nanotechnology has the potential to enhance agricultural practices by leveraging the unique properties of nanomaterials, such as their small size and effective penetration capabilities. These properties make nanoparticles (NPs) a promising tool for addressing agricultural and environmental challenges such as, helping to reduce chemical runoff and fertilizer loss through controlled release of essential micronutrients, targeted disease and pest control, and precise nutrient delivery to plants. − For more than a decade, a wide range of metal-based (MBNs) and carbon-based (CBNs) nanomaterials have been identified as efficient nanocarriers for delivering agrochemicals and nucleic acids to plants. , Among them, carbon nanotubes (CNTs), a rigid and rod-like nanomaterial, have shown potential utility in genetic engineering methods due to their exceptional ability to deliver foreign DNA into plant cells. − A system to efficiently deliver genetic material to plant cells is particularly sought after for use in a variety of gene editing applications: from crop improvement via CRISPR/Cas-mediated trait modification, to functional genomics studies and the development of transgenic plants with enhanced resistance to pests, diseases, or environmental stressors. −

Though a breakthrough material, CNTs are now the subject of hot debate regarding their safety, use, risk of release into the environment. − There is currently an urgent push to find safer nanomaterial alternatives to CNTs, with preference given to materials that are biodegradable, low in toxicity, and cost-effective. , One promising direction involves leveraging organic waste valorization to develop circular and sustainable materials, offering a viable alternative to CNTs in terms of environmental compatibility and resource efficiency. Numerous biodegradable and naturally derived nanomaterials, often made from stock materials with Generally Recognized as Safe (GRAS) status by the United States Food and Drug Administration, present promising alternatives. Of these, plant-derived polymers have shown strong potential as NPs for use in agricultural applications including seed coatings, soil treatments, and the targeted delivery of antifungal agents, hormones, and insecticides. ,, However, it is uncertain whether these natural polymers exhibit properties comparable to those of their highly effective CBNs and MBNs counterparts, particularly regarding their size and uniformity. The efficacy of CNT uptake has been speculated to be due to their small scale and high aspect ratio, allowing them to pass the plant cell wall’s extremely narrow size exclusion limit of ∼20 nm. NPs and their conjugates can be administered to plants through different exposure routes such as leaves, stems, roots, and seeds using techniques such as foliar application, hydroponics, soil incorporation, or agar medium. In foliar applications, points of entry into plant tissues include cuticular pores and stomatal openings, which each have size exclusion limits of 2 nm and 10–50 nm, respectively. Later, cellular internalization and subsequent translocation of NPs in plants have been suggested to be through endocytosis, pore formation, mediated by carrier proteins, or through plasmodesmata, all which have their own size limitations. These size exclusion thresholds exclude many polymeric NPs, which are often greater than 100 nm in diameter. ,

Despite this, there has been a recent shift in attention to soft, flexible nanomaterials for use with plants. Multiple reports have demonstrated internalization of >100 nm polymeric NPs in a variety of plant systems, particularly that within the roots of plants. ,, For example, polystyrene nanoparticles as large as 200 nm have been shown to successfully enter and accumulate within the roots of wheat plants, and up to 2 μm in the roots of lettuce plants. Nanoliposomes approximately 100 nm in diameter have demonstrated efficient delivery of essential therapeutic components in crops via foliar application. Charged polystyrene NPs with a mean diameter of 200 nm can penetrate and accumulate into the root epidermis of Arabidopsis thaliana. Recently, we demonstrated that both zein and cellulose nanocrystals (CNCs) that were considerably larger than 100 nm in at least one dimension (159 ± 10 nm in diameter for zein, and 212.9 ± 43.1 in length and 3.2 ± 0.8 nm in height for CNCs) are capable of entering plant cells and can deliver plasmid DNA (pDNA), resulting in successful expression of GFP for up to 7 days. This ability of polymer-based NPs to bypass the size exclusion limit is often attributed to their mechanical flexibility, allowing the NPs to deform and gain access to these small entry points. The wheat-derived polysaccharide arabinoxylan (AX) used in this study features a branched polymeric structure composed of two pentose sugars: arabinose and xylose. Its inherently dendritic architecture, combined with self-assembling properties and an abundance of hydroxyl groups along the backbone, makes AX a promising candidate for biomolecule delivery applications. As AX can be derived from a variety of feedstock sources ranging from wheat flour to spent brewer’s grains, there is an opening for potential value-addition using cereal waste products to develop an AX-based plant nanocarrier platform. −

Here, for the first time, we present experimental evidence that AX-based nanocarriers can enter plant cells and mediate the direct delivery of chemical compounds or nucleic acids in treated cells or whole plants. Previously, Sarker et al. proposed a laboratory-scale method for extracting and characterizing AX from wheat bran and demonstrated its chemical modification for DNA complexation, but they did not employ its use in vitro or in vivo. Using the exact same material generated within this previously reported work, AX nanoparticles were functionalized with glycidyl trimethylammonium chloride (GTMAC) to produce a positively charged formulation (AX+), enabling electrostatic interaction with negatively charged nucleic acids. A separate formulation of AX+ was conjugated with fluorescein isothiocyanate (AX–FITC) and used in preliminary uptake studies to assess the internalization of dye in plant cells. The untagged AX+ was employed later for complexation with pDNA containing the fluorescent GFP gene (Figure ). Our experiments confirmed the ability of AX-based nanocarriers to deliver dyes and DNA to plant cells by the detection of fluorescence and expression of GFP transcripts. The proposed AX-mediated delivery platform offers a cost-effective and sustainable “green” alternative, making it particularly well-suited for transient gene-editing applications and highly attractive for industrial use. Given that AX constitutes approximately 86.4% of the total hemicellulose content of wheat bran, its abundance and biocompatibility position it as an affordable, sustainable, and safe substitute for CBNs and MBNs for the direct delivery of cargo to plant cells.

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Synthesis of AX conjugates and experimental design. (A) AX was modified with GTMAC under alkaline conditions to yield cationic AX+. AX+ was used for assessing potential phytotoxicity and impact on the transcriptome of tomato seedlings. (B) AX was grafted to FITC to generate AX–FITC via dibutyltin dilaurate-catalyzed reaction in DMSO. Tobacco callus cells and leaves were incubated in a 1 mg/mL dispersion of AX–FITC overnight to determine potential uptake via confocal microscopy. (C) AX+ was mixed with pDNA encoding GFP to form pDNA–AX polyplexes. pDNA–AX was introduced into plant cells via overnight incubation, vacuum infiltration, or leaf injection. Subsequent expression of GFP gene products was determined confocal microscopy and RT-qPCR.

Materials and Methods

Biological Material

pGFPGUSPlus was a gift from Claudia Vickers and was provided by Addgene (Addgene plasmid # 64401; https://www.addgene.org/64401/). Plasmid DNA was isolated from the Escherichia coli culture using a QIAGEN Plasmid Mini Kit (Qiagen Inc. Valencia, CA). Tomato seeds (cv. ‘Micro-Tom’) were purchased from Reimer Seeds Inc. (Saint Leonard, MD). Tobacco seeds (cv. ‘Havana’) were purchased from Victory Seed Company. The tobacco callus culture that was used in experiments was established previously in the Khodakovskaya laboratory.

Synthesis and Characterization of GTMAC-Modified AX (AX+) and AX–FITC Nanoconjugates

Commercially available AX derived from wheat flour was used to generate both AX–FITC and cationic AX species. In a previously reported synthesis, the chemical conversion of AX to AX+ involved a reaction of AX with GTMAC in the presence of sodium hydroxide (NaOH) aqueous solution. The reaction was continued for 5 h at 50 °C. The samples were characterized by 1H NMR spectroscopy and Fourier transform infrared spectroscopy (FTIR), following our earlier work. For labeling AX with a fluorescent dye, FITC, the dye was reacted with AX in the presence of dibutyltin dilaurate in dimethyl sulfoxide (DMSO) at 50 °C for 5 h (Figure A) as previously reported by Cho et al.

Preparation and Characterization of pDNA–AX Polyplexes

Using a fabrication method similar to a previously reported method, AX+ NPs were fabricated by dissolving them from dry powder at a ratio of 1 mg/mL in sterile double-distilled water by constant stirring set to 700 rpm at room temperature for 1.5 h. pDNA–AX conjugates were prepared by incubating pDNA containing GFP gene with the fabricated cationized arabinoxylan for 30 min. To find the binding stoichiometry, AX+ and pDNA were mixed at different ratios: 1 μg of pDNA + 3 μg of AX+ (1:3), and 1 μg of pDNA + 5 μg of AX+ (1:5). These conjugates were then loaded into a 1% agarose gel added with ethidium bromide (Invitrogen by Thermo Fisher Scientific) to understand the binding ability of AX+ NP with the pDNA. The samples were mixed with the DNA Gel Loading Dye Blue (Thermo Scientific) and loaded into the wells alongside a 1 kb Plus DNA Ladder (Invitrogen by Thermo Fisher Scientific). Electrophoresis was performed at 100 V for 30 min, and bands were visualized using a UV transilluminator (Bio-Rad, CA, USA).

Dynamic Light Scattering (DLS) Analysis

Polyplexes, prepared at 1:3 and 1:5 ratios of DNA and AX+ polymer, respectively, were subjected to DLS analysis for the determination of particle sizes and zeta-potentials, which were measured using a Zetasizer NanoZS (Malvern Instruments, Herrenberg, Germany), equipped with a helium–neon laser with an excitation wavelength of 633 nm. The equipment was set to auto mode, and the viscosity and refractive index of water were measured.

TEM Analysis

Each polyplex sample (1:3 and 1:5 DNA:AX) was put for two min on a 300-mesh Formvar-carbon coated copper grid (Ted Pella Inc., Redding, California, USA), then wicked off using filter paper. Phosphotungstic acid 0.1%, pH adjusted to 7–8, was put onto the grid and left to stand for two min before being wiped off, and the grid allowed to air-dry. Images were captured at the NDSU Electron Microscopy Core using a JEOL JEM-1400Flash transmission electron microscope (JEOL USA, Peabody, Massachusetts, USA).

Treatment of Tobacco Samples Using Incubation, Injection, and Vacuum Infiltration Methods

Incubation of Tobacco Samples with Conjugates

Tobacco leaf tissues and tobacco callus culture (callus) were incubated with a solution of pDNA–AX polyplex and left overnight at room temperature with continuous shaking at 120 rpm. Samples were then washed 8 times using sterile double-distilled water and analyzed using confocal microscopy.

Injection of Young Tobacco Leaves

Tobacco leaves were injected with a solution of AX+ and pDNA using a syringe by making a puncture in the leaves and pushing the entire conjugate solution through the puncture. Seedlings were allowed to grow further for 3, 5, and 7 days postinjection.

Vacuum Infiltration of Tobacco Seedlings

Whole tobacco seedlings were immersed in the conjugate solution, and a vacuum of – 0.8 bar was applied. After the vacuum was released, the leaves imbibed the surrounding solution, likely through the stomata. Seedlings were allowed to grow further for 3, 5, and 7 days after vacuum infiltration.

Confocal Microscopy

Analysis of samples was done with an LSM 880 laser microscope at the University of Arkansas for Medical Sciences, UAMS). The visualization of AX–FITC and GFP protein was performed at 488 nm. All the confocal images produced were analyzed for intensity of fluorescence using Zen Lite 3.0 software, and the datasets were compared by single-factor ANOVA, α = 0.05 in either Microsoft Excel (AX–FITC conjugates) or R software (pDNA–AX conjugates).

RT-qPCR Detection of GFP Gene Expression in Exposed Tobacco Samples

RNA was isolated from unexposed tobacco samples (seedlings, leaves, and tobacco callus culture), using a Plant RNeasy Qiagen Kit (Qiagen Inc. Valencia, CA), and the samples exposed to AX+ (control), pDNA, and pDNA–AX. The cDNA was generated from 1 μg of total RNA using a SuperScript III FirstStrand Synthesis System (Invitrogen, USA) with a dT20 oligonucleotide as a primer, according to the manufacturer’s protocol. cDNA samples were diluted to 200 ng/μL and used for real-time quantitative PCR analysis with PowerUp SYBR Green Master Mix (Applied Biosystems, Lithuania) in CFX96 Real-Time System (Bio-Rad, Singapore). To confirm expression of the GFP gene in plant material treated with pDNA–AX, the GFP gene was amplified using primers 5′-GGA­TCC­ATG­GTG­AGC­AAG­GGC­GAG­GAG­CTG-3′ (f) and 5′-TTA­CTT­GTA­CAG­CTC­GTC­CAT­GCC­GAG­AGT-3′ (r). The reaction mixture contained 10 μL of SYBR green, 0.1 μM primer, 200 ng of cDNA, and DNase-free water to a final volume of 20 μL.

Germination of Tomato Seeds Exposed to Different Doses of AX+

For germination experiments, the seeds were surface sterilized by immersing the seeds in 70% ethanol for 1 min, followed by soaking in 50% bleach solution on continuous shaking for 10 min, and then rinsed with sterile water 8–10 times to remove any bleach. Subsequently, the tomato seeds were grown on the MS media supplemented with a range of AX+ concentrations: 100, 250, and 500 mg/L. In a separate experiment, tomato seeds were coated with AX+ by airbrush with concentrations of 100 and 500 mg/L and allowed to grow for 3 weeks under controlled growth chamber conditions (8 h dark (22 °C)/16 h light (22 °C) cycle). Germination data were consistently collected every day, and the photos were collected at intervals of 7 days. Fresh/dry weights and root/shoot lengths were recorded for 21-day-old, germinated tomato seedlings. Untreated tomato seeds and seedlings germinated from the untreated seeds were used as appropriate controls. Datasets were compared by single-factor ANOVA, α = 0.05 in Microsoft Excel, and the results were presented as means and standard errors.

RNA-Seq of Tomato Seedlings Exposed to AX+

Total RNA was extracted from 21-day-old control (untreated) seedlings and seedlings exposed to 100, 250, and 500 mg/L AX+ by using the RNeasy Plant Mini Kit (Qiagen Inc. Valencia, CA). The quality of RNA was measured using an ND-2000 instrument (NanoDrop Technologies). RNA samples were sent to Novogene Co., Ltd. (Sacramento, CA) for RNA-Seq RNA-Seq data analysis was provided by Novogene, which included sample quality control, library construction, library quality control, sequencing, data quality control, and bioinformatics analysis. RNA sequencing was performed via an Illumina platform (Illumina, San Diego, CA).

Validation of RNA-Seq Analysis Using RT-qPCR

RNA was isolated from tomato plants exposed to AX+, as described above. The cDNA was generated from 1 μg of total RNA using a SuperScript III FirstStrand Synthesis System (Invitrogen, USA) with a dT20 oligonucleotide as a primer, according to the manufacturer protocol. cDNA samples were diluted and used for RT-qPCR analysis with the SsoAdvanced Universal SYBR Green PCR master mix (Bio-Rad, USA) in a CFX Opus 96 Real-Time PCR detection system (Bio-Rad, Singapore). The tomato genes Sugar Transporter ERD6-like 16 (PTHR48021:SF37) and Aquaporin TIP3-2-Related (PTHR45665:SF23) were amplified (Figure S6). To amplify Sugar Transporter ERD6-LIKE 16, the primers 5′- TGT­ACA­CAC­TAT­TGA­ACTC­CCTG-3′ (f) and 5′-ATC­TCG­TGG­TCA­ATA­GAG­CGA-3′ (r) were used. To amplify Aquaporin TIP3-2-Related, the primers 5′-TGC­AGA­CCT­CAT­AGAG­GGGA-3′ (f) and 5′-CGC­CCT­GGA­TAG­ACAA­CCAC-3′ (r) were used. House-keeping gene (18S) was amplified for all samples using the primers 5′-GAA­AGT­TGG­GGG­CTCG­AAGA-3′ (f) and 5′-CCT­AAA­AGC­AAC­ATCC­GCCG-3′ (r) (Table S5).

Four independent biological samples were used in the analysis. For each biological sample, three technical replicates were run. The real-time PCR data was analyzed using the “ΔΔCt” method to obtain the relative mRNA expression of each sample. −

Results and Discussion

Covalent Attachment of FITC (AX–FITC), and GTMAC (AX+) to AX and Their Use in Biological Experiments

To develop a plant-compatible delivery platform, AX conjugates for both tracking and nucleic acid delivery applications were synthesized. For the initial investigation into cellular uptake of AX-based nanocarriers, AX was converted in AX+ using GTMAC as a source of quaternary nitrogen (Figure A) as previously described to enable electrostatic complexation with pDNA. AX+ was labeled with FITC by reacting the dye with AX+ in dimethyl sulfoxide in the presence of dibutyltin dilaurate at 50 °C for 5 h (Figure B) as previously reported by Cho et al. The product was purified by dialysis and analyzed with UV–vis spectroscopy (Figure S1). The AX–FITC conjugate exhibited a slightly red-shifted and broadened absorbance peak relative to FITC, suggesting successful conjugation, whereas AX+ did not exhibit any substantial absorbance (Figure S1). Tobacco cells and leaves were later incubated in AX–FITC and analyzed with confocal microscopy to initially probe the potential for instantaneous uptake. To determine whether AX+ had negative effects on germination, AX+ was reserved for later use both as a sprayed seed treatment and as an additive to growth media supporting tomato seedlings. AX+ was mixed with pDNA to create pDNA–AX polyplexes, which were used to evaluate the ability of AX-based NPs to deliver genetic material into plants through various methods including passive incubation, vacuum infiltration, and direct leaf injection (Figure C). Thorough characterization of feedstock AX and AX+ including elemental, FTIR, and 1H NMR analyses are available in the publication wherein the materials were originally generated by Sarker et al.

Passive Delivery of AX–FITC to Tobacco Cells and Leaf Tissue via Incubation

To evaluate AX uptake in plant cells, AX–FITC was applied to leaves of 1-month-old tobacco seedlings and tobacco callus cells in the form of a 1 mg/mL dispersion. Both whole leaves and callus cells were incubated with AX–FITC conjugate for 24 h. All samples were later imaged using fluorescent confocal microscopy and digitally assessed for a fluorescence signal after thorough washing to remove any noninternalized, residual conjugate material (Figure A,B). The fluorescence signal observed within cells was then digitally measured and quantified. Both the tobacco callus culture and leaves treated with AX–FITC conjugates had a detectable fluorescence signal within cells, suggesting successful internalization (Figure C,D). Minimal signals were detected in AX+ and water-treated samples for both callus cells and leaves. Incubation with AX–FITC resulted in substantially higher fluorescence than FITC alone, with AX–FITC being 44.01% higher than FITC in leaves and 99.95% higher in callus cells.

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Assessment of AX–FITC conjugate internalization in tobacco callus cells and leaves after simple overnight incubation. (A, B) Confocal microscopy visualization of fluorescence signal in callus cells (A) and leaves (B) treated with water, FITC, AX+, and AX–FITC. (C, D) Mean fluorescence intensity of callus cells (C) and leaves (D) treated with AX+, FITC, and AX–FITC. n = 3. Scale bars = 20 μm. Statistical significance was determined by one-way ANOVA, α = 0.05 (** = p ≤ 0.01, *** = p ≤ 0.001).

Based on the intense fluorescence signal from within both callus and leaf cells, it seems that AX nanocarriers may penetrate cell walls and deliver material into plant cells. In treated leaves, the highest points of fluorescence intensity were observed in or around the stomata (Figure B). This observation may indicate that passive foliar uptake of AX NPs primarily occurs through stomatal flooding instead of cuticle penetration. To ensure that the observed fluorescence was due to true internalization rather than residual unbound FITC, we next employed a more stringent test of delivery using pDNA encoding GFP. Beyond having real applications in genetic engineering, this approach enables internalization to be confirmed not only through confocal microscopy but also through the presence of GFP transcripts.

Determination of Loading Capacity and Characterization of pDNA–AX Polyplexes

Before the biological experiments with pDNA and AX+ could begin, the binding ratio of AX+ and pDNA first had to be determined, and the resulting polyplex had to be characterized. AX+ NPs were electrostatically complexed with negatively charged pDNA by mixing at pDNA:AX+ ratios of 1:3 and 1:5 based on the loading capacity determined by an electrophoretic mobility shift assay (Figure A). Based on the lack of a band and fluorescence signal contained within the gel wells, it is apparent that both 1:3 and 1:5 pDNA:AX+ ratios are effective for tight electrostatic association between pDNA and AX+.

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Confirmation and characterization of pDNA–AX polyplexes. (A) Gel imaging of the AX+ bound with pDNA (35s-eGFP-Nos) at different ratios. 1:3 pDNA:AX+ and 1:5 pDNA:AX+ were loaded into a 1% agarose gel and assessed for mobility against that of 1 μg of free pDNA. (B–D) Characterization of pDNA–AX conjugates: (B, C) unimodal distributions of (B) hydrodynamic diameter and (C) ζ-potential; (D) TEM image of 1:3 pDNA–AX polyplex (scale bar = 200 nm).

Dynamic light scattering (DLS) analysis was conducted to determine both the size and the surface charge of the 1:3 and 1:5 polyplexes. The average hydrodynamic diameter of the pDNA–AX polyplexes was 168.2 nm for the 1:3 ratio, and 218.0 nm for the 1:5 ratio (Figure B). These results suggest that increasing the AX+ content in the polyplex does indeed increase the overall size of the NPs slightly. The zeta potentials of each ratio were determined to be −11.5 and 3.17 mV for the 1:3 ratio and 1:5 ratio, respectively; the higher positive surface charge corresponds to the higher content of the AX+ polymer (Figure C). Transmission electron microscopy (TEM) imaging of the 1:3 ratio polyplex revealed stable spherical NPs (Figure D). The 1:3 ratio was used in the following biological experiments due to the smaller average diameter compared to the 1:5 ratio.

Passive Delivery of pDNA–AX to Tobacco Callus Cells and Leaves Using the Incubation Method

To confirm whether AX-based nanocarriers can penetrate plant cell walls and deliver cargo within plant cells, AX+ NP were selected to enable electrostatic association between the NP and negatively charged nucleic acids, as described in Figure C. The ability of AX+ to traverse plant cell walls was assessed by incubating pDNA–AX with both tobacco callus cells and tobacco leaves. In this experiment, we aimed to understand whether the simple incubation of callus cells or leaves with pDNA–AX would be sufficient for cellular internalization (Figure A,B). In our experiments, pDNA–AX treated tobacco callus cells showed stronger fluorescence than all other callus treatments: 52.09% stronger than free pDNA and 70.05% and 67.92% more potent than water and AX+ treated samples, respectively (Figure C). In leaves, pDNA–AX treatments showed stronger fluorescence compared to other treatments: 53.20% stronger than free pDNA, and 94.31% and 116.06% more potent than water and AX-treated samples, respectively (Figure D).

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Visualization and detection of GFP transcripts in callus tobacco cells and leaves treated using the incubation method. (A, B) Confocal microscopy visualization of fluorescence signal in tobacco callus cells (A) and tobacco young leaves (B) incubated with water, AX+, free pDNA, and pDNA–AX. Scale bars = 20 μm. (C, D) Mean fluorescence intensity of treated callus cells (A) and leaves (B). n = 3. (E, F) RT-qPCR analysis of GFP mRNA abundance in treated callus cells (E) and leaves (F) 24 h postincubation. Statistical significance was determined by one-way ANOVA, α = 0.05 (* = p ≤ 0.05, ** = p ≤ 0.01, *** = p ≤ 0.001).

Real-time PCR (RT-qPCR) analysis of pDNA–AX treated callus cells and leaves corroborated the confocal microscopy results by confirming GFP gene expression in both tissue types, as evidenced by the presence of GFP transcripts in the treated tissues (Figure E,F). In contrast, negligible GFP expression was detected in samples treated with AX alone, pDNA alone, or water, indicating that pDNA–AX enabled the successful delivery of pDNA into plant cells. However, both the fluorescence intensities observed via confocal imaging and the GFP transcript levels were low, suggesting limited uptake of pDNA–AX by using the incubation method. To enhance delivery efficiency, we explored additional treatment approaches involving external physical forces, such as leaf injection and vacuum infiltration. Both methods, previously described as efficient strategies for improving plant transformation, are often used alongside Agrobacterium-mediated transformation to enhance bacterial uptake into leaves and increase the transformation success rate.

Enhancing Uptake of pDNA–AX to Tobacco Seedlings and Leaves Using Vacuum Infiltration and Leaf Injection

Leaf infiltration, including both vacuum and syringe-based approaches, is a foundational technique in plant biology. ,, These methods enable the introduction of compounds such as dyes, nucleic acids, or nanoparticles into the intercellular spaces of leaf tissue. Infiltration relies on a pressure differential, either by vacuum or manual force, to drive materials into the apoplast. This technique has been widely used to study gene expression, plant-microbe interactions, and delivery efficiency in plant biotechnology applications.

Here we use injection and vacuum infiltration of tobacco tissues to enhance pDNA–AX uptake by plant cells without relying on biological carriers such as Agrobacterium or cost-prohibitive methods like biolistics. Using a pipet, one-month-old tobacco seedlings were abaxially injected with water, AX+, free pDNA, or pDNA–AX and analyzed using confocal microscopy 5 days post-treatment and RT-qPCR at 3, 5, and 7 days post-treatment (Figure ). Leaves injected with water, AX+ alone, or free pDNA showed no detectable fluorescence, whereas those treated with pDNA–AX displayed a strong GFP signal (Figure A,B). The green fluorescence in pDNA–AX treated leaves was 173.57% higher than in leaves injected with free pDNA. To further validate the internalization and expression of pDNA–AX, RT-qPCR analysis was performed at 3, 5, and 7 days following injection (Figure C–E). GFP transcript levels in pDNA–AX treated leaves peaked at 5 days post-treatment (Figure D). By day 7, the expression declined to levels comparable to the background signal observed with free pDNA (Figure E).

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Visualization and confirmation of GFP expression in tobacco leaves injected with pDNA–AX. Comparative confocal images of tobacco leaves injected with H2O, AX+, pDNA, pDNA–AX 5 days post-treatment. (A) Confocal microscopy imaging of 1-month-old tobacco leaves injected with either water, AX+, free pDNA, or pDNA–AX. Scale bars = 20 μm. (B) Measured mean fluorescence intensity of injected leaves. n = 3, each biological replicate generated six data points. (C–E) RT-qPCR analysis of GFP mRNA abundance in injected leaves (C) 3, (D) 5, and (E) 7 days postinjection. Statistical significance was determined by one-way ANOVA and Tukey’s HSD test, α = 0.05 (ns = not significant, *** = p ≤ 0.001, **** = p < 1 × 10–16).

To determine if vacuum infiltration enabled effective uptake of pDNA–AX into cells of seedlings, 14-day-old tobacco seedlings were infiltrated with pDNA–AX in a vacuum chamber. Five days postinfiltration, seedlings were taken for confocal microscopy to detect the presence of GFP fluorescence signal, and at 3, 5, and 7 days post-treatment, samples were analyzed for the presence of GFP transcripts using RT-qPCR (Figure ). Similar to the results of the leaf injection experiments, only pDNA–AX treated leaves successfully exhibited a substantial green signal beyond that of chlorophyll autofluorescence (Figure A). pDNA–AX leaves fluoresced 106.47% stronger than leaves injected with free pDNA (Figure B) and exhibited relatively steady levels of GFP transcript expression over 3, 5, and 7 days postinfiltration (Figure C–E). Both experiments with leaf injection and vacuum infiltration confirmed that pDNA–AX successfully induced GFP expression in tobacco cells and that both injection and infiltration methods of treatments are fully sufficient for enhancing the uptake of pDNA complexed with AX+.

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Visualization and confirmation of GFP expression in tobacco seedlings vacuum infiltrated with pDNA–AX. Comparative confocal images of tobacco seedlings vacuum infiltrated with H2O, AX+, pDNA, pDNA–AX 5 days post-treatment. (A), Confocal microscopy imaging of 14-day-old tobacco seedlings vacuum infiltrated with either water, AX+, free pDNA, or pDNA–AX. Scale bar = 20 μm. (B), Measured mean fluorescence intensity of vacuum infiltrated seedlings. n = 3, each biological replicate generated six data points. (C-E), RT-qPCR analysis of GFP mRNA abundance in seedlings (C) 3, (D) 5, and (E) 7 days after vacuum infiltration. Statistical significance was determined by one-way ANOVA and Tukey’s post hoc test, α = 0.05 (* = p ≤ 0.05, ** = p ≤ 0.01, *** = p ≤ 0.001, **** = p ≤ 0.0001).

A combination of confocal microscopy and gene expression analysis demonstrated that AX NPs over 200 nm in diameter can successfully deliver genetic material to plant cells and result in the successful expression of transgenes. This evidence challenges the prevailing assumption that NP size strictly limits cellular uptake in plants. Our results align with recent studies reporting the uptake of polymeric NPs larger than 100 nm, potentially facilitated by stomatal entry as well as structural features within the cell wall. It is important to highlight that while the AX+ used in this study has been cationized, the 1:3 pDNA–AX polyplex possesses a slightly negative surface charge (Figure B,C). In animal cells, the membrane-disruptive properties of cationic polymers and nanoparticles are well-documented, often facilitating intracellular delivery. Similarly, in bacteria, the disruption of membranes by cationic molecules forms the basis for the development of many cationic bactericidal agents. However, in plants, there have been some conflicting reports regarding NP surface charge and their uptake and translocation in different tissues. While there are many reports of foliar uptake of highly cationic NP, there is some evidence to suggest that once internalized, they do not translocate effectively due to their affinity for negatively charged cell surfaces. , Interestingly, there are now multiple reports of anionic NP having higher mobility in plants compared to positively charged NP. − Zhang et al. investigated the foliar uptake of polymeric NP with different surface charges and aspect ratios. In their work, they determined that while the positively charged NP were found to be closer associated with the epidermis of leaves, the negatively charged NP were more readily translocated within the plant. Plant transformation is an inherently low-efficiency process that could drastically benefit from a highly mobile, low toxicity nanocarrier. AX+ has already been demonstrated to be an appropriate material to conjugate with CRISPR/Cas9 DNA, and may be a greatly beneficial tool for the plant genetic engineering sector.

In this study, we demonstrate that the method of conjugate delivery is critical for maximizing the pDNA uptake efficiency in plant cells. Simple incubation of plant material with pDNA–AX results in only minimal transgene expression (Figure ), which may be insufficient for many biotechnological applications. In contrast, applying external force, such as with vacuum infiltration or direct injection, significantly enhances transformation efficiency (Figures and ). Both techniques have been previously shown to facilitate the uptake of loaded cargo and DNA with CNTs. , Both vacuum infiltration and injection may induce microdamage to plant cell walls, potentially creating additional entry pathways for NP uptake. Notably, GFP expression remains stable up to 7 days after vacuum infiltration (Figure E). However, when pDNA–AX is delivered via leaf injection, GFP expression declines by day 7, suggesting that transgene expression mediated by AX+ as a nanocarrier is at least transient. Transient expression has been widely recognized as a valuable tool in biotechnology and is entirely sufficient for CRISPR/Cas9 gene editing applications. − However, although AX+ appears to be an effective nanocarrier for use with plants, it is critical to ascertain whether AX+ imposes any toxicity on the plants that will be treated.

Assessing the Potential Toxicity of AX+ NPs through Phenotypic and Genotypic Analysis

To explore the potential of AX+ for novel agricultural applications, its interactions with plants must be thoroughly investigated at both physiological and molecular levels. A key consideration is whether AX+, as a nanosized material, induces phytotoxic or genotoxic effects when applied at specific concentrations. We exposed tomato seeds and seedlings representing an actual agricultural crop to relatively high doses of AX+. We then assessed plant responses through standard germination tests, phenotypic analysis of germinated seedlings, and comprehensive transcriptomic profiling using RNA-Seq technology.

As a plant-originated biomolecule, AX is not anticipated to possess strong phytotoxic properties. AX was tested for toxicity in animals earlier, where AX extracted from psyllium husk has been demonstrated to present no acute toxicity in mammals. However, it is well-known that materials can possess unique properties at a nanoscale that are not expected for the original bulk material, and charged particles can induce biological effects different from those of their neutral counterparts. Previously, we demonstrated that nanoscale carbon-based materials elicit biological responses in planta that are absent from non-nanoscale carbon (activated carbon). Additionally, multiple sources have reported that positively charged NP elicit higher cytotoxicity compared to negatively charged NP. , To determine if AX+ elicits any effects on tomato seed germination, AX+ was applied in three doses to seeds as a direct air spray or as a supplement to the MS agar growth media (Figure ). It was noted that AX+ did not impose any delay of germination after 14 days when applied as a direct spray to tomato seeds at 100 or 500 mg/L doses (Figure A,B). Similarly, when AX+ was added as a supplement to standard MS agar media, no inhibition of germination was observed at both tested doses (Figure C,D).

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Germination of tomato seeds in the presence of AX+ applied as air spray or supplement to growth medium. (A, B) Images (A) and germination rate (B) of tomato seeds sprayed with AX+ water solutions (100 mg/L and 500 mL) over 2 weeks postapplication. n = 10. (C, D) Images of seedlings germinated from seeds (C) and germination rate (D) of tomato seeds incubated on MS medium (control) and MS supplemented with 100, 250, and 500 mg/L AX+ over 2 weeks. Error bars represent SE, Statistical significance was determined by one-way ANOVA, α = 0.05.

The seedlings cultivated on MS media supplemented with AX+ (Figure ) were further assessed phenotypically (Figure S2). Supplementation of the growth medium with AX+ did not result in a significant reduction of the length of roots and stems of exposed tomato seedlings (Figure S2A). However, a statistically significant and dose-dependent decrease in the total fresh biomass of the AX-exposed seedlings was noted (Figure S2C). No biomass reduction was observed for seedlings grown from the seeds exposed to AX+ through sprayed applications. Overall, we concluded that relatively high doses of AX+ (250 mg/L, 500 mg/L) and long-term exposure to AX+ may cause some toxicity to exposed plants. However, for experiments utilizing pDNA–AX, a concentration of 75 μg/mL was used, which is substantially lower than the threshold that induces negative effects. If AX+ is intended for the delivery of specific chemicals and agents essential for plant agriculture, it is crucial to evaluate and test the potential adverse effects associated with high-dose applications. To further assess the possible phytotoxicity of AX+, we examined whether AX+ treatments influence the molecular processes in exposed plants. Therefore, RNA-Seq was used to analyze the total transcriptome of tomato seedlings subjected to AX+ exposure.

Assessing the Transcriptomic Influence of AX+ on Exposed Tomato Plants

Investigating how AX+ influences the total transcriptome of exposed plants is essential for understanding the potential risks tied to the use of nanomaterials in agricultural and environmental applications. This level of study may provide insights into the whole gene network and signaling pathways affected by nanomaterials, − and can also lead to the identification of particular transcripts associated with nanotoxicity. , Here, we aimed to understand whether AX+ can significantly influence the transcriptome of tomato seedlings germinated and grown in the presence of AX+ for 21 days. The tomato (cv. ‘Micro-Tom’) was chosen for this task because it serves as a model species for molecular research, with a fully sequenced transcriptome and accessible databases. Figure describes a summary of the results of the RNA-Seq principal component analysis (PCA) of the sequenced RNAs revealed that exposure to AX+ resulted in substantial differences in total gene expression between groups of untreated seedlings and groups of seedlings exposed to AX+ (Figure A). Overall, 16729 differently expressed genes (DEGs) were identified as common across all samples (Figure B). Notably, groups exposed to higher AX+ concentrations (250 and 500 mg/L) displayed greater divergence from the control group compared to those exposed to the lowest dose (100 mg/L) (Figure A). These dose-dependent effects highlight the significant impact of AX+ on the tomato transcriptome.

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Transcriptomic analysis of tomato seedlings grown on MS mediums supplemented with AX+ in 100, 250, 500 mg/L doses. (A), PCA score plot of the gene expression profile for seedlings grown on plain MS, or grown on MS medium supplemented with either 100, 250, or 500 mg/L AX+ for 21 days. n = 3. (B) Venn diagram of total identified differently expressed genes (DEGs) across all samples. (C-E), KEGG pathway enrichment analysis of seedlings grown on MS mediums supplemented with (C) 100 mg/L, (D) 250 mg/L, and (E) 500 mg/L AX+. Dot size corresponds to the number of genes (count) associated with each pathway, while the color gradient reflects the adjusted p value (padj), ranging from indigo (p = 1.00) to red (p = 0.00). The x axis represents the gene ratio, calculated as the proportion of input genes associated with each pathway to the total number of genes in the dataset. “BS”: biosynthesis.

124 DEGs were associated with both 250 and 500 mg/L treatments (Figure B). 1778 DEGs were substantially changed compared to the control (log2 fold change > 1.5) across all treatments. Of these, 170 DEGs were from the 100 mg/L treatment, 647 from the 250 mg/L treatment, and 961 from the 500 mg/L treatment, suggesting that with an increased concentration of AX+, more genes are dysregulated. The top 20 KEGG pathways that were differentially regulated in 100, 250, and 500 mg/L-treated samples against that of control, including gene counts for each pathway, are shown in Figures S3 and S4). The most significantly impacted metabolic pathways from AX+ exposure were pathways associated with phenolic compound biosynthesis and plant hormone signaling (Figures C–E, S3, and S4).

A list of the top 100 most-effected DEGs and volcano plot visualization for all concentrations of AX+ are available in Tables S1–S6 and Figure S5. From these top 100 most dysregulated genes, 29 of them were found to be common between the 250 and 500 mg/L AX+ seedlings (Table S4). Notably, in these 29 DEGs, the trend of dysregulation was typically conserved between concentrations and often amplified in the 500 mg/L group compared to the 250 mg/L group (Table S4). A majority of these DEGs were downregulated (22) rather than upregulated (7) and include genes that play a key role in flavonoid, secondary metabolite, and alkaloid biosynthesis, such as chalcone synthase B (Table S4). Though there is a definite, substantial, dose-dependent impact of AX+ on the tomato transcriptome, it is important to note that the phenotypical differences observed in the germinating seedlings were rather slight; the only parameter that was impacted was the fresh biomass (not dry weight), and this effect was only observed in the two highest concentrations of AX+ (Figure S2). Validation of RNA-Seq data was determined using RT-qPCR to confirm similar expression levels of the genes Aquaporin TIP3-2-Related and Sugar Transporter ERD6-Like 16 in the transcriptomic data (Figure S6). Primers used for validation are available in the Supporting Information (Table S5).

The use of biodegradable soft nanopolymers derived from plant waste as nanocarriers holds great potential for advancing plant agriculture and plant genetic engineering. These nanopolymers offer a sustainable and eco-friendly approach to delivering agrochemicals, nutrients, and genetic materials to plants with high efficiency and minimal environmental impact. In plant agriculture, such nanocarriers can improve the targeted and controlled release of fertilizers, pesticides, and growth regulators, reducing chemical waste and enhancing crop productivity. , Their biocompatibility ensures that they do not cause toxicity to plants, soil, or surrounding ecosystems, making them a safer alternative to synthetic carriers. Their unique nature allows them to penetrate plant tissues without causing damage, facilitating efficient gene transfer without the need for conventional, often invasive, expensive and labor-intensive methods such as Agrobacterium-mediated transformation. − or biolistic delivery. Furthermore, their ability to be functionalized with specific biomolecules enables precise targeting, improving the success rate of genetic modifications.

Here, we have provided experimental evidence that a natural polymeric nanomaterial derived from wheat bran (AX) can efficiently deliver nucleic acids (pDNA) and covalently bound dyes (FITC) into plant cells with intact cell walls. The relatively simple conjugation of AX+ with DNA and covalent conjugation of AX with dye, combined with the absence of significant negative effects on plant growth and gene expression, highlight AX-based nanocarriers as a promising option for agrochemical and nucleic acid delivery. This capability can potentially enhance crop protection and nutrient uptake while reducing the need for excessive chemical application. Integrating waste-derived AX into agricultural practices supports the principles of circular agriculture, enhancing sustainability by reducing the reliance on synthetic chemical inputs. Additionally, in plant genetic engineering, AX presents a promising vehicle for delivering DNA, RNA, or gene-editing tools, such as CRISPR/Cas directly into plant cells. This innovative approach has the potential to streamline plant transformation processes, significantly reducing both time and costs, while providing a more efficient and environmentally sustainable alternative for crop improvement.

Supplementary Material

jf5c13415_si_001.pdf (1MB, pdf)

Acknowledgments

The authors would like to acknowledge and thank Scott Payne and Jayma Moore from the Advanced Imaging and Microscopy Laboratory at NDSU and Ms. Jimli Goswami from the Department of Civil, Construction and Environmental engineering, NDSU for electron microscopy and UV-Vis experiment, respectively. The authors appreciate Claudia Vickers for donating plasmid DNA pGFPGUSPlus used in experiments, as provided by Addgene. The authors are grateful to the UAMS Digital Microscopy Core Laboratory and personally to J. Kamykowski for providing access to UAMS confocal microscopy. The foundational infrastructure for this research was previously established with funding from USDA-NIFA (ARFI 2020-04096 to M.K.). This work also was partially supported by the National Science Foundation under NSF EPSCoR Track-1 Cooperative Agreement OIA #1946202 (to M.Q.).

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jafc.5c13415.

  • Complementary physicochemical, phenotypic, and molecular evidence for experiments involving AX+ and AX–FITC, including spectroscopic analysis of FITC-AX compared with free FITC (Figure S1); phenotypic assessments to quantify root and shoot growth along with fresh and dry biomass under multiple AX+ concentrations (Figure S2); transcriptomic profiling through KEGG pathway enrichment analyses of the top 20 up- and downregulated pathways across 100, 250, and 500 mg/L treatments (Figures S3 and S4) alongside volcano plots visualizing global differential gene expression patterns (Figure S5); targeted RT-qPCR validation to measure relative transcript levels of aquaporin TIP3-2-related and sugar transporter ERD6-like 16 genes normalized to 18S (Figure S6); gene-level datasets listing the top 100 differentially expressed genes at each concentration (Tables S1–S3); genes shared between higher-dose treatments (Table S4); and primer sequences used for expression analyses (Table S5) (PDF)

#.

Department of Materials Science and Engineering, University of Central Florida, Orlando, FL 32816, USA

†.

M.R. and K.V. contributed equally to this work.

The authors declare no competing financial interest.

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