Plant Microtechnique with Resin: Towards Plant Histolomics

TL;DR

This paper introduces a new resin-based method for plant histology that improves image quality and enables quantitative analysis of plant tissues.

Contribution

The novel resin-based microtechnique and histolomics framework enable reproducible, quantitative plant histology.

Findings

The resin-based method improves section stability and image reproducibility compared to traditional paraffin embedding.

A trichrome stain and adhesive treatment enhance contrast and allow detailed visualization of tissues and organelles.

A workflow using MATLAB and Photoshop enables morphometric and compositional analysis of plant histolomes.

Abstract

Plant microtechnique involves a sequence of skill-intensive histological procedures that often yield poorly reproducible images and limited quantitative information. Nevertheless, it provides an essential cellular and tissue context needed to understand biological functions. In this work, we present an optimized resin-based microtechnique that replaces paraffin embedding, incorporates a chemically activated adhesive treatment for glass slides, and develop a trichrome stain for resin sections. All these improvements enhanced section stability and image reproducibility, enabled a broader color palette with sharp contrast of tissues, cells and organelles, and selected ultrastructural features using light microscopy. Based on these preparations, a quantitative micrograph analysis workflow was developed based on image segmentation and feature extraction using MATLAB (R2024a) and Adobe Photoshop (CS6). This approach enables the measurement of a wide range of morphometric and compositional features, generating structured histological datasets that we refer to as plant histolomes. As an illustrative application, this workflow was applied to leaves from several model plants species and integrated multiple anatomical traits into a composite feature, the “C4 Kranz-anatomy level”, enabling quantitative comparison along the C3-C4 anatomical transition. The resin-based microtechnique and the histolomics framework developed in this work provide a robust and reproducible basis for the quantitative plant histology, bridging classical microscopy with a data-driven tissue analysis.

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Full text

1. Introduction

Omics approaches have been widely used for the quantitative and large-scale study of diverse molecular components [1,2]. Molecular abundances are often correlated with tissue, organ-, or whole-plant phenotypes [3]. However, the biological activity of molecules depends on tissue, cellular, and subcellular structures, as well as their spatial localization, which often are overlooked or poorly studied when inferring molecular functions from omics data [4,5].

Plant histology is a descriptive discipline that studies the microscopic structures and composition of plant tissues, with the aim of understanding their organization, development, and function [6,7]. It provides the structural and spatial context that complements the understanding of molecular functions [4]. However, plant histology tends to lack reproducibility and is mostly qualitative, hindering precise integration with omics data [7,8]. These limitations arise from skill-dependent microtechnique methods and the lack of systematic strategies for quantitative micrograph analysis.

Plant microtechnique is a methodological discipline that encompasses the procedures required to prepare plant samples for microscopic examination and downstream histological interpretation. Classic microtechnique includes sampling, fixation, dehydration, paraffin embedding, sectioning, affixing (attachment to glass slides), staining, immunostaining, dehydration, mounting (resin and coverslip placement), and microscopy observation [9]. Compared with in vitro omics methods, microtechnique is more artisanal, which limits its reproducibility [10]. Reproducibility is mainly affected during the most skill-dependent manual steps, which occur form affixing through mounting. During these steps, sections must be carefully handled, adhered to glass slides, and exposed to multiple wet treatments. Paraffin sections are fragile and they may wrinkle or tear during affixing and may detach or become scratched during subsequent wet steps on glass slides [11]. Moreover, paraffin is removed after affixing, leaving the tissue section eve more vulnerable. As a result, artifacts may be introduced, distorting the histological results.

Successful section handling during skill-dependent steps depends on a careful balance of variables such as tissue type, section thickness, adhesive treatment of glass slides (gluing), affixation conditions, subsequent wet-step conditions, and operator skill. Plant sections are more challenging to obtain and handle than animal sections due to high water content, rigid cell walls, fibers, and hard crystals that easily disrupt soft paraffin sections during cutting [11]. Thicker sections are generally easier to handle and less prone to tearing during cutting, but they inherently reduce cellular and subcellular resolution due to overlapping focal planes [12]. Gluing treatments typically consist of slide coatings with gelatin, casein, albumin, or polylysine solutions [13,14,15]. However, these methods exhibited limited adhesive performance, generated uneven and intense background staining, and are prone to scratching, increasing the likelihood of artifacts [16]. During affixation, sections are placed in a warm water bath to flatten, collected onto a glass slide, and heated at 30 °C overnight to promote adhesion [14]. Sections are then deparaffinized and subjected to wet procedures such as staining, immunostaining, and mounting. During these steps, sections may detach from the glass slide if adhesion is insufficient or may be accidentally scratched. Ultimately, success heavily relies on paraffin mechanical stability; slide adhesiveness, and operator skill.

Replacing paraffin with a harder, permanent embedding medium and improving glass-slide gluing could enhance microtechnique robustness and reproducibility. LR White, an acrylic resin commonly used in transmission electron microscopy (TEM), is a harder embedding medium that is also compatible with light microscopy [17]. It has shown excellent performance for producing thin and ultrathin sections of plant leaves containing hard calcium oxalate crystals while preserving ultrastructural detail [18]. Its hydrophilic nature allows sections to be analyzed without resin removal, maintaining mechanical support throughout the skill-dependent steps.

Glass-slide preparation for microtechnique can be divided into two steps: cleaning and gluing. Thorough glass-slide cleaning is critical to eliminate residues from manufacturing and handling, such as oils, dirt, and smudges, which may cause artifacts or poor section adhesion. Various strong acidic, basic, and oxidizing solutions have been formulated to deeply clean and simultaneously activate glass slides, including MeOH/HCl, H_2_SO_4_/K_2_CrO_4_ (sulfochromic), H_2_O_2_/H_2_SO_4_ (piranha), CH_3_COOH, H_2_SO_4_, and NaOH [19,20,21]. These solutions clean beyond water and soap by dissolving or oxidizing contaminants. Activation generates hydroxyl groups that react with (3-aminopropyl)triethoxysilane (AES) during a downstream silanization process (gluing), yielding a positively charged glass surface [22]. Unlike classic protein-based coatings, which provide an uneven, non-covalent adhesive layer, AES-based gluing provides a more uniform, covalent-bonded adhesive layer.

Staining and immunostaining are fundamental wet methods in microtechnique for assessing morphology, general composition, and protein localization [9]. Johansen’s staining has long served as a reference method in plant microtechnique, method that has undergone several modifications since 1940 [9,23,24,25,26]. It relies on safranin, which stains lignin- and suberin-rich cell walls magenta, and fast green, which stains cytoplasmic proteins turquoise [27,28,29]. Johansen’s staining visualizes global tissue structure and composition, whereas immunostaining reveals the spatial localization of specific proteins [4]. On the other hand, several dyes have been used to stain plant resin sections, including light green, Coomassie blue, acid fuchsin, safranin, iodine, periodic acid–Schiff (PAS), methyl violet, and toluidine blue [30,31,32]. However, to our knowledge, the effectiveness of Johansen’s staining on resin sections remains unclear.

After successful microtechnique execution, many researchers remain uncertain about how to extract meaningful information from micrographs. Histological interpretation is often limited to qualitative descriptions of abnormalities or immunolabel localization. A systematic and quantitative histological analysis would allow researchers to fully exploit micrograph information. Histological features can be described in terms of morphology, morphometry, and composition: morphology refers to qualitative structure; morphometry quantifies abundance, size, and shape; and composition reflects the overall or specific molecular composition of cellular structures [33,34,35]. Staining supports boundary-based morphological and morphometric analyses, as well as color-based compositional analyses, while immunostaining enables analysis of localization and abundance of specific molecules. These analyses are traditionally performed manually and visually. However, digital tools—such as color measurement and image segmentation—allow these features to be converted into quantitative data far exceeding what can be achieved by eye alone [36].

Overall, integrating a robust microtechnique with a systematic image-analysis workflow would enable the generation of reproducible images suitable for obtaining histological datasets. This would allow quantitative comparison of plant histological features across species and/or conditions, and bridge the gap between omics data and plant histology. Here, the term “histolomics” was introduced and refers to a descriptive, data-driven discipline that studies the microscopic structure and composition of tissues in a systematic, large-scale manner, enabling quantitative tissue phenotyping and characterization of tissue responses to different stimuli. Likewise, the “histolome” was defined as the set of qualitative annotations and quantitative features extracted from one or a small number of representative, high-resolution histological sections that collectively capture the structural and compositional diversity of a plant organ. To our knowledge, this is the first work to propose plant histolomics. An analogous discipline known as “histomics” exists; however, it primarily focuses on mining digital pathology images using machine-learning algorithms to extract histological features from hematoxylin- and eosin-stained animal tissues [37,38]. Moreover, histolomics does not address microtechnique optimization, which is a central part of the present work.

Here, we present a novel resin-based plant microtechnique with streamlined and robust skill-dependent steps, increasing the reproducibility of histological images. A strategy for quantitative image analysis of multiple histological features using MATLAB and Adobe Photoshop is also presented. This workflow was validated across several C_3_ and C_4_ plant leaves commonly used in plant research, as well as amaranth seeds and seedlings.

2. Results

2.1. Glass-Slide Adhesive Treatment

To reduce section detachments during microtechnique wet steps, previously reported methods for glass-slide cleaning, activation, and AES-based gluing were evaluated to identify the most suitable workflow for resin sections. Several cleaning and activation solutions were tested, which originally were developed for silicon wafers and glass slides, such as MeOH/HCl, H_2_SO_4_/K_2_CrO_4_ (Sulphocromic), H_2_O_2_/H_2_SO_4_ (piranha), CH_3_COOH, H_2_SO_4_, and NaOH [19,20]. Next, the effectiveness of cleaning and activation of the glass-slide was evaluated by measuring the water contact angle. Sulphocromic and piranha solutions produced the best results, yielding the highest wettability and therefore the lowest contact angles. Activated slides were subsequently “glued” with AES. All treatments showed a decrease in wettability and a corresponding increase in contact angle after gluing (Supplementary Figure S1a,c). Finally, the effectiveness of the gluing treatment was tested by affixing a resin section to each slide and subjecting it to strong washing steps. Again, sulphocromic- and piranha-activated slides performed best, retaining the entire section, whereas other treatments resulted in partial or complete section detachment (Supplementary Figure S1b).

2.2. Trichome Staining for Resin Sections

To support the interpretation of the obtained results, an overview of Amaranthus cruentus leaf histology is shown in Figure 1 and Figure 2. Johansen’s reference staining did not work on resin sections; therefore, it was redesigned and expanded to a trichrome method. Toluidine blue, Johansen’s, and Coomassie’s staining were tested and the colors acquired by cell walls (CW), protoplasm (PR), and resin backgrounds were measured in HSB (Hue, Saturation, Brightness) color space.

Hue values depended on the dye, saturation reflected staining performance, and brightness was mainly determined by microscope and camera settings and remained similar across micrographs. Toluidine blue produced monochromatic staining (similar hues), and Johansen’s produced polychromatic but uneven and poorly saturated colors (Figure 3). Achieving different hues between cell structures is essential for identification and digital segmentation, as is maximizing saturation relative to the unstained resin background. By testing combinations, we found that Coomassie’s solvent enabled staining of resin sections with Coomassie dye itself as well as with Johansen’s dyes (Safranin and Fast Green), producing polychromatic, uniform, and well-saturated colors (Figure 3). We therefore reformulated Johansen’s staining and added a third dye—Lugol’s iodine—resulting in a trichrome staining (Figure 4).

Trichrome staining revealed the morphology and general composition of amaranth leaves. Individually, Fast Green stained the protoplasm turquoise, Safranin stained most tissue magenta, and iodine specifically stained starch granules mauve. None of the dyes stained vacuoles or the resin background, and Fast Green and Safranin did not stain starch granules (Supplementary Figure S2). Because the protoplasm (PR) comprises the cytosol and all organelles, including the vacuole (Figure 1), the term proteoplasm (PT) was introduced to refer specifically to the turquoise (proteinaceous) fraction: the cytosol and all organelles except the vacuole and starch granules.

A synergistic effect emerged in the trichrome combination: Safranin selectively stained cell walls, and both hue and saturation values increased. Proteoplasm shifted from turquoise to blue hues, while cell walls shifted into magenta and reddish hues (Supplementary Figure S2). The cell wall and proteoplasm colors in the epidermis, mesophyll, and bundle sheath were measured. Proteoplasm hues were similar across tissues, but bundle sheath cell walls displayed magenta, reddish, and even orange hues, whereas epidermis and mesophyll cell walls displayed only magenta. The bundle sheath also contained both stained and unstained starch granules, a pattern confirmed by TEM, which revealed electron-dense and electron-lucent granules (Figure 5).

The order and timing of dyes were crucial for achieving selective coloration. Fast Green–stained sections could be counterstained with Safranin for 5–10 s; longer exposure caused Safranin to replace Fast Green (Supplementary Figure S3). Conversely, Safranin-stained sections could not be properly counterstained with Fast Green (Supplementary Figure S4). Iodine staining—whether used alone or as a counterstain—showed unpredictable intensity, ranging from absent to intermediate or intense, even between serial sections from the same sample on the same slide. We could not identify the underlying cause of this variability. However, within each individual section, staining intensity was always uniform, regardless of whether it was absent, intermediate, or intense (Supplementary Figure S5).

Some technical considerations are important for optimal staining and micrograph interpretation. The orange hue in bundle sheath cell walls was most evident with a 40× objective and a diaphragm aperture equivalent to 10×. Mounting with Entellan improved sharpness immediately and continued to improve as it dried. Iodine staining may fade over days to months; thus, micrographs should be acquired within one to three days after mounting. Finally, common artifacts associated with resin sections must be recognized for accurate interpretation (Supplementary Figure S6).

2.3. High-Resolution Resin-Based Microtechnique

The resin-based microtechnique allowed light microscopy to resolve small organelles and even ultrastructures typically visible only by TEM. Trichrome staining identified cell walls in magenta to orange, proteoplasm in turquoise, and starch granules in mauve. Differences in proteoplasm saturation enabled identification of cytosol, chloroplasts, mitochondria, nuclei, and nucleoli (Supplementary Figures S7 and S9). An intensely iodine-counterstained paradermal section revealed two layers in the epidermal cell wall—an internal orange layer and an external magenta layer. The same section revealed additional ultrastructural features, including membranes within vacuoles, chloroplast-associated protein granules, rod-shaped organelles of unknown identity, and vesicles involved in cell-wall material transport (Supplementary Figure S10).

2.4. Histolomic Analysis

To analyze micrographs obtained using the resin-based microtechnique, a four-step workflow was developed, which includes visual analysis, color analysis, micrograph segmentation, and data analysis.

Visual analysis consisted of classical qualitative identification of tissues and morphology (Figure 1 and Figure 2), with the added identification of organelles and other features enabled by the high resolution of resin sections. Comparing young and senescent amaranth leaves, whole-micrograph differences included decreased proteoplasm, increased starch granules, and rounding of palisade mesophyll cells in senescent leaves. At higher magnification, additional differences were observed: reduced cell wall thickness, appearance of starch granules in palisade mesophyll, and loss of cytosol (Supplementary Figure S11). Color analysis involved measuring the color of each structure and assigning hue ranges for segmentation (Figure 5).

Micrograph segmentation digitally isolated cellular structures into layers for further analysis (Supplementary Video). The color contrast provided by the trichrome staining enabled segmentation based on color (composition) and boundaries. We semi-automatically segmented cell walls, proteoplasm, and unstained elements (vacuoles, starch granules, background) based on hue and saturation. Proteoplasm could be further segmented into low-saturation components (cytosol and nucleus) and high-saturation components (mainly chloroplasts and mitochondria). Layers clarified tissue composition and spatial patterns, but the thin, discontinuous mesophyll cell walls prevented automated boundary-based morphometric analysis (Supplementary Figure S12). Therefore, we manually segmented each cell. Segmented micrographs for young and senescent leaves are shown in Supplementary Figures S13 and S14.

Data analysis involved defining and measuring primary and derived morphometric and compositional features. Morphometric features included number, size, perimeter, area, thickness, and circularity; compositional features involved semi-quantitative densitometric analysis of stained molecules (proteins, lignin/suberin, starch, and immunolabels). Senescent leaves showed smaller and more rounded palisade cells, reduced cell wall thickness, decreased proteoplasm, and significantly increased starch content, whereas lignin content did not differ (Supplementary Figure S15). Intense iodine counterstaining enabled additional segmentation of starch granules, as well as inner and outer cell wall layers, based on iodine-driven color shifts (Supplementary Figure S16).

2.5. Resin-Based Microtechnique and Histolomics Validation

The resin-based microtechnique yielded reproducible images across species and tissues. To test its versatility and the potential of histolomic analysis, we applied it to three C_3_ leaves (Arabidopsis thaliana, Triticum aestivum, and Glycine max) and two C_4_ leaves (Zea mays, and Amaranthus cruentus). The dye selectivity matched that observed in amaranth leaves (Figure 6 and Figure 7 and Supplementary Figures S17–S19). Histolomic analysis revealed a number of similarities and differences between the leaf sections: the same fundamental tissues but different morphology, morphometry and composition.

To integrate these data into a physiologically and evolutionarily meaningful metric, we propose a composite feature termed the C_4_ Kranz level. This feature, shown in Table 1, integrates primary and derived morphometric and compositional features associated with C_4_ Kranz anatomy [5], serving as an illustrative example of the utility of histolomic analysis.

Microtechnique was also applied to amaranth seed germination and cotyledon maturation, revealing organelle development and wall thickening (Supplementary Figures S20 and S21). Finally, the trichrome staining also worked on paraffin sections, even without deparaffinization (Supplementary Figure S22).

The images shown in Figure 6 and Figure 7 and in Supplementary Figures S11 and S17–S19 are segmented images in which the intercellular space layer and the resin background outside the tissue section were omitted for visualization and analysis. The corresponding original, unsegmented micrographs are provided in Supplementary Figure S23 to allow verification of staining homogeneity and to ensure full transparency regarding all image-processing steps applied during segmentation.

2.6. Immunostaining of Trichrome-Stained Resin Sections

Trichrome staining is compatible with immunostaining, enabling precise protein localization. We performed immunostaining on amaranth leaf sections to detect the RuBisCO large subunit (RbcL). BCIP/NBT development produced an intense blue signal and did not interfere with trichrome staining, resulting in a tetrachrome staining. Micrographs were segmented according to hue ranges, and protein and RbcL concentrations were quantified. RbcL was present in nearly all tissues but was most abundant in the bundle sheath and mesophyll; the bundle sheath contained approximately twice as much RbcL as the mesophyll (Figure 8).

3. Discussion

3.1. Improving Robustness and Reproducibility of Plant Microtechnique

The resin-based microtechnique streamlined and improved the robustness of skill-dependent steps. Resin embedding provided continuous mechanical support, reducing tearing and scratching and thereby better preserving tissue structure (Supplementary Figures S22 and S23). Glass-slide cleaning, activation, and AES gluing ensured reliable section adhesion; Piranha- or sulphocromic-activated slides completely eliminated section detachment issues (Supplementary Figure S1). These advantages reduced failure rates during skill-dependent steps, thereby increasing robustness and reproducibility. Because of the toxicity of Cr^6+^, Piranha solution is preferable. Notably, classic methods overlook slide activation and recommend minimal gluing time, which in our experience is insufficient [39]. Activation also cleaned slides deeply, reducing dirt artifacts.

Present trichrome staining method (Figure 4) reduced wet steps, time, reagents, toxicity, and variability compared with classical methods [9,23,24,25,26]. Johansen’s staining involves many steps, large reagent volumes, and toxic or explosive compounds such as picric acid [9]. Our method reduces approximately 18 wet steps to 8, shortens processing time from hours to minutes, and decreases the number of reagents from nine to six, thereby reducing the risk of section detachment and mechanical damage. It also relies on small volumes of accessible, less-toxic reagents, enabling easy disposal and preventing reagent reuse, which further improves reproducibility (Supplementary Video).

However, the trichrome staining method still relies on two hazardous reagents for glass-slide cleaning and silanization: piranha solution and (3-aminopropyl)triethoxysilane (AES). Piranha solution can be reused multiple times until its oxidizing capacity is exhausted; because glass slides are pre-cleaned with water and detergent, its effective working lifetime is expected to be long. Both reagents must be disposed of in accordance with local regulatory guidelines. Among them, AES represents the primary chemical waste generated by our workflow.

3.2. Structural and Compositional Information Rendered by Trichrome Staining

Trichrome staining revealed the molecular composition of cell structures. Fast Green reportedly stains proteins and is used to quantitatively stain SDS–PAGE gels and nitrocellulose membranes [27,28]. Safranin stains lignin, suberin, nuclei, and chromosomes, although its effectiveness in staining chromatin is not well-documented [29,40,41]. Iodine reacts with polysaccharides, with the highest affinity for amylose [42,43]. Thus, trichrome staining was consistent with the reported molecular specificity of the dyes, except for the nucleus and chromatin, which stained turquoise (Figure S8).

T. aestivum (Supplementary Figure S19) and Z. mays (Figure 6) nuclei showed heterochromatin, but only those of T. aestivum stained magenta. Safranin can interact with DNA [42]. However, it was observed that safranin alone stained almost all tissue structures; its specificity was defined by counterstaining with fast green (Supplementary Figure S2). Therefore, there should be specific molecules responsible for the safranin staining in T. aestivum heterochromatin. Similarly, trichrome staining revealed stainable and non-stainable starch granules (Figure 5). The amylose content could explain the differences. However, the unstained granules were whiter and more electrolucent than the background in light and electron microscopy. On the other hand, the evidence is suggestive but not conclusive if non-stained granules are the same as electrolucent granules. We can only conclude that the amaranth leaf has two types of starch granules with different compositions and interactions with LR white resin. Strikingly, both types can be formed in the same chloroplast.

Safranin counterstaining with iodine causes a shift from magenta to reddish and orange hues, differentiating the molecular composition of the cell wall. Our evidence suggests that this process occurs gradually and selectively, beginning in the bundle sheath cell wall (Figure 5) and progressing until most cell walls across all tissues undergo a color shift (Supplementary Figure S16). Yet, the epidermal wall showed an internal orange wall and an external thicker magenta wall. Furthermore, color segmentation showed that all cell walls retained a small magenta portion (Supplementary Figure S16).

The plant cell wall consists of multiple layers arranged from the exterior to the interior: the middle lamella, primary wall, secondary wall, and plasma membrane. The primary wall is mainly composed of cellulose, hemicellulose, proteins, and pectins, whereas the secondary wall is enriched in cellulose, hemicellulose, and lignin [44]. In epidermal cells, additional outer layers of cutin and wax are present [45]. The primary wall is typically thin (approximately 0.1 µm), while the lignified secondary wall is substantially thicker, ranging from ~2 to 10 µm [46].

The evidence suggests that safranin stains both lignified and non-lignified cell walls, with color differences reflecting wall type and thickness: orange-stained walls correspond predominantly to thicker secondary walls, whereas magenta-stained walls correspond to thinner primary walls. Safranin is known to stain lignin and suberin. Lignin can form complexes with iodine that alter its UV–visible absorption spectrum, while the interaction between iodine and suberin remains poorly understood [47,48,49].

The interaction between lignin and iodine may therefore explain the color shift observed in lignified secondary walls, while non-lignified primary walls and cutinized epidermal walls remain magenta. However, this interpretation is complicated by the observation that the ring structures of tracheids, which are also lignified, did not exhibit a comparable shift toward orange hues [33] (Supplementary Figure S10).

Together, these observations indicate that iodine counterstaining introduces additional layers of compositional information for cell walls and starch granules beyond that provided by the reference Johansen’s staining. The chemical basis of iodine interactions with these structures, as well as their potential compositional heterogeneity, remains incompletely understood. Further research is needed to clarify these interactions and reduce variability in iodine staining intensity.

3.3. Resin-Based Microtechnique Resolution

The resin-based microtechnique (RBM) resolved tissues, cells, organelles, and ultrastructural features, enabling large-scale histolomic analysis. The smallest structures resolved were membranes within bundle sheath vacuoles (≈336–560 nm), which were confirmed by TEM to correspond to cytosolic invaginations (Supplementary Figures S8 and S10). These dimensions approach the resolution limit of widefield light microscopy (≈200 nm in XY and 500–700 nm in Z) [50] and overlap with the ultrastructural information typically accessible at low TEM magnification. This “bridge” between light and electron microscopy underpins the histolomic potential of RBM: TEM can be used to characterize ultrastructure in detail, while high-resolution light microscopy enables quantitative analysis across large tissue areas. In addition, the resin section thickness (≈500 nm) remains within the depth of field (z resolution) of widefield microscopy, minimizing out-of-focus interference and conceptually resembling confocal optical sectioning. Thus, trichrome staining of resin sections complements high-resolution fluorescence approaches, which are restricted to labeled targets. Achieving and preserving plant paraffin sections at comparable thicknesses throughout the skill-dependent steps is considerably more challenging than with resin sections.

However, resolution is not determined solely by optical performance and tissue preservation; it also depends on sufficient color contrast between cellular structures to allow their discrimination (Supplementary Figure S12). Trichrome staining resolved multiple structures based on their differential responses to the dyes, an effect further enhanced by the single focal plane provided by thin resin sections (Supplementary Figures S10 and S16). In addition, microtechnique relies on accurate color description, as color has qualitative interpretative relevance in histology. The HSB color space effectively separates dye identity (hue), staining performance (saturation), and microscope acquisition settings (brightness) (Figure 3). Hue differences enabled segmentation of up to four distinct structures; however, color overlap—analogous to fluorescence bleed-through—should be avoided to ensure reliable segmentation (Figure 8).

3.4. Histolomics Approach

Digital micrographs should be analyzed beyond visual inspection. Primary and derived morphometric and compositional features captured broad histological differences between C_3_ and C_4_ leaves (Figure 6 and Figure 7; Supplementary Figures S17–S19). However, to move beyond classical qualitative histological descriptions toward meaningful quantitative comparisons, composite features—such as the C_4_ Kranz anatomy level—must be developed and standardized.

Plants are broadly classified as C_3_ or C_4_ based on leaf anatomy and the spatial distribution of photosynthetic enzymes, particularly RuBisCO. C_3_ leaves exhibit small, inconspicuous bundle sheath cells and randomly distributed chloroplasts, whereas C_4_ leaves are characterized by enlarged bundle sheath cells arranged in a wreath-like Kranz anatomy and containing large, centripetally or centrifugally positioned chloroplasts (Figure 1). In C_3_ plants, RuBisCO is localized in mesophyll cells and is directly exposed to atmospheric oxygen, leading to photorespiration and reduced photosynthetic efficiency. In contrast, C_4_ plants spatially isolate RuBisCO within bundle sheath cells and operate CO_2_-concentrating mechanisms that suppress photorespiration and enhance carbon fixation. This anatomical and biochemical specialization represents an evolutionary adaptation to declining atmospheric CO_2_ and rising O_2_ levels and emerged through a multistep transition involving intermediate anatomical and enzymatic states [5].

To study this multistep, histology-related process, we selected and weighted primary and derived features associated with C_4_ Kranz anatomy (Table 1). The weighting was performed by assigning a “feature score” that represents the relative importance of each histological feature for C_4_ photosynthesis. These weights reflect evolutionary and functional considerations derived from comparative analyses of C_3_–C_4_ transitions [5]. Bundle sheath (BS) size ratio and BS organelle size ratios represent the relative area of BS cells and their organelles compared with the total area of BS and mesophyll tissues, and were each assigned a weight of 20%. The presence of a photosynthetically competent BS tissue with large chloroplasts constitutes a primary requirement for C_4_ photosynthesis and represents the initial transition from C_3_ to proto-Kranz anatomy.

The centripetal or centrifugal positioning of mitochondria and chloroplasts, which characterizes the subsequent transition from proto-Kranz to C_2_ photosynthesis, was weighted at 15% for each feature. Wreath-like tissue organization—defined by enlarged BS cells forming a continuous ring tightly surrounded by one or two mesophyll cell layers—represents another key anatomical step and was assigned a weight of 15%.

The ultimate functional objective of C_4_ Kranz anatomy is the insulation of RuBisCO from atmospheric O_2_, thereby minimizing photorespiration and integrating the major histological features characteristic of C_4_ tissues. This insulation is achieved through thickened and lignified bundle sheath (BS) cell walls and reduced contact between BS cells and intercellular air spaces. Accordingly, BS cell wall (CW) size ratio (BS CW area relative to the total BS and mesophyll CW area), BS CW lignification (relative lignin enrichment in BS cell walls compared with mesophyll cell walls), and BS air insulation (the proportion of BS–mesophyll contact perimeter relative to the total BS perimeter, including interfaces with intercellular spaces) were used to quantitatively describe RuBisCO insulation from the air. Each of these features was assigned a weighting of 5%.

This composite feature indicates how far a given plant deviates from fully developed Kranz anatomy, relative to the species exhibiting the most advanced Kranz anatomy in this study (Figure 6 and Figure 7; Supplementary Figures S17–S19). At present, it is intended as an illustrative example demonstrating how meaningful quantitative metrics can be extracted from histological images. Weighting histological features according to their quantitative contribution to physiological performance—such as photosynthetic efficiency—would substantially strengthen this approach compared with subjective weighting alone. In addition, the spatial distribution of photosynthetic enzymes represents an important determinant that should be incorporated in future iterations. Despite these limitations, the Kranz anatomy level provides a valuable first framework for the quantitative description and comparison of complex histological features. Where direct quantitative weighting is difficult, consensus-based conventions within the plant research community may facilitate standardized comparisons. Similar strategies could be extended to quantify tissue responses to environmental stress, genetic perturbations, nutrient availability, or developmental stage.

This histolomic analysis challenged the classification of amaranth as a C_4_ species. In amaranth leaves, RbcL, wall thickness, and lignin content increased progressively from palisade mesophyll and spongy mesophyll to bundle sheath. Although the leaf displayed near-ideal Kranz anatomy, yet its RuBisCO distribution resembled that of C_2_ plants [5]. While histolomics seeks to condense complex anatomical features into quantitative metrics, it must also retain the individuality of each cell. In A. thaliana, we found a previously unreported protein-rich vascular bundle proteinaceous cell (VBPC) with smooth morphology and ≈26% of the vascular bundle protein content (Supplementary Figure S17), suggesting a specialized function.

3.5. Current Limitations and Future Work

The RBM enables more robust and streamlined handling during dexterity-dependent steps and provides improved section stability, resolution, and reproducibility compared with paraffin sections under the conditions tested here. Nevertheless, RBM also presents limitations and should be regarded as complementary to, rather than a replacement for, classical paraffin-based microtechnique. Resin sections are more prone to wrinkling during slide affixation than paraffin sections, and RBM requires the use of an ultramicrotome, which is not routinely available in many basic histology laboratories. In addition, paraffin embedding remains particularly advantageous for producing long, continuous serial sections, which are essential for developmental and ontogenetic studies. The performance of paraffin-based methods has also been extensively validated across a wide range of plant organs, including highly lignified tissues, whereas the applicability of RBM to such materials requires further systematic evaluation [9,14]. Accordingly, RBM does not aim to replace the long-standing expertise and methodological strengths of paraffin histology, but rather to provide a robust and reproducible alternative for quantitative histology and image-based analyses that support the development of histolomic approaches.

Color management is critical for capturing and representing true and reproducible colors. Standardized color references, such as Macbeth charts or neutral gray patches, are widely used in photography but have limited applicability in micrograph color management. In addition, the diversity of microscopy setups introduces substantial variability in acquisition conditions, making histolomic comparisons most reliable within the same experiment rather than across datasets generated by different laboratories. At present, adjusting white balance and exposure using the unstained resin background represents the most generally applicable approach. The development of dedicated microscopy setups would substantially improve the robustness of inter-laboratory histolomics comparisons.

More feature descriptors and AI based image segmentation and analysis algorithms should be developed. Improved shape descriptors are needed, as circularity failed to differentiate tissues (Supplementary Figure S15). Positional features—such as the location of organelles within bundle sheath cells—could refine functional interpretations. Manual segmentation is time-consuming limiting sample size and thus statistical power; deep-learning approaches could automate segmentation and measurement of large areas, fully leveraging the resolution of resin-based microtechnique.

The histolomics approach requires dedicated data hosting to enable systematic reporting. Micrographs would benefit from an online platform offering interactive annotations and well-organized datasets, analogous to genomic browsers, to facilitate exploration, comparison, and reuse of histological data (Supplementary Figure S11).

4. Conclusions

Although omics disciplines have advanced through rapidly evolving technologies [1], plant histology has largely relied on techniques developed nearly a century ago [9]. The resin-based microtechnique presented here provides robust and streamlined procedures for preparing plant leaves, seeds, and seedlings for microscopy, yielding reproducible, high-quality images. Building on this foundation, histolomic analysis introduces a systematized digital framework for quantitative interrogation of histological images, enabling large-scale tissue phenotyping and the analysis of structural responses to environmental and physiological stimuli. Although substantial technological challenges remain before histolomics can be broadly adopted, this work outlines a practical roadmap toward that goal. We present the first histolome datasets of commonly studied C_3_ and C_4_ plants, offering a new perspective on plant structure, composition, and their inherent biological complexity and beauty

5. Materials and Methods

5.1. Plant Material

Seeds of Amaranthus cruentus L. cv. Amaranteca (amaranth), Arabidopsis thaliana (Arabidopsis), Triticum aestivum (wheat), Glycine max (soybean), and Zea mays (maize) were germinated in 500 g of BM2 HP germination substrate (Berger, Saint-Modeste, QC, Canada). After 4 weeks, one leaf per plant was harvested. For A. cruentus, one leaf was harvested at 4 weeks (young) and one at 8 weeks (senescent). In addition, during germination of A. cruentus, seedlings were harvested at different stages—from seed to seedlings with mature cotyledons.

5.2. General Sample Processing and Microscopy

Resin embedding was performed using standard methods with some modifications [14,51]. Approximately 3 × 4 mm samples were collected from a primary vein, and the leaf blade. In addition, one seed, one germinating seed, and complete cotyledons at various developmental stages were sampled from A. cruentus. Samples were fixed in 3% glutaraldehyde, 2% formaldehyde, and 0.5% Tween 20 in PBS (5–10 times the sample volume) under vacuum for 20 min, then shaken for 1 h, all on ice. Fixation was continued overnight at 4 °C, followed by three PBS rinses, each with shaking for 1 h at room temperature.

Samples were dehydrated in 30%, 50%, 70%, 95%, and absolute ethanol, with shaking for 1 h at each step. Pre-embedding in LR White resin:absolute ethanol (1:1) was performed overnight with shaking, followed by inclusion in pure resin (#14381, Electron Microscopy Sciences, Hatfield, PA, USA) for another night. Pre-embedding can be shortened to 4 h for light, porous samples such as leaves, and maintained overnight for dense samples such as seeds. Typically, all samples were left overnight to ensure complete and homogeneous resin penetration. Samples were then transferred to gelatin capsules #2 (70103, Electron Microscopy Sciences, Hatfield, PA, USA) filled with pure resin and polymerized at 55 °C for 48 h.

For light and electron microscopy, thin and ultrathin sections, respectively, were obtained using a PowerTome PC RMC (Boeckeler Instruments Inc., Tucson, AZ, USA). Samples measuring 3 × 4 mm can be difficult to handle; alternatively, slightly larger samples (≈5 × 6 mm) can be collected, and the excess trimmed with a scalpel or microscissors prior to encapsulation. It is important that the sample does not exceed these dimensions so that it can be properly positioned within the capsule. Additional details on embedding, sectioning, and staining are shown in Supplementary Video.

Thin sections were transferred to slides using a Perfect Loop, affixed on a hotplate at 70 °C for 2 min, and subjected to the different procedures described below. Micrographs were acquired using an Axio Imager M2 microscope (Carl Zeiss Microscopy, LLC, White Plains, NY, USA) equipped with a Canon T5i camera (Canon, Ota-ku, TYO, Japan). White balance was adjusted using the unstained resin background as a reference, and any subsequent contrast or color saturation adjustments were applied uniformly to all images prior to export in JPG format.

Ultrathin sections were placed on Formvar–carbon support film, 150 copper mesh (#FCF150-Cu-50, Electron Microscopy Sciences, Hatfield, PA, USA), and counterstained with 2% uranyl acetate for 10 min, followed by 2% lead citrate for 4 min. Samples were examined with a JEM-200 CX transmission electron microscope at 100 kV (JEOL Ltd., Akishima, TYO, Japan) equipped with a digital camera (SIA, Duluth, GA, USA).

Paraffin embedding was performed using standard methods with some modifications [14,52]. An A. cruentus leaf sample was fixed and dehydrated in ethanol as described above, except with two absolute ethanol steps. The sample was infiltrated in absolute ethanol:xylene (1:1) for 1 h, xylene for 1 h, xylene:paraffin (Tissue-Tek VIP 4005, Sakura Finetek, Torrance, CA, USA) (1:1) for 1 h at 60 °C, paraffin 1 for 1 h at 60 °C, and finally paraffin 2 for 1 h at 60 °C. Sections of 10 µm thickness (CUT 6062, SLEE, Nieder-Olm, Germany) were obtained, placed in a water bath at 32 °C, collected on slides, and affixed on a hotplate at 60 °C for 2 min. Sections were then stained and micrographs were acquired.

5.3. Glass Slide Adhesive Treatment

We developed a glass slide treatment for microtechnique based on previously reported cleaning and activation methods for silicon wafers and glass slides [19,20,21]. Glass slide treatment consisted of three steps: washing, chemical activation, and silanization (gluing). Slides (2947-25X75, Corning Incorporated, Corning, NY, USA) were first washed by soaking them in 2% soap (Hyclin Plus Neutral, Hycel, Guadalajara, JAL, Mexico) for 30 min, after which both sides were scrubbed with 15% soap using a streak-free fiber pad (3068257, Scotch-Brite, 3M, St. Paul, MN, USA). They were then rinsed with tap water followed by Milli-Q water.

Six cleaning/activation solutions were tested: MeOH:HCl 1:1 (MeOH–HCl), sulfochromic mixture (K_2_Cr_2_O_7_ ≈ 4% in concentrated H_2_SO_4_, forming H_2_CrO_4_), piranha solution (30 mL concentrated H_2_O_2_ in 70 mL concentrated H_2_SO_4_), glacial acetic acid (CH_3_COOH), concentrated sulfuric acid (H_2_SO_4_), and 1 M sodium hydroxide (NaOH). The sulfochromic mixture was prepared by forming a paste of K_2_Cr_2_O_7_ with a small amount of water, followed by slow addition of sulfuric acid. All solutions are hazardous; piranha solution and sulfochromic mixture are strong oxidizers that can react violently with organic matter, and the sulfochromic mixture is also highly toxic and must be handled and disposed of according to safety regulations.

Slides were immersed in staining jars containing each solution for 30 min at room temperature. A control group received no activation. After treatment, slides were rinsed three times with Milli-Q water, blotted on paper towels, and dried in an oven at 60 °C for 30 min.

For silanization, dried slides were immersed in 4% (3-aminopropyl)triethoxysilane (AES, #440140, Sigma-Aldrich, St. Louis, MO, USA) in absolute ethanol for 1 h at room temperature. They were then drained and incubated in 1 mM acetic acid in absolute ethanol for 10 min, rinsed with Milli-Q water, and dried again at 60 °C for 1 h. After the final rinse and drying, slides were stored in dust-free containers to avoid microscopy artifacts.

Water contact angle was measured by placing each slide on a flat, level surface positioned between a backlit flash (Neewer S-400N, Shenzhen, GD, China) and a camera (Canon T5i with EF 100 mm f/2.8L Macro IS USM lens, Canon, Ota-ku, TYO, Japan). A 1 µL Milli-Q water droplet was applied to the slide, allowed to stand for 30 s, and photographed. Contact angles were quantified in Adobe Photoshop CS6 using the ruler tool.

To evaluate slide adhesiveness under harsh conditions, thin (500 nm) cross-sections of A. cruentus leaf were affixed to the slides at 70 °C for 2 min and subjected to three aggressive washing procedures. The first consisted of proteinase K digestion (10 µg/mL in PBS), followed by denaturation at 95 °C for 2 min and a 10 s rinse with a wash bottle. The second involved heating in water at 95 °C for 15 min followed by a 10 s water-jet rinse. The third involved heating in methanol at 95 °C for 3 min followed by a 10 s water-jet rinse. Sections were stained with 0.5% safranin (in water) and imaged. During incubations, slides were covered with microtube lids to prevent drying (Supplementary Video).

5.4. Trichrome Staining for Resin Sections

We performed experimental assays to evaluate the performance of commonly used dyes and Johansen’s staining on LR White resin section. Paradermal sections were stained using different methods. Toluidine Blue O (70103, Ted Pella, Inc., Redding, CA, USA) was applied for 5 min and rinsed with water. Johansen’s method was performed as described in Supplementary Figure S2. Coomassie Blue (0.025% Coomassie Brilliant Blue in 40% MeOH and 10% acetic acid) was applied for 5 min and rinsed with water. Safranin (1% safranin O in 40% MeOH and 10% acetic acid) was applied for 10 s, rinsed with water, followed by Coomassie Blue for 10 s and a water rinse. Fast Green (1% fast green FCF in 40% MeOH and 10% acetic acid) was applied for 5 min, rinsed with water, followed by Coomassie Blue for 2 min 30 s and a rinse with water. Finally, Fast Green was applied for 5 min, rinsed with water, then safranin for 5 s and rinsed with water. After the final rinse, slides were dried on a hotplate at 70 °C for 2 min and mounted with acrylic resin (Entellan, Merck) and a coverslip.

Because Johansen’s staining was ineffective on resin sections, we developed a novel trichrome staining method by reformulating the safranin and fast green dyes and incorporating iodine (Supplementary Figure S2). Sections shown in Supplementary Figures S4–S6 were stained as indicated in Figure 3b, with modifications to staining steps, order, and times as described in each figure. Sections in the remaining figures (except Figure 1 and Figure 2) were stained following Figure 3b without modification.

5.5. Immunohistochemical Staining

Trichrome staining reveals tissue structures and their general composition based on the resulting color patterns. However, localization of specific proteins requires immunostaining. Immunostaining was performed using standard methods [14,53]. Resin sections underwent antigen retrieval with 100 µL of proteinase K at 50 µg/mL in PBS at 37 °C for 30 min. Sections were rinsed with TBST (20 mM Tris, 150 mM NaCl, 1% Tween 20, pH 7.6). Blocking was performed using 200 µL of 1% BSA in TBST (blocking solution) at 37 °C for 30 min. After removing the blocking solution, 200 µL of rabbit anti-RbcL (RuBisCO large subunit) antibody (Agrisera antibodies, Vännäs, SE-AC, Sweden) diluted 1:500 in blocking solution was added. Sections were incubated at 37 °C for 1 h, and then rinsed twice with TBST for 5 min each.

Alkaline phosphatase–conjugated goat anti-rabbit antibody (Merk-Millipore, Burlington, MA, USA), 200 µL diluted 1:500 in blocking solution, was added and incubated at 37 °C for 30 min. Sections were rinsed twice with TBST, and color development was carried out with freshly prepared BCIP/NBT solution at 37 °C for 2 h. BCIP/NBT solution consisted of 0.02% BCIP and 0.03% NBT in alkaline phosphatase buffer (100 mM Tris, 100 mM NaCl, 5 mM MgCl_2_, 0.05% Tween 20, pH 9.5). Finally, sections were rinsed twice with TBST, twice with Milli-Q water, and dried on a hotplate at 70 °C for 2 min.

A negative control was performed without the anti-RbcL antibody, and a positive sample was stained using the trichrome staining. The incubation steps were carried out by covering the section with a microtube lid, as shown in Supplementary Video.

5.6. Digital Analysis Workflow

We developed a three-step image-analysis workflow based on MATLAB (R2024a) and Adobe Photoshop (CS6). First, we sampled the colors produced by trichrome staining in distinct tissue structures to define their corresponding ranges in the HSB color space. Second, these color ranges were used to segment cellular structures in MATLAB, and the resulting segmented layers were exported. Finally, the exported layers were used in Adobe Photoshop for quantitative analysis of tissue structure and composition. The complete workflow required approximately one day per micrograph.

5.6.1. Histolomic Analysis—Color Measurement

Different areas (n = 30) of the micrographs were measured using the Adobe Photoshop CS6 “eyedropper” tool with a sample size of 3 × 3 pixels, and statistical analyses were performed with GraphPad Prism 9.

5.6.2. Histolomic Analysis—Semi-Automatic Image Segmentation with Matlab R2024a

Micrographs were segmented using the “Color Thresholder” tool in the HSV color space. Segmented images were exported in black and white. To ensure accurate adjustments, HSV values were monitored in the segmentation function and sliders were tuned until the desired values were obtained. The final functions are in the Supplementary Data S1. Images were exported at a resolution close to the original micrograph (see Supplementary Video).

5.6.3. Histolomic Analysis—Manual Image Segmentation and Morphometric and Compositional Analysis with Adobe Photoshop CS6

Morphometric analysis: A micrograph layer with a layer mask was generated for each element to be measured. The mask density was adjusted to 50%, and each element was manually contoured using the Brush tool. The workspace was calibrated using the “Set Measurement Scale” tool and the micrograph’s scale bar. Selections of each element were retrieved through the layer masks and measured using the “Record Measurements” function. Data were exported and analyzed in GraphPad Prism 9.

Compositional analysis was carried out by densitometric quantification of lignin, proteins, and immunolabel. The micrograph was converted to grayscale/negative, and a baseline adjustment was applied. The black-and-white segmented image was resized to match the micrograph and used as a mask. Image analysis was then performed on the masked grayscale/negative micrograph, and the resulting data were saved for statistical analysis in GraphPad Prism 9.

A detailed tutorial on image segmentation and analysis using Matlab and Adobe Photoshop CS6 is presented in the second part of Supplementary Video.