Assessing Structural Traits in Triticum aestivum and Zea mays for C₃ and C₄ Photosynthetic Differentiation Using Free-hand and Semi-thin Sections

Photosynthesis is a fundamental process where light energy is converted into chemical energy, serving as the cornerstone of terrestrial trophic networks. The majority of plants follow the C₃ pathway of photosynthesis, where the primary photosynthetic product is the three-carbon compound glycerate 3-phosphate. C₃ photosynthesis evolved over 2 billion years ago in the atmosphere abundant in CO₂ and low in O₂¹. The key photosynthetic enzyme ribulose 1,5 bisphosphate carboxylase/oxygenase (Rubisco), which evolved under these conditions, is suboptimal for current low CO₂ high O₂ conditions as it competitively reacts with O₂, initiating photorespiration². Photorespiration is a wasteful pathway that consumes energy instead of producing it and releasing CO₂ as a byproduct. Consequently, it is crucial to maintain high CO₂ concentration around Rubisco in chloroplasts to prevent oxygenation^3,4. Due to the inability of C₃ plants to concentrate CO₂, there is a significant CO₂ drawdown from ambient air to chloroplasts, curbing photosynthesis and affecting plant growth and biomass production^2,5,6.

In C₃ plants, photosynthesis is limited by the entrance of CO₂ through stomata, its diffusion through the mesophyll, and the biochemical activity of photosynthetic enzymes. The entry of CO₂ is first limited by stomatal conductance, which is controlled by environmental conditions such as air temperature and humidity. Then CO₂ diffuses through the leaf's internal gaseous and liquid phase to Rubisco⁷. In C₃ plants, all stages of photosynthesis occur in the chloroplasts of mesophyll cells, and plants need to maintain a constant influx of CO₂ from the atmosphere into chloroplasts. The dependence of CO₂ availability in chloroplasts on stomatal openness, mesophyll architecture, and individual cell and chloroplast characteristics leaves plants susceptible to environmental constraints that eventually affect photosynthesis, like low water availability and high temperatures^7,8,9,10, particularly highlighting their vulnerability to climate change conditions¹¹.

Given the challenges posed by the inefficiencies of the C₃ pathway, as well as limitations in maintaining optimal CO₂ levels and susceptibility to environmental factors, in certain plants, another pathway, the C₄ photosynthesis pathway, has evolved. Characteristically, C₄ plants have two spatially separated biochemical pathways; the initial CO₂ fixation occurs in mesophyll cells by phosphoenolpyruvate carboxylase, which has a higher affinity for CO₂ than Rubisco and also lacks oxygenation activity. The formed C₄ product is further transported to bundle sheath cells, where it is decarboxylated, and CO₂ is again released and fixed by Rubisco (C₃ photosynthesis)^12,13,14. The greater affinity of PEP carboxylase to CO₂ and thick cell walls of bundle sheath cells allows CO₂ concentration in bundle sheath cells, and thus, C₄ plants minimize the photorespiration by spatially segregating CO₂ fixation and the Calvin cycle. The adoption of the C₄ pathway showcases nature's adaptive response to environmental constraints, offering insights into potential strategies for improving crop productivity and resilience in changing climate conditions¹⁵.

The specialized anatomy of the leaf structure in C₄ plants is characterized by veins surrounded by enlarged vascular bundle sheath cells containing chloroplasts and with a radial arrangement of mesophyll cells in an outer ring patterning around bundle sheath cells. The mesophyll cells are in close proximity to the bundle sheath cells, which enables a rapid and continuous transport of metabolites between the two cell types. This cell's arrangement is typical of C₄ plants and is referred to as Kranz anatomy¹⁶. In C₃ species, mesophyll cell specialization and disposition can vary, but bundle sheath cells are distinctly smaller and have a few chloroplasts or no chloroplasts at all. Specific Kranz anatomy allows concentrating CO₂ in chloroplasts in bundle sheath cells where the C₃-carboxylating enzyme Rubisco is located, effectively hindering photorespiration^{4, 17, 18}. Despite its seemingly complex arrangement, these changes have occurred independently multiple times in the evolution of angiosperms, indicating that it is a relatively feasible evolutionary pathway^19,20,21, and various taxa have been shown to be at an intermediate stage between C₃ and C₄ carbon metabolism, referred to as C₃-C₄ or C₂, having abilities to concentrate and re-assimilate CO₂^22,23,24,25.

Many C₄ plants are crops with high economic importance, and genetically engineering C₃ crops, like rice, to improve their climate resilience and secure the yield has been a topic of interest in the last decades^26,27. However, the engineering efforts call for a detailed understanding of C₄ specialized anatomy and how it controls photosynthesis^2,28.

Establishing the C₄ Kranz anatomy is a prerequisite to achieving the ambitious goal of engineering C₄ photosynthesis into C₃ crops²⁵. However, the current understanding of the regulation of Kranz anatomy and methods for quickly screening its key anatomical traits is limited, making it difficult to identify hybrid species. Previous studies have shown that key traits regulating photosynthetic efficiency in C₃ and C₄ plants include the interveinal distance, the diameter of the bundle sheath complex, and the size of bundle sheath cells^14,29. These traits can be easily screened using free-hand sections and quantitatively analyzed using semi-thin sections. Here, we describe the method of assessing the traits that allow for C₃ and C₄ anatomical differentiation diagnostics through free-hand cross and light microscopy, namely bundle sheath area, interveinal distance, and vein frequency.

1. Plant growth conditions

1. Select wheat (Triticum aestivum L. Var. Winter Wheat, "Fredis") and maize (Zea mays L. Var. Saccharata, "Golden Bantam") as representative C₃ and C₄ plants, respectively. Grow both plant species in an environment-controlled growth chamber from seeds.
2. Maintain relative humidity at 55% with the air temperature at 25 °C/18 °C (day/night) with a 12 h light period. Maintain photosynthetic photon flux density (Q) at plant surface at 1200 µmol/m²/s to represent their natural growing conditions. Grow all plants at an ambient CO₂ concentration of 400 µmol/mol.
3. Water plants every 2^nd day with distilled water and fertilize with Hoagland's solution 10 days after seedling emergence. Grow the plants at the given conditions for 60 days and use fully expanded leaves for microscopic analyses.

2. Preparation and viewing of free-hand sections

1. Select 3 mature non-senescent leaves from each plant and keep them in distilled water. Using a new single-sided razor blade, make one leveling cut to form a right angle with the leaf surface to ensure the sections are transverse cross-sections made perpendicular to the leaf surface.
2. Place the leaf specimen on a glass plate on a piece of dental wax to stabilize the sample and avoid slipping. Cut by moving the blade straight down on the leaf specimen to cut cleanly through cells. Keep the cross-sections in distilled water.
3. Mount the samples on a glass slide by placing them on a water droplet and covering them with a glass coverslip for microscopical viewing and imaging. Avoid trapping air bubbles.
4. View the slide under a compound light microscope at 10x and 20x magnification. Save images as per the software guide, along with the relevant scale.

3. Preparation of semi-thin sections

1. Cut sections of approximately 3 mm x 5 mm in size from intercostal areas alongside the midrib from leaves from both Z. mays and T. aestivum on a glass plate as in step 2.2. Ensure that the length of the cut section runs along the grain of the leaf.
2. Place the plant material in a syringe with 1 mL of fixation buffer (4% glutaric aldehyde in 0.1 M phosphate buffer, pH = 6.9, Table 1).
3. Hold the syringe vertically, remove the air, and, holding a finger on the tip, pump the syringe to create a vacuum. Repeat until the samples lie at the bottom of the syringe, indicating successful infiltration.
4. Transfer the contents of the syringe to a labeled glass vial and store at 4 °C for 48 h before the next step (Table 2).
5. For each vial, gently remove the fixative with a pipette and add the phosphate buffer until the sample is covered. Leave for 30 min. Repeat this step 3x.
6. Remove the phosphate buffer with a pipette and add the osmium tetroxide (CAUTION) until the sample is covered. Samples and other organic matter will turn black. Use double gloves. Leave for 1 h.
7. Remove the previous solution with a pipette and cover the samples with distilled water. Leave for 30 min Repeat this step 3x and use double gloves.
8. Remove the previous solution with a pipette and cover the samples with the 50/50 distilled water-ethanol solution. Leave for 30 min. Repeat this step with distilled water-ethanol solutions in the following series: 30/70, 20/80, 10/90, and 2x at 100 % ethanol. Samples in any of these solutions can be left overnight at 4 °C.
9. Remove the previous solution with a pipette and cover the samples with 1/3 resin-ethanol solution. Put the samples on a mixer plate and leave for 2 h. Repeat this step with resin-ethanol solutions in the following series: 1/2, 1/1, 2/1, 3/1, and 2x at 100% resin. Samples in any of these solutions can be left overnight at 4 °C.
10. Cover the cavities of a flat embedding mold (16 cavities, cavity dimensions: 14 L x 4.8 W x 3 D (mm)) with 100% resin and place the samples on one end of the cavity. Prepare pencil-written paper labels for each sample and place them on the other end of the cavity.
11. Completely fill all the cavities with 100% resin and straighten the samples and labels if they moved. Cover the mold with embedding film and polymerize in an oven as per acrylic resin product guidelines.
12. Set the polymerized block with the sample in the specimen holder of the ultramicrotome. Trim the block using a rough glass knife until the tissue becomes visible and excess resin is eliminated.
13. Cut semi-thin sections transversely (1 µm) using a glass or diamond knife. Collect the sections using a metal inoculation loop from the water surface and place them on a glass slide. Dry the slide on a hot plate (60 °C) to fix the sections onto the glass.
14. Stain the fixed sections with toluidine blue (1% aqueous solution) for 5 s before rinsing with distilled water and drying again on the hot plate.
    NOTE: Ensure the chemical fixation process is performed under a fume hood. Glass knives should be replaced periodically to ensure sharp cuts that allow for sections without tears or distortions.

4. Imaging of samples

1. Imaging of fresh leaf sections
   1. Set up the compound light microscope with the accompanying software as per the manual.
   2. Place the glass slide on the stage. Adjust the eyepiece and view the section at 10x to obtain an overview, then at 20x and 40x objectives.
   3. At each objective, navigate the Live channel, focus on the area of interest, and locate the bundle sheath cells around the vein.
   4. Once focused, click the Freeze button and add the scale by pressing the Scale Bar button in the Tools tab. Save the image in .TIF format for image analysis.
2. Imaging of resin-infiltrated sections
   1. Set up the automated light microscope in connection with the accompanying software on the computer.
   2. Load the transparent channel and place the slide on the microscope stage. Focus on the section using a 40x objective magnification and adjust the brightness to ensure all cell structures are visible.
   3. Set the location area of the section using the navigator function to ensure the entire section is imaged and click the Stitch button.
   4. Click Done and then Start to allow the microscope to image the entire section. Open the image analysis software and open the tiles taken by the microscope.
   5. Using the Align feature, ensure that the tiles do not overlap. After generating the image, adjust the positioning, brightness, and rotation using the Adjust tab.
   6. Print the scale on the image before saving in .TIF format.
3. Anatomical measurements using ImageJ software
   1. Open ImageJ software and load the image by dragging it into the window.
   2. Using the Line tool, calibrate the scale as follows: Draw a line over the scale bar, choose Analyze > Set Scale, change the known distance to the length of the scale bar, and change the unit of length to the correct unit.
   3. Measure the prospective trait by selecting the Line Tool, drawing a line across the area of interest, and pressing the M key.
   4. For area measurements, select the Polygon Selection Tool and draw around the tissue of interest. Conclude by pressing the M key. The measurements show up as a pop-up window, which can be copied to a spreadsheet for further data analysis.

Figure 1A shows the correct orientation for sectioning the leaf for both fresh sectioning and light microscopy. The method for cutting fresh sections using a single-sided razor and a dental wax sheet can be seen in Figure 1B. The resulting sections are shown in Figure 1C.

Figure 2 shows free-hand sections of representative leaves of the C₃ plant T. aestivum and the C₄ plant Z. mays. Bundle sheath cells are visible around the vein, with maize having rounder and greener cells, indicating a higher presence of chloroplasts. It is possible to obtain leaf thickness and total bundle sheath area from these images; however, further characterization of the mesophyll is not possible with this method.

Figure 3 shows semi-thin cross-sections of T. aestivum and Z. mays. With this method, all tissues of the leaf are visible and measurable. Kranz's anatomy is evident in Z. mays, with mesophyll cells arranged around the veins. Bundle sheaths are enlarged and filled with chloroplasts, notably concentrated towards the mesophyll cells. This allows a rapid exchange of metabolites between bundle sheath cells and mesophyll cells. From these images, the following leaf structural traits were measured: interveinal distance between small and large veins, bundle sheath diameter, total bundle sheath area, veins area, fraction of bundle sheath cells area, bundle sheath cells area, vein frequency within 500 µm, and leaf thickness. Box plots in Figure 4 show the difference in such traits between T. aestivum and Z. mays. All but one trait, the interveinal distance between large veins, are significantly different at a 95% confidence interval with a p-value < 0.05.

Figure 4 shows a collection of box plots that represent the anatomical trait differences between T. aestivum and Z. mays, exemplifying the differences between C₃ and C₄ bundle sheath structures. There is a significant distinction between the interveinal distances between the crop species, the diameter of the bundle sheath (under the same growing conditions), the fraction of bundle sheath cells and non-photosynthetic material (i.e., veins), and vein frequency in a 500 µm section width - all of which are strongly indicative of what is expected between C₃ and C₄ leaf anatomy.

Figure 5 shows some examples of mistakes that are likely to occur when either cutting or preparing the samples. The leaf anatomy in these images is masked or deformed and is thus not reliable for characterization. Overstaining with toluidine blue can occur in sections cut with the ultramicrotome in which the stain is left for too long, thus obscuring the cell structure (Figure 5A,B). Common errors can occur during the infiltration protocol in which the resin does not fully penetrate the leaf tissues (Figure 5C,D). Leaf cross-sections that are cut along the grain result in oblong cells in which the mesophyll and bundle sheath cells cannot be located nor correctly measured (Figure 5F,G,H).

Statistical analyses were conducted in R (version 4.3.1) to create boxplots. Leaf anatomical traits were measured in triplicate (3 leaves per plant, 3 plants per species).

Figure 1: Correct method for cutting fresh leaf cross sections. (A) The orientation of leaf sections in order to obtain a cross-section. (B) Technique for sectioning leaves using a single-sided razor blade on a piece of flat dental wax. (C) The resulting cross-sections appear as thin leaf slivers that are ready for mounting on a glass slide and viewing under the microscope. Please click here to view a larger version of this figure.

Figure 2: Free-hand sections of representative leaves. (A) The C₃ plant Triticum aestivum (400x magnification); and (B) the C₄ plant Zea mays at (200x magnification). Cell type is indicated by arrows, where V = vein, BSC = bundle sheath cell, and MC = mesophyll cell. Please click here to view a larger version of this figure.

Figure 3: Semi-thin cross sections. (A) Zea mays and (B) Triticum aestivum to show the differences in bundle sheaths and chloroplast positions in C₄ species with Kranz anatomy (A) and C₃ species without Kranz anatomy (B). Abbreviations: Tl = leaf thickness, BS = bundle sheath, SV = small vein, ID = interveinal distance, and LV = large vein. Please click here to view a larger version of this figure.

Figure 4: Box-plots showing the difference in anatomical traits between Triticum aestivum and Zea mays. (A) interveinal distance between small veins, (B) interveinal distance between large veins, (C) bundle sheath diameter, (D) total bundle sheath area, (E) veins area, (F) fraction of bundle sheath cells area, (G) bundle sheath cells area, (H) vein frequency within 500 µm and (I) leaf thickness. Please click here to view a larger version of this figure.

Figure 5: Examples of common mistakes that might occur in tissue processing, indicated by red arrows. Overstaining in samples embedded in LR White acrylic resin (A) Triticum aestivum and (B) Zea mays; tears in resin in sections embedded in Spurr's epoxy resin (C) Ceratozamia robusta and (D) Dioon mejiae; T. aestivum free-hand sections showing (E) cross-section and (F) longitudinal section; bias in sectioning due to oblique cuts in (G) Annona muricata and (H) Encephalartos villosus. Please click here to view a larger version of this figure.

Table 1: Recipe for the fixative solution.

Table 2: Resin infiltration protocol.

In this article, we discuss both the quantitative and qualitative methods of measuring leaf anatomy and ways in which they can be optimized. Furthermore, the methodology is applied to representative crop species so as to determine which anatomical traits are most useful in distinguishing between C₃ and C₄ cross-sections. Understanding these traits is essential as hybrid species, termed C₂ photosynthesis, is becoming a more promising avenue of research. As of now, only one crop species, Diplotaxis tenuifolia (arugula), has been identified to use C₂ photosynthesis, but it is likely that there are more than what current records indicate²⁵. High levels of intraspecific photosynthetic diversity and plasticity are present within C₂ lineages. This implies that confirming the occurrence of C₂ photosynthesis necessitates multiple lines of evidence, such as CO₂ compensation point, ultrastructural analyses, and immunohistochemistry³⁰.

Free-hand and semi-thin sections have been used in identifying and measuring specific plant tissues for many years, but this article aims to discuss the particular use case of anatomical leaf structures that affect photosynthetic differentiation between C₃ and C₄ crop plants. There are caveats to both free-hand and semi-thin sectioning that ought to be considered. Free-hand sections, while quick to obtain, are likely to result in leaf sections with inconsistent thicknesses, hindering quantitative analyses. This can be avoided by using specialized equipment for the sectioning of fresh plant material (e.g., vibratome or cryostat). The thicker sections from hand-sectioning can result in multiple layers of cells atop each other, blurring the cells and, thus, the anatomy. If the purpose of sectioning is purely qualitative, freehand sections offer the advantage of chemical processing of plant tissues (i.e., histochemistry)³¹. This allows for the qualitative indication of the presence of particular compounds in plant cells. Conversely, semi-thin sections almost always result in consistently thick sections. However, the processes are longer, delicate, and thus prone to error. Issues that may arise, as well as ways to avoid them, are discussed further.

Fixing
Fresh material must be used for both free-hand and semi-thin sections, i.e., leaves from plants that are sufficiently hydrated. Sections from wilted leaves do not have structural integrity as the cells have a decreased turgor pressure. When obtaining semi-thin sections from resin-embedded material, it is critical to ensure that leaf sections are fixed under a vacuum so as to replace the air inside the leaf. This allows for structural integrity and osmosis during the infiltration process. If this step is not done correctly, cells are likely to collapse, leading to cross-sections that do not accurately depict the real leaf structure and subsequently incorrect anatomical measurements³².

Conversely, high vacuum pressure can damage leaves with soft tissues (like cotyledons), resulting in shrunk cells or the destruction of the samples. This can be avoided by using an appropriate vacuum pump setting or by skipping it altogether and just soaking the samples in the fixative solution. If the latter is preferred, samples can be cut at all sides, as to ease the infiltration of the fixative into the tissue.

Pre-Infiltration
Cell wall thickness and sclereid tissue are the most influential anatomical traits to the permeation of solutions that allow for optimal fixation and infiltration of the specimen. The purpose of sclereids in plant tissue is mechanical support and protection, which can impact the infiltration process³³. As these tissues have highly thickened lignified cell walls, permeation of solutions can be hindered. Samples that have infiltration issues result in semi-thin sections with broken cells and tears in the resin (Figure 5C,D). Longitudinal sectioning results in distorted cells, causing biases in anatomical analyses and measuring of leaf traits such as cell size and cell density (Figure 5F-H).

Resin Infiltration
Quality sectioning is vital for the observation of anatomical tissues, which are directly affected by the medium in which the specimen is embedded. The most common embedding media used in plant histology are paraffin wax, epoxy, and acrylic resins^34,35. Wax-embedding requires preparation of samples by chemical fixative, and infiltration with liquid wax, which are hardened at room temperature. These cuts typically result in sections that are 5-10 µm in thickness, which, as demonstrated here, can be achieved just as well by hand (Figure 2) and require just unfixed fresh tissue. In this instance, the arrangement of cells around the bundle sheath is vital to distinguishing between C₃ and C₄ plants, which requires the use of a more robust and rigid embedding media. Epoxy resin (Spurr's resin³⁶) and acrylic resin (LR White) were tested both for hardness, longevity, sample infiltration, as well as sectioning durability in this study. Epoxy resin has been used in the embedding of biological material for decades³⁷. Acrylic resins are relatively new and were initially considered for embedding plant specimens due to their resilience over time as, unlike epoxy resins, they do not discolor as a result of UV exposure and are less prone to shrinkage³⁸. LR White acrylic resin (hard grade) was selected for its non-toxicity and UV-stable formula. Acrylic resins tend to be considerably less viscous than epoxy resins, which allows for quicker sample infiltration, and the hardness of the material allows for cleaner cutting with a microtome³⁹. Figure 5C,D shows the results of epoxy resin in leaf specimens with a large amount of structural tissue, where tears are inevitable due to the softness of epoxy resin, as compared with acrylic resin (Figure 3).

Ultramicrotomy
Trimming away excess resin may create cracks in the resin that penetrate into the sample. To avoid this, it is vital to consider the density and brittleness of the resin chosen for embedding. Cutting deeper into the section should avoid any artifacts caused by hand or rough knife trimming. The sample should be aligned perpendicularly to the sample holder and validated by increasing the magnification through the eyepiece, as well as viewing a fixed and stained section. Plant cells not cut at a perfectly right angle appear as tapered cells whereby one end is larger than the other⁴⁰.

Post-staining
Toluidine blue (1% in aqueous solution) was used to stain semi-thin sections cut on the ultramicrotome. If the stain is left to dry on the section too long, it results in overstaining, causing an oversaturation of the section when imaged (Figure 5A). When the stain is not filtered well, it results in undissolved particles drying on the section, hindering the visualization of the anatomical structures (Figure 5B). This issue can be avoided by using a fine filter when making toluidine blue solution as well as limiting exposure of the section to the stain. Samples are dried on a hot plate at 60 °C prior to staining to ensure that the section bonds to the glass microscope slide, but should not be exposed for more than 10 s to toluidine blue before being rinsed and dried again⁴¹.

Considerations on anatomical measurements
The measurements shown in this paper are just an example of the many anatomical traits that can be quantitatively assessed using semi-thin sections, including a fraction of intercellular airspace, average mesophyll thickness (without vascular tissue), and mesophyll surface area exposed to intercellular air spaces per unit of leaf area⁴². However, while bundle sheath cell size is an important indicator to differentiate between the C₃ and C₄ metabolism, bundle sheath cell wall thickness is another important trait that cannot be viewed based on light microscopy alone, as it determines the degree of leakiness of bundle sheath cells, i.e., the amount of CO₂ back diffusing into mesophyll cells^42,43. High leakiness reduces the photosynthetic efficiency, as a large fraction of energy used for CO₂ fixation by PEP carboxylase is lost^44,45,46. For such ultrastructural analyses, Transmission Electron Microscopy and image analyses are required. Although only light microscopy was considered for the present study, the protocol includes the treatment with osmium tetroxide, a contrasting and fixing agent which is a cornerstone for TEM microscopy. Polymerized samples can thus still be used for TEM analysis whenever the need arises, even if not considered at the time of preparation.

Although anatomical diagnosis can easily detect Kranz-like structures, the presence of C₄ can further be diagnosed by gas-exchange measurements which can show typical C₄ physiology, i.e., low CO₂ compensation point, limited O₂ inhibition of photosynthesis and different carbon isotope composition^2,4,47.

It should also be noted that Kranz anatomy is not necessary for C₄ photosynthesis to develop, as shown by the discovery of single-cell C₄ plants^48,49,50. Anatomical analysis alone may, therefore, not be sufficient to detect C₄ physiology in less studied taxa. The coupling of anatomical characterization with physiological analysis offers powerful tools for investigating leaf traits and their structure-function relationship in both crops and wild plants.