To prepare thin sections, the collected wood samples must be cleaned of resin and other extractants, and then split into pieces small enough to accommodate the subsequent processing steps. Before splitting, we recommend measuring the tree-ring widths for easier subsequent dating verification of the anatomical cross-sections. Proper preparation involves three key aspects: ensuring that the pieces are of a size suitable for handling and compatible with the rotary microtome, properly aligning the wood samples to maintain consistent orientation throughout the process, and clearly labeling each piece to facilitate easy identification and re-orientation during subsequent steps. This ordered approach accounts for the limitation of maximum sample dimensions and re-orientation constraints associated with using a rotary microtome.

When collecting new wood material, it’s crucial to prioritize intact and properly oriented samples ( Figure 3 ), ensuring that micro-sections can be taken perpendicular to the orientation of the conduits.

For obtaining cores, use well-sharpened increment borers (e.g., 3-Thread Increment Borers Haglöf, Sweden), preferably powered by a cordless drill (Metabo Cordless drill BS 18 LTX BL Q I with a PowerX3 torque setting adapter, Metabo, Germany) for constant rotation to reduce micro-cracks (Gartner et al., 2023). Opt for larger cores (8-12 mm in diameter) to increase the chances of obtaining untwisted and intact wood cores. Wrapping the cores in paper or inserting them in paper straws avoids molding, reduces bending, and allows the wood to shrink without cracks during drying. Avoid gluing cores onto wood holders, as glue entering the cell conduits will obstruct paraffin infiltration. For 5 mm cores, consider collecting an additional core in the same radial direction for tree-ring width measurements. Separating the tree-ring width measurement process will simplify the preparation of the anatomical samples.

When dealing with stem sections, e.g., from subfossil wood, prioritize selecting samples (cookies or wedges) with well-oriented cutting planes and of manageable sizes (i.e., about 5-10 cm thickness). Sufficiently thick stem sections facilitate the preparation of an adequate radial strip which should not be thicker than 1 cm in tangential width and longitudinal height to fit into the mold. For fragile wood, prevent uncontrolled splitting by gluing the surface of the wood disk on a thick paper sheet or cardboard using silicone glue (e.g., Bau-Silikon transparent, OBI, Switzerland) that will not infiltrate the cell conduits due to its viscosity. Ensure the radial strip is sawn perfectly perpendicular to the conduit’s longitudinal axis, preferably using a band saw (e.g., Hammer Bandsäge N2-35, Felder group, Switzerland) with 0.6 mm deep fine blades (e.g., FLEX-BACK-Sägeblatt Felder group, Switzerland) to maintain a clean-cut surface with unfilled lumina and reduce damage to the cell walls. Keep an adjacent radial strip for tree-ring width measurements and dating. Additional orientation adjustments can be made by sanding the sample if needed.

If not done previously, perform the tree-ring width measurements required for cross-dating. Preferably, perform these measurements on the additional adjacent sample collected earlier using traditional measuring systems, such as a microscope connected with a moving table, or using novel imaging-based approaches (e.g., https://youtu.be/8q78USQksXY) together with software such as CooRecorder (Maxwell and Larsson, 2021). However, if there is not enough wood available and tree-ring widths must be measured on the same core used for wood anatomy, measurements must be conducted without mutual interference. Therefore, tree-ring width measurements should be conducted on cut surfaces rather than sanded ones to facilitate infiltration through unobstructed cell lumina. This is best done on species with clearly distinguishable ring borders to avoid filling the conduits with chalk, and on untwisted cores where the fiber direction remains constant along the core. For both large (>8 mm) and standard (5 mm) cores, both the upper and lower cross-sectional surfaces can be cut using the WSL core microtome (Gartner and Nievergelt, 2010; Gartner et al., 2015) with disposable NT blades (Light-Duty A type NT BA-170) to facilitate paraffin infiltration. Ensure that no more than 1/3 of the core diameter is removed on either side, but enough to obtain a perfectly cross-sectional surface of at least ~2 mm tangential width with open conduits, and to keep the sample well-anchored within the paraffin block.

Once the wood cores or the radial strips have been prepared, they still need to be cleaned from resins and other alcohol-soluble extractants, which might interfere with paraffin infiltration and the sectioning. The removal of extractants is achieved through a 24-hour Soxhlet extraction with 96% ethanol using a Soxhlet apparatus (standard 500 ml Soxhlet extractor with dimroth condenser, and 1000 ml round flask, e.g., Carl Roth GmbH + Co, Germany; with an electric heater, ELET36002-18, VWR International). It is important to note that ink-based labels on the sample might be washed out or hidden because the samples (especially if subfossil) may turn dark during the process. To avoid this, labels should be marked with a water-resistant soft pencil (e.g., Stabilo aquarellable All 8008, Galaxus), and a picture of the samples before the insertion in the Soxhlet extractor additionally helps label reassignment if needed. Silicon-glued paper can be kept on fragile samples since it does not interfere with the extraction process and the infiltration of paraffin in the tissue processor (see next steps).

A clear and consistent sample labeling strategy is crucial for effective wood processing and subsequent data analysis. Since wood samples are often split into sub-pieces for lab processing and imaging, and potentially further divided into sub-images depending on the analysis system used, it is essential to adopt a labeling system that can track the order and identity of sub-pieces and images throughout the entire workflow, from sample preparation to data analysis.

At the WSL laboratory, a standardized labeling system has been developed to facilitate tracking samples and their fragments during processing. This system ensures that the sample’s origin can be traced, even as the material is divided and imaged in successive stages. The labeling structure includes key elements separated by an underscore to simplify data handling: siteID_speciesID_tree/sampleID_sectionNr_imageNr (with siteID and speciesID being optional, depending on the study design). For example, the label “YAM_LASI_224_01_1” refers to the first image of the first thin-section closest to the bark from tree Nr. 224, a Larix sibirica from Yamal in northern Siberia. SectionNr and imageNr are added incrementally as the sample is processed, maintaining chronological order from bark to pith ( Figure 4 ).

For both cores and radial strips, the length of the split sub-pieces should preferably be around 3 cm long to allow them to fit into the embedding cassettes (Simport Histocette M490 and Simport Macrocette M512, VWR International) and ensure enough margins for correct positioning within the mold for paraffin embedding (37 x 24 x 05 mm for 5 mm cores and 37 x 24 x 09 mm for big/deep sample, e.g., embedding dish, stainless steel, Biosystems, Nussloch, Germany). To ensure that each annual ring is present from beginning to end in at least one of the adjacent pieces it is essential to split the sample diagonally across at least two ring boundaries with a sharp cutter knife (e.g., NT Cutter A-300 RP) and a wooden mallet (e.g., LUX Holzhammer Comfort 70 mm, OBI, Switzerland). Additionally, marking the fiber direction on the tangential side of each sub-sample will ensure a reference for a correct orientation within the paraffin block. For cores with surfaces prepared on both the lower and upper sides, mark the side intended for sectioning.

Paraffin infiltration and embedding stabilize the samples during sectioning by filling all hollow spaces, ensuring that the physical dimensions remain intact throughout processing. To achieve complete infiltration, we first submerge the samples in water and place them in a vacuum chamber connected to a pump (e.g., 2 Gal (8L) Vacuum Chamber Kit, Shenzhen Haocheng Instrument Co., Ltd, China). We run 4 to 5 cycles of applying a vacuum in the chamber until the water starts to boil and then release the vacuum. To infiltrate woody tissues with molten paraffin (Paraplast™, melting point 56°C, Biosystems, Nussloch, Germany) we use a tissue processor (Leica TP 1020, Biosystems, Nussloch, Germany) ( Figure 5 ). During this stepwise process, water is first removed from the tissue by increasing the purity of ethanol and subsequently, the ethanol is removed from the tissue using UltraClear (J.T Baker UltraClear™), a substitute and more environmentally friendly and less harmfull for humans clearing agent than the commonly used Xylene. Paraffin infiltration occurs as the samples are immersed in a bath of liquid paraffin, with a vacuum pump employed to enhance the rate of paraffin penetration into the tissues. Shaking additionally improves paraffin infiltration into the tissue. An example of the detailed infiltration program, including the duration of each step, is presented in Figure 5 .

After tissue infiltration, the sample is embedded within a paraffin block to facilitate fixing it in the microtome holder and to further stabilize it during sectioning. This is achieved with an embedding station (e.g., Leica HistoCore Arcadia H, Biosystems, Nussloch, Germany), where the still-warm wax-infiltrated tissues are placed in the mold of previously labeled cassettes, which are then filled with warm molten paraffin. As the surrounding paraffin cools, it solidifies and provides a hard, stable, and solid block to support sectioning. To prevent the infiltrated samples from cooling faster than the surrounding paraffin and causing discontinuities (cracks) between the sample and the paraffin block, this process should be performed relatively quickly. If cracks occur, the sample embedding in the paraffin block should be repeated. Proper sample orientation in the mold is crucial, which is facilitated by the previous marking of fiber direction. If wood surface cutting was done perpendicular to the fiber direction, this will result in a parallel alignment of the transverse plane of the wood sample with the block surface for sectioning. Since the surface at the bottom of the mold will become the cutting plane, the sample plane selected for thin sectioning should be positioned facing the bottom of the mold. For silicon-paper-glued subfossil material, this means that the sample is positioned with the paper side facing up. Moreover, each specimen should be positioned so that the blade strikes it diagonally rather than parallel to the tree rings. This angle ensures that both the dense latewood and the less dense earlywood are cut simultaneously. This is usually achieved by placing the sub-sample obliquely within the mold. For the positioning press the sub-sample to the bottom of the mold with straight pointed tweezers (e.g., Tweezers straight pointed, 130 mm, Carl Roth GmbH + Co, Germany) until the paraffin starts to solidify. Large cassettes of 10 mm height, as used for the infiltration of large samples, should be replaced by equally labeled small cassettes (5 mm high) without lids to match the clamp size of the microtome and better stabilize the sample. To accelerate the solidification of the paraffin block, the embedded samples can be placed on a cooling plate (e.g., Leica HistoCore Arcadia H, Biosystems, Nussloch, Germany) for 20 minutes and/or stored in a regular refrigerator at ~ 4-6°C for 30 minutes until they can easily be extracted from the mold. Use abrupt and forceful pull to separate the paraffin block from the cassette.

After embedding, excess paraffin surrounding the solidified block on the sides of the cassette should be removed with a disposable blade (e.g., Light-Duty A type NT BA-170) to ensure a stable grip within the microtome clamp (universal cassette clamp 14 0502 37999, Biosystems, Nussloch, Germany).

To prepare for sectioning ( Figure 6 ), the wood surface of the sample needs to be exposed by removing the top layer of paraffin and wood. This process called trimming, precutting, or facing, is performed with the rotary microtome (HistoCore AUTOCUT, 14051956472, Biosystems, Nussloch, Germany) using either low- (FEATHER^® N35, Biosystems, Nussloch, Germany) or high-profile (Micros HP, Biosystems, Nussloch, Germany) disposable blades, with a cutting step of 5-7 µm to avoid damage to the dry and hard samples until the desired sample surface is entirely exposed. The primary difference between high-profile and low-profile disposable blades lies in their geometry and intended applications. High-profile blades are larger in height and have a steeper cutting angle, providing greater visibility and clearance, which is beneficial for broader or more aggressive cutting tasks. In contrast, low-profile blades have a flatter design, allowing for closer contact with the sample surface and offering finer precision, making them ideal for precision sectioning. The blade angle directly affects cutting efficiency and sample quality, with high-profile blades excelling in robust tasks and low-profile blades offering better control and stability for obtaining thin, high-quality sections for anatomical studies.

For trimming, set a blade clearing angle of 5°. The blade can be used as long as no artifacts such as stripes from notches in the blade are appearing. Standard 5 mm cores with unprepared wood surfaces before infiltration and embedding should be pre-cut to a depth not exceeding 1/3 of their diameter. This ensures that at least 2/3 of the core remains in the block, thus avoiding tissue that is not properly infiltrated by paraffin. To remove accumulated cutting waste from the blade holder, use a brush (Pelican Hair brushes no. 23, size 2-3) with bottom-up movements, i.e. away from the blade edge, to avoid trimming the brush hairs. To facilitate the cleaning of the microtome and the working place, place a moist sheet of lab paper at the bottom front of the microtome to easily remove the waste generated at the end of the trimming process.

Before thin sectioning, it is recommended to immerse the samples in water and store them at 4-6 °C in the refrigerator for several hours (preferably overnight) to harden the paraffin and soften the wood. These conditions are necessary to ease sectioning and prevent the paraffin from sticking to the microtome blade holder or brush, thereby enabling the creation of a cohesive “ribbon” of subsequent thin sections.

For sectioning ( Figure 7 ), low-profile N35 disposable microtome blades (FEATHER^® N35, Biosystems, Nussloch, Germany) are typically used. In some cases as for denser woods, more stable high-profile blades may be preferred (Micros HP, Biosystems, Nussloch, Germany). The typical blade wear is 10 samples. Set the blade clearing angle at 5°. To maximize the blade usable duration, move the blade systematically, working from one end to the other. The ideal section thickness for achieving optimal staining intensity and contrast in QWA measurements (e.g., using ROXAS) is 12 µm. Thicker sections (i.e., 14-16 µm) are not recommended due to their greater weight, which increases the likelihood (up to 70%) of losing the section during the subsequent staining process. While sectioning, ribbons of generated sections are produced and collected. If the ribbon does not form well, ensure that the block is still sufficiently cold. Cracks in the ribbon may indicate damaged or used blades. Even slight contact with metallic items, such as the tweezer tips or the ferrule (metal band) of the brush, can damage the blade surface. Hard impurities in the sample can also cause these cracks. Moving or changing the blade, as well as flipping the sample in the microtome clump, may resolve any issues.

To collect the selected sections from the ribbon (from 1 to 3 sections), use pointed-tipped dissection needle (e.g., Dissecting needle straight, Carl Roth GmbH + Co, Germany) to first transfer the selected ribbon floating on top of a 38°C water bath (VWR^® W20, Histology Water Bath, VWR International) with the smooth (shiny) side down. The warm water helps the floating ribbon (floatation) to stretch and ensure it is completely flat. Examine each section as it floats and inspect for imperfections when selecting the target section. The target section can then be caught and positioned on a pre-labeled and thinly albumin-coated microscope slide (SuperFrost^® slides with 90° ground edges, Biosystems, Nussloch, Germany) by holding it with a brush while gently lifting the slide from below the floating section. Therefore, one drop of Albumin (Protein glycerol, P049.1, Carl Roth GmbH + Co, Germany) is applied to the slide surface with a gentle single wiping movement to obtain a homogeneous thin layer (without gaps or excess), which keeps the thin section sticking to the slide during the staining process. The slides should also be labeled accordingly with a pencil or a chemical-resistant printed sticker (Labid-technologies N0FTT-149C1-2WH or Biosystems 85-1086-00, Nussloch, Germany; printed with the Zebra ZD621 label printer using the Zebra ZD621 software, Zebra online store). If the slide scanner supports it, consider using QR codes for labeling. Don’t use solvable ink to prevent the labels from fading or dissolving during the staining process. Additionally, as the warmed paraffin has become sticky, the brush should only be very gently held on the sample during the collection of the target section to prevent damaging the thin section. Once the target thin section adheres to the slide, drain them vertically for a brief time to remove excess water and place the slide on the slide drier surface of the water bath until all water has evaporated. The slides can then be gathered in slide racks (glass rack for 10 slides with metal handle and staining jar, VWR International). Slides with thin sections can be stored in slide racks or slide boxes (Slide Boxes with Lid, VWR International) until staining. The samples in the paraffin blocks can be archived in an embedding carton box (Diastore Block 12, Histocom, Switzerland) and thus remain available in case the procedure needs to be repeated.

To complete the process, the thin sections must be deparaffinized and stained to reveal the underlying tissue structures and facilitate quantitative image analysis ( Figure 8 ). Additionally, they must be permanently fixed for secure handling, such as imaging and long-term storage. To melt and remove the paraffin from the prepared slides, place the thin sections in slide-racks and put them in an oven (BINDER FD 115 drying and heating chambers with forced convection, Faust, Switzerland) at 60°C for at least 2 hours before staining but no longer than 24 hours.

The staining process is performed under a fume hood with slide-racks in staining dishes using a mixture of 0.8% Safranin and 0.5% Astrablue [according to (Gärtner and Schweingruber, 2013)] to stain the lignin red and the cellulose blue. The process consists of subsequent baths to dissolve and remove the remaining paraffin with Xylene, to remove the Xylene with Ethanol, and to stain the thin sections. After staining, the samples are rinsed with water, dehydrated with Ethanol, and then prepared for fixation with Euparal (Euparal 7356.1, Carl Roth GmbH + Co, Germany). A staining protocol with relative bath durations is provided in Figure 8 . The process can be performed in sequence with multiple slide-racks. The bath durations have been optimized according to the thickness of the thin section to guarantee sufficient contrast for further image analysis.

Sample fixation occurs immediately after staining. To fix the thin sections in Euparal, carefully dry the thin section after dehydration in 96% ethanol by gently patting the slide with a paper towel. Then, apply Euparal on top of the thin section using a pipette (Pasteur pipettes without cotton plug with bulb, Carl Roth GmbH + Co, Germany). For large thin sections covered by a 50 mm long cover glass (Cover Slips, VWR International), use two large drops. To reduce air bubbles, the applied Euparal drops should be allowed to flow down the inclined thin section and cover it completely before placing the cover glass. Fixation is achieved by placing the slide between a magnet (e.g., Ferrite magnets Y35 20mm diameter 5mm height, Supermagnete) and an iron plate (e.g., Ferrous metal plate, DIY/Home improvement store). Additionally, one could use a layer of polyethylene (e.g., LDPE Tubular film 200mm diameter 100µm thickness, Elke-Plastic, Switzerland) to protect the iron plate and magnets from the excessive fixing agents. Leave the slides in a laboratory fume hood for one to two days before removing the magnets. The process can be accelerated by putting the magnet-covered slides for 2-4 hours in a regular oven at 60°C. After drying, and before imaging, clean the excessive fixing agent with 96% ethanol to remove any potential dirt obstructing the imaging. If large quantities of fixing agents cover the cover glass, first use razor blades (Single Edge Stainless Steel Teflon Coated Blades, Biosystems, Switzerland) to carefully remove it, ensuring that the generated dust is also removed. If this repeatedly occurs, reduce the fixing agent applied. Since the staining and fixing process for an entire batch takes several hours, it is advisable to schedule this procedure on a day separate from sample sectioning.

The fixed, labeled, and cleaned slides are prepared for imaging using a high-volume automated slide scanner (Zeiss Axio Scan.Z1 slide scanner, Carl Zeiss, Germany) ( Figure 9 ). This advanced equipment efficiently acquires high-quality images of entire thin sections under consistent and controlled conditions. Specifically, bright-field imaging is employed with a 10x objective lens, maintaining constant illumination for a high resolution of 2.2675 pixels per micron. After producing a 3-dimensional map of the section surface to account for unavoidable waviness using autofocus, the entire section is digitized with a z-stack around the surface map and stitched to a flat image, thus ensuring sharp image quality. Slide IDs can be read directly by the system from the QR code on the label, provided by a text file list, or manually adjusted after scanning. For alternative imaging methods using common optical microscopes, including guidelines for image capture, focusing, illumination, and stitching, we refer readers to established protocols presented in von Arx et al., (2016). This reference details step-by-step recommendations to ensure high-quality imaging, even in the absence of slide scanners, as well as software tools for stitching images when scanning entire sections.

To ensure compatibility with ROXAS software for cell anatomical measurement, the collected images of thin sections undergo a specific processing procedure. A custom macro is used to place overlapping cropping rectangles on the whole-section image, ensuring each tree ring in the overlapping regions is fully represented in only one rectangle. This strategic placement selects the best-quality parts of the sections, avoiding issues such as cracks and disrupted tissue. The cropping rectangles adhere to defined dimensions, with a maximum length of 32000 pixels along the radial tree stem axis and approximately 5000 pixels in width (around 2.2 mm of wood) along the tangential axis. JPG images are then batch-exported as defined by the cropping rectangles, with the pith side oriented at the top. To maintain a clear hierarchical organization, the newly exported JPG images are labeled by appending an incremental number (ImageID) to the thin section label (cf. step 2). This numbering system progresses sequentially from the bark to the pith, facilitating subsequent analysis.