Serial section videography (SSV): A low‐cost protocol for generating 3D reconstructions of internal plant structure

Abstract

Premise

Analyzing structural changes along the length of an organ provides insight into its development. However, traditional histological methods are limited by intensive procedures and size restrictions. Micro‐computed tomography (microCT) enables non‐destructive internal imaging along the length of an organ, but high cost, technical complexity, and limited accessibility hinder widespread application. Here, we describe serial section videography (SSV), a new low‐cost technique for generating three‐dimensional (3D) reconstructions of internal plant anatomy using serial sectioning and open‐source software.

Methods and Results

SSV was applied to four fern rhizomes with varied gross morphology and diverse vascular architectures. Specimens were sectioned using a sliding microtome or a handheld blade, and imaged using either a digital camera or smartphone setup. Images were aligned using Fiji and segmented using 3D Slicer. The SSV method enabled continuous visualization of internal stem anatomy over several centimeters and is adaptable to both laboratory and field settings.

Conclusions

This protocol offers an alternative to microCT for generating 3D anatomical reconstructions, enabling researchers to examine development and structural variation across organs with minimal equipment and software. This accessible protocol reduces technical and financial barriers and is particularly well‐suited for comparative studies of vascular tissues, advancing the study of plant anatomy and development.

Given that plants are iterative organisms with largely immobile cells, analyzing structural changes along the length of an organ offers valuable insights into developmental processes. For this reason, it has long been the goal of botanists to reconstruct the three‐dimensional (3D) structure of internal plant tissues.

However, investigating the internal structure across extended lengths of plant organs such as stems, petioles, or roots presents certain challenges. For instance, traditional histology requires lengthy infiltration (Ruzin, 1999) and is largely restricted to small tissue fragments. Historically, 3D reconstruction of micrographs necessitated near‐continuous serial sectioning and camera lucida drawings, as illustrated in the vascular network of Pteridium aquilinum (L.) Kuhn by Tansley and Lulham (1904). Recently, advanced imaging techniques, including micro‐computed tomography (microCT), offer promising alternatives for non‐destructive, high‐resolution anatomical reconstruction of plant material (Piovesan et al., 2021; Suissa et al., 2023). However, microCT has significant drawbacks: it is expensive, technically demanding, and frequently limited to specialized research facilities. Scan times may extend over several hours, and access to the necessary instrumentation is not widely available. Moreover, without vertical stitching software, the challenge of examining internal structure across long lengths of tissues remains an issue with microCT. All of these factors collectively hinder their widespread use in comparative or field‐based anatomical and developmental studies.

Recent methods have been leveraged to generate 3D reconstructions of plant tissues at the anatomical level (Miki et al., 2023). These, and analogous approaches, have been used to reconstruct the anatomy of small regions of plants such as root tips, and even paleobotanical specimens (Matsunaga et al., 2017). These approaches allow for low‐cost, confocal‐like images to be generated for cell‐level analysis. However, we lack a standardized analogous approach for 3D reconstruction of internal plant structure across a long length of material.

To address these limitations, we adapt and expand upon the serial sectioning and videography approach first used by Zimmerman and Tomlinson for palm stem anatomy (Zimmermann and Tomlinson, 1965) and more recently with cycad petiole anatomy (Tomlinson et al., 2018). The serial section videography (SSV) protocol presented here offers a method for rapid serial sectioning and developmental videography to visualize internal structure over extended segments of plant tissue. By capturing sequential images of the cut face during sectioning—whether with a sliding microtome in the lab or a handheld razor blade in the field—and using open‐source software for alignment and reconstruction, our approach provides a new, accessible, low‐cost, and rapid protocol for generating developmental videography and 3D reconstructions of internal plant anatomy and micromorphology.

METHODS AND RESULTS

Sample collection and fixation

Four fern species with varying rhizome morphology and vascular anatomy were selected to assess this protocol. These included: Sitobolium punctilobulum (Michx.) Desv., Goniophlebium formosanum (Baker) Rödl‐Linder, Onoclea sensibilis L., and Lorinseria areolata (L.) C. Presl. Rhizomes were collected and cleaned by first removing the excess soil with a gentle wash in the sink. Roots were trimmed using a razor blade or pruners, and portions of the petioles were removed, leaving only the base of the petiole attached to the rhizome. Any soil left on the rhizome was removed using a soft‐bristle toothbrush under cold running water while gently scrubbing to avoid damaging the rhizome. Rhizomes for each species were either sectioned fresh from living material or fixed in formalin acetic alcohol (FAA) following standard protocol (Ruzin, 1999). Rhizomes were then placed in FAA for 24 h to preserve the tissue structure. After a day in the fixative, rhizomes were rinsed with 70% ethanol three times. Rhizomes were then placed in 50‐mL Falcon tubes or scintillation vials, which were filled with 70% ethanol and stored.

Sectioning and photography

The experimental setup to acquire images included a sliding microtome, camera, and camera mount (Appendix 1). We used a Leica SM2010 R sliding microtome (Leica Microsystems, Wetzlar, Germany), a standard camera mount, and a Sony Alpha 6000 digital camera (Sony Group Corporation, Tokyo, Japan) with a Tokina FiRIN 100 mm f/2.8 FE Macro lens (Kenko Tokina USA, Huntington Beach, California, USA) (Figure 1A). The methodology is described below for laboratory use, but the same technique can be applied using a smartphone with an attached 10–20× macro phone lens (e.g., Apexel APL‐HB100mm; Apexel, Littleton, Colorado, USA), steady hands, and a razor blade.

Figure 1.

SSV experimental setup. (A) Standard setup showing sliding microtome, camera mount, and lighting. (B) Rhizome placement in adjustable clamp. (C) Camera mount placement. (D) Alignment of the camera with the organ.

Prior to sectioning, the Sony Alpha 6000 was mounted onto a camera stand with the lens angled downwards. Rhizomes were tightly fastened into an adjustable cassette clamp on the sliding microtome to ensure stability and positioned with the shoot apex towards the bottom and the base of the rhizome towards the top (Figure 1B). After the rhizome was stabilized, the camera was manually focused, and the imaging parameters were manually set to ensure constant lighting between images (Figure 1C, D). Lighting fixtures were attached to the camera stand to ensure proper exposure. Fixtures were rotated and moved based on what visually appeared to provide the highest quality lighting.

Rhizomes were placed such that their position in the field of view was consistent throughout imaging. While this step was important, downstream digital adjustments were made when analyzing the image stacks to adjust images that were unaligned. Because scale bars were not used, the diameter of the rhizome was measured using digital calipers and recorded in a lab notebook prior to sectioning. This allowed for a global setting of the scale in downstream analysis (Figure 2A). To ensure smooth sectioning and consistent image quality, rhizomes were kept hydrated during sectioning using 70% ethanol for fixed material or H₂O for fresh material (Figure 2B). Blade selection occurred based on rhizome size and tissue toughness: low‐profile blades (Leica DB80LS) were used for smaller, softer rhizomes, while high‐profile blades (Leica DB80HS) were used for larger, tougher rhizomes. Section thickness varied between 50–150 µm depending on the rhizome; thicker rhizomes with high amounts of sclerenchyma were sectioned with high‐profile blades and thicker intervals, whereas thinner rhizomes with more parenchyma were sectioned with low‐profile blades at thinner intervals. Section thickness was kept consistent throughout the duration of each individual experiment.

Figure 2.

SSV setup details. (A) Measure organ to set the digital scale during analysis. (B) Ensure organ stays hydrated during sectioning (70% ETOH if fixed, H₂O if fresh). (C) For long, flimsy organs, stabilize with styrofoam. (D) To ensure high‐quality images, use remote shutter release.

Because rhizomes were sometimes too long to secure with the cassette clamp, we used a makeshift styrofoam mount to hold against the rhizome to ensure stability during sectioning (Figure 2C). Keeping the rhizome stable during sectioning was important to prevent movement between each section and to ensure consistent slice thicknesses. After each section, the microtome blade was locked, and the blade was covered with the safety cap. Images were captured using a remote shutter release to minimize any movement of the camera and ensure focus during imaging (Figure 2D). The camera parameters varied between rhizomes depending on the contrast and light intensity; however, the camera parameters were kept constant throughout each experiment. We set up the camera in Aperture Priority mode with an initial set of parameters as follows: aperture (f/8.0), ISO (200), and automatic shutter speed.

Rhizome sectioning and image capturing were continued until one to three nodes were sectioned (Figure 3). A single experiment yielded an image stack of approximately 100–300 serial images. Any remaining portion of the rhizome that was not sectioned was placed in a scintillation vial containing 70% ethanol for preservation. Individual sections left on the sliding microtome (Figure 1) were collected and placed in a scintillation vial in 70% ETOH for downstream histology.

Figure 3.

Examples of serial sections along the length of a rhizome showing departure of leaf traces. Sections move acropetally from left to right, from base towards tip. (A) Sitobolium punctilobulum, (B) Lorinseria areolata, (C) Onoclea sensibilis, (D) Goniophlebium formosanum.

Image alignment and videography

The image files were downloaded to a local desktop, and image stacks of serial sections were then uploaded into Fiji (Schindelin et al., 2012). Images were first downsized by 0.6× using the Scale function to reduce overall size. Image stacks were aligned using the Linear Stack Alignment with SIFT plugin, accessible via Plugins → Registration → Linear Stack Alignment with SIFT. This tool employs scale‐invariant feature transform (SIFT) to detect and match features across images, enabling alignment. Key parameters were generally kept at default values unless the image dimensions warranted adjustment. In most cases, we aligned image stacks once, but in the case of highly unaligned image stacks, alignment was conducted 3–5 times on previously aligned image stacks. In Fiji, the scale was set in the image stacks, and projection distances were set by inputting these manually into the Properties section of the image stacks. The image stacks were then saved as AVI files at 15 frames per second (Videos 1, 2, 3, 4).

Video 1.

Download video file ^{(8.8MB, mp4)}

Aligned developmental video of Lorinseria areolata.

Video 2.

Download video file ^{(2.6MB, mp4)}

Aligned developmental video of Onoclea sensibilis.

Video 3.

Download video file ^{(2.6MB, mp4)}

Aligned developmental video of Goniophlebium formosanum.

Video 4.

Download video file ^{(4.5MB, mp4)}

Aligned developmental video of Sitobolium punctilobulum.

Segmentation in 3D Slicer

Image stacks were imported into 3D Slicer for segmentation and visualization. This was done either directly via the SlicerMorph plugin or by first exporting the image stack from Fiji as an NRRD file. Segmentation followed standard 3D Slicer protocols (https://training.slicer.org/). Rhizome tissues were first segmented to create a rhizome mask, followed by segmentation of vascular tissues to generate a vascular mask. Both were segmented using the paint and interpolate approach in the Segment Editor tool: approximately every 5–15 slices were manually annotated, and the software interpolated the labels between these sections. Final 3D reconstructions of the rhizome and vascular tissue volumes were generated using the show3D module (Figure 4A–D).

Figure 4.

3D reconstructions of rhizomes and vascular architecture for (A) Lorinseria areolata, (B) Sitobolium punctilobulum, (C) Goniophlebium formosanum, (D) Onoclea sensibilis.

CONCLUSIONS

Here, we introduce the SSV method, a new, low‐cost protocol for generating developmental videos and 3D reconstructions of micromorphology. This protocol can be conducted in the laboratory with a high‐end sliding microtome and digital camera, or in the field with a smartphone and razor blade. In addition to low cost and quick results, this approach has the added benefit of allowing the user to generate these data from much longer samples than would be possible using traditional histological methods or microCT (without vertical stitching). Importantly, this protocol uses open‐source software, which is available to anyone. It can be used in conjunction with temporary stains, such as phloroglucinol, in species with hard‐to‐observe internal structure.

Compared to microCT, traditional histology, or histological 3D analysis (Miki et al., 2023), this protocol does not allow users to investigate cell‐level detail. Instead, SSV allows users to investigate internal micromorphology and development. Like the work of Zimmermann and Tomlinson (1965), SSV is particularly useful for examining vascular architecture across the length of an organ. This method provides a powerful and inexpensive tool for studying vascular development, the relationship between vascular architecture in stems and lateral organs, and the structure of nodes and internodes. Moreover, as anatomical traits (such as vascular area) can vary along an organ (Isebrands and Larson, 1977; Herrera et al., 2015), SSV can provide insight into changes in the area and patterning of the vasculature within an individual organ in comparative studies. SSV can replicate the anatomical detail seen in some low‐resolution microCT scans without the need for advanced instrumentation, making it useful for large‐scale, comparative, or field‐based studies. By generating developmental videography along the length of an organ and lowering the technical and financial barriers to creating 3D reconstruction of plant organs, this method expands our capacity for developmental and structural studies in plants.

AUTHOR CONTRIBUTIONS

J.S.S. developed the methods, analyzed the data, and wrote the manuscript. G.R. sectioned plant material, analyzed data, and drafted the manuscript. Both authors approved the final version of the manuscript.

Appendix 1. Protocol for conducting the serial section videography method.

Materials

If done in the laboratory with a sliding microtome:

• 1.Plant material
• 2.Sliding microtome
• 3.Low‐ or high‐profile blade
• 4.Camera
• 5.Macro lens
• 6.Camera stage
• 7.Remote shutter release
• 8.Digital calipers or ruler
• 9.Squeeze bottle
• 10.Computer downloaded with Fiji and 3D Slicer

If done in the field:

• 1.Plant material
• 2.Razor blade (single‐edge)
• 3.Camera or smartphone with hand lens attachment
• 4.Digital calipers or ruler
• 5.Computer downloaded with Fiji and 3D Slicer

Methodology

Sample collection and fixation

• 1.Collect plant material.
• 2.Clean material: Remove excess soil or debris with a gentle wash in the sink, trim roots or lateral organs.
• 3.Fixation: Place plant material in formalin acetic alcohol (FAA) for 24 h to preserve tissue structure (optional).
• 4.Rinse: After fixation, rinse material with 70% ethanol three times.
• 5.Storage: Store material in 50‐mL Falcon tubes or scintillation vials filled with 70% ethanol.

Sectioning and photography (in the lab)

• 1.Set up equipment: Mount the camera onto a camera stand with the lens angled down.
• 2.Position the organ: Fasten the organ into the adjustable cassette clamp on the sliding microtome. Measure a reference point for downstream scale setting.
• 3.Focus the camera: Manually focus the camera and set imaging parameters to ensure constant lighting.
• 4.Sectioning: Use appropriate blades based on organ size and tissue toughness; maintain consistent section thickness (50–150 µm).
• 5.Capture images: Use a remote shutter release to minimize movement; capture images until one to three nodes are sectioned.

Sectioning and photography (in the field)

• 1.Hold organ steady in hand or on lab bench.
• 2.Slice sections of equal thickness using a single‐sided blade. Care should be taken to minimize variation in slice thickness.
• 3.Take successive images of the cut end of the organ. Care should be taken to maintain the same orientation while imaging the organ. Use a camera with a macro lens or a smartphone with an attached 10× hand lens to image.

Image alignment and videography

• 1.Download images: Transfer the images from the camera's SD card (or smartphone) to a computer.
• 2.Upload to Fiji: Load the image stack into Fiji software.
• 3.Downsize images: Use the Scale function to reduce the overall size by 0.6×.
• 4.Align images: Use the Linear Stack Alignment with SIFT plugin for alignment.
• 5.Set scale: Set the scale in the image stacks using digital caliper measurements.
• 6.Save as video: Save the aligned image stacks as AVI files at 15 frames per second.
• 7.Save the image stack as an NRRD file.

Segmentation in 3D Slicer

• 1.Import image stack: Load the image stacks into 3D Slicer via the SlicerMorph plugin or as NRRD files.
• 2.Segment tissues: Use the Segment Editor tool to create masks for the external tissue of the organ and vascular tissues or internal tissues of interest.
• 3.Interpolate labels: Manually annotate every 5–15 slices; interpolate labels between sections.
• 4.Generate 3D reconstructions: Use the show3D module to create final 3D reconstructions.

Potential complications and points to consider

Sectioning:

• 1.Care should be taken when clamping the organ in the sliding microtome to avoid applying too much pressure.
• 2.Vertical positioning of the organ in the sliding microtome is up to the discretion of the user. The position of the organ should be considered in order to maximize the desired length while maintaining stability of the structure. For instance, if too long of a portion is placed in the clamp, it may become flimsy and unable to be sectioned properly.
• 3.Maintaining the rhizome in a consistent position throughout image capture is essential for proper orientation during the video and image alignment. The user will want to minimize any positional shifts.
• 4.It is important to maintain consistent sectioning thickness within each experiment for the downstream analysis of image stacks.
• 5.Caution should be taken when placing hands near the organ or sliding microtome blade to prevent any injury that could occur.
• 6.If users desire to keep sections for downstream embedding and traditional histology, they can fill the sliding microtome reservoir with 70% ETOH to catch each section that falls during the sectioning process. This does not, however, maintain the serial order of the sections. If users would like to keep the serial order, sections can be individually collected after each slice and placed in an individual vial of 70% ETOH.
• 7.Maintaining the microtome in optimum condition after each use is crucial. To ensure proper care, any remaining debris should be dusted off using a brush, the microtome tray should be detached and debris thrown away, and all surfaces should be wiped down with 70% ethanol.
• 8.Blades can be used until they become dull; they will often last through several organs. The cutting edge can also be shifted along the blade's length, which will extend the overall use time of the blade.

Image acquisition and alignment:

• 1.Camera settings should be set to the user's specifications considering the available light intensity and contrast of the plant material being imaged. Ensuring the camera is in manual focus, aperture, shutter speed, and ISO will ensure consistent images across the sample.
• 2.Users should adjust the alignment parameters to best fit their image stacks.
• 3.Sectioning in the field will lead to larger discrepancies between slices and potentially different slice thickness. This may cause issues with quantitative analyses in the Z‐projection; however, it does not drastically affect the reconstructions qualitatively.

DATA AVAILABILITY STATEMENT

All data are provided in the manuscript.