# Confocal fluorescence microscopy reveals subtle lignification variations in a Scots pine blue ring

Ciara Greaves, Ignacy Bonter, Alan Crivellaro, Clive Oppenheimer, Harriet Kempson, Aleksei Potapov, Sandra Metslaid, Maris Hordo, Regino Kask, Ahto Kangur, et al.

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## To cite this version:

Ciara Greaves, Ignacy Bonter, Alan Crivellaro, Clive Oppenheimer, Harriet Kempson, et al. Confocal fluorescence microscopy reveals subtle lignification variations in a Scots pine blue ring. Dendrochronologia, 2025, Dendrochronologia, 90, pp.126301. 10.1016/j.dendro.2025.126301. hal-05034849

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# Confocal fluorescence microscopy reveals subtle lignification variations in a Scots pine blue ring

Ciara Greaves<sup>a, *</sup>, Ignacy Bonter<sup>b</sup>, Alan Crivellaro<sup>c,d</sup>, Clive Oppenheimer<sup>a</sup>, Harriet Kempson<sup>b</sup>, Aleksei Potapov<sup>e</sup>, Sandra Metslaid<sup>e</sup>, Maris Hordo<sup>e</sup>, Regino Kask<sup>e</sup>, Ahto Kangur<sup>e</sup>, Simon Hawkins<sup>f</sup>, Corentin Spriet<sup>g</sup>, Andrew D. Friend<sup>a,h</sup>, Jim Haseloff<sup>b</sup>, Alma Piermattei<sup>c,d</sup>

## ARTICLE INFO

### Keywords:

- Blue rings

- Confocal microscopy

- Double staining

- Lignin

- Tree rings

- Xylem

## ABSTRACT

Blue rings (BRs) are anomalies in tree rings revealed by a double staining technique. They have been thought to reflect reduced cell wall lignification during ephemeral end-of-growing-season cold periods, often following significant volcanic eruptions. BRs have been distinguished visually according to qualitative descriptions and are considered a promising proxy for cold extremes at the intra-seasonal scale. However, reduced cell wall lignin content has never been successfully measured in BRs, limiting understanding of their relationship with thermal (or other) conditions. Further, difference in lignin content between BRs and 'standard' rings has not been confirmed independently of the double staining method. Here, using confocal microscopy of samples of Scots pine, we show that BRs are indeed associated with marked reduction of lignin in latewood cells. We also identify subtle variations in lignin across rings that are not well resolved by the double staining method. We draw on these findings to propose a new framework to visualise and measure lignin changes in tree rings. Our findings emphasise the potential of BRs as a paleoproxy of intraseasonal climate variability.

## 1. Introduction

A blue ring (BR) is considered to identify reduced cell wall lignification in tree rings, exposed in transverse thin wood anatomical sections treated with a double staining technique, whereby thin wood sections are treated with a solution of Safranin and Astra-Blue (Gerlach, 1984; Schweingruber, 2007). Single or multiple tangential layers of blue-stained xylem cells (i.e. axial tracheids) typically occur in the latewood and have been observed in several conifer species (Piermattei et al., 2015, 2020; Tardif et al., 2020; Büntgen et al., 2022). They are thought to indicate a response to abrupt and abnormal cooling at the end of the growing season (Gindl et al., 2000; Piermattei et al., 2015), which other tree-ring parameters such as latewood density and ring width may not capture (Gindl et al., 2000; Piermattei et al., 2020; Greaves et al.,

2023), thereby offering new insights into ephemeral climate extremes of the past.

While we adopt the definition of a BR as a continuous layer of less lignified axial tracheids (Piermattei et al., 2015; Crivellaro et al., 2018), BRs often present differently, and we expect that within a BR there may be considerable heterogeneity in lignin abundance. Though attempts have been made to describe BRs qualitatively based on their visual appearance (Montwé et al., 2018; Piermattei et al., 2020), the lignin content in BRs has yet to be measured or differentiated. Concealed lignin variability may offer additional insights into past climatic phenomena, such as variations in the intensity or duration of cooling events, surpassing the binary BR categorisation employed in previous studies. Further, while the interpretation of BRs as signifying lignin reduction is widely accepted, it has not been verified.

<footnote><sup>a</sup> Department of Geography, University of Cambridge, Cambridge, UK</footnote>

<footnote><sup>b</sup> Department of Plant Sciences, University of Cambridge, Cambridge, UK</footnote>

<footnote><sup>c</sup> Department of Agricultural, Forest and Food Sciences, University of Torino, Grugliasco, Italy</footnote>

<footnote><sup>d</sup> Forest Biometrics Laboratory, Faculty of Forestry, "Stefan cel Mare" University of Suceava, Romania</footnote>

<footnote><sup>e</sup> Chair of Forest and Land Management and Wood Processing Technologies, Institute of Forestry and Engineering, Estonian University of Life Sciences, Tartu, Estonia</footnote>

<footnote><sup>f</sup> Université de Lille, Sciences et Technologies, CNRS, UMR 8576-UGSF-Unité de Glycobiologie Structurale et Fonctionnelle, Lille F-59000, France</footnote>

<footnote><sup>g</sup> Université de Lille, CNRS, Inserm, CHU Lille, Institut Pasteur de Lille, US 41 - UAR 2014 - PLBS, Lille F-59000, France</footnote>

<footnote><sup>h</sup> Conservation Research Institute, University of Cambridge, Cambridge, UK</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: ccg39@cam.ac.uk (C. Greaves).</footnote>
Several techniques have been developed to quantify and characterise cell wall lignification in plants, providing both absolute and relative measurements (Hatfield and Fukushima, 2005; Asshoff and Hättenschwiler, 2006; Lupoi et al., 2015). A disadvantage of some of these methodologies, such as chemical assays or bulk spectroscopy, is that they are destructive and/or do not resolve internal variability, providing instead a mean lignin content for the sample (Baldacci-Cresp et al., 2020). For tree-ring research, where subtle changes throughout the ring may capture aspects of environmental and ecological variability within a growing season, an approach capable of revealing spatial detail is desirable. Techniques such as Raman spectroscopy and UV microspectrophotometry have been used to measure lignin in various tree species (Röder et al., 2004; Agarwal, 2006; Lehringer et al., 2008; Gierlinger et al., 2010; Miyafuji et al., 2017; Fredriksson et al., 2018). Raman spectroscopy has also been applied to BRs but interference from other aromatic compounds hindered identification of lignin in these Mountain pine (*Pinus uncinata* Mill.) samples (Piermattei et al., 2020).

Another approach is confocal microscopy (Donaldson et al., 2010), which can reveal lignin from its autofluorescence, or in conjunction with other fluorescent stains, without interference from other compounds (Dickson et al., 2017). Baldacci-Cresp et al., (2020) used it to study a range of plant material, including the xylem of poplar (*Populus tremula* x *alba*). Their samples were stained only with Safranin, which binds to lignin, thus fluorescently labelling lignified xylem cell walls (Bond et al., 2008; Mottiar et al., 2020). Baldacci-Cresp et al., (2020) then used a macro in ImageJ software to calculate the ratio of red-to-green fluorescence in micrographs, thereby mapping the amount of lignin within cell walls. Our aim was to apply this approach to conifer tree rings for the first time and, in turn, independently verify the reduction of lignin in BRs and resolve fine-scale variations in lignin content. Using samples of Scots pine (*Pinus sylvestris* L.) we compare the lignin content within the latewood of a BR and that of a 'standard' (fully lignified) ring.

## 2. Materials and methods

We used samples of Scots pine collected by Metslaid et al. (2011) from an Estonian hemiboreal forest in which BRs were previously identified for the year 1976 by Greaves et al. (2023). This 1976 BR was associated with uncommonly low September-October temperatures (the end of the growing season). Our sample was a south-facing radius cut from a disk at 2.5 m in height up the tree, the radius was ~1.5 cm wide across its transverse section and 2 cm thick along its axial section. Sub-samples, spanning five rings, including the 1976 BR, were isolated (Greaves et al., 2023). Using a WSL Lab microtome (Gärtner et al., 2015a), multiple 20-µm-thick transverse cross-sections were cut from the selected region of the radii (Gärtner and Nievergelt, 2010; Gärtner et al., 2015b). Two staining procedures were used: the traditional double staining method to visualise the BR and Safranin staining for the Baldacci-Cresp et al. (2020) confocal imaging protocol.

The first group of thin sections were double stained with Safranin and Astra-Blue as a reference for the study (Gerlach, 1984; Schweingruber, 2007), whereby lignified cells appear red and less lignified cells blue – this is the typical method for identifying BRs (Piermattei et al., 2015). The Safranin solution contained 0.1 g of Safranin powder dissolved in 100 ml of distilled water, while the Astra-Blue solution comprised of 0.5 g of Astra-Blue powder combined with 100 ml of distilled water and 2.0 ml of acetic acid. The two solutions were combined in a ratio of 1:4 (one part of Safranin and four parts Astra-Blue) (Piermattei et al., 2020). The thin sections were stained for five min, then washed with distilled water, and then with an ethanol series (50%, 75%, then 99%) to remove any excess stain. The double-stained thin sections were placed on a glass slide, mounted in water, and enclosed by a glass cover slip. The highest quality thin section, e.g., with least damaged cell walls, was selected for imaging with a compound microscope at 40 × magnification.

The other group of thin sections were stained for 10 min in a Safranin

solution, derived from dissolving 0.2 g of Safranin powder in 100 ml of 50% ethanol solution. Thin sections were subsequently rinsed in a 50% ethanol solution for 10 min and then in distilled water for 15 min. The Safranin-stained thin sections were finally mounted in water on a glass slide and enclosed with a glass cover slip. Again, the best thin section was selected for imaging. Mounting samples in water under a glass slide demands immediate confocal imaging, i.e., before desiccation occurs. The use of gene frames was attempted but the sample floated, and a flat surface was unavailable, preventing high-resolution imaging. We note that other non-fluorescing mounting agents might preserve the sample for longer.

Images of the Safranin-stained thin section were obtained with a Leica SP8 confocal microscope. Sequential 400 Hz bidirectional scanning was undertaken in 16-bit mode with hybrid detectors (HyD) in standard mode, using a water immersion 40 × objective. Two white light laser (WLL) lines were used to excite the sample using sequential scanning, with the 'between lines' setting. The first set, with excitation line at 488 nm (green), was adjusted to 66.5% relative power with a gain of 500 and a collection window (emission bandpass) of 540–560 nm. While the second set, with excitation line at 560 nm (red), was set to 25% relative power with a gain of 222.5 and a collection window of 570–600 nm (Baldacci-Cresp et al., 2020). WLL was set to 70% power both to optimise image quality and avoid damage to the sample and microscope. The excitation lines were tested between 10% and 70% relative to the total WLL power to find optimum settings for our Scots pine samples, i.e., yielding images with wide dynamic range and without saturation. The Z-step, i.e., the vertical distance between sequential optical sections in a 3D image stack, was set to 1–1.5 µm, depending on sample thickness and to obtain a consistent number of optical slices through the sample. Additionally, intensity values from three adjacent scan lines were averaged to improve signal-to-noise. Five image files (Series 8, 11, 18, 19 and 20) of different regions of our standard ring (control) and five regions (Series 3, 4, 6, 7 and 10) of the BR were generated.

The image files obtained from the confocal microscope were processed in ImageJ (Baldacci-Cresp et al., 2020). Within ImageJ, stack viewing was configured to 'hyperstack', allowing multiple image planes and channels to be amalgamated. Additionally, the 'colourised' function was used to assign colours to the raw greyscale images to enhance visual interpretation. The image channels were organised with the green channel preceding the red channel. Subsequently, a Z Project at maximum intensity was generated. This yields a composite image emphasising the brightest pixel values from a stack of images along the z-axis. The images were rotated so that the ring boundary was perpendicular to the direction of cell growth and cropped to ten vertical rows of latewood cells and three rows of earlywood cells, representing regions of ~300 µm × 400 µm. Then, the 'Safranin ratio' ImageJ macro, developed by Baldacci-Cresp et al. (2020), was run to produce ratiometric images, i.e., the ratio of red-to-green light emitted by Safranin-stained lignified cell walls. An automated brightness threshold was chosen to distinguish cellular structures from empty spaces (Baldacci-Cresp et al., 2020), thereby excluding voids (e.g., xylem lumen) from subsequent statistical and image analyses of lignin content. The largest region of cells between rays (which were not relevant to our study) was selected for subsequent analysis.

Profiles of mean red-to-green light ratios through the latewood and earlywood of the ring were generated by calculating the mean red-to-green ratio for each vertical column of pixels across the images. A profile was plotted for each image, and each profile was fitted with two horizontal lines representing the mean latewood and earlywood red-to-green light ratios. Due to the automated thresholding, there was some minor variability in pixel values between images, so the plots were not stacked. Additionally, regions of interest were selected from the largest area of cells between rays in the latewood of each image, for which the mean red-to-green ratio was computed.

Additionally, to facilitate visual interpretation and intercomparison
of the images, we remapped pixel values using a look-up table (LUT), whereby red-to-green ratios range from 0 (purple), indicating no lignin, to 2 (red), showing a high degree of lignification (Baldacci-Cresp et al., 2020).

### 3. Results

The 1976 BR is visualised with the original Safranin and Astra-Blue double-staining method in Fig. 1 A. In the 1976 ring, the earlywood cells and the first half of the latewood are red-stained, except for blue-stained ray cells throughout the latewood (Fig. 1 A). The final rows of axial tracheids of the latewood have blue-stained secondary cell walls and red-stained middle lamellae and primary cell walls (Fig. 1B). The latewood of the 1975 ring is entirely red-stained (Fig. 1 A and C).

The confocal microscope image in Fig. 1D reveals lignin deficiency in the 1976 BR latewood compared with the 1975 standard ring latewood (Fig. 1E). In both images, the latewood middle lamellae and primary cell walls fluoresce prominently, indicating the presence of lignin. However, there is more fluorescence for the 1975 latewood (Fig. 1E) than the 1976 latewood (Fig. 1D). The secondary cell walls of the latewood in the 1976 ring (Fig. 1D) are darker and less visible than those of the standard ring (Fig. 1E), demonstrating reduced lignin in the BR.

Fig. 2A and B further reveal lignin changes as represented by scaled red-to-green ratios, which increase with increasing lignin concentrations. The BR (1976) latewood secondary cell walls (Fig. 2B) are frequently yellow and green with some blue, representing a marked reduction in lignin content compared with the red and orange of the standard (1975) latewood (Fig. 2A). The middle lamellae and primary

cell walls of the latewood and earlywood consistently exhibit high lignin abundance in both images. However, the red (higher lignin) middle lamellae and primary cell walls are more apparent in the standard (1975) ring latewood and subsequent (1976) earlywood (Fig. 2A). Further images are provided in the Supplementary materials.

Fig. 2C and D display profiles of raw red-to-green ratio changes (i.e. lignin abundance), within the region of interest of the corresponding standard ring and BR images. We note that the amplitude of the fluctuations in the ratiometric profile in the latewood is consistently greater in the BR compared with the standard ring. This difference is attributed to a greater contrast in ratio values between the BR secondary and primary cell walls and middle lamellae. Fig. 2C (the standard ring) indicates a consistent red-to-green ratio of ~1.5 until the latewood/earlywood boundary, whereupon it increases in the earlywood cells of the next ring (~1.6). Fig. 2D shows lignin changes in the 1976 BR. The ratio initially troughs at ~1.2, then rises steeply at ~300 pixels, reaching ~1.6 at the latewood/earlywood boundary. It peaks at ~1.7 at ~1050 pixels, in the initial early wood cells, then declines to ~1.4. Further examples illustrating the same distinction between standard ring and BR but to varying degrees are provided in the Supplementary material. In four of the five BR profiles, the ratio increases until the earlywood/latewood boundary and then declines in the subsequent earlywood cells, the exception being the profile of Series 10 where the ratio is stable in the earlywood cells of the following ring (see Supplementary material). All five of the standard ring profiles exhibit a relatively consistent red-to-green ratio in the latewood followed by a rise in the ratio in the earlywood cells of the next ring (1976).

Mean values of latewood red-to-green ratios for the five standard

<figure><img src="image_5.png" /><figcaption>**Fig. 1.** (A) Zoomed image of a 20-µm-thick cross-section of Scots pine double stained with Safranin and Astra-Blue, revealing a latewood BR (less lignified axial tracheid cell walls) for the year 1976 (Greaves et al., 2023). (B and C) Images of the double stained sections latewood/earlywood boundary at 1976/1977 and 1975/1976 with zoom. (D and E) Safranin-stained samples under a confocal microscope, wherein regions with higher lignin abundance fluoresce more and are brighter in the image. Imaging and display conditions were consistent for each image. Scale shown is approximate.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Ratiometric images of Scots pine standard ring (A) (Series 8) and BR (B) (Series 6) latewood followed by earlywood of the following ring. Scale shown is approximate. The LUT colour scale ranges from a red-to-green ratio of 0 (purple) to a ratio of 2 (red), and the white box represents the region for which profile data are taken. (C and D) Line graphs representing profiles of red-to-green light ratios (representing lignin abundance) along with horizontal lines indicating respective mean latewood and earlywood red-to-green ratios, and a dashed line indicating the latewood/earlywood boundary. (E and F) Box plots representing the median (horizontal grey line), interquartile ranges (box), range excluding outliers (whiskers) and outliers (circles) of the latewood ratios generated from all the standard ring and BR images.</figcaption></figure>

ring images analysed vary from 1.47–1.67 with a mean of 1.58 ± 0.098 (1 sigma standard deviation; Fig. 2E). Mean red-to-green ratios of the latewood for the five BR images analysed vary from 1.27–1.42 with a mean of 1.36 ± 0.058 (Fig. 2F). The mean red-to-green ratio for the BR

is 2.21 standard deviations below that for the standard ring. As seen in the box plots (Fig. 2E and F), the BR latewood has a markedly lower ratio than the standard ring, with no BR values lying within the range for the standard ring. More variability in the ratios is seen in the standard ring
images, which have a range of 0.204 compared with 0.155 for the BR samples.

We note that the BR cell walls show evidence for some loss of structural integrity. For example, in Fig. 2B, some fraying and broken cell walls can be seen. However, lignin (red-to-green ratio) profiles are well replicated (see Supplementary material) in our images and differences in lignin content between standard rings and BRs exceed any noise introduced by this fraying.

## 4. Discussion and conclusions

Our confocal microscopy results show pronounced reductions in latewood lignin content in BRs compared with standard rings, validating previous interpretations of BRs (Piermattei et al., 2015, 2020; Crivellaro et al., 2022).

Our profiles of lignin variations throughout the tree rings also reveal details not captured by the double-staining technique (Fig. 2C and D), opening the potential to obtain new insights into the relationship between xylogenesis and environmental conditions. Notably, lignin content rises in the final layers of BR latewood cells. Additionally, we show a positive step change in lignin in the earlywood cells of the subsequent ring to the standard ring, confirming findings from other studies that identify a higher proportion of lignin in earlywood cells compared to latewood cells (Fredriksson et al., 2018). In contrast, although the lignin content of earlywood cells increases in the ring following a BR initially (compared to the latewood), it subsequently decreases. These trends are both well replicated in the images analysed (see Supplementary material).

While the overall lignin reduction in BRs was expected, other aspects of our findings raise questions concerning the still poorly understood mechanism of lignification (Rathgeber et al., 2016; Pérez-de-Lis et al., 2024). The recovery of lignin content in the final outermost layers of latewood cells in our 1976 sample is not resolved by the double staining procedure, which reveals blue-stained secondary cell walls for the final rows of latewood cells (Fig. 1 A). The processes behind this increase in lignin in the final latewood cells are unclear. Possible explanations include environmental factors, an adaptive mechanism, or use of reserves from the growing season to lignify these final rows of latewood cells. Another consideration is the reduced size of the terminal tracheids: since their cell wall area is smaller (Gindl et al., 2000), less time may be required for differentiation. Additionally, the composition of lignin types, e.g., p-hydroxyphenyl, guaiacyl, and syringyl subunits (Antonova et al., 2019), could influence the lignification dynamics in the terminal tracheids. The reduction in earlywood lignin content trend in the 1977 ring (Fig. 2D) (following the 1976 BR) is also unexpected. It could reflect a cold start to the growing season, but a more interesting possibility is that it indicates a depletion of carbon allocation resources.

We recognise that our study is limited to analysis of a single BR (1976) and one standard ring (1975), questioning its representativeness. However, it offers a template for further studies on lignification patterns across different tree rings and species. BRs have presented elsewhere in different forms visually and have also been associated with varying climatic and environmental contexts. Variability in abiotic or biotic factors may result in tree rings with alternative lignin profiles. We hope that our findings will stimulate further research, for instance in calibrating ratiometric values obtained by fluorescence microscopy to absolute measures of lignin abundance. This could be achieved by preparing and imaging slides with known quantities of commercially available lignin, thus providing a standard for comparison.

Our study focused on the 1976 BR from Scots pine grown in Estonia, associated with anomalously cold September-October temperatures (Greaves et al., 2023). Future research should explore lignin abundance in BRs across different species and environmental conditions, as well as its variability within the same species or individual plants. Such investigations will enhance our understanding of the relationship between thermal conditions and lignin abundance, thereby expanding the

potential for BRs as a proxy for within-growing season climate variability.

## Funding sources

C.G received funding from Natural Environment Research Council - United Kingdom Research and Innovation. I.B received funding from the Herchel Smith Fund. This study was supported by the BB/L014130/1 for the OpenPlant Synthetic Biology Research Centre to J.H. A.Piermattei was funded by the Agritech National Research Center and received funding from the European Union Next-Generation EU (PIANO NAZIONALE DI RIPRESA E RESILIENZA (PNRR) - MISSIONE 4 COMPONENT 2, INVESTIMENTO 1.4 - D.D. 1032 17/06/2022, CN00000022). This study was supported by the Estonian Ministry of Education and Research (project SF0170014s08), Estonian Science Foundation (Grant 8890), Järvseja Training and Experimental Forest Centre, Estonian Environmental Investment Centre, Estonian University of Life Sciences ASTRA project Value-chain based bio-economy (supported by the European Union, European Regional Development Fund). A.D.F acknowledges support from the Natural Environment Research Council [grant number NE/W000199/1].

## Data statement

The data produced in this study is available in the Supplementary information file.

## CRediT authorship contribution statement

**Friend Andrew D.:** Writing – review & editing, Supervision, Conceptualization. **Bonter Ignacy:** Writing – review & editing, Writing – original draft, Methodology, Investigation, Data curation, Conceptualization. **Haseloff Jim:** Supervision, Funding acquisition. **Crivellaro Alan:** Writing – review & editing, Writing – original draft, Supervision, Investigation, Data curation, Conceptualization. **Spriet Corentin:** Writing – review & editing, Validation, Software, Resources, Methodology. **Greaves Ciara:** Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Software, Resources, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization. **Potapov Aleksei:** Writing – review & editing, Investigation. **Metslaid Sandra:** Writing – review & editing, Investigation. **Piermattei Alma:** Writing – review & editing, Writing – original draft, Visualization, Validation, Project administration, Methodology, Investigation, Formal analysis. **Oppenheimer Clive:** Writing – review & editing, Writing – original draft, Supervision, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Kempson Harriet:** Software, Conceptualization. **Kangur Ahto:** Investigation. **Hawkins Simon:** Writing – review & editing, Validation, Software, Resources, Methodology. **Hordo Maris:** Investigation. **Kask Regino:** Investigation.

## Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgements

C.G and I.B designed the study. A.Potapov, S.M, M.H, R.K and A.K conducted the fieldwork to collect samples. C.G undertook the slide preparation, image analysis and data analysis. I.B carried out the confocal imaging. H.K, S.H, C.S, A.Piermattei and A.C provided technical support. C.G prepared the first draft with input from A.Piermattei, A.C, I.B and C.O. All other authors contributed to improving the article. We are grateful to the two anonymous reviewers for their constructive
comments on the manuscript.

## Appendix A. Supporting information

Supplementary data associated with this article can be found in the online version at doi:10.1016/j.dendro.2025.126301.

## Data availability

The data produced in this study is available in the Supplementary information file.

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