Fabien Baldacci-Cresp
email: fabien.baldacci-cresp@univ-lille.fr

Corentin Spriet
email: cspriet@univ-lille.fr

Laure Twyffels
email: laure.twyffels@ulb.ac.

Anne-Sophie Blervacq
email: sophie.blervacq@univ-lille.fr

Godfrey Neutelings
email: godfrey.neutelings@univ-lille.fr

Marie Baucher
email: mbaucher@ulb.ac.be

Simon Hawkins
email: simon.hawkins@univ-lille.fr

A rapid and quantitative safranin-based fluorescent microscopy method to evaluate cell wall lignification

SUMMARY

One of the main characteristics of plant cells is the presence of the cell wall located outside the plasma membrane. In particular cells, this wall can be reinforced by lignin, a polyphenolic polymer that plays a central role for vascular plants, conferring hydrophobicity to conducting tissues and mechanical support for upright growth. Lignin has been extensively studied by a range of different techniques including anatomical and morphological analyses using dyes to characterize the polymer localization in situ. With the constant improvement of imaging techniques, it is now possible to revisit old qualitative techniques and adapt them to obtain efficient, highly resolutive, quantitative, fast and safe methodologies. In this study, we revisit and exploit the potential of fluorescent microscopy coupled to safranin-O staining to develop a quantitative approach for lignin content determination. The developed approach is based on ratiometric emission measurements and the development of an ImageJ macro. To demonstrate the interest of our methodology compared to other commonly-used lignin reagents, we showed that safranine-O staining was used to evaluate and compare lignin contents in previously characterized A.

thaliana lignin biosynthesis mutants. In addition, the analysis of lignin content and spatial distribution in the Arabidopsis laccase mutant also provided new biological insights into the effects of laccase gene down-regulation in different cell types. Our safranine-O based methodology, also validated for flax, maize and poplar, significantly improves and speed-up anatomical and developmental investigations of lignin, which we hope will contribute to new discoveries in many areas of cell wall plant research.

SIGNIFICANCE STATEMENT (2 sentences 75 words max)

Lignin has long been the subject of extensive study as it plays a major role in plant growth and defense and is an important factor affecting the quality of plant biomass and plant-derived products. The development of an easy and quantitative confocal microscopy method to provide detailed spatial information about lignin content at the cell wall level will prove a valuable addition to the toolbox of lignin analytical methods for both fundamental and applied research.

INTRODUCTION

A main characteristic of the plant cell is the presence of a cell wall. This wall allows plant cells to acquire a certain rigidity essential for the maintenance of a strong intracellular osmotic pressure that contributes to maintaining the plant stem upright. Depending upon cell types, this wall can be mainly composed of up to three layers. The first one is the middle lamella, a thin structure mainly composed of pectins and shared by two adjacent cells. This layer is the pedestal on which the two other structuresthe primary and the secondary cell walls -can be developed. The primary wall is the first to be deposited and is mainly composed of cellulose, hemicelluloses, pectins and proteins [START_REF] Carpita | Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth[END_REF]. In certain tissues, plant cells acquire a secondary wall that is deposited between the primary cell wall and the plasma membrane. Like primary cell walls, secondary cell walls are rich in cellulose, but generally show higher hemicellulose and lower pectin content providing greater mechanical strength and reduced extensibility. This secondary cell wall, generally much thicker than the primary wall and made of several layers (S1, S2 and S3), is mainly observed in cells from vascular and supporting tissues. The formation of the secondary cell wall is, in most cases, associated with the deposition of lignin that confers hydrophobicity and rigidity to the cell wall [START_REF] Boerjan | Lignin biosynthesis[END_REF]. Lignin is a phenolic polymer composed of 3 main units, p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units derived fom pcoumaryl alcohol, coniferyl alcohol and sinapyl alcohol, respectively. The biosynthesis of lignins is initiated during the formation of the S1 secondary cell wall layer and newly formed lignin is first observed in the cell corners and middle lamella before spreading progressively, to the primary cell wall and different layers (S1, S2 and S3) of the secondary cell wall [START_REF] Terashima | Heterogeneity in formation of lignin-XI: An autoradiographic study of the heterogeneous formation and structure of pine lignin[END_REF]. Given the major importance of ligninboth for the plant and economically -it is not surprising that the biosynthesis/polymerization and degradation of this polymer have been extensively studied for a very long time [START_REF] Boerjan | Lignin biosynthesis[END_REF].

One of the major challenges of recent and future years is to improve our ability to characterize, both quantitatively and qualitatively, lignin in plant cell walls. In terms of quantitative analysis, three main types of wet chemistry methods are traditionally used: acetylbromide assays (Morrison, 1972b, Morrison, 1972a), the acidic method [START_REF] Van Soest | Use of detergents in the analysis of fibrous foods. II. A rapid method for the determination of fibre and lignin[END_REF] or the Klason method [START_REF] Johnson | The spectrophotometric determination of lignin in small wood samples[END_REF].

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the Klason method seems more suitable and efficient on woody species, the acetylbromide and the acidic methods are more suitable for herbaceous species. In these species, the acetyl bromide method is faster and presents a better recovery of lignin [START_REF] Fukushima | Comparison of acetyl bromide lignin with acid detergent lignin and Klason lignin and correlation with in vitro forage degradability[END_REF].

Various types of methods are also available for determining lignin chemical composition (i.e.

determination of S, G and H contents and calculation of S/G ratios). There include chemical degradation techniques, such as thioacidolysis [START_REF] Lapierre | Thioacidolysis of Poplar Lignins: Identification of Monomeric Syringyl Products and Characterization of Guaiacyl-Syringyl Lignin Fractions[END_REF] and nitrobenzene oxidation [START_REF] Monties | Lignins[END_REF], as well as derivatization methods and spectroscopic methods based on NMR [START_REF] Mansfield | Whole plant cell wall characterization using solution-state 2D NMR[END_REF] or using pyrolysis-GC/MS coupling [START_REF] Wagner | Exploring lignification in conifers by silencing hydroxycinnamoyl-CoA:shikimate hydroxycinnamoyltransferase in Pinus radiata[END_REF].

If these qualitative and quantitative approaches are powerful, they all present the same drawback: they are destructive methods that require grinding and/or extraction of cell wall polymers/lignins from the plant sample being analyzed. As a result, the data obtained represent an overall mean value for a whole organ/tissue and thus do not take into account the fact that lignin content and composition vary between different tissues, cells and cell wall layers. To get around this problem and to obtain information on lignin localization more rapidly, researchers have exploited different histochemical methods including basic fushin and auramine staining (epifluorescence observations) [START_REF] Dharmawardhana | Characterization of vascular lignification in Arabidopsis-Thaliana[END_REF][START_REF] Pesquet | Novel markers of xylogenesis in zinnia are differentially regulated by auxin and cytokinin[END_REF] revealing global lignin deposition, as well as more 'specific' tests such as phloroglucinol staining (Weisner reagent) revealing lignin hydroxycinnamaldehyde functions [START_REF] Vallet | Histochemistry of Lignin Deposition during Sclerenchyma Differentiation in Alfalfa Stems[END_REF][START_REF] Liljegren | Phloroglucinol stain for lignin[END_REF], and Maüle staining providing indications on the S/G ratio [START_REF] Meshitsuka | Studies on the mechanism of lignin color reaction. XIII. Maule color reaction[END_REF]. Another approach based on safranin/alcian blue staining, the FASGA method [START_REF] Tolivia | Fasga: A New Polychromatic Method for Simultaneous and Differential Staining of Plant Tissues[END_REF], coupled to an adapted image workflow [START_REF] Legland | Histological quantification of maize stem sections from FASGA-stained images[END_REF] allows a quantification of the histology of maize based on lignin-containing tissues detection but does not permit a strict quantification of cell wall lignin content.

Other approaches based on micro-vibrational spectroscopy (IR and Raman) were successfully adapted to plant biology and more particularly to in situ cell wall characterization [START_REF] Gierlinger | Revealing changes in molecular composition of plant cell walls on the micron-level by Raman mapping and vertex component analysis (VCA)[END_REF][START_REF] Agarwal | Raman imaging to investigate ultrastructure and composition of plant cell walls: distribution of lignin and cellulose in black spruce wood (Picea mariana)[END_REF][START_REF] Agarwal | FT-Raman Spectroscopy of Wood: Identifying Contributions of Lignin and Carbohydrate Polymers in the Spectrum of Black Spruce (Picea mariana)[END_REF][START_REF] Gierlinger | New insights into plant cell walls by vibrational microspectroscopy[END_REF]. They allow scientists to obtain qualitative, quantitative and spatial data on lignin. However, microspectroscopy approaches are generally very difficult to calibrate in terms of acquisition analysis and are also time consuming for both machine use

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5 and data analyses. Overall, such methodology is not adapted for the rapid measurement of lignin content in a tissue/sample compared to a reference and can therefore represent an inconvenience for functional genomics approaches that can involve the analyses of large numbers of samples (e.g. mutant populations, biological replicates).

In 2008, Bond and co-authors [START_REF] Bond | Safranine fluorescent staining of wood cell walls[END_REF] demonstrated that the azo dye safranin-O, commonly used with a counterstain to color lignified cell walls red in botanical samples examined by transmitted light in the visible spectrum, also presents an interesting fluorescence characteristic. Using a range of different plant samples made up of cells with high or low lignin levels these authors showed that the safranin-O emission spectrum depends on lignin quantitythe higher the lignin content of the stained tissue, the more intense the red part of the emission spectrum. While providing a novel way to distinguish lignin-rich from cellulose-rich cell walls without the need for counterstaining, their approach remained qualitative and did not allow for statistical comparison of lignin levels in different samples, We therefore decided to develop a technique that would i) generate high definition spatial information about lignin localization, ii) provide quantitative information that can be analyzed to support statistically significant differences in lignin content between samples, and iii) be fast, easy and low-risk for the experimenter.

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RESULTS

Safranin emission spectra depends on lignin content

Bond and co-authors [START_REF] Bond | Safranine fluorescent staining of wood cell walls[END_REF] demonstrated that the safranin emission spectrum changes according to cell wall lignin content. Higher lignin amount induces a relative increase in the red part of the emission spectrum. This effect can be clearly seen in the safranin-O spectra (under 488 nm excitation) obtained from different cell walls (zones 1-4) in a stem cross-section from a 2-month-old flax plant (Fig. 1). We used flax as an appropriate model plant to develop the safranin-based methodology since the stem of this species contains two populations of cells with highly contrasted secondary cell wall lignification profiles: xylem cells with a highly lignified secondary walls and cortical bast fibers with poorly lignified secondary cell walls [START_REF] Day | Lignification in the flax stem: evidence for an unusual lignin in bast fibers[END_REF]. This allowed us to observe different levels of cell wall lignification within the same histological section as shown with the toluidine blue staining in Fig. 1a.

Analysis of the Safranin fluorescence emission spectra under 488 nm excitation of different flax stem cell types (Fig. 1b) indicated that the maximum peak intensity is 570 nm regardless of cell type analyzed.

Spectral analysis shows that the more lignified the cell wall is, the higher the emission spectrum in the orange and red region (570 -600 nm) (Fig. 1b). In contrast, the "green" part of the spectrum (520 -560 nm) remains stable indicating that it is the orange/red part of the spectrum that discriminates between the different cell walls analyzed. Beyond 600 nm, the variability is lost.

Based on these observations, we decided to develop a ratiometric emission spectrum approach for evaluating cell wall lignification levels that would be robust, both within a histological section and between histological sections from different provenances (e.g. species, genotype, age).

Method development: from sample preparation to image treatment.

We firstly developed a standardized methodology designed to limit time consumption and risk to the manipulator without altering result quality (Method S1 and Fig. S1). The aim of the first (facultative)

step with EtOH alcohol is to fix and conserve samples (up to several weeks in 70 % EtOH) for later analysis as well as to limit the autofluorescence of pigments (e.g. chlorophyll) that can interfere with

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7 safranin fluorescent emission. The time necessary for this step must be defined depending upon the considered sample but an overnight incubation is usually suitable for most samples. As described later, it is important to have flat cross-sections and, in this case, samples are sectioned with a vibroslicer as thinely as possible (30-100µm) to obtain the best morphology. However, since the methodology is based on epifluorescence the thickness of the cut will not unduly influence image acquisition. Next, a 10 min staining step in safranin-O alcoholic solution (0.2 % in 50 % EtOH) is performed at room temperature in the dark with shaking. This step is followed by a short rinse with 50% EtOH to remove safranin excess on the tube/plate, a 15 min rinse with 50% EtOH and 2 x 10 min H2O rinses under shaking in the dark to remove excess safranin. Samples are subsequently mounted in distilled water and analyzed with a confocal microscope.

Before starting any comparative confocal analyses, it is important to obtain safranin spectra of lignified and non-lignified cell walls from the plant sample so as to determine invariable "green" channel and variable "red" channel values. Then, for each series of observations on a given sample, it is necessary to calibrate laser intensity and time acquisition. The more these two values are low, the less there will be a risk of safranin spectra shift as discussed later. Since our methodology is based on fluorescence quantification, the chosen laser intensity and time acquisition must provide enough fluorescence intensity per pixel whilst avoiding saturation. Finally, it is mandatory to select a flat area of the sample for observation.

Spectrum acquisition provides images of the type presented in Fig. 2a, b and c. The corresponding image file is then treated with the developed ImageJ macro. This macro applies a region of interest (ROI) selection, a noise threshold subtraction, a 32-bit conversion and a median radius filter. Next, it defines the highest intensity value, applies a threshold and converts the background to NaN. At this moment there are two possibilities for the user: leave the automatic threshold selection, or manually correct this threshold so as to make it more coherent with your knowledge of your sample morphology. Finally, the ratio calculation is made and a ratiometric image is generated as presented in Fig 2c.

Default settings allow the user to obtain an image representation on a color scale from purple to red in a ratio range from 0 (purple) to 2 (red). In our context, black pixels correspond to NaN pixels without

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ratiometric value (under the threshold, not considered in measurement even if selected). Purple pixels correspond to pixels with ratiometric value close to 0. In the case where the default range does not lead to useful image representation, it can be restricted or enlarged (Fig. S2). It is important to be careful at this stage since a valid comparison of different images requires that they are all built using the same range.

In addition to the ratiometric image, the macro also allows a quantitative representation. For this, the user selects the ROI on the ratiometric image and makes a measurement with image J (Fig. 2d). The same method is then applied to further ROIs allowing statistical analysis.

Biological validation of the method in Arabidopsis

In order to test the robustness of our technique and its capacity to generate biologically relevant information on cell wall lignin content we applied it to different model plant species. We first evaluated the ratiometric technique in different lignin mutants from Arabidopsis thaliana. The lignin biosynthetic pathway has been extensively studied in this species over the last 20 years and modifications in lignin content/structure have been well characterized in a number of different mutants making them ideal test objects for our technique. Our results (Fig. 3) show that the ratiometric values of fiber cell walls in cinnamyl alcohol dehydrogenase (CAD) and caffeic acid O-methyl transferase (COMT) (Fig. 3b,e) mutants are not significantly different from those of WT plants (Fig. 3a,d). While both mutants contain similar amounts of lignin to WT plants, cad mutants possess lignin with a higher aldehyde content and comt mutants show an altered S/G lignin monomer ratio [START_REF] Van Acker | Lignin biosynthesis perturbations affect secondary cell wall composition and saccharification yield in Arabidopsis thaliana[END_REF]. The fact that the ratiometric values remain unchanged suggests that our safranin ratiometric technique is insensitive to differences in lignin structures. In contrast, similar analyses on the f5h mutant (Fig. 3c) revealed a significant increase in the red/green ratiometric value indicating higher lignin content (9 %) in agreement with previous lignin chemical analyses [START_REF] Van Acker | Lignin biosynthesis perturbations affect secondary cell wall composition and saccharification yield in Arabidopsis thaliana[END_REF]. Taken together, these results indicate that the safranin ratiometric technique reflects cell wall lignin content but is not affected by lignin composition making it a powerful in situ analytical method for lignin quantitative analysis.

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As another example we investigated the lac4-2 lac17 double mutant previously characterized by Berthet and co-authors [START_REF] Berthet | Disruption of LACCASE4 and 17 results in tissue-specific alterations to lignification of Arabidopsis thaliana stems[END_REF]. Destructive wet chemistry analysis (acetyl bromide) indicated that this mutant contains 40 % less lignin in floral stems as compared to WT plants and shows weaker staining of interfascicular fibers with the Weisner reagent. The application of our technique to floral stem cross-sections from this mutant enabled us to both confirm previous observations and obtain more detailed information about the effect of the mutation on the cell wall phenotype (Fig. 4). Firstly, mutant interfascicular fiber cell walls are thickened, but have less well-organized inner secondary cell walls in comparison with WT sections (Fig. 4a andb). Secondly, lignification appears to be restricted to the middle lamella and primary cell wall of mutants compared to WT fibers that also possess lignified secondary cell walls. In addition to these observations that confirm the results obtained by Berthet et al., our method also allowed us to obtain quantification on the amount of lignin reduction in fiber cell walls.

Examination of average ratiometric values (Fig. 4c) indicated that mutant fiber lignin content was reduced by about 50 % compared to WT cell walls. These values are higher than the reduction in Klason lignin values (30 -40 %) determined by Berthet and co-workers but were obtained from analysis of the whole stem that contains both interfascicular fibers and fiber bundles.

The differences between the values obtained by safranin and wet chemistry of whole stems could suggest that the reduction in cell wall lignin is higher in fibers compared to vascular bundles and we therefore decided to analyze cell walls in the latter region. The analysis of cell walls in vascular bundles (Fig. 4d -f) show that there is no significant difference in mean cell wall lignin levels between WT and mutant plants when overall mean values for the bundle are taken into account. This result suggests that the 30-40 % reduction in Klason lignin values reported by Berthet and coworkers is due to a proportionally greater reduction in fibers compared to vascular bundles and indicates that the lac4-2 lac17 double mutation affects the lignification process in different ways depending upon the tissue considered.

The safranin ratiometric approach also allowed us to obtain new information about the effect of the mutation at the cell/cell wall layer level. For example, when the maximum (and not the mean) ratio values obtained from the lignified zones of WT fibers (middle lamella, primary cell wall, secondary cell wall) and mutant fibers (middle lamella, primary cell wall) are considered (Fig. 4c) the values indicate

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10 that lignin levels in middle lamella and primary cell walls are comparable. Taken together these results

indicate that the lac4-2 lac17 mutation primarily affects secondary cell wall lignification (but not middle lamella/primary cell wall lignification) and is associated with increased secondary cell wall thicknesspotentially in an attempt to compensate the loss of rigidity due to reduced lignification. The ratiometric approach also generates intriguing information about the effect of the mutation on the lignification process in vessels. Indeed, our results would suggest that the down-regulation of the Lac4 and Lac17

genes is not associated with a decrease in overall vascular bundle lignin content, but is, in fact, associated with increased lignification in xylem vessel cell walls and decreased lignification in fiber cell walls within the vascular bundle. Interestingly, mutant vascular bundles appear to contain a higher proportion of xylem fibers with poorly-lignified walls thereby underlining the existence of cell specific lignification programs.

In conclusion, the use of our safranin-based imaging approach allowed us to obtain both quantitative and precise spatial information about cell wall lignin content in a single experiment and on the same sample thereby providing a considerable advantage over chemical analysis (quantitative data, but loss of spatial information) or other histochemical reagents such as Weisner (spatial information, but no quantitative data).

Biological validation of the method in other plant species

The applicability of our method was used to investigate cell wall lignin in the flax lbf1 mutant that is characterized by the presence of bast fibers with a secondary cell wall lignin content of around 17 % compared to 5 % in WT plants [START_REF] Chantreau | Ectopic lignification in the flax lignified bast fiber1 mutant stem is associated with tissue-specific modifications in gene expression and cell wall composition[END_REF]. These values were obtained by destructive wet chemistry (acetyl bromide) and mask a strong spatial heterogeneity. As shown in Figure 5, the safranin ratio of bast fibers varies from 0 to 7 unit (safranin ratio mean of 1.2 unit corresponding to 2.4 times more lignin signal than WT) in a first developmental stage of lignification in 4-week-old plants (Fig. 6b andd) and from 0 to 15 (safranin ratio mean of 2.2; 4.4 times more lignin signal than WT) in a more advanced developmental stage of lignification (fig. 5c andd).

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Lignin content was also evaluated in the cell walls of maize, a monocotyledon plant. On stem cross sections various tissues with different lignin levels were observed with a 4x objective (Fig. 6a). Nonlignified parenchyma cell walls appear in black i.e. under the detection threshold of our methodology.

Hypodermis cell walls are colored in blue indicating low lignin levels (safranin ratio = 0.53) in comparison with the more heavily lignified cell walls of the epidermis (safranin ratio =0.93) and vascular bundle sheath that appear red (safranin ratio = 0.98) (Fig. 6a). This approach is complementary to an approach based on the bioinformatic treatment of FASGA stained stem sections that allows genotyping based on the surface of lignification [START_REF] Legland | Histological quantification of maize stem sections from FASGA-stained images[END_REF]. The safranin ratio methodology also allows a better resolution as it is independent of the section thickness that can become a limiting point when the cell density and therefore the opacity to the transmitted light are too important, as it occurs in the vicinity of large metaxylem vessels (Figure 6b). Sclerenchyma cell walls (red) are more lignified than vessels (blue to yellow) or phloem cell walls (purple, blue) (Figure 6b). Also, the development and the associated lignification appears polarized with 5 to 6 lignified cell layers on the protoxylem side against 1 to 3 cell layers on the phloem side. Altogether, these results confirm that the safranin approach can be easily applied to both monocots and dicots and provides highly detailed ratiometric quantitative data on cell wall lignin levels in comparison with global wet chemistry analyses which are longer and more laborious to set up.

The finesse of analyzes beyond simple quantification.

The described safranin method is not only easy to use, but also provides a high resolution spatial information on cell wall lignin that can usually only be obtained by using more time-consuming and fastidious techniques such as light/SEM immunolocalisation or Raman spectroscopy [START_REF] Gierlinger | New insights into plant cell walls by vibrational microspectroscopy[END_REF][START_REF] Kiyoto | Immunolocalization of 8-5' and 8-8' linked structures of lignin in cell walls of Chamaecyparis obtusa using monoclonal antibodies[END_REF][START_REF] Day | Lignification in the flax stem: evidence for an unusual lignin in bast fibers[END_REF][START_REF] Simon | Lignification and Advances in Lignin Imaging in Plant Cell Walls[END_REF].

An example of high-resolution using the safranin technique is illustrated in Figure 7 showing the results of an analysis of flax stem xylem. In contrast to other classical histological analyzes (eg. phloroglucinol-HCl) that suggest a uniform lignification of xylem cell walls [START_REF] Day | Lignification in the flax stem: evidence for an unusual lignin in bast fibers[END_REF]Huis et al., 2012), our methodology indicates that periclinal cell walls often contain higher amounts of lignin than anticlinal cell walls (Fig. 7a). Similarly, examination of a pit between two fiber tracheids at 60x magnification

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12 (Fig. 7b) provides an image quality and detailed information that approaches that obtained by electron microscopy [START_REF] Fromm | Lignin distribution in wood cell walls determined by TEM and backscattered SEM techniques[END_REF]. The obtained image reveals the localized high lignin levels in the vicinity of the pit as well as in the compound middle lamella between the two cells.

Technical controlslimits to be respected.

As in most imaging approaches, it is important to control different technical aspects that can affect the acquired signal and may lead to the generation of artifacts and false interpretations.

A first parameter is related to sample thickness and the focal plane at which observations are made. The lower the focal plane (i.e. the deeper within the section that one goes), the more incident laser energy and the resulting emission energy will be decreased. The extent to which this occurs is likely to be different depending upon the wavelength considered. To characterize this phenomenon in our methodology, we analyzed green and red channel fluorescence intensities of fiber cell walls at different focal planes in a section of safranin-O stained poplar xylem (Fig. 8). Our results show that the fluorescence intensity of the two channels is identical for the first 9 optical sections (upper to lower) and increases from around 70% of maximum intensity in the first optical section to 100% at the 9 th optical section. From the tenth optical section onwards, signal intensity decreases for the 2 channels but to a greater extent for the green channel compared to the red channel. These results indicate that the good benchmark for undertaking analysis is when the sample appears focused in the green channel.

An additional requirement resulting from this phenomenon is that the sample prepared is flat. As shown in the Figure 8, an uneven sample when analyzed at a given z-depth will give a different result depending upon the tissue thickness crossed. In Figure 8b, at the surface of the sample section, only the xylem appears in blue (Fig. 8b,i) whereas deeper in the section, other tissues appear first in blue and then in yellow and the initial xylem which appears in yellow and then redder (Fig 8b, ii to iv).

A second parameter concerns the fluorophore's resistance to bleaching. Since the technique depends on a ratiometric approach, the absolute intensity level does not matter, and it is more important to check the ratiometric stability. To evaluate this parameter, we subjected images of a 3-month-old poplar stem cortical fiber stained by safranin-O to increasing (intensity and accumulation time per pixel) excitation 13 (Fig. 9). For this, we exposed the sample at a laser intensity of 5% (Fig. 9a-c) and 100% (Figure 9d-f) on continuous acquisition (20s / acquisition -512x512 pixels image). At 5% laser intensity (Fig. 9a-c), differences in the percentage red/green ratio between the first and the fifteenth passage were of the order of 4%, whereas this value increased to around 35 % when 100% laser intensity was used (Figure 9d-f, g) resulting in a very different visual image depending upon total (intensity and time) laser energy used (Fig. 9h). Such a result indicates that excessive excitation of safranin affects its spectral properties and provokes a photoinduced spectral shift in favor of green wavelengths. Analyses should therefore be performed at low (power and/or time) excitation levels in order to keep any potential changes in the red/green ratio to a minimum.

DISCUSSION

Lignin is the third most abundant biopolymer on Earth and plays a vital role in many aspects of plant life (1). The presence and structure of this polymer also have an important impact on the quality of a wide range of different plant-based resources and is therefore of major economic importance. Together with other cell wall polymers, lignin also represents a major sink for atmospheric carbon and its biosynthesis therefore contributes to counteracting climate change [START_REF] Bastin | The global tree restoration potential[END_REF]. It is therefore not surprising that scientists have developed a wide range of different techniques for studying this important polymer.

Many different chemical and physical techniques are available to quantify and structurally characterize lignin in plant cell walls [START_REF] Simon | Lignification and Advances in Lignin Imaging in Plant Cell Walls[END_REF] and have been successfully used on both highly lignified tissues from woody plants and on more weakly lignified samples such as those found in herbaceous species.

However, many of these dosage methods are destructive and do not take into account the variability in lignin quantity and structure that occurs between different cell types and/or cell wall layers. As a result, the potential role of genes in the regulation of lignin biosynthesis and deposition at the cellular level is often ignored or simplified. This aspect is particularly important given recent reports demonstrating that

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14 the deposition of lignin in the cell wall is either under the spatial control of particular proteins localized in the cell wall [START_REF] Yi Chou | Distribution, mobility, and anchoring of lignin-related oxidative enzymes in Arabidopsis secondary cell walls[END_REF][START_REF] Hosmani | Dirigent domain-containing protein is part of the machinery required for formation of the lignin-based Casparian strip in the root[END_REF][START_REF] Tobimatsu | Lignin polymerization: how do plants manage the chemistry so well?[END_REF].

In contrast to destructive quantitative techniques, a number of in situ non-destructive techniques including classic light microscopy histochemistry, UV microscopy, TEM immunolocalisation and Raman spectroscopy can also be used to analyse cell wall lignin [START_REF] Simon | Lignification and Advances in Lignin Imaging in Plant Cell Walls[END_REF].

Of these, it is perhaps the Maüle and Wiesner staining methods that are the most widely used light microscope approaches for analysing lignin in situ [START_REF] Monties | Lignins[END_REF]. The Wiesner reagent (phloroglucinol-HCl) colours lignified cell walls in red and the Maüle reaction brings information about the S/G ratio. Both of these methods are relatively rapid, easy to use and accessible to most labs that possess a standard light microscope. However, they do not provide quantitative data thereby preventing statistical comparisons of cell wall lignin levels, either within the same sample, or between different samples. In addition, care should also be taken with the Wiesner reagent as it only highlights the aldehyde groups present in the polymer. In this paper we developed a simple and rapid confocal fluorescence imaging approach based on safranin-O histochemistry and that is able to provide both detailed spatial information and quantitative data on cell wall lignin levels.

Safranin-O is a basic azo dye that binds to the acidic lignin polymer (it is believed to form pi-pi interactions with the polymer [START_REF] Stockert | Histochemical mechanism for the orthochromatic staining and fluorescence reaction of lignified tissues[END_REF]. A comparison of safranin-O ratiometric images and statistical analysis of derived quantitative data indicated statistically significant differences in agreement with previous chemical analysis thereby confirming that the safranin-O method can be used to analyze cell wall lignin levels in situ [START_REF] Van Acker | Lignin biosynthesis perturbations affect secondary cell wall composition and saccharification yield in Arabidopsis thaliana[END_REF]. Similar analyses of Arabidopsis cad and comt mutants indicated no statistically significant differences in cell wall lignin levels also in agreement with previous chemical analysis ( [START_REF] Van Acker | Lignin biosynthesis perturbations affect secondary cell wall composition and saccharification yield in Arabidopsis thaliana[END_REF]. Although lignin content is unchanged in these two mutants, they do show important modifications in lignin structure (cad: increased G units, decreased S units, increased S-unit aldehydes;

comt: almost total elimination of S units, severely decreased S/G ratio). The fact that these differences did not affect the ratiometric images strongly suggests that the safranin-O based method is insensitive to such changes in lignin structure. As such, it represents a powerful complement to Wiesner staining (sensitive to aldehyde content) and Maüle staining (sensitive to S unit content) for characterizing lignin.

C O N F I D E N T I A L 16
Another important advantage of our method is that it enables quantification for individual cell walls/cell wall layers thereby enabling a much more precise understanding of the effects of mutations on the lignification process in plants. This point was demonstrated by our analysis of the lac4-2 lac17 double mutant previously characterized by . Comparison of ratiometric images from WT and mutant plants enabled us to observe and demonstrate the existence of statisticallysignificant changes in cell wall lignin levels at both the tissue (xylem vs interfascicular fiber) and cell (vessels vs xylem fiber) levels. Somewhat intriguingly, these analyses also suggested that vessel lignin levels were higher in mutants compared to WT plants.

CONCLUSION

The rapidly increasing availability of relatively low-cost and powerful confocal microscopes is providing scientists with the possibility to revisit old histochemical methods and to develop more efficient, safer, faster and more reproducible approaches, with a better spatial resolution. In this paper we have presented the combination of safranin-O staining coupled to confocal microscopy and an automated image treatment with the free software ImageJ that allows researchers to obtain quantitative data on cell wall lignin content very precisely and to undertake statistically valid comparisons between different cells, either within the same section or between different samples. The toolbox (method plus provided Image J macro) we describe in this paper will create a field of opportunities for modern histology in lignin imaging. It can be used to improve and complete the existing classical staining procedures and other approaches and will contribute to a better understanding of how lignification is regulated at the cellular level.

Page 16 of 38

SUBMITTED MANUSCRIPT The Plant Journal C O N F I D E N T I A L EXPERIMENTAL PROCEDURES

Plant material and growth conditions

For this study, 4 model plants were used: Arabidopsis thaliana, Linum usitatissimum (flax), Populus tremula x P. alba (poplar) and Zea mays (maize). For Arabidopsis thaliana, wild-type (Col-0, N60000), and selected 'lignin' mutant lines (comt1, cad6, f5h1 [START_REF] Vanholme | A Systems Biology View of Responses to Lignin Biosynthesis Perturbations in Arabidopsis[W][END_REF] and lac4-2 lac17 double mutant) were used [START_REF] Berthet | Disruption of LACCASE4 and 17 results in tissue-specific alterations to lignification of Arabidopsis thaliana stems[END_REF]. For L. usitatissimum, the Diane fiber variety (WT and lbf1 mutant) was used [START_REF] Chantreau | Ectopic lignification in the flax lignified bast fiber1 mutant stem is associated with tissue-specific modifications in gene expression and cell wall composition[END_REF]. For P. tremula x alba, the 717-1B4 WT line (INRA clone), was used. For Z. mays, an indeterminate forage genotype was used. A. thaliana, L. usitatissimum and Z.

mays were grown in soil using a 16h/8h photoperiod, 23°C/20°C temperature, 35-65% hygrometry in a greenhouse with natural light completed with sodium to maintain light intensity at 100 µmol.m -2 .s -1 . A.

thaliana plants were harvested at stage 6. Populus plants were grown in soil for 2.5 months (60 cm high) in a phytotron with a 16h/8h photoperiod, 24°C/21°C temperature, 35-65% hygrometry with white light (HPI Master Plus Philips, metal halid) to maintain light intensity at 120 µmol.m -2 .s -1 .

Instruments

Plant stem cross-sections -80 µm -were made using Microm HV640 and Leica S100 vibroslicers. All the image acquisitions were performed on a LSM710 Confocal (Zeiss) equipped with Argon laser and 541 nm laser and a Nikon A1R confocal equipped with Argon laser and 561 nm laser.

Image J -Fiji macro step development

An Image J -Fiji macro was developed by successive automated steps with possible user intervention at a key point. The steps of the process are as follows : // split channels; // selection of region of interest and subtraction of background noise; // 32 bit conversion and apply median filter radius 1; // set the green image that has the highest intensity value; // apply a threshold and convert the background to NaN;

// wait for user to define threshold; // calculate the ratio (plugin ratioplus) ; // copy aaa_lut_fret.lut in the lut field of the Fiji install and run; // Set ratio range; // Measures.

Macro downloading address: https://nextcloud.univ-lille.fr/index.php/s/rKDBBEM3fznRa6Z

C O N F I D E N T I A L C O N F I D E N T I A L C O N F I D E N T I A L C O N F I D E N T I A L

SAFRANIN-O SOLUTION:

• Ethanol [final 50% (w/v)],

• Safranin-O [final 0.2% (w/v)]

• Ultrapure water to the final volume Mix the solution well for at least 15min until you get everything completely dissolved.

PREPARATION OF THE SOLUTION AND ITS STORAGE

• Safranin-O is purchased as a powder and dissolved in ethanol 50% solution.

• To prepare the staining solutions, weigh the indicated amount of the corresponding dye powder, dissolve it in ethanol 50% solution by mixing the solution on the magnetic stirrer in a small flask for at least 15min.

• To store the solutions, wrap them in aluminium foil to avoid exposing them to light at 4°C.

Note 1: if possible prefer fresh safranin-O solution.

Note 2: for long term conservation, a 10x solution can be prepared, split in aliquots and keep several months in the dark at -20°C.

SAMPLE TREATMENT

• Collect samples, maximum 0.5 cm length.

• Place them in 70% EtOH overnight with shaking at 4°C to fix and remove pigments that can impact the fluorescence acquisition.

• Remove alcohol solution. • Coat samples in moult with agarose solution (3 to 4%).

• Perform section -ideally from 30µm to 100µm -on a vibroslicer and place them on EtOH 50% solution until staining. For this step and depending on your samples and the vibroslicer you must adapt section parameters (speed, frequency, amplitude section) to obtain the best morphology. Better the samples morphology is kept better will be the results.

Note: the same procedure can be envisaged with other embedding as paraffin or PEG to use microtome instead of vibroslicer to obtain better morphology. In this case the global procedure will be longer. However, the gain in morphological quality is justified only for fragile tissues.

SAFRANIN-O STAINING

• Prepare fresh 0.2% Safranin-O.

• Place vibrosliced samples in safranine-O solution for 10min under shacking at RT at obscurity.

• Remove all safranine-O solution.

• Rinse one time in ethanol 50% solution for 10min under shacking at RT at obscurity.

• Remove all the solution.

• Rinse two time in ultrapure water for 15min under shacking at RT at obscurity.

• Mount samples in water for imaging.

Note 1: For the staining and for rinsing it's important to have a big volume of solution compared to sample number and to incubate in container allowing good homogenization under shaking.

Note 2: Adapt the flushing time if necessary. Young lignified tissues or tissues with low lignin content don't support too high decolouring time.

Note 3: Never mount your samples in glycerol or glycerol-containing mounting medium because safranin-O is highly soluble in glycerol.

CONFOCAL ANALYSIS

For the first analysis: • Perform a comparative spectra analysis between lignified and non-lignified tissue. This step is essential to determine the peak of maximum intensity, the invariable area of the spectrum (green part, generally between 520nm and 560 nm) and the area of the spectrum sensitive to lignin quantity (yellow to red part, generally between the maximum intensity peak and 630 nm).

• For the spectral analysis use a 488nm excitation coupled with acquisition from 500nm to 700nm. Warning: limit 488 nm laser power and pixel time exposition to avoid spectra alteration as presented in this study.

Current analysis:

• To optimize results and limiting 488nm laser using to avoid spectral shift, the acquisition was developed with 2 independent track with two different laser line excitation.

Track 1-green channel -ex: 488nm, em: 530-560 nmlimit 488 nm laser power (1-5% on recent laser).

Track 2-red channel -ex: 561nm, em: 570-600 nm.

• Considering the semi-quantitative comparison between independent samples, settings of the reference sample must limit light range acquisition to 70% of the maximum tones (about 2900 tones for 12 bit image acquisition).

• Acquire sequentially both tracks.

Note: Acquisition parameters depend on the confocal microscope used. The important point is to well define the green and the red channel and adapt the parameters in function of.

IMAGE TREATMENT

• For the image treatment use the Fiji install completed with the developed macro available to this address: (https://nextcloud.univlille.fr/index.php/s/rKDBBEM3fznRa6Z).

• Open the Fiji app

• Open the macro file with Fiji.

• Open the image.

• Run the macro and follow the instructions.

The result is a ratiometric image (representing ratio range from 0 to 2). Note 2: Two parameters of the macro can be adjusted depending on the acquisition.

-The "rooling" must be adapted according to image resolution and size of the analysed biological structure which vary with the objective used and the zoom used for the acquisition.

-The ratio range representation is set from 0 to 2 (purple to red). If the variation is more or less important, the ratiometric range can be adjusted to optimize representation. In this case, the same range must be applied to all image included in the comparison. Modify the range don't modify raw pixel values.

  Using a range of plant samples characterized by the presence of cells showing different amounts of cell wall lignin, Bond and co-workers demonstrated that cell walls/cell wall regions containing high lignin levels (e.g. cell corners of pine wood) exhibited red fluorescence in safranin-O-stained samples observed by confocal microscopy, while non-lignified cell walls (cotton fibers, poplar wood G-layers) fluoresced green. This same study also demonstrated that the red fluorescence in safranin-O stained samples was spatially correlated with high lignin autofluorescence in non-stained samples confirming the correlation between lignin quantity and red fluorescence.Since a number of different factors can affect the intensity of acquired signals, we firstly undertook a number of experiments to standardize sample preparation and analysis. Our results indicated that sample depth and laser intensity should all be carefully controlled to avoid the generation of artifacts during image acquisition and facilitate valid comparative studies. We also demonstrated that a simple fixation in 70 % EtOH either before or after section preparation was compatible with the technique thereby facilitating sample collection and processing.The development of a novel macro in the widely used ImageJ software enabled the generation of a ratiometric image and the subsequent quantification of relevant cell wall lignin levels in selected areas.It should be emphasized that the ratiometric images and derived quantitative data do not represent absolute values such as those obtained by Klason or acetyl bromide analyses (e.g. lignin level expressed as a % of dry cell wall residue). Moreover, since safranin fluorescence can be modulated by a number of different factors (dye concentration, incubation and washing time, sample thickness and observation depth, photobleaching) it is extremely difficult to establish absolute correlation curves. Nevertheless, the technique does provide quantitative values that allow statistically valid comparisons to be madeeither between different cell walls/cell wall layers in the same sample, or indeed between cell walls/cell wall layers from different samples (e.g. mutant vs WT plants). This point was demonstrated by our analysis of the previously characterized Arabidopsis f5h lignin mutant that has increased lignin content.

Figure 1 .Figure 2 .Figure 3 .

 123 Figure 1. Cell wall associated safranin spectra. a) Flax stem cross-section stained with toluidine blue

Figure 4 .

 4 Figure 4. Biological validation of the method in Arabidopsis lac4-2 lac17 mutant. From a) to b) are

Figure 5 .Figure 6 .

 56 Figure 5. Biological validation of the method in other plant species: flax. From a) to c) are represented

Figure 7 .Figure 8 .

 78 Figure 7. Flax xylem cell walls: the finesse of the method. a) Safranin ratiometric images of xylem cross

Figure 9 .

 9 Figure 9. Effects of laser energy (intensity and time) on cell wall images and green/red ratio. The effects

  if the macro is completely automated, the user has the possibility to act on the threshold applied to image treatment. Let the automatic threshold choice except if you consider that the morphology doesn't correspond to your scientific knowledge of the model studied.

ACKNOWLEDGEMENTS

We thank the TisBio facilities and the FRABio research federation for providing the technical environment conducive to achieving this work. We thank Fabienne Guillon for her critical reading of the manuscript. This study was funded by grants from the French Region Hauts de France and the EU (CPER/FEDER project ALIBIOTECH) to F.B.-C. and by the Fonds de la Recherche Scientifique (FRS-FNRS) research project T.0004.14 to F.B.-C and M.B. We thank Prof. Wout Boerjan for providing the A.thaliana lines used in this study. We sincerely thank all those who believed in this project, trusted us and supported us in our approach. Thanks to Dr. Nathalie Bessodes without whom this project could not have been possible. I dedicate this work to my daughters Laura and Rosalie.

Method resume

The complete method is described in the supplementary file Methods S1. A resume scheme of the method is presented in the supplementary figure S1.

SUPPORTING INFORMATION LEGENDS


Bruno Alies

Mohamed A Ouelhazi

Amit Patwa

Julien Verget

Laurence Navailles

Valérie Desvergnes

Philippe Barthélémy

Cytidine-and guanosine-based nucleotide-lipids

Hybrid nucleotide-lipids composed of a lipid covalently attached to purine and pyrimidine nucleobases exhibit supramolecular properties. The novel cytidine and guanosine derivatives are promising bioinspired materials, which can act as supramolecular gelators depending on both the nucleobase and the presence of salts. These supramolecular properties are of broad interest for biomedical applications.

Introduction

Supramolecular materials derived from biological structures such as peptides, 1,2 carbohydrates, 3 or nucleosides 4 are attractive for various applications, 5,6 including green chemistry, 7,8 materials science, drug delivery, which are biocompatible structures composed of a lipid covalently attached to a nucleotide, are desirable candidates for the implementation of supramolecular biomaterials as they easily engage noncovalent interactions. 18,19 Nucleotides-based lipids, in particular, are unique in that they have multiple possibilities for non-covalent interactions depending on the nature of the bases and can self-associate into stable different type of selfassembly, such as ribbons, bilayers, fibers etc. In previous studies, we reported the self-assembly properties of several nucleotide lipids (NLs) featuring 3'-monophosphate thymidine or adenosine head groups covalently attached to 1,2dipalmitoyl-sn-glycerol. 20,21,22 Interestingly, biocompatible hydrogels, resulting from the self-assembly of thymidine-3'-(1,2-dipalmitoyl-sn-glycero-3-phosphate) (diC16-3′-dT) salts, were found to be effective soft materials for in vivo injection and implantation. 23 The modulation of the polar head structure, in particular the nature of the base could influence the aggregation properties of the NLs. Therefore, we elected to design NLs with a double dipalmitoyl glycerol phosphate linked to the 3'secondary hydroxyl of the four natural nucleosides. Indeed, the diverse hydrogen bond donor-acceptor sites in all four nucleobases (A: Adenine, T: Thymine C: Cytosine, and G: Guanine) are prone to form self-base pairing interactions that stabilize supramolecular self-assemblies. 24,25 It is therefore valuable to explore the self-assembly properties of A, C, T and G based nucleotide-lipids as supramolecular building blocks. Specifically, we report the synthesis and the physicochemical studies of a series of nucleoside-3'-monophosphate NLs featuring palmitic double chains and A, T, C, G nucleobases. Several examples of nucleoside-based amphiphiles have been reported, 26,27,28 however to the best of our knowledge, there are no previous examples of Cytosine and Guanosine nucleoside-3′-monophosphate amphiphiles.

Results and discussion

In this study, we prepared four purine and pyrimidine nucleotide-lipids via a phosphoramidite strategy. The supramolecular assemblies obtained with A, T, C, G nucleotide lipids either in the absence or presence of sodium chloride are reported (Fig. 1). Depending on both the conditions and the nature of the base the stabilisation of supramolecular hydrogels was observed. Likewise, as a proof of concept, in vitro drug release experiments show that a controlled delivery of a drug can be obtained thanks to the structure of the nucleolipid based gels.

Several strategies have been used to synthesize nucleotide lipids, including coupling reactions of monoalkyl phosphate with nucleoside and enzyme catalysis, 29 or via phosphoramidite synthetic pathway. The novel NLs, diC16-3′-dG 1a and diC16-3′-dC 1b, were synthesized using a similar approach to that used for the synthesis of diC16-3'-LNA-A 30 (see ESI, Scheme S1). This approach uses standard procedures, routinely adopted in oligonucleotide synthesis via phosphoramidite chemistry in the solid phase and adapted to solution. Briefly, commercially available isobutyryl protected guanosine phosphoramidite 2a (isobutyryl-dG-CE phosphoramidite, see SI, Scheme S1) and acetyl protected cytosine phosphoramidite 2b (Ac-dC-CE phosphoramidite, see SI, Scheme S1) were deprotected with methylamine solution in THF to achieve dG-CE phosphoramidite 3a and dC-CE phosphoramidite 3b, respectively. These phosphoramidites (3a and 3b) were coupled with 1,2-dipalmitoyl-sn-glycerol in the presence of a tetrazole solution in acetonitrile (weakly acidic condition) to provide intermediate coupling product. The resulting phosphite intermediates were oxidized, with iodine solution in THF/pyridine/water mixture, to obtain cyanoethyl protected phosphate intermediate. After deprotection of the dimethoxytrityl (5'-DMTr) group under acidic conditions (trichloroacetic acid in DCM), the removal of the cyanoethyl chain was achieved under basic conditions (triethylamine) to Please do not adjust margins Please do not adjust margins yield diC16-3′-dG 1a and diC16-3′-dC 1b as their triethylammonium salts (for detailed synthetic procedure, see SI, Scheme S1). The NLs featuring thymidine (diC16-3′-dT 1c) 31 and adenosinne (diC16-3′-dA 1d), 32 were synthesized according to the literature procedure.

Spectroscopy studies

In order to study the inter-nucleobase interactions in the NLs self-assemblies UV-visible and Circular Dichroism experiments were performed. Figure 2 show the UV spectra of each NLs in aqueous (dashed line) or NaCl solution (dotted line) with their correspondent deoxynucleosides (solid line). It is well known that UV spectra of aromatic rings are sensitive to π-π interaction, when two or more rings are mutually close they give rise to hypochromic effect. A decrease of about 20 % in the absorbance values was observed for diC16-3'-dA, diC16-3'-dT and diC16-3'-dG compared with deoxyadenosine, thymidine and deoxyguanosine, respectively. In the case of diC16-3'-dC, no significant decrease in absorbance and slight bathochromic shift (5 nm) were observed compared to deoxycytidine. In order to get further structural information, circular dichroism (CD) of NLs and deoxynucleosides was also investigated (Fig. 2, right scale). Compared to deoxyadenosine (solid line), the CD spectra of diC16-3'-dA exhibited more pronounced peaks at 225 and 280 nm and the appearance of a band at 240 nm. With diC16-3'-dT, the band at 270nm is similar to thymidine. Preparation in NaCl (red dotted line) intensified the band. DiC16-3'-dG exhibited the appearance of two new peaks, a positive band at 240 nm and a negative band at 260 nm in addition to the peak at 225 nm observed in both CD spectra. With purine NLs (diC16-3'-dA & diC16-3'-dG) large differences are seen between NLs and deoxynucleosides (Fig. 2 dashed vs solid lines). In NaCl solutions, the diC16-3'-dC CD band at 266nm exhibited a substantial shift and increase compared to diC16-3'-dC prepared in water solution (dashed green vs dotted green lines). These new bands were indicative of the formation of chiral structures. It is hypothesized that in the case of well-hydrated alkali metal salts electrostatic repulsions between the negatively charged phosphate would occur, leading to the formation of low curvature aggregates. In turn, less hydrated salts would result in higher curvature assemblies due to the stronger interaction among nucleosides. In this direction, we compared the UV and the CD spectra of NLs in pure water and in pseudo-physiological conditions (NaCl: 154 mM equivalent to 0.9% w/v). Except for diC16-3'-dG and diC16-3'-dA, which were not soluble in pseudophysiological conditions, diC16-3'-dT and diC16-3'-dC did not show any significant change in their UV spectra. Whereas, in CD spectroscopy diC16-3'-dT exhibited a slight increase in CD intensity but for diC16-3'-dC, a large increase in CD intensity was clearly observed with a red-shift to higher wavelength (Fig. 2, bottom right, dashed vs dotted line). A different type of structure resulting from intermolecular nucleobase interactions may explain this abrupt change. These results indicate that in addition to the hydrophobic effect through the dipalmitoyl double chains, inter-stacking and H-bonding between nucleobases plays a major role in stabilizing NLs aggregates. In addition, the counter ion used in these systems strongly impact on the supramolecular organizations of diC16-dT and diC16-3'-dC.

Taken together, UV-vis and CD experiments show that the diC16-3'-dX series form supramolecular assemblies, which can be modulated in the presence of NaCl.

Salt effects on self-assemblies

We further investigated the supramolecular changes of NLs either in the presence or absence of NaCl. Dynamic Light Scattering (DLS) measurements were collected both in water and in NaCl conditions. The mean diameters and the polydispersity index (PDI) of the assemblies are reported as well as the size distributions graphs (Table 1 down andFigure ESI 1). In a first series of experiments, NLs were dispersed in pure water under sonication. Results obtained by DLS measurements indicated that sonication was not suitable for a good dispersion of pyrimidines NLs (diC16-3'-dC and diC16-3'-dT) giving a highly polydisperse size distribution. In contrast, for purine NLs (diC16-3'-dG and diC16-3'-dA), a single distribution was observed with a mean diameter at around 220 nm for both purine NLs with a relatively low PDI (0.23). The dispersion of NLs was also investigated in the absence of sonication. Surprisingly, in this condition, opposite behaviours are observed. Indeed, pyrimidine NLs (diC16-3'-dC and diC16-3'-dT) show a single distribution whereas purine based NLs (diC16-3'-dG and diC16-3'-dA) were not well dispersed as observed by DLS. These differences in behaviour illustrate the importance of the nucleobase in the dispersion of NLs. In NaCl conditions, size distributions of diC16-3'-dC and diC16-3'-dT (only these NLs were soluble in these conditions) showed the presence of a large size distribution with the appearance of larger structures ranging between 180 to 1000 nm, supporting the hypothesis that fibers formations occurred in the presence of well-hydrated alkali metal. The results of the DLS experiments indicated that the size of the supramolecular structures depends on the nature of the NL nucleobase. In the case of diC16-3'-dG and diC16-3-dA, bearing purine nucleobases, higher sizes of the supramolecular assemblies were observed with a tendency to form one size distribution compared with NLs possessing pyrimidine nucleobases.

To characterize the thermal reversibility of the hydrogels, Differential Scanning Calorimetry (DSC) experiments were performed with a total of three heating and cooling cycles in pure water and in NaCl conditions (Fig. 3). Heating and cooling cycles results from four independent runs of thermal analysis are reported in Table 1 (up). DSC experiments showed main phases transitions (Tm) at various temperatures depending of nucleobases and conditions (Fig 3). In water, transition occurs (heating/cooling) at 15/14, 36/32, 20/19 and 35/27°C for diC16-3'-dA, diC16-3'-dT, diC16-3'-dG, and diC16-3'-dC, respectively. The Tm observed for NLs are similar to that for the natural di palmitoyl-glycerol phosphocholine (DPPC) as well as synthetic di palmitoyl uridine phosphocholine (DPUPC). 33 A clear hysteresis between heating and cooling cycle was observed for diC16-3'-dC (8°C) and diC16-3'-dT (4°C), whereas only 1°C difference occurred with diC16-3'-dG and diC16-3'-dA (See Fig 3 and Table 1 up). DiC16-3'-dG and diC16-3'-dA exhibited other small transitions. In NaCl, the main phase transition temperatures increased for all NLs and reached 39/37, 40/38, 42/39 and 42/40°C for diC16-3'-dA, diC16-3'-dT, diC16-3'-dG, and diC16-3'-dC, respectively without any additional peaks. Interestingly, a lower hysteresis between heating and cooling cycle was observed with diC16-3'-dC and diC16-3'-dT (1°C) in NaCl conditions. Note that for diC16-3'-dC and diC16-3'-dT in water, broad peaks in heating and cooling cycle were detected; this tendency did not appear in NaCl conditions. In This main phase transition disparity between nucleobases (purine-pyrimidine) disappeared in NaCl conditions where all NLs reached approximately the same thermal reversibility at around 40°C. This observation indicates that the counter ion (Na + ) possess a stronger impact on the melting temperature of purine derivatives (diC16-3'-dA and diC16-3'-dG) than pyrimidine analogues (diC16-3'-dC and diC16-3'-dT). Although purines bases were able to give hydrogen bonding, in water condition, we observed that the transition temperatures of diC16-3'-dA and diC16-3'-dG were dramatically lower than pyrimidine NLs. This could be explained by a higher steric hindrance with purine NLs causing difficulties to self-organise.

In the presence of a well-hydrated cation (Na + ), more stable assemblies could be reached suggesting that Na + would stabilize the supramolecular structure.

Rheology and gel behaviors

In order to further characterize the physicochemical properties of NLs at a macroscopic level, we performed gel formation experiments (Fig. 1B). Depending of the conditions (water or NaCl), striking differences were observed. In presence of NaCl, a strong diC16-3'-dC gel was formed (Fig. 1B left), whereas diC16-3'-dG precipitated without forming any gels. Following these qualitative tests, the mechanical properties of new cytidine and guanosine nucleotide-lipids based gels were analyzed using oscillatory rheology to characterize the Linear Visco-Elastic Regime (LVER) and the strength of the gels. Figure 4 shows the oscillatory strain sweeps for cytidine and guanosine nucleotide-lipids based gels at 5% and 10%, respectively. The LVER, in which the moduli are independent of the applied strain, of the new compounds were determined up to 100%. For all hydrogels investigated, the elastic G' consistently exceeds the viscous G" moduli, indicating the dominant elastic character of the nucleotidelipid based Low Molecular Weight Gelators (LMWGs). In the case of diC16-3'-dC gels, the LVER was determined either in the absence or presence of NaCl (0.9% w/v). The breaking points measured by moduli inversion were found at 1 and 5% and the G' moduli of 600 Pa and 2400 Pa, respectively, indicating that the presence of NaCl strongly impacts on the diC16-3'-dC gels rheological properties. For diC16-3'-dG, due to the lack of solubility in the presence of NaCl, the LVER was determined only in water at 10%. At this concentration, the G derivative shows similar properties to the diC16-3'-dC at 5% in water (G'= 800 Pa, moduli inversion at 5%). For diC16-3'-dT and diC16-3'-dC in NaCl conditions, the gel-sol transition occurred instantaneously. Note that in pure water an interstate transition between the gel to solution was observed. This gel-solution transition difference was consistent with DSC results where broad peaks were observed in water conditions contrary to sharps peaks obtained in NaCl conditions (Fig. 3, right panels). Such behaviour could explain the broad peaks obtained in heating and cooling and the large hysteresis obtained for diC16-3'-dC and diC16-3'-dT in pure water. These results also indicated that the phase transition behaviour depends mainly on the nucleobases type, the selfbinding affinity between nucleobases and the environmental conditions, notably, the counter ion used.

Drug release experiments

Due to their unique properties, supramolecular gels are currently under investigation as drug delivery matrix. Controlling the kinetics of drug release from such soft materials via the molecular structure of gelators is highly desirable since it would allow for a modulation of drug release depending on the external conditions. To illustrate the usefulness of the novel nucleolipids in this field, in-vitro drug diffusion studies were carried out using diazepam as a drug. Diazepam is a hydrophobic compound belonging to the benzodiazepine family, which are used as an anticonvulsant in the treatment of epilepsy. The data of percentage drug release are shown in Fig. 5. For the kinetic study following plots were made: cumulative % drug release versus time for both diC16-3'-dC and diC16-3'-dG in the presence or absence of NaCl. As a general comment, a slower release is observed for both nucleolipids in the absence of NaCl (T 1/2 > 400 minutes for both compounds). After addition of NaCl, a faster release of diazepam was observed for diC16-3'-dG (T 1/2 = 20 min) compared to diC16-3'-dC (T 1/2 = 400 min), indicating that the nature of the base is playing a major role in the release mechanism from the supramolecular gels. Interestingly, these results demonstrate that the kinetic of the drug release can be modulated via both the nature of the base and the presence of salts.

Conclusion

In summary, a facile route for the synthesis of nucleoside-3'-monophosphate featuring the four natural nucleobases has been reported. Similarly, to thymidine and adenosine 3'nucleotide lipids previously published, both Cytosine and Guanosine derivatives reported in this contribution exhibit self-assembly properties. Taking together, the physico-Fig. 4. Oscillatory strain sweeps for cytidine and guanosine nucleotide-lipids based gels at 5% and 10%, respectively. Elastic, (G', red) and viscous (G", blue) moduli versus strain in percent. Left) LVER of diC16-3'-dC (5%, w/v) in water (solid lines) or with 0.9% NaCl (symbols). Right) LVER of diC16-3'-dG (10%, w/v) in water.

chemical studies indicate that the self-assembly properties depend strongly on both the conditions and the nature of nucleobases.

The presence of NaCl induced NLs-NLs interactions, but the specific supramolecular structures achieved give rise to different states at the macrocopic level depending on the nucleobase. The control of the lipid selfassemblies via the molecular code of the four natural nucleotides represent a powerful strategy, which could be used to construct biocompatible soft materials such as gels. As illustrated with the drug release experiments the kinetic of release can be controlled by both the nature of the nucleobase and the presence of salts. Their nucleic acid controlled selfassembly is likely to greatly extend the potential of lipid-based materials for biomedical applications, including drug delivery.

Experimental Section General Procedures for synthesis

All the compounds were purchased from Sigma-Aldrich, Fluka and Alfa Aesar unless otherwise mentioned. Solvents for reactions were purchased from Sigma-Aldrich in the highest quality and from VWR for other uses. All the reactions were run under nitrogen atmosphere unless otherwise stated. Analytical thin layer chromatography (TLC) was performed on pre-coated silica gel F254 plates with fluorescent indicator from Merck. The detection of compounds was accomplished using a UV light (254 nm) and visualized on TLC plates by subrsequent spraying with 10 % conc. H 2 SO 4 solution in ethanol, followed by heating. Column chromatography was performed with flash silica gel (0.04-0.063 mm) from Merck. All the compounds were characterized using 1 H, ). The coupling constants J are given in Hertz (Hz); the peak multiplicity is reported as The coupling constants J are given in Hertz (Hz); the peak multiplicity is reported as follows: s = singlet, brs = broad singlet, d = doublet, t = triplet, m = multiplet. High resolution electronspray ionization mass spectra (HR ESI-MS) were performed by the CESAMO (Bordeaux, France) on a QSsat Elite mass spectrometer (applied Biosystems). The instrument is equipped with an ESI source and spectra were recorded in negative mode. The electrospray needle was maintained at 4500 V and operated at room temperature. Samples were introduced by introduced by injection through a 10 μL sample loop into a 200 μL/min flow of methanol from the LC pump. For more details, see ESI.

Preparation mode of diC16-3'-dX

DiC16-3'-dA, diC16-3'-dT, diC16-3'-dC, diC16-3'-dG were synthesized by our laboratory (see details in ESI). The simple procedure was followed for all compounds. Generic name of diC16-dX is given to specificity either DiC16-3'-dA, diC16-3'-dT, diC16-3'-dC or diC16-3'-dG. Exact mass around 6 mg of diC16-dX was weighted on 0.1 mg precision balance and dissolved and mixed in dichloromethane into 10mL volumetric flask. Aliquots were then prepared by transferring 500µL into glass vial. Aliquots were then evaporated until dryness and stored at -20°C. Hydration of the lipid film is accomplished simply by adding 3 mL mΩ water or 3mL of a NaCl solution (0.9% w/v; 154mM) The obtained solutions were then heated for 30 minutes and sonicate for 5 min at temperature of 60°C. Finally, the resulting solutions were directly assessed by either Dynamic Light Scattering, CD or UV-Vis Spectroscopy.

CD / UV-Vis Spectroscopy

UV-Vis and CD spectra of diC16-3'-dX was performed on JASCO J-1500 Circular Dichroism (optical path length=1cm) in quartz cuvette coupled with a Peltier device. All measurements were carry out at 25°C.

Differential Scanning Calorimetry

Differential Scanning Calorimetry (DSC) was realized using METTLER TOLEDO. A total of three heating and cooling cycles (5-65°C) on 2% w/w in pure water and in NaCl conditions at a constant rate of 0.5°C min -1 and 100µL aluminium crucible. Heating and cooling results of thermal analysis are reported in Table 1.

Dynamic Light Scattering

Dynamic Light Scattering (DLS) measurements were performed using Zetasizer 3000 HAS MALVERN at 25°C with 400 µL of a solution containing 100 µM of diC16dX prepared as explained above. Measurements of Table 1 down have been performed in water with 5 minutes sonication for purine nucleolipids (diC16-3'-dG and diC16-3'-dA) and without sonication for pyrimidines nucleolipids (diC16-3'-dC and diC16-3'-dT).

Gel Formation

Water or a NaCl solution (0.9% w/v ;154mM) were added to diC16-3'-dC or diC16-3'-dG to obtain a solution at 2% w/v of diC16-3'-dX. Then the solution was heated at 60°C during 30 minutes and sonicate the 5 last minutes. The solutions were allowed to cool down at room temperature and gel formation were observed Please do not adjust margins Please do not adjust margins (diC16-3'-dC water / NaCl & diC16-3'-dG / water) or not (diC16-3'-dG NaCl). See Figure 1B.

Drug release experiments 6.0 mg of either diC16-3'-dG and diC16-3'-dC were accurately weighted in 3 mL UV cuvette. 200 µL of diazepam solution at 500 µM are added in the cuvette. Cuvette is sealed and heated at 60°C for 30 min then to cool down at room temperature for 30 min. Appearance of gel is visually checked as solution become opalescent and viscous. Cuvette is set in a Jasco V630 spectrophotometer and UV spectra (400 nm to 200 nm) is recorded every 2 min after careful addition (without disturbing the gel at the bottom of the cuvette) of 1600 µL of either water or 154 mM NaCl solution. In this setting, the beam of the spectrophotometer pass thought the bulk (above the gel) allowing to follow the kinetic of diazepam release. Absorbance at 313 nm (specific for diazepam, epsilon = 1700 cm -1

.M -1

) was subtracted from baseline at 350 nm and converted into percentage.

Rheology experiments

Rheological measurements were carried out on a Malvern Kinexus Pro+ rheometer with steel coneplate geometry (diameter: 40mm, angle: 1°). The lower plate is equipped with a Peltier temperature control system, and all samples were studied at 25 ± 0.01°C unless indicated otherwise. A solvent trap was used to ensure homogeneous temperature and to prevent water evaporation. The hydrogels were heated at 65°C and the resulting liquid was placed into the rheometer and subjected to sinusoidal oscillations. The experimental conditions to determine the linear visco elastic regime (LVER) were determined by performing an amplitude strain sweep from 0.01 to 100% at an angular frequency of 1Hz (6.283rad.s -1

).

Conflicts of interest and acknowledgments

There are no conflicts to declare. This work was realized under the frame of the Laboratory of Excellence AMADEus with the reference ANR-10-LABX-0042-AMADEUS, operated by the "Agence Nationale de la Recherche" under the program "Initiative for Excellence IdEx Bordeaux" (ANR-10-IDEX-0003-02). The work was also supported by INSERM and CNRS. Aurore Beaurepaire, Candice Delcourt, Alexandra Gaubert are gratefully acknowledged for their help with circular dichroism, UV-vis and rheology experiments, respectively. The authors thank the BIC Center for technical assistance during TEM observations and Marc Biran from the CRMSB.

Notes and references

Fig. 1 .

 1 Fig. 1. A) Chemical structure of the nucleotide-lipid (NLs). B) Nucleotide lipid derivatives were incubated either in the absence or presence of sodium chloride. The novel C and G nucleotide lipid aqueous suspensions exhibit different macroscopic behaviors, the cytosine NL stabilizes hydrogels in the presence or NaCl, whereas the guanosine derivative forms a gel in the absence of NaCl.

Fig. 2 .

 2 Fig.2. UV (scale on left axis) and CD (scale on right axis) spectra of NLs prepared in water (dashed line), in 0.9% w/v NaCl solution (diamond) with their deoxynucleosides (solid line). The nature of the nucleic bases is indicated on the graphs. DiC16-3'-dG and diC16-3'-dC are not soluble in NaCl solution therefore their spectra are not displayed. All spectra were recorded at 100 μM in room temperature.

Tables 1 .

 1 Up) Transition temperatures by DSC of NLs obtained by DSC. The results reported are the average of four independent experiments. ± symbol indicated the standard deviation. ΔT(NaCl -Water) emphasize the increase of transition temperatures in the presence of NaCl (0.9% w/v). Down) Z-average, diameters and polydispersity index obtained by DLS for each diC16-3'-dX.

Fig. 3 .

 3 Fig. 3. DSC experiments of diC16-3′-dA (top left blue), diC16-3′-dT (top right red), diC16-3′-G (bottom left black) & diC16-3′-C (bottom right green) in the absence (solid lines) or presence of 0.9% (w/v) NaCl (dotted lines). Cooling (positive signal) and heating (negative signal) cycle are displayed. Conditions are arbitrary separated and only one cycle is shown for clarity reason.

13 C and 31 P 1 H 31 P

 1331131 Nuclear Magnetic Resonance (NMR) spectroscopy. These NMR spectra were recorded (either in CDCl 3 obtained from Eurisotop) on BRUKER Avance DPX-300 ( at 121.49 MHz). The chemical shifts (δ) are given in parts per million (ppm) relatively residual solvent peaks (CHCl 3 : 1 H: 7.26, 13 C: 77.16

Fig. 5 .

 5 Fig. 5. In vitro drug release experiments showing the cumulative release of diazepam from the gel (% drug release) versus time (in minutes). Kinetics of release for diC16-3'-dC (left) and diC16-3'-dG (right) with the addition of water or sodium chloride solution (NaCl 0.9% w/v).

Transition Temperature (°C) Heating Cycle ΔT(NaCl -Water) diC16-3'-dX Water NaCl

	diC16-3'-dA	14.2 ±0.2	38.3 ±0.4	24
	diC16-3'-dT	40.0 ±3.6	41.0 ±1.5	1
	diC16-3'-dG	19.8 ±0.1	40.9 ±0.8	21
	diC16-3'-dC	35.4 ±0.6	41.5 ±0.8	6
	diC16-3'-dX	Water	Cooling Cycle	NaCl	ΔT(NaCl -Water)
	diC16-3'-dA	13.8 ±0.1	35.3 ±1.5	22
	diC16-3'-dT	34.6 ±6.0	38.5 ±0.3	4
	diC16-3'-dG	18.6 ±0.1	38.4 ±0.3	20
	diC16-3'-dC	27.5 ±2.4	40.1 ±1.1	13
		Z-average (nm) / Particles diameters (nm)	PolyDispersity Index
	diC16-3'-dX			With sonication
	diC16-3'-dG			186 / 238		0.232
	diC16-3'-dA			199 / 220		0.234
		Without sonication
	diC16-3'-dC			99 / 105		0.089
	diC16-3'-dT			81 / 85		0.193