# Exploring the influence of wild silks composition on dyeing with turmeric and indigo: spectroscopic insights

Nathan Ferrandin-Schoffel, Sophie Cersoy, Sophie Desrosiers, Annabel Vallard

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Nathan Ferrandin-Schoffel, Sophie Cersoy, Sophie Desrosiers, Annabel Vallard. Exploring the influence of wild silks composition on dyeing with turmeric and indigo: spectroscopic insights. Spectrochimica Acta Part A: Molecular and Biomolecular Spectroscopy [1994-..], In press, 10.1016/j.saa.2025.126019 . hal-04983661

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# Exploring the influence of wild silks composition on dyeing with turmeric and indigo: spectroscopic insights

Nathan Ferrandin-Schoffel<fnref n="1" />,<fnref n="2" />,<fnref n="3" />,* , Sophie Cersoy<fnref n="1" />, Sophie Desrosiers<fnref n="4" />, Annabel Vallard<fnref n="2" />

**Keywords** Cocoon; Degumming; Sericin; Calcium oxalate; FTIR; UV-Vis

## Abstract

"Wild" silks, produced by various silkworm species, are often valued in textile production, but present greater challenges in dyeing compared to the "domestic" mulberry silk from *Bombyx mori*. This study investigates the reasons behind these challenges, focusing on key components such as fibroin, sericin proteins, and calcium oxalate crystals. The degumming process, which removes sericin, generally enhances dye penetration; however, its efficiency varies with the type of silk and its chemical composition. Using Fourier-transform infrared (FTIR), Raman spectroscopy, and UV-Visible spectroscopy, we characterized the composition of cocoon samples from 11 different silkworms, both with and without a preliminary degumming step, and assessed their dyeing behavior with turmeric and indigo dyes. FTIR provided insights into the molecular structure and physico-chemical interactions within the silk fibers, while UV-Visible spectroscopy quantified dye sorption. Our findings demonstrate that effective degumming is crucial for dye uptake, with greater degumming loss often leading to a higher and more homogenous dye sorption. In some cases, the degumming procedure did not completely remove sericin, the latter of which not only physically impedes dye penetration into the silk mass but also forms physicochemical interactions with the dye, influencing the overall sorption capacity. Additionally, the presence of calcium oxalate crystals - particularly in wild silks like *Antheraea paphia* - can further impede dye sorption. Corroborating images obtained using optical digital microscopy supported these findings.

<footnote n="1">¹Centre de Recherche sur la Conservation (CRC), Muséum national d'Histoire naturelle, CNRS, ministère de la Culture, 36 rue Geoffroy-Saint-Hilaire 75005 Paris, France</footnote>

<footnote n="2">²Centre Asie du Sud-Est (CASE, UMR 8170), EHESS, 2 Cours des Humanités 93300 Aubervilliers, France</footnote>

<footnote n="3">³(Present address) Sciences et Ingénierie de la Matière Molle (SIMM, UMR 7615), ESPCI Paris - PSL, 10 rue Vauquelin 75005 Paris, France</footnote>

<footnote n="4">⁴Centre de Recherches Historiques (CRH, UMR 8558), EHESS, 54 boulevard Raspail 75006 Paris, France</footnote>

<footnote>*Corresponding author: nathan.ferrandinschoffel@lacambre.be</footnote>
# 1. Introduction

Silkworm silk is a natural protein fiber, with its earliest recorded use in textile manufacturing dating back approximately 6000 years in China [1]. The primary species used in the silk industry is the mulberry silkworm *Bombyx mori* (Linnaeus, 1758), which is native to Asia and has been domesticated there. However, other species of silkworms, often referred to as wild or non-mulberry, comprising in particular Lepidoptera from the Saturniidae family, have been used by local craftspeople worldwide since at least the 3rd millennium BCE [2]. These include, among others, *Antheraea yamamai* (Guérin-Méneville, 1855), *Antheraea paphia* (Linnaeus, 1758), *Antheraea pernyi* (Guérin-Méneville, 1855), *Antheraea assamensis* (Helfer, 1837), *Samia cynthia* (Drury, 1773), *Samia ricini* (Jones, 1791), *Borocera madagascariensis* (Boisduval, 1833), *Attacus atlas* (Linnaeus, 1758), *Cricula trifenestrata* (Helfer, 1837) and *Anaphe panda* (Boisduval, 1847). Among these species, attempts were made in the 19th century to introduce *Antheraea pernyi* and *Samia cynthia* into Western industrial silk textiles production [2]. This introduction was driven not only by the need to replace domestic silkworms affected by pebrine, a disease that devastated silkworm farms in Europe, but also by the desire to offer new types of materials to the international luxury market. These silks, each differing biochemically [3–5], exhibit unique characteristics compared to *Bombyx mori* silk, including differences in color, luster, texture, mechanical strength and elasticity [6,7].

Throughout history and across various silk-producing regions, dyers often cite the complexity of dyeing wild silks [8]. Figure 1 illustrates the differences in color rendering in a Japanese silk crepe dyed after weaving, which comprises both mulberry and non-mulberry silk filaments. In the late 19th century, Thomas Wardle (1831-1909), a British textile entrepreneur associated with William Morris and the Arts & Crafts movement [9,10], reported from his extensive experimental studies conducted in India that dyeing "tussar" silks (a generic term used to refer to silks produced by various *Antheraea* species from the Saturniidae family) results in a coloration at least twice as weak compared to domestic silk [6]. Wardle hypothesized that Indian tasar silk (*Antheraea paphia* specifically) was more difficult to dye than mulberry silk due to its lower solubility, as he conducted experiments with zinc chloride solutions [6]. Despite his pioneering work [6,11–13], the physicochemical causes behind these observations have not been thoroughly investigated, and they remain inadequately understood.
<figure><img src="image_2.png" /><figcaption>Figure 1. Silk crepe with resist dyeing post-weaving (Japan, 19th century), composed of fine filaments from *Bombyx mori* (domestic silk), which absorb dye well, and flat, thicker filaments from a wild silk (presumably *Antheraea yamamai*), which remain undyed. The bat patterns appear white due to the resist dyeing. Fabric detail (top) and two magnifications (bottom): x50 (left) and x250 (right). © S. Desrosiers</figcaption></figure>

The hypothesis proposed for this research is that the differences in color rendering after dyeing are attributable to the varying compositions of silk cocoons. While all silks are primarily composed of proteins, specifically fibroin and sericin, their amino acid sequences [4,5] and crystallinity [14–16] vary among species. Additionally, the quantities of calcium oxalate, which can be abundant in some wild cocoons and are absent in *Bombyx mori*, also differ [15,17]. Calcium oxalate crystals, which can take the form of cubes or needle-like raphides [15,17,18], are presumed to not be directly biosynthesized by silkworms. Instead, their presence in silk is attributed to the silkworm's diet and the excretion of minerals
during cocoon spinning [15]. These crystals are present in many plants, where they serve functions in calcium regulation, plant protection, and metal detoxification [19]. Hypothetically, these crystals could make the degumming process (partial or total) more difficult, as they may limit the porosity of cocoon and the diffusion of solvents [20], and/or they may chemically bind to the sericin proteins removed during this process. Given that adhesive sericin surrounds the structural fibroin proteins and binds them together [21], the presence of calcium oxalate crystals could reduce the permeability of the silk, thereby inhibiting the attachment of dyes within the fibrous network. To test this hypothesis, it is crucial to investigate the impact of cocoon degumming on its composition - particularly for wild silks, where this remains poorly understood - and ultimately assess how these factors influence dyeing effectiveness.

We investigated the impact of silk composition, with and without a preliminary degumming step, on dyeing efficiency. Several silk cocoons from different sericigenous species, originating from Asia and Madagascar, were collected and characterized. Degumming and dyeing protocols were applied under consistent conditions for a relevant comparison, using well-documented chemical compounds and dyes. The choice of dyes was limited to turmeric and indigo, mentioned in Wardle's writings [6,12], without a mordanting step, as studying mordant effects would require a separate investigation. The common and straightforward boiling-off in alkaline soap solution process [22–29] was used for degumming the silk cocoons. A buffer consisting of sodium bicarbonate (NaHCO₃) and sodium carbonate (Na₂CO₃), both commonly used in the alkali degumming process and often used in conjunction with soap [4,23,25,27,28,30,31], was introduced into the soap solution to maintain a constant pH across all samples, within an ideal range of 9.5-10.5. Indeed, a lower pH makes degumming less effective, while a higher pH can degrade the fibroin proteins of the silk [23,29]. The effectiveness of the dyeing step was studied by examining the silk samples using digital microscopy, as well as UV-Vis absorbance of the dye baths to estimate the amount of dye adsorbed/absorbed onto/into the silk samples. Additionally, Raman and Fourier-transform infrared (FTIR) spectroscopies were employed to enable a comprehensive study of the silk components [15,32–34], including calcium oxalate [15,34], with the aim of characterizing the physicochemical interactions formed with the dye molecules.

## 2. Experimental

### 2.1. Materials

#### 2.1.1. Silk samples

The collected silk cocoons are listed in Table 1. A commercial *B. mori* white silk thread (Artifilum) sourced from China was used as a reference degummed sample. Photographs of the cocoons
and the thread is provided in the Supporting information (Figure S1). As shown in the photograph, the cocoons differ notably in color and size. Additionally, upon handling, some cocoons, such as those of *A. paphia* and *A. atlas*, are significantly stiffer, while others, like those of *A. assamensis* and the uniquely net-like cocoons of *C. andrei* and *C. trifenestrata*, are much softer. Notably, the *A. atlas* cocoon stands out not only for its impressive size but also for being considerably thicker than the other cocoons (thickness not measured).

<table><caption>Table 1. Natures and origins of the silk cocoons studied.</caption><thead><tr><th>Species</th><th>Family</th><th>Designation</th><th>Historical origin [2,35]</th><th>Country of collection</th><th>Supplier</th></tr></thead><tbody><tr><td>Bombyx mori<br>(white and yellow cocoons)</td><td>Bombycidae</td><td>Mulberry</td><td>China<br>(Zhejiang)</td><td>Thailand</td><td>CNRS, Centre Asie du Sud-Est, France</td></tr><tr><td>Antheraea<br>yamamai</td><td>Saturniidae</td><td>Tensan</td><td>Japan</td><td>France</td><td>National Museum of Natural History (MNHN), France</td></tr><tr><td>Antheraea paphia<br>(or Antheraea mylitta)</td><td></td><td>Tasar</td><td>India</td><td>India</td><td>essiespinner.etsy.com</td></tr><tr><td>Antheraea pernyi</td><td></td><td>Tussah</td><td>China</td><td>France</td><td>National Museum of Natural History (MNHN), France</td></tr><tr><td>Antheraea<br>assamensis</td><td></td><td>Muga</td><td>India<br>(Assam)</td><td>India</td><td>essiespinner.etsy.com</td></tr><tr><td>Cricula<br>andrei</td><td></td><td>Cricula</td><td>South-East Asia</td><td>N/A</td><td>National Museum of Natural History (MNHN), France</td></tr><tr><td>Cricula<br>trifenestrata</td><td></td><td>Cricula</td><td>South-East Asia</td><td>N/A</td><td>National Museum of Natural History (MNHN), France</td></tr><tr><td>Samia cynthia<br>(advena)</td><td></td><td>Eri</td><td>United States of America</td><td>N/A</td><td>National Museum of Natural History (MNHN), France</td></tr><tr><td>Samia ricini<br>(white and red cocoons)</td><td></td><td>Eri</td><td>India</td><td>India</td><td>essiespinner.etsy.com</td></tr><tr><td>Attacus atlas</td><td></td><td>Fagara</td><td>South-East Asia</td><td>Thailand</td><td>Office pour les insectes et leur environnement (OPIE), France</td></tr><tr><td>Borocera<br>madagascariensis</td><td>Lasiocampidae</td><td>Landibe</td><td>Madagascar</td><td>Madagascar</td><td>Soies de Marie [36]</td></tr></tbody></table>

### 2.1.2. Dyes, soap and other chemicals

Fresh turmeric rhizomes were sourced from Thailand and obtained from the Thanh Binh Jeune grocery store (Paris, France). Synthetic indigo powder (VAT Blue 1) was purchased from Kremer. A well-characterized commercial solid soap [37] was used for degumming and washing the silks after dyeing. Sodium hydroxide was purchased from VWR Chemicals Prolabo, while sodium hydrosulfite,
sodium bicarbonate and calcium oxalate were sourced from Sigma-Aldrich. Sodium carbonate was purchased from Fluka analytical.

## 2.2. Degumming

For each species, a cocoon was cut into small pieces of a few millimeters and immersed in deionized water containing a 1.0 g/L 1:1 (wt/wt) NaHCO₃/Na₂CO₃ buffer and 25 wt% of soap, with a liquor ratio of 1:50, in 4 mL screw-top vials. The pH of the aqueous stock solution at room temperature was 9.93 ± 0.02 (measured with a Mettler Toledo MA235 pH-meter). The mixture was brought to a boil (95-100 °C) for 1 hour. The silk samples were then thoroughly rinsed with boiling deionized water (three times), followed by cold deionized water (three times). After wringing, the samples were air-dried overnight, gently heated at 40 °C in an oven for 1 hour, and then placed in an environmental chamber to equilibrate in the adsorption regime at 23 °C and 50% relative humidity (RH) for at least 24 hours before weighing. The degumming loss was calculated as follows (eq. 1):

$$ \text{Degumming loss (\%)} = \frac{w_b - w_a}{w_b} \times 100 \quad (\text{eq. 1}) $$

where w_b and w_a are the silk weights at 23 °C and 50% RH before and after the degumming process, respectively. Samples were weighed in triplicate, and the average degumming loss values are reported with the standard deviations.

## 2.3. Dyeing

The protocols, applied to cut small silk pieces of a few millimeters, were established based on training from the French textile artisan and dyer Marie Foyer [36], as well as a literature review [38-42]. For both turmeric and indigo dyes, all dyed samples were wrung out and washed with warm soapy deionized water. They were then soaked in warm deionized water (approximately 40°C) for 20 minutes before being wrung out again and left to air dry overnight.

### 2.3.1. Turmeric

A 50 g/L stock turmeric solution was prepared. Rhizomes were chopped into small pieces of around 1 cm, crushed, and placed in deionized water for maceration for 10 minutes at room temperature. The solution was brought to a boil (95°C – 100°C) for 1 hour and then filtered using a sieve. The pH of the solution at room temperature was 6.46 ± 0.02 (pH-meter Mettler Toledo MA235). Weighed at 23°C / 50% RH, the silk samples were dyed at 80 °C for 1 hour in 1.5 mL Eppendorf tubes, with a liquor ratio of 1:20.
### 2.3.2. Indigo

A 5.0 g/L stock indigo solution was prepared. 500 mg of indigo powder was allowed to macerate for 15 minutes in 100 mL of deionized water. 500 mg of sodium hydroxide was gently added, and the solution was heated to 60°C. Subsequently, 1.5 g of sodium hydrosulfite was added. The mixture was stirred and cooled to 40°C, resulting in a characteristic yellow-green solution with a sheen. The pH of the solution at room temperature was 8.07 ± 0.02 (pH-meter Mettler Toledo MA235). Weighed at 23°C / 50% RH, the silk samples were dyed at 40 °C for 20 min in 1.5 mL Eppendorf tubes, with a liquor ratio of 1:20.

## 2.4. Optical microscopy and photography

The samples were examined using a Keyence VHX-1000 digital microscope, illuminated in transmission or reflection, with 50×, 100× and 200× objectives. The diameters of individual fibroin filaments, which together form silk fibers, were measured using ImageJ software on images captured at 200× magnification. For each sample, ten measurements were averaged, and the uncertainties are reported as standard deviations. Photographs of the samples on a black background (Figures S1 and S2) were taken with a Nikon D800 camera.

## 2.5. Spectroscopy

### 2.5.1. FTIR

Fourier-transform infrared (FTIR) absorption spectra were recorded using a Nicolet 6700 spectrometer (Thermo) equipped with a diamond crystal attenuated total reflection (ATR) Smart Endurance module, scanning between 500 and 4000 cm⁻¹. A total of 128 scans were recorded for each spectrum, with a resolution of 4 cm⁻¹. Up to three spectra were obtained for each sample. Spectral decomposition into Gaussian components was performed using the OMNIC software. The average relative intensities I<sub>r</sub> (eq. 2, eq. 3 and eq. 5) are reported with the standard deviations.

### 2.5.2. Raman

Raman spectra of silk samples were recorded using a confocal inVia microspectrometer (Renishaw) equipped with a 785 nm laser source, covering an extended spectral range of 100 - 3200 cm⁻¹. For each spectrum, a 50× objective was used, with an exposure time of 10 seconds and 3 to 5 accumulations. The laser power at the objective ranged from 2.0 to 10.6 mW.
Additional Raman spectra were obtained using a 1064 nm laser source with a portable i-Raman EX microspectrometer (B&W TEK), spanning a spectral range of 100 - 2500 cm⁻¹. The use of the 1064 nm laser was necessary to overcome the excessive fluorescence observed in several silk samples and dyes when using the 785 nm laser. For these measurements, a 100× objective was employed, with an integration time of 10 to 30 seconds and 3 to 5 accumulations. The intrinsic laser power ranged from 45 to 450 mW.

#### 2.5.3. UV-Vis

The fresh aqueous turmeric dye solutions were centrifuged at 14,000 rpm for 2 minutes. The clear supernatants were analyzed by UV-Visible spectroscopy, using a 10 mm quartz cuvette. Spectra were immediately recorded using the Cary 5000 UV-Vis-NIR double-beam spectrophotometer over a wavelength range of 190 - 800 nm. Absorbance values were obtained with a 1 nm step size and a scan speed of 600 nm/min. Spectra were recorded in duplicate, and the average absorbance values are reported with the standard deviations. To prevent signal saturation, all turmeric dye solutions were diluted by a factor of 40 for UV-Vis analysis.

UV-Vis analysis was specifically performed for turmeric, as indigo suspensions were not clear or homogeneous, given that indigo is insoluble in water.

### 3. Results and Discussion

To investigate variations in silk composition resulting from degumming and dyeing, we conducted a spectroscopic characterization of the collected cocoon samples at three stages: first, on the raw cocoons; subsequently, after degumming; and finally, on the dyed samples, with or without prior degumming. The dyeing experiments with turmeric and indigo were performed on eight types of silk cocoons: white *B. mori*, yellow *B. mori*, white *S. ricini*, red *S. ricini*, *A. assamensis*, *A. paphia*, *B. madagascariensis*, and *A. atlas*. To provide additional data for a comprehensive understanding of the process, degumming was also applied to five additional cocoon types, available in smaller quantities: *A. pernyi*, *A. yamamai*, *S. cynthia*, *C. andrei*, and *C. trifenestrata*.

Due to limited sample availability, only one to two cocoons per species were used and characterized across all stages of the study (raw, degummed, and dyed), and further studies are required to investigate potential intra-species composition variations.
3.1. FTIR characterization of the silk cocoons

<figure><img src="image_3.png" /><figcaption>Figure 2. (a) FTIR spectra of the outer and inner walls of white *B. mori*, white *S. ricini*, and *B. madagascariensis* cocoons, as well as monohydrated calcium oxalate. (b) Focus on the 760 – 1440 cm⁻¹ region. I and II: absorption bands of amides I and II. O: absorption bands of calcium oxalate. (AG)n: absorption band of polyalanine-glycine β-sheets. (A)n: absorption band of polyalanine β-sheets.</figcaption></figure>

Figure 2 displays the FTIR spectra of the inner and outer walls of white *B. mori*, white *S. ricini*, and *B. madagascariensis* cocoons (spectra of all the other collected cocoons are provided in the Supporting information, Figure S3). The amide I (1620 cm⁻¹) and II (1515 cm⁻¹) bands, characteristic of proteins (fibroin and sericin) [15,43], are intense in the spectra of all cocoons (Figure 2a) except for the outer wall of *B. madagascariensis*, where only a weak amide II band is detected. This is due to a very intense band at 1605 cm⁻¹ that obscures the amide I band. This intense band corresponds to calcium oxalate, as confirmed by comparison with the reference FTIR spectrum of the salt (Figure 2a), indicating its high abundance on the outer wall of this cocoon. As evidenced by the presence of bands O at 1315
cm⁻¹ and 780 cm⁻¹ [15,34] (Figure 2b), the white *S. ricini* cocoon also contains calcium oxalate, though in smaller quantity, particularly on its outer wall. In accordance with descriptions in literature [15,17], *B. mori* lacks calcium oxalate.

An evident distinction between the FTIR spectra of *B. mori* silk and those of wild silks is the consistent presence of a band (A)ₙ at 965 cm⁻¹ in the latter (Figure 2b and Figure S3). This band is highlighted in Figure 2b for *S. ricini* and the inner wall of *B. madagascariensis* (where the outer wall's intense calcium oxalate signal largely obscures the silk signals) and is attributed to the polyalanine ((A)ₙ) sequences. These sequences correspond to the rocking deformation vibration of CH₂ and are characteristic the β-sheets of fibroin proteins in most wild silks [15,34,44]. The absence of this band in the FTIR spectrum of *B. mori* is due to its protein β-sheets being predominantly composed of polyalanine-glycine (AG)ₙ sequences [15,45] (band (AG)ₙ at 975 cm⁻¹, Figure 2b).

To allow a comparative analysis of the composition between the inner and outer walls of the cocoons and of the intensities of different bands, all FTIR spectra were decomposed. The spectral decomposition was conducted over the wavenumber range of 1280 - 1730 cm⁻¹, as illustrated in Figure 3, which shows the example of the outer wall spectrum of white *S. ricini*, with band assignments based on a comprehensive literature review [5,15,33,44,46,47]. A semi-quantitative analysis of calcium oxalate and the proportion of amorphous versus crystalline regions was performed by calculating the following relative intensities Iᵣ (eq. 2 and 3):

$$I_r(\text{CaC}_2\text{O}_4)(\%) = \frac{A(1315)}{A(\text{amide II})} \quad \text{and} \quad I_r(\text{am/cr}) = \frac{A(1635)}{A(1620)} \quad (\text{eq. 2 and 3})$$

where A(X) represents the area (absorbance) of the FTIR band at wavenumber X in cm⁻¹. The area of the band at 1315 cm⁻¹, corresponding to calcium oxalate (CaC₂O₄), was normalized to the amide II band (components at 1510 and 1520 cm⁻¹ in Figure 3) to allow for a relevant comparison among the silk samples. This normalization approach follows the methodology described by Gheysens et al. (2011) [17]. The amide I band was used to estimate the proportion of amorphous regions (1635 cm⁻¹) relative to the crystalline regions defined by the β-sheets (1620 cm⁻¹) in the silk proteins analyzed. Calculating Iᵣ (am/cr) offers an initial approximation of the proportion of sericin proteins relative to fibroin proteins, given that sericin is predominantly amorphous, whereas fibroin is semi-crystalline. Consequently, a higher Iᵣ (am/cr) value suggests a greater proportion of sericin. This remains an approximation, as the crystallinity index of fibroin proteins can differ among various silk cocoons, affecting the Iᵣ (am/cr) values. For instance, according to Vyas and Shukla (2016) [16], the crystallinity index of *B. mori* fibroin is higher (~ 75%) compared to that of *S. ricini* and *A. paphia* (~ 65%).
<figure><img src="image_4.png" /></figure>

<table><thead><tr><th>Position (cm<sup>-1</sup>)</th><th>Assignments</th><th>References</th></tr></thead><tbody><tr><td>1695</td><td>Amide I, β-turns</td><td>[5,15,33]</td></tr><tr><td>1635</td><td>Amide I, random coil</td><td>[5,15,33,47]</td></tr><tr><td>1620</td><td>Amide I, β-sheets</td><td>[5,15,33,47]</td></tr><tr><td>1520</td><td>Amide II, β-sheets and random coil</td><td>[15,33,47]</td></tr><tr><td>1510</td><td>Amide II, β-sheets and random coil</td><td>[15,33,47]</td></tr><tr><td>1445 *</td><td>δ<sub>as</sub> (CH<sub>3</sub>), δ (CH<sub>2</sub>)</td><td>[15,33,44,46,47]</td></tr><tr><td>1400 *</td><td>w (CH<sub>2</sub>), δ (CH<sub>3</sub>)</td><td>[15,33,44,47]</td></tr><tr><td>1370 *</td><td>δ<sub>s</sub> (CH<sub>3</sub>), δ (CH<sub>2</sub>)</td><td>[15,33,46,47]</td></tr><tr><td>1330 *</td><td>δ<sub>s</sub> (CH<sub>3</sub>), δ (CH)</td><td>[15,33,45–47]</td></tr><tr><td>1315</td><td>Calcium oxalate</td><td>[5,15]</td></tr></tbody></table>

Figure 3. Decomposition of the FTIR spectrum of the outer wall of the white *S. ricini* cocoon over the range 1280-1730 cm⁻¹ (linear baseline, Gaussian components), with band assignments. δ: bending; δ<sub>as</sub>: asymmetric bending; δ<sub>s</sub>: symmetric bending; w: wagging. *: FTIR bands with assignments that are debated in the literature cited.

Intensity I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) was calculated for all cocoons on both the inner and outer walls (Figure 4a), except for *C. andrei* and *C. trifenestrata* as their unique net-like structures do not allow distinguishing between the two distinct faces (see photographs on Figure S2). The results show that calcium oxalate is more abundant on the outer wall in all cases, with particularly high concentrations observed for *A. atlas*, *A. paphia* and *B. madagascariensis*, where it is 5 to 11 times more abundant compared to the inner walls. The outer wall of *B. madagascariensis* is especially notable, with a calcium oxalate relative intensity of 30%, the highest among all samples. This suggests that calcium oxalate is primarily excreted early in the cocoon formation process, as the outer layer is constructed before the inner layer by the silkworms. Conversely, some wild silk cocoons, such as *A. pernyi*, *S. cynthia*, *C. andrei*, and *C. trifenestrata*, contain either similar amounts or even less calcium oxalate compared to *B. mori*. The reason for the lack of calcium oxalate excretion in *B. mori*, despite its diet of mulberry leaves that contain this compound [48], remains unclear. The low or absent calcium oxalate excretion in these wild silk moths might also be attributed to genetic factors or other unknown reasons.
<figure><img src="image_5.png" /><figcaption>Figure 4. Relative intensities (a) I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) and (b) I<sub>r</sub> (am/cr), and (c) diameters of individual fibroin filaments on the inner and outer walls of the cocoons. The walls of *C. andrei* and *C. trifenestrata* are indistinguishable.</figcaption></figure>

White crystals are observed using optical microscopy on the outer surface of certain cocoons, particularly those of *B. madagascariensis* and *A. paphia*, where they are notably abundant (Figure 5b,c). These crystals were attributed to calcium oxalate, in agreement with the highest values of I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) determined by FTIR for these two cocoons (Figure 4a).
<figure><img src="image_6.png" /><figcaption>Figure 5. Digital optical microscope images of the (a-c) outer and (d-f) inner walls of white *B. mori*, *B. madagascariensis* and *A. paphia* panels (g-i) present the same samples after degumming, with (i) showing the outer wall of the *A. paphia* cocoon which does not become fluffy after degumming.</figcaption></figure>

Intensity $I_r$ (am/cr) was calculated for the same 13 cocoons, except for the outer walls of *A. atlas*, *A. paphia* and *B. madagascariensis* (Figure 4b). For these three samples, the proportion of calcium oxalate is such that its characteristic band at 1605 cm⁻¹ can significantly distort the calculation of the areas of the Gaussian components of the amide I band in the same frequency region (Figure S3a). Therefore, calculations were performed for all other samples, where $I_r$ (CaC₂O₄) ≤ 3.5%, an arbitrary threshold chosen to ensure that $I_r$ (am/cr) values would not be significantly impacted by the calcium oxalate interference. A key observation from this study is that, unlike $I_r$ (CaC₂O₄), a larger $I_r$ (am/cr) is not consistently observed on the outer wall of the cocoons. This indicates that sericin is not always found in a greater proportion on the outer wall, as is the case for *B. mori* [23]. For example, *A. assamensis* demonstrates a case where the $I_r$ (am/cr) value is more than five times higher on the inner wall compared to the outer wall. Additionally, some species, such as *B. mori* and *B. madagascariensis*, show significantly greater intensities than others, like *A. pernyi*, *A. yamamai*, and *S. cynthia*. This variation may be attributed to differences in the proportion of sericin present in the silk cocoons. Since sericin is removed, either partially or entirely, during the degumming process, this aspect is further explored in the “3.2. Degumming of the silk samples” section.
The diameter of fibroin filaments (also called fibroin “brins”), which constitute the silk fibers and are bound together by sericin (e.g., two fibroin filaments per silk fiber in *B. mori* [21]), was measured on both the inner and outer walls of each cocoon (Figure 4c). The filament diameter exhibits significant variation between cocoons, ranging from less than 15 µm (*B. mori* and *B. madagascariensis*) to over 32 µm (*A. paphia*), consistent with the literature [18]. Furthermore, for some cocoons, such as *B. mori* (white and yellow), *S. ricini* (white and red), and *A. assamensis*, the filament diameter is slightly wider on the outer wall compared to the inner wall. In contrast, no difference is observed in the other species, and an inverse trend is even seen for *A. paphia* and *A. yamamai*. One possible explanation for these differences could be the variation in sericin content between the walls (inner vs outer), with thicker filaments potentially corresponding to higher sericin content. However, there is no clear correlation between the relative intensity $I_r$ (am/cr) (Figure 4b), which has been associated with sericin content, and filament diameter (Figure 4c). Moreover, as discussed in section “3.2. Degumming of the silk samples”, the sericin content and its removal had minimal impact on the fibroin filament size observed under the digital microscope. Thus, the observed differences between the cocoons and their walls are likely due to inherent size variations between the fibroin filaments themselves.

## 3.2. Degumming of the silk samples

The impact of degumming on each cocoon's composition was investigated by measuring the mass loss and comparing the FTIR spectra of the silk samples before and after the process was applied.

### 3.2.1. Degumming loss

The degumming process significantly affects the mass loss and visual appearance of silk cocoons, varying widely across different species (Figure 6). The degumming loss ranged from 5% to 36%, reflecting the varying sericin content and the efficiency of its removal. *B. mori* cocoons exhibited a high degumming loss of approximately 32%, indicating a potentially complete extraction of sericin, aligning with typical measurements for this species [5,21,49,50]. Several wild silk cocoons, such as *A. yamamai*, *B. madagascariensis*, *C. andrei*, and *C. trifenestrata*, also showed significant mass loss, over 25%. In contrast, cocoons from *S. cynthia* and *A. atlas* experienced much lower degumming losses, under 10%, indicating less sericin removal and/or inherently lower sericin content. Moreover, some cocoons exhibited a noticeable change in appearance after degumming (Figure S2), not only losing their natural colors (notably yellow *B. mori* and *A. yamamai*), but also appearing fluffier, as highlighted by crosshatched bars in Figure 6. This fluffiness results from the extensive removal of sericin, which normally acts as a gum and binds the fibroin filaments together. This effect is illustrated by the digital optical microscope images, such as for white *B. mori* and *B. madagascariensis* samples, which show
loosened silk fibroin filaments, allowing light to pass through (Figure 5g,h). In contrast, in cocoons like *A. paphia* (Figure 5i) and *A. atlas*, which are the stiffest ones upon handling, the fibroin filaments stuck together, indicating incomplete sericin removal. This difference in appearance and fibroin filaments adhesion underscores the variability in sericin removal efficiency across different silk types.

<figure><img src="image_7.png" /></figure>

Figure 6. Degumming loss of the cocoon samples. Crosshatched bars represent samples that exhibited a fluffy appearance after degumming. Solid bars represent samples that did not show a noticeable change in appearance after degumming.

The factors that influence the values of degumming loss in silk cocoon samples include: (1) the sericin content within the cocoon, (2) the presence of other extractable compounds, such as calcium oxalate, and (3) the strength of physicochemical interactions or bonds between fibroin, sericin, and other extractable compounds. The current literature on these factors, especially concerning wild silks, is limited. In this study, we examined these factors using FTIR, involving calculating relative intensities $I_r$ (am/cr) and $I_r$ ($CaC_2O_4$) (Figure 8), and Raman spectroscopy.

### 3.2.2. FTIR characterization

Changes were observed via FTIR following degumming. For the *B. mori* cocoon (Figure 7a), a distinct band 1 at 1695 cm⁻¹ emerges clearly after degumming, and notable variations are observed within wavenumber ranges 2 [1370-1415] cm⁻¹ and 3 [1010-1090] cm⁻¹. These changes were attributed to the removal of sericin. The amorphous component of the amide I band around 1635 cm⁻¹ diminishes, reflecting sericin's primarily amorphous nature. This reduction allows for better distinction of band 1 at 1695 cm⁻¹, indicative of the β-turn secondary structure in fibroin proteins [15,33]. The changes in wavenumber ranges 2 (associated with C-H and O-H bending) and 3 (linked to C-OH stretching) are consistent with the presence of serine residues abundant in sericin [5,15,32,46], whose removal during degumming decreases these spectral features. The post-degumming FTIR spectrum of *B. mori* is closely aligned with that of a reference commercial silk thread (devoid of sericin) (Figure 7a), showing that the
degumming process effectively removed sericin, corroborated by the high degumming loss value noted (Figure 6).

<figure><img src="image_8.png" /></figure>

Figure 7. FTIR spectra of (a) the *B. mori* thread, undegummed (outer wall) and degummed *B. mori* cocoon, and undegummed and degummed *A. paphia* cocoon (outer wall), and of (b) the white residues from the degumming baths, to be compared to pure calcium oxalate, solid soap and the experimental mixture of calcium oxalate and solid soap. O: absorption bands of calcium oxalate. (AG)n: absorption band of polyalanine-glycine β-sheets. (A)n: absorption band of polyalanine β-sheets. S-O: characteristic absorption bands of the partial reaction product between calcium oxalate and solid soap. S: characteristic absorption bands of solid soap.

The changes observed in the FTIR spectra at wavenumber ranges 1, 2 and 3 due to degumming are also visible for all non-mulberry silk samples, though to a lesser extent (Figure S4, and *B. madagascariensis* in Figure 7a). These reduced variations may be attributed to a lower initial quantity of sericin proteins in these silks, species differences in protein sequences, and/or less efficient extraction of sericin during the degumming process. Additionally, for all cocoons containing calcium oxalate, white particles formed in the degumming bath. Concurrently, three new bands S-O appeared in the FTIR
spectra of the degummed samples at 1470, 1540 and 1575 cm⁻¹ (Figure S4, and *B. madagascariensis* in Figure 7a). The white residues were analyzed by FTIR to determine their composition (black spectrum in Figure 7b). The results showed that these residues are partially composed of calcium oxalate (bands O at 660, 780, 885, 950, 1315, and 1605 cm⁻¹) and solid soap (bands S at 720, 1420, 1465, and 1550 cm⁻¹), along with the three bands S-O. To investigate whether these bands were due to a reaction between calcium oxalate and solid soap, a mixture of the two compounds was prepared in deionized water (several milligrams of calcium oxalate and soap placed in several milliliters of water), and boiled for 1 hour, following the degumming protocol. The resulting FTIR spectrum of the mixture (green spectrum in Figure 7b) confirmed the presence of the three bands S-O, indicating a partial reaction between the two substances. This reaction likely involves a cation exchange between calcium oxalate (Ca²⁺) and the conjugate bases of the fatty acids present in the soap, such as palmitate sodium (Na⁺), which is listed in the list of the commercial soap ingredients [37]. Palmitate calcium exhibits characteristic FTIR bands at these frequencies [51,52]. This suggests that calcium oxalate is not completely removed during degumming, with some crystals reacting and remaining in the silk despite the final rinsing step. This finding is consistent with expected behavior of oxalic acid, which has two pKa values at 1.2-1.5 and 3.8-4.4 under ambient conditions [53]. The alkaline degumming solution (pH = 9.93) is not sufficiently effective in dissolving oxalate crystals, as oxalic acid is soluble in water (220 g/L at 25 °C [53]), unlike calcium oxalate [19].

Consistent with the expected outcomes of cocoon degumming and the partial or total removal of sericin, a consistent decrease in I~r~ (am/cr) was observed across all samples, ranging from slight to significant (Figure 8a). The substantial drop in I~r~ (am/cr) for *B. mori* (both white and yellow) of more than 76% aligns with the potential total extraction of sericin. Similarly, I~r~ (am/cr) decreased significantly by over 91% for *A. assamensis*, *C. andrei*, and *C. trifenestrata*, in agreement with a total or near-total removal of sericin. The initial I~r~ (am/cr) values for *A. pernyi*, *A. yamamai*, and *S. cynthia* were notably lower than those for other species, suggesting a potentially lower sericin content in these samples. Interestingly, for *S. ricini* (both white and red cocoons), which initially had relatively high I~r~ (am/cr) values, the decrease following degumming was notably smaller (between 7 % and 41 %) compared to other samples with similar initial values. This suggests that sericin in *S. ricini* is only partially removed, possibly due to stronger physicochemical interactions or bonds with fibroin in this species.
<figure><img src="image_9.png" /><figcaption>Figure 8. (a) I<sub>r</sub> (am/cr), (b) diameters of individual fibroin filaments, and (c) I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) of the cocoons (averages of values from the inner and outer walls), before and after degumming. I<sub>r</sub> (am/cr) values were not calculated for *A. atlas* (undegummed), *A. paphia* (undegummed), *A. yamamai* (undegummed) and *B. madagascariensis* (undegummed and degummed) because for these cocoons, I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) > 3.5 %, significantly impacting the calculation of I<sub>r</sub> (am/cr) and rendering it irrelevant.</figcaption></figure>

Except for white *B. mori* and *C. trifenestrata* (out of the 13 cocoon samples), the degumming step, involving partial or complete removal of sericin, does not lead to a reduction in the diameter of the fibroin filaments, as observed through digital microscopy images (Figure 8b). Instead, the diameter either remained unchanged or even increased slightly in several cases. As a result, no correlation was found between the almost consistent decrease in I<sub>r</sub> (am/cr) (Figure 8a), associated with sericin removal, and the variation in filament thickness. While digital optical microscopy enables the observation of
fibroin filament detachment during sericin removal (e.g., Figure 5), it does not allow for direct characterization of the sericin, focusing instead on the fibroin itself. The reasons behind the slight increase in filament diameter observed in some cocoon samples remain unclear and warrant further investigation. Hypothetically, these changes could be attributed to a mechanical loosening of the fibroin filaments during degumming, a creation of charges within the proteins due to the applied alkaline solution, leading to electrostatic repulsion, or the compound resulting from the reaction between calcium oxalate and residual soap, which may remain on the fibroin filaments.

Excluding samples with initially low or negligible levels of calcium oxalate (I<sub>r</sub> < 1%), a significant decrease in I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) occurred in all cases, although it was less pronounced for *A. yamamai* (Figure 8c). Furthermore, there is no correlation between degumming loss and initial calcium oxalate content (Figure 9a). This finding reflects the complex interplay of factors specific affecting calcium oxalate content in the samples: (1) some crystals may be extracted, (2) others may remain unextracted, and (3) some crystals may react with the soap in the degumming solution, resulting in a product that partially remains in the silk while another part is extracted, forming white aggregates in solution. Additionally, the *B. madagascariensis* sample, which has by far the highest calcium oxalate content, exhibits a very significant degumming loss (36%), comparable to that of *B. mori* (white and yellow, 32%). Assuming that *B. madagascariensis* does not contain significantly more sericin than *B. mori*, this suggests that a higher calcium oxalate content does not necessarily hinder the efficiency of degumming and the extraction of sericin proteins.

Furthermore, as a general trend, a higher I<sub>r</sub> (am/cr) before degumming, indicating a greater initial sericin content, correlates with a higher degumming loss (Figure 9b). This observation supports the understanding that samples with more sericin lose more mass during the degumming process. Additionally, the correlation between degumming loss and sericin loss, rather than initial sericin content, was further investigated using the relative variation calculated by eq. 4 (Figure 9c):

$$ \Delta(\text{am/cr}) (\%) = \frac{I_r (\text{am/cr}) (\text{before degumming}) - I_r (\text{am/cr}) (\text{after degumming})}{I_r (\text{am/cr}) (\text{before degumming})} \quad (\text{eq. 4}) $$

Although the highest mass losses due to degumming align with the highest theoretical sericin losses (i.e., the highest Δ(am/cr) values for *B. mori* and for *C. andrei* and *C. trifenestrata*), this trend does not clearly apply to all samples. This lack of correlation is not unexpected, as degumming loss also logically depends on the initial sericin content, as previously indicated (Figure 9b). For instance, *S. cynthia* (X) exhibits a high Δ(am/cr) value (62%), suggesting significant theoretical sericin extraction, yet it has one of the lowest degumming losses (7%). This discrepancy is attributed to the initial sericin content before degumming (I<sub>r</sub> (am/cr) = 1.3), which is among the lowest compared to other samples (Figure 9b). Other observations remain unexplained and warrant further investigation. For example, despite similar mass losses (~15%) after degumming, the sericin content in white *S. ricini* (X, Δ(am/cr)
= 41%) decreased significantly more than in its red counterpart (x, Δ(am/cr) = 7%), even though their initial Ir (am/cr) values before degumming are very similar (Figure 9b), indicating comparable sericin contents. This difference in behavior could be explained by variations in the chemical composition of the cocoons, including the structure of fibroin and sericin proteins, crystallinity, and the amount of gum present. Such differences might stem from biological variability within the species or environmental factors affecting the silkworms, such as diet, humidity, and temperature.

<figure><img src="image_10.png" /></figure>

Figure 9. Degumming loss of the cocoons plotted against the pre-degumming values of (a) I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) and (b) I<sub>r</sub> (am/cr), and against (c) Δ(am/cr) (averages of values from the inner and outer walls). I<sub>r</sub> (am/cr) values were not calculated for undegummed *A. atlas*, *A. paphia*, *A. yamamai* and *B. madagascariensis* because for these cocoons, I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) > 3.5 %, significantly impacting the calculation of I<sub>r</sub> (am/cr) and rendering it irrelevant.

3.2.3. Raman characterization

The Raman spectra are almost identical before and after degumming (785 and 1064 nm, Figure S5), making it impossible to identify sericin-related bands or precisely track variations in calcium oxalate content as effectively as FTIR. The only significant change observed was in the yellow cocoon of *B. mori*, where the β-carotene signals (1005, 1160, 1190, 1215 and 1525 cm<sup>-1</sup>), which are responsible for its coloration [54,55], disappeared (Figure S5). This disappearance is attributed to the extraction of this pigment during degumming, as β-carotene is likely physically and chemically associated with sericin [54], which is almost entirely removed during the process. As a result, the degummed sample turned white (Figure S2), which is consistent with the removal of the pigment.
### 3.3. Dyeing of the silk samples

#### 3.3.1. Visual and microscopic inspections

The homogeneity of dyeing, defined here as the absence of color disparities on the sample surface observable to the naked eye and under an optical microscope (×50 to ×200), varies depending on the species, the wall considered (inner or outer), and the degumming step. Table 2 illustrates this variation using synthetic indigo dye. The apparent homogeneity of turmeric dye is more challenging to observe due to the natural color of some samples.

<table><caption>Table 2. Homogeneity of indigo dye applied to undegummed and degummed silkworm cocoons, determined by visual inspection and optical microscopy (magnification up to ×200), with microscopic images of *B. mori* and *B. madagascariensis*. Photographs at the macroscopic scale are provided in the Supporting Information (Figure S2).</caption><thead><tr><th rowspan="2">Cocoon sample</th><th colspan="2">Undegummed</th><th colspan="2">Degummed</th></tr><tr><th>Inner wall</th><th>Outer wall</th><th>Inner wall</th><th>Outer wall</th></tr></thead><tbody><tr><td><i>B. mori</i> (white)</td><td>+++</td><td>+</td><td>+++*</td><td></td></tr><tr><td><i>B. mori</i> (yellow)</td><td>+</td><td>+</td><td>+++*</td><td></td></tr><tr><td><i>S. ricini</i> (white)</td><td>+++</td><td>-</td><td>+++*</td><td></td></tr><tr><td><i>S. ricini</i> (red)</td><td>++</td><td>-</td><td>+++*</td><td></td></tr><tr><td><i>A. assamensis</i></td><td>++</td><td>+</td><td>++</td><td>+++</td></tr><tr><td><i>A. paphia</i></td><td>++</td><td>+</td><td>++</td><td>++</td></tr><tr><td><i>B. madagascariensis</i></td><td>++</td><td>+</td><td>+++*</td><td></td></tr><tr><td><i>A. atlas</i></td><td>++</td><td>-</td><td>++</td><td>+</td></tr></tbody></table>

- : Very high heterogeneity visible to the naked eye, with a large portion of the surface remaining undyed.

+ : Moderate to high heterogeneity visible to the naked eye, with several areas of the surface either

undyed or more weakly dyed. ++ : Heterogeneity visible only under optical microscopy, with several

fibroin filaments either undyed or more weakly dyed. +++ : Homogeneous dyeing to the naked eye and

under optical microscopy. * : Sample exhibiting a fluffy appearance following degumming, with indistinguishable walls.

<figure><img src="image_11.png" /></figure>

Dyeing looks more uniform on degummed samples, likely because the partial or total removal of sericin enhances dye uptake, as sericin is known to hinder dye efficiency [6,25]. This is especially true for fluffy degummed samples, where sericin extraction is presumed to be complete or nearly so. Furthermore, dyeing tends to be more homogeneous on the inner walls of cocoons. In some instances, such as *B. mori* (Figure 4b) [23], this may result from a lower amount of sericin on the inner wall. In
other cases, such as *B. madagascariensis* (Table 2) and *A. paphia* (Figure S6), digital optical microscopy reveals that calcium oxalate crystals on the cocoon's surface, especially on the outer wall, which contains large quantities, remain white after dyeing, while the protein fibers become colored. This suggests a lack of physico-chemical interactions between calcium oxalate and the dyes, leading to apparent dye heterogeneity. Moreover, without degumming, dyeing is more uneven on the surface of yellow *B. mori* and red *S. ricini* cocoons compared to their white counterparts (Table 2). This could be due to the presence of natural pigments, namely β-carotene in *B. mori* and unidentified pigments in *S. ricini*, which may interfere with dye adherence. Similarly, other natural pigments, such as methyl isoeugenol in *A. assamensis* [56], might impede dyeing efficiency.

### 3.3.2. Raman characterization

The characteristic signals of turmeric dye were not detected in any of the dyed silk samples, either by FTIR (Figure S7), or Raman spectroscopy (Figures S8 and S9), either at 785 or 1064 nm. In contrast, the characteristic Raman signals of synthetic indigo were clearly identified in the dyed silks (Figures S8 and S9), without any observable wavenumber shift. However, Raman is less effective for detailed study of the interactions between the dye and the silk compared to FTIR, which allows for more detailed semi-quantitative *in situ* analysis of the dye.

### 3.3.3. FTIR characterization

The FTIR signals of indigo are detected with varying intensity depending on the sample (Figure S10 and Figure 10). For instance, the indigo signals are more pronounced on the outer wall of undegummed *A. paphia* compared to the inner wall of undegummed white *S. ricini*. Additionally, the indigo signals are more intense in the spectrum of *S. ricini* than in the fluffy silk of degummed white *B. mori* cocoons (Figure 10). Moreover, due to the final washing step of the dyeing procedure ("2.3. Dyeing" section), the reaction product between soap and calcium oxalate forms in the samples containing the mineral (S-O bands in Figure 10), similarly to what occurs following the degumming step (Figure 7a).
<figure><img src="image_12.png" /><figcaption>Figure 10. FTIR spectra of indigo and of undyed and dyed (with synthetic indigo) white *B. mori*, white *S. ricini*, and *A. paphia* cocoons. UD: undegummed sample. D: degummed sample. S-O: absorption bands characteristic of the reaction product between calcium oxalate and solid soap. i<sub>1480</sub>: absorption band at 1480 cm<sup>-1</sup> of indigo used for the calculation of I<sub>r</sub> (indigo).</figcaption></figure>

Some shifts were observed, notably an intense band of indigo that originally appears at 1059 cm<sup>-1</sup> for the reference dye, found at 1069 cm<sup>-1</sup> in the silks (Figure 10). This shift, partly attributed to the C=O (rocking) bonds of indigo [57], indicates the existence of physicochemical interactions (e.g., hydrogen bonds, Van der Waals forces) between the dye and the silks. The shift associated with the C=O bonds (involving two ketones in the molecule) is more pronounced (shift of +10 cm<sup>-1</sup>) compared to that due to the N-H bonds (two secondary amines in the molecule) (shift of +5 cm<sup>-1</sup>) observed at 1120 - 1125 cm<sup>-1</sup> (N-H rocking + C-N stretching [57]). This suggests that interactions between the dye and the silks may be predominantly with the ketones rather than the amines. The specific molecular binding sites on the various protein sequences of the different silks are not well understood; hence, for instance, a more detailed study using Nuclear Magnetic Resonance (NMR) could be considered to elucidate these interactions further.

To study and compare more precisely the proportion of indigo on the inner and outer walls of the cocoons (whether degummed or not), FTIR spectra were decomposed in the wavenumber range of 1470 - 1730 cm<sup>-1</sup> (Figure 11a, exemplified by the spectrum of the outer wall of an undegummed white *B. mori* cocoon). A semi-quantitative analysis was then conducted, based on the calculation of the relative intensity as described in eq. 5:

$$I_r (\text{indigo}) (\%) = \frac{A(1480)}{A(\text{amide II})} \qquad (\text{eq. 5})$$
where A(1480) represents the area (absorbance) of the FTIR band at 1480 cm⁻¹ due to indigo (denoted as *i*₁₄₈₀ in Figure 10). This well-resolved band, distinct from the signals associated with silk, corresponds to both the stretching of C-C bonds and the deformation of C-H and N-H bonds of indigo [58]. The amide II band (components at 1515 and 1540 cm⁻¹ in Figure 11a) was chosen for normalization over the amide I band because indigo does not exhibit significant FTIR signals in the amide II region (Figure 10). All *I*<sub>r</sub> (indigo) intensities are reported in Figure 11b,c.

<figure><img src="image_13.png" /><figcaption>Figure 11. (a) Decomposition of the FTIR spectrum for the outer wall of undegummed indigo-dyed white *B. mori* cocoon in the 1470-1730 cm⁻¹ range (linear baseline, Gaussian components). (b,c) Relative intensity *I*<sub>r</sub> (indigo) on the inner and outer walls of (b) undegummed and (c) degummed cocoons (white bars: samples exhibiting a fluffy appearance following degumming, with indistinguishable walls).</figcaption></figure>
With the sole exception of the inner wall of *A. atlas* (for reasons yet to be elucidated), the intensity $I_r$ (indigo) is significantly lower, often markedly so, after degumming (Figure 11). For degummed samples exhibiting a fluffy appearance (white bars in Figure 11), the values are all among the lowest ($I_r \le 3\%$). This reduction is attributed to the partial or complete removal of sericin, which allows indigo dye molecules to physically penetrate the sample more easily as the fibroin filaments are no longer adhered together. The extent of dye uptake within the mass would depend on the nature and abundance of the residual sericin layer surrounding the individual filaments, explaining the variations in $I_r$ (indigo) values among samples, as well as between undegummed samples. In samples where fibroin filaments remain adhered, the dye solution penetrates less effectively, resulting in the dye molecules accumulating primarily on the inner and outer surfaces of the cocoons, leading to heterogeneous dyeing as described in “3.3.1. *Visual and microscopic inspections*” section. This disparity explains why indigo dye is more readily detected on undegummed samples (i.e., those rich in sericin) using attenuated total reflectance (ATR), a surface analysis technique.

Furthermore, in several cases, for a same silk sample, the $I_r$ (indigo) intensity is higher on the inner wall compared to the outer wall, as observed, for example, in undegummed white and yellow *B. mori* (Figure 11b). This difference could be correlated with a lower sericin content on the inner wall, as also discussed in “3.3.1. *Visual and microscopic inspections*” section. Given that sericin glue acts as a barrier to dye penetration, dye molecules likely entered the sample more easily through the inner wall, where they accumulated. This accumulation occurs because the adhesive sericin protein gradually becomes more abundant towards the outer layers, limiting further penetration and distribution of the dye molecules.

Interestingly, the calcium oxalate content does not appear to correlate with the $I_r$ (indigo) intensity (Figure S11). This is evident, for instance, when comparing the intensities on the inner and outer walls of undegummed *A. paphia* and *B. madagascariensis* (Figure 11b). Although the outer wall of each cocoon contains much more calcium oxalate than the inner wall (Figure 4a), the $I_r$ (indigo) values show minimal differences, or none. This finding suggests that the presence of calcium oxalate, which according to microscopic observations does not interact with the dyes (images provided with Table 2), does not impede the dye's penetration into the silk fibers.

### 3.3.4. UV-Visible characterization

Aqueous turmeric dye solutions visibly decolorize after application to the silks, likely due to the sorption of the dye, particularly curcumin [39,59]. Therefore, it was deemed relevant to conduct UV-Vis absorbance measurements on the clear post-dyeing solutions to determine the concentrations of residual dye. Indigo suspensions were excluded from UV-Vis analysis due to the dye's insolubility in
water. The absorbance spectrum of the turmeric stock solution, diluted 40-fold in water (as the signal of the stock solution itself was saturated under the experimental conditions) shows a peak at 276 nm (Figure 12a), attributed to curcumin [59]. This peak was used to calculate the concentration ratio C(x)/C<sub>st</sub>, where C(x) represents the dye concentration in the post-dyeing solution and C<sub>st</sub> is the concentration of the stock solution. This ratio theoretically corresponds to the fraction of dye not absorbed by the silk.

To establish a correlation between absorbance and concentration, variously diluted solutions (duplicates, diluted factors: 40, 100, 200, 250, 400, and 1000 from the stock solution) were heated to 95-100°C for 1 hour as controls without silk samples. The absorbance was then measured using UV-Vis spectroscopy (Figure 12b). A calibration curve (absorbance as a function of C(x)/C<sub>st</sub>) was developed to quantify the amount of dye sorbed by the silks.

<figure><img src="image_14.png" /><figcaption>Figure 12. (a) Absorbance spectrum of the turmeric dye stock solution diluted by a factor of 40. (b) Absorbance values of the turmeric dye solution plotted against the dilution factor of the stock solution, used for calibration. (c,d) Percentage of dye sorbed from turmeric dye solutions by (c) undegummed and (d) degummed cocoons, calculated based on UV-Vis absorbance measurements.</figcaption></figure>

However, the UV-Vis spectra of solutions applied to undegummed cocoon samples of red *S. ricini*, *A. assamensis*, *B. madagascariensis*, and *A. atlas* showed unassigned bands near the critical wavelength (~276 nm) (Figure S12), complicating the accurate measurement of absorbance at this peak. Consequently, calculations for these four undegummed samples were not performed. The unassigned bands might result from natural dyes present in these cocoons, which are extracted during degumming
and dyeing processes. These natural dyes are unknown, except for *A. assamensis*, where the golden color is partly due to methyl isoeugenol, which has an intense absorbance band at 285 nm [56].

The percentage of sorbed dye for each turmeric-dyed cocoon sample, both degummed and undegummed, is presented in Figure 12c,d. Among the undegummed cocoons (Figure 12c), *A. paphia* exhibits the lowest dye sorption, with only 1% of the dye sorbed. In contrast, its degummed equivalent (Figure 12d) shows much higher dye sorption at 14%. For the degummed samples, *B. mori* and *B. madagascariensis* exhibit the highest dye sorption, with percentages exceeding 30%. The other degummed silks display relatively similar dye sorption percentages, ranging from 14% to 23%. Interestingly, the degumming procedure does not consistently enhance dye sorption: for *B. mori* (white and yellow), the dye sorption percentage remains largely unchanged after degumming. Moreover, in the case of white *S. ricini*, the applied degumming process actually reduces dye sorption. These results suggest that while sericin may physically impede dye penetration into the silk mass, it can still form physicochemical interactions with the dye, influencing the overall sorption capacity.

To better interpret the UV-Vis data, the analysis of which captures the global quantity of dye sorbed, the percentage of dye sorbed was plotted against the degumming loss for each cocoon sample (Figure 13a). A clear correlation was found: greater degumming loss corresponds to enhanced dye uptake, regardless of the cocoon sample. Additionally, for the degummed samples, the percentage of dye sorbed was plotted against the intensity $I_r$ (am/cr) (Figure 13b), obtained by FTIR measurements, which, unlike UV-Vis, are surface-specific and detect residual sericin and dye interactions at the cocoon surface. For samples that developed a fluffy appearance after degumming, namely *B. mori* and *S. ricini*, the percentage of sorbed dye decreases as $I_r$ (am/cr) increases, indicating that higher residual sericin content hampers dye uptake. Conversely, *A. assamensis*, *A. paphia* and *A. atlas*, which showed minimal change in appearance during degumming and contain relatively low sericin content, demonstrated low dye sorption (circled group in Figure 13b). These observations align with the conclusions drawn from the indigo-dyed samples and the calculation of FTIR intensities $I_r$ (indigo) (see "3.3.3. *FTIR characterization*"). These findings suggest that physical factors influence dyeing efficiency and sorption. In degummed samples where the fibroin filaments are no longer agglutinated (fluffy samples), the dye solution can penetrate more easily. In these cases, dye uptake is likely influenced by the presence of a heterogeneous layer of residual sericin coating the individual filaments. In contrast, in samples where the fibroin filaments remain adhered together (circled group in Figure 13b), the dye solution's penetration is limited, resulting in fewer available dye binding sites and a tendency for the dye to fix primarily on the outer surface of the cocoons.
<figure><img src="image_15.png" /><figcaption>Figure 13. Percentage of dye (curcumin) sorbed during dyeing as a function of (a) degumming loss and (b) I<sub>r</sub> (am/cr) (averages of the inner and outer wall values) before dyeing for each degummed cocoon sample. The I<sub>r</sub> (am/cr) values were not calculated for *B. madagascariensis*, as for this degummed sample, I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) > 3.5%, significantly impacting the calculation of I<sub>r</sub> (am/cr) and rendering it irrelevant.</figcaption></figure>

There is no apparent correlation between dye sorption and calcium oxalate content (Figure S13).
For the fluffy degummed samples, I<sub>r</sub> (CaC<sub>2</sub>O<sub>4</sub>) does not seem to influence dye sorption: *B. madagascariensis*, which retains the highest concentration of calcium oxalate crystals after degumming (Figure 8c), sorbs the dye as effectively as *B. mori*, which has none. This aligns with the fluffy appearance of degummed *B. madagascariensis*, indicating substantial sericin removal, thus separating the fibroin filaments into individual ones and facilitating dye penetration into the mass. However, as noted through the microscopic examination of the samples (Table 2), the presence of calcium oxalate crystals, and potentially the reaction product between calcium oxalate and the soap (section "3.2.2. FTIR characterization"), may reduce the number of available binding sites on the sample surfaces, thereby affecting the amount of dye adsorbed. This is especially evident in undegummed *A. paphia*, where the proportion of sorbed dye was nearly negligible (Figure 12c).

**4. Conclusions**

We investigated the dyeing behavior of various silkworm cocoons, with a focus on their
composition and the degumming process, as well as the role of sericin and calcium oxalate in dye
sorption. Our findings highlight several key points and suggest avenues for further research. Firstly, the
results emphasize the significant impact complexity of the degumming process on the dye uptake of
wild silk cocoon fibers. While degumming generally enhances dye penetration by removing sericin, the
efficiency and outcome of the process vary depending on the sericin content and the specific
characteristics of each species. Notably, our study revealed that a greater degumming loss tends to
facilitate dye uptake. However, in cases like *A. paphia*, where the appearance does not change
substantially after degumming, the presence of calcium oxalate appeared to impede dye sorption, possibly due to incomplete removal of sericin.

Despite these insights, some questions remain unanswered regarding the precise mechanisms underlying degumming and dyeing processes. Future research should aim to address these questions, particularly by employing advanced analytical techniques. For instance, Nuclear Magnetic Resonance (NMR) spectroscopy in the solid phase could be tested to finely characterize the silks composition and the nature of the interactions between dyes and silk components. This approach could provide deeper insights into the physical and chemical bonds involved, potentially clarifying the role of specific protein sequences in the dyeing process. Additionally, destructive quantitative analyses, such as thermogravimetric analysis, EDTA extraction, and protein quantification techniques like Lowry or Biuret assays, could be used to quantify calcium oxalate, and fibroin and sericin contents in silk samples. Such studies would enhance the understanding of the degumming process and contribute significantly to the broader field of silk dyeing.

Moreover, the influence of natural pigments in wild silks on dye adhesion remains an open question. Characterizing these pigments could reveal their role in the dyeing behavior of the silk fibers and offer new insights into their interactions with dyes.

Looking ahead, it is crucial to extend these investigations beyond raw cocoons to include studies on dyed silk threads and fabrics. This could be particularly valuable in the context of preserving colored silks, as understanding the interactions between dyes and silk components at a finer scale could inform conservation strategies. In this regard, the impact of degumming on the micro- and macroscale mechanical properties of silk fibers, as well as the effect of mordants on fiber integrity and aging, remains to be explored. Investigating these aspects could provide practical insights into the long-term stability and preservation of wild silk textiles, a field that remains underexplored. Furthermore, our study was limited to turmeric and indigo dyes; thus, future research should explore the interaction of silk with a broader range of dyes, including acid dyes often used in dyeing factories, to fully understand the factors influencing dye uptake and retention across different silk types.

## Acknowledgments

The authors thank all those who kindly provided silk samples and advice, namely textile craftswoman Marie Foyer, entomologists Rodolphe Rougerie (MNHN, Paris) and Hervé Guyot (Office pour les insectes et leur environnement, France), and anthropologists Laurence Douny (Max Planck Institute for the History of Science, Berlin) and Nicolas Césard (MNHN, Paris). Heartfelt thanks are expressed to Marie Foyer for training in dyeing techniques. Oulfa Belhadj, Alice Gimat and Bertrand
Lavédrine (CRC) and Ludovic Bellot-Gurlet and Aline Percot (MONARIS, Paris) are warmly thanked
for technical contribution and fruitful discussions.

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CRediT authorship contribution statement

**Nathan Ferrandin-Schoffel**: Conceptualization, Investigation, Validation, Visualization, Writing - original draft. **Sophie Cersoy**: Resources, Conceptualization, Supervision, Writing - review and editing.

**Sophie Desrosiers**: Resources, Conceptualization, Writing - review and editing. **Annabel Vallard**: Funding acquisition, Project administration, Resources, Conceptualization, Writing - review and editing.

Funding

This work was supported by ANR "Matters and culture: the wide world of wild silks" (WILDSILKS-ANR-19-CE27-0020 / AAPG2019) funded by the French Agence nationale de la recherche.

Statements & Declarations

Competing Interests: the authors declare that they have no conflict of interest and no relevant financial or non-financial interests to disclose.