# Variation in the hierarchical structure of lignin-blended cellulose precursor fibers

Jiliang Liu, Jenny Bengtsson, Shun Yu, Manfred Burghammer, Kerstin Jedvert

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Jiliang Liu, Jenny Bengtsson, Shun Yu, Manfred Burghammer, Kerstin Jedvert. Variation in the hierarchical structure of lignin-blended cellulose precursor fibers. International Journal of Biological Macromolecules, 2023, 225, pp.1555-1561. 10.1016/j.ijbiomac.2022.11.211. hal-05087954

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# Variation in the hierarchical structure of lignin-blended cellulose precursor fibers

Jiliang Liu<sup>a, *</sup>, Jenny Bengtsson<sup>b</sup>, Shun Yu<sup>c</sup>, Manfred Burghammer<sup>a</sup>, Kerstin Jedvert<sup>b</sup>

## ARTICLE INFO

### Keywords:

Hierarchical structure

Lignin-cellulose precursor fibers

Scanning X-ray microdiffraction

## ABSTRACT

Regenerated cellulose fibers have been considered as potential precursor fibers for carbon fibers because of their balanced cost and performance. Increased attention has been paid to blending lignin with the regenerated cellulose to generate precursor fibers which render good mechanical properties and higher carbon yield. The mechanical properties of carbon fibers have been found closely correlated to the structure of precursor fibers. However, the effects of lignin blending on molecular- and morphological structure of the precursor are still unclear. This study aims at clarifying the structural information of lignin-cellulose precursor fibers from molecular level to mesoscale by scanning X-ray microdiffraction. We present the existence of a skin-core morphology for all the precursor fibers. Increase of lignin content in precursor fiber could reduce the portion of skin and cause obvious disorder of the meso- and molecular structure. By correlating structural variations with lignin blending, 30% lignin blending has been found as a potential balance point to obtain precursor fibers maintaining structural order together with high yield rate.

## 1. Introduction

Carbon fibers (CFs) are increasingly being used as reinforcing agents in lightweight composites, but their use is limited by high price and reliance on fossil fuel raw materials. Most commercial CFs are manufactured from petroleum-derived polymer – polyacrylonitrile (PAN) – precursor fibers (PFs). The current financial cost of CFs, of which approximately half is the cost of PAN, limits the wide-spread use of CFs [1]. Furthermore, there is increasing concern about the environmental cost of the petroleum-related PAN synthesis [2–4]. These factors are motivating the exploration of alternatives to PAN for production of CFs.

Regenerated cellulose fibers are promising low-cost alternatives for precursor fibers (PFs) for the production of CFs. They have high tenacity and modulus, and maintain high mechanical performance during carbonization [5]. Compared to PAN, regenerated cellulose fibers are less expensive and have a lower carbon footprint. Lignin-cellulose blends also show promise as an even lower-cost candidate for PFs. Lignin is a large amorphous macromolecular byproduct of wood pulping that has high carbon content (~60 %). Although lignin does not form fibers directly, lignin-cellulose blends exhibit good molecular orientation with promising mechanical properties and high carbon yield,

making them excellent candidates as PFs. There are extensive studies on the stabilization and carbonization of lignin-cellulose blended PFs [6–8] but little information on the effect of lignin-cellulose blends on the molecular structure of regenerated cellulosic PFs. PF with a high lignin content enables a high carbon yield during CF manufacturing. However, a high lignin content also disrupts the crystallinity and molecular orientation of cellulose within the PF, which will have detrimental effect on the mechanical properties of the CF [9–11]. Thus, the lignin content must be balanced with regard to both carbon yield and molecular orientation.

Studies of pure cellulose fibers have shown that structural order, such as crystallinity and degree of orientation, are important determinants of their physical properties, e.g., better tensile stress, modulus, and elongation. Transmission electron microscopy (TEM) is a common imaging technique for studying the molecular structure of cellulose fibers [12]. However, due to the limited penetration depth of the electrons, thin samples need to be prepared in order to be suitable for TEM. Micrometer scale morphology of regenerated cellulose fibers can be revealed by scanning electron microscopy (SEM) or atomic force microscopy (AFM) [13–16]. However, these techniques are limited to the determination of surface morphology, with internal structure only

<footnote><sup>a</sup> European Synchrotron Radiation Facility, ESRF, Grenoble 38000, France</footnote>

<footnote><sup>b</sup> RISE Research Institutes of Sweden, Materials and Production, Mölndal 43153, Sweden</footnote>

<footnote><sup>c</sup> RISE Research Institutes of Sweden, Bioeconomy and Health, Lund 22370, Sweden</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: jiliang.liu@esrf.fr (J. Liu).</footnote>
visible after sectioning, which could introduce distortions leading to artifacts regarding structural information. As an alternative, scanning X-ray microdiffraction (SXM) can fill the multiscale gap by characterizing in detail the “skin-core” morphology of regenerated cellulose fibers [13–15]. Analysis of individual microdiffraction patterns enables quantitative characterization of crystallite dimensions, orientation, and overall crystallinity at the microscale. Scattering in the small-angle regime (SAXS) can provide information on the distribution of microvoids and the alignment of cellulose elemental fibrils (CEFs) [17].

In this work we demonstrate that SXM applied to lignin–cellulose precursors from different blends can reveal the effects of lignin blending on the multi-level structure of regenerated cellulose fibers. Our results unravel the impact of lignin blending on the “skin-core” morphology of PFs; providing a fundamental understanding of the variation of the molecular architecture among different lignin blended PFs. The structural variation induced by changes in lignin content identifies a potential critical point for lignin blending, where the molecular orientation is maintained together with a high carbon content of the fiber.

## 2. Experiments and methods

### 2.1. Sample preparation

Lignin–cellulose precursor fibers were prepared from softwood Kraft lignin (received from Backhammar Pilot Plant (Backhammar, Sweden). For further characterization of the lignin used we refer to our previous publication [10]) and softwood Kraft dissolving pulp, with an intrinsic viscosity of 465 mL/g (measured according to ISO 5351:2010) (Georgia Pacific Cellulose, USA). Lignin and cellulose were dissolved simultaneously in 1-ethyl-3-methylimidazolium acetate (EMIMAc, Aldrich 95 %) at 70 °C, for 1 h in a closed reactor with overhead stirring at 30 rpm. Solutions were prepared with 8–16 wt% solid content, with 8 wt% cellulose and 0–8 wt% lignin, i.e. 0–50 wt% lignin to cellulose ratio in the solution (Table 1). Deaeration was performed at 60 °C and below 100 mbar pressure for 15 h. The solution was spun using bench-scale spinning equipment, customized by RISE Research Institutes of Sweden, consisting of a piston pump, a coagulation bath (7 L), and a take-up roll. The temperature of the solution was set to 40 °C. The spinneret had 33 orifices with a diameter of 150 µm and an aspect ratio (L/D) of 3. The solution was filtered through a 5 µm metal fleece filter and extruded with a fixed extrusion velocity of 4 m/min. The extruded filaments were led via an air gap of 1 cm into a coagulation bath of deionized water at a temperature of 2–5 °C. The take-up speed during spinning was set to 24 m/min to achieve a draw ratio of 6. The draw ratio was maximized with respect to achieving stable spinning for all lignin-cellulose ratios. It was found in a previous study that lignin-blending enables high draw ratios during spinning, therefore, the pure cellulose solution was limiting the used draw ratio in this study [18].

### 2.2. Scanning X-ray microdiffraction experiments

SXM experiments were carried out at the Microfocus beamline ID13 at European Synchrotron Radiation Facility (ESRF) with an X-ray energy of 13 KeV (wavelength 0.953 Å). Beryllium Compound refractive

<table><caption>**Table 1**
Comparison of prepared solution [18].</caption><thead><tr><th>Sample name</th><th>Cellulose, wt %</th><th>Lignin, wt %</th><th>Lignin, ratio %</th><th>Total solids content, wt%</th></tr></thead><tbody><tr><td>Lignin 0 %</td><td>8</td><td>0</td><td>0</td><td>8</td></tr><tr><td>Lignin 10 %</td><td>8</td><td>0.9</td><td>10</td><td>8.9</td></tr><tr><td>Lignin 33 %</td><td>8</td><td>4</td><td>33</td><td>12</td></tr><tr><td>Lignin 50 %</td><td>8</td><td>8</td><td>50</td><td>16</td></tr></tbody></table>

lenses (CRL) focused the monochromatic beam down to a size of 2.5 µm at the sample position. An ion chamber and a circular guard aperture of 20 µm in diameter were put upstream of the sample to reduce the diffuse background scattering. The total flux was ~10¹² photons/s. Downstream of the sample, a beam stop, blocking the direct beam, and a 2D photon counting Eiger X 4 M detector (DECTRIS, 2167 × 2070 pixels of size 75 µms) were positioned. The sample-to-detector distance was calibrated to 164.7 mm using an alpha-Al₂O₃ powder standard. The samples were fly-scanned on a mesh grid of 30 × 30 points having a step size of 2.5 µm. The exposure time was 50 milliseconds. Individual PFs were glued on a frame with empty window and aligned to focus position of the microbeam. The frame was mounted on a motor motion stage for fly-scanning.

### 2.3. Data processing of Scanning X-ray microdiffraction

A customized processing program written in Python was used for all data. The packages, PyFAI [19], silx, and Imfit [20] were used for fast integration, visualization, and fitting. Fibers from regenerated cellulose (cellulose II) are composed of crystallites together with some disordered (“amorphous”) regions [14,15]. The cellulose II crystallites are monoclinic with $a = 8.1$ Å, $b = 9.03$ Å, $c = 10.31$ Å, $\gamma = 117.10°$ [21]. The equatorial intensity profile provided quantitative structural information, such as lattice constant, crystallinity index (CI) and crystallite domain size. The lattice constants of lignin 0 %, shown in Table 2, exhibit a moderate discrepancy from previous reports of cellulose II crystals [21]. Calculations of crystallinity of cellulose and crystallite domain size have been made in numerous ways and remain open to discussion [22–24]. Deconvolution of multi-Voigt functions and an asymmetrical amorphous function have been used for fitting the scattering power of reflections.

The micro-SAXS has been extensively used to characterize the mesoscale architecture of PFs, such as the distribution of microvoids and arrangement of CEFs within PF. As shown in Fig. S8a, we attempt to determine the porosities of regenerated fibers by a Guinier plot at a Q range of 0.01–0.1 Å⁻¹ [25]. The core region of 100 % cellulose regenerated fiber exhibits a structure factor peak at Q ~0.15 Å⁻¹. This structure factor was interpreted as due to interference among inter-fibrils.

## 3. Results

### 3.1. Morphological structure determined by SXM

Individual PF with different lignin blending ratio were scanned with the 2.5 µm X-ray beam. SXM data includes both small- and wide-angle X-ray scattering (SAXS–Q range from 0.01 to 0.5 Å⁻¹ and WAXS–Q range from 0.5 to 4 Å⁻¹). SAXS provides mesoscale structural information on the 10–100 Å scale, WAXS provides information on structural features over the range 1–10 Å. The degree of preferred orientation (DPO) represents the deviation of CEFs from the fiber axis, which is a critical parameter for the structural order of CEFs within PFs. The PFs containing highly oriented CEF exhibited good mechanical properties, such as high tenacity and elastic modulus [26]. These good mechanical properties are preserved after carbonization. The DPO reflects the structural integrity of oriented CEFs. As shown in Fig. S1, the DPO was determined by the FWHM of Lorentz functions used to fit the azimuthal profile of featured reflections of cellulose. As shown in Fig. 1b, the variation of the DPO across the pure cellulose fiber indicates that the

<table><caption>**Table 2**
Comparison of lattice constant of cellulose II crystal.</caption><thead><tr><th>Lattice constant</th><th>a</th><th>b</th><th>c</th><th></th></tr></thead><tbody><tr><td>Cellulose II [21]</td><td>8.01 Å</td><td>9.03 Å</td><td>10.31 Å</td><td>117.10°</td></tr><tr><td>Fitted results</td><td>7.90 Å</td><td>9.40 Å</td><td>10.46 Å</td><td>117.10°</td></tr></tbody></table>
<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Scanning X-ray microdiffraction of a pure cellulose fiber (lignin 0%). a) optical microscope image of cellulose fiber. b) map of degree of preferred orientation shows a skin-core morphology. c) the microdiffraction pattern from the core region of the fiber, inset is the zoom-in of the SAXS region. 1D equatorial intensity profile is azimuthally integrated from 2D diffraction patterns. d) the microdiffraction pattern from the skin region of the fiber prepared in a manner similar to (c).</figcaption></figure>

fiber has a distinct “skin” with a higher DPO and a core with a lower but uniform DPO. This “skin-core” morphology is consistent with previous studies of pure cellulose fibers using SXM [27,28]. The differences in the diffraction pattern of core, Fig. 1c, and from skin, Fig. 1d, indicates that the *meso* and molecular structures of the skin differ significantly from the core. The higher DPO at the “skin” indicates the CEF are better organized in the skin than in the core.

### 3.2. Effect of lignin on molecular properties of the CEFs

The structural features of cellulose elemental fibrils (CEFs), i.e., DPO, CI, the lattice constant of unit cell and the domain size of cellulose crystallite, have been determined by SXM WAXS-analysis. This analysis identified an association between structural variations of these properties and the changes of lignin content.

<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Variation of molecular structure of PFs at microscopic scale. a)–d) optical image of PFs having different lignin content for scanning. While PFs with 0 % lignin are transparent, blending of lignin dyes the PFs. e)–h) maps of the degree of preferred orientation for PFs. The hotter color (red) indicates a better orientation preference, in contrast, the colder color (blue) implies that fibrils are less oriented. i)–l) maps of crystallinity index of fibers having different lignin content.</figcaption></figure>
Maps of DPO, Fig. 2e-h, show that while the “skin-core” morphology is preserved, compared to the pure cellulose fiber, for all blends, an increase of lignin content within PFs will significantly reduce the DPO of CEFs [27,28].

Another parameter that will influence the mechanical properties of the resultant CFs is CI of PFs. The CI provides a quantitative representation of the ratio between CEF and amorphous content (composed by lignin and amorphous cellulose) for the PFs. PFs adequate for production of CFs with desired mechanical properties should contain enough lignin to achieve good carbon yield after carbonization, but not cause too much disruption of the cellulose crystallinity [29]. Thus, it is important to obtain a quantitative correlation of lignin content (mass fraction) and cellulose crystallinity to identify the optimal blend ratio for PFs with a good yield and desirable properties.

Maps of CI, as shown in Fig. 2i-l, show a strong anticorrelation with lignin content and exhibit “skin-core” morphology, with a higher CI in “skin”. We qualitatively compare the variation of lignin content with CI to identify their synergistic relation. By comparing the variation of mass fraction of cellulosic materials with the CI, we find that the CI decreases rapidly already at low concentration lignin-blending, i.e. up to ~30 % lignin, c.f. Fig. 3. However, then the CI seems to be less altered when the lignin concentration is increased above ~30 %. This implies that including lignin will not only contribute to mass fraction of amorphous content but also increase the ratio of amorphous cellulose to cellulose content. However, at a lignin content of 33 % or greater, lignin becomes the predominate amorphous component and causes little increase of amorphous cellulose.

The variations of crystal domain size also support 33 % as a critical blending ratio. Fig. 4a shows that the domain size of the (0 0 4) cellulose reflection that corresponds to the fibrillar length of CEF, decreases from 139.8 ± 8.8 Å for 0 % lignin to 126.4 ± 11.9 Å, 127.4 ± 7.8 Å, 127.3 ± 9.9 Å for lignin content of 10 %, 33 % and 50 %, respectively. The variations of in-plane, normal to (110) and (020) crystal plane, crystallite domain size of CEF exhibit a similar trend. From Fig. 4b, it can be found that domains sizes corresponding to the (1 1 0) and (0 2 0) reflections are decreasing with increasing lignin content. For 0 % lignin PF, the domain sizes of (1 1 0) and (0 2 0) reflections are ~31 Å and ~35 Å. Whereas for 50 % lignin PF, the domain size drops to ~28 Å and ~31 Å, respectively. Therefore, while an increase in lignin content will induce additional amorphous cellulose, the amount of amorphous

<figure><img src="image_6.png" /><figcaption>Fig. 3. The correlation of lignin content and crystallinity index (CI). Cellulose content (CC), theoretical mass content of cellulose and shown as black solid diamond, is calculated as (1 - lignin content). Curve of CC is fitted by 2nd order polynomial. CI with standard deviation for skin and core has been shown as green and blue solid circle, similar polynomial fitting used to obtain curves of CI. To compare the variations between CC and CI, the origin of curve of CC has been shifted to the origin of curves of CI (black dash curve).</figcaption></figure>

cellulose is still much lower than the lignin content, especially at lignin concentrations of 33 % and greater.

In summary, WAXS data from the PFs indicate that an increase of lignin content will cause disorder in CEFs in both transverse and longitudinal directions of the PFs, which will lead to a reduction in degree of preferred orientation and CI. Based on our findings, lignin becomes the dominate amorphous component in the fibers when the concentration of lignin exceeds ~30 %.

### 3.3. Mesoscale structural features revealed by SAXS

During spinning process, cellulose molecules first form elongated crystallites-CEFs, then CEFs pack together as a fibril matrix containing microvoids. In Fig. 5a, we observe that, in the core region, the 0 % lignin PF shows significant interference at a Q of 0.15 Å⁻¹ for the equatorial intensity profile. This pattern corresponds to a structure factor caused by inter-fibril interference from highly ordered CEFs within fibrils matrix. A free 2D cylinder model has been introduced to quantify the inter-fibril distance within the PF. Fitting results indicate that the inter-fibril distance of 0 % lignin PF is ~44 Å, with a cylinder model having a diameter of ~20 Å. The (1 1 0) domain size of CEF of 0 % lignin PF is 31 Å. The discrepancy between the diameter of the cylinder model and the estimated domain size may come from the irregular shape of CEFs.

The interference functions become smeared out with an increase in lignin content in the core region, cf. Fig. 5a. This indicates that adding lignin causes increase in the disorder of CEFs within the fibril matrix.

The arrangement of CEFs within the skin region is different from core. Fig. S9, shows that there is a significant difference in SAXS pattern for the core and skin region of 0 % lignin PF, which disappears for 50 % lignin PF. Hence, in contrast to the core region, the lignin content has less impact on SAXS in “skin”. This could be due to leaching out of lignin from the “skin” during fiber formation [30]. For example, coagulation results in a lignin poor region close to the fiber surface [5]. Thus, “skin” may be composed almost entirely of cellulose for all PFs, regardless of the added lignin content. Continuous electron density contrast between CEFs and microvoids lead to surface scattering of SAXS [17,27,31]. The intensity profiles in the Porod region (Q range from 0.01 to 0.1 Å⁻¹) in the log-log plot of Fig. 5b come from the surface scattering between CEFs and microvoids. Microvoids have elongated morphology [17,31]. The disorder of the fibril matrix causes great polydispersity in the diameters of microvoids. Averaging scattering from these microvoids lead to intensity profiles exhibited as power of Q at Porod region. Average diameters of microvoids could be estimated by radius of gyration (Rg) using a Guinier plot,

$$I(Q) = I(0) / Q \times \exp(-Q^2 R_s^2 / 3). \quad (1)$$

over a Q range of 0.01 to 0.1 Å⁻¹ [25]. As shown in Fig. 5b, the equatorial intensity profiles of SAXS of all PFs show similar distribution regardless of lignin content. From Fig. 5c, the radius of gyration calculated from pattern of core regions has been found grown with increase of lignin content.

## 4. Discussion

### 4.1. The “skin-core” morphology of PFs

A “skin-core” morphology has been found previously for viscose, lyocell and other man-made cellulose fibers [13–16,28]. An increased DPO, and CI, have been attributed to the stress on the fiber surface in the coagulation bath. Using scanning microdiffraction, we have demonstrated that this specific morphology occurs in all PFs from blends of lignin-cellulose. Due to the higher polymer concentration in solution during spinning, the incorporation of lignin into PFs will in this case result in an increase in the fiber diameter from 17 to 25 µm. However, the thickness of the skin gradually reduces with increasing lignin
<figure><img src="image_7.png" /><figcaption>Fig. 4. The effects of lignin blending on transversal and longitudinal dimensions of CEFs. a) Crystal domain size of (0 0 4) reflections for different lignin content. b) Crystal domain size of (1 1 0) and (0 2 0) reflections for different lignin content. Increase of lignin content will reduce both transversal and longitudinal dimensions of CEFs.</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>Fig. 5. a) SAXS data from core regions of PFs from different lignin blends. There is an obvious interference diffraction pattern at 0.15 Å⁻¹ for 0 % lignin (see Fig. S9). The increase of lignin content will smear the interference pattern in the SAXS data. b) SAXS of the skin region of PFs from lignin blends. No significant difference of SAXS patterns is observed for different PFs. c) the Q range of 0.03–0.1 area fitted with a Guinier plot. These results indicate that increase of lignin content will increase the Rg of microvoids.</figcaption></figure>

content. Data in Fig. 2 demonstrate that the thickness of the skin could vary from 2 to 7 µm. To avoid overestimating the thickness caused by the skin morphology and higher CI in the skin region, scans with better resolution (0.25 µm) have been applied to 0 % and 50 % lignin PFs. As shown in Fig. S7, the thickness of the skin region for lignin 0 % PF is less than 5 µm, compared to the diameter of 17 µm. The thickness of the skin region gradually decreases to less than 2 µm for 50 % lignin PF, which has a diameter around 25 µm. The maps of CI and DPO indicate that the cellulose phase is better organized in skin region than in the core for all PFs. Molecular orientation is likely to have an impact on the mechanical properties of CFs. Thus, the thickness of the skin region could affect the performance of CFs.

## 4.2. The effects of lignin content on the cellulose crystallite structure of PFs

We have found that increasing the lignin content will cause substantial reductions in CI and DPO of PFs. Lignin is an amorphous macromolecule that shows no preferred molecular orientation. An increase of amorphous content will reduce the tensile modulus [5]. In addition, increase of lignin content may disrupt the molecular organization of cellulose within PFs. This is consistent with previous studies of cellulose–lignin fibers [9] and cellulose–chitosan fibers [32]. The subtle changes of lattice constant and domain size of cellulose crystallites

indicates that the inclusion of lignin impedes the formation of a homogeneously ordered structure of cellulose. In sum, blending of lignin will increase the fraction of amorphous cellulose within PFs, but the amorphous cellulose content remains lower than lignin content. At a lignin content higher than ~30 %, increase of amorphous content is predominately due to lignin.

## 4.3. The effects of lignin content on mesoscale structure of PFs

SAXS from the core of pure cellulose PF exhibits an obvious inter-fibril pattern at a Q of ~0.15 Å⁻¹. This pattern spreads out with increase of lignin content, indicating an increase in disorder. Increase of inter-fibril distance and disorder of CEF within fibril matrix could lead to a shift and flattening of the interference pattern. Elongated microvoids within the fibril matrix have been reported to have radii varying from 10 to 70 Å for lyocell regenerated cellulose fibers [17]. SEM images of a lignin-cellulose fiber spun from NMMO-solutions fiber also showed micropores [5]. Here, Rg of the microvoids of different PFs were shown to increase in size from ~30 to ~50 Å on addition of lignin. The growth of Rg of microvoid moderates once lignin content reaches 33 %. Maps of Rg in Fig. S10 suggest there are microvoids with random size distributed in the skin of all PFs. This is consistent with SEM studies of the skin region of lyocell fibers [14]. In summary, an increase in lignin content causes disorganization of the CEF matrix and an increase in porosity
within the PFs.

### 4.4. Mechanical properties of PF from different blends

Fiber morphology and fine structure are critical factors impacting the performance of CFs. Regenerated cellulose fibers, having well-ordered structure and high purity, have been considered as potential PFs [1]. While pure cellulose PFs exhibit a highly ordered structure, the conversion rate of carbonization is not ideal. To overcome the problem of low yield rate of pure cellulose PFs, lignin blending has been introduced. The mechanical properties of CFs are strongly correlated with properties of the PFs [10,18]. Mechanical properties of lignin-cellulose PFs may be affected by dissolubility, spinnability, draw ratio etc. [5]. To achieve good solubility, we used ionic liquid, a less toxic solution, to dissolve kraft pulps and kraft lignin. Lignin-cellulose regenerated from ionic liquid solution exhibits a nonlinear variation of mechanical properties with the level of lignin blending. Tenacity decreased significantly from

30 % lignin to 50 % lignin content. This is consistent with our findings on the critical mass fraction for lignin blending based on structural variation of the fibers [9]. Under conditions of 40 °C solution temperature and draw ratio of 6, birefringence and tensile modulus of PFs are reduced with increasing lignin content. The tensile modulus is reduced from 18.5 GPa for 0 % lignin, to 10 GPa for 50 % lignin [26]. This is consistent with the observation that an increase of lignin will lower the degree of preferred orientation of CEF and thereby decrease the mechanical strength of the fiber. The observed decrease in tensile modulus is likely due to the disorder of inter-fibrillar and intra-fibrillar structure of CEF within PFs [26].

### 4.5. The variations of hierarchical structure of PF from different blends

Fig. 6 shows a schematic of the hierarchical structure of lignin-cellulose blended PFs. High spatial resolution of SXM enabled the characterization of the skin-core morphology for all PFs, where the

<figure><img src="image_9.png" /><figcaption>**Fig. 6.** The impact of lignin blending on hierarchical structure of PFs. a) The CEF is composed by cellulose chains and form a fibril matrix within PF. Both CEFs and fibril matrix of pure cellulose PF have fine structure. b) The PF blended lignin will reduce the thickness of skin region. The lignin will impregnate the CEF matrix and cause disorganization of CEF. The cellulose chains within CEF become disorder as well with the incorporation of lignin.</figcaption></figure>
"skin" exhibits enhanced cellulosic crystallinity compared to the core region. As shown in Fig. 6b, increase of lignin content within PFs reduces the thickness of the skin. In the cell wall of plant, lignin content causes the disorder of cellulose fibril matrix [33,34]. For regenerated cellulose fiber, adding lignin content also prevents the CEFs from forming an organized matrix within the PF and causes the disorder of crystal structure of CEF.

## 5. Conclusion

Here, we report studies of the precursor fiber from different blends of lignin and cellulose by SXM. SXM is a powerful technique that enables non-invasive investigation of the hierarchical structure of PFs. Alteration of the multi-scale structure of PFs significantly impacts their mechanical properties, such as tensile strength, degree of orientation etc., which is expected to further affect the mechanical properties of CFs. The X-ray analysis reveals that lignin alters the fiber morphology and the cellulose orientation and crystallinity. Based on the findings, structural features, such as CI and domain size of PFs with different lignin concentrations, ~30 % lignin content could be an optimal point for achieving the beneficial increase in carbon content from lignin while still maintaining a high molecular orientation and crystallinity of the cellulose. Thus, the correlation between lignin content and structural order of PF, revealed in this study, could provide an important guide for the optimal choice of blends to generate suitable PF for the desired CF.

## CRediT authorship contribution statement

Jiliang Liu and Manfred Burghammer design and conducting the experiment; Jenny Bengtsson and Kerstin Jedvert prepare the precursor materials; Jiliang Liu, Yu Shun and Manfred Burghammer develop method and analyze the X-ray diffraction data. Jiliang Liu write the manuscript, all authors review and edit the manuscript.

## Declaration of competing interest

The authors declare that they have no conflict of interest.

## Acknowledgement

Vinnova is gratefully acknowledged for financial support, grant nr. 2020-00831. We gratefully thank Dr. Hanna Ulmefors for her productive comments on our paper. S.Y. acknowledge Swedish Research Council (VR) financial support nr. 2018-06378. This is an ID13 inhouse project.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.ijbiomac.2022.11.211.

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