Structural changes during heterogeneous sulfation
and following homogenization of cotton cellulose

Xijun Wang · Pan Chen* · Yu Ogawa ·
Yoshiharu Nishiyama* · Haisong Qi*

Received: date / Accepted: date

**Abstract** We used to consider native cellulose to be composed of microfibrils continuously crystalline along the chain direction with constant lateral dimension. To sulfate the surface of cellulose microfibrils, we activated cotton fibers by pre-swelling in water or DMF followed by immersion in an excess amount of sulfamic acid in dimethylformamide (DMF) in heterogeneous conditions. The degree of sulfation was followed by elemental analysis and conductometric titration, while the structural changes associated with the sulfation and the homogenization were investigated by infrared spectroscopy, solid-state NMR, and X-ray diffraction. The initial sulfation of the cellulose surface was well described as a first order reaction with a kinetic constant of 2×10⁻³s⁻¹ at 80°C using 3.6% sulfamic, indicating that about half of the available hydroxy groups were sulfated within the first 5 minutes. A water pre-swelling step, followed by solvent exchange with DMF, increased the amount of available hydroxy groups by up to a factor of two. At long reaction times, there was a gradual increase in the degree of sulfation beyond the initially available surface by up to 20% during the two hours reaction investigated. FTIR solid-state NMR spectra both suggest the reaction of surface primary hydroxy groups with sulfamic acid. While the apparent lateral crystallite size of cotton is unaffected by sulfation treatment, X-ray diffraction suggested that homogenization resulted in cellulose nanofibers (CNFs) with a crystallite width below 3 nm, much smaller than the initial cellulose microfibrils (6-7 nm). This apparent crystallite splitting during homogenization suggests that cotton microfibrils have an intertwined polycrystalline structure, rather than being monocrystalline along the whole length.

<footnote>X. Wang · H. Qi qihs@scut.edu.cn
State Key Laboratory of Pulp and Paper Engineering, South China University of Technology, Guangzhou, 510640, China</footnote>

<footnote>P. Chen panchen@bit.edu.cn
Beijing Engineering Research Centre of Cellulose and Its Derivatives, School of Materials Science and Engineering, Beijing Institute of Technology, 100081, Beijing, P.R. China</footnote>

<footnote>Y. Ogawa · Y.Nishiyama yoshiharu.nishiyama@cermav.cnrs.fr
Univ. Grenoble Alpes, CNRS, CERMAV, 38000 Grenoble, France</footnote>
**Keywords** surface sulfation · cellulose microfibrils · cotton · sulfamic acid · degree of substitution · FTIR · NMR · X-ray diffraction

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# 1 Introduction

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We used to consider native cellulose to be composed of unique crystalline structural entities known as microfibrils that typically have nanometer widths (Nishiyama, 2009). This structure is formed due to the concerted polymerization of multiple chains during biosynthesis. There is a strong correlation between the synthase arrangement and the cross-sectional shape of the microfibrils, but the presence of hemicellulose in the synthesis environment also affects the cross-section due to the strong affinity with cellulose, polluting the surface to impede further co-crystallization (Thomas et al., 2013, 2014). In higher plants, all the cellulose synthase complexes observed hitherto are of rosette type constituted of 6 granules (Read and Bacic, 2002), each composed of 3 synthases (Sethaphong et al., 2013). Consequently, the basic unit of cellulose microfibrils in higher plants can be assumed to be composed of 18 chains, although the arrangement of the secondary wall forming synthase has never been visualized. Cellulose from some higher plants, such as cotton hairs or ramie bast fibers, show sharp X-ray diffraction patterns that can only be explained by much larger cross-section, typically 5-6 nm wide corresponding to a much greater number of cellulose chains, on the order of 100 (Nishiyama, 2009). Nevertheless, in the current model, we consider each microfibril to be a single entity that is continuously crystalline along its micron-scale length.

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The structure of cellulose microfibrils, with the inner part of crystal inaccessible to solvent, leads to a possibility of “heterogeneous” chemical modification at the nano-scale, where only the surface of these microfibrils is modified, preserving the chemical and physical stability of the crystalline core while drastically modifying the external chemical properties (Heise et al., 2022; Tardy et al., 2021). One such modification is the introduction of charged groups (Saito et al., 2007; Tong et al., 2021; Noguchi et al., 2017). For example, Lepoutre et al. (1973) grafted polyacrylonitrile onto never dried pulp and hydrolyzed to acrylates, obtaining a highly swollen pulp and a colloidal dispersion of microfibrils. A simpler method was later discovered using TEMPO-mediated oxidation in aqueous media, which selectively oxidizes primary alcohols on the microfibril surface into carboxyl groups (Saito et al., 2007). Gentle mechanical treatments can readily disperse microfibrils into the suspension. Other surface esterification methods introducing charged groups also work in a similar way Sehaqui et al. (2016); Beaumont et al. (2020, 2021).

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A simple sulfuric acid treatment can be used to introduce sulfate half esters onto the surface of cellulose microfibrils, but also leads to hydrolysis of the microfibrils into shorter fragments(Beck-Candanedo et al., 2005; Elazzouzi-Hafraoui et al., 2008). In this case, the resulting fragments are generally lateral aggregates of crystalline elements instead of individual crystals.

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An alternative to sulfuric acid treatment is sulfation in aprotic solvents such as N, N-dimethylformamide (DMF) (Briois et al., 2013; Kuznetsov et al., 2018). Wagenknecht et al. studied sulfation of cellulose in the solution state using an N₂O₂-DMF system, but the same reaction can be carried out in a heterogeneous system by swelling cellulose fibers using DMF, as first reported for sulfation of highly crystalline cellulose for dispersion in water (Wagenknecht et al., 1991).
Heterogeneous sulfation of cellulose fibers results in fibrous colloidal particles with high surface charge, which can be used for many potential applications such as dispersion aids of carbon nanotubes, Mxene, or silver nanowires, (Feng et al., 2022; Wang et al., 2023; Liu et al., 2022; Li et al., 2022), demonstrating the relevance of this process for functional materials.

Although numerous studies have employed this surface modification, the reaction kinetics and the associated structural aspects have not yet been understood. According to the single-crystalline microfibril model, the available reaction sites on the microfibril surface should be determined by the starting sample, and the reaction should follow first-order kinetics. Indeed, our previous study on the sulfation of chitin showed a clear first order reaction to chitin(Wang et al., 2022a). For cellulose fibers, recent studies on TEMPO oxidation kinetics are also based on this hypothesis, but the experimental results do not exactly follow a first order kinetics, similar to the case in the previous studies(Guan et al., 2019; Saito and Isogai, 2004). To gain further insight into the heterogeneous esterification of higher plant fibers, and whether the reaction can be explained as a simple surface modification of single-crystalline microfibrils, we followed the kinetics of sulfation on cotton fibers using DMF-sulfamic acid system and analyzed the structure of products before and after dispersion into water.

## 2 Experimental section

*Materials* Cotton fibers were collected by our laboratory. N,N-dimethylformamide (DMF, 99.5%) and sulfamic acid were purchased from Maclean Biochemical Technology Co., Ltd (Shanghai, China). Sodium hydroxide (NaOH, 99.7%) and Sulfuric acid (H₂SO₄, 98%) were purchased from Guanghua Sci-Tech Co., Ltd (Guangdong, China). DMF was stored under anhydrous conditions using a 3A molecular sieve. All other chemical reagents were used as received.

*Pretreatment* a) DMF swelling. Dry cotton fibers were immersed in DMF with 3A molecular sieves and shaken for 6 hours at 300 rpm at 25°C, followed by filtration to remove the excess DMF. b) Water activation. Dry cotton fibers were placed into a glass bottle with a screw cap, and deionized water was then added to cover the fibers. The bottle was then shaken overnight under the same conditions. Excess water was then removed by filtration. The wet cotton fibers were dispersed in DMF-containing molecular sieves and shaken under the same conditions for 4 hours. The solvent was further exchanged by successive suction and filtration repeated 5 times.

*Sulfation reaction* The sulfation process was carried out based on our previous work (Wang et al., 2022b). Briefly, a flask containing 1.8 g of sulfamic acid in a 50 mL DMF solution was placed in an oil bath and heated to 80 °C. Dry cotton fibers (1.0 g) were then added to the solution. To study the reaction kinetics, the duration of the reaction was varied within the range of 5-120 minutes.

At the end of the reaction, the sulfated fibers were collected by filtration using a PTFE membrane (pore size 0.2 µm). The excess sulfamic acid was removed by resuspending the fibers in fresh DMF (typically 1 g fibers in 50 mL solvent). The fibers were then collected by filtration after stirring for 5 mins, and the operation
was repeated for five times. The sample was then further washed with water to remove DMF. This was done by repeatedly suspending in deionized water followed by centrifugation at 10 000 g for 10 min. In this step, the precipitate was diluted by about 3 times and the operation was repeated 20 times.

**Ion exchange** The $NH_4^+$ was exchanged to $H^+$ by immersing in $H_2SO_4$. First, 0.6 g of sulfated fibers were immersed in 80 mL of 5 mM $H_2SO_4$ solution. The mixture was then shaken at 250 rpm for 2 hours at 25 °C. Subsequently, the acid solution was removed by centrifugation at 7000 g for 10 minutes. For complete ion exchange, this step was repeated 10 times and with a five-fold dilution each time. Finally, the excess $H^+$ was removed with deionized water by repeated centrifugation until the conductivity of the supernatant remained constant at 26 µScm$^{-1}$.

**Homogenization** The sulfated cotton fibers were further diluted to approx. 0.2% and dispersed using a high-pressure homogenizer (model Mini, Noozle Fluid Technology Co., Ltd., Shanghai, China) by passing the suspension through a 200 µm zirconia nozzle at 80 MPa for 5 times. To prevent nozzle blockage due to the increasing viscosity of the suspension, deionized water in varying amounts was added after each homogenization cycle. As a consequence, the concentration of the obtained nanofiber suspension ranges from 0.5% to 0.1%.

**Optical microscopy** The rinsed fibers were put on a glass slide, covered with a coverslip, and observed using a polarizing microscope BX53M (Olympus, Japan) under crossed-polarizers and with the insertion of a retardation plate.

**Elemental analysis** The element contents of carbon, hydrogen, nitrogen, and sulfur were measured using an elemental analyzer (Vario EL cube, Elementar, Germany). Elemental analysis was performed on approx. 5 mg of freeze-dried sulfated sample. The signal intensity was calibrated using sulfanilamide to obtain the element weight fraction. The degree of sulfation, DS, is defined as the number of sulfate groups per glucose residue and was calculated from the weight fractions of carbon and sulfur, $\phi_C$ and $\phi_S$ as

$$DS_{elem} = \frac{\phi_S 6(= \text{number of C/residue}) \times 12(= \text{molar mass of C})}{\phi_C 32(= \text{molar mass of S})} \quad (1)$$

**Conductometric titration** The surface charge density of the CNFs was determined by conductometric titration according to the previous work (Pingrey and Hsieh, 2022). Briefly, 40 mL of 0.5 g/L sulfated CNFs suspension was titrated with 10 mM NaOH, by sequentially adding 0.1 mL aliquots of base solution using a pipette. The number of sulfate groups was estimated from the moles of NaOH required to reach the first equivalence point (i.e. minimum conductivity).

**Fourier-transform infrared spectroscopy** Fourier-transform infrared (FTIR) spectra were recorded using single-bounce attenuated total reflectance on a diamond prism using a Spectrum Two spectrometer (PerkinElmer). Freeze-dried samples kept at ambient conditions were pressed against the diamond prism, and the reflected intensity was compared to the measured intensity without the sample. The registered spectra were analyzed using “specio” and “scikit” Python modules for
spectral decomposition and presentation. The absorption was multiplied by the wave number (ATR correction) values for presentation as an approximation to the transmission-mode measurement equivalence.

**Solid-state NMR spectroscopy** Solid-state ¹³C NMR spectroscopy was performed using a Bruker Avance III 400 MHz spectrometer (¹³C frequency of 100.6 MHz) with 12 kHz magic angle spinning (MAS). The ¹³C chemical shifts were calibrated with respect to that of the glycine carboxyl group (176.03 ppm). The ¹H radio frequency field strength was set to give a 90° pulse duration of 2.5 µs. The ¹³C radio frequency field strength was obtained by matching the Hartman-Hahn conditions at 60 kHz. For cross-polarization (CP), a contact time of 2 ms, 35 ms acquisition, and a recycle delay of 2 s was used. For the single pulse (SP) measurement, the 90° pulse duration was 4 µs the recycle delay was 64 s.

**X-ray diffraction** X-ray diffraction was measured using a point-collimated X-ray beam from a Cu anode X-ray tube mounted on an X-ray generator. Freeze-dried samples were pressed using a hand device with flat edges and trimmed on both sides with a sharp blade to make 1 mm-thick blocks. The thickness direction is perpendicular to the pressing. The sample is mounted on a sample holder with a pinhole so that the incident beam is perpendicular to the pressing. The scattered X-ray was recorded on an imaging plate placed in a vacuum chamber with the sample. The imaging plate, after exposure, was read with a bioimaging analyzer (BAS-1800II, Fujifilm) and numerically treated to obtain X-ray photon counts at each pixel and to obtain line profiles using an in-house program.

**Turbidimetry** Suspension turbidity $\tau$ (cm⁻¹) was determined by UV-vis transmission spectroscopy. In a typical measurement, the suspensions of the nanofibers were diluted to 0.5 mg/mL, and spectroscopy was performed in a quartz cuvette with path length of 1 cm using a UV-2600i spectrophotometer (Shimadzu, Japan) at the wavelengths of 290–900 nm. The fiber mass-length ratio $\mu$ (Da/cm) was estimated by fitting the turbidity spectra to the following equation (Tanaka et al., 2017):

$$ \tau(\lambda) = \frac{88\pi^3 n c}{15\lambda^3 N_A} \left(\frac{dn}{dc}\right)^2 \mu \quad (2) $$

where c is fiber concentration, $\lambda$ is the wavelength, n is the refractive index of the solvent, $N_A$ is Avogadro's number, and $dn/dc$ is the fiber refractive index increment, which was assumed to be 0.135 mL/g in this case. Assuming the square cross-section and density of crystal, the width of the fibril $d_t$ in nm can then be estimated as: (Shimizu et al., 2016)

$$ d_t = 10^{-5} \sqrt{\mu} \quad (3) $$

**Atomic force microscopy** Nanofiber suspensions diluted to about 0.01 % were put onto freshly cleaved mica and dried at ambient temperature overnight. The observation was made using Dimension Icon (Bruker Billerica MA, Germany) in tapping mode.
<figure><img src="image_1.png" /><figcaption>**Fig. 1** Schematic diagram of CNF generation from cotton cellulose through reaction with sulfamic acid followed by homogenization</figcaption></figure>

## 3 Results

### 3.1 Sulfation of cotton cellulose

Fig. 1 shows the process of the CNF generation. First, the cotton microfibrils were pre-treated by DMF or water to improve the activation of the C6-OH groups on the fiber surface. According to our previous work, the sulfation between cellulose and sulfamic acid mainly takes place on the surface of the microfibrils, resulting in cellulose containing negative sulfate half-ester groups (Wang et al., 2022b). To produce CNFs, the sulfated microfibrils were then subjected to a high-pressure microfluidic nanodisperser. Due to the electrostatic repulsion provided by the sulfate half-ester groups, the individual nanofibers were colloidally stable in water.

The fibers show some fibrillation and ruptured ends but roughly conserve the integrity of the fiber morphology of cotton fibers after preparation by cutting mill, and have the morphology of a collapsed tube typical of cotton fibers (Fig. 2a). After sulfation and rinsing with water, the width of the fibers hugely increases, and less collapse can be observed (Fig. 2b-f), but the integrity of the fibers is intact.

### 3.2 Evolution of DS as a function of time

Fig. 3 shows the degree of substitution as a function of reaction time. Systematic differences between DS values obtained from titration and elemental analysis were observed, which could be attributed to the different combustion efficiency in the elemental analysis with respect to the sulfanilamide we used as a standard. A similar observation of higher DS values using elemental analysis compared to conductometric titration and the number of accessible hydroxy groups was also observed in our previous study on chitin sulfation (Wang et al., 2022a).

As the sulfation of the surface hydroxy groups occurs in a large excess of sulfamic acid, the reaction is expected to follow first order kinetics. The first three
<figure><img src="image_2.png" /><figcaption>Fig. 2 Rinsed cotton fibers (a) and fibers sulfated for 5 min (b), 15 min (c), 30 min (d), 60 min (e), and 120 min (f) after water activation. Observation under crossed-polarizer and a retardation plate.</figcaption></figure>

DS values (up to 30 min) were therefore used to fit a first order kinetic function:

$$DS = A(1 - \exp(\kappa t)), \qquad (4)$$

where $\kappa$ is the kinetic constant and A the fraction of reactive site per residue. The fitting curves are indicated by solid lines in Fig. 3. The kinetic constants were mostly in the range of 0.12-0.13 min⁻¹ suggesting that half of the initially available hydroxy groups are sulfated in the first 5 minutes under the reaction conditions used (3.6% sulfamic aid, 80°C.) At long reaction times (beyond 50 min), the degree of sulfation does not appear to level off, but instead slowly continues to increase, suggesting additional reaction processes.

## 3.3 Infrared spectra

The FTIR spectra of cotton fibers with increasing reaction time (Fig. 4) show intense peaks at 812, 867, 1223, and 1420 cm⁻¹, consistent with the presence of sulfate half ester groups (Feng et al., 2021; Pingrey and Hsieh, 2022). Furthermore, the peak at 895 cm⁻¹ commonly assigned to the surface primary hydroxy groups of native cellulose (Åkerholm et al., 2004) disappears in the early stages of the reaction, especially for samples activated by water. For the DMF-activated samples, the sulfate peaks increase monotonically with reaction time, whereas for the water-activated samples, some of the sulfate peaks (812, 867, 1223, 1420, 3050 cm⁻¹) reach a maximum and decrease in water-activated samples, while the peak
<figure><img src="image_3.png" /><figcaption>**Fig. 3** Degree of substitution as a function of reaction time measured by conductometric titration (blue circles) and elemental analysis (orange circles) for water-activated (a) and DMF-activated (b) samples. The solid curves indicate fitting to a first-order kinetic equation using data up to 30 min.</figcaption></figure>

at 2485 cm⁻¹ continued to increase. This peak at 2485 cm⁻¹ is less pronounced in the DMF-activated samples.

After sample homogenization, the spectral features significantly changed for all samples. The peaks at 1420, 2485, and 3050 cm⁻¹ disappeared, while the peak at 895 cm⁻¹ reappeared (Fig. 5). The appearance of the 895 cm⁻¹ peak suggests the creation of a significant amount of new surfaces.

## 3.4 NMR spectra

Fig. 6 shows solid-state NMR spectra for the cotton fiber samples in dry-state, wet-state, sulfated for 5 min with water activation, and after homogenization, normalized by the integration of the whole signal intensities. As previously reported, wetting the cotton results in the appearance of distinct relatively sharp peaks in the high-field components of C4 (83-85 ppm) and C6 (61-63 ppm) as-
<figure><img src="image_4.png" /><figcaption>**Fig. 4** Infrared spectra of a) DMF-activated and b) water-activated cotton as a function of reaction time and c) non-negative factorization of the whole spectra series, and d) subspectra calculated from c) calculated as A-0.4B-0.12water and B-0.8A-0.08water. The subplot shows the zoom of the 800-900 cm⁻¹ region.</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 5** Infrared spectra of homogenized sulfated cellulose from a) DMF-activated and b) water-activated cotton as a function of reaction time.</figcaption></figure>

signed typically to the signals from the crystal surface (VanderHart and Atalla, 1984). The high-field component of the C6 at 61.2 ppm decreases in the first stage of sulfation at 5 min, corroborating the findings from FTIR spectra, and indicating sulfation on the primary hydroxy groups.

Fig. 7 shows the SP-MAS spectra representing the mobile part of the system at different stages. The general feature of the unmodified cotton is similar to α cellulose reported by (Tang et al., 1999), with the C6 at around 62 ppm becoming less prominent when sample is wet. This mobile C6 drastically decreases after sulfation, and a new peak emerges at 68.5 ppm, probably corresponding to the C6 with the sulfation on O6. Again, there were no peaks corresponding to the sulfation of O2 and O3. Note that the C2 and C3 peaks in native cellulose occurred between 70 and 80 ppm.

X-ray diffraction gives anisotropic patterns when the incident beam is parallel to the sample, as the cellulose chains tend to lie perpendicular to the compression direction, causing the 1-10, 110, and 200 equatorial reflections to appear stronger in this direction. There is little change in the diffraction pattern after sulfation, but after homogenization the diffraction pattern became very blurred. Line scans along the compression direction (horizontal) are shown in Fig. 8 with a fitting assumed Gaussian peak profiles. The diffraction profile in the equatorial direction after homogenization cannot be well fitted with crystalline peaks of native cellulose and
<figure><img src="image_6.png" /><figcaption>**Fig. 6** 13C CP/MAS NMR spectra of a) dry, and b) wet cotton and fiber samples after c) 5 minutes or d) 120 minutes sulfation, and freeze-dried sample after homogenization of the 5-minute sulfated sample.f)-i)are difference spectra b)-a), c)-b), d)-c) and e)-d).</figcaption></figure>

<table><caption>**Table 1** CNF width versus reaction time for DMF- and water-activated cotton cellulose fibers, estimated by turbidimetry.</caption><tr><td>reaction time (min)</td><td>5</td><td>15</td><td>30</td><td>60</td><td>120</td></tr><tr><td>water activation (nm)</td><td>9.7</td><td>8.4</td><td>6.7</td><td>5.4</td><td>4.1</td></tr><tr><td>DMF swelling (nm)</td><td>11.2</td><td>9.8</td><td>8.1</td><td>6.6</td><td>5.7</td></tr></table>

a monotonic background only. Instead, the fitting requires an additional broad peak component centered at around 20° (corresponding to a d-spacing of 0.44 nm). Even when this broad component is added to the model function, the line broadening of the crystalline peaks corresponds to a crystallite size smaller than 3 nm.

The average CNF widths estimated from turbidity spectra (Carr Jr. and Hermans, 1978; Tanaka et al., 2017) are shown in Table 1. The estimated width of the nanofibrils is systematically larger than the crystallite size calculated from the X-ray diffraction line broadening, suggesting a bundling of multiple fibrils into one fibrillar object at the nanoscale.

The typical height of nanofibers observed with AFM was about 4 nm (Fig. 9), irrespective of the sulfation time, which is the limit size estimated by turbidimetry. It is unclear if the nanofibers have similar widths and heights due to the limited
<figure><img src="image_7.png" /><figcaption>**Fig. 7** 13C SP/MAS NMR spectral changes by wetting, sulfation, and homogenization</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>**Fig. 8** X-ray diffraction of a) starting cellulose, b) sulfated for 5 minutes after water activation, c) homogenized. The compression direction of the sample is horizontal on the diffraction pattern. The intensity profile in the horizontal direction is shown as a function of scattering angle (°) overlaid on the diffraction pattern with a peak fitting assuming Gaussian profiles.</figcaption></figure>

resolution and if the morphology observed in the limited zone is representative of
the suspension since selective migration can occur during the sample preparation.

## 4 Discussion

*The site of reaction and accessibility* Water activation followed by a solvent ex-change is widely used to increase the reactivity of cellulose from the solid phase. It is believed to expose more hydroxy groups that can then react in the non-aqueous medium. Although DMF and water are expected to swell cotton fibers to a similar degree, the reactivity of water-activated cotton fibers is significantly higher
<figure><img src="image_9.png" /><figcaption>**Fig. 9** AFM height images of nanofibers obtained by homogenization of cotton fibers sulfated for (a) 5 min, (b) 15 min, (c) 30 min (d) 60 min (e) 120 min after water activation. The inset in the bottom left corner of the image corresponds to the height profile of the line on the figure, with the peaks labeled corresponding to the numbers on the image.</figcaption></figure>

than for DMF-activated fibers, exposing 10% more hydroxy groups according to elemental analysis or 50% more according to conductometric titration. This effect of water swelling on the cellulose structure can be readily seen in the solid-state NMR spectra difference in dry- and wet-state, where the C6 resonance peak at 65 ppm decreases and 61.2 increases on wetting. The peak at 65 ppm is commonly attributed to the *tg* conformation of O6, which is unique in the crystalline phase of native cellulose(Horii et al., 1983; Briois et al., 2013). The resonance up-field at 61 ppm is attributed to the more relaxed *gg* conformation on the surface(Horii et al., 1983). Wetting might also increase the mobility of the surface hydroxy groups that would reduce the CP-MAS signal, so the spectral change gives the lower bound of the actual change in conformation.

When the sample was sulfated for 5 minutes after solvent exchange from water, the peak at 61.2 ppm was drastically reduced, indicating that the surface primary hydroxy groups were consumed in the first sulfation reaction. This observation is consistent with the FTIR spectra, where the 895 cm⁻¹ band, assigned to the primary alcohol of the surface chains, almost disappears in the first 5 minutes of reaction, similar to the 61.2 ppm signal in the NMR spectra. The carbon next to the sulfated hydroxy groups shows a 5-6 ppm downfield shift to the unsubstituted version in solution. When sulfated, the C2 and C3 peaks are expected to appear in the 80-81 ppm region. No intensity change can be observed in this region, while 68-69 ppm signal is increased after sulfation, as seen by the negative peaks in Fig. 6g and 6h. Thus, it seems that the sulfation happens predominantly at the O6 position of the microfibril surface. In the initial step, the surface O6 at 61 ppm is decreased,
<figure><img src="image_10.png" /><figcaption>**Fig. 10** Schematic illustration of possible internal structures of a microfibril where a chain can be on the crystal surface for a segment and in a crystalline core for another segment. A) cross- over between an elementary unit and a neighboring unit. B) twisted elementary unit co-crystallizing with neighboring ones. The crystallite size, as estimated from the X-ray peak broadening, is larger than the elementary unit.</figcaption></figure>

but at prolonged sulfation, O6 that was initially in *tg* conformation is partially consumed as the positive peak of 6h attests. This explains the departure from the first order kinetics, which assumes a fixed number of reactive sites; instead, new reactive sites are slowly created during the prolonged reaction.

The crystallite size estimated from the cotton sample is about 6-6.5 nm from the broadening of X-ray diffraction, corresponding to about 10-12 chains along each side. The microfibril is expected to have exposed (110) and (1-10) surfaces with chain spacing of about 5.4 and 6 Å(Nishiyama, 2009). One can consider a very simple squarish cross-sectional model constituted of 10 times 12 chains. In this model (10-1+12-1)x2 chain sides are exposed on the surface for 120 chains. For each side of the chain, O6 is pointing out every other residue from the surface so that the accessible O6 would be 20/120 = 16.7% of the O6 groups in the whole cross-section. A limiting value of DS = 0.17 is expected in this case, which is slightly lower than the value 0.22 of water-activated cotton evaluated by conductometry and higher than 0.15 for DMF activated version. It is overall in agreement with the picture of crystalline microfibril with the significant difference in reactivity or accessibility between crystalline surface and interior.

*The meaning of lateral size decrease by homogenization* The most striking results are the considerable decrease in crystallite size and the creation of new unsulfated surfaces observed after homogenization. This is suggested by the reappearance of 895 cm⁻¹ peak in infrared spectra of all sulfated samples, a significant decrease of crystalline C4 signal at 89 ppm in CP-MAS spectra, and significant line broadening of X-ray diffraction. All those experimental results point to the splitting of
microfibrils into much smaller fibrils similar to wood cellulose. Indeed, the behavior
is very similar to the TEMPO-oxidized wood pulp, where diffraction broadening
can be observed after mechanical dispersion (Daicho et al., 2018). In the case of
wood pulp, the discussion was based on the fact that in the intact cell wall, wood
microfibrils were composed of 2-3 nm wide microfibrils. The microfibril would
stick against neighboring microfibrils during the pulping process, where lignin and
some hemicellulose were removed, leading to a larger apparent crystallite width.
In the case of cotton, the microfibril width is already significant in the nascent
state without being dried (Hirai et al., 1990; Tsuji et al., 1992). Still, the basic
unit of biosynthesis might be the 18 chains that co-crystallize in the absence of
other substances in the cell wall that can be separated by mechanical treatment
after loosening due to surface modification. If the surface chains are always on
the surface throughout the whole chain length, there would be modified chains on
the surface and unmodified chains in the core. This would lead to a dispersion
of microfibrils without a decrease in lateral size. The sub-fibrillation would imply
that a chain can be on the surface at one locus and inside the crystalline core at
another as described in Figure 10. This also corroborates with the fact that when
surface esterified, there seems to be no distinct population of esterified chains and
non-esterified chains (Beaumont et al., 2021)

5 Conclusion

The reaction of sulfamic acid in DMF with cotton fibers follows first-order kinetics with respect to the available surface in the first step of the reaction. Sulfation extends further into the fiber structure but with little impact on the crystal structure of the native cellulose. Mechanical homogenization of the sulfated fibers leads to a suspension of CNFs crystallite width below 3 nm, much smaller than the initial microfibrils (6-7 nm). This suggests an internal structure of microfibril in which a single chain along its length can alternate its position in the cross-section, at the surface, and inside the crystalline core.

**Acknowledgements** The authors are grateful for the financial support for this work by the Guangdong Basic and Applied Basic Research Foundation (2020A1515110594, 2023B1515040013) and State Key Laboratory of Pulp & Paper Engineering (2022C01). P.C. acknowledges financial support from the Beijing Natural Science Foundation (2232064) and the National Natural Science Foundation (52373097). We also thank Dr. Weiying Li from SCUT/State Key Laboratory of Pulp and Paper Engineering for the elemental analysis. We thank Dr. Thomas Parton from the University of Cambridge for fruitful discussions and advice on the manuscript.

Conflict of interest

The authors declare that they have no conflict of interest.

6 Ethics approval and consent to participate

Not applicable.
7 Consent for publication

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All authors agree to publish.

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8 Availability of data and materials

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All original data are available at https://github.com/yoshi-CERMAV/cellulose-2023 together with the python code for the data treatment. The corresponding Jupyter notebook is "https://github.com/yoshi-CERMAV/cellulose-2023/cellulose2023.ipynb".

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9 Competing interests

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The authors declare no competing interest.

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10 Funding

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China Guangdong Basic and Applied Basic Research Foundation (Q.H.: 2020A1515110594, 2023B1515040013), China State Key Laboratory of Pulp & Paper Engineering Grant(Q.H.: 2022C01), Beijing Natural Science Foundation (P.C.: 2232064), National Natural Science Foundation (P.C.: 52373097).

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11 Authors' contributions

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Y. N., Q. H., and P. C designed the experiments. X.W. prepared the samples, carried out the kinetics study, and turbidimetry, Y. O. carried out the solid-state NMR measurement, Y. N. carried out X-ray and FTIR measurements, analyzed the data, and wrote the manuscript. All authors reviewed the manuscript.

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