Cryo-ET comparison of the hierarchical ultrastructure of silkworm, spider, and artificial silk fibers

Domain: chemistry
Published: 2026-03-07
Editorial outcome: minor_revision

Abstract
Spider and silkworm silks are renowned for their exceptional mechanical properties, which arises from their
ultrastructural organization. However, this architecture remains incompletely understood. Here, we apply cryo-
electron tomography to examine the hierarchical organization of silkworm, spider, and artificial silks. In
silkworm silk, we observe nanofibrils of ~3.6 nm in diameter, interconnected by abundant bridges and
representing the smallest fibrillar features currently accessible by cryo-ET. These nanofibrils align with the fiber
axis and are organized into a herringbone pattern, with stacked layers building the micron-scale filament. Spider
silk displays densely packed nanofibrils with near-perfect axial alignment and minimal voids. In contrast,
silkworm silk shows regionally heterogeneous gaps, whereas artificial silk lacks the ordered packing
characteristic of natural materials. These observations provide a structural basis for understanding silk formation
and may guide future biomimetic fiber design.

Introduction
Natural silk, primarily derived from silkworms and spider major ampullate gland, exhibits excellent mechanical
properties, including high tensile strength and toughness. This gives them not only tensile strength comparable to that
of high-performance steel or Kevlar but also exceptional toughness1-3. For millennia, silk has been prized in textiles
for its smooth texture, luster, and strength. More recently, due to its extraordinary mechanical properties, as well as
biocompatibility, biodegradability, and eco-friendly manufacturing methods, silk fibers are becoming a candidate
material for biomedical engineering4-7, flexible electronic wearables8,9 and data storage medium10, attracting
increasing attention11-20.

The superior mechanical properties of silkworm silk and spider silk are closely tied to their ultrastructure,
specifically the morphology, size and hierarchical assembly of the fundamental building blocks within the fibers.
Decades of research have focused on understanding how silk proteins, stored in a soluble state in the silk glands,
transform into insoluble solid fibers with distinct mechanical properties. Two primary models, liquid crystal
spinning21-25 and micellar spinning26, have sought to explain this process. The liquid crystalline model proposes
that spinning dopes in the spider gland and duct adopt a nematic phase, characterized by rod-like structures that
are considered aggregates of spherical silk
proteins21,23. In contrast, the micelle model suggests that natural silk
fibroin (NSF) forms micelles, approximately 100–200 nm in diameter, in solution due to its amphiphilic primary
sequence26,27. As the NSF concentration increases, these micelles coalesce into globules ranging from 0.8 to 15
µm in diameter, which subsequently align under shear forces to form fibers26,28.

Despite these models, the precise structural units and assembly process of silk proteins in silk glands remain
largely elusive. Our recent studies29 have shown a multi-scale controllable self-assembly spinning mechanism
based on the NSF in silkworm spinning dope, suggesting that flexible nanofibrils may represent key building
elements, approximately 4 nm in diameter, rather than the previously suggested micellar structure26,28,30,31 or rod-
like structure formed by aggregation of globular fibroin molecules23. Within the silk glands, these fibroin
nanofibrils initially exhibit random orientation, but intriguingly, they align into a herringbone pattern in the
anterior silk gland near the spinneret. This raises questions about whether this ordered structure persists in the
final silk fibers or undergoes further transformation during the spinning process. Thus, it is essential to
investigate the ultrastructure of silk fibers for a comprehensive understanding of the spinning process.

In addition to natural silk fibers, artificial silks have attracted increasing interest due to their potential for scalable
production and customizable properties. Artificial silk is typically produced by regenerated silk fibroin16,32 or
recombinant silk proteins13-15,18,20 using techniques such as wet spinning13,15,16,18-20,32, dry spinning31,33,
electrospinning14, interface wiredrawing34 or microfluidic spinning35,36. These processes aim to fabricate
artificial silk fibers with structures and properties comparable to those of natural silk, but the properties of
resulting artificial silk fibers are particularly influenced by factors such as protein sequence, spinning dope
concentration, flow rate, humidity, post-spin stretching, buffer composition, etc11,12,33. The ultrastructure within
artificial silk is even less understood. Understanding these structural discrepancies is crucial for improving
artificial silk production and bridging the performance gap between synthetic and natural materials.

Investigating the ultrastructure of silk fibers from the atomic to the macroscopic scales remains a major challenge.
At the atomic level, previous studies using X-ray diffraction (XRD), Raman spectroscopy, neutron scattering
and solid-state nuclear magnetic resonance have established that silk is a semicrystalline polymer. Its crystalline
regions are primarily composed of β-sheets formed by polyalanine, contributing to silk's strength, while the
amorphous regions, rich in glycine, provide the toughness of the silk fibers37-41. However, a consensus on the
hierarchical structure of silk from the nanoscale to the macroscale is still lacking. Various models based on
atomic force microscope (AFM)42-47, scanning electron microscope (SEM)48 and Anderson light localization49
suggest the presence of nanofibrils with diameters ranging from 20 to 220 nm. Nevertheless, techniques like
transmission X-ray microscopy50 and ultramicrotomy51 have failed to detect nanofibrils, with some studies
indicating silk fibers consist of spherical units, 10-200 nm in diameter, the size of which varies depending on the
type of silk26,30,51-53. Neutron scattering and small-angle XRD40,54 suggest a long-range organization in silk fibers,
although its nature and relationship to the protein sequence remain unclear.

Despite significant progress, many aspects of the supramolecular structure and assembly process of silk remain
unresolved. One major challenge is the lack of suitable techniques for studying the native structure of silk fiber
in vivo. Furthermore, conventional sample preparation methods, such as chemical fixation, dehydration, resin-
embedding and heavy metal staining, often introduce artifacts that result in overly condensed structures,
obscuring the fundamental units of silk. The intrinsically dense structure of silk further complicates efforts to
achieve its high-resolution ultrastructural characterization.

In this study, we combined cryo-focused ion beam (cryo-FIB) milling with state-of-the-art cryo-electron
tomography (cryo-ET) to investigate the ultrastructure of force-drawn silk fibers from silkworms, spiders, and
that of artificial silk. Cryo-FIB is a cutting-edge technique used to prepare high-quality samples for structural
and ultrastructural analysis, particularly in the context of cryo-ET. This method preserves the native structure,
enables high-resolution imaging by producing ultra-thin sections, and minimizes artifacts typically associated
with conventional sample preparation methods. Our study uncovered the morphological and hierarchical
structure of fibroin within silk fibers with unprecedented level. The results revealed that the fundamental
structural unit of silk is nanofibrils approximately 3.6 nm in diameter, which share the same morphology as those
found in the silk gland. These nanofibrils are arranged in parallel along the fiber axis, forming a herringbone
pattern. Multiple layers of this herringbone pattern stack to form the final micron-scale fiber, indicating that the
herringbone pattern formed near the spinneret remains stable during fiber formation. In spider silk, a more
condensed structure adorned with nanofibrils is clearly observed, with their long axes tightly aligned in parallel
along the flow direction, leaving no visible gaps. In contrast, silkworm silk exhibits larger, regionally varying
gaps between nanofibrils, while artificial silk lacks the highly ordered arrangement found in natural silks. In
summary, gaining a deeper understanding of silk's ultrastructure provides valuable insights into replicating and
improving upon the natural spinning processes of silkworms and spiders, guiding the development of polymer-
based materials with tunable properties.

Results
Structure of fibroin extracted from silk glands of silkworm
Fibroin, a key component of silk, is understood to exist either as a spherical macromolecular complex with a
molecular weight of up to 2.4 MDa, composed of a fibroin heavy chain, light chain, and P25 in a molar ratio of
6:6:155, or as a rod-like structure formed by the elongation and aggregation of globular fibroin23,30,52. In this study,
NSFs were extracted from the lumen of posterior silk gland of silkworms. Gel filtration chromatography and 4-
16% SDS-PAGE analysis showed that NSFs are a macromolecular complex formed by the fibroin heavy chain,
light chain, and P25 subunits (Supplementary Fig. 1a). Interestingly, 3-16% BN-PAGE analysis showed two
distinct bands for freshly extracted NSFs, with higher molecular weight bands gradually transforming into lower
molecular weight bands upon the addition of amphipol. This transformation displayed a dose-dependent
relationship with amphipol concentration (Supplementary Fig. 1b). Metal shadowing analysis confirms that
(Supplementary Fig. 1c-e).

For improved sample uniformity, NSFs were treated at a 1:6 molar ratio for 72 hours (Supplementary Fig. 1b)
and rapidly frozen via plunge-freezing. Cryo-ET revealed that NSFs exhibit typical fibrous protein
characteristics, displaying high flexibility with a diameter of 4.0 ± 0.6 nm and a length of 32.7 ± 16.4 nm (mean
± SD) (Fig. 1, a-c). While the diameters of individual nanofibers were highly uniform, the lengths varied
significantly (Fig. 1c). Subtomogram averaging further revealed that fibroin nanofibers appear as a string of
beads structure (Fig. 1d, Supplementary Fig. 2), and metal ions are essential for maintaining this structure29.

Most studies suggest that the spherical structures in nanofibrils represent either fibroin monomers 23,56 or fibroin
aggregates26,30,31,52,57-59. The fibroin heavy chain consists of 12 repeat sequences and 11 linker sequences, with
the repeat sequences containing a high proportion of GAGAGS and GAGAGY motifs that govern the mechanical
properties of silk fibers, and the linker sequences showing high conservation (Supplementary Fig. 3), which
provides evidence for the concerted evolution of silk protein genes60,61. Since AlphaFold 3 can only predict
polypeptide structures up to 5000 amino acids62, we deleted one repeat and one linker sequence from fibroin
heavy chain (Δ2303-2621) to predict its structure. The results indicated that the fibroin heavy chain (Δ2303-
2621) can fold into a spherical structure measuring approximately 10.6 nm × 10.7 nm × 15.1 nm (Fig. 1e), which
is significantly larger than the 4 nm × 4 nm × 4 nm spherical structures observed in nanofibrils (Fig. 1d). Due to
the absence of homologous sequences, the three-dimensional (3D) structure of fibroin heavy chain predicted by
AlphaFold 3 lacked accuracy and confidence. Therefore, while this model does not accurately represent the true
structure of the fibroin heavy chain, it provides a useful reference for estimating the size of fibroin. That is, if
the fibroin heavy chain is folded into a tight spherical structure, its size is approximately 10.6 nm × 10.7 nm ×
15.1 nm, while the actual spherical structure in individual nanofibrils is much smaller than predicted size.
Therefore, we hypothesize that the smaller globular structures observed in nanofibrils represent individual
domains formed by segments of the fibroin molecule. Unfortunately, due to the flexibility of NSF nanofibrils,
obtaining high-resolution 3D structures through single-particle reconstruction remains challenging.
Consequently, it is still unclear how many spherical domains an individual fibroin nanofibril consist of.

Structure of fibroin in silkworm silk
To determine whether the morphology of NSF nanofibrils changes during silk formation, we reconstructed the
morphology and organization of silk proteins within silk fibers from third-instar silkworm. Using cryo-FIB
milling, the lamella with the thickness ≤ 100 nm were qualified for decreasing the overlapping signal in the 2D
image during the data collection (Supplementary Fig. 4, a-b). The lamella was transferred to a Titan Krios cryo-
electron microscope (cryo-EM) for data collection (Supplementary Fig. 4c). In the fibroin region inside the silk
fibers，nanofibrils are oriented parallel to the fiber axis, with consistent widths, whereas the sericin region lacked
a distinct nanofibril structure (Fig. 2, a-b, Supplementary Fig. 4d). Interestingly, an unknown macromolecular
structure was observed in the sericin region (Fig. 2b), but this macromolecular is absent in the sericin region of
cocoon silk. It is therefore speculated that this structure might be a specific macromolecule expressed during the
early larval stage of silkworms. Despite the overall parallel arrangement of nanofibrils in the fibroin regions, we
found that the nanofibril distribution was uneven, with tightly packed areas and looser regions (Fig. 2, a-b). The
imperfect parallel arrangement of nanofibrils resulted in gaps within the silk fibers, which could be a contributing
factor to fiber fractures during stretching63,64. In the densely packed areas, nanofibrils were laterally associated,
forming bridges (Fig. 2, c-f) that may be relevant to mechanical coupling between neighboring nanofibrils.

A small set of nanofibrils were picked from the tomogram, aligned, and averaged (Supplementary Fig. 5a), We
found that the fibroins in silk also appeared as nanofibrils with diameters of 3.6 ± 0.8 nm and lengths of 81.9 ±
38.5 nm (mean ± SD) (Fig. 2, g-h), morphologically similar to those extracted from silk glands in vitro (Fig. 1d).
The results indicate that the morphology of fibroin nanofibrils appears broadly similar as they move through the
lumen of silk gland to form silk fibers. Our measurements likely overestimate nanofibril length, as tightly
connected nanofibrils cannot always be distinguished. Notably, such small diameters for nanofibrils have not
been previously reported for silk.

Structure of silk protein in spider silk and artificial silk
There are significant differences in mechanical properties between silkworm silk and spider silk, particularly
spider dragline silk, which surpasses silkworm silk and even outperforms the best synthetic fibers like Kevlar65.
Using metal shadowing detection, we found that the morphology of natural spidroin (NSP) in the lumen of the
spider’s major ampullate gland (Araneus ventricosus) closely resembled the fibroin of silkworms, appearing as
a string of beads (Fig. 3, a-c). However, the molecular weight of NSP was lower than that of silkworm NSF (Fig.
3d). Recent studies have shown that the silk from major ampullate glands of spiders contain 18 proteins66.

To further investigate the relationship between the ultrastructure of animal silk and its mechanical properties, we
manually pulled fresh silks from the spider and used cryo-FIB milling to prepare lamella ≤ 100 nm thick
(Supplementary Fig. 6, a-c). Then, the samples were transferred to a cryo-EM for high-resolution 3D imaging
(Fig. 3e, Supplementary Fig. 6d). Cryo-EM revealed numerous nanofibrils arranged parallel to the long axis of
the spider silk fibers. Unlike the loose arrangement of nanofibrils in silkworm silk, the nanofibrils in spider silk
were tightly packed and well-oriented with no visible gaps (Fig. 3f), resulting in a much denser fiber structure
with higher nanofibril density compared to silkworm silk (Supplementary Fig. 7a). This tight packing may
explain why spider silk exhibits superior mechanical properties over silkworm silk.

We also prepared artificial silk fibers using the interface wiredrawing method34 (Supplementary Fig. 8, a-b) and
obtained lamella ≤ 100 nm thick via cryo-FIB milling (Supplementary Fig. S8, c-d). In striking contrast, although
both natural and artificial silk are composed of NSF, artificial silk exhibited minimal nanofibril orientation and
a non-uniform phase distribution (Fig. 3h, Supplementary Fig. 8e). These findings suggest that the buffer
composition during spinning significantly affects the orientation degree and density of the nanofibrils within silk
fibers, and these factors play a crucial role in determining their mechanical properties.

Structural organization of silkworm silk
To investigate whether silk fibers contain ordered structures similar to or higher than the herringbone pattern29
observed near the spinneret in the anterior silk gland，we introduced a deep learning strategy-based method
(REST) to establish the relationship between low-quality and high-quality density, applying this knowledge to
restore signals in cryo-ET (Supplementary Fig. 5b)67. The results show that this method improves the signal-to-
noise ratio of tomographic slices and reduces some artifacts caused by the excessive density of silks and the
missing data region in the 3D reconstruction of sample (missing wedge) (Fig. 4, a-b). Notably, we observed
numerous regularly arranged dot-like structures in the XZ view of the silk fibers (Fig. 4, a-b, bottom). By tracking
each dot along the y axis, we determined that each dot corresponds to a nanofibril, which were densely stacked
on the YZ view (Fig. 4, c-d). This arrangement is consistent with the herringbone patterns found near the
spinneret in the anterior silk gland (Fig. 4e)29. Previous studies29,68 identified an axis within the herringbone
pattern, possibly a pseudo-axis formed by interactions involving the N-terminal domain of the fibroin heavy
chain of fibroin. In our experiments, we observed both tightly connected regions and blank regions within the
same herringbone pattern. Interestingly, a distinct axis perpendicular to the nanofibrils was also evident in the
blank regions of the herringbone pattern (Supplementary Fig. 9a), indicating that there may be other proteins
involved in the formation of the herringbone pattern.

Recent studies propose that the herringbone patterns of fibroin may actually resemble a 3D bottlebrush
structure68. Statistical analyses of the distances between points along the z-axis and x-axes revealed that the
spacing between points along the z-axis is significantly smaller and more uniform than that along the x-axis
(Supplementary Fig. 7b). This finding indicates that the nanofibrils within the silk are anisotropically arranged
rather than simply stacked in parallel. Such an arrangement contributes to enhancing the mechanical properties
of the material69. Moreover, they are organized into a herringbone pattern, where multiple layers of this
herringbone pattern are stacked along the x-axis to ultimately form micro-sized silk fibers (Fig. 4f). Obviously,
the final microscale fiber does not exist as a compressed bundle of aligned nanofibrils, and no higher-order
organization was observed beyond the herringbone pattern. This finding suggests that the herringbone pattern,
formed near the spinneret in the anterior silk gland, remains intact throughout silk fiber formation and is not
replaced by more complex arrangements. Notably, the anisotropic arrangement likely affects the mechanical
properties of silk fibers in the x and z directions.

Methods
Ethical Statement
No ethical approval was required for this study, as silkworms (Bombyx mori) and spiders (Araneus
ventricosus) are not subject to institutional animal care and use regulations.

Experimental animal preparation
Silkworms (Bombyx mori) were reared to the third larval stage at room temperature, fed a diet of mulberry leaves.
Twenty adult Araneus ventricosus spiders (body length 1.1-1.7 cm) were collected from the wild in Sichuan and
Beijing, China, between August and October. The spiders were housed individually in separate containers and
provided with moths and water.

Preparation of natural silk fibroin, natural spidroin, and cryo-vitrification
Silkworm larvae were reared at room temperature until reaching the early wandering stage, after which they
were frozen using liquid nitrogen and stored at -80 ºC. Silk glands were dissected from the frozen silkworms,
and the cells on the surface of the posterior of middle silk gland (PMSG) were manually peeled off using forceps.
Then, the spinning drop was transferred to the extraction buffer (50 mmol·L-1 NaCl, 50 mmol·L-1 Bis-Tris, 0.5
mmol·L-1 EDTA, pH 7.5) to dissolve the NSF, resulting in a NSF solution. NSF solution was mixed with
amphipol in extraction buffer at a mass ratio of 1:6. After incubation at 4ºC for 72 h, 3 μL of the mixture was
loaded onto a glow-discharged holey grid (GIG, 1.2/1.3, Au) and vitrified by flash-plunging into liquid ethane
using a Vitrobot Mark IV (FEI, USA). Blotting time, force level and humidity were set to be 7 s, 0 and 100%,
respectively.

Adult spiders were anesthetized at 4℃ for 30 min and then dissected on ice to isolate the major ampullate glands.
The silk glands were rinsed three times with phosphate buffered saline (PBS), and the outer epithelial cell layer
was carefully removed using fine forceps. The gel-like spinning dope was gently extracted and transferred into
PBS to prepare the NSP solution.

Biochemial analysis of natural silk fibroin and natural spidroin
The homogeneity and composition of NSF were assessed by size-exclusion chromatography (SEC) and sodium
dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). SEC was performed using a Superose 6
increase 3.2/300 column (GE Healthcare, USA). Protein fractions were subsequently analyzed by SDS-PAGE
and visualized by Coomassie Brilliant Blue staining.

The component of NSP was examined by SDS-PAGE. A homemade linear gradient SDS-PAGE gel (3%-16%)
was prepared using a Hoefer SG 30 gradient maker (Thermo Fisher Scientific, USA). The concentrations of NSF
and NSP were determined by measuring absorbance at 280 nm (A280) with a NanoDrop 2000C
spectrophotometer (Thermo Fisher Scientific, USA).

NSF solution was incubated with amphipol (Anatrace, USA) at mass ratio of 1:3, 1:6, 1:9, respectively, at 4ºC.
Following different incubation periods, the samples were mixed with native-PAGE loading buffer (Invitrogen,
USA) and subjected to electrophoresis at 4ºC under the following conditions: 90 V for 30 min, followed by 175
V for 90 min. The gels were stained with Coomassie Brilliant Blue. A linear gradient blue native polyacrylamide
gel electrophoresis (BN-PAGE) gel (3%-16%) was cast following Witting’s protocol77, with a Hoefer SG 30
gradient maker (Thermo Fisher Scientific, USA).

Metal shadowing
Metal shadowing was performed following established procedures29,78. For metal shadowing imaging of NSP
and NSF solutions extracted respectively from spider and silkworm, a 5 μL solution (30 μg·mL-1) was applied
to glow-discharged copper grids for 2 min. Excess solution was carefully removed using blotting paper. For
metal shadowing imaging of NSF in silkworm spinning dope and NSP in spider pinning dope, a glow-discharged
copper grid was placed on the spinning dope and gently pressed with tweezers to avoid any artifact formation.
NSF was dehydrated through a graded ethanol series (0, 25%, 50%, 75%, 100%) for 4 min each step. The samples
were then shadowed with tungsten using a DV-502B high vacuum evaporator (Denton Vacuum, USA) and air-
dried. A tungsten wire (8 cm in length) was clamped between two electrodes at a distance of 3.8 cm, with a
separation of 9.3 cm from the center of the specimen platform. The angle between tungsten wire and the sample
was approximately 10°. The NSF samples were rotated during evaporation for 14.5 min. Transmission electron
microscopy was performed using a Tecnai Spirit (FEI, USA).

Preparation of artificial silkworm silk, force-drawn silk, and cryo-vitrification of silk
samples
Artificial silkworm silk was prepared by the interface wiredrawing as previously described34 with modifications.
Briefly, 3 μL buffer (50 mmol·L-1 Citric acid, 50 mmol·L-1 Bis-tris propane, 16% w/v Polyethylene glycol 3,350,
pH 5.0) was placed on a glass plate, followed by the addition of an equal volume of NSF solution (3 mg·mL-1).
The solutions were quickly mixed with a 10 μL pipette tip, and the resulting artificial silks were immediately
placed on the glow-discharged mesh grids for subsequent analysis.

Fresh silkworm and spider silks were obtained by the forcible silking method. Third-instar silkworms and size-
matched spiders were gently immobilized using adhesive tape and placed under a stereomicroscope. Silk fibers
were then carefully drawn from the spinnerets using fine-tipped tweezers and immediately transferred onto glow-
discharged mesh grids. Third-instar larvae were selected due to their relatively small silk diameter (~3–5 μm),
which facilitates cryo-vitrification.

To ensure the physiological relevance of the fibers studied, the reeling speed was manually controlled to match
the natural spinning rates of silkworms (4-15 mm s-1) and spiders (10-20 mm s-1)79. To prevent dehydration, 3
μL of distilled water was added to the grid prior to silk application. Excess liquid was blotted with Whatman
filter paper for 6 s, and then the grids were rapidly plunge-frozen into liquid ethane using an EM GP system
(Leica Microsystems, Germany). Vitrified grids were stored in liquid nitrogen until further used.

Cryo focused ion beam milling and Cryo-ET data acquisition
Cryo-focused ion beam (cryo-FIB) milling was performed using a Helios NanoLab 600i Dual Beam SEM
(Thermo Fisher Scientific, USA), as previously described80. The goal was to prepare lamellae with a final
thickness of ≤ 100 nm. Frozen grids were transferred with the cryo-transfer shuttle into the SEM chamber via a
Quorum pp3000T cryo-transfer system (Quorum Technologies, UK) at -180℃. During milling, the angle
between the FIB and the silk surface was maintained at 5°- 10°, with milling performed from both sides to create
vitrified silk lamellae. The ion beam accelerating voltage was set at 30 kV, and the current ranged from 0.43 nA
(rough milling) to 10 pA (fine milling). The fine lamellae had a thickness of less than 100 nm.

Cryo-ET data acquisition was carried out using a Titan Krios transmission electron microscope fitted with a K2
detector. Overview tilt series of silks lamellae were acquired under a magnification of ×105,000, resulted in a
physical pixel size of 1.36 Å, under counting mode. Before data collection, the pre-tilt of the sample was
determined visually, and the pre-tilt was set to be 10° or -9° to match the pre-determined geometry caused by
loading grids. And the tilt range were set to be between -60° to +40° for -10° pre-tilt or -40 to +60° for +10° pre-
tilt, with a 3° step, resulting in 33 tilts and 99 electrons per tilt series. The slit width was set to be 20 eV, with the
refinement of zero-loss peak after collection of each tilt series, and nominal defocus was set to be -3.5 to -4.5 μm.
Fro the fibroin extracted from silk glands, the tilt range were set to be between -55° to +55° with a 3° step,
resulting in 35 tilts and 115 electrons per tilt series. All tilt series used in this study were collected using a dose-
symmetry strategy-based beam-image-shift facilitated acquisition scheme, by in-house developed scripts within
SerialEM software. The rest of the collection details are consistent with those mentioned above and are
summarized in Supplementary Table 1.

Tomogram reconstruction and sub-volume averaging of the fibroin
Motion correction and contrast transfer function (CTF) estimation were carried out in Warp; tilt series alignment
was carried out in IMOD 4.8.49 and AreTomo 2. For the fibroin extracted from silk glands, 77 tomograms were
binned 4 and apply the deconvolution method in Warp to denoise in order to facilitate the particle picking. To
pick filaments, CrYOLO 1.6.081 was employed to detect the straight filaments in the XY planes of the tomograms.
These segments were extracted with a box size of 40 pixels (218 Å) using Warp. The reference was used a
featureless cylinder using Dynamo 1.1.332. The sub-volumes were then aligned and classified using RELION
4.0. This averaged structure has an estimated resolution of 24 Å based on the ‘gold-standard’ FSC with 0.143
criterion. The details see supplementary Fig. 2.

For the fibroin within fibers, due to the low signal-to-noise ratio (SNR) in lamella, tomograms were reconstructed
using a SIRT-like filter and binned 2, as implemented in IMOD 4.8.49. Particles picking was performed using
CrYOLO 1.6.0 on the filtered tomograms. These segments were extracted with a box size of 90 pixels (245 Å)
using RELION 3.0. The reference was used a featureless cylinder using Dynamo 1.1.332. The sub-volumes were
then aligned and classified using RELION 3.0. This averaged structure has an estimated resolution of 22 Å based
on the ‘gold-standard’ FSC with 0.143 criterion. The details see supplementary Fig. 5a.

Restoring the tomograms by deep learning
Due to the dense arrangement of silk fibers and the presence of a significant missing wedge effect 3D especially
in XZ plane, we employed the REST method67 for tomogram restoration. First, we generated a ground truth by
randomly distributing the averaged map (many fibers) to densely arrange in the particle box. Then employing
random rotations and shifts (high-quantity datasets), noise and missing wedge artifacts were then added to the
ground truth to form the low-quantity datasets. These data were paired and then were used as training pairs for
REST training. The trained model was subsequently utilized to recover the raw tomograms from the weighted
back-projection (WBP) reconstruction. The details see supplementary Fig. 5b.

Measurement and Visualization
Visualization and figure generation were performed in UCSF ChimeraX v1.3 and IMOD 4.8.49. The diameter
and length of fibroin nanofibrils were measured manually using the measure tool in IMOD 4.8.49 and Fiji 2.32.
To analyze the density of spider and silk fibers, we projected the same number of XY slices into a single 2D
projection keep the same volume. The resulting projection was then binarized, and the proportion of the fibers
in space was quantified by calculating the ratio of white pixels (representing fibers) to the total number of pixels
in the projection.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_2": "In this manuscript, the authors utilized cryo-FIB milling and cryo-electron tomography to investigate and compare the structures of silk fibers from the silkworm, spider, and artificial silks. The results revealed previously unknown structural details, including the fundamental structural unit and arrangement, and provided new insight into the structural basis underlying the differences in the mechanical properties of the silkworm silk fibers and spider silk fibers. The manuscript focuses on an interesting topic that may attract readers from various scientific backgrounds. The data and analysis are solid. The manuscript The manuscript is well-organized and clearly written. I enjoyed reading it and think it is almost ready to be published as is in Nature Communication.\nI have only some minor suggestions for the authors to consider.\nThe manuscript studied and compared the structural details of silk fibers from silkworms, spiders, as well as artificial silks. The introduction provided background information about previous structural studies on silkworm silks and spider silks, but not artificial silks. It would be helpful to provide some brief background information about artificial silks, e.g., how they are made and their differences compared to natural silks if known. The structural difference between silkworm silks and spider silks is used to explain the better mechanical properties of spider silk fibers compared to that of the silkworm. While the authors also studied and revealed the structural difference between natural spider silk and artificial silk, can the structural difference be linked to the difference between natural spider silk and artificial silks?\nPage 6, Line 204: “(Fig. 4, a b, bottow)” should be “(Fig. 4, a b, bottom)”", "reviewer_3": "The paper titled “Cryo-ET Analysis Unveils the Minimum Structural Unit and Hierarchical Ultrastructure of Silk Fibers in Silkworm and Spider” describes the ultrastructure of silkworm silk, spider silk, and artificial silk analyzed using cryo-electron tomography. I find such type of work interesting and important for the field of silk biomaterials. It is evident that the authors have put significant effort into acquiring remarkable images using cryo-electron microscopy.\nBelow, I outline points that could be clarified to improve the paper:\n1. Title and Main Message: I believe the title does not accurately reflect the paper's main message and should be revised accordingly. The primary finding, as stated in the abstract, is the identification of differences in the arrangement of nanofibrils among spider silk fibers, silkworm silk fibers, and artificial silk fibers: “In spider silk, a dense structure featuring nanofibrils is clearly visible, with their long axes tightly aligned parallel to the flow direction, leaving no discernible gaps. In contrast, silkworm silk exhibits larger, regionally varying gaps between nanofibrils, while artificial silk lacks the highly ordered arrangement characteristic of natural silks.” (Page 1, lines 24-27). In previous work from the same group/corresponding author (https://doi.org/10.1016/j.scib.2023.12.050, also listed in this manuscript as reference 29), the smallest fibrillar unit of approximately 4 nm arranged in a herringbone pattern was reported. Thus, my understanding is that the finding of this work is not a structural unit or a “new pattern,” but rather the differences between the three types of silk. If this is correct, then the title should be changed accordingly, and the abstract should emphasize this point.\n2. Methodological Concerns: I am confused by the authors' use of various manipulative methods while simultaneously claiming to operate under purely native conditions where the protein remains unchanged. May be some details were not provided/mentioned? The authors should explain this point, specifically addressing the following:\no Why they applied shear to a shear-sensitive protein and concluded that the protein remained intact.\no Why they added an additional polymer without explaining how they differentiate between the protein and the polymer.\no Why they froze the entire silk gland and how they prevented ice nucleation in this process.\no How they utilized FIB milling and managed to avoid graphitization, considering that silk has low thermal conductivity and may burn even under cryo-FIB conditions.\nCo-reviewer:\nOverall, I find the paper interesting, and it is clear that significant work and effort have gone into acquiring these results. It presents important findings regarding the size and morphology of silk nanofibrils under different conditions. The use of FIB to create lamellae for TEM imaging is a crucial point, as it emphasizes their effort to maintain the native structure of silk fiber nanostructures.\nHowever, I am not fully convinced that the results are reliable. The bottom line is that they did not measure any material in its native state; they consistently altered it in some way. Even the silk fibers used were forced reeled from the animal. Additional comments can be found throughout the main text and in the methods section.\nI strongly disagree with many of the sample preparation choices made by the authors. Most of the techniques employed are likely to change and disrupt the native structure and state of the protein. Bulk freezing, dehydration with ethanol, and the addition of other polymers to the fibroin solution are known to alter fibroin structure and conformation. At the very least, these methods do not represent the native forms of the protein found inside the silk gland.\nFurthermore, from my personal experience, it is essential for the authors to provide proof that the silk fibers were not damaged during FIB milling. Even under gentle milling settings and cryogenic conditions, the protein can change structure and heat up significantly.\nSpecific Comments:\n• Page 3, line 108: Applying shear and flow to a shear-sensitive protein is inadvisable.\n• Page 4, line 115: If another polymer (amphipol) is added to the sample, how can the authors determine what they observe under the TEM microscope? Is it fibroin or the second polymer? How can they ensure that the second polymer does not alter the fibroin structure or macrostructure?\n• Page 4, line 115: 72 hours is a lengthy time to store NSF; the protein structure can be completely altered in that time.\n• Page 4, line 147: Are they discussing artificially spun fibers? Is it forced spinning from silkworms? This should be clarified. Why didn't they use natural silk fibers? What is a \"molted silkworm\"?\n• Page 4, line 148: Every FIB lamella preparation of silk needs to be validated with proof that it was not carbonized (e.g., through EDS or diffraction). Silkworm silk fibers have very low thermal conductivity, so during FIB milling, they accumulate energy and can reach high temperatures (>1000°C), leading to pyrolysis and carbonization.\n• Page 5, line 161: They barely discuss these bridges. What are their diameters? What do they believe they are composed of?\n• Page 5, lines 166-168: This statement is incorrect. Multiple studies have shown that the protein undergoes significant changes along the silk gland.\n• Page 6, lines 211-212: I do not understand how they interpret the \"perpendicular axis\" and how it correlates with the macro- fiber structure.\n• Page 9, line 311: This freezing process destroys the samples! Freezing such large samples results in ice crystal formation, which forces the sample into an unnatural morphology. Additionally, ice crystal formation exerts shear on the silk proteins, altering their structure and fibrillation.\n• Page 9, line 314: This purification process involves passing the samples through porous scaffolds and applying high shear to the proteins, significantly changing their native non-fibrillated state.\n• Page 9, line 328: Why did the authors add amphipol? Do they understand what impact it will have on fibroin?\n• Page 9, line 329: Again, 72 hours is a long time to store NSF and expect it to remain unchanged.\n• Page 10, line 334: How did they obtain spider silk proteins? This information is not mentioned.\n• Page 10, lines 338-339: The authors state that NSF has been dehydrated in ethanol… WHY? Dehydration with ethanol will completely alter the protein structure!\n• Page 10, line 348: Why did they add the buffer? Can't they spin it without buffer? It is NSF; they should be able to spin it easily.\n• Page 10, line 352: “Forced silking” will also change the structure!\n• Page 10, line 358: Everything done using FIB milling needs to be checked to ensure that the material has not been altered due to thermal effects caused by the beam. Even under gentle milling settings and cryogenic conditions, silkworm silk fibers can heat up and form graphene structures. To ensure this is not the case, they should use EDS to check the elemental composition of the lamellae and/or perform electron diffraction."}, "author_response": "Response to reviewers\nManuscript ID: NCOMMS-24-81232\nTitle: Cryo-ET Analysis Unveils the Minimum Structural Unit and Hierarchical\nUltrastructure of Silk Fibers in Silkworm and Spider\nJournal: Nature Communications\nReviewer #2 (Remarks to the Author):\nIn this manuscript, the authors utilized cryo-FIB milling and cryo-electron\ntomography to investigate and compare the structures of silk fibers from the silkworm,\nspider, and artificial silks. The results revealed previously unknown structural details,\nincluding the fundamental structural unit and arrangement, and provided new insight\ninto the structural basis underlying the differences in the mechanical properties of the\nsilkworm silk fibers and spider silk fibers. The manuscript focuses on an interesting\ntopic that may attract readers from various scientific backgrounds. The data and\nanalysis are solid. The manuscript is well-organized and clearly written. I enjoyed\nreading it and think it is almost ready to be published as is in Nature\nCommunications.\nResponse: We sincerely appreciate the reviewer’s recognition of our research.\nThe comments are valuable and very helpful for us to revise and improve the paper,\nand important guidance for our research. We have tried our best to improve the\nmanuscript and made corresponding revisions, and we hope to meet with approval.\nRegarding the comments and suggestions, we would like to respond point-by-point as\nfollows.\nI have only some minor suggestions for the authors to consider.\nThe manuscript studied and compared the structural details of silk fibers from\nsilkworms, spiders, as well as artificial silks. The introduction provided background\ninformation about previous structural studies on silkworm silks and spider silks, but\nnot artificial silks. It would be helpful to provide some brief background information\nabout artificial silks, e.g., how they are made and their differences compared to\nnatural silks if known.\nResponse: We thank the reviewer for this insightful suggestion.\nWe agree that providing background information on artificial silks would improve the\ncontextual framework of our study, especially since artificial silk is also one of the\nkey subjects of structural comparison in our work.\nTo address this, we have added a new paragraph to the Introduction, which outlines\nthe methods commonly used to fabricate artificial silk (e.g., wet spinning,\nelectrospinning, and microfluidic spinning), the typical materials employed (e.g.,\nregenerated silk fibroin and recombinant silk proteins), and the structural and\nmechanical differences between artificial and natural silks reported in the literature.\nThis addition helps clarify why artificial silk is included as a comparative target in our\nstudy and underscores the significance of elucidating its ultrastructural characteristics.\nRevisions in the Manuscript:\nA new paragraph has been added to the Introduction (lines 62–72 in the revised\nmanuscript) as follows:\n“In addition to natural silk fibers, artificial silks have attracted increasing interest\ndue to their potential for scalable production and customizable properties. Artificial\nsilk is typically produced by regenerated silk fibroin1,2 or recombinant silk proteins3-7\nusing techniques such as wet spinning1-4,6-8, dry spinning9,10, electrospinning5,\ninterface wiredrawing11 or microfluidic spinning12,13. These processes aim to fabricate\nartificial silk fibers with structures and properties comparable to those of natural silk,\nbut the properties of resulting artificial silk fibers are particularly influenced by\nfactors such as protein sequence, spinning dope concentration, flow rate, humidity,\npost-spin stretching, buffer composition, etc10,14,15. The ultrastructure within artificial\nsilk is even less understood. Understanding these structural discrepancies is crucial for\nimproving artificial silk production and bridging the performance gap between\nsynthetic and natural materials.”\nThe structural difference between silkworm silks and spider silks is used to explain\nthe better mechanical properties of spider silk fibers compared to that of the silkworm.\nWhile the authors also studied and revealed the structural difference between natural\nspider silk and artificial silk, can the structural difference be linked to the difference\nbetween natural spider silk and artificial silks?\nResponse: We appreciate the reviewer’s thoughtful and important comment.\nEstablishing a direct and quantitative link between the structural differences and\nmechanical properties of natural spider silk, silkworm silk, and artificial silks is\nindeed an important direction, and it remains a key focus of our ongoing research.\nIn the current study, our primary aim was to resolve long-standing controversies\nsurrounding the fundamental building blocks and hierarchical ultrastructure of natural\nsilkworm and spider silks. While we also analyzed the ultrastructure of artificial silks,\nwe did not measure their mechanical properties in this work, which limits our ability\nto directly correlate structural differences with performance differences between\nnatural and artificial silks.\nNevertheless, the distinct ultrastructural features we observed between artificial and\nnatural silks strongly suggest that spinning conditions, particularly the composition of\nthe spinning buffer, play a crucial role in determining the nanostructure of the\nresulting fibers (lines 208-210). Since mechanical properties of silks are intimately\ntied to their hierarchical structure, we believe these observations provide indirect but\nvaluable insights into how processing conditions may influence the functionality of\nartificial fibers.\nImportantly, beyond resolving structural controversies, this study also contributes a\nhigh-resolution methodological framework that can be applied in future investigations\naiming to systematically connect spinning conditions, structural organization, and\nmechanical performance of silk fibers. We have added discussion on this point in the\nrevised manuscript (lines 301–305) to clarify the implication of our findings in the\ncontext of artificial silk development\nRevisions in the Manuscript:\nA new sentence has been added to the Discussion section (lines 301–305 in the\nrevised manuscript) as follows:\n“Furthermore, a comparison of the ultrastructure between artificial and natural\nsilkworm silk reveals that, although both are derived from NSFs, differences in buffer\ncomposition during the spinning result in marked variations in fiber ultrastructure.\nThese structural differences, in turn, significantly influence the mechanical properties\nof the resulting silk fibers.”\nPage 6, Line 204: “(Fig. 4, a b, bottow)” should be “(Fig. 4, a b, bottom)”\nResponse: We are particularly grateful to the reviewer for pointing out the mistakes\nthat we had not noticed.\nWe have revised “Fig. 4, a b, bottow” as “Fig. 4, a b, bottom” in the revision.\nReviewer #3 (Remarks to the Author):\nThe paper titled “Cryo-ET Analysis Unveils the Minimum Structural Unit and\nHierarchical Ultrastructure of Silk Fibers in Silkworm and Spider” describes the\nultrastructure of silkworm silk, spider silk, and artificial silk analyzed using\ncryo-electron tomography. I find such type of work interesting and important for the\nfield of silk biomaterials. It is evident that the authors have put significant effort into\nacquiring remarkable images using cryo-electron microscopy. Below, I outline points\nthat could be clarified to improve the paper:\nResponse: We thank the reviewer for the positive and encouraging evaluation of our\nwork. We are grateful that the reviewer found our study both interesting and\nsignificant for the field of silk biomaterials, and we appreciate your recognition of the\neffort invested in acquiring the cryo-EM data.\nIn response to the reviewer's constructive comments, we have carefully addressed\neach of your specific concerns and suggestions in the detailed point-by-point replies.\nWhere appropriate, we have clarified experimental procedures, expanded explanations\nin the Methods and Discussion sections, and revised the manuscript to improve clarity\nand rigor.\nWe believe that these revisions have strengthened our manuscript and enhanced the\nclarity of our findings. We sincerely appreciate the reviewer's insightful comments,\nwhich have helped us improve both the technical details and the overall presentation\nof our work.\nWe hope that the revised version of our manuscript meets the reviewer's expectations\nand look forward to the further evaluation.\n1: Title and Main Message: I believe the title does not accurately reflect the paper's\nmain message and should be revised accordingly. The primary finding, as stated in the\nabstract, is the identification of differences in the arrangement of nanofibrils among\nspider silk fibers, silkworm silk fibers, and artificial silk fibers: “In spider silk, a\ndense structure featuring nanofibrils is clearly visible, with their long axes tightly\naligned parallel to the flow direction, leaving no discernible gaps. In contrast,\nsilkworm silk exhibits larger, regionally varying gaps between nanofibrils, while\nartificial silk lacks the highly ordered arrangement characteristic of natural silks.”\n(Page 1, lines 24-27). In previous work from the same group/corresponding author\n(https://doi.org/10.1016/j.scib.2023.12.050, also listed in this manuscript as reference\n29), the smallest fibrillar unit of approximately 4 nm arranged in a herringbone\npattern was reported. Thus, my understanding is that the finding of this work is not a\nstructural unit or a “new pattern,” but rather the differences between the three types of\nsilk. If this is correct, then the title should be changed accordingly, and the abstract\nshould emphasize this point.\nResponse: We sincerely thank the reviewer for this thoughtful and constructive\ncomment regarding the main message and title of our manuscript.\nOur study aims to showcase the fundamental building blocks and hierarchical\nultrastructure of silk fibers while also comparing the differences among these silks.\nWe agree that the comparative analysis of nanofibril arrangements among spider silk,\nsilkworm silk, and artificial silk fibers represents one of the central findings of this\nstudy. While this work reconfirms the nanofibril unit and herringbone pattern\npreviously observed16, the current study provides crucial advances in resolving and\ncomparing these architectures directly in situ across different silk types, under\nnear-native conditions using cryo-FIB and cryo-ET. Specifically, we directly\nvisualized the smallest structural units identified to date within the fiber and, for the\nfirst time, discovered their anisotropic arrangement in silkworm silk fibers.\nAlthough the basic structural unit and its arrangement identified in silkworm silk\nfibers are consistent with the morphology of silk proteins extracted from the silk\ngland and the higher-order structures observed at the terminal region of the anterior\nsilk gland in our previous work, these features had not previously been reported\nwithin silk fibers. Therefore, this current study provides strong evidence under more\nnative conditions to address the ongoing debate regarding the fundamental structural\nunit in silkworm and spider silk. At the same time, the comparative analysis of\npacking density, nanofibril arrangement, and hierarchical organization across the three\nsilk types constitutes another key contribution of this work, offering new structural\ninsights into the mechanical performance differences between them.\nIt is worth mentioning that this study was initially motivated by critiques from some\ncolleagues who questioned whether the nanofibrillar morphology we previously\nobserved in silk proteins extracted from the silk gland accurately represented their\nnative state, raising concerns that extraction conditions might have altered the protein\nstructure. Moreover, the widely cited micelle model (Jin et al., Nature, 2003) is\nprimarily based on scanning electron microscopy (SEM) and atomic force microscopy\n(AFM) observations, which reveal morphologies that differ substantially from the\nnanofibrillar morphology we reported for extracted silk proteins (Song et al., Science\nBulletin, 2024). In light of this, the primary objective of the present study was to\nelucidate the ultrastructure structure inside silkworm silk fibers—including the\nfundamental structural unit and their arrangement—under near-native conditions, and\nto compare these ultrastructures with those found in spider silk and artificial silk\nfibers. This work aims to provide important references for deciphering the natural\nspinning mechanism and developing biomimetic artificial fibers with superior\ncomprehensive performance.\nOn the other hand, due to the highly dense nature of silk fibers, even high-resolution\nSEM or AFM techniques struggle to reveal their internal structure clearly,\nhighlighting the need for more advanced imaging approaches. By employing the\ncombined cryo-FIB and cryo-ET technique, we made meaningful progress in\naddressing this challenge. However, it is regrettable that, due to the extremely dense\nstructure of spider silk, we were unable to investigate its internal hierarchical\norganization in greater detail. Thus, our comparative analysis among these silk fibers\nremains relatively preliminary, limited to basic metrics such as packing density. At\npresent, we are conducting ongoing ultrastructural analyses of seven different spider\nsilk fiber types, which, given the significant differences in their mechanical properties,\nmay provide further insights into the internal structural variations among spider silks.\nFinally, we sincerely thank the reviewer once again for the valuable suggestions,\nwhich have helped us clarify the main contributions of the current study and provided\nimportant guidance for our future research.\n2: Methodological Concerns: I am confused by the authors' use of various\nmanipulative methods while simultaneously claiming to operate under purely native\nconditions where the protein remains unchanged. May be some details were not\nprovided/mentioned? The authors should explain this point, specifically addressing\nthe following:\n⚫ Why they applied shear to a shear-sensitive protein and concluded that the protein\nremained intact.\nResponse: We thank the reviewer for raising this important concern and appreciate\nthe opportunity to clarify our experimental approach.\nWe agree that gel filtration chromatography, like all chromatographic processes,\ninvolve fluid flow and may introduce a certain degree of shear stress, which could, in\nprinciple, influence the morphology of shear-sensitive proteins such as silk fibroin.\nHowever, in our study, the sole purpose of performing gel filtration was to analyse the\nhomogeneity and composition of NSF. Therefore, even if minor shear-induced\nmorphological changes occurred during this purification step, they do not affect the\nvalidity of the subunit identification, which was the only purpose of that procedure.\nImportantly, we emphasize that gel filtration chromatography or any other\nshear-inducing purification steps was not used in the preparation of samples for\nvitrification, metal shadowing, or cryo-electron tomography (cryo-ET).\nWe acknowledge that the methodology description in the original manuscript may\nhave caused confusion. In the revised version, we have clarified the experimental\nworkflow and explicitly stated which samples underwent gel filtration and for what\npurpose, in order to avoid misunderstanding and improve the transparency and\nreadability of the manuscript (see materials and methods section, lines 346–349).\nWe sincerely appreciate the reviewer’s careful reading and insightful comment, which\nprompted this important clarification of our methods.\n⚫ Why they added an additional polymer without explaining how they differentiate\nbetween the protein and the polymer.\nResponse: We sincerely thank the reviewer for this insightful and important\ncomment.\nThe polymer referred to in this context is amphipol, a short-chain amphipathic\npolymer widely used in structural biology to replace detergents at the transmembrane\nsurface of membrane proteins, thereby stabilizing their structure in aqueous\nsolution17,18. Amphipol is chemically distinct from silk fibroin and is known not to\nform fibrillar or ordered structures on its own in solution.\nNonetheless, we fully agree with the reviewer that clearly distinguishing between\nprotein and polymer contributions in our data is essential for clarity. In response to\nthis concern, we have included additional metal shadowing electron microscopy\nresults in the revised manuscript (lines127-129, supplementary Fig. 1c-e) that\ncompare the morphology of samples with and without amphipol. These results show\nthat amphipol alone does not form any nanofibril structures under the experimental\nconditions used (Fig. 1a), while NSF retains its characteristic nanofibrillar\nmorphology even in the absence of amphipol (Fig. 1b-c). Thus, the nanofibril\nstructures observed in our experiments can be confidently attributed to NSF and not to\namphipol.\nFig. 1 Effect of amphipol on the morphology of NSF. (a) Metal shadowing image of amphipol\nalone (0.18 mg mL-1). (b) Metal shadowing image of NSF (0.03 mg mL-1) without amphipol. (c)\nMetal shadowing image of NSF mixed with amphipol at a mass ratio of 1:6 in the extraction\nbuffer (50 mmol L−1 NaCl, 50 mmol L−1 Bis-Tris, 0.5 mmol L−1 EDTA, pH 7.5), with a final NSF\nconcentration of 0.03 mg mL−1. Scale bar: 200 nm.\nThe addition of amphipol in selected assays was motivated by its ability to stabilize\nNSF under in vitro conditions, particularly during sample handling and preparation.\nNSF is known to be highly unstable once extracted, and tends to aggregate or\nprecipitate rapidly—even at low temperatures and in the absence of freezing19-22. In\nour previous work16, we performed extensive screening and found that both amphipol\nand digitonin significantly enhance the solubility and stability of NSF (Fig. 2a-b).\nTurbidity assays and circular dichroism (CD) analysis further confirmed that NSF\nremained stable for at least 336 hours in the presence of digitonin (Fig. 2c-e).\nMoreover, CD analysis showed that digitonin did not appreciably alter the random\ncoil structure of freshly extracted NSF (Fig. 2f). Although amphipol similarly\nstabilizes NSF in solution, its high absorbance interferes with CD spectroscopy.\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 2. Digitonin and Amphipol stabilized NSF in vitro. (a) A280 of NSF over time in the\npresence of MSP1D1 or BSA. (b) A280 of NSF over time in the presence of detergents. (c)\nNSF turbidity analysis. The inset (left) showed that NSF was stable in the presence of digitonin\nafter 336 h. The inset (right) showed the aggregation of NSF in the absence of digitonin after\n336 h. (d) CD spectra of NSF in the presence of digitonin over time. (e) CD spectra of NSF in the\nabsence of digitonin over time. (f) CD spectra of NSF in the presence or absence of digitonin.\nTaken together, these findings validate the use of amphipol and digitonin as essential\nstabilizing agents that do not introduce structural artifacts or interfere with the\ninterpretation of NSF morphology. All structural analyses and tomographic\nreconstructions were performed on samples in which amphipol was either excluded or\nits influence empirically shown to be negligible.\nWe once again thank the reviewer for this valuable comment, which helped us\nsignificantly improve the clarity and rigor of our methodological description in the\nrevised manuscript.\n⚫ Why they froze the entire silk gland and how they prevented ice nucleation in this\nprocess.\nResponse: We sincerely thank the reviewer for this thoughtful and important\nquestion.\nThe decision to freeze the entire silkworm was primarily based on practical\nconsiderations. Rearing silkworms is time-consuming, as each lifecycle spans\napproximately 60 days, and the availability of fresh mulberry leaves becomes highly\nlimited during the winter season. To ensure experimental continuity and access to NSF,\nwe opted to freeze whole silkworms and subsequently purify NSF from their silk\nglands as needed, thereby eliminating the need for continuous rearing.\nWe acknowledge the general concern that freezing biological specimens may induce\nirreversible structural changes, such as precipitation or denaturation. Indeed, previous\nstudies and our own observations indicate have shown that extracted NSF undergoes\nirreversible precipitation when frozen in liquid nitrogen23,24, likely due to\nshear-induced aggregation during the freeze-thaw cycle25,26. It is well known that\nfreezing and thawing induce regenerated silk fibroin (RSF) or NSF to form porous\nstructures in vitro27-29. However, it is important to emphasize that the solution\nmicroenvironment of RSF or NSF in vitro differs significantly from its native\nphysiological environment within the silk gland lumen. The physiological conditions\ninside the gland—including specific ion concentrations, pH, and potential stabilizing\nbiomolecules—appear to play a protective role in preserving the integrity of NSF\neven under freezing conditions. Moreover, the intact silkworm body may further\nbuffer the silk gland against rapid temperature fluctuations and mechanical stress,\nthereby reducing the likelihood of ice crystal-induced damage.\nIn support of this, our previous work16 demonstrated that soluble NSF could be\nsuccessfully extracted from silk glands of frozen silkworms (Fig. 1a-b), suggesting\nthat the physiological environment within the lumen of silk gland—including ions, pH,\nand possibly unknown protective factors—may be essential roles in preventing NSF\ndenaturation or coagulation. Furthermore, SDS-PAGE analysis revealed that NSFs\npurified from the posterior silk gland (PSG), the posterior region of middle silk gland\n(PMSG), the middle region of middle silk gland (MMSG) and the anterior region of\nmiddle silk gland (AMSG) of frozen silkworms exhibited identical electrophoretic\nmobility and molecular composition as NSF extracted from the anterior silk gland\n(ASG) of living silkworm (Fig. 1b-c).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024) under CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 1. Purification of NSF from silk gland. (a) Silk gland of the silkworm. The silk gland is\ndivided into ASG, MSG and PSG. MSG is further divided into AMSG, MMSG and PMSG. (b)\nNSF was purified from PSG, AMSG, MMSG and PMSG of frozen silkworm, respectively, and\nthen separated by a linear 4%-16% gradient SDS-PAGE. (c) Top. A linear 4%-16% gradient\nSDS-PAGE analysis of NSF purified by ions exchange chromatography. M, marker; CR, the crude\nextraction of ASG of living silkworm; F1, the flow-through fraction after S cation exchange media;\nS, cation exchange media; F2, the flow-through fraction after Q anion exchange media; Q, anion\nexchange media. Bottom, NSF purified by size exclusion chromatography. The inset showed the\npurity of NSF valued by 4%-16% SDS-PAGE. Fractions 1 and 2 were pooled for CD spectra and\nmetal shadowing, respectively.\nMetal shadowing confirmed the formation of nanofibrils in all these samples, which\nclosely resembled those derived from living silkworms (Fig. 2a). In addition, circular\ndichroism (CD) spectra revealed NSFs from PSG, PMSG, MMSG and AMSG of\nfrozen silkworms exhibited secondary structures comparable to those of NSF from\nASG of living silkworms (Fig. 2b).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024) under CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 2. The morphology and structure of NSF. (a) Metal shadowing of NSF from PSG, PMSG,\nMMSG, AMSG of frozen silkworm and ASG of living silkworm. Scale bar, 200 nm. (b) CD\nspectra of NSF from PSG, PMSG, MMSG, AMSG of frozen silkworm and ASG of living\nsilkworm.\nTaken together, these results provide strong evidence that NSF extracted from frozen\nsilkworms retains its native morphology, molecular composition, and structural\nfeatures, thereby validating the use of frozen specimens in our study without\ncompromising the reliability of our findings. Once again, we sincerely appreciate the\nreviewer’s insightful comments.\n⚫ How they utilized FIB milling and managed to avoid graphitization, considering\nthat silk has low thermal conductivity and may burn even under cryo-FIB conditions.\nResponse: We thank the reviewer for this very important and insightful question.\nWe fully acknowledge the potential concern that focused ion beam (FIB) milling,\neven under cryogenic conditions, could induce structural alterations in sensitive\nbiological materials, particularly protein-based fibers. This issue was carefully\nconsidered during our experimental design, especially regarding the reliability of\nultrastructural features observed in lamellae. However, although we lacked access to\nadvanced analytical tools (such as EDS) and knowledge reserves to directly evaluate\npotential damage (e.g., carbonization) in lamellae, we performed structural\ncross-validation using an independent experimental approach. Specifically, we\nconducted single-particle cryo-electron tomography (cryo-ET) and subtomogram\naveraging on NSF extracted from silk gland that had not undergone FIB processing.\nRemarkably, the basic structural units observed in the FIB-milled lamellae closely\nmatched those seen in the cryo-ET reconstructions from extracted NSF samples,\nproviding strong biochemical and morphological evidence that the ultrastructure\nwithin the lamellae was well preserved and not artificially induced during FIB\nmilling.\nFurthermore, as shown in Fig. 2a of the manuscript, the lamella included both\nfibroin-rich and sericin-rich regions. Notably, the sericin-rich region (left side) lacked\nnanofibrillar organization, while the fibroin-rich region (right side) displayed dense\nnanofibrillar networks. Since sericin is a globular protein and is not naturally fibrillar,\nthe absence of nanofibrils in this region argues strongly against the possibility that\nnanofibrils in the fibroin region are artifacts introduced by the FIB processing. This\ninternal spatial distinction within the same lamella supports the conclusion that the\nnanofibrillar organization observed in the fibroin region is intrinsic and not\nbeam-induced.\nAdditionally, recent advances in cryo-FIB milling technology have demonstrated that,\nwhen carefully implemented, this technique can preserve native biological structures\nwith high fidelity and has been widely used in in situ cryo-ET for cellular and\nmacromolecular imaging30-40.\nIn our study, we employed a state-of-the-art Thermo Fisher Helios NanoLab 600i\nDualBeam cryo-FIB/SEM system specifically optimized for cryo-electron microscopy\nsample preparation. Several key precautions were taken to minimize beam-induced\nheating and structural damage:\nUse of fresh, hydrated silk fibers:\nThe silk fibers used for cryo-FIB milling were freshly drawn from the silkworm\nspinneret, and retained intrinsic moisture. The presence of water enhances local\nelectrical conductivity and thermal dissipation, significantly reducing sample charging\nand heat accumulation during ion beam exposure. This moisture content helped\nmitigate thermal damage and the risk of graphitization, in contrast to dried cocoon\nsilk.\nCryogenic conditions throughout the FIB process:\nAll FIB milling procedures were conducted under deep cryogenic conditions (< –\n180 °C) using the Helios NanoLab system. At such low temperatures, biological\nsamples are vitrified, effectively preserving their native structures while dramatically\nreducing thermal diffusion and decomposition. This approach effectively eliminates\nthe risk of burning or graphitization, even in materials with low thermal conductivity\nlike silk.\nTriple-layer protective platinum coating:\nPrior to milling, we deposited three successive layers of protective platinum (Pt) on\nthe fiber surface. These coatings:\n· Enhanced surface conductivity,\n· Provided shielding from direct ion beam exposure,\n· Distributed both charge and heat more evenly,\n· Prevented sputtering and beam-induced structural degradation.\nLow current milling during final thinning steps:\nFor the final thinning of the lamellae, we employed extremely low ion beam currents\n(10 pA) to minimize any residual heat generation. This is a widely accepted approach\nin cryo-FIB protocols to prevent damage in sensitive specimens.\nCollectively, these precautions ensured that sample preparation was conducted with\nminimal physical or thermal disturbance. Most importantly, the consistent\nmorphological features observed between cryo-ET reconstructions from FIB-milled\nlamellae and independently prepared NSF samples strongly supports the conclusion\nthat cryo-FIB milling did not introduce significant structural artifacts or thermal\ndamage in our study.\nOnce again, we sincerely thank the reviewer for raising this critical point, which has\nhelped us improve the rigor and clarity of our methodological description.\nCo-reviewer\nOverall, I find the paper interesting, and it is clear that significant work and effort\nhave gone into acquiring these results. It presents important findings regarding the\nsize and morphology of silk nanofibrils under different conditions. The use of FIB to\ncreate lamellae for TEM imaging is a crucial point, as it emphasizes their effort to\nmaintain the native structure of silk fiber nanostructures.\nResponse: We sincerely thank the reviewer for the positive and encouraging\ncomments.\nWe are grateful that the reviewer found our study interesting and recognized the\nsignificance of our findings regarding the size and morphology of silk nanofibrils. We\nalso greatly appreciate the acknowledgment of the substantial experimental effort\ninvolved. These comments are highly motivating and have been instrumental in\nhelping us further refine and improve the manuscript.\nHowever, I am not fully convinced that the results are reliable. The bottom line is that\nthey did not measure any material in its native state; they consistently altered it in\nsome way. Even the silk fibers used were forced reeled from the animal. Additional\ncomments can be found throughout the main text and in the methods section.\nResponse: We sincerely thank the reviewer for this insightful and important\ncomment.\nThe concern raised by reviewer touches on a fundamental challenge that has long\nexisted in silk research, namely the inherent difficulty in probing ultrastructure in\ntruly native states without any perturbation. We fully agree that despite our best\nefforts to preserve native-like conditions, certain experimental manipulations, such as\nfreezing, addition of amphipol, and fiber reeling, are actually indispensable and have\nan impact on the morphology of NSF. However, we have taken extensive measures to\ndemonstrate the reliability of our results. Specifically, we addressed three key steps\nthat may be viewed as perturbative:\nAddition of amphipol to preserve NSF in solution:\nThe polymer referred to in this context is amphipol, a short-chain amphipathic\npolymer widely used in structural biology to replace detergents at the transmembrane\nsurface of membrane proteins, thereby stabilizing their structure in aqueous\nsolution17,18. Amphipol is chemically distinct from silk fibroin and is known not to\nform fibrillar or ordered structures on its own in solution.\nNonetheless, we fully agree with the reviewer that clearly distinguishing between\nprotein and polymer contributions in our data is essential for clarity. In response to\nthis concern, we have included additional metal shadowing electron microscopy\nresults in the revised manuscript (lines127-129, supplementary Fig. 1c-e) that\ncompare the morphology of samples with and without amphipol. These results show\nthat amphipol alone does not form any nanofibril structures under the experimental\nconditions used (Fig. 1a), while NSF retains its characteristic nanofibrillar\nmorphology even in the absence of amphipol (Fig. 1b-c). Thus, the nanofibril\nstructures observed in our experiments can be confidently attributed to NSF and not to\namphipol.\nFig. 1 Effect of amphipol on the morphology of NSF. (a) Metal shadowing image of amphipol\nalone (0.18 mg mL-1). (b) Metal shadowing image of NSF (0.03 mg mL-1) without amphipol. (c)\nMetal shadowing image of NSF mixed with amphipol at a mass ratio of 1:6 in the extraction\nbuffer (50 mmol L−1 NaCl, 50 mmol L−1 Bis-Tris, 0.5 mmol L−1 EDTA, pH 7.5), with a final NSF\nconcentration of 0.03 mg mL−1. Scale bar: 200 nm.\nThe addition of amphipol in selected assays was motivated by its ability to stabilize\nNSF under in vitro conditions, particularly during sample handling and preparation.\nNSF is known to be highly unstable once extracted, and tends to aggregate or\nprecipitate rapidly—even at low temperatures and in the absence of freezing19-22. In\nour previous work16, we performed extensive screening and found that both amphipol\nand digitonin significantly enhance the solubility and stability of NSF (Fig. 2a-b).\nTurbidity assays and circular dichroism (CD) analysis further confirmed that NSF\nremained stable for at least 336 hours in the presence of digitonin (Fig. 2c-e).\nMoreover, CD analysis showed that digitonin did not appreciably alter the random\ncoil structure of freshly extracted NSF (Fig. 2f). Although amphipol similarly\nstabilizes NSF in solution, its high absorbance interferes with CD spectroscopy.\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 2. Digitonin and Amphipol stabilized NSF in vitro. (a) A280 of NSF over time in the\npresence of MSP1D1 or BSA. (b) A280 of NSF over time in the presence of detergents. (c)\nNSF turbidity analysis. The inset (left) showed that NSF was stable in the presence of digitonin\nafter 336 h. The inset (right) showed the aggregation of NSF in the absence of digitonin after\n336 h. (d) CD spectra of NSF in the presence of digitonin over time. (e) CD spectra of NSF in the\nabsence of digitonin over time. (f) CD spectra of NSF in the presence or absence of digitonin.\nTaken together, these findings validate the use of amphipol and digitonin as essential\nstabilizing agents that do not introduce structural artifacts or interfere with the\ninterpretation of NSF morphology. All structural analyses and tomographic\nreconstructions were performed on samples in which amphipol was either excluded or\nits influence empirically shown to be negligible.\nFreezing of whole silkworms for sample preservation:\nThe decision to freeze the entire silkworm was primarily based on practical\nconsiderations. Rearing silkworms is time-consuming, as each lifecycle spans\napproximately 60 days, and the availability of fresh mulberry leaves becomes highly\nlimited during the winter season. To ensure experimental continuity and access to NSF,\nwe opted to freeze whole silkworms and subsequently purify NSF from their silk\nglands as needed, thereby eliminating the need for continuous rearing.\nWe acknowledge the general concern that freezing biological specimens may induce\nirreversible structural changes, such as precipitation or denaturation. Indeed, previous\nstudies and our own observations indicate have shown that extracted NSF undergoes\nirreversible precipitation when frozen in liquid nitrogen23,24, likely due to\nshear-induced aggregation during the freeze-thaw cycle25,26. It is well known that\nfreezing and thawing induce regenerated silk fibroin (RSF) or NSF to form porous\nstructures in vitro27-29. However, it is important to emphasize that the solution\nmicroenvironment of RSF or NSF in vitro differs significantly from its native\nphysiological environment within the silk gland lumen. The physiological conditions\ninside the gland—including specific ion concentrations, pH, and potential stabilizing\nbiomolecules—appear to play a protective role in preserving the integrity of NSF\neven under freezing conditions. Moreover, the intact silkworm body may further\nbuffer the silk gland against rapid temperature fluctuations and mechanical stress,\nthereby reducing the likelihood of ice crystal-induced damage.\nIn support of this, our previous work16 demonstrated that soluble NSF could be\nsuccessfully extracted from silk glands of frozen silkworms (Fig. 3a-b), suggesting\nthat the physiological environment within the lumen of silk gland—including ions, pH,\nand possibly unknown protective factors—may be essential roles in preventing NSF\ndenaturation or coagulation. Furthermore, SDS-PAGE analysis revealed that NSFs\npurified from the posterior silk gland (PSG), the posterior region of middle silk gland\n(PMSG), the middle region of middle silk gland (MMSG) and the anterior region of\nmiddle silk gland (AMSG) of frozen silkworms exhibited identical electrophoretic\nmobility and molecular composition as NSF extracted from the anterior silk gland\n(ASG) of living silkworm (Fig. 3b-c).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 3. Purification of NSF from silk gland. (a) Silk gland of the silkworm. The silk gland is\ndivided into ASG, MSG and PSG. MSG is further divided into AMSG, MMSG and PMSG. (b)\nNSF was purified from PSG, AMSG, MMSG and PMSG of frozen silkworm, respectively, and\nthen separated by a linear 4%-16% gradient SDS-PAGE. (c) Top. A linear 4%-16% gradient\nSDS-PAGE analysis of NSF purified by ions exchange chromatography. M, marker; CR, the crude\nextraction of ASG of living silkworm; F1, the flow-through fraction after S cation exchange media;\nS, cation exchange media; F2, the flow-through fraction after Q anion exchange media; Q, anion\nexchange media. Bottom, NSF purified by size exclusion chromatography. The inset showed the\npurity of NSF valued by 4%-16% SDS-PAGE. Fractions 1 and 2 were pooled for CD spectra and\nmetal shadowing, respectively.\nMetal shadowing confirmed the formation of nanofibrils in all these samples, which\nclosely resembled those derived from living silkworms (Fig. 4a). In addition, circular\ndichroism (CD) spectra revealed NSFs from PSG, PMSG, MMSG and AMSG of\nfrozen silkworms exhibited secondary structures comparable to those of NSF from\nASG of living silkworms (Fig. 4b).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024) under CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 4. The morphology and structure of NSF. (a) Metal shadowing of NSF from PSG, PMSG,\nMMSG, AMSG of frozen silkworm and ASG of living silkworm. Scale bar, 200 nm. (b) CD\nspectra of NSF from PSG, PMSG, MMSG, AMSG of frozen silkworm and ASG of living\nsilkworm.\nTaken together, these results provide strong evidence that NSF extracted from frozen\nsilkworms retains its native morphology, molecular composition, and structural\nfeatures, thereby validating the use of frozen specimens in our study without\ncompromising the reliability of our findings. Once again, we sincerely appreciate the\nreviewer’s insightful comments.\nForce-drawn fibers from silkworm and spider:\nIt is important to note that, in nature, both silkworm cocoon formation and spider web\nweaving involve the natural pulling or drawing of silk proteins from the gland. The\nnatural spinning speeds are approximately 4-15 mm s-1 for silkworms and 10-20 mm\ns-1 for spiders41. In our study, we carefully calibrated the forced reeling speed to fall\nwithin these physiological ranges as closely as possible, thereby minimizing\nmechanical stress and preserving native structural features.\nTo further confirm that forced reeling does not introduce structural artifacts, we also\nperformed cryo-FIB milling and cryo-electron tomography on silk fibers extracted\ndirectly from silkworm cocoons. The lamellar ultrastructure of cocoon silk was\nconsistent with that observed in freshly drawn silk (Fig. 5), supporting the reliability\nof our approach. However, due to pronounced image drift during tilt-series acquisition\nof lamellae prepared from dried cocoon silk, we ultimately excluded these samples\nfrom tomographic data collection. We speculate that this instability arose from the\ncomplete dehydration of cocoon silk, which compromises its electrical conductivity.\nThe resulting charge accumulation during imaging likely caused the observed drift.\nIn contrast, freshly drawn silkworm silk and spider silk retain residual moisture,\nwhich enhances conductivity and stabilizes imaging during cryo-ET. Thus, our choice\nto use freshly reeled silks for tomographic analysis was driven primarily by their\nsuperior performance under cryo-imaging conditions.\nFig. 5. Comparison of the ultrastructure of silkworm cocoon silk and freshly drawn silk. (a)\nLow-magnification projection image of the freshly drawn silk from third-instar silkworm. (b)\nLow-magnification projection image of silkworm cocoon silk.\nTaken together, we have systematically addressed each potential source of artifact and\nprovided extensive experimental evidence demonstrating that the ultrastructure,\nmolecular composition, and secondary structure of silk proteins are preserved\nthroughout each key step of our workflow. While we acknowledge that no in vitro\nmethod is entirely free of manipulation, we believe that our carefully controlled\nworkflow and multiple levels of internal validation ensure that the structures reported\nhere are reliable and faithfully reflect native silk protein assemblies.\nI strongly disagree with many of the sample preparation choices made by the authors.\nMost of the techniques employed are likely to change and disrupt the native structure\nand state of the protein. Bulk freezing, dehydration with ethanol, and the addition of\nother polymers to the fibroin solution are known to alter fibroin structure and\nconformation. At the very least, these methods do not represent the native forms of the\nprotein found inside the silk gland.\nResponse: We sincerely appreciate the reviewer’s critical insight regarding the\npotential structural perturbations caused by our sample preparation protocols.\nWe understand the concern that procedures such as bulk freezing, ethanol dehydration,\nand the addition of amphiphilic polymers may affect the native structure of silk\nfibroin. Below, we address each point in detail, with supporting evidence from our\nown data and prior work.\nFreezing of whole silkworms for sample preservation:\nThe decision to freeze the entire silkworm was primarily based on practical\nconsiderations. Rearing silkworms is time-consuming, as each lifecycle spans\napproximately 60 days, and the availability of fresh mulberry leaves becomes highly\nlimited during the winter season. To ensure experimental continuity and access to NSF,\nwe opted to freeze whole silkworms and subsequently purify NSF from their silk\nglands as needed, thereby eliminating the need for continuous rearing.\nWe acknowledge the general concern that freezing biological specimens may induce\nirreversible structural changes, such as precipitation or denaturation. Indeed, previous\nstudies and our own observations indicate have shown that extracted NSF undergoes\nirreversible precipitation when frozen in liquid nitrogen23,24, likely due to\nshear-induced aggregation during the freeze-thaw cycle25,26. It is well known that\nfreezing and thawing induce regenerated silk fibroin (RSF) or NSF to form porous\nstructures in vitro27-29. However, it is important to emphasize that the solution\nmicroenvironment of RSF or NSF in vitro differs significantly from its native\nphysiological environment within the silk gland lumen. The physiological conditions\ninside the gland—including specific ion concentrations, pH, and potential stabilizing\nbiomolecules—appear to play a protective role in preserving the integrity of NSF\neven under freezing conditions. Moreover, the intact silkworm body may further\nbuffer the silk gland against rapid temperature fluctuations and mechanical stress,\nthereby reducing the likelihood of ice crystal-induced damage.\nIn support of this, our previous work16 demonstrated that soluble NSF could be\nsuccessfully extracted from silk glands of frozen silkworms (Fig. 1a-b), suggesting\nthat the physiological environment within the lumen of silk gland—including ions, pH,\nand possibly unknown protective factors—may be essential roles in preventing NSF\ndenaturation or coagulation. Furthermore, SDS-PAGE analysis revealed that NSFs\npurified from the posterior silk gland (PSG), the posterior region of middle silk gland\n(PMSG), the middle region of middle silk gland (MMSG) and the anterior region of\nmiddle silk gland (AMSG) of frozen silkworms exhibited identical electrophoretic\nmobility and molecular composition as NSF extracted from the anterior silk gland\n(ASG) of living silkworm (Fig. 1b-c).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 1. Purification of NSF from silk gland. (a) Silk gland of the silkworm. The silk gland is\ndivided into ASG, MSG and PSG. MSG is further divided into AMSG, MMSG and PMSG. (b)\nNSF was purified from PSG, AMSG, MMSG and PMSG of frozen silkworm, respectively, and\nthen separated by a linear 4%-16% gradient SDS-PAGE. (c) Top. A linear 4%-16% gradient\nSDS-PAGE analysis of NSF purified by ions exchange chromatography. M, marker; CR, the crude\nextraction of ASG of living silkworm; F1, the flow-through fraction after S cation exchange media;\nS, cation exchange media; F2, the flow-through fraction after Q anion exchange media; Q, anion\nexchange media. Bottom, NSF purified by size exclusion chromatography. The inset showed the\npurity of NSF valued by 4%-16% SDS-PAGE. Fractions 1 and 2 were pooled for CD spectra and\nmetal shadowing, respectively.\nMetal shadowing confirmed the formation of nanofibrils in all these samples, which\nclosely resembled those derived from living silkworms (Fig. 2a). In addition, circular\ndichroism (CD) spectra revealed NSFs from PSG, PMSG, MMSG and AMSG of\nfrozen silkworms exhibited secondary structures comparable to those of NSF from\nASG of living silkworms (Fig. 2b).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024) under CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 2. The morphology and structure of NSF. (a) Metal shadowing of NSF from PSG, PMSG,\nMMSG, AMSG of frozen silkworm and ASG of living silkworm. Scale bar, 200 nm. (b) CD\nspectra of NSF from PSG, PMSG, MMSG, AMSG of frozen silkworm and ASG of living\nsilkworm.\nTaken together, these results provide strong evidence that NSF extracted from frozen\nsilkworms retains its native morphology, molecular composition, and structural\nfeatures, thereby validating the use of frozen specimens in our study without\ncompromising the reliability of our findings. Once again, we sincerely appreciate the\nreviewer’s insightful comments.\nEthanol dehydration in metal shadowing:\nEthanol, as an organic solvent, can in some cases alter protein structure when added\ndirectly to protein solutions. However, in this study, the ethanol dehydration step used\nin the metal shadowing experiment was performed after the protein samples had been\nadsorbed onto the carbon film of the EM grids. At this stage, the samples were already\npartially immobilized, and ethanol dehydration served primarily to enhance adhesion\nbetween the protein and the carbon support, thereby minimizing sample loss during\nsubsequent grid transfer and drying steps. In addition, ethanol treatment helps reduce\nsurface tension on the grid, facilitating smoother handling during processing.\nTo directly evaluate whether ethanol dehydration affects the ultrastructure of NSF, we\nconducted control experiments in which metal shadowing was performed both with\nand without the ethanol dehydration step. As shown in Fig. 3, NSF exhibited similar\nfibrillar morphology under both conditions, indicating that ethanol dehydration does\nnot introduce discernible artifacts in terms of ultrastructural preservation.\nFig. 3. The effect of ethanol dehydration on the morphology of NSF. (a) Metal shadowing of NSF\nwith ethanol dehydration. (b) Metal shadowing of NSF without ethanol dehydration. Scale bar: 200 nm.\nAddition of amphipol to preserve NSF in solution:\nThe polymer referred to in this context is amphipol, a short-chain amphipathic\npolymer widely used in structural biology to replace detergents at the transmembrane\nsurface of membrane proteins, thereby stabilizing their structure in aqueous\nsolution17,18. Amphipol is chemically distinct from silk fibroin and is known not to\nform fibrillar or ordered structures on its own in solution.\nNonetheless, we fully agree with the reviewer that clearly distinguishing between\nprotein and polymer contributions in our data is essential for clarity. In response to\nthis concern, we have included additional metal shadowing electron microscopy\nresults in the revised manuscript (lines127-129, supplementary Fig. 1c-e) that\ncompare the morphology of samples with and without amphipol. These results show\nthat amphipol alone does not form any nanofibril structures under the experimental\nconditions used (Fig. 4a), while NSF retains its characteristic nanofibrillar\nmorphology even in the absence of amphipol (Fig. 4b-c). Thus, the nanofibril\nstructures observed in our experiments can be confidently attributed to NSF and not to\namphipol.\nFig. 4 Effect of amphipol on the morphology of NSF. (a) Metal shadowing image of amphipol\nalone (0.18 mg mL-1). (b) Metal shadowing image of NSF (0.03 mg mL-1) without amphipol. (c)\nMetal shadowing image of NSF mixed with amphipol at a mass ratio of 1:6 in the extraction\nbuffer (50 mmol L−1 NaCl, 50 mmol L−1 Bis-Tris, 0.5 mmol L−1 EDTA, pH 7.5), with a final NSF\nconcentration of 0.03 mg mL−1. Scale bar: 200 nm.\nThe addition of amphipol in selected assays was motivated by its ability to stabilize\nNSF under in vitro conditions, particularly during sample handling and preparation.\nNSF is known to be highly unstable once extracted, and tends to aggregate or\nprecipitate rapidly—even at low temperatures and in the absence of freezing19-22. In\nour previous work16, we performed extensive screening and found that both amphipol\nand digitonin significantly enhance the solubility and stability of NSF (Fig. 5a-b).\nTurbidity assays and circular dichroism (CD) analysis further confirmed that NSF\nremained stable for at least 336 hours in the presence of digitonin (Fig. 5c-e).\nMoreover, CD analysis showed that digitonin did not appreciably alter the random\ncoil structure of freshly extracted NSF (Fig. 5f). Although amphipol similarly\nstabilizes NSF in solution, its high absorbance interferes with CD spectroscopy.\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 5. Digitonin and Amphipol stabilized NSF in vitro. (a) A280 of NSF over time in the\npresence of MSP1D1 or BSA. (b) A280 of NSF over time in the presence of detergents. (c)\nNSF turbidity analysis. The inset (left) showed that NSF was stable in the presence of digitonin\nafter 336 h. The inset (right) showed the aggregation of NSF in the absence of digitonin after\n336 h. (d) CD spectra of NSF in the presence of digitonin over time. (e) CD spectra of NSF in the\nabsence of digitonin over time. (f) CD spectra of NSF in the presence or absence of digitonin.\nTaken together, these findings validate the use of amphipol and digitonin as essential\nstabilizing agents that do not introduce structural artifacts or interfere with the\ninterpretation of NSF morphology. All structural analyses and tomographic\nreconstructions were performed on samples in which amphipol was either excluded or\nits influence empirically shown to be negligible.\nIn summary, we have carefully validated each sample preparation step to ensure that\nthe structural and morphological features of NSF are faithfully preserved. Wherever\nfeasible, we have provided direct experimental evidence to rule out potential artifacts\nintroduced by freezing, ethanol dehydration, or amphipol addition. We thank the\nreviewer once again for raising these important considerations, which prompted us to\nclarify and strengthen our methodological descriptions in the revised manuscript.\nFurthermore, from my personal experience, it is essential for the authors to provide\nproof that the silk fibers were not damaged during FIB milling. Even under gentle\nmilling settings and cryogenic conditions, the protein can change structure and heat\nup significantly.\nResponse: We thank the reviewer for this very important and insightful question.\nWe fully acknowledge the potential concern that focused ion beam (FIB) milling,\neven under cryogenic conditions, could induce structural alterations in sensitive\nbiological materials, particularly protein-based fibers. This issue was carefully\nconsidered during our experimental design, especially regarding the reliability of\nultrastructural features observed in lamellae. However, although we lacked access to\nadvanced analytical tools (such as EDS) and knowledge reserves to directly evaluate\npotential damage (e.g., carbonization) in lamellae, we performed structural\ncross-validation using an independent experimental approach. Specifically, we\nconducted single-particle cryo-electron tomography (cryo-ET) and subtomogram\naveraging on NSF extracted from silk gland that had not undergone FIB processing.\nRemarkably, the basic structural units observed in the FIB-milled lamellae closely\nmatched those seen in the cryo-ET reconstructions from extracted NSF samples,\nproviding strong biochemical and morphological evidence that the ultrastructure\nwithin the lamellae was well preserved and not artificially induced during FIB\nmilling.\nFurthermore, as shown in Fig. 2a of the manuscript, the lamella included both\nfibroin-rich and sericin-rich regions. Notably, the sericin-rich region (left side) lacked\nnanofibrillar organization, while the fibroin-rich region (right side) displayed dense\nnanofibrillar networks. Since sericin is a globular protein and is not naturally fibrillar,\nthe absence of nanofibrils in this region argues strongly against the possibility that\nnanofibrils in the fibroin region are artifacts introduced by the FIB processing. This\ninternal spatial distinction within the same lamella supports the conclusion that the\nnanofibrillar organization observed in the fibroin region is intrinsic and not\nbeam-induced.\nAdditionally, recent advances in cryo-FIB milling technology have demonstrated that,\nwhen carefully implemented, this technique can preserve native biological structures\nwith high fidelity and has been widely used in in situ cryo-ET for cellular and\nmacromolecular imaging30-40.\nIn our study, we employed a state-of-the-art Thermo Fisher Helios NanoLab 600i\nDualBeam cryo-FIB/SEM system specifically optimized for cryo-electron microscopy\nsample preparation. Several key precautions were taken to minimize beam-induced\nheating and structural damage:\nUse of fresh, hydrated silk fibers:\nThe silk fibers used for cryo-FIB milling were freshly drawn from the silkworm\nspinneret, and retained intrinsic moisture. The presence of water enhances local\nelectrical conductivity and thermal dissipation, significantly reducing sample charging\nand heat accumulation during ion beam exposure. This moisture content helped\nmitigate thermal damage and the risk of graphitization, in contrast to dried cocoon\nsilk.\nCryogenic conditions throughout the FIB process:\nAll FIB milling procedures were conducted under deep cryogenic conditions (< –\n180 °C) using the Helios NanoLab system. At such low temperatures, biological\nsamples are vitrified, effectively preserving their native structures while dramatically\nreducing thermal diffusion and decomposition. This approach effectively eliminates\nthe risk of burning or graphitization, even in materials with low thermal conductivity\nlike silk.\nTriple-layer protective platinum coating:\nPrior to milling, we deposited three successive layers of protective platinum (Pt) on\nthe fiber surface. These coatings:\n· Enhanced surface conductivity,\n· Provided shielding from direct ion beam exposure,\n· Distributed both charge and heat more evenly,\n· Prevented sputtering and beam-induced structural degradation.\nLow current milling during final thinning steps:\nFor the final thinning of the lamellae, we employed extremely low ion beam currents\n(10 pA) to minimize any residual heat generation. This is a widely accepted approach\nin cryo-FIB protocols to prevent damage in sensitive specimens.\nCollectively, these precautions ensured that sample preparation was conducted with\nminimal physical or thermal disturbance. Most importantly, the consistent\nmorphological features observed between cryo-ET reconstructions from FIB-milled\nlamellae and independently prepared NSF samples strongly supports the conclusion\nthat cryo-FIB milling did not introduce significant structural artifacts or thermal\ndamage in our study.\nOnce again, we sincerely thank the reviewer for raising this critical point, which has\nhelped us improve the rigor and clarity of our methodological description.\nSpecific Comments:\nPage 3, line 108: Applying shear and flow to a shear-sensitive protein is inadvisable.\nResponse: We thank the reviewer for raising this important concern and appreciate\nthe opportunity to clarify our experimental approach and the terminology used in the\nmanuscript.\nWe agree that gel filtration chromatography, like all chromatographic processes,\ninvolve fluid flow and may introduce a certain degree of shear stress, which could, in\nprinciple, influence the morphology of shear-sensitive proteins such as silk fibroin.\nHowever, in our study, the sole purpose of performing gel filtration was to analyse the\nhomogeneity and composition of NSF. Therefore, even if minor shear-induced\nmorphological changes occurred during this purification step, they do not affect the\nvalidity of the subunit identification, which was the only purpose of that procedure.\nImportantly, we emphasize that gel filtration chromatography or any other\nshear-inducing purification steps was not used in the preparation of samples for\nvitrification, metal shadowing, or cryo-electron tomography (cryo-ET).\nWe acknowledge that the methodology description in the original manuscript may\nhave caused confusion. In the revised version, we have clarified the experimental\nworkflow and explicitly stated which samples underwent gel filtration and for what\npurpose, in order to avoid misunderstanding and improve the transparency and\nreadability of the manuscript (see materials and methods section, lines 346–349).\nWe sincerely appreciate the reviewer’s careful reading and insightful comment, which\nprompted this important clarification of our methods.\nPage 4, line 115: If another polymer (amphipol) is added to the sample, how can the\nauthors determine what they observe under the TEM microscope? Is it fibroin or the\nsecond polymer? How can they ensure that the second polymer does not alter the\nfibroin structure or macrostructure?\nResponse: We sincerely thank the reviewer for this insightful and important\ncomment.\nThe polymer referred to in this context is amphipol, a short-chain amphipathic\npolymer widely used in structural biology to replace detergents at the transmembrane\nsurface of membrane proteins, thereby stabilizing their structure in aqueous\nsolution17,18. Amphipol is chemically distinct from silk fibroin and is known not to\nform fibrillar or ordered structures on its own in solution.\nNonetheless, we fully agree with the reviewer that clearly distinguishing between\nprotein and polymer contributions in our data is essential for clarity. In response to\nthis concern, we have included additional metal shadowing electron microscopy\nresults in the revised manuscript (lines127-129, supplementary Fig. 1c-e) that\ncompare the morphology of samples with and without amphipol. These results show\nthat amphipol alone does not form any nanofibril structures under the experimental\nconditions used (Fig. 1a), while NSF retains its characteristic nanofibrillar\nmorphology even in the absence of amphipol (Fig. 1b-c). Thus, the nanofibril\nstructures observed in our experiments can be confidently attributed to NSF and not to\namphipol.\nFig. 1 Effect of amphipol on the morphology of NSF. (a) Metal shadowing image of amphipol\nalone (0.18 mg mL-1). (b) Metal shadowing image of NSF (0.03 mg mL-1) without amphipol. (c)\nMetal shadowing image of NSF mixed with amphipol at a mass ratio of 1:6 in the extraction\nbuffer (50 mmol L−1 NaCl, 50 mmol L−1 Bis-Tris, 0.5 mmol L−1 EDTA, pH 7.5), with a final NSF\nconcentration of 0.03 mg mL−1. Scale bar: 200 nm.\nThe addition of amphipol in selected assays was motivated by its ability to stabilize\nNSF under in vitro conditions, particularly during sample handling and preparation.\nNSF is known to be highly unstable once extracted, and tends to aggregate or\nprecipitate rapidly—even at low temperatures and in the absence of freezing19-22. In\nour previous work16, we performed extensive screening and found that both amphipol\nand digitonin significantly enhance the solubility and stability of NSF (Fig. 2a-b).\nTurbidity assays and circular dichroism (CD) analysis further confirmed that NSF\nremained stable for at least 336 hours in the presence of digitonin (Fig. 2c-e).\nMoreover, CD analysis showed that digitonin did not appreciably alter the random\ncoil structure of freshly extracted NSF (Fig. 2f). Although amphipol similarly\nstabilizes NSF in solution, its high absorbance interferes with CD spectroscopy.\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 2. Digitonin and Amphipol stabilized NSF in vitro. (a) A280 of NSF over time in the\npresence of MSP1D1 or BSA. (b) A280 of NSF over time in the presence of detergents. (c)\nNSF turbidity analysis. The inset (left) showed that NSF was stable in the presence of digitonin\nafter 336 h. The inset (right) showed the aggregation of NSF in the absence of digitonin after\n336 h. (d) CD spectra of NSF in the presence of digitonin over time. (e) CD spectra of NSF in the\nabsence of digitonin over time. (f) CD spectra of NSF in the presence or absence of digitonin.\nTaken together, these findings validate the use of amphipol and digitonin as essential\nstabilizing agents that do not introduce structural artifacts or interfere with the\ninterpretation of NSF morphology. All structural analyses and tomographic\nreconstructions were performed on samples in which amphipol was either excluded or\nits influence empirically shown to be negligible.\nWe once again thank the reviewer for this valuable comment, which helped us\nsignificantly improve the clarity and rigor of our methodological description in the\nrevised manuscript.\nPage 4, line 147: Are they discussing artificially spun fibers? Is it forced spinning\nfrom silkworms? This should be clarified. Why didn't they use natural silk fibers?\nWhat is a \"molted silkworm\"?\nResponse: We sincerely thank the reviewer for raising these important questions and\nallowing us to clarify the sample source used in our study.\nIn the section titled \"In situ structure of fibroin in silkworm silk\", the silk fibers\nanalyzed were not artificially spun fibers, but rather freshly drawn from living\nthird-instar silkworms. The term “3-molted silkworm” used in the original manuscript\nrefers to larvae that have undergone three molts, corresponding to the fourth-instar\nstage in the Bombyx mori life cycle. In fact, the samples were obtained from\nthird-instar silkworms. We sincerely apologize for this error and are grateful to the\nreviewer for pointing it out. To improve clarity and accuracy, we have replaced this\nterm with “third-instar” in the revised manuscript.\nIn this study, we did not use mature cocoon silk for two primary reasons:\nTechnical limitations associated with cryo-lamella preparation and imaging:\nDried cocoon silk tends to introduce significant image drift during tilt-series\nacquisition in cryo-ET. We attribute this to the complete dehydration of cocoon silk,\nwhich leads to poor electrical conductivity and charge accumulation during imaging.\nIn contrast, silk freshly drawn from third-instar silkworms and spiders retains\nsufficient moisture to facilitate vitrification, enhances electrical conductivity, and\nenables higher quality cryo-FIB lamella preparation and data acquisition.\nSample thickness requirements for cryo-FIB: Efficient vitrification requires that\nsample thickness not exceed 10 µm. Cocoon silk fibers typically range from 20–30\nµm in diameter, making them difficult to vitrify during cryo-fixation. In contrast, silk\nfibers freshly drawn from third-instar silkworms generally measure 3–5 µm in\ndiameter, making them ideal for high-resolution cryo-EM analyses.\nFinally, to ensure the physiological relevance of the fibers studied, we carefully\ncontrolled the pulling rate during reeling to approximate the natural spinning speeds\nof silkworms (4-15 mm s-1) and spiders (10-20 mm s-1)41. We therefore believe that\nthe silk fibers freshly drawn from third-instar silkworms and spiders in our study\nclosely represent the native state of silk proteins as secreted and aligned during the\nnatural spinning process.\nWe have revised the manuscript to clarify these points and are grateful to the reviewer\nfor these thoughtful comments, which have helped us improve the accuracy and\nprecision of our descriptions.\nPage 4, line 148: Every FIB lamella preparation of silk needs to be validated with\nproof that it was not carbonized (e.g., through EDS or diffraction). Silkworm silk\nfibers have very low thermal conductivity, so during FIB milling, they accumulate\nenergy and can reach high temperatures (>1000°C), leading to pyrolysis and\ncarbonization.\nResponse: We sincerely thank the reviewer for highlighting this critical concern\nregarding potential carbonization during cryo-FIB milling, especially given the\ninherently low thermal conductivity of silkworm silk fibers.\nWe fully acknowledge the potential concern that focused ion beam (FIB) milling,\neven under cryogenic conditions, could induce structural alterations in sensitive\nbiological materials, particularly protein-based fibers. This issue was carefully\nconsidered during our experimental design, especially regarding the reliability of\nultrastructural features observed in lamellae. However, although we lacked access to\nadvanced analytical tools (such as EDS) and knowledge reserves to directly evaluate\npotential damage (e.g., carbonization) in lamellae, we performed structural\ncross-validation using an independent experimental approach. Specifically, we\nconducted single-particle cryo-electron tomography (cryo-ET) and subtomogram\naveraging on NSF extracted from silk gland that had not undergone FIB processing.\nRemarkably, the basic structural units observed in the FIB-milled lamellae closely\nmatched those seen in the cryo-ET reconstructions from extracted NSF samples,\nproviding strong biochemical and morphological evidence that the ultrastructure\nwithin the lamellae was well preserved and not artificially induced during FIB\nmilling.\nFurthermore, as shown in Fig. 2a of the manuscript, the lamella included both\nfibroin-rich and sericin-rich regions. Notably, the sericin-rich region (left side) lacked\nnanofibrillar organization, while the fibroin-rich region (right side) displayed dense\nnanofibrillar networks. Since sericin is a globular protein and is not naturally fibrillar,\nthe absence of nanofibrils in this region argues strongly against the possibility that\nnanofibrils in the fibroin region are artifacts introduced by the FIB processing. This\ninternal spatial distinction within the same lamella supports the conclusion that the\nnanofibrillar organization observed in the fibroin region is intrinsic and not\nbeam-induced.\nAdditionally, recent advances in cryo-FIB milling technology have demonstrated that,\nwhen carefully implemented, this technique can preserve native biological structures\nwith high fidelity and has been widely used in in situ cryo-ET for cellular and\nmacromolecular imaging30-40.\nIn our study, we employed a state-of-the-art Thermo Fisher Helios NanoLab 600i\nDualBeam cryo-FIB/SEM system specifically optimized for cryo-electron microscopy\nsample preparation. Several key precautions were taken to minimize beam-induced\nheating and structural damage:\nUse of fresh, hydrated silk fibers:\nThe silk fibers used for cryo-FIB milling were freshly drawn from the silkworm\nspinneret, and retained intrinsic moisture. The presence of water enhances local\nelectrical conductivity and thermal dissipation, significantly reducing sample charging\nand heat accumulation during ion beam exposure. This moisture content helped\nmitigate thermal damage and the risk of graphitization, in contrast to dried cocoon\nsilk.\nCryogenic conditions throughout the FIB process:\nAll FIB milling procedures were conducted under deep cryogenic conditions (< –\n180 °C) using the Helios NanoLab system. At such low temperatures, biological\nsamples are vitrified, effectively preserving their native structures while dramatically\nreducing thermal diffusion and decomposition. This approach effectively eliminates\nthe risk of burning or graphitization, even in materials with low thermal conductivity\nlike silk.\nTriple-layer protective platinum coating:\nPrior to milling, we deposited three successive layers of protective platinum (Pt) on\nthe fiber surface. These coatings:\n· Enhanced surface conductivity,\n· Provided shielding from direct ion beam exposure,\n· Distributed both charge and heat more evenly,\n· Prevented sputtering and beam-induced structural degradation.\nLow current milling during final thinning steps:\nFor the final thinning of the lamellae, we employed extremely low ion beam currents\n(10 pA) to minimize any residual heat generation. This is a widely accepted approach\nin cryo-FIB protocols to prevent damage in sensitive specimens.\nCollectively, these precautions ensured that sample preparation was conducted with\nminimal physical or thermal disturbance. Most importantly, the consistent\nmorphological features observed between cryo-ET reconstructions from FIB-milled\nlamellae and independently prepared NSF samples strongly supports the conclusion\nthat cryo-FIB milling did not introduce significant structural artifacts or thermal\ndamage in our study.\nOnce again, we sincerely thank the reviewer for raising this critical point, which has\nhelped us improve the rigor and clarity of our methodological description (Lines\n432-436).\nPage 5, line 161: They barely discuss these bridges. What are their diameters? What\ndo they believe they are composed of?\nResponse: We sincerely thank the reviewer for this thoughtful comment.\nAs noted, the original manuscript included only limited discussion of the bridge\ndensities observed between nanofibrils. This is primarily because we currently lack\ndirect biochemical data or high resolution structural evidence to confidently determine\ntheir molecular composition. These bridges may represent silk-associated proteins or\nintermolecular linkages between fibroin nanofibrils. However, without definitive data,\nsuch interpretations remain speculative.\nAdditionally, these densities are relatively poorly resolved in the reconstructed\ntomograms, making it technically challenging to accurately determine their\ndimensions. Therefore, we have chosen to describe these features objectively, without\nover-interpreting their structural characteristics. A simplified schematic has been\nincluded to help visualize their relative positioning within the nanofibrillar\narchitecture.\nWe agree that these inter-fibrillar structures are intriguing and likely biologically\nsignificant. Future studies incorporating biochemical labeling or targeted probes may\nhelp elucidate their composition and function. We appreciate the reviewer’s insightful\ncomment, which has helped us refine the scope and clarity of our discussion.\nPage 5, lines 166-168: This statement is incorrect. Multiple studies have shown that\nthe protein undergoes significant changes along the silk gland.\nResponse: We thank the reviewer for pointing this out.\nWe fully acknowledge that numerous prior studies have demonstrated that silk fibroin\nundergoes significant secondary structural transitions—particularly from random coil\nto β-sheet structures—as it progresses along the silk gland26,42-45. However, in our\nstudy, we focus specifically on the morphology of the fundamental structural units\nwithin the silk fibers, rather than their secondary structure. Both our in vitro extraction\nand in situ imaging analyses consistently reveal that fibroin exists in the form of\nnanofibrils. In our previous work16, we also showed that this nanofibrillar morphology\nis preserved across different regions of the silk gland.\nBased on these observations, we concluded that the morphology of fibroin nanofibrils\nremains largely unchanged as they transit through the silk gland. This morphological\nconsistency does not contradict the well-documented changes in secondary structure.\nPage 6, lines 211-212: I do not understand how they interpret the \"perpendicular axis\"\nand how it correlates with the macro-fiber structure.\nResponse: We sincerely thank the reviewer for this thoughtful comment.\nThe “perpendicular axis” we refer to likely represents a structural element or factor\ninvolved in the formation of the herringbone patterns. Previous studies have proposed\nthat such an axis may arise from interactions between the N-terminal domains of\nopposing nanofibrils16. However, in our current study, we observed that this slender\ncentral axis remains clearly visible even in regions where surrounding nanofibrils are\nabsent. This observation suggests that the axis is unlikely to result solely from\nN-terminal interactions between fibroin molecules, and instead may involve\nadditional molecular components or organizing elements responsible for herringbone\npattern formation.\nImportantly, we previously reported abundant herringbone patterns in the anterior\nregion of the silk gland near the spinneret of the silkworm16. In the present study, we\nfurther show that these characteristic structures are retained in the native silk fiber\nitself. Notably, we also observed similar herringbone patterns in the third limb of the\nduct in spider silk gland, suggesting that this morphological feature is conserved\nacross distinct silk-producing species.\nThese findings underscore a key distinction between natural and artificial spinning\nprocesses. Specifically, it suggests that hierarchical structures such as herringbone\npatterns are pre-organized during the natural spinning process, rather than being\nformed post-extrusion through mechanical drawing, as is common in artificial fiber\nproduction.\nPage 9, line 311: This freezing process destroys the samples! Freezing such large\nsamples results in ice crystal formation, which forces the sample into an unnatural\nmorphology. Additionally, ice crystal formation exerts shear on the silk proteins,\naltering their structure and fibrillation.\nResponse: We sincerely thank the reviewer for this thoughtful question.\nThe decision to freeze the entire silkworm was primarily based on practical\nconsiderations. Rearing silkworms is time-consuming, as each lifecycle spans\napproximately 60 days, and the availability of fresh mulberry leaves becomes highly\nlimited during the winter season. To ensure experimental continuity and access to NSF,\nwe opted to freeze whole silkworms and subsequently purify NSF from their silk\nglands as needed, thereby eliminating the need for continuous rearing.\nWe acknowledge the general concern that freezing biological specimens may induce\nirreversible structural changes, such as precipitation or denaturation. Indeed, previous\nstudies and our own observations indicate have shown that extracted NSF undergoes\nirreversible precipitation when frozen in liquid nitrogen23,24, likely due to\nshear-induced aggregation during the freeze-thaw cycle25,26. It is well known that\nfreezing and thawing induce regenerated silk fibroin (RSF) or NSF to form porous\nstructures in vitro27-29. However, it is important to emphasize that the solution\nmicroenvironment of RSF or NSF in vitro differs significantly from its native\nphysiological environment within the silk gland lumen. The physiological conditions\ninside the gland—including specific ion concentrations, pH, and potential stabilizing\nbiomolecules—appear to play a protective role in preserving the integrity of NSF\neven under freezing conditions. Moreover, the intact silkworm body may further\nbuffer the silk gland against rapid temperature fluctuations and mechanical stress,\nthereby reducing the likelihood of ice crystal-induced damage.\nIn support of this, our previous work16 demonstrated that soluble NSF could be\nsuccessfully extracted from silk glands of frozen silkworms (Fig. 1a-b), suggesting\nthat the physiological environment within the lumen of silk gland—including ions, pH,\nand possibly unknown protective factors—may be essential roles in preventing NSF\ndenaturation or coagulation. Furthermore, SDS-PAGE analysis revealed that NSFs\npurified from the posterior silk gland (PSG), the posterior region of middle silk gland\n(PMSG), the middle region of middle silk gland (MMSG) and the anterior region of\nmiddle silk gland (AMSG) of frozen silkworms exhibited identical electrophoretic\nmobility and molecular composition as NSF extracted from the anterior silk gland\n(ASG) of living silkworm (Fig. 1b-c).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 1. Purification of NSF from silk gland. (a) Silk gland of the silkworm. The silk gland is\ndivided into ASG, MSG and PSG. MSG is further divided into AMSG, MMSG and PMSG. (b)\nNSF was purified from PSG, AMSG, MMSG and PMSG of frozen silkworm, respectively, and\nthen separated by a linear 4%-16% gradient SDS-PAGE. (c) Top. A linear 4%-16% gradient\nSDS-PAGE analysis of NSF purified by ions exchange chromatography. M, marker; CR, the crude\nextraction of ASG of living silkworm; F1, the flow-through fraction after S cation exchange media;\nS, cation exchange media; F2, the flow-through fraction after Q anion exchange media; Q, anion\nexchange media. Bottom, NSF purified by size exclusion chromatography. The inset showed the\npurity of NSF valued by 4%-16% SDS-PAGE. Fractions 1 and 2 were pooled for CD spectra and\nmetal shadowing, respectively.\nMetal shadowing confirmed the formation of nanofibrils in all these samples, which\nclosely resembled those derived from living silkworms (Fig. 2a). In addition, circular\ndichroism (CD) spectra revealed NSFs from PSG, PMSG, MMSG and AMSG of\nfrozen silkworms exhibited secondary structures comparable to those of NSF from\nASG of living silkworms (Fig. 2b).\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 2. The morphology and structure of NSF. (a) Metal shadowing of NSF from PSG, PMSG,\nMMSG, AMSG of frozen silkworm and ASG of living silkworm. Scale bar, 200 nm. (b) CD\nspectra of NSF from PSG, PMSG, MMSG, AMSG of frozen silkworm and ASG of living\nsilkworm.\nTaken together, these results provide strong evidence that NSF extracted from frozen\nsilkworms retains its native morphology, molecular composition, and structural\nfeatures, thereby validating the use of frozen specimens in our study without\ncompromising the reliability of our findings. Once again, we sincerely appreciate the\nreviewer’s insightful comments.\nPage 9, line 314: This purification process involves passing the samples through\nporous scaffolds and applying high shear to the proteins, significantly changing their\nnative non-fibrillated state.\nResponse: We thank the reviewer for raising this important concern and appreciate\nthe opportunity to clarify our experimental approach and the terminology used in the\nmanuscript.\nWe agree that gel filtration chromatography, like all chromatographic processes,\ninvolve fluid flow and may introduce a certain degree of shear stress, which could, in\nprinciple, influence the morphology of shear-sensitive proteins such as silk fibroin.\nHowever, in our study, the sole purpose of performing gel filtration was to analyse the\nhomogeneity and composition of NSF. Therefore, even if minor shear-induced\nmorphological changes occurred during this purification step, they do not affect the\nvalidity of the subunit identification, which was the only purpose of that procedure.\nImportantly, we emphasize that gel filtration chromatography or any other\nshear-inducing purification steps was not used in the preparation of samples for\nvitrification, metal shadowing, or cryo-electron tomography (cryo-ET).\nWe acknowledge that the methodology description in the original manuscript may\nhave caused confusion. In the revised version, we have clarified the experimental\nworkflow and explicitly stated which samples underwent gel filtration and for what\npurpose, in order to avoid misunderstanding and improve the transparency and\nreadability of the manuscript (see materials and methods section, lines 346–349).\nWe sincerely appreciate the reviewer’s careful reading and insightful comment, which\nprompted this important clarification of our methods.\nPage 9, line 328: Why did the authors add amphipol? Do they understand what impact\nit will have on fibroin?\nResponse: We thank the reviewer for raising this important point.\nThe polymer referred to in this context is amphipol, a short-chain amphipathic\npolymer widely used in structural biology to replace detergents at the transmembrane\nsurface of membrane proteins, thereby stabilizing their structure in aqueous\nsolution17,18. Amphipol is chemically distinct from silk fibroin and is known not to\nform fibrillar or ordered structures on its own in solution.\nNonetheless, we fully agree with the reviewer that clearly distinguishing between\nprotein and polymer contributions in our data is essential for clarity. In response to\nthis concern, we have included additional metal shadowing electron microscopy\nresults in the revised manuscript (lines127-129, supplementary Fig. 1c-e) that\ncompare the morphology of samples with and without amphipol. These results show\nthat amphipol alone does not form any nanofibril structures under the experimental\nconditions used (Fig. 1a), while NSF retains its characteristic nanofibrillar\nmorphology even in the absence of amphipol (Fig. 1b-c). Thus, the nanofibril\nstructures observed in our experiments can be confidently attributed to NSF and not to\namphipol.\nFig. 1 Effect of amphipol on the morphology of NSF. (a) Metal shadowing image of amphipol\nalone (0.18 mg mL-1). (b) Metal shadowing image of NSF (0.03 mg mL-1) without amphipol. (c)\nMetal shadowing image of NSF mixed with amphipol at a mass ratio of 1:6 in the extraction\nbuffer (50 mmol L−1 NaCl, 50 mmol L−1 Bis-Tris, 0.5 mmol L−1 EDTA, pH 7.5), with a final NSF\nconcentration of 0.03 mg mL−1. Scale bar: 200 nm.\nThe addition of amphipol in selected assays was motivated by its ability to stabilize\nNSF under in vitro conditions, particularly during sample handling and preparation.\nNSF is known to be highly unstable once extracted, and tends to aggregate or\nprecipitate rapidly—even at low temperatures and in the absence of freezing19-22. In\nour previous work16, we performed extensive screening and found that both amphipol\nand digitonin significantly enhance the solubility and stability of NSF (Fig. 2a-b).\nTurbidity assays and circular dichroism (CD) analysis further confirmed that NSF\nremained stable for at least 336 hours in the presence of digitonin (Fig. 2c-e).\nMoreover, CD analysis showed that digitonin did not appreciably alter the random\ncoil structure of freshly extracted NSF (Fig. 2f). Although amphipol similarly\nstabilizes NSF in solution, its high absorbance interferes with CD spectroscopy.\nReproduced from Song, K. et al. Decoding silkworm spinning programmed by pH and metal ions. Sci. Bull. 69, 792-802 (2024)\nunder CC BY 4.0 (https://creativecommons.org/licenses/by/4.0/).\nFig. 2. Digitonin and Amphipol stabilized NSF in vitro. (a) A280 of NSF over time in the\npresence of MSP1D1 or BSA. (b) A280 of NSF over time in the presence of detergents. (c)\nNSF turbidity analysis. The inset (left) showed that NSF was stable in the presence of digitonin\nafter 336 h. The inset (right) showed the aggregation of NSF in the absence of digitonin after\n336 h. (d) CD spectra of NSF in the presence of digitonin over time. (e) CD spectra of NSF in the\nabsence of digitonin over time. (f) CD spectra of NSF in the presence or absence of digitonin.\nTaken together, these findings validate the use of amphipol and digitonin as essential\nstabilizing agents that do not introduce structural artifacts or interfere with the\ninterpretation of NSF morphology. All structural analyses and tomographic\nreconstructions were performed on samples in which amphipol was either excluded or\nits influence empirically shown to be negligible.\nWe once again thank the reviewer for this valuable comment, which helped us\nsignificantly improve the clarity and rigor of our methodological description in the\nrevised manuscript.\nPage 9, line 329: Again, 72 hours is a long time to store NSF and expect it to remain\nunchanged.\nResponse: We appreciate the reviewer’s concern regarding the stability of NSF\nduring the 72-hour incubation period.\nBased on our longstanding efforts to resolve the overall structure of fibroin using\nsingle-particle cryo-EM, we have found that achieving a homogenous sample is\ncritical for high-quality structural analysis. Through extensive optimization, we\ndetermined that treating NSF with amphipol for 72 hours significantly improveds\nsample homogeneity, making it more suitable for cryo-EM.\nTo ensure that this extended incubation does not introduce structural artifacts, we\nperformed a series of control experiments. Turbidity measurements, CD spectroscopy,\nand metal shadowing EM imaging all consistently showed that NSF incubated with\namphipol or digitonin remained both structurally and morphologically stable for up to\n15 days at 4°C. In contrast, untreated NSF rapidly underwent conformational changes,\neventually aggregating into β-sheet-rich precipitates16.\nThese results provide strong experimental evidence that the 72-hour incubation in the\npresence of amphipol preserves the native state of NSF while enhancing its suitability\nfor structural studies, without introducing detectable alterations.\nPage 10, line 334: How did they obtain spider silk proteins? This information is not\nmentioned.\nResponse: We thank the reviewer for pointing out the missing information regarding\nthe preparation of spider silk proteins.\nWe apologize for this oversight. In the revised manuscript, we have added a detailed\ndescription in the Materials and methods section. Briefly, the spinning dope of major\nampullate silk gland was extracted from Nephila pilipes spider as follows (Lines\n341-344).\n“Adult spiders were anesthetized at 4℃ for 30 min and then dissected on ice to isolate\nthe major ampullate glands. The silk glands were rinsed three times with phosphate\nbuffered saline (PBS), and the outer epithelial cell layer was carefully removed using\nfine forceps. The gel-like spinning dope was gently extracted and transferred into PBS\nto prepare the NSP solution.”\nWe have revised the manuscript to include this information and thank the reviewer for\npointing out this important detail.\nPage 10, lines 338-339: The authors state that NSF has been dehydrated in ethanol…\nWHY? Dehydration with ethanol will completely alter the protein structure!\nResponse: We thank the reviewer for raising this important concern.\nEthanol, as an organic solvent, can in some cases alter protein structure when added\ndirectly to protein solutions. However, in this study, the ethanol dehydration step used\nin the metal shadowing experiment was performed after the protein samples had been\nadsorbed onto the carbon film of the EM grids. At this stage, the samples were already\npartially immobilized, and ethanol dehydration served primarily to enhance adhesion\nbetween the protein and the carbon support, thereby minimizing sample loss during\nsubsequent grid transfer and drying steps. In addition, ethanol treatment helps reduce\nsurface tension on the grid, facilitating smoother handling during processing.\nTo directly evaluate whether ethanol dehydration affects the ultrastructure of NSF, we\nconducted control experiments in which metal shadowing was performed both with\nand without the ethanol dehydration step. As shown in Fig. 3, NSF exhibited similar\nfibrillar morphology under both conditions, indicating that ethanol dehydration does\nnot introduce discernible artifacts in terms of ultrastructural preservation.\nFig. 3. The effect of ethanol dehydration on the morphology of NSF. (a) Metal shadowing of NSF\nwith ethanol dehydration. (b) Metal shadowing of NSF without ethanol dehydration. Scale bar: 200 nm.\nPage 10, lines 348-349: Why did they add the buffer? Can't they spin it without buffer?\nIt is NSF; they should be able to spin it easily.\nResponse: We thank the reviewer for this important question.\nIn addition to natural silk fibers, artificial silks have attracted increasing interest due\nto their potential for scalable production and customizable properties. Artificial silk is\ntypically produced by regenerated silk fibroin1,2 or recombinant silk proteins3-7 using\ntechniques such as wet spinning1-4,6-8, dry spinning9,10, electrospinning5, interface\nwiredrawing11 or microfluidic spinning12,13. These processes aim to fabricate\nartificial silk fibers with structures and properties comparable to those of natural silk,\nbut the properties of resulting artificial silk fibers are particularly influenced by\nfactors such as protein sequence, spinning dope concentration, flow rate, humidity,\npost-spin stretching, buffer composition, etc10,14,15.\nIn this study, we employed an interface wiredrawing technique to fabricate artificial\nsilk fibers, which requires an optimized spinning buffer to facilitate fiber formation.\nAlthough the protein used in our study is NSF, once it is extracted from the silk gland\nlumen, it no longer retains the natural physiological environment necessary for\nspontaneous fiber assembly. Without the addition of an appropriate spinning buffer,\nit is extremely difficult to generate continuous and uniform silk fibers using interface\nwiredrawing. Hence, the buffer is not intended to alter the protein itself, but rather to\ncompensate for the loss of the natural glandular environment and provide the minimal\nconditions required to trigger and stabilize fiber formation in vitro. The ultrastructure\nwithin artificial silk is even less understood. Understanding these structural\ndiscrepancies is crucial for improving artificial silk production and bridging the\nperformance gap between synthetic and natural materials.\nPage 10, line 352: “Forced silking” will also change the structure!\nResponse: We appreciate the reviewer’s concern regarding the potential structural\nimpact of forced silking\nIt is important to note that, in nature, both silkworm cocoon formation and spider web\nweaving involve the natural pulling or drawing of silk proteins from the gland. The\nnatural spinning speeds are approximately 4-15 mm s-1 for silkworms and 10-20 mm\ns-1 for spiders41. In our study, we carefully calibrated the forced reeling speed to fall\nwithin these physiological ranges as closely as possible, thereby minimizing\nmechanical stress and preserving native structural features.\nTo further confirm that forced reeling does not introduce structural artifacts, we also\nperformed cryo-FIB milling and cryo-electron tomography on silk fibers extracted\ndirectly from silkworm cocoons. The lamellar ultrastructure of cocoon silk was\nconsistent with that observed in freshly drawn silk (Fig. 5), supporting the reliability\nof our approach. However, due to pronounced image drift during tilt-series acquisition\nof lamellae prepared from dried cocoon silk, we ultimately excluded these samples\nfrom tomographic data collection. We speculate that this instability arose from the\ncomplete dehydration of cocoon silk, which compromises its electrical conductivity.\nThe resulting charge accumulation during imaging likely caused the observed drift.\nIn contrast, freshly drawn silkworm silk and spider silk retain residual moisture,\nwhich enhances conductivity and stabilizes imaging during cryo-ET. Thus, our choice\nto use freshly reeled silks for tomographic analysis was driven primarily by their\nsuperior performance under cryo-imaging conditions.\nFig. 5. Comparison of the ultrastructure of silkworm cocoon silk and freshly drawn silk. (a)\nLow-magnification projection image of the freshly drawn silk from third-instar silkworm. (b)\nLow-magnification projection image of silkworm cocoon silk.\nTaken together, we have systematically addressed each potential source of artifact and\nprovided extensive experimental evidence demonstrating that the ultrastructure,\nmolecular composition, and secondary structure of silk proteins are preserved\nthroughout each key step of our workflow. While we acknowledge that no in vitro\nmethod is entirely free of manipulation, we believe that our carefully controlled\nworkflow and multiple levels of internal validation ensure that the structures reported\nhere are reliable and faithfully reflect native silk protein assemblies.\nPage 10, line 358: Everything done using FIB milling needs to be checked to ensure\nthat the material has not been altered due to thermal effects caused by the beam. Even\nunder gentle milling settings and cryogenic conditions, silkworm silk fibers can heat\nup and form graphene structures. To ensure this is not the case, they should use EDS\nto check the elemental composition of the lamellae and/or perform electron\ndiffraction.\nResponse: We sincerely thank the reviewer for raising this important and technically\ninsightful concern.\nWe fully acknowledge the potential concern that focused ion beam (FIB) milling,\neven under cryogenic conditions, could induce structural alterations in sensitive\nbiological materials, particularly protein-based fibers. This issue was carefully\nconsidered during our experimental design, especially regarding the reliability of\nultrastructural features observed in lamellae. However, although we lacked access to\nadvanced analytical tools (such as EDS) and knowledge reserves to directly evaluate\npotential damage (e.g., carbonization) in lamellae, we performed structural\ncross-validation using an independent experimental approach. Specifically, we\nconducted single-particle cryo-electron tomography (cryo-ET) and subtomogram\naveraging on NSF extracted from silk gland that had not undergone FIB processing.\nRemarkably, the basic structural units observed in the FIB-milled lamellae closely\nmatched those seen in the cryo-ET reconstructions from extracted NSF samples,\nproviding strong biochemical and morphological evidence that the ultrastructure\nwithin the lamellae was well preserved and not artificially induced during FIB\nmilling.\nFurthermore, as shown in Fig. 2a of the manuscript, the lamella included both\nfibroin-rich and sericin-rich regions. Notably, the sericin-rich region (left side) lacked\nnanofibrillar organization, while the fibroin-rich region (right side) displayed dense\nnanofibrillar networks. Since sericin is a globular protein and is not naturally fibrillar,\nthe absence of nanofibrils in this region argues strongly against the possibility that\nnanofibrils in the fibroin region are artifacts introduced by the FIB processing. This\ninternal spatial distinction within the same lamella supports the conclusion that the\nnanofibrillar organization observed in the fibroin region is intrinsic and not\nbeam-induced.\nAdditionally, recent advances in cryo-FIB milling technology have demonstrated that,\nwhen carefully implemented, this technique can preserve native biological structures\nwith high fidelity and has been widely used in in situ cryo-ET for cellular and\nmacromolecular imaging30-40.\nIn our study, we employed a state-of-the-art Thermo Fisher Helios NanoLab 600i\nDualBeam cryo-FIB/SEM system specifically optimized for cryo-electron microscopy\nsample preparation. Several key precautions were taken to minimize beam-induced\nheating and structural damage:\nUse of fresh, hydrated silk fibers:\nThe silk fibers used for cryo-FIB milling were freshly drawn from the silkworm\nspinneret, and retained intrinsic moisture. The presence of water enhances local\nelectrical conductivity and thermal dissipation, significantly reducing sample charging\nand heat accumulation during ion beam exposure. This moisture content helped\nmitigate thermal damage and the risk of graphitization, in contrast to dried cocoon\nsilk.\nCryogenic conditions throughout the FIB process:\nAll FIB milling procedures were conducted under deep cryogenic conditions (< –\n180 °C) using the Helios NanoLab system. At such low temperatures, biological\nsamples are vitrified, effectively preserving their native structures while dramatically\nreducing thermal diffusion and decomposition. This approach effectively eliminates\nthe risk of burning or graphitization, even in materials with low thermal conductivity\nlike silk.\nTriple-layer protective platinum coating:\nPrior to milling, we deposited three successive layers of protective platinum (Pt) on\nthe fiber surface. These coatings:\n· Enhanced surface conductivity,\n· Provided shielding from direct ion beam exposure,\n· Distributed both charge and heat more evenly,\n· Prevented sputtering and beam-induced structural degradation.\nLow current milling during final thinning steps:\nFor the final thinning of the lamellae, we employed extremely low ion beam currents\n(10 pA) to minimize any residual heat generation. This is a widely accepted approach\nin cryo-FIB protocols to prevent damage in sensitive specimens.\nCollectively, these precautions ensured that sample preparation was conducted with\nminimal physical or thermal disturbance. Most importantly, the consistent\nmorphological features observed between cryo-ET reconstructions from FIB-milled\nlamellae and independently prepared NSF samples strongly supports the conclusion\nthat cryo-FIB milling did not introduce significant structural artifacts or thermal\ndamage in our study.\nOnce again, we sincerely thank the reviewer for raising this critical point, which has\nhelped us improve the rigor and clarity of our methodological description (Lines\n432-436).\nReferences\n1. Wang, J. X. et al. Artificial superstrong silkworm silk surpasses natural spider\nsilks. Matter 5, 4396-4406 (2022).\n2. Ha, S. W., Tonelli, A. E. & Hudson, S. M. Structural studies of Bombyx mori\nsilk fibroin during regeneration from solutions and wet fiber spinning.\nBiomacromolecules 6, 1722-1731 (2005).\n3. Schmuck, B. et al. High-yield production of a super-soluble miniature spidroin\nfor biomimetic high-performance materials. Mater. Today 50, 16-23 (2021).\n4. Qi, X. M. et al. 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Biochemistry-Us\n43, 14080-14087 (2004)."}, {"round": 2, "reviewer_comments": {"reviewer_2": "The revised manuscript by Song et al. has thoroughly addressed all my previous concerns. The added paragraph in the introduction and the updated discussion have improved the manuscript. Overall, this version of the manuscript demonstrates significant research effort with substantial data, solid analysis, and rigorous methodology. The revised version is ready for publication in Nature Communications.", "reviewer_3": "The authors have made visible efforts to address the previous comments. The revised manuscript is clearer, with improved methodological explanations and a more detailed discussion. The new paragraph on artificial silk and clarifications about cryo-FIB milling, amphipol use, and freezing procedures strengthen the technical transparency. However, several conceptual and interpretative issues remain only partially resolved. In particular, the paper continues to overstate its novelty regarding the “minimum structural unit” and makes claims about “in-situ” or “near-native” conditions that are not experimentally justified. While the Cryo-ET dataset itself is technically impressive and of potential importance for the silk field (furthermore, in my view, the work is impressive and deserve to be published in Nature communication journal), but the manuscript still requires substantial textual revision before it can be accepted.", "reviewer_author_response": "Response: Thank you very much for your constructive suggestions and guidance throughout the review process. We are grateful for your time and effort in evaluating our work, and we sincerely appreciate your recognition of the improvements made in the revised manuscript."}, "author_response": ""}]

Decision letter