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This naturally occurring protein protects your hair, skin, and nails

Structure and Function

  • Supplemental Keratin

Associated Conditions

Risks and side effects.

Keratin is a protein in the cells on the surface of the skin . The fingernails, hair , and skin need keratin to grow, function, and stay healthy.

Cosmetic treatments to improve hair and nail health are often enriched with keratin. Keratin also occurs naturally in some foods and can be taken in supplement form as well.

This article will explain the different types of keratin, health conditions related to overproduction of keratin, and the possible risks of keratin-enriched cosmetic products.

Types of Keratin

There are 54 types of keratin produced by the body. Of these, 28 are considered type I and 26 belong to type II.

Type I keratins are categorized as being the smaller and more acidic type of keratin. They work to promote and maintain the health of the of epithelial cells on the outside of the body, i.e., areas that are exposed to the outside environment. Type I keratins are the major component of hair and nails. They are also found inside the mouth, urinary tract, vagina, and anus.

Type II keratins are larger than their type I counterparts and have a neutral pH. They are found in the internal epithelial cells, which line various structures and organs contained within the body, including the lungs, kidneys, liver, and digestive system.


Alpha-keratins are the exclusive form of keratin found in humans and in the wool of other mammals. The structure of the alpha-keratin is fibrous and helical, and both I and II keratins can fall under the category of alpha.


Beta-keratins are categorized as polypeptide chains and are only found in birds and reptiles, although those species can also possess alpha-keratins. Both alpha and beta keratins help these animals maintain the composition of their claws, scales, beaks, skin, and feathers.

In its natural form, keratin appears as long structures, with filaments arranged in bundles or fibers. These fibers are made up of individual keratin molecules that are cross-linked together, forming a tough and resilient protein. This is what gives keratin the ability to provide structure and protection to the areas of the body it inhabits.

Keratin cannot be dissolved in water, solvents, acids, or alkalines, so its structure remains largely intact when exposed to many of the body's natural chemicals.

Uses of Supplemental Keratin

Many industries have used keratin or other vitamin supplements that promote production of keratin as a form of maintaining or improving hair, skin, and nail health. The beauty industry in particular advertises keratin as a way to combat damaged hair.

Hair Treatments and Products

Hair is made up of 90% keratin, which is why it is often a component in hair treatments that are used for shinier and stronger tresses.

Keratin hair treatments provided in a salon are chemical protein treatments (sometimes called Brazilian blowouts) that reduce frizz and make the hair look shiny and silky.

A stylist will first wash your hair with a special shampoo and then apply a liquid keratin solution to your hair in small sections. Then your hair will be blown dry. The stylist will use a flat iron to seal the keratin solution.

For optimal results, you should avoid washing your hair for a few days.

Depending on the type of keratin used, the results will vary widely. Studies have found isolated animal keratin to be less efficient overall than a synthetic version of human keratin.

Many people also use shampoos and conditioners infused with keratin oil to make their hair healthier.

Keratin used in hair treatments is often cultivated from animal sources, such as feathers, hooves, and wool, and therefore are not appropriate for people who follow a vegan lifestyle.

Biotin is a B vitamin that has also soared in popularity because it is said to have a positive impact on the body’s ability to synthesize proteins such as keratin, thus leading to healthier skin, hair, and nails.  

Soluble Keratin

Keratin is not an easily dissolvable protein. A soluble form of the protein is available and has been targeted towards athletes who wish to supplement their protein intake for athletic performance.

Food Sources of Keratin

Nutrients such as biotin, vitamin A, and zinc can boost keratin production in the body. Some food sources rich in these nutrients include:

  • Sweet potatoes
  • Sunflower seeds

Hyperkeratosis is an umbrella term for a variety of skin conditions that result from the excessive production of keratin proteins. This can be caused by inflammation, genetics, or for unknown reasons.

Forms of hyperkeratosis include:

  • Keratosis pilaris ("chicken skin") : Keratosis pilaris occurs when keratin clogs pores and blocks hair follicles, giving the skin a bumpy appearance. It is not dangerous in any way.
  • Actinic keratosis : This skin condition is very common and is associated with sun exposure. It causes lesions on the body that can feel like rough sandpaper. The lesions are considered a precursor to skin cancer. Your healthcare provider may monitor your skin and/or treat the lesions.
  • Epidermolytic hyperkeratosis : This form of hyperkeratosis is inherited and it is present in infants at birth.
  • Lichen planus : This is a type of inflammatory disorder that most commonly affects the flexor (inner) surfaces of the arms and legs. It looks like a purple rash with flat, itchy areas.
  • Calluses and corns : Calluses and corns appear on the foot, especially where there is daily pressure and friction. This creates micro-injuries that lead to an overproduction of keratin, causing lesions to form.
  • Chronic folliculitis : Hyperkeratosis can cause chronic folliculitis, which is an inflammation of the hair follicles that occurs when the follicles become blocked.
  • Atopic dermatitis (eczema) : This is a chronic inflammatory skin condition that causes an itchy rash to appear on the skin. 
  • Psoriasis and psoriasiform dermatitis : Psoriasiform dermatitis is a chronic condition that causes thick, silvery scales to appear on the skin. Psoriasis is the most common form of psoriasiform dermatitis.
  • Lichenoid dermatitis : Lichenoid dermatitis is a chronic skin condition that causes itchy, red scales to appear on the skin. It can appear on any part of the body but is most common on the scalp, face, neck, arms, and legs.
  • Ichthyoses : This is a group of skin disorders characterized by scaly, red, dry, itchy skin. The condition is usually genetic but some medical conditions and medications can also cause ichthyosis.  

Treatment of hyperkeratosis depends on the specific condition:

  • Keratosis pilaris is treated with exfoliants containing salicylic acid and retinols. Creams containing urea are also effective. These products are available over-the-counter.
  • For lichen planus , a corticosteroid cream may be prescribed by your healthcare provider.
  • Actinic keratosis treatments include prescription creams (such as Effudex, Aldera, and others), cryosurgery with liquid nitrogen, photodynamic therapy, chemical peels, and CO2 laser ablation. Very thick lesions may be removed with curettage or scraping followed by heating the area to ensure the lesion has been destroyed.
  • Epidermolytic hyperkeratosis may be treated with sea salt baths, oral retinoids (and possibly experimental treatments involving gene therapy).
  • Calluses and corns : Calluses and corns are usually removed manually. They can be treated at home by soaking the affected area and rubbing the callus or corn with a pumice stone or emery board. Thicker calluses and corns may need to be removed by a healthcare provider with a surgical blade.
  • Atopic dermatitis : Treatment may include topical creams or ointments such as a corticosteroid or tacrolimus ointment, light therapy, or oral or injectible prescription medication. Taking good care of your skin by bathing regularly and choosing gentle cleansers and moisturizers can also help improve symptoms.  
  • Psoriasis : Psoriasis can't be cured, but it can be treated with topical medications such as steroid creams and non-steroid alternatives such as anthralin. Oral medications and light therapy can also help.  
  • Ichthyoses : There is no cure for ichthyosis, but the symptoms can be controlled with hydrating creams and ointments and/or oral or topical retinoids.   

Although there is not a lot of evidence suggesting that using keratin by itself is dangerous to hair, skin, and nail health, the chemicals that may be added to keratin hair treatments can have adverse effects. Formaldehyde exposure has been a problem for those who use hair products with keratin regularly.

The use of formaldehyde in these products can lead to health issues, including:

  • Itching and stinging eyes
  • Nose and throat irritation
  • An allergic reaction
  • Itchy skin with or without a rash
  • Scalp irritation that may present with burns or blisters
  • Mood changes
  • Hair loss and damage

Extended exposure to formaldehyde has also been shown to have carcinogenic (cancer-causing) effects.

Opt for keratin treatments that are free of unwanted chemicals and substances such as formaldehyde to avoid unnecessary risks to your overall health.

Keratin is a naturally occurring protein in the body that is found in the hair, skin nails, mouth, and internal organs. It plays a key role in providing structure and protection to the skin and tissues.

Certain foods contain minerals and vitamins that boost the production of keratin. It is also available in supplement form and in various over-the-counter shampoos, conditioners, and cosmetics. Professional keratin treatments are available in salons to make the hair stronger and shinier.

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Antończak PP, Hartman-Petrycka M, Garncarczyk A, Adamczyk K, Wcisło-Dziadecka D, Błońska-Fajfrowska B. The effect of callus and corns removal treatments on foot geometry parameters, foot pressure, and foot pain reduction in women . Appl Sci . 2023;13(7):4319. doi:10.3390/app13074319

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By Angelica Bottaro Angelica Bottaro is a professional freelance writer with over 5 years of experience. She has been educated in both psychology and journalism, and her dual education has given her the research and writing skills needed to deliver sound and engaging content in the health space.

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  • Open access
  • Published: 19 November 2022

Keratin-mediated hair growth and its underlying biological mechanism

  • Seong Yeong An 1 ,
  • Hyo-Sung Kim 2 ,
  • So Yeon Kim 1   nAff7 ,
  • Se Young Van 1 ,
  • Han Jun Kim 2   nAff8 ,
  • Jae-Hyung Lee   ORCID: orcid.org/0000-0002-5085-6988 3 ,
  • Song Wook Han 4 ,
  • Il Keun Kwon 5 ,
  • Chul-Kyu Lee 6 ,
  • Sun Hee Do   ORCID: orcid.org/0000-0003-2372-5269 2 &
  • Yu-Shik Hwang   ORCID: orcid.org/0000-0001-6197-7542 1  

Communications Biology volume  5 , Article number:  1270 ( 2022 ) Cite this article

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  • Biomaterials – cells
  • Stem-cell niche

An Author Correction to this article was published on 22 December 2022

A Publisher Correction to this article was published on 22 December 2022

This article has been updated

Here we show that intradermal injection of keratin promotes hair growth in mice, which results from extracellular interaction of keratin with hair forming cells. Extracellular application of keratin induces condensation of dermal papilla cells and the generation of a P-cadherin-expressing cell population (hair germ) from outer root sheath cells via keratin-mediated microenvironmental changes. Exogenous keratin-mediated hair growth is reflected by the finding that keratin exposure from transforming growth factor beta 2 (TGFβ2)-induced apoptotic outer root sheath cells appears to be critical for dermal papilla cell condensation and P-cadherin-expressing hair germ formation. Immunodepletion or downregulation of keratin released from or expressed in TGFβ2-induced apoptotic outer root sheath cells negatively influences dermal papilla cell condensation and hair germ formation. Our pilot study provides an evidence on initiating hair regeneration and insight into the biological function of keratin exposed from apoptotic epithelial cells in tissue regeneration and development.


Keratin is a cytoskeletal protein that forms intermediate filaments within epithelial cells and participates in maintaining the strength of the cells 1 . It is a major protein found within the hair that contributes to its mechanical strength 2 . Human hair consists of three main layers: the medulla in the center of the hair, the cortex surrounding the medulla, which contains a fiber mass mainly consisting of keratin protein, and the cuticle, the outer layer of the hair shaft 3 . During hair growth, dermal papilla (DP) cells secrete various paracrine factors to induce migration of stem cells from the bulge region of the outer root sheath (ORS) to the upper region of the follicle, and the migrated cells become transit amplifying cells, which then undergo differentiation into matrix cells. Hair growth is initiated by cortical cells differentiated from matrix cells located in the follicle bulb region, and a large amount of keratin is synthesized mainly in the cortex 4 , 5 , 6 . Deposition and rearrangement of keratin filaments are followed by the assembly of keratin-associated proteins and intracellular deposited keratin in spindle-shaped epithelial cells of the cortex, and the assembly is stabilized by the formation of inter- and intra-molecular disulfide bonds 7 . At the stages of the anagen-catagen transition of the hair cycle, apoptosis of cells begins to appear in the epithelial strand, and then the apoptotic cells are phagocytosed by macrophages and neighboring epithelial cells. Ultimately, keratin remains the main protein in the hair 8 , 9 , 10 , 11 .

In our previous study, mouse models with full-thickness dorsal excisional wounds were used to assess the effect of keratin-based hydrogels on wound healing 12 . Interestingly, hair growth was observed only in areas treated with keratin hydrogel, along with accelerated wound healing, which led us to study the biological function of hair-derived keratin in hair regeneration. In this study, intradermal injection of human hair-derived keratin promoted hair follicle formation and subsequent hair growth in a mouse model. Extensive research has been conducted to elucidate the biological function of keratin as an intermediate filament participating in intracellular signaling pathways and the mechanical role of keratin, and as an intracellular scaffold modulating cell stiffness and morphology in response to microenvironmental changes 13 , 14 . However, the control of cellular behavior via the extracellular interaction of keratin with cells has not been well studied, especially in hair growth. Hence, the underlying mechanism through which keratin stimulates hair growth was examined by studying the interaction of keratin with DP cells and ORS cells, which are known as the main types of cells that regulate hair growth and regeneration, and the whole experimental procedure is illustrated in supplementary fig.  1 . DP condensation and the generation of a P-cadherin expressing cell population (hair germ, HG) were induced via the extracellular interaction of DP and ORS cells with keratin. Furthermore, DP condensation and P-cadherin-expressing HG formation were mediated by spatial exposure from TGFβ2-induced apoptotic ORS cells and deposition of keratin, which represents the role of injected exogenous keratin in a similar manner as the exposed keratin from apoptotic ORS cells during hair cycle. Our pilot study provides a possible explanation that keratin is not only a structural protein of hair but also a factor that induces hair regeneration.

Intradermal injection of hair-derived keratin promotes hair growth

First, we performed in vivo experiments in mice to evaluate keratin-mediated hair growth by injecting hair-derived keratin into the hair-removed dorsal skin area. The dorsal area of C57BL/6 mice was shaved with an animal clipper, and 0.5% (w/v) and 1.0% (w/v) keratin were injected. Twenty-eight days after keratin injection, hair growth and hair follicle formation in keratin-injected C57BL/6 mice were analyzed for the number and stage of hair follicles. We found that hair growth and the formation of anagen follicles were promoted, compared to non-treated mice (Fig.  1a–d ). Only a single injection of keratin resulted in much higher hair growth compared to the control, and almost equivalent hair growth compared to minoxidil, applied every day for 28 days. In addition, in situ RNA hybridization showed an increase in the Lgr5-positive cell populations in the lower bulge region and hair follicles after keratin injection (Supplementary Fig.  1 ). Such keratin-mediated hair growth was confirmed in a separate mouse study, and the promoting effect of keratin injection on hair growth was verified. There were no differences in the number of hair follicles formed between mice injected with keratin and those treated with minoxidil (Supplementary Fig.  2a–c ). However, the number of anagen follicles and the size of hair follicles were found to be increased in keratin-injected mice compared to minoxidil-treated mice (Supplementary Fig.  2d–f ). Keratin injection-mediated hair growth was observed throughout the surface of the back skin of mice and mixed stages of hair follicles were also found, which might be due to the dispersion of keratin solution after injection, and the injected keratin remained up to 2 weeks after injection (Supplementary Fig.  3 ).

figure 1

a Images of hair growth on the back skin of mice at day 1, day 14, and day 28 after injection of keratin. b Histological images of the back skin of mice at 4 weeks after injection of keratin. c , d Graphical representation and quantification of hair follicles with different hair cycle stages in skin sections of mice ( n  = 12 sections, in six mice; mean ± standard deviation (s.d.)). * P ,0.05, indicates a difference between control and keratin-injected groups. Scale bars, 200 μm. e Graphical representation of DP cell condensation in the presence of keratin. * P ,0.01, indicates a difference between control and keratin-treated. ( n  = 6; mean ± standard deviation (s.d.)); Control, non-treated DP cell; keratin-treated, DP cells in the presence of keratin. f Condensation of DP cell by immunofluorescent staining in the presence of keratin; 4′,6-diamidino-2-phenylindole (DAPI), blue; SOX2, alkaline phosphatase (ALPase), FGF10, BMP6, red; β-catenin, CD133, and FGF7, green. Scale bars, 100 μm.

Keratin induces condensation of DP cells and germ formation from ORS cells in vitro

To understand how injected keratin induces hair follicle formation and growth, we studied the interaction of keratin with DP and ORS cells, which are known to be the main cells participating in hair follicle formation 4 , 5 , 6 , 15 . The most distinct characteristic of DP cells exposed to keratin for 3 days was their condensation to form spherical aggregates (Fig.  1e, f ) with high expression levels of β-catenin, SOX2, CD133, and alkaline phosphatase (ALPase) (Fig.  1f ), which is a molecular identity signature reflecting the hair-inductive property of DP cells 16 , 17 , and high expression levels of FGF7, FGF10, and BMP6 (Fig.  1f ), reflecting paracrine factors controlling hair growth 18 (keratin-mediated condensation of DP cells on day 1 and time-lapse images are presented in Supplementary Fig.  4 ). However, the growth of DP cells was suppressed upon exposure to keratin (Supplementary Fig.  5a ), and relatively lower Ki67 expression and BrdU incorporation were observed in keratin-treated DP cells (Supplementary Fig.  5b–d ). Condensed DP cell aggregates contained cells expressing Ki67, and β-catenin was expressed in the contact region between cells within DP cell aggregates (Supplementary Fig.  6 ), in which DP cells remained viable during culture (Supplementary 7). Such keratin-mediated condensation of DP cells was observed when cells were seeded at different cell densities on Matrigel (Supplementary Figs.  8 and 9 ), and no difference in DP cell condensation was found when they were treated with different keratin concentrations (Supplementary Fig.  10 ). RNA sequencing analysis showed downregulation of the expression levels of genes associated with cell division and upregulation of mRNAs encoding proteins related to integrins, growth factors, migration, and extracellular matrix organization (Supplementary Fig.  11a, b ). Real-time qPCR analysis showed upregulation of various genes, such as CD133, SOX2, corin, SHH , versican , β -catenin, BMP6, FGF7 , and FGF10 , known as a molecular identity signature reflecting the hair-inductive property of DP cells 16 , 17 , 18 , 19 (Supplementary Fig.  12 ). In addition, the effect of keratin on maintenance of the condensed DP cell aggregates was analyzed. The DP cell spheroids showed higher levels of gene expression, indicating the hair-inductive property of DP cells as compared to a DP cell monolayer (Fig.  2a ). The spherical shape of DP cell aggregates was consistently maintained, showing high levels of expression of β-catenin, SOX2, CD133, ALPase, FGF7, FGF10, and BMP6 (Fig.  2b–d , Supplementary Fig.  13 ) in the presence of keratin.

figure 2

a Schematic illustration of microwell-mediated DP cell spheroid formation and images of DP cell spheroids within microwells and retrieved DP cell spheroids from microwells. Graphical representation of DP cell property-related gene expressions of DP cells and DP cell spheroids. * P ,0.01, indicates a difference between DP cells and DP cell spheroids; 2D, DP cells; 3D, DP cell spheroids. Scale bars, 100 μm. ( n  = 4; mean ± standard deviation (s.d.)). b Images of replated DP cell spheroids in the presence of keratin by observation using a light microscope. Scale bars, 100 μm. c Images of replated DP cell spheroids in the presence of keratin by immunofluorescent staining; DAPI, blue; SOX2, CD133, FGF10, BMP6, red; β-catenin, ALPase, FGF7, green. Scale bars, 100 μm. d Graphical representation of DP cell property-related molecular expressions of replated DP cell spheroid culture in the presence of keratin; ELISA. * P ,0.01, indicates a difference between control and keratin-treated. ( n  = 3; mean ± standard deviation (s.d.)).

ORS cells formed colonies within a few hours of exposure to keratin, and subsequently formed strand-like extended structures by day 3 (Fig.  3a, b ). The proliferation of ORS cells was suppressed upon exposure to keratin, as indicated by decreased BrdU incorporation (Supplementary Fig.  14 ). High β-catenin expression, known to occur during migration, further differentiation of stem cells in the ORS region 6 , and a local cell population expressing P-cadherin, known to be a marker of secondary HG formation 20 , 21 , were observed along with extended structures in keratin-treated ORS cells (Fig.  3c , Supplementary Fig.  15 ). In addition, keratin-treated ORS cells showed lower expression levels of CD34 than untreated ORS cells but maintained high levels of SOX9 expression (Fig.  3c ). With reduced expression of CD34, the Lgr5+ cell population, which is known to participate in hair germ formation 22 , 23 , 24 , emerged in keratin-treated ORS cells (Supplementary Fig.  16 ). In addition, real-time qPCR analysis showed upregulation of the expression of various genes, such as SOX9, EDAR, FOXN1 25 , MSX2 26 , SHH 27 , EFNB1 , ITGA6 28 and β -catenin , indicating matrix and shaft differentiation of stem cells from the ORS bulge region (Supplementary Fig.  17 ). Furthermore, RNA sequencing analysis of keratin-treated ORS cells revealed upregulation of mRNA expression levels of acidic hair keratins, mainly KRT31, KRT33B, KRT34 , and KRT37 (Supplementary Fig.  18a ). We also observed increased expression of KRT34 and β-catenin proteins (Supplementary Fig.  18b ). These findings imply that hair keratin-mediated alterations in the protein and gene expression profiles indicate germ formation and further differentiation of ORS cells.

figure 3

a Time-lapse images of ORS cells in the presence of keratin; black arrows indicate colony formation. Scale bars, 100 μm. b Image of ORS cells and quantification of ORS cell growth in the presence of keratin. * P ,0.01, indicates a difference between control and keratin-treated. Scale bars, 100 μm. ( n  = 6; mean ± standard deviation (s.d.)); ORS outer root sheath. c In vitro germ formation of ORS cell by immunofluorescent staining in the presence of keratin; DAPI, blue; phalloidin, integrin β1, red; P-cadherin, CD34, β-catenin, SOX9, green. Control confluent, ORS cell culture at confluent cell density under ORS culture medium; keratin-treated confluent, ORS cell culture at confluent cell density in the presence of keratin. Scale bars, 100 μm.

TGFβ2 induces apoptosis of ORS cells, keratin release and deposition, mediating condensation of DP cells

The findings described above demonstrated that the exposure to keratin induced the condensation of DP cells and the formation of P-cadherin-expressing germs of ORS cells, which led us to ask whether the observed interaction of DP and ORS cells with keratin might be related to a biological event that occurs during hair cycling. During the anagen-catagen transition stage, TGFβ2 is synthesized by DP cells stimulated by dihydrotestosterone and is spatiotemporally localized in the lower part of the hair bulb at the catagen stage, thus suppressing the proliferation of epithelial cells but inducing caspase-mediated apoptosis 11 . Therefore, we hypothesized that exposure to keratins derived from apoptotic ORS cells during hair cycling might drive DP cell condensation and secondary HG formation through interaction with DP and ORS cells. To address this question, we induced apoptosis of ORS cells by treatment with TGFβ2 and characterized microenvironmental changes, such as the release or deposition of keratin. Apoptosis array analysis showed the upregulation of apoptosis-related markers, including Bax, caspase-3, cytochrome C, and SMAC, in ORS cells treated with TGFβ2 (Fig.  4a , Supplementary Fig.  19 ). Extended structures composed of spindle-shaped ORS cells developed only in the presence of TGFβ2; annexin V-and TUNEL-positive apoptotic cells were mainly found in the extended structures of TGFβ2-treated ORS cells, and elevated expression levels of caspase-3 and massive deposition of keratin were observed along the extended structures (Fig.  4b ), which was confirmed by western blot analysis (Supplementary Fig.  20 ). To determine whether the released or deposited keratin derived from TGFβ2-induced apoptotic ORS cells could influence DP cell condensation, the condensation activity of DP cells was tested by direct contact co-culture and culture in conditioned medium. Local condensation of DP cells with the formation of spherical cell colonies was observed in the concentric region of the extended structures in the TGFβ2-treated ORS cell layers (Fig.  4c , Supplementary Fig.  21a ). The conditioned medium collected from TGFβ2-treated ORS cell cultures contained relatively higher levels of keratin, and DP cell condensation into spherical cell aggregates was distinctly increased following the culture of DP cells in the conditioned media (Fig.  4d , Supplementary Fig.  21b ). These results indicate that the deposition or release of keratin from TGFβ2-induced apoptotic ORS cells could regulate the induction of DP cell condensation.

figure 4

a Graphical quantification of apoptosis array of ORS cells and TGFβ2-treated ORS cells. b TGFβ2-induced apoptosis and its following keratin exposure of ORS cells by immunofluorescent staining; DAPI, blue; phalloidin, keratin 34, red; annexin V, keratin 34, TUNEL, caspase 3, green. Scale bars, 200 μm. c Images of DP cell condensation on TGFβ2-treated ORS cell layers. Co-culture of cell tracker-treated DP cells (red) on TGFβ2-treated ORS cell layers. Immunofluorescent image; E-cadherin, green; DAPI, blue. Scale bars, 200 μm. d DP cell condensation under conditioned medium collected from TGFβ2-treated ORS cell culture. Western blot image of released keratin 34 from ORS cell culture and TGFβ2-treated ORS cell culture. Immunofluorescent image; ALPase, red; β-catenin, green; DAPI, blue; Control-DP medium, DP culture medium; Control-ORS Medium-TGFβ2, ORS medium including TGFβ2; CM from TGFβ2-treated ORS, conditioned medium collected from TGFβ2-treated ORS cell culture; Keratin treatment, DP medium containing 1(w/v)% keratin. Scale bars, 50 μm.

Keratin release and deposition through caspase-6-mediated keratin degradation stimulates condensation of DP cells

Keratin fragmentation occurs during apoptosis of epithelial cells 29 ; intracellular insoluble keratin is fragmented by caspases during apoptosis and released as soluble fragments 30 . In our study, apoptosis array analysis showed a two-fold increase in caspase-3 expression levels in TGFβ2-treated ORS cells, and another study reported that type I keratin, including hair keratin, contains a cleavage site, VEVD, for caspase-6 31 . When hair keratin was digested with caspase-3 and caspase-6, keratin fragments were generated only in hair keratin digested by caspase-6 (Fig.  5a ). Furthermore, higher gene and protein levels of caspase-6 and its cleaved form (active caspase-6) were observed in TGFβ2-treated ORS cells (Fig.  5b, c ). These findings imply that the release or deposition of keratin derived from TGFβ2-treated ORS cells through caspase-6-mediated proteolysis can influence DP cell condensation. To test this, we silenced the expression of caspase-6 in ORS cells by siRNA transfection and examined the levels of released keratin (Fig.  5d ). We observed lower levels of released keratin in caspase-6-knockdown ORS cells in the presence of TGFβ2 (Fig.  5e ), and lower condensation activity of DP cells cultured in conditioned media collected from caspase-6-knockdown ORS cell cultures or co-cultured on the caspase-6-knockdown ORS cell layer in the presence of TGFβ2 (Fig.  5f–h ). Caspase-6-knockdown ORS cells developed relatively spread structures even in the presence of TGFβ2, which were different from the extended structures of TGFβ2-treated non-knockdown ORS cells. TUNEL-positive apoptotic cells were found in both the spread and extended structures, and keratin deposition was influenced by caspase-6 expression levels (Fig.  5h , Supplementary Fig.  22 ). We then immunodepleted keratin from the conditioned medium of TGFβ2-treated ORS cell cultures using a column containing anti-human type I + II hair keratin antibody-conjugated beads assay and examined its effect on the condensation of DP cells. The elimination of keratins in the conditioned media was confirmed (Fig.  6a ), and there was no substantial difference in the levels of growth factors contained in the conditioned media before and after immunodepletion (Fig.  6b , Supplementary Fig.  23 ). The increase in TGFβ2 in the conditioned medium was derived from the exogenous TGFβ2 added to induce apoptosis of ORS cells, and TGFβ2 did not influence DP cell condensation by itself (Supplementary Fig.  24 ). The immunodepletion assay showed that the removal of keratins suppressed DP cell condensation (Fig.  6c ), which reflects the functional role of keratins in DP condensation.

figure 5

a SDS-PAGE images of caspase 3- and caspase 6-mediated keratin degradation. b Caspase 6 mRNA expression in TGFβ2-treated ORS cells by real-time qPCR. P ,0.01, indicates a difference between ORS cells (control) and TGFβ2-treated ORS cells (TGFβ2). n  = 4; mean ± standard deviation (s.d.)). c Western blot images of caspase 6 expression in TGFβ2-treated ORS cells. d Caspase 6 mRNA expression in ORS cell culture in the presence of TGFβ2 and caspase 6-knockdown ORS cell culture in the presence of TGFβ2 by real-time-qPCR; Control, ORS cells; TGFβ2, ORS cell culture in the presence of TGFβ2; TGFβ2+siRNA, caspase 6-knockdown ORS cell culture in the presence of TGFβ2. * P ,0.01, indicates a difference between control and experimental group. # P ,0.01, indicates a difference between ORS cell culture in the presence of TGFβ2 and caspase 6-knockdown ORS cell culture in the presence of TGFβ2. n  = 5; mean ± standard deviation (s.d.)). e Western blot image of keratin in conditioned medium collected ORS cell culture in the presence of TGFβ2 and caspase 6-knockdown ORS cell culture in the presence of TGFβ2. f DP cell condensation activity in conditioned medium collected from caspase 6-knockdown ORS cell culture in the presence of TGFβ2; DP medium, basic DP medium; ORS Medium; basic ORS medium; ORS CM, conditioned medium collected from ORS cell culture; ORS CM + TGFβ2, conditioned medium collected from ORS cell culture in the presence of TGFβ2, ORS CM + TGFβ2+siRNA, conditioned medium collected from caspase 6-knockdown ORS cell culture in the presence of TGFβ2. * P ,0.01, indicates a difference between DP cell culture in DP medium and DP cell culture in other culture media and conditioned media. # P ,0.01, indicates a difference between DP cell culture in conditioned medium collected from ORS cell culture in the presence of TGFβ2 and DP cell culture in conditioned medium collected from caspase 6-knockdown ORS cell culture in the presence of TGFβ2. n  = 4; mean ± standard deviation (s.d.)). g Images of DP cell condensation in conditioned medium collected from caspase 6-knockdown ORS cell culture in the presence of TGFβ2, and DP cell condensation by immunofluorescent staining; phalloidin, red; β-catenin, green; DAPI, blue; Scale bars, 100 μm. h Images of DP cell condensation on caspase 6-knockdown ORS cell layer in the presence of TGFβ2. Co-culture of cell tracker-treated DP cells (red) with TGFβ2-treated ORS cell layers. Scale bars, 200 μm. Immunofluorescent image of caspase 6-knockdown ORS cells in the presence of TGFβ2: Phalloidin, E-cadherin, red; KRT34, TUNEL, caspase 6, green; DAPI, blue; Scale bars, 100 μm.

figure 6

a Western blot image of the keratin-eliminated conditioned medium and the keratin-bound beads. b Graphical quantification of growth factor content of the conditioned medium and the keratin-eliminated conditioned medium collected from TGFβ2-treated ORS cell culture using antibody array for growth factors. c DP cell condensation activity in keratin-eliminated conditioned medium of TGFβ2-treated ORS cell culture using a column containing anti-human type I + II hair keratin antibody-conjugated beads. Immunofluorescent image; phalloidin, red; β-catenin, green; DAPI, blue; DP media, DP culture medium; TGFβ2-ORS CM, conditioned medium collected from TGFβ2-treated ORS cell culture; TGFβ2-ORS CM (IgG-column), conditioned medium collected from TGFβ2-treated ORS cell culture and then treated with normal IgG-conjugated beads; 1% Keratin, DP medium containing 1(w/v)% keratin; TGFβ2-ORS CM (keratin Ab-column), keratin-eliminated conditioned medium collected from TGFβ2-treated ORS cell culture. Graphical representation of DP cell condensation activity. * P ,0.05 and ** P ,0.01, indicate a difference between DP media and experimental groups. # P ,0.01, indicates a difference between TGFβ2-ORS CM (IgG-column) and TGFβ2-ORS CM (keratin Ab-column). Scale bars, 100 μm. ( n  = 4; mean ± standard deviation (s.d.)). d Germ formation of ORS cells and KRT31/KRT34 knockdown ORS cells. Western blot image of keratin content of conditioned media and cell lysates from negative control siRNA-transfected ORS cell culture in the presence of TGFβ2 and KRT31/KRT34 knockdown ORS cells culture in the presence of TGFβ2. Image of ORS cells and KRT31/KRT34 knockdown ORS cells in the presence of TGFβ2. Germ formation of ORS cells and KRT31/KRT34 knockdown ORS cells in the presence of TGFβ2 by immunofluorescent staining; phalloidin, red; P-cadherin, RUNX1, KRT34, green; DAPI, blue. Scale bars, 200 μm.

Spatial deposition of keratin derived from TGFβ2-induced apoptotic ORS cells induces germ formation

In addition to keratin-mediated condensation of DP cells, the ability of keratin released from TGFβ2-induced ORS cells to induce germ formation was tested Contrary to DP cell condensation, the keratin released in conditioned media was not effective in generating aP-cadherin-expressing cell population, which was confirmed by an immunodepletion assay (Supplementary Fig.  25 ). This result prompted us to ask whether secondary HG formation by cells expressing P-cadherin is influenced by spatially deposited keratin, which is caused by the spatiotemporal apoptosis of ORS cells. The expression of TGFβ2, which is restricted to the outermost ORS cell layer in the anagen phase, has been reported to be upregulated spatiotemporally in the boundary region between germinal matrix cells and DP cells in the lower bulb region during late anagen and catagen 8 . Therefore, to study the spatial deposition of keratin derived from TGFβ2-induced apoptotic ORS cells and its effect on germ formation by ORS cells, the time-course effect of TGFβ2 treatment on expression levels of caspase-6 protein, keratin deposition, and germ formation was characterized. The extended structures were progressively developed in the TGFβ2-treated ORS cell layers over the cultivation time, and theP-cadherin-expressing germ was spatially developed in the TGFβ2-treated ORS cell layers (Supplementary Fig.  26 ). In addition, immunocytochemical staining of the TGFβ2-treated ORS cell layers showed that a population of RUNX1-and P-cadherin-positive cells, representative markers of germ formation 10 , 32 , 33 , 34 , emerged in the concentric region of the extended structures, and the caspase-6-expressing apoptotic cell population and the keratin-deposited area also increased over time in the extended structures (Supplementary Fig.  26a ), and such highly intensified staining was confirmed by flow cytometric analysis (Supplementary Fig.  26b ). Next, to consider the effect of spatial deposition of keratin on the formation of P-cadherin-expressing germs in vitro, the expression of KRT31/KRT34 in ORS cells was downregulated by siRNA transfection. Downregulation of the expression of keratin in both conditioned media and ORS cells after KRT31/KRT34 downregulation was confirmed (Fig.  6d ). In contrast to the well-developed stranded structures in the ORS cell layers, the KRT31/KRT34- knockdown ORS cells did not form extended structures even in the presence of TGFβ2, and keratin deposition and the emergence of RUNX1-and P-cadherin-expressing ORS cell populations were markedly suppressed in the KRT31/KRT34-knockdown ORS cell culture in the presence of TGFβ2 (Fig.  6d ). KRT31/KRT34 downregulation did not influence cell growth or the generation of Lgr5+ and P-cadherin+ cell populations in ORS cells (Supplementary Figs.  27 and 28 ).

In vivo knockdown of keratin expression suppresses anagen hair follicle formation and hair growth

The in vitro data from studies on the interaction of keratin with DP and ORS cells and the release and deposition of keratin from TGFβ2-induced apoptotic ORS cells have shown a pivotal role of keratin in controlling DP condensation and HG formation. Finally, to determine whether downregulation of KRT31/KRT34 expression can suppress hair follicle formation and hair growth in vivo, keratin expression in mice was temporarily downregulated by intravascular lipofectamine-mediated delivery of KRT31/KRT34 siRNAs. RT-PCR analysis showed effective KRT31/KRT34 downregulation of KRT31/KRT34 mRNA expression on day 7 (Fig.  7a ). Furthermore, it was found that KRT31/KRT34 downregulation inhibited hair growth compared to the control (Fig.  7b ). Notably, dysregulation of hair follicle cycling was observed following downregulation of KRT31/KRT34 in mice; and histological analysis of hair follicle sections showed a strong suppression of the formation of anagen follicles, with no appearance of anagen follicles in 56% of skin tissue sections following KRT31/KRT34 downregulation in mice on day 7 (Fig.  7c ). In addition, the formation of catagen follicles was relatively reduced on day 7 upon downregulation of KRT31/KRT34 in mice with or without exogenous keratin injection (Fig.  7c ). An anagen bulb containing a population of cells expressing P-cadherin was hardly seen in immunohistological sections of KRT31/KRT34 knockdown mice (Fig.  7d , Supplementary Fig.  29 ). Emergence of Lgr5-positive cell population and molecular expression of KRT34 were also distinctly decreased in KRT31/KRT34 knockdown mice on day 7 (Supplementary Figs.  30 and 31 ). In contrast, relatively higher Lgr5 -positive staining was observed in exogenous keratin-injected KRT31/KRT34 knockdown mice on day 7 in comparison with KRT31/KRT34 knockdown mice (Supplementary Fig.  30 ), and an additional injection of hair-derived keratin after KRT31/KRT34 siRNA transfection allowed the hair follicles to enter the anagen phase and regrow hair, similar to the controls. No obvious histological differences were found in the formation of hair follicles and hair growth between the control skin and keratin-injected skin of KRT31/KRT34 knockdown mice after 2 weeks (Fig.  7e ). Furthermore, the formation of P-cadherin-positive germs and strong expression of β-catenin were observed in the region of anagen hair follicles in sections of control skin and keratin-injected skin of KRT31/KRT34 knockdown mice (Fig.  7d , Supplementary Fig.  32 ), and strong staining for KRT34 was found in the ORS region surrounding the DP, which corresponds to the caspase-6-positive region (Supplementary Fig.  32 ). Interestingly, it was found that the region stained positive for caspase-6, KRT34, and P-cadherin moved upward into the hair shaft region of the expanded hair follicles (Supplementary Fig.  32 ).

figure 7

a Graphical representation of KRT31 and KRT34 mRNA expressions in mice on day 7 after KRT31/KRT34 knockdown. Control, mice injected with negative control siRNA-loaded lipofectamine; siRNA, KRT31/KRT34 knockdown mice; siRNA+KRT, KRT31/KRT34 knockdown, and hair keratin-injected mice; * P ,0.05, indicates a difference between control and experimental groups. ( n  = 4, in 4 mice; mean ± standard deviation (s.d.)). b Images of hair growth on the back skin of mice on day 3, day 7, day 10, and day 14 after KRT31/KRT34 siRNA transfection. c Histological images of the back skin of mice on day 7 after KRT31/KRT34 siRNA transfection and intradermal injection of keratin. Graphical representation and quantification of hair follicle formation in skin sections of mice ( n  = 36 sections, in 8 mice; mean ± standard deviation (s.d.)). * P ,0.01, indicates a difference between control group and experimental groups. Scale bars, 200 μm. d Immunohistochemical images of the back skin of mice on day 7 after KRT31/KRT34 siRNA transfection and intradermal injection of keratin; β-catenin, red; P-cadherin, green; DAPI, blue. Scale bars, 20 μm. e Histological images of the back skin of mice on day 14 after KRT31/KRT34 siRNA transfection and intradermal injection of keratin. Graphical representation and quantification of hair follicle formation in skin sections of mice ( n  = 23 sections, in five mice; mean ± standard deviation (s.d.)). * P ,0.01, indicates a difference between control group and experimental groups. Scale bars, 20 μm.

Our study shows that intradermal injection of human hair-derived keratin promotes hair growth with enhanced formation of anagen hair follicles and an increase in the size of hair follicles. Hair growth is controlled by the interactions between two distinct cell types: mesenchyme and epithelial cells, while DP and stem cells from the bulge region of the ORS participate in hair follicle formation 4 , 5 , 6 , 15 . The in vivo injection of keratin-induced follicle formation and hair growth. This effect could be associated with keratin-mediated DP cell condensation and germ formation via its interaction with cells, as evidenced by the strong expression of various signature molecules, such as β-catenin and P-cadherin, which are highly expressed during hair follicle formation 4 , 5 , 6 , 15 , in keratin-treated DP ORS cell cultures. Secondary HG expressing P-cadherin emerges at the telogen stage and leads to the first stage of hair regeneration 32 . The interaction of P-cadherin with β-catenin plays an important role in maintaining the anagen phase of the hair cycle 35 , and β-catenin-expressing cells migrated from the bulge region of the ORS undergo further differentiation in the follicle region 6 . Keratin-treated ORS cells showed morphological changes, such as spindle shape, strong expression of β-catenin, and reduced expression of CD34, also a population of P-cadherin-expressing cells emerged. With the loss of CD34 expression, an Lgr5-expressing cell population was generated in keratin-treated ORS cells. CD34-positive stem cells have been reported to convert directly into P-cadherin-expressing HG cells 33 , and CD34-positive stem cells migrate downward to the lower bulge region and convert to Lgr5-positive stem cells, which participate in HG formation during the telogen stage 24 . In addition to keratin-mediated ORS cell differentiation, treatment with exogenous keratin induces DP cell condensation. Epithelial fibroblast growth factor 20 (Fgf20) is known to control dermal condensate morphogenesis 36 ; hence, we evaluated the presence of potent growth factors, such as FGF20, in hair-derived keratin to influence cellular behavior. Western blot analysis showed no presence of FGF20 in the keratin extract used in this study, and application of MALDI-TOF mass spectroscopy indicated that the extracted protein was purely keratin (Supplementary Fig.  33 ). Along with their differentiation, growth of ORS cells was stopped in the presence of keratin, but CCK-8 analysis, which measures mitochondrial dehydrogenase activity, showed a temporal increase in growth on day 1 of keratin treatment (Supplementary Fig.  14 ). This change in cellular metabolic activity by keratin treatment needs to be studied further. Hair regeneration is processed by hair follicles undergoing repeated cycles of anagen (hair growth stage), catagen (regression stage), and telogen (relative rest stage) 37 . During the telogen phase, secondary HG progressively appears at the base of the follicular epithelium. At that point, HG cells form a cell cluster and are activated to begin hair regeneration 10 , and DP cells undergo condensation to form a follicular papilla beneath the secondary HG. The interaction between the DP condensate and secondary HG leads to the formation of new hair follicles by enveloping the DP with downwardly extended epithelial cells 10 , 32 , 38 . However, despite considerable progress in understanding the cellular interactions that control hair growth, it is not clear how secondary HG formation and DP condensation, the key biological events causing hair regeneration, are initiated at the beginning of a new hair cycle.

Hair growth in keratin-injected mice, keratin-mediated condensation of DP cells, and the formation of P-cadherin expressing germs, assessed by in vitro cell study, led us to explore the biological function of keratin in hair regeneration as a pilot study. At the anagen-catagen transition stages of the hair cycle, the local deposition of TGFβ2 in the lower region of the follicle is restricted due to the spatiotemporal secretion of TGFβ2 produced by DP cells 11 , which is consistent with the spatial gradient of apoptosis of epithelial cells 9 . Spatiotemporally localized TGFβ2 induces apoptosis of ORS cells in the lower part of the hair bulb at the catagen stage, resulting in the expression of caspase and its-mediated fragmentation of insoluble keratin into soluble keratin fragments 8 , 9 , 10 , 11 , 29 . Although the relationship between TGFβ2 expression and the intrinsic property of DP cells related to condensation is not well known, our data showed that TGFβ2 expression was downregulated during DP cell condensation and rapidly upregulated during the dispersion of condensed DP cells (Supplementary Fig.  34a, b ). Mesenchyme condensation such as dermal condensates is promoted by BMP signaling and transient downregulation of TGF-β signaling, showing an antagonistic relationship 18 , 39 , 40 . The condensed DP cells might maintain the property of DP cells 38 to drive hair growth by releasing paracrine factors, which can induce stem cell activation and differentiation during the anagen phase, and may induce spatiotemporal apoptosis of adjacent ORS cells via increased expression of TGFβ2 during the anagen to catagen transition. In our study, it was shown that local DP cell condensation and germ formation in TGFβ2-induced apoptotic ORS cells depend on the exposure of keratin via caspase-6 expression and consequently its-mediated keratin exposure from apoptotic cell death, as evidenced by the suppressed DP cell condensation and germ formation in caspase-6-knockdown or KRT31/KRT34-knockdown ORS cell culture, even if there was no distinct difference in TGFβ2-mediated ORS cell apoptosis. From these in vitro findings, it could be inferred that spatially increased keratin exposure, following gradual apoptosis during the regression stage, might provide a cue to derive germ formation and new hair cycle initiation from telogen to anagen, which spatial keratin deposit could be identified at upper void space unoccupied cells in the newly formed hair follicle (Supplementary Fig.  35 ) In addition, apoptosis-related keratin exposure also could be identified by similar spatial expressions of apoptosis-markers such as Annexin V and active caspase 3 and KRT 34 in the developing hair follicles containing proliferating cells (Supplementary Fig.  36 ).

Finally, to determine the biological function of keratin in vivo, the effect of downregulating KRT31/KRT34 mRNA expression on hair growth in mice was evaluated. Exogenous keratin injection in KRT31/34 knockdown mice resulted in a relatively reduced formation of catagen follicles on day 7. This might be due to the temporal inhibition of keratin expression accompanied by stem cell differentiation into the matrix and shaft during the anagen phase, which might influence catagen formation. In addition, the formation of anagen hair follicles and hair growth were suppressed in mice with temporal downregulation of KRT31/KRT34, which could be recovered by intradermal injection of additional exogenous keratin. With the poor formation of anagen follicles in KRT31/KRT34 knockdown mice, P-cadherin and Lgr5-expressing cell population was scarcely observed in telogen follicles in these mice (Supplementary Figs.  29 and 30 ). In vitro KRT31/34 knockdown ORS cells did not show any change of ability for cell growth and differentiation into P-cadherin and Lgr5-expressing cells (Supplementary Figs.  27 and 28 ), and such P-cadherin and Lgr5-expressing cell population could be formed by injecting exogenous keratin into KRT31/34 knockdown mice even though less than control mice (Supplementary Figs.  29 and 30 ). These findings indicate that alteration of hair keratin gene expression might influence hair growth following stem cell differentiation into the matrix and shaft, and hair cycle transitions might be controlled by keratin-mediated microenvironmental change. However, the reduced anagen follicle formation could not be explained in KRT31/34 knockdown mice model starting at synchronized telogen phase, which hair cycle-dependent keratin expression and apoptosis-related keratin exposure even during telogen to anagen transition need to be studied further. The expression of hair keratins is not restricted to the anagen phase, showing cell growth and differentiation-mediated hair keratin production, and are found at all stages of the hair cycle 41 , 42 , 43 .A distinct epithelial cell population expressing Bcl-2 in secondary HG and DP was found during the telogen-anagen transition, which shows differential susceptibility to apoptosis 44 . Such programmed cell death-related cellular processes were also detected during telogen, and the stimulation of autophagy following programmed cell death initiated the telogen-anagen transition 45 . In addition, A study using transgenic mice overexpressing an anti-apoptotic gene reported that the inhibition of the apoptotic death of ORS cells even during anagen resulted in the early termination of hair follicle stem cell activation and proliferation, whereas the initiation of a new hair cycle was postponed by inhibiting the apoptotic death of ORS cells during telogen 46 . These studies suggested that telogen might not be the only resting phase in hair growth, but also an activating phase, including DP condensation and secondary HG formation 47 , 48 . These studies indicate that spatiotemporal apoptosis during the hair cycle can be an essential process in controlling hair regeneration, and our findings show that keratin can be an important factor influencing hair growth. However, these pilot observations require follow-up studies of hair keratin expression and its apoptosis-related exposure during the hair cycle using genetically modified ORS cells equipped with an on-off expression system for keratin expression and a xenograft mouse model to determine the in vivo mechanisms.

Despite in vitro and in vivo studies of keratin-mediated hair growth, the mechanism by which keratin induces DP condensation and hair germ formation remain unclear. In this study, DP cell condensation was also induced on Matrigel (Supplementary Fig.  9 ), and we found a decrease in the hardness of Matrigel in the presence of keratin. Hence, we tested the keratin-mediated change in the hardness of Matrigel, and keratin treatment resulted in a partial disintegration of Matrigel (Supplementary Fig.  37 ), which might influence cell and matrix interactions. In addition, loss of vinculin, which participates in local cell adhesion, was found in keratin-mediated DP cell condensation (Supplementary Fig.  38 ), and highly decreased expression of vinculin was observed in keratin-treated ORS cells (Supplementary Fig.  39 ). A recent report showed that mechanical instability of cells to ECM contact was a factor in controlling activation of hair follicle stem cells in the bulge region, which was proved by the finding that loss of vinculin allowed hair follicle stem cells to escape quiescence and forced the initiation of a new hair cycle 49 . Keratin exposure from TGFβ2-induced spatiotemporal apoptotic ORS cells might influence the mechanical properties of the microenvironment and cell-to-ECM interactions, which might be a cue to drive DP cell condensation and activation of hair follicle stem cells participating in hair germ formation. However, further studies on keratin-mediated mechanotransduction are required.

Taken together, the results presented in this study reveal that hair regeneration is regulated by keratin-mediated germ formation and DP condensation through biological events including TGFβ2-induced ORS cell apoptosis and the spatial exposure of keratin derived from apoptotic ORS cells (Supplementary Fig.  40 ). Our pilot study indicated that keratin is not only a major structural component of hair, but can also play a functional role in the induction of HG formation and DP condensation, facilitating entry into a new hair cycle. In conclusion, considering the biological function of keratin in hair growth, our study suggests that keratin can be a potent biomaterial for developing therapeutic agents for hair loss treatment. Understanding how cellular behavior is regulated by spatiotemporal keratin exposure from apoptotic epithelial cells can provide additional insight into deciphering cellular interactions between epithelial cells and mesenchyme in the morphogenesis of other tissues.

Experimental design

This study aimed to unravel the biological functions of keratin in hair growth. First, to determine the activity of hair growth, mouse back skin hair was removed, and hair follicle formation and hair regrowth were evaluated after intradermal injection of hair-derived keratin (Fig.  1b–d ). Next, to define the interaction of keratin with major cells participating in hair growth, cellular behaviors such as DP cell condensation and germ formation were studied by treating in vitro DP cells and ORS cell cultures with keratin (Fig.  1e, f , Figs.  2 and  3 ). The results (Figs.  1 – 3 ) led to the hypothesis that keratin-induced hair growth could be closely related to a biological cascade that occurs during the hair cycle with regard to the release of keratin from TGF-β2-induced apoptotic ORS cells at the stages of the anagen-catagen transition. To characterize keratin release from apoptotic ORS cells or deposition, apoptosis of ORS cells and the subsequent keratin release from TGF-β2-treated ORS cells and deposition were evaluated. Direct co-culture of DP cells with TGF-β2-treated ORS cells and DP cell culture in conditioned medium collected from TGF-β2-treated ORS cell culture was done to evaluate the effect of released or deposited keratin on DP cell condensation (Fig.  4 ). Following DP cell condensation induced by released and deposited keratin, apoptosis-related caspase 3 and caspase 6 expression, and caspase-mediated keratin degradation was characterized. The release and deposition of keratin through caspase-mediated keratin degradation in TGF-β2-treated apoptotic ORS cells and its effect on DP cell condensation were evaluated by in vitro siRNA-mediated downregulation of caspase 6 mRNA expression in TGF-β2-treated apoptotic ORS cells (Fig.  5 ). To confirm the effect of keratin released from TGF-β2-treated apoptotic ORS cells on DP cell condensation, keratin was eliminated from the conditioned medium collected from TGF-β2-treated cell culture by immunodepletion, and DP cell condensation was evaluated by culturing DP cells in keratin-depleted conditioned medium (Fig.  6a–c ). To prove the effect of keratin released from TGF-β2-treated apoptotic ORS cells on DP cell condensation, keratin 31 (KRT31) and keratin 34 (KRT34) expression were downregulated in ORS cells by in vitro KRT31/KRT34 siRNA transfection. Then P-cadherin-expressing germ formation by ORS cells was observed in KRT31/KRT34-knockdown ORS cell culture in the presence of TGF-β2 to evaluate the effect of keratin spatially deposited on germ formation (Fig.  6d ). Finally, to study the role of keratin in hair follicle formation and hair growth in vivo, KRT31/KRT34 were downregulated using Invivofectamine KRT31/KRT34 siRNA transfection in mice (Fig.  7 ). The whole experimental procedure is illustrated in Supplementary Fig.  41 .

Cell culture

Human ORS cells (ORS; CEFO, CB-ORS-001: 36 years old female) and human DP cells (DP; CEFO, CB-HDP-001: 54 years old male) were purchased and expanded in each human ORS cell growth medium (CEFO, CB-ORS-GM) and human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 according to the manufacturer’s instructions. Cultures were fed every two days and passaged by treatment with 0.25% trypsin/EDTA (Gibco, 25200056), and the expanded DP cells within 5 passages and ORS cells within 3 passages were used in this study.

Human hair keratin extraction

Human hair keratin was extracted by slightly modified Sindai method 12 , and kindly provided by Gapi Bio. Briefly explaining, human hair was washed using general detergent and delipidized with chloroform (JUNSEI CHEMICAL, 28560-0350):methanol (Merck Millipore, 106009) (2:1, v/v) for 24 hr at room temperature. The delipidized hair was oxidized with 2(w/v)% peracetic acid (Sigma-Aldrich, 269336) for 12 hr at 37 °C. The hair was reacted with Shindai solution (5 M urea (Sigma-Aldrich,U5378), 2.4 M thiourea (Sigma-Aldrich, T7875), 5% 2-mercaptoethanol (Sigma-Aldrich, M6250), 24 mM trizma base (Sigma-Aldrich, T1503), pH 8.5) for 72 hr at 50 °C. After reaction, The mixture was centrifuged at 3500 rpm for 20 min and supernatant was dialyzed (12–14 kDa cut-off, Spectra/Por 4 dialysis membrane, 132706) against deionized water for 5 days with three changes in water a day. Solution of hair keratin was centrifuged at 3500 rpm for 20 min and supernatant was lyophilized by freeze-dryer.

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)

Characterization of human hair keratin was performed by SDS-PAGE analysis.100 μg of human hair keratin was dissolved in 13 μl of DBPS and mixed with 5 μl of LDS (Invitrogen, B0007). and 2 μl of sample reducing agent (Invitrogen, B0009). Each sample was denatured by heating at 75 °C for 10 min before loading into precast Bolt 4 to 12% Bis-Tris Mini Protein Gels (Invitrogen, NW04122BOX). Electrophoresis was carried out at a constant voltage of 200 V for 22 min in 1× MES Running Buffer (Invitrogen, B0002). Subsequently, separated proteins were stained with SimplyBlue SafeStain (Invitrogen, LC6060).

Matrix-assisted laser desorption ionization-time of flight mass spectrometer

Protein spots were enzymatically digested in-gel using modified porcine trypsin (Promega modified). Gel pieces were washed with 50% acetonitrile to remove SDS, salt, and stain. Washed and dehydrated spots were then vacuum dried to remove solvent and rehydrated with trypsin(8–10 ng/μl) solution in 50 mM ammonium bicarbonate pH 8.7 and incubated 8–10 h at 37°C. Samples were analyzed using the BRUKER autoflex maX with LIFTTM ion optics. Both MS and MS/MS data were acquired with a SMARTBEAM LASER with 2 kHz repetition rate, and up to 4000 shots were accumulated for each spectrum. MS/MS mode was operated with 2 keV collision energy; air was used as the collision gas such that nominally single collision conditions were achieved. Although the precursor selection has a possible resolution of 200, in these studies of known single-component analytes a resolution of 100 was utilized. Both MS and MS/MS data were acquired using the instrument default calibration, without applying internal or external calibration. MS/MS ions searches were performed with the license Mascot for in-house use.

Human hair keratin-mediated hair growth test in mice

In vivo mouse studies for keratin-mediated hair growth were done separately at Chemon Inc and Konkuk University.

For in vivo studies, six-week-old C57BL/6 male mice were purchased from Orientbio Inc. (Gyeonggi-do, Korea). This study was performed within the animal facility area of Chemon Inc in Gyeonggi bio center, and the animals were housed in a room that was maintained at a temperature of 23 ± 3 °C and a relative humidity of 55 ± 15%, with artificial lighting from 08:00 to 20:00, 150–300 Lux of luminous intensity. Throughout the experimental period, the temperature and humidity of animal room were measured every hour with a computer-based automatic sensor, and as a result of measurements, there were no deviations to have adverse effect to the result of study. Animals were offered irradiation-sterilized pellet food for lab animal (Teklad certified irradiated global 18% protein rodent diet; 2918 C, Envigo, UK). Underground water disinfected by ultraviolet sterilizer and ultrafiltration was given via water bottle, ad libitum . Examination of water was performed by an authorized Gyeonggido Institute of Health & Environment (Suwon, Gyeonggi-do, Republic of Korea), and there were no factors that could affect results. All animal experiments were performed according to Chemon Inc’s standard operating procedures. The study protocol was approved by the Institutional Animal Care and Use Committee of Chemon Inc, which is accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International (Approval No.: 2018-08-011). To examine the hair growth promoting effect of keratin according to keratin concentration 0.5(w/v)% and 1.0 (w/v)% keratin in phosphate-buffered saline (PBS; Gibco, 10010023) was used. Animals were anesthetized by intraperitoneally administration of zoletyl and rumpoon mixture (4:1, v/v) at the dose of 1 mL/kg. The dorsal area was shaved with an animal clipper. Upon shaving the mice all of the hair follicles were synchronized in the telogen stage, showing pink color. The wound-free animals were selected and weighed. The selected animals were distributed in a randomized manner so that the average weight of each group was distributed as uniformly as possible according to the weighted weight. All animals were randomized into 5 groups based on different topical applications: normal control, DPBS administration, 0.5% keratin administration, 1% keratin administration, and 3% minoxidil as a positive control. The test substance was administered intradermally, the clinically planned route, and the positive control substance was applied tropically. The keratin groups were administered once at day 1, and the positive control substance was administered once/day, 5 times/week, and 4 weeks. The intradermal administration was divided into 2 sites in the dorsal part of animals inhaled with isoflurane using a 0.3 mL insulin syringe (31 G), and divided into 75 μl at 1 site. Topical administration was applied to the back of the animal, which was corrected for 0.15 mL using a 1 mL syringe, rubbed 10 times with a glass rod, and applied evenly. The mice were sacrificed after 24 weeks.

For another in vivo studies, male C57BL/6 mice were used, which were purchased from YoungBio (Samtako, 1404957265). The mice were housed under controlled condition at a temperature of 23 ± 2 °C, humidity of 50 ± 5%, and light-dark cycle of 12 h. Mice were provided with a laboratory diet and water ad libitum. All animal experiments were approved by the Institutional Animal Care and Use Committee of Konkuk University (KU18159, KU19066), and procedures on animals were performed in accordance with the relevant guidelines and regulations. The hair on the dorsal skin of mice was shaved repeatedly using an electric clipper to synchronize the hair follicle cycle. Before treatment, the dorsal hair was completely removed using the commercial hair removal cream Veet® (Reckitt Benckiser, 62200809951). To examine the hair growth-promoting effect of keratin, 1.0 (w/v)% keratin in phosphate-buffered saline (PBS; Gibco, 10010023) was used. 10-week old mice were shaved repeatedly to synchronize the hair cycle and randomly assigned to three groups: Neg. Con group; 3% Minoxidil group with daily topical application of 100 μl of 3% Minoxidil (Minoxyl® 3%; Hyundai Pharm, Co., Seoul, Korea); 1.0 (w/v)% Keratin group with intradermal injection of 100 μl of 1.0 (w/v)% keratin once. The mice were sacrificed after 2 weeks.

In vivo dispersion test of fluorescent dye-conjugated keratin

Human hair keratin was labeled with the fluorescent probe Alexa 488 (Alexa Fluor™ 488 Protein Labeling Kit, Invitrogen, A10235), using a procedure provided by the manufacturer. Briefly, human hair keratin was dissolved in DPBS and sodium bicarbonate (Component B) was added. Subsequently, the keratin solution was reacted at reactive dye (Component A) and room temperature for 1 h. Labeled human hair keratin was purified using purification columns, and injected intradermally into SCID nude mice. In vivo fluorescence signal was observed by the in vivo optical imaging system (Perkin Elmer, IVIS Spectrum), and the fluorescence signal was observed and imaged up to 14 days.

Interaction assay of DP cells with keratin

DP cells were seeded at a density of 2 × 10 4 cell/cm 2 on 12 well and six-well non-treated tissue culture plate (SPL LIFE SCIENCES, 32012, 32006). The DP cells were adjusted to be stable for 1 day in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 prior to keratin treatment. After 1 day of adjustment, DP cells were cultured in human DP growth medium containing 1.0(w/v)% keratin or not. The morphological change of DP cells in the presence of keratin was observed under inverted fluorescent microscopy (Olympus IX71), and the number of condensed DP cell aggregates was counted. Cell proliferation upon keratin treatment was measured using Cell Counting Kit-8 (Dojindo Molecular Technologies, CK04-20). DP cells were seeded on 12 well non-treated tissue culture plate (SPL LIFE SCIENCES, 32012) at a seeding density of 1 × 10 4 cells/cm 2 , and cultured in human DP growth medium (CEFO, CB-HDP-GM) containing 1.0(w/v)% keratin or not in a humidified atmosphere of 5% CO 2 at 37 °C, and the medium was refreshed every two days. At specific time points (1, 3, and 5 days), each well had 10 μl of the Cell Counting Kit-8 solution added and then was incubated at 37 °C for 2 h. Cell proliferation assays were performed in a 96-well plate reader by measuring the absorbance at a wavelength of 450 nm. In a parallel study, BrdU incorporation assay was done by BrdU Cell Proliferation ELISA Kit (Abcam, ab126556). The analysis was performed in accordance with the manufacturer’s instructions. In addition, LIVE/DEAD™ visibility/cytotoxicity kit (Invitrogen, L3224) was used for the LIVE/DEAD assay. Before the LIVE/DEAD assay, 5 μl of calcein AM (Component A) and 20 μl of EthD-1 (Component B) were added to 10 ml of DPBS to prepare a LIVE/DEAD assay working solution. Cells were washed by DPBS, and LIVE/DEAD assay working solution was added. After incubation at room temperature for 10 min, the stained cells were observed, and images were taken using inverted fluorescent microscopy (Olympus IX71).

For DP cell condensation assay according to different cell seeding density, DP cells were seeded at a seeding density of 5 × 10 3 cell/cm 2 , 1 × 10 4 cell/cm 2 and 2 × 10 4 cell/cm 2 on 6 well non-treated tissue culture plate (SPL LIFE SCIENCES, 32006). The DP cells were adjusted to be stable for 1 day in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 prior to keratin treatment. After 1 day of adjustment, DP cells were cultured in human DP growth medium containing 1.0 (w/v)% keratin or not. The number of condensed DP cell aggregates was counted using inverted fluorescent microscopy (Olympus IX71).

Interaction assay of DP cells with keratin on matrigel

For DP cell condensation assay on matrigel, the matrigel was diluted at 1:2 volume ratio in ice-cold serum-free DMEM media (CORNING, 10-013-CV), and gelation was done by incubating at 37 °C for 30 min. DP cells were seeded at a seeding density of 2 × 10 4 cell/cm 2 on top of matrigel, and the DP cells on matrigel were adjusted to be stable for 1 day in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 prior to keratin treatment. After 1 day of adjustment, DP cells were cultured in human DP growth medium containing 1.0(w/v)% keratin or not. Cell viability of DP cells on matrigel was visualized using two-color fluorescence cell viability assay, EthD-1/ Calcein AM live/dead assay (Invitrogen, L3224), which was done according to the manufacturer’s instruction, and the morphological change and live/dead stage of DP cells on matrigel in the presence of keratin was observed under inverted fluorescent microscopy (Olympus IX71).

Indirect analysis of physical property of matrigel

1 ml of Matrigel (Growth factor reduced basement membrane matrix, Corning, CLS356231) was mixed to contain 100,000,000 FluoSpheres (Invitrogen, F13080). 500 μl of Matrigel mixed with FluoSpheres was added to the insert of the transwell (Corning, CLS3460) and incubated at 37 °C for >1 h. Matrigel reacted to various conditions in the presence or absence of cells or human hair keratin. FluoSpheres emitted by changes in the physical properties of Matrigel were measured by fluorescence density by spectrofluorometer (Biotec Ex 430/ Em 465).

Interaction assay of ORS cells with keratin

ORS cells were seeded at 2 × 10 4 cell/cm 2 on 12 well and six-well tissue culture plate (SPL LIFE SCIENCES, 30012, 30006). The ORS cells were adjusted to be stable for 1 day in human ORS cell growth medium (CEFO, CB-ORS-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 prior to keratin treatment. After 1 day of adjustment, ORS cells were cultured in human DP growth medium containing 1.0(w/v)% keratin or not. The morphological change of ORS cells in the presence of keratin was observed under inverted fluorescent microscopy (Olympus IX71) and time-lapse images were captured. Cell proliferation upon keratin treatment was measured using Cell Counting Kit-8 (Dojindo Molecular Technologies, CK04-20). ORS cells were seeded on 12 well tissue culture plate (SPL LIFE SCIENCES, 30012) at a seeding density of 1 × 10 4 cells/cm 2 , and cultured in human ORS cell growth medium (CEFO, CB-ORS-GM) containing 1.0(w/v)% keratin or not in a humidified atmosphere of 5% CO 2 at 37 °C, and the medium was refreshed every two days. At specific time points (1, 3, and 5 days), each well had 10 μl of the Cell Counting Kit-8 solution added and then was incubated at 37 °C for 2 h. Cell proliferation assays were performed in a 96-well plate reader by measuring the absorbance at a wavelength of 450 nm. In a parallel study, BrdU staining was done by immunocytochemical staining with mouse anti-BrdU antibody (Santa Cruz Biotechnology, sc-32323, diluted 1:100). In addition, BrdU incorporation assay was done by BrdU Cell Proliferation ELISA Kit (Abcam, ab126556) according to the manufacturer’s instructions. The final concentration of BrdU in the cell culture medium was 10 µM, and BrdU was added before 24 h of assay. DNA hydrolysis was additionally performed by 1 M HCl.

RNA extraction and sequencing

To perform transcriptome sequencing (RNA-Seq) analysis of DP cells and ORS cells, total RNA was extracted from the ORS and DP cells in the absences of keratin and in the presence of keratin. Quality and integrity of the extracted total RNA was assessed by BioAnalyzer and the standard illumina sequencing system protocol (TruSeq Stranded mRNA LT Sample Prep Kit) have been used to make libraries for RNA-Seq. Around 300 bp fragments were isolated using gel electrophoresis, amplified by PCR and sequenced on the Illumina HiSeq 2500 in the paired-end sequencing mode (2 × 10 1  bp reads).

RNA-seq read processing and differential gene expression analysis

Quality of the raw sequencing reads were assessed, and qualified raw sequencing reads were aligned to the human genome reference hg19 using TopHat alignment tool (v2.1.0) [PMID: 23618408]. Uniquely and properly mapped read pairs have been used for further analysis. Gene annotation information was downloaded from Ensembl (release 75) biomart ( http://www.ensembl.org/ ). To evaluate expression levels of genes, the RPKM (reads per kilobase of exon per million mapped reads) measurement unit was used [PMID: 18516045] and the fold change between two samples (untreated and treated with keratin) was calculated based on the calculated RPKM. DESeq2 R package [PMID: 20979621] was used to identify differentially expressed genes between undifferentiated neural stem cell and the differentiated dopaminergic neuron. Differentially expressed genes were defined as those with changes of at least twofold between samples at a false discovery rate (FDR) of 5%.

DP cell spheroid formation and maintenance assay of the replated DP cell spheroids

For DP cell spheroid formation, cell spheroids as a micro tissue unit were generated by docking DP cells into polyethylene glycol (PEG) microwell array with 450 µm in diameter. PEG microwells were fabricated by microfabrication procedures, reported previously 50 . Micropatterns with 450 µm in diameter were generated on a silicon wafer using SU-8 photoresist (MicroChem Corp, USA). Poly(dimethylsiloxane) (PDMS) molds were generated by mixing silicone elastomer base solution and curing agent (Sylgard 184, Essex Chemical, USA) in a 10:1 ratio and pouring on the patterned silicon master. The generated PDMS stamps were used to mold PEG microwells. Hydrogel microwells were fabricated using micromolding of poly(ethylene) glycol (PEG)-diacrylate (1000 Da) mixed with 1% (w/w) of the photoinitiator Irgacure 2959 (Ciba Specialty Chemicals Corp., Tarrytown, NY). Glass substrates were treated with 3-(Trimethoxysilyl) propylmethacrylate (TMSPMA) (Sigma, USA) for 30 min and baked at 70 °C overnight. A microfabricated PDMS stamp was placed on the PEG monomer solution on a treated glass slide. The monomers were crosslinked by exposing to UV light (350–500 nm wavelength, 100 mW/cm 2 ) for 27 sec. After peeling the PDMS stamp from the substrate, the microwell structures were washed with ethanol and 1× DPBS overnight before using them to culture cells. In all, 200 μL of cell suspension (1 × 10 6 cells per mL) was spread on PEG microwells mounted on a glass slide, and undocked cells were removed by gentle washing with PBS after 30 mins of incubation. DP cell spheroid was formed within PEG microwell by incubating the cell-docked microwell in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 for 1 day. The formed DP cell spheroids were retrieved from PEG microwell by gentle agitation, and then replated on 12 well and six-well tissue culture plate (SPL LIFE SCIENCES, 30012, 30006). The replated DP cell spheroids were adjusted to adhere on tissue culture plate for 1 day in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 prior to keratin treatment. After 1 day of adjustment, DP cells were cultured in human DP growth medium containing 1.0(w/v)% keratin or not. The morphological change of DP cell spheroids in the presence of keratin was observed under inverted fluorescent microscopy (Olympus IX71) and CLSM (Confocal Laser Scanning Microscopy, Nikon, D-ECLIPSE C1).

TGFβ2-mediated ORS cell apoptosis and co-culture with DP cells

ORS cells were seeded at 2 × 10 5 cell/cm 2 on 12 well tissue culture plate (SPL LIFE SCIENCES, 30012) to make confluent ORS cell layer. The ORS cells were adjusted to be stable for 1 day in human ORS cell growth medium (CEFO, CB-ORS-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 day of adjustment, ORS cells were cultured in human DP growth medium containing 100 ng/ml TGFβ2 (PeproTech, 100-35B) for 5 days, and the media was refreshed every day.

To evaluate DP cell condensation in direct co-culture of DP cells and TGFβ2-treated ORS cells, DP cells were stained with a cell tracker (Red CMTPX, Invitrogen, C34552) according to manufacturer’s instruction. The stained DP cells were seeded at a density of 2 × 10 4 cell/cm 2 on confluent TGFβ2-treated ORS cell layer which cultured for 5 days prior to co-culture, and cultured in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 and 2 days of co-culture, DP cell condensation was observed under inverted fluorescent microscopy (Olympus IX71).

To evaluate DP cell condensation under conditioned media from TGFβ2-treated ORS cell layer, the conditioned media were collected from TGFβ2-treated ORS cell layer after 5 days of culture. DP cells were seeded at a density of 2 × 10 4 cell/cm 2 on 12 well and 6 well non-treated tissue culture plate (SPL LIFE SCIENCES, 32012, 32006). The DP cells were adjusted to be stable for 1 day in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 day of adjustment, DP cells were cultured under various culture media; human DP growth medium, human ORS cell growth medium containing 100 ng/ml TGFβ2, conditioned medium collected from TGFβ2-treated ORS cell layer and human DP growth medium containing 1(w/v)% keratin. After 1 and 2 days of culture, DP cell condensation was observed under inverted fluorescent microscopy (Olympus IX71).

Immunodepletion study

To study the role of keratin released from TGFβ2-induced apoptotic ORS cells in DP condensation and germ formation of ORS cells, the released keratin in conditioned media from TGFβ2-treated ORS cell layer culture was removed by immunodepletion method. First, antibodies-conjugated beads were prepared as follows; 150μl of nProtein A Sepharose (GE Healthcare, 17528001) was incubated with 400μl of guinea pig anti-Type I + II Hair Keratins antibody (PROGEN, GP-panHK) or guinea pig normal IgG (Sigma-Aldrich, I4756), as another negative control, for 18 h at 4 °C. Non-specific binding was prevented with blocking buffer containing 1% bovine serum albumin (BSA; Sigma-Aldrich, A9418) in TBS (Tris-Buffered Saline; Biosesang, TR2005-000-74) with 0.1% Tween 20 (Duchefa Biochemie, P1362.1000) for 3 h at 4 °C. The conditioned media were collected from TGFβ2-treated ORS cell layer cultured for 5 days, and 30 ml of the conditioned media were mixed with 75μl antibodies-conjugated beads, and then incubated with gentle shaking overnight at 4 °C. After incubation, antibodies-conjugated beads were removed by passing the mixture through a Centrifuge Columns (Thermo Scientific, 89898).

To evaluate DP cell condensation under keratin-removed conditioned media, DP cells were seeded at a density of 2 × 10 4 cell/cm 2 on 12 well and six-well non-treated tissue culture plate (SPL LIFE SCIENCES, 32012, 32006). The DP cells were adjusted to be stable for 1 day in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 day of adjustment, DP cells were cultured under various culture media; human DP growth medium, conditioned medium collected from TGFβ2-treated ORS cell layer, conditioned medium collected from TGFβ2-treated ORS cell layer and incubated with guinea pig normal IgG-conjugated beads, conditioned medium collected from TGFβ2-treated ORS cell layer and incubated with guinea pig anti-Type I + II Hair Keratins antibody-conjugated beads, and human DP growth medium containing 1(w/v)% keratin. After 1 and 2 days of culture, DP cell condensation was observed, and the number of condensed DP cell aggregates was counted using inverted fluorescent microscopy (Olympus IX71).

To evaluate P-cadherin expressing germ formation of ORS cells under keratin-removed conditioned media, ORS cells were seeded at a density of 2 × 10 5 cell/cm 2 on 12 well tissue culture plate (SPL LIFE SCIENCES, 30012) to make confluent ORS cell layer. The ORS cells were adjusted to be stable for 1 day in human ORS cell growth medium (CEFO, CB-ORS-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 day of adjustment, ORS cells were cultured under various culture media; human ORS cell growth medium, conditioned medium collected from ORS cell layer, conditioned medium collected from TGFβ2-treated ORS cell layer, conditioned medium collected from TGFβ2-treated ORS cell layer and incubated with guinea pig normal IgG-conjugated beads, conditioned medium collected from TGFβ2-treated ORS cell layer and incubated with guinea pig anti-Type I + II Hair Keratins antibody-conjugated beads, and human ORS cell growth medium containing 1(w/v)% keratin. After 3 days of culture, P-cadherin expressing germ formation of ORS cells was characterized by immunocytochemical staining.

Caspase-3 and caspase-6-mediated hair keratin digestion assay

1(w/v)% hair keratin was dissolved in the reaction solution composed of 50 mM HEPES (Gibco, 15630-080), 50 mM NaCl (JUNSEI CHEMICAL, 19015-1250), 0.1% CHAPS (Sigma-Aldrich, C3023), 10 mM EDTA (Sigma-Aldrich, 03609), 5% glycerol (SAMCHUN CHEMICALS, G0274) and 10 mM DTT (Sigma-Aldrich, 43815) at pH 7.2. 5U/ml Casase-3 (Enzo, ALX-201-059) or 5U/ml Casase-6 (Enzo, ALX-201-060) was added to the reaction solution containing hair keratin and incubated at 37 °C for 0, 1, 3, and 24 h. After the reaction, samples were denatured at 70 °C for 10 min in LDS sample buffer (Invitrogen, B0007). Equal amounts of denatured samples were loaded in pre-casted 4–12% Bis-Tris Plus Gels (Invitrogen, NW04120BOX), and the electrophoresis was done by running at 200 V for 22 min. The gel was rinsed three times with distilled water for 5 min each and stained by SimplyBlue SafeStain (Invitrogen, LC6060). After 1 hr of staining, the gel was rinsed using distilled water until the background was removed thoroughly, and then images of the gel was obtained using a commercialized scanner (Canon, TS8090).

In vitro caspase-6 gene knockdown study

To evaluate the effect of caspase-6-mediated keratin degradation during TGFβ2-induced ORS cell apoptosis on keratin release or deposition and DP condensation, caspase-6 gene expression in ORS cells was silenced by caspase-6 siRNA transfection. Capase-6 siRNA duplex (Bioneer, 839-1) or negative control siRNA duplex (Bioneer, SN-1012) was diluted in 250 μl Opti-MEM (Gibco, 31985062) to make a final concentration of 100 nM. 3.5 μl Lipofectamine 2000 (Invitrogen, 11668-030) was mixed in 250 μl Opti-MEM, and the mixture was incubated for 5 min at room temperature. The diluted caspase-6 siRNA duplex and diluted Lipofectamine 2000 were mixed and incubated for 20 min at room temperature. Before transfection, ORS cells were seeded at a density of 2 × 10 5 cell/cm 2 on 12 well tissue culture plate (SPL LIFE SCIENCES, 30012) to make confluent ORS cell layer. The ORS cells were adjusted to be stable for 1 day in human ORS cell growth medium (CEFO, CB-ORS-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 day of adjustment, human ORS cell growth medium (CEFO, CB-ORS-GM) was changed with fresh same medium without serum. The capase-6 siRNA/Lipofectaminr 2000 mixture or negative control siRNA/Lipofectamine 2000 mixture was added to ORS cell culture, and incubated for 5 hr at 37 °C. After transfection, the medium was changed with a fresh medium containing serum, and the transfected ORS cells were cultured in the presence of 100 ng/ml TGFβ2 or in the absence of TGFβ2 for 5 days.

To evaluate DP cell condensation in direct co-culture of DP cells and TGFβ2-treated/caspase-6-silenced ORS cells, DP cells were stained with a cell tracker (Invitrogen, C34552) according to manufacturer’s instruction. The stained DP cells were seeded at a density of 2 × 10 4 cell/cm 2 on confluent TGFβ2-treated/caspase-6-silenced ORS cell layer which was cultured for 5 days prior to co-culture, and cultured in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 2 days of co-culture, DP cell condensation was observed under inverted fluorescent microscopy (Olympus IX71).

To evaluate DP cell condensation under conditioned media from TGFβ2-treated/caspase-6-silenced ORS cells, the conditioned media were collected from TGFβ2-treated/caspase-6-silenced ORS cell culture after 5 days. DP cells were seeded at a density of 2 × 10 4 cell/cm 2 on 12 well and 6 well non-treated tissue culture plate (SPL LIFE SCIENCES, 32012, 32006). The DP cells were adjusted to be stable for 1 day in a human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 day of adjustment, DP cells were cultured under various culture media; human DP growth medium, human ORS cell growth medium containing 100 ng/ml TGFβ2, conditioned medium collected from TGFβ2-treated/negative control siRNA-transfected ORS cell layer, conditioned medium collected from TGFβ2-treated/negative control siRNA-transfected ORS cell layer containing additional 100 ng/ml of TGFβ2, and conditioned medium collected from TGFβ2-treated/caspase-6-silenced ORS cell layer. After 1 and 2 days of culture, DP cell condensation was observed under inverted fluorescent microscopy (Olympus IX71).

In vitro KRT31/KRT34 gene knockdown study

To evaluate the effect of KRT31/KRT34 gene knockdown during TGFβ2-induced ORS cell apoptosis on keratin release or deposition and germ formation of ORS cells, KRT31 and KRT34 gene expressions in ORS cells were silenced by KRT31/KRT34 siRNA transfection. KRT31 siRNA duplex (Bioneer, 3881-1) and KRT34 siRNA duplex (Bioneer, 3885-1) or negative control siRNA duplex (Bioneer, SN-1002) were diluted in 250 μl Opti-MEM (Gibco, 31985062) to make a final concentration of 100 nM. 3.5-μl Lipofectamine 2000 (Invitrogen, 11668-030) was mixed in 250 μl Opti-MEM, and the mixture was incubated for 5 min at room temperature. The diluted KRT31/KRT34 siRNA duplex and diluted Lipofectamine 2000 were mixed and incubated for 20 min at room temperature. Before transfection, ORS cells were seeded at a density of 2 × 10 5 cell/cm 2 on 12 well tissue culture plate (SPL LIFE SCIENCES, 30012) to make confluent ORS cell layer. The ORS cells were adjusted to be stable for 1 day in human ORS cell growth medium (CEFO, CB-ORS-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 1 day of adjustment, human ORS cell growth medium (CEFO, CB-ORS-GM) was changed with fresh same medium without serum. The KRT31/KRT34 siRNA/Lipofectamine 2000 mixture or negative control siRNA/Lipopectamin 2000 mixture was added to ORS cell culture, and incubated for 5 hr at 37 °C. After incubation, the medium was changed with a fresh medium containing serum, and the transfected ORS cells were cultured in the presence of 100 ng/ml TGFβ2 for 5 days, and morphological change was observed under inverted fluorescent microscopy (Olympus IX71).

In order to evaluate the activity of KRT31/KRT34 knockdown ORS cells for cell growth and differentiation, cell proliferation was tested by Cell Counting Kit-8 up to 5 days after gene knockdown. Each well-containing cells was replaced with 100 μl of DMEM and 10 μl of Cell Counting Kit-8 solution, and incubated at 37 °C for 1 h. Cell proliferation assays were performed in a 96-well plate reader by measuring the absorbance at a wavelength of 450 nm. In addition, BrdU was added to each well-containing cells to observe BrdU incorporation activity by immunocytochemical staining. BrdU was added as final concentration of 10 μM, and the treated cells were cultured for 24 h. After cell fixation and permeabilization for immunofluorescent staining, DNA hydrolysis was performed using 1 M Hcl. Subsequent procedures were performed in accordance with the procedures of Immunocytochemical staining.

Apoptosis and growth factor antibody array

TGFβ2-induced ORS cell apoptosis was evaluated by comparative analysis using human apoptosis antibody array (Abcam, ab134001), and the comparative analysis of growth factors present in the conditioned medium collected from TGFβ2-treated ORS cell culture in immunodepletion study were done using human growth factor antibody array (Abcam, ab134002) according to manufacturer’s instructions.

For apoptosis array analysis, cell lysates were prepared as follows; cells on culture plate were washed with PBS three times. After washing, cells were incubated with the 1× Lysis buffer at 4 °C for 30 min, and lysates were centrifuged at 12,000 rpm for 10 min to remove the debris. After centrifuge, the supernatant was collected and used for the apoptosis array analysis. Following process of apoptosis antibody array was done according to manufacturer’s instructions. Dot blots on membrane were analyzed using the plugin Protein Array Analyzer ( http://image.bio.methods.free.fr ) on ImageJ ( https://imagej.nih.gov/ij/ ).

For growth factor antibody array analysis, in the immunodepletion study, conditioned media were collected from TGFβ2-treated ORS cell culture and prepared as follows; the collected conditioned media were centrifuged to remove debris, and the supernatant was collected. The supernatant was concentrated ~20 times by Amicon Ultra centrifugal filter units (Millipore, Z717185). The concentrated conditioned media were used for growth factor antibody array analysis and following process of growth factor antibody array was done according to manufacturer’s instructions. Dot blots on membrane were analyzed using the plugin Protein Array Analyzer ( http://image.bio.methods.free.fr ) on ImageJ ( https://imagej.nih.gov/ij/ ).

Western blot analysis

Molecular expressions of KRT34 and β-catenin in hair keratin-treated ORS cells, keratin content at protein level in conditioned medium collected from TGFβ2-treated ORS cell culture, the keratin content in keratin-removed condition medium collected from TGFβ2-treated ORS cell culture in immunodepletion study, keratin content at protein level in conditioned medium collected from KRT31/KRT34-silenced ORS cell culture or molecular keratin expression in KRT31/KRT34-silenced ORS cell, molecular expressions of caspase 6 in TGFβ2-treated ORS cells and keratin content at the protein level in conditioned medium collected from caspase 6-silenced ORS cell culture were evaluated by western blot analysis.

Proteins from various conditioned medium were concentrated about twentyfold by Amicon Ultra centrifugal filter units (Millipore, Z717185), and cells were lysed on ice for 30 min in 100 µl ice-cold RIPA lysis buffer (Millipore, 20–188) containing a protease inhibitor cocktail (Roche, 4693116001), and then lysates were centrifuged at 12,000 rpm to remove debris. Samples were denatured at 70 °C for 10 min in LDS sample buffer (Invitrogen, B0007), and equal amounts of denatured samples were loaded in pre-casted 4–12% Bis-Tris Plus Gels (Invitrogen, NW04120BOX), and the electrophoresis was done by running at 200 V for 22 min. After electrophoresis, the proteins in the gel were transferred to PVDF membranes (Bio-Rad, 1620174) using electrophoretic transfer cell (Bio-Rad, 1703930). Immunoblotting for the membranes was carried out as follows; the membranes were incubated in TBS containing 5% skim milk (bioWORLD, 30620074-1) at room temperature for 60 min. The membranes were further incubated in TBS containing 1% skim milk and primary antibodies for overnight at 4 °C. The primary antibodies used in western blot analysis were as follows; rabbit Anti-KRT34 (LifeSpan BioSciences, LS‑B15620, diluted 1:1000), guinea pig Anti-Type I + II Hair Keratins (PROGEN, GP-panHK, diluted 1:1000), mouse anti-FGF-20 (Santa Cruz, sc-398722, diluted 1:1000), rabbit anti-Annexin V (Abcam, ab14196, diluted 1:2000), rabbit anti-Caspase-3 (Cell Signaling Technology,8G10, diluted 1:2000) and Mouse Anti-GAPDH (Abcam, ab8245, diluted 1:5000). After incubation with primary antibodies, the membranes were incubated with following secondary antibodies in TBS containing 1% skim milk at room temperature for 2 hr; HRP conjugated goat anti-guinea pig IgG (Abcam, ab97155, diluted 1:10000), HRP conjugated mouse anti-rabbit IgG (Santa Cruz Biotechnology, sc-2357, diluted 1:5000) and HRP conjugated goat anti-mouse IgG (Santa Cruz Biotechnology, sc-2005, diluted 1:5000). At each process, the membranes were washed three times with TBS containing 0.1% Tween 20 (Duchefa Biochemie, P1362.1000) for 10 min. The membranes were treated with ECL substrate (Bio-Rad, 1705061) to visualize signal, and the signals on the membranes were transferred to X-ray film (AGFA, CP-BU New).

Real-time quantitative polymerase chain reaction (RT-qPCR)

The gene expressions indicative of DP cell’s intrinsic property and TGFβ2 gene expressions of DP cell spheroids and the replated DP cell spheroids were evaluated byRT-qPCR. The gene expressions indicative of DP cell’s intrinsic properties were evaluated by RT-qPCR. Total RNA was extracted from DP cells and DP Cell spheroids using Hybrid-R (GeneAll, 305-101). cDNAs were synthesized by reverse transcription reaction using a CycleScript RT PreMix (Bioneer, K-2044). After cDNA synthesis, cDNAs were mixed with SYBR kit (PhileKorea, QS105-10) and primers (Table  S1 ). The mixture was reacted by thermal cycler (QIAGEN, Rotor-Gene Q) and analyzed by the program provided (Rotor-Gene Q Series Software, V1.7). The whole process was carried out according to the manufacturer’s instructions.

TGFβ2 gene expressions of DP cell spheroids and the replated DP cell spheroids were evaluated by RT-qPCR. DP cell spheroids were generated by PEG microwell-mediated condensation as previously described. The formed DP cell spheroids were retrieved from PEG microwell, and then replated on 6 well tissue culture plate (SPL LIFE SCIENCES, 30006). The replated DP cell spheroids were cultured in human DP growth medium (CEFO, CB-HDP-GM) at 37 °C in a humidified atmosphere containing 5% CO 2 . After 0, 1, 3, 5, and 7 days of culture, total RNA was extracted from DP cell spheroid, and RT-qPCR was done by the previously described same method. The primer used in this experiment was noted in Table  S1 .

Indirect enzyme-linked immuno-sorbent assay (ELISA)

The molecular expressions indicative of DP cell’s intrinsic property from the replated DP spheroids cultured in the presence of keratin were evaluated by ELISA. Replated DP spheroids cultured in the presence of keratin were harvested and lysed with RIPA lysis buffer (Millipore, 20–188) containing a protease inhibitor cocktail (Roche, 4693116001). Lysates were centrifuged for 15 min at 12,000 rpm to remove the debris, and the supernatant was collected. The lysates were diluted to a final concentration of 20 μg/ml in DPBS (Dulbecco’s Phosphate-Buffered Saline; Gibco, 14190-144). Antigens were coated on wells in a 96-well ELISA plate (Invitrogen, 44-2404-21) by loading 50 μl of the diluted lysate, and the wells were incubated at 4 °C for 18 hr. The plate was washed three times with 200 μl of DPBS, and non-specific binding of antibodies was blocked by treating with DPBS containing5(w/v)% bovine serum albumin (BSA; Sigma-Aldrich, A9418) for 2 hr at room temperature. Primary antibodies such as rabbit anti-β-actin (Abcam, ab8227, diluted 1:1000), rabbit anti-β-catenin (Abcam, ab16051, diluted 1:1000), rabbit anti-FGF7 (Santa Cruz Biotechnology, sc-7882, diluted 1:200), Goat anti-FGF10 (Santa Cruz Biotechnology, sc-7375, diluted 1:200) and goat anti-BMP6 (Santa Cruz Biotechnology, sc-7406, diluted 1:200) were diluted in DPBS containing 1(w/v)% BSA, and added 100 μl to each well. The plate was incubated at room temperature for 2 hr for the reaction of antibodies with antigens. After incubation with primary antibodies, the wells were washed with DPBS, and 100 μl of HRP conjugated secondary antibodies (anti-rabbit IgG-HRP (Santa Cruz Biotechnology, sc-2004) or anti-goat IgG-HRP (Santa Cruz Biotechnology, sc-2033)) was added to each well, and then was incubated for 2 hr at room temperature. After incubation with secondary antibodies and following washing with DPBS. 100 μl of TMB substrate solution (Thermo Scientific, N301) was added to each well and incubated for 30 min at room temperature. The reaction was stopped by 100 μl of the stop solution (2 M sulfuric acid; Sigma-Aldrich, 258105), and the absorbance of samples was measured at 450 nm on a microplate spectrophotometer reader (Bio-Rad, Benchmark Plus, BR170-6930).

In vivo KRT31/KRT34 gene knockdown study

To confirm the effect of keratin on hair growth, KRT31/KRT34 was silenced by lipofectamine-mediated delivery of KRT31/KRT34 siRNAs. First, Invivofectamine complex for KRT31/KRT34 siRNA delivery was prepared as follows; siRNAs of KRT31 (Bioneer, 16660-1), KRT34 (Bioneer, 16672-1) and negative control (Bioneer, SN-1003) were purchased, and siRNAs of KRT31 and KRT34 were dissolved in RNase-free water as each24 mg/ml concentration respectively. The two solutions, KRT31 siRNA and KRT34 siRNA, were combined as 1:1 volume ratio to be 12 mg/ml of final concentration. 12 mg of negative control siRNA was also dissolved in 1 ml of RNase-free water. siRNAs-Invivofectamine (Thermo Fisher Scientific, IVF3005) complex was prepared by the manufacturer’s instruction, and 0.5 mg/ml of the complex was prepared finally prior to injection to mice. For in vivo study, 6-week-old mice were shaved repeatedly to synchronize the hair cycle and randomly assigned to three groups: Con group with IV injection of 200 μl of negative control siRNA injection; siRNA group with IV injection of 200 μl of KRT31/KRT34 siRNA injection; siRNA+Keratin group with KRT31/KRT34 siRNA injection (IV, 200 μl) and intradermal injection of total 100 μl of keratin a day after first siRNA injection. For each group, mice were sacrificed at either day 7 or day 14. Pictures of the back skin were taken on day 3, 7, 10, and 14 to examine the hair growth. The silencing of KRT31/KRT34 gene expressions was confirmed by RT-qPCR.

Total RNA was extracted from the cells using Hybrid-R (GeneAll, 305-101). cDNAs were synthesized by reverse transcription reaction using a CycleScript RT PreMix (Bioneer, K-2044). After cDNA synthesis, cDNAs were mixed with SYBR kit (PhileKorea, QS105-10) and primers (Table  S2 ). The mixture was reacted by a thermal cycler (QIAGEN, Rotor-Gene Q) and analyzed by the program provided (Rotor-Gene Q Series Software, V1.7). The whole process was carried out according to the manufacturer’s instructions.

Histological analysis

The skin tissues were fixed with 10% neutral-buffered formalin (BBC Biochemical, 0141). The tissues were embedded in paraffin and sectioned at 4-μm thickness, followed by staining with hematoxylin and eosin for histological analysis. The number of hair follicles in each cycle and the diameter of anagen hair follicles was quantified in multiple fields on perpendicular sections at ×100 magnification.

Immunocytochemical staining

DP cells and ORS cells were fixed for 10 min in 3.7% paraformaldehyde (Sigma-Aldrich, F8775), and were permeabilized in 0.2% Triton X-100 (Sigma-Aldrich, T9284). Non-specific binding was blocked by treating with 4% BSA (Sigma-Aldrich, A9418). Cells were incubated in primary antibody diluents (GBI Labs, E09-500) containing the following primary antibodies for overnight at 4 °C; rabbit anti-β-catenin (Abcam, ab16051, diluted 1:100), rabbit anti-SOX2 (Cell Signaling Technology, 3579 S, diluted 1:200), rabbit anti-CD133 (Abcam, ab16518, diluted 1:50), mouse anti-integrin β1 (Santa Cruz Biotechnology, sc-59829, diluted 1:50), rabbit anti-P-cadherin (Cell Signaling Technology, 2189 S, diluted 1:50), mouse anti-E-cadherin (Abcam, ab1416, diluted 1:100), mouse anti-alkaline phosphatase (Abcam, ab126820, diluted 1:100), mouse anti-RUNX1 (Santa Cruz Biotechnology, sc-365644, diluted 1:50), rabbit anti-KRT34 (LifeSpan BioSciences, LS‑B15620, diluted 1:100), rabbit anti-FGF7 (Santa Cruz Biotechnology, sc-7882, diluted 1:50), goat anti-FGF10 (Santa Cruz Biotechnology, sc-7375, diluted 1:50), goat anti-BMP6 (Santa Cruz Biotechnology, sc-7406, diluted 1:50), rabbit anti-CD34 (Abcam, ab81289, diluted 1:100), rabbit anti-SOX9 (Abcam, ab185966, diluted 1:100), rabbit anti-Annexin V (Abcam, ab14196, diluted 1:100), rabbit anti-caspase-3 (Abcam, ab13847, diluted 1:100), rabbit anti-caspase-6 (Abcam, ab52951, diluted 1:100), mouse anti-BrdU (Invitrogen, MA3-071, diluted 1:100), rabbit anti-Ki67 (Cell Signaling Technology, 9027 S, diluted 1:100), rabbit anti-Lgr5 (Abcam, ab219107, diluted 1:100) and rabbit anti-Vinculin (Abcam, ab129002, diluted 1:100). After incubation with primary antibodies, cells were washed three times with DPBS and were incubated with the following secondary antibodies for 1 hr at room temperature; Alexa Fluor 488-conjugated goat anti-rabbit IgG (Invitrogen, A-11034, diluted 1:200), Alexa Fluor 594-conjugated goat anti-rabbit IgG (Invitrogen, A-11012, diluted 1:200), Alexa Fluor 594-conjugated goat anti-mouse IgG (Invitrogen, A-11032, diluted 1:200) and Alexa Fluor 488-conjugated goat anti-mouse IgG (Invitrogen, A-11001, diluted 1:200). With secondary antibodies, actin was stained using rhodamine phalloidin (Invitrogen, R415, diluted 1:400), and Tunel staining was performed using the Turner Enzyme (Roche, 11767305001) and Tunel Label (Roche, 11767291910). Briefly, cells were washed twice with DPBS (Gibco, 14190-144) and incubated at 37 °C for 1 hr with 200 μl of TUNEL reaction mixture (Turner Enzyme: Tuner Label, 1:9, v/v). After incubation with secondary antibodies, phalloidin, and Tunel reaction mixture, cells were washed with DPBS three times and counterstained with DAPI (Sigma-Aldrich, D9542). Finally, stained cells were observed under an inverted fluorescence microscope (OLYMPUS, IX71) and captured.

Immunohistochemical staining

Paraffin-embedded tissue sections were deparaffinized by treating xylene three times for 5 min and rehydrated in graded ethanol (100%, 95%, 90%, 80%, and 50%). Antigen retrieval step was performed by incubating the sections in sodium citrate buffer (10 mM Sodium Citrate; Biosesang, C2004, 0.05% Tween 20; Duchefa Biochemie, P1362.1000, pH 6.0) at sub-boiling temperature for 20 min. Non-specific binding of antibody was blocked by treating with blocking buffer containing 1% normal horse serum (Abcam, ab7484) and 5% BSA (Sigma-Aldrich, A9418) for 30 min at room temperature. After blocking, tissue sections were incubated in primary antibody diluents (GBI Labs, E09-500) containing the following primary antibodies for overnight at 4 °C; goat anti-P-cadherin (R&D Systems, AF761, diluted 1:50), rabbit anti-β-catenin (Abcam, ab16051, diluted 1:50), rabbit anti-KRT34 (Biorbyt, orb628339, diluted 1:100), guinea pig anti-type I + II hair keratins (PROGEN, GP-panHK, diluted 1:50), mouse anti-Ki67 (Abcam, ab279653, diluted 1:50), rabbit anti-Annexin V (Abcam, ab14196, diluted 1:100), rabbit anti-Caspase-3 (Abcam, ab13847, diluted 1:50), and rabbit anti-active Caspase-3 (Abcam, ab32042, diluted 1:50). When the host species of primary antibodies were a mouse, endogenous mouse Ig of the tissue was blocked with Mouse on Mouse blocking agent (Abcam, BMK-2202) according to the manufacturer’s instruction. After washing tissue sections with DPBS three times, were incubated with the following secondary antibodies overnight at 4 °C; donkey Alexa Fluor 488-conjugated anti-goat IgG (Invitrogen, A-11055, diluted 1:200), goat Alexa Fluor 594-conjugated anti-rabbit IgG (Invitrogen, A-11012, diluted 1:200), goat Alexa Fluor 488-conjugated anti-rabbit IgG (Invitrogen, A-11034, diluted 1:200) and goat Alexa Fluor 647-conjugated anti-guinea pig IgG (Abcam, ab150187, diluted 1:200). After incubation with secondary antibodies, tissue sections were washed with DPBS three times, counterstained with DAPI (Sigma-Aldrich, D9542) and mounted with mounting medium (Sigma-Aldrich, F4680). Finally, images were acquired using an inverted fluorescence microscope (OLYMPUS, IX71).

In situ RNA hybridization

In situ hybridization (ISH) was performed with the RNAscope (Advanced Cell Diagnostics, ACD), including the RNAscope 2.5 Duplex reagent kit and RNAscope probes. ISH was performed manually with an RNAscope 2.5 HD Duplex assay (Chromogenic) in accordance with the manufacturer’s instructions for Formalin-Fixed Paraffin-Embedded (FFPE) tissue. RNA probes hybridizing to mouse LGR5 RNA (ACD, 312178 -C2) and mouse CD34 RNA (ACD, 319168 -C2) were hybridized to the tissues. Steps of amplification and detection were conducted as recommended by the RNAscope 2.5 HD duplex detection kit user manual (ACD, 322500-USM). Tissue sections were observed under a standard light microscope (OLYMPUS, IX71) and captured.

Flow cytometric analysis

Molecular expressions of P-Cadherin and RUNX1 for were analyzed by flow cytometry. ORS cells were cultured in the presence of TGFβ2 and harvested by incubating with 0.25% Trypsin/EDTA for 10 min at 37 °C. Cells were recovered by centrifugation and fixed with 0.01% formaldehyde in DPBS for 15 min at room temperature. Cells were incubated for 30 min at room temperature with primary antibodies such as Mouse anti-RUNX1 (Santa Cruz Biotechnology, sc-365644, diluted 1:50) or Mouse anti-P-Cadherin (R&D Systems, FAB861G, diluted 1:100). Cells were resuspended in 100 μl of diluted fluorochrome-binding secondary antibody (Alexa Fluor 488-conjugated anti-mouse IgG, A-11001). Washing with DPBS was performed in each step. Flow cytometric analysis was done using a FACSCanto (BD Biosciences, USA), and analysis of flow cytometry data was done with FlowJo V10 program.

Statistics and reproducibility

All values obtained from in vitro and in vivo analysis are presented as the mean ± standard deviation (SD). Statistically, differences were identified by two-sided Student’s t test or one-way ANOVA parametric test. A P value of <0.05 was considered significant. The number of samples per independent experiment is described in the legends.

Reporting summary

Further information on research design is available in the  Nature Portfolio Reporting Summary linked to this article.

Data availability

All data that support the findings of this study are available from the corresponding author upon reasonable request. Full-length uncropped original western blots used in the manuscript are shown in Supplementary Fig.  42 . The numerical data that make up the all graphs in the paper are shown in Supplementary Data  1 – 6 . The transcriptome sequencing data (RNA-Seq) have been deposited at NCBI GenBank under BioProject ID PRJNA576064 (BioSample SAMN12924151–SAMN12924158). The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the jPOST partner repository with accession numbers PXD037830 (JPST001909).

Change history

22 december 2022.

A Correction to this paper has been published: https://doi.org/10.1038/s42003-022-04366-w

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This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korean government (MSIT) (NRF-2016R1D1A1B03931933) and by the Bio & Medical Technology Development Program of NRF funded by the Korean government (MSIT) (no. 2017M3A9E4048170), and by a fund (2018ER610300) by Research of Korea Centers for Disease Control and Prevention.

Author information

So Yeon Kim

Present address: Department of Dental Hygiene, College of Health Science, Cheongju University, Cheongju, 360-764, Republic of Korea

Han Jun Kim

Present address: Terasaki Institute for Biomedical Innovation, Los Angeles, CA, 90064, USA

Authors and Affiliations

Department of Maxillofacial Biomedical Engineering, College of Dentistry, Kyung Hee University, Seoul, 02447, Republic of Korea

Seong Yeong An, So Yeon Kim, Se Young Van & Yu-Shik Hwang

Department of Veterinary Clinical Pathology, College of Veterinary Medicine, Konkuk University, 120 Neungdong-ro, Gwangjin-gu, Seoul, 05029, Republic of Korea

Hyo-Sung Kim, Han Jun Kim & Sun Hee Do

Department of Oral Microbiology, College of Dentistry, Kyung Hee University, Seoul, 02447, Republic of Korea

Jae-Hyung Lee

KeraMedix Inc, # 204, Open Innovation Bld, Hongryeung Bio-Cluster, 117-3 Hoegi-ro, Dongdaemun-gu, Seoul, 02455, Republic of Korea

Song Wook Han

Department of Dental Materials, College of Dentistry, Kyung Hee University, Seoul, 02447, Republic of Korea

Il Keun Kwon

Headquarters of New Drug Development Support, Chemon Inc. 15 F, Gyeonggi Bio Center, Cheongju, Gyeonggi-do, 16229, Republic of Korea

Chul-Kyu Lee

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S.Y.A. performed most of the experiments and wrote the paper. S.Y.K. extract and purified human hair-derived keratin. S.Y.V. extract and purified human hair-derived keratin and analyzed gene expressions using real-time-qPCR. H.S.K. carried out in vivo silencing experiment and histological analysis. H.J.K. carried out in vivo mouse experiment and histological analysis. J.H.L. carried out RNA sequencing and data analysis. S.W.H. and I.K.K. discussed the results of the experiments and commented on the manuscript. C.K.L. carried out in vivo mouse experiment and histological analysis. Y.S.H. and S.H.D. directed the project, and Y.S.H. drafted the manuscript with input from all authors.

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Correspondence to Sun Hee Do or Yu-Shik Hwang .

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An, S.Y., Kim, HS., Kim, S.Y. et al. Keratin-mediated hair growth and its underlying biological mechanism. Commun Biol 5 , 1270 (2022). https://doi.org/10.1038/s42003-022-04232-9

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what is bio keratin

Keratin definition and example

​​​​​​​​​​​​​​Keratin n., plural: ​​​​​​​​​​​​​​keratins [ˈkɛrətɪn] Definition: Structural protein found in hair, nails, and outer layer of skin

Table of Contents

Keratin is a fibrous protein naturally present in hair, skin, and nails. In hair care, it serves as a protective and structural element, enhancing strength and smoothness while reducing frizz. Keratin treatments aim to replenish and fortify hair’s natural keratin content, resulting in healthier, more manageable locks. Read on to learn more about keratin, its definition, examples, genetics, structure, and significance in biology and medicine.

What Is Keratin?

Keratin is a natural fibrous structural protein that forms an integral part of human hair, nails, and skin and is characterized by its intricate secondary structure. Comprising alpha helices , this alpha keratin forms a robust framework, which is a fibrous and structural protein consisting of polypeptide chains that intertwine to create a sturdy and resilient framework.

The amino acids that make up keratin also change depending on where it is found and what it does. Cysteine residues are especially important because they make cystines ( a type of chemical bond when disulfide bonds link cysteine residues together covalently ). Keratin is very stable because of the cysteines in it.

Keratin is insoluble in warm as well as in hot water and is unaffected by proteolytic enzymes . Fully hydrated keratin filaments (about 16 percent water) are 10 to 12 percent longer than those that are dehydrated.

keratin filaments SEM cross-section of fingernail

Watch this vid about ​​​​​​​​​​​​​​a type of keratin, alpha-keratin:

Biology definition: Keratin is a fibrous structural protein abundant in hair, nails, skin, feathers, hooves, horns, and so on. Keratins are made up of coiled polypeptide chains and when they combine they form supercoils. Keratins protect epithelial cells from damage.

Skin cells that are constantly exposed to pressure and rubbing leads to the formation of calluses. The skin thickens and the epidermal cells undergo cornification. Cornification is a process in which a keratinized layer of epidermis forms and serves as an epidermal barrier. During cornification, keratin fills up the cell resulting in the loss of cytoplasmic organelles and the cessation of metabolism. Ultimately, the fully keratinized cells undergo programmed cell death.

Overview: Keratin is one of the types of scleroproteins or  fibrous proteins .  Scleroproteins , in turn, are one of the three major types of proteins (the other two are spheroproteins and membrane proteins ). Scleroproteins are characterized by their long protein filaments. They act as structural proteins. They are usually water-insoluble. Apart from keratin, other examples of scleroproteins are collagen , elastin , and fibroin.

Etymology: from German “keratin”, from Ancient Greek “κέρας” (kéras), meaning “horn” +‎ -in.

Examples Of Occurrence

Keratin filaments are widespread within keratinocytes throughout the entirety of the cornified layer of the epidermis. Additionally, they are found in every epithelial cell. For instance, antibodies against keratin 8, keratin 5, and keratin 14 react with murine thymic epithelial cells. Antibodies of this type have been utilized as fluorescent markers in a genetic study of the thymus of rats.

Both vertebrates have keratin in their bodies. The slime threads of hagfish and the hair fiber, nails, wool, claws, epidermis, horns, and hooves of mammals all contain keratin.

Only sauropsids, which include all living reptiles and birds, have keratin. Various animal parts, including beaks, feathers, and talons of birds, as well as the scales, nails, and claws of reptiles, are embedded in them.

In the antlers of animals like the impala, keratin covers a bone core.

Horns of the impala

On chromosomes 12 and 17 , there are two distinct clusters containing a total of 54 functional keratin genes. This suggests that their genesis can be attributed to a series of gene duplications on these particular chromosomes. The keratin genes are clustered on different chromosomal loci and are distributed across multiple chromosomes.

The expression of keratin genes is tissue-specific and regulated by various factors, including developmental stages, environmental cues, and cellular differentiation . Keratins play a critical role in maintaining the structural integrity of various epithelial tissues. Mutations in keratin genes lead to different disorders such as epidermolysis hyperkeratosis, Areata Alopecia, Keratosis pharyngitis, and Simplex Epidermolysis Bullosa .

Protein Structure

During the years 1982 and 1983, Elaine Fuchs and Israel Hanukoglu made a significant breakthrough by uncovering the primary sequences of keratins. These sequences provided compelling evidence for the presence of two distinct yet closely related keratin families, known as type I and type II keratins . Hanukoglu and Fuchs put forth a conceptual model suggesting that keratins and intermediate filaments proteins contain a central domain that extends across 310 amino acid residues. This domain is comprised of four segments anticipated to adopt an alpha-helical structure , alongside three concise linker segments predicted to assume a beta-turn conformation.

Type 1 and 2 Keratins

Within the human genome, 54 keratin genes have been categorized based on their functional roles. Among these, 28 are found in type 1 and the remaining 26 in type 2.

  • Type I keratins are typically smaller and more acidic proteins. This type is essential for maintaining the integrity of epithelial cells.
  • Type II keratins are relatively massive, neutral pH proteins.

Disulfide bridges

Apart from intramolecular and intermolecular hydrogen bonds, keratins exhibit a notable abundance of the amino acid cysteine, containing sulfur, which is crucial for forming disulfide bridges. These bridges impart enhanced stiffness and lasting strength through enduring, thermally stable linkages, similar to how non-protein sulfur bonds enhance the stability of vulcanized rubber.

Roughly 14% of the human hair cuticle consists of cysteine. When hair and epidermis burn, they release distinct odors due to volatile sulfur compounds. The substantial disulfide bonding makes keratins insoluble in the absence of certain dissociating or reducing agents.

Comparatively, the keratins present in hair, which are more pliable and supple, possess few inter-chain disulfide bridges as compared to the keratins found in the nails, horns, and claws of mammals, which tend to be more rigid.

Hair and α-keratins are constructed from individual protein strands coiled in an α-helical manner which are then intertwined into superhelical chains that can undergo further coiling.

In reptiles and birds, the disulfide bridges within their keratins serve to anchor and fortify their β-pleated sheets, which are subsequently intricately intertwined.

Filament formation

Keratin filament formation is the process of assembling keratin proteins into complex structures within cells , contributing to the strength of tissues like skin, hair, and nails. This involves the arrangement of keratin molecules into dimers, tetramers, and higher-order structures, guided by alpha-helical segments and disulfide bridges formed by sulfur-containing amino acids. This process is regulated by factors such as tissue type, cellular differentiation, and environmental cues. The resulting filaments provide tissues with mechanical resilience against stress.

Microscopy of keratin filaments inside cells

Keratins can be classified into two categories:

  • Neutral-Basic

These keratins are associated with specific occurrences in various tissues.

  • For instance, keratin 1 and keratin 2, along with keratin 9 and keratin 10, are found in the stratum corneum and keratinocytes .
  • Keratin 3 and keratin 12 are present in the cornea
  • Keratin 4 and keratin 13 are associated with stratified epithelium.
  • Keratin 5, along with keratin 14 and keratin 15, is found in the stratified epithelium as well.
  • Keratin 6, keratin 16, and keratin 17 are linked to squamous epithelium.
  • Keratin 7 and keratin 19 are present in ductal epithelia , and keratin 8, keratin 18, and keratin 20 are associated with simple epithelium .
  • The distribution of these keratins in specific tissues showcases their diverse roles in maintaining tissue integrity and function.

Data Source: Shoaib Zaheer of Biology Online


Cornification is the formation of an epidermal barrier consisting of a horny layer of the epidermis . The production of keratin characterizes it. In addition to keratin production, cornification involves the production of minor proline-rich proteins and transglutaminase.

Under the plasma membrane , this results in the development of a cornified cell envelope. During the final phases of cornification, the cells shed their organelles, including their nuclei , and fill with keratin. Consequently, this results in a shutdown of metabolism . The completely keratinized cells eventually undergo programmed cell death .

On the skin’s surface, cells undergo cornification, particularly epidermal cells, and become nearly impermeable. The result is an epidermal barrier that is both rigid and elastic. Persistent scratching and pressure on the cornified layer result in further thickening and callus formation. However, since epidermal cells are continually shed and replaced, calluses may dissolve when pressure is removed.

It is a fibrous material composed of keratin proteins, typically produced by certain insects and spiders. but their evolutionary connection to keratins found in vertebrates remains ambiguous.

Typically, spider silk measures between 1 and 2 micrometers (µm) in thickness, in contrast to roughly 60 µm for human hair and even greater dimensions for specific mammals.

The valuable characteristics of silk fibers, both in biological and commercial contexts, are intricately tied to how multiple protein chains are structured. These chains are structured into compact, crystalline segments of varying dimensions, intermingled with pliable, amorphous regions where the chains intertwine randomly. This structural pattern is somewhat reminiscent of what occurs with man-made polymers like nylon, which was developed to emulate silk.

In the case of silk obtained from hornet cocoons, it comprises pairs of structures that are approximately 10 µm in width. These structures consist of central cores and outer coatings, and they can be stacked in layers, sometimes up to 10 layers thick. These layers can also form variedly shaped plaques. Interestingly, fully grown hornets employ silk not only as a material for creating intricate structures but also as an adhesive, much like how spiders use silk.

Hoof glue and horn glue, both of which are formed from partially hydrolyzed keratin, are animal-based proteins. These proteins are broken down through a process called hydrolysis, which involves the use of heat and chemicals to break the complex protein structures into smaller fragments.

Hoof glue is derived from the hooves of animals, typically cattle. It has been used for centuries in woodworking, bookbinding, and other crafts due to its strong adhesive properties, ease of use, and ability to bond well with various materials.

Horn glue, similarly, is derived from the keratin protein found in animal horns. The process of producing horn glue is similar to that of hoof glue. Horn glue has been historically used for various applications, such as in the production of musical instruments, artwork, and even as an adhesive in ancient construction.

Clinical Significance

Protein-based biomaterials have surfaced as a promising substitute due to their inherent capacity for cellular interaction, structural reinforcement, and intercellular communication. Over the past century, advancements in the extraction, purification, and analysis of keratin proteins from sources like wool, feathers, antlers, and other animal-derived materials have paved the way for the creation of a foundation for keratin-oriented biomaterials.

With attributes akin to substances that the body produces naturally, keratins display inherent biological activity and compatibility.

Isolated keratins possess the ability to self-assemble into structures that play a role in cellular recognition and behavior modulation. It is these attributes that have led to the innovation of keratin-based biomaterials with applications spanning wound healing, pharmaceutical delivery, tissue engineering, trauma care, and medical apparatus.

Extensive research delves into the historical context of keratin investigation and the current landscape of novel methodologies in medical domains such as medical science, tissue engineering, regenerative medicine, and pharmaceutical delivery, all focused on comprehending keratin’s functionality within the human body.

Certain conditions can lead to abnormal keratin development, encompassing keratoderma, hyperkeratosis, and keratosis. Various ailments, such as athlete’s foot and ringworm, stem from fungi that feed on keratin.

Keratins play a pivotal role in modern hair cosmetics. They help maintain the strength and integrity of hair, reduce frizz, and hair loss, and promote healthy hair growth. Keratin treatments (e.g., Brazilian Blowouts) have gained popularity in hair salons and beauty supply stores due to their ability to transform frizzy and damaged hair into smoother and more manageable locks.

These treatments typically involve the application of a liquid keratin solution to the hair, which is then sealed using a flat iron. The heat helps in bonding the keratin to the hair, making the hair healthier in appearance.

Hair keratin treatment

It’s worth noting that some traditional keratin hair treatments (chemical protein treatments) cause formaldehyde exposure, which raised concerns about exposure to harmful chemicals. In response, newer formulations have emerged, focusing on safer methods for achieving the desired results.

Additionally, keratin supplements and hair products like keratin oils, and conditioners infused with keratin aim to support hair health from within, providing the building blocks necessary for keratin production and cell growth.

As science continues to explore the benefits of keratin, innovations like feather keratin hydrolysates obtained from sustainable sources have introduced environmentally friendly options that further contribute to the hair care landscape. However, excessive use of keratin solutions and treatments can lead to the accumulation of keratin residues that may eventually result in keratin clogs, impeding hair growth and health.

How does keratin treatment work on hair? 

Human keratin is a natural protein found in human hair. Keratin in hair, thus, has a vital role in maintaining hair strength and resilience. Keratin treatments are effective on hair due to the protein’s ability to bind to the hair shaft, filling in gaps and creating a smoother surface.

This results in reduced frizz, increased shine, and improved manageability. These effects are often achieved, though, by delaying hair washing after about 48 hours. This time frame is essential to ensure longer-lasting outcomes as it would allow the keratin molecules in the treatment to properly adhere to the hair and continue the process of polymerization.

Remember how the keratins are formed by disulfide bridges as mentioned earlier? This is one of the reasons shampoos that are sulfate-free are recommended for in washing keratin-treated hair. Sulfates, especially strong ones, can break down disulfide bridges in keratin proteins. The effects, however, eventually weaken over time, lasting typically around three months.

Diagnostic use

The manifestation of keratin aids in recognizing the epithelial source of anaplastic malignancies. Carcinomas, thymomas, sarcomas, and trophoblastic tumors are among the malignancies that exhibit keratin expression.

Moreover, the specific sequence of keratin subtypes being expressed allows for the anticipation of the primary tumor’s source when assessing metastatic instances. Cholangiocarcinoma typically expresses CK7, CK8, and CK18, whereas hepatocellular carcinomas typically express CK8 and CK18. However, metastases of colorectal carcinoma express CK20 but not CK7.

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  • BiologyOnline. (2023). Cornification. Retrieved 14 August, 2023, from https://www.biologyonline.com/dictionary/cornification
  • Gregersen., E. (2023). keratin. Retrieved 14 August, 2023, from https://www.britannica.com/science/polymer/Synthetic-polymers
  • Sarma, A. (2022). Biological importance and pharmaceutical significance of keratin: A review. International Journal of Biological Macromolecules.
  • Vedantu. (2023). Keratin. Retrieved 14 August, 2023, from https://www.vedantu.com/biology/keratin

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Last updated on August 18th, 2023

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Preparation and applications of keratin biomaterials from natural keratin wastes

  • Mini-Review
  • Published: 26 March 2022
  • Volume 106 , pages 2349–2366, ( 2022 )

Cite this article

  • Rong-Rong Yan 1 ,
  • Jin-Song Gong 1 ,
  • Chang Su 1 ,
  • Yan-Ling Liu 1 ,
  • Jian-Ying Qian 1 ,
  • Zheng-Hong Xu 2 , 3 &
  • Jin-Song Shi 1  

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Keratin is a kind of natural polymer that is abundant in feathers, wool, and hair. Being one of the natural biomolecules, keratin has excellent biological activity, biocompatibility, biodegradability, favorable material mechanical properties, and natural abundance, which exhibit significant biological and biomedical application potentials. At present, the strategies commonly used for preparing keratin from hair, feathers, wool, etc. include physical, chemical, and enzymatic methods. The present article mainly reviews the structure, classification, preparation methods, and the main biological applications of keratin, and these applications cover wound healing, hemostasis, targeted release of tissue engineering drugs, and so on. It is expected to lay the foundations for its future in-depth investigations and wide applications of keratin biomaterials.

• There are several pathways to prepare biologically active keratin from wool, feathers, and human hair, etc

• Promoting blood coagulation by keratin is related to the adhesion and activation of platelets and the aggregation of fibrin

• The biological applications of keratin, including wound healing and tissue engineering, are summarized

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what is bio keratin

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This work was supported by the National Key Research and Development Program of China (No. 2021YFC2100400), the National Natural Science Foundation of China (No. 21978116), the Fundamental Research Funds for the Central Universities (No. JUSRP22047), and the Ningxia Hui Autonomous Region Key Research & Development Plan (No. 2019BCH01002).

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Rong-Rong Yan, Jin-Song Gong, Chang Su, Yan-Ling Liu, Jian-Ying Qian & Jin-Song Shi

National Engineering Research Center for Cereal Fermentation and Food Biomanufacturing, School of Biotechnology, Jiangnan University, Wuxi, 214122, People’s Republic of China

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Jiangsu Provincial Research Center for Bioactive Product Processing Technology, Jiangnan University, Wuxi, 214122, People’s Republic of China

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RRY and JSG conceived this review. RRY, CS, YLL, and JYQ collected all references. RRY wrote the manuscript draft. JSG, ZHX, and JSS edited the manuscript. All the authors read and approved the manuscript.

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Yan, RR., Gong, JS., Su, C. et al. Preparation and applications of keratin biomaterials from natural keratin wastes. Appl Microbiol Biotechnol 106 , 2349–2366 (2022). https://doi.org/10.1007/s00253-022-11882-6

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Engineering with keratin: A functional material and a source of bioinspiration

Benjamin s. lazarus.

1 Materials Science and Engineering Program, University of California San Diego, 9500 Gilman Drive, La Jolla, CA, USA

Charul Chadha

2 Department of Mechanical Science and Engineering, University of Illinois Urbana-Champaign, Champaign, IL, USA

Audrey Velasco-Hogan

Josiane d.v. barbosa.

3 Department of Materials, University Center SENAI CIMATEC, Salvador, Brazil

Iwona Jasiuk

Marc a. meyers.

4 Department of Mechanical and Aerospace Engineering, University of California San Diego, San Diego, CA, USA

5 Department of Nanoengineering, University of California San Diego, San Diego, CA, USA

Keratin is a highly multifunctional biopolymer serving various roles in nature due to its diverse material properties, wide spectrum of structural designs, and impressive performance. Keratin-based materials are mechanically robust, thermally insulating, lightweight, capable of undergoing reversible adhesion through van der Waals forces, and exhibit structural coloration and hydrophobic surfaces. Thus, they have become templates for bioinspired designs and have even been applied as a functional material for biomedical applications and environmentally sustainable fiber-reinforced composites. This review aims to highlight keratin's remarkable capabilities as a biological component, a source of design inspiration, and an engineering material. We conclude with future directions for the exploration of keratinous materials.

Graphical abstract

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Biomaterials; Engineering materials; Materials design


Keratin is a ubiquitous biological polymer comprising the bulk of mammalian, avian, and reptilian epidermal appendages, including nails, hair, the outer layer of skin, feathers, beaks, horns, hooves, whale baleen, claws, scales, hagfish slime, and gecko pads ( Fraser et al., 1972 ; McKittrick et al., 2012 ; Meyers et al., 2008 ; Wang et al., 2016a ). The omnipresence of keratin-based materials in biological systems leads to a broad range of characteristics and functions, from the impact resistance of hooves and horns to lightweight yet stiff feathers that resist buckling under aerodynamic loads. Other examples include krill filtering by whale baleen, the protective scales of a pangolin, and even the reversible dry adhesive mechanism in gecko setae that allows these lizards to climb smooth vertical walls ( Huang, 2018 ; Labonte et al., 2016 ; Meyers et al., 2013 ). The ability of keratinous materials to perform diverse functions is derived from their ingenious structuring and tunability across many length scales. The broad array of architectures and their corresponding functions have led to the development of several keratin-inspired structures with tailored properties. Thus, keratin's structural diversity serves as a design template for the next generation of engineered materials.

Additionally, keratin has desirable intrinsic properties (biocompatibility, response to hydration, stiffness, strength, and other attributes). As a readily available and renewable material, it has been utilized as a raw material in fiber-reinforced composites. One aspect of keratin that deserves note is that it is comprised of keratinocytes after they undergo apoptosis and consists, for the most part, of ‘dead’ structures. Therefore, keratinous materials do not have the self-healing capability of living tissues such as bone. In bone, cells embedded in the structure tackle damage by repairing the torn or broken tissue.

Herein, we aim to concatenate keratin's performance as a multifunctional biological material, its use in the development of bioinspired structures, and its utilization in engineered systems. This review is organized as follows. The rest of the introduction summarizes keratin's general structure and properties as a basis for understanding the diversity of keratin-based systems. The next section, entitled “ Bioinspired Materials Based on Keratinous Systems ” highlights how the various structures found in keratin-based materials guide bioinspired designs across a broad range of functions (mechanical, lightweight, reversible adhesion, thermal, structural colors, and hydrophobicity). The third section, “ Keratin as a Material for Engineered Systems ” discusses how keratin's intrinsic material properties are harnessed for various engineering applications, focusing on biomaterials and fiber-reinforced composites. Although there are nearly twenty existing reviews of keratin and keratin-based materials, many of them focus on its structure and properties ( Bradbury, 1973 ; Chapman, 1969a ; Marshall et al., 1991 ; McKittrick et al., 2012 ; Norlén, 2006 ; Wang et al., 2016a ; Wang and Sullivan, 2017 ), use as a biomaterial for biomedical applications ( Donato and Mija, 2019 ; Feroz et al., 2020 ; Rouse and Van Dyke, 2010 ; Shavandi et al., 2017 ), or extraction techniques ( Chilakamarry et al., 2021 ; Donato and Mija, 2019 ; Feroz et al., 2020 ; Khosa and Ullah, 2013 ; Shavandi and Ali, 2019 ), and there are only a few reviews that acknowledge keratin-based bioinspired materials ( McKittrick et al., 2012 ; Wang et al., 2016a ) ( Table 1 ). None of the reviews that include bioinspiration are recent; much progress has been accomplished that warrants an updated review. This timely review illustrates how keratin obtains its vast range of functionalities from its structure and intrinsic properties and how these features are used to develop bioinspired and engineered materials. We conclude with recommendations on the future directions for keratin applications and bioinspired designs.

Summary of keratin review papers

Structure of keratin

The term keratin originates from the Greek word ‘kera,’ which means horn. Historically, keratin denoted proteins extracted from modifications of skin such as horns, claws, and hooves. However, with an increased understanding of its structural and chemical characteristics, keratin now refers to all intermediate filament (IF)-forming proteins with specific physicochemical properties that are produced in any vertebrate epithelium ( Bragulla and Homberger, 2009 ). These proteins form the bulk of cytoplasmic epithelial and epidermal appendageal structures (i.e., hair, wool, horns, hooves, and nails) ( Wang et al., 2016a ). They are also present inside cells as IFs, which provide structural stiffness, together with actin fibers and microtubules; we will not include this form in this review. This review will use the term “keratin” to describe this material at the nanoscale (macrofibrils) and below. In contrast, “keratinous material” will be used to describe the larger scale structures that are composed of these keratin fibers.

Keratins are broadly classified as having either α- or β-ultrastructures ( Figure 1 ). Typically, mammalian keratin is found in the α-keratin form, while avian and reptilian keratins are β-keratin types; however, one mammal, the pangolin, is known to have both α- and β-keratin domains in its scales ( Wang et al., 2016c ). Like all biological materials, both α- and β-keratinous materials form hierarchical structures with geometries ranging from the atomic scale to the macroscale, as shown in Figures 1 and ​ and2. 2 . Both α- and β-keratin are built from amino acids at the atomic level. In α-keratin, the amino acids form a right-handed α-helix secondary protein structure stabilized by hydrogen bonds ( Burkhard et al., 2001 ; Fraser et al., 1976 ; Pace and Scholtz, 1998 ; Rojas-Martínez et al., 2020 ; Singamneni et al., 2019 ). These protein structures, also referred to as polypeptide chains, are approximately 45 nm in length and form the basic building block of an IF at the sub-nanoscale. Two polypeptide chains twist together in a left-handed rotation to form a dimer, referred to as coiled-coil ( Crick, 1952 ). The dimers are also approximately 45 nm in length and have a diameter of ∼2 nm. It is believed that the coiled-coil structure increases the stability of the filament compared to a single α-helix ( Chou and Buehler, 2012 ). Terminal segments of the dimer constitute an amorphous head and a tail domain. Both the head and tail regions aid in the dimer's self-assembly. The two coiled-coil dimers then aggregate together to form a tetramer which bonds lengthwise (with disulfide bonds) to create protofilaments. Two protofilaments align to form a protofibril. Four protofibrils then connect to create an IF ( Bray et al., 2015 ). The IFs, which are ∼7 nm in diameter for α-keratin, are crystalline and are embedded in an amorphous keratin matrix. Crystalline IFs and the amorphous matrix form IF-matrix composites, which act as a basic structure for macrofibrils (∼400-500 nm in diameter). In literature, keratins are often considered short fiber-reinforced biopolymers consisting of an amorphous matrix and crystalline fibers (IFs)( McKittrick et al., 2012 ).

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Comparison between the atomic scale and sub-nanoscale of α- and β-keratin

Both α- and β-keratin, composed of amino acids, are similar at the atomic scale. The secondary protein structures are distinct for α (helix)- and β (sheet)-keratin at the sub-nanoscale. The subsequent polypeptide chains both form dimers which assemble into protofilaments and finally intermediate filaments. At the scale of IFs, both structures converge despite the differences in their diameters.

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Once α and β keratin form IFs, their general structure converges before splitting at larger length scales

The IFs embed in an amorphous matrix which then forms macrofibrils. These macrofibrils fill dead pancake-shaped keratinocyte cells, which stack on top of each other forming lamellae. From there, the structure of each keratinous system diverges to fulfill its specific function better. On the micro-, meso-, and macroscale, a vast range of designs and configurations are formed from the keratinous building blocks.

Similar to α-keratin, β-keratin is composed of amino acids at the atomic scale and has a comparable hierarchical order (dimer to protofilament to IF) at the sub-nanoscale. The most significant difference, compared with α-keratin, is that β-keratin has a different secondary protein structure characterized by pleated β-sheets ( Toni et al., 2007 ). In β-keratins, the antiparallel peptide chains are positioned side-by-side to form a rigid planar surface. These surfaces are slightly bent with respect to each other, creating a pleated arrangement ( McKittrick et al., 2012 ). The planarity of the peptide bond and the lateral hydrogen bonding accounts for the formation of the pleated sheet ( Fraser and Parry, 2011 ). Similarly to α-keratin, the β-sheet self-assembles into a dimer, which forms the basis of the distorted β-sheet (called a protofilament). Protofilaments align to form the β-keratin IF, which is ∼3 nm in diameter. For β-keratin, the terminal sections of the polypeptide proteins wrap around the filaments to form the amorphous matrix. Besides the differences between the α- and β-keratin at the sub-nanoscale, both keratin types form similar hierarchical structures up to the nanoscale ( Figure 2 ). At the microscale, keratinous materials' architecture diverges for different organisms to optimize their structures for their specialized functions.

At the nanoscale, the IFs are embedded in an amorphous matrix in both α- and β-keratins. This IF-matrix nanocomposite structure subsequently groups to form macrofibrils (∼400-500 nm in diameter) and then fibers (∼6 μm). Variations in the IF alignment, volume fraction, orientation, and matrix properties account for the wide range of mechanical properties of keratin-based structures. Keratinocytes are the once living cells that are filled with keratin fibers. Their formative boundaries encapsulate the orientation and can vary across organisms or locations within a specific organism. When stacked together, the keratinocytes form a layered structure at the microscale due to their inherent directional growth from the follicle. In some systems such as the horse hoof wall, woodpecker beak, pangolin scale, and bighorn sheep horn, the interface between neighboring keratin cells exhibits a wavy sutured morphology. Through their layered growth, keratin cells form laminated sheets. The hierarchical structure of many keratinous systems begins to diverge at this scale. This layered structure is a defining feature of keratin-based materials. The laminated sheets organize themselves into different arrangements at the mesoscale. For example, the laminated structure in some horns and hooves is characterized by embedded microtubules, whereas the lamellae in hair cuticles have an overlapping configuration. Even more so, at these larger length scales, some keratinous materials begin forming cellular solids such as the foamy centers of quills and feather shafts. The divergence of the structure at the meso and macroscales for each organism will be explained in greater detail in Section 2.

There are also morphological differences among different keratinocytes: in hair, they are elongated along the axis (one dimension much larger than the other two); in pangolin scales and many other places, they are pancake-shaped, with one dimension much smaller than the other two. There seems to be a preponderance of suture structures at the mesoscale. The surface of a cortical cell in human hair after tension exhibits a suture-like structure, which increases the contact area of cortical cells and therefore increases the adhesion between adjacent cells and decreases splitting of hair along the axis. This suture structure is also found in the pangolin scale. It has a width between 250 and 450 nm and creates an interlocking effect. This structure has been studied and generalized by the Ortiz group ( Li et al, 2011 , 2012 , 2013 ; Lin et al., 2014a , 2014b ). Figure 3 shows the suture structures in hair and pangolin scale.

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Intercellular suture structures are present on the surface of many keratinocytes

(A) Human hair. Reproduced with permission ( Yu et al., 2017a ). Copyright 2017, Elsevier.

(B) Pangolin scales. Reproduced with permission ( Wang et al., 2016c ). Copyright 2016, Elsevier.

To fully capture the hierarchical structure of keratin, computational models have been developed for each length scale. Starting with the fundamental building blocks of amino acids, these models aim to analyze the mechanical properties and arrangement of molecules in IFs at the sub-nanoscale ( Chou and Buehler, 2012 ; Qin et al, 2009 , 2011 ; Qin and Buehler, 2011 ). Chou and Buehler (2012) pioneered this effort by reconstructing heterodimers from an entire amino acid sequence of keratin proteins using molecular dynamics simulations. The geometric dimensions of the reconstructed dimer matched well with the experimental observations. Using this model, they compared keratin's mechanical properties with and without disulfide bonds and concluded that the disulfide bonds improve keratin's durability and strength ( Chou and Buehler, 2012 ). This feature is similar to the one in elastomers, where vulcanization introduces sulfur bonds between the chains and increases the performance markedly. Qin et al. (2009) , from the same Buehler group, discussed the hierarchical structure of IFs and analyzed each hierarchical level's influence on the IF's mechanical properties. They divided the hierarchical structure of IFs at the atomic scale and sub-nanoscale ( Figure 1 ) into additional eight hierarchical levels. Using molecular dynamics modeling, they concluded that each hierarchical level demonstrates a distinct deformation mechanism, which enables keratin to sustain prominent deformation at higher length scales (beyond the nanoscale) ( Qin et al., 2009 ). The dominant mechanisms at each hierarchical level and their description are summarized in Table 2 . In another paper, Chou et al. (2015) demonstrated how information from atomic-scale models could be utilized to predict human hair's mechanical properties at the mesoscopic scale through a bottom-up approach ( Chou et al., 2015 ).

Key hierarchical levels and their corresponding mechanisms

Mechanical properties of keratin

The polymeric nature of keratin lends itself to a wide range of mechanical properties that vary according to its amino acid composition, structure, and hydration level ( Bertram and Gosline, 1987 ; Fraser and Parry, 2014 ; Greenberg and Fudge, 2013 ; McKittrick et al., 2012 ; Wang et al., 2016a ). The amino acid sequence and corresponding residues dictate the availability of disulfide bridges. The amino acid cysteine has a thiol group which allows for a covalently bonded di-sulfide bond to be formed with another cysteine further along the chain and creates a fold in the protein. Chou and Buehler (2012) showed that keratin's hardness is strongly correlated with the density of sulfur cross-links ( Chou and Buehler, 2012 ). A low amount of sulfur indicates soft keratins (outer layer of skin, i.e., stratum corneum). In contrast, a high amount of sulfur leads to hard keratins (e.g., hair, nails, feathers, hooves) ( Chou and Buehler, 2012 ; Parbhu et al., 1999 ; Smack et al., 1994 ).

Based on the structural arrangement described in the previous section, keratin's amino acid chains can either curl into helices (α-configuration) or bond side-by-side into pleated sheets (β-configuration). The molecular arrangement associated with the alignment of IFs directly influences the mechanical properties of keratinous materials ( McKittrick et al., 2012 ). The stress-strain curve of a typical α-keratinous material consists of three distinct regions: linear elastic region, yield region, and post-yield region, as shown in Figure 4 A. Figure 4 A decomposes the contributions of both the IFs and the matrix to the properties of α-keratin fibers. The linear elastic region extends approximately up to a 2% strain. In this region, the stress increases linearly with an increase in strain ( Chapman, 1969b ). Beyond 2% strain, the keratinous material enters the yield region in which it reaches critical stress beyond which the coiled-coil region of the α-keratin helices begins to unravel into the β-pleated sheet structure exhibited by β-keratin ( Cao, 2002 ; Kreplak et al., 2004 ; Paquin and Colomban, 2007 ). As a result, the stress-strain curve exhibits a large plateau. X-ray diffraction studies have shown that microfibrils open at various points and increase in length during the conversion ( Bendit, 1957 ). However, atomic-scale simulations have demonstrated that the structure of the dimer assembles in a specific sequence ( Chou and Buehler, 2012 ). The low increment in stress in the yield region can be explained by the Ciferri model ( Ciferri, 1963 ). Ciferri proposed that the low increment in stress is due to thermodynamic equilibrium existing between α-and β-structures. The α- and β-keratins coexist in equilibrium at a constant stress value dependent on temperature but not on each state's relative quantities. The plateau region exists up to ∼30% strain, beyond which the material enters the post-yield region, where the stress again increases with an increase in strain. The rise in stress can be attributed to the coupling between the matrix and IFs. Even though the α-keratin continues to convert to β-keratin until 70-80% of strain, the matrix starts resisting deformation at ∼30% strain and thus begins to bear additional stress. As a result, a sharp rise in tangent modulus is observed ( Ciferri, 1963 ).

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Mechanical properties of keratin and keratinous materials

(A) Idealized stress-strain curve of α-keratin showing three distinct regions. This is a representative curve and does not take into account factors like viscoelasticity or structural deformation mechanisms. Still, it does highlight the plateau yield region and the range of these three phases of deformation. Reproduced with permission ( McKittrick et al., 2012 ). Copyright 2012, Springer.

(B) Spring and dashpot configuration of the two-phase model that is used to incorporate the hydration-induced viscoelasticity of the amorphous matrix.

(C) Tensile stress-strain curves of bird feathers and claws test at different humidities at a strain rate of 0.11 min −1 . Adapted with permission ( Taylor et al., 2004 ). Copyright 2004, Springer.

(D) Effect of strain rate on biopolymers' strength (whale baleen, hair, pangolin) and the synthetic polymer PMMA. Reproduced with permission ( Wang et al., 2019 ). Copyright 2018, Wiley.

Several attempts have been made to capture the mechanical properties of keratin analytically. The most notable ones are the two-phase model proposed by Feughelman ( Feughelman, 1959 ) and the Hearle-Chapman model ( Chapman, 1969b ; Hearle, 1967 ). The initial two-phase model of Feughelman was later modified to incorporate additional features of keratin. In this revised model, the keratinous material comprises two phases: C and M. Phase C denotes long water-impenetrable and relatively rigid cylindrical rods. These rods are embedded in a water-absorbing matrix called phase M. Phase C represents a coiled-coil part of the polypeptide chain in α-keratin. This phase has lower sulfur content to interact with water. Phase M consists of non-helical parts of α-keratin (like its head and tail) and matrix structure surrounding polypeptide chains. These parts have higher sulfur content and can absorb water, giving rise to viscoelastic behavior in keratin. According to this model, the initial region (named the linear elastic region by earlier, less complex studies) of the stress-strain curve for α-keratin can be represented by a spring and dash-pot model ( Figure 4 B) where a spring (with a spring constant of E f ) is in parallel with another spring (with a spring constant, E M ) and dashpot (with viscosity, η). The spring constant, E f , represents Young's modulus of the crystalline phase and therefore does not depend on moisture content. The E M and η represent the properties of a viscoelastic amorphous matrix dependent on moisture and temperature. As evident from the spring-dashpot model, the non-linear viscoelastic behavior of keratin in the Hookean region is due to the matrix phase described as a weak “gel” structure ( Feughelman, 2002 ). As the gel structure is extended at a fixed rate, the bonds progressively break down. If the extension is ceased, the broken bonds re-form rapidly in equilibrium.

In the yield region, the α-helices in the crystalline phase C are extended to the fiber structure's total length. As a result, they start unfolding to β-units at a nearly constant stress, governed by a thermodynamic equilibrium between α- and β-units. Most of the force applied to the keratinous material in this region is resisted by the IFs, whereas phase M resists only a small force that is nearly constant. The viscosity contributes to the time constant for the relaxation and provides resistance to folding and unfolding of α-helices.

When the α-helices transition to β-pleated sheets in the yield region, they extend in length. Figure 5 A shows a full period of the α-helical structure consisting of the atomic sequence ( -CCNCCNCCNCC-) ; its length is 0.52 nm. When this helix is fully rectified and extended ( Figure 5 B), its length becomes 1.39 nm. However, the assembly of polypeptides is such that a folded β-pleated sheet is formed; this reduces the length to 1.2 nm. Thus, the nominal strain of the α to β be calculated and is equal to 1.34. However, it is rarely achieved experimentally, and other processes are thought to take place.

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Full period (one rotation, corresponding to -CCNCCNCCNCC-) for α-helix (0.52 nm) and corresponding distance for β-pleated sheet (1.2 nm)

The stretched β configuration with the same chain (-CCNCCNCCNCC-) has a length of 1.39 nm. The formation of pleats reduces the length to 1.2 nm. The theoretical strain corresponding to full transformation is equal to 1.34; this is seldom achieved in real cases. Reproduced with permission ( Yu et al., 2017a ). Copyright 2017, Elsevier.

At a larger spatial scale, the IFs parallel to each other start moving closer together, jamming the still unfolding α-helices against the matrix phase, which consists of globular matrix proteins. Owing to the increase in length when the α-helices transition to β-pleated and the jamming of proteins in the matrix, further extension of the material distorts matrix proteins. As a result, the matrix starts carrying more load resulting in an increase in stress with strain. The above is the essence of the Feughelman model.

Chapman ( Chapman, 1969b ) and Hearle et al. ( Hearle, 1967 ) independently extended the two-phase model to explain the zonal unfolding of α-helices in microfibrils by considering the effect of mechanical coupling between the fibril and matrix. They assumed that the single fibril is of infinite length. The matrix never enters the yield region and therefore behaves elastically as it bears only a small portion of the total force. Based on this model, they derived the equations to predict stresses and strains in different regions as shown below. We use subscripts f and M for the fiber and matrix, respectively.

Hookean region: E M ≪ E f thus, the stress is taken by the fibril

At the yield point

Yield region: (once the transition of α-fibrils has started)

End of post yield region

where 1 E p = 1 E f + 1 E M − ( σ c − σ e ) / ( 2 ε b )

E M is Young's modulus of the matrix, E f is an initial fibril modulus, σ and ε are the total stress and strain, respectively, in the material. σ c is equal to the critical stress at which the unfolding of α fibrils begins. ε 2 is the strain due to unfolding of α fibrils, σ e is the equilibrium stress for the transition between α to β fibrils, ε M is the strain in the matrix, ε b is the strain associated with α to β transition, and E p is the effective modulus in a post-yield region. The detailed derivation for the above equations is given in Hearle and Chapman ( Chapman, 1969b ; Hearle, 1967 ).

In general, α-keratin has a high tensile fracture strain, primarily due to the stretching and sliding of the polymer chains across many length scales. The hagfish slime threads have the highest tensile breaking strain of 2.2 when tested in seawater ( Fudge et al., 2003 ). Despite large tensile breaking strains, there are significant variations in tensile strength across species due to structural orientation, hydration, and composition ( Fudge et al., 2003 ). The tensile strength ranges from 2 MPa in the stratum corneum to 225 MPa in human hair to 530 MPa in the hagfish's dry slime threads. Mechanical properties of keratinous materials also depend on the orientation and volume fraction of IFs: greater alignment of the IFs results in a higher tensile strength along the alignment direction. Thus, the tensile strength of human hair (where all the IFs in the cortex are aligned with the hair axis) is higher than that of human nails (where there are three layers in which the IFs are oriented at 90° to each other).

The degree of hydration dramatically influences the mechanical properties of keratin. Increasing humidity and water content decreases the stiffness, strength, and hardness ( Collins et al., 1998 ; Johnson et al., 2017 ; Liu et al., 2016a ; Wang et al., 2016a ) This behavior, summarized in Table 3 , is attributed to the interaction of water molecules with the amorphous matrix, which breaks stabilizing hydrogen bonds and increases the mobility of the fibers within the matrix ( Winegard and Fudge, 2010 ). In equine hoofs, Young's modulus drops an order of magnitude between dry and hydrated conditions ( Bertram and Gosline, 1987 ; Kasapi and Gosline, 1999 ). This increase in ductility in hydrated keratinous samples is associated with a higher tensile strain but lower tensile stress. Thus, hydration has a drastic effect on strength. The feather, for example, sees its tensile strength more than halved from 221 MPa to just 106 MPa when placed in 0% relative humidity (RH) environment vs. 100% RH environment ( Taylor et al., 2004 ). These trends can be seen in Figure 4 C, which shows the stress-strain curves of bird feathers (rachis) and claws under tension at different relative humidities. Additionally, the pangolin scale has been shown to exhibit a decrease in hardness with hydration, from 314 MPa to 148 MPa in dry and hydrated states, respectively ( Liu et al., 2016a ). Other systems like whale baleen, porcupine quill, horn, and claws also see drastic reductions in strength with increasing hydration.

Mechanical properties of keratinous systems at various humidity levels

∗% RH = % Relative humidity, perp. = perpendicular to longitudinal axis of the tubules, para. = parallel to longitudinal axis of the tubules.

There are apparent variations in the mechanical properties of different keratinous systems. For example, the hoof, which has reinforced tubules, exhibits Young's modulus of 14.6 GPa at 0% RH, more than three times that of fingernails, claws, and feathers under the same humidity conditions. Even keratinous materials found in similar organisms, such feathers and claws, have noticeably different mechanical behaviors. These variations can also be observed in Figure 4 C. These differences can result from deviations in both chemical composition (i.e., mineralization, degree of crystallinity, etc.) or structure (porosity, lamellar arrangement, fiber orientation, etc.)

Keratin is known to be highly strain-rate sensitive, which is related to its viscoelasticity and viscoplasticity, i.e., its time-dependent response ( Yu et al, 2017a , 2017b ). This is a typical behavior of polymers. Figure 4 D shows the strength ( σ ) versus strain rates ( ε ˙ ) on a log-log scale for whale baleen, hair, pangolin scales, and a synthetic polymer (polymethyl methacrylate [PMMA]); the similarity is evident. The strain rate sensitivities “m” (defined as d ( l o g σ ) d ( l o g ε ˙ ) ) for biological materials (hair, pangolin, whale baleen) are comparable to those of PMMA, a synthetic polymer. In the case of whale baleen, the strain-rate sensitivity of the dry samples (m ≈ 0.02–0.03) is significantly lower than that of the hydrated ones (m ≈ 0.09–0.11). This difference is attributed to the hydrated specimens' increased viscosity, enabled by the water molecules penetrating the amorphous matrix and plasticizing it. In the dry specimens, the effect of the mineral phase becomes stronger.

The general trend for keratinous materials is that increasing strain rate increases stiffness and strength, while decreasing the breaking strain ( Johnson et al., 2017 ; Kasapi and Gosline, 1996 ; Seki et al., 2005 ; Song et al., 2009 ). Thus, most keratin materials undergo an elastic to ductile-plastic to brittle transition with an increasing strain rate, as was shown for the toucan rhamphotheca ( Seki et al., 2005 ) and pangolin scales ( Wang et al., 2016c ). This rate-dependent behavior has important implications for impact resistance, suggesting that these materials can withstand greater stresses under dynamic conditions and have different failure mechanisms than quasi-static conditions. The embrittlement at high strain rates is an important consideration.

Keratin is also one of the toughest biological materials, as seen in Figure 6 ( Wang et al., 2016a ). This characteristic is due primarily to its hierarchical structure. As demonstrated by Qin et al. (2009) , different hierarchical levels can undergo distinct deformations that enable keratin to absorb larger amounts of energy before failure ( Qin et al., 2009 ). The matrix is primarily responsible for distributing the applied loads during large deformations, while the fibers carry the most load and serve to arrest cracks. Some keratinous materials have optimized mesoscale features, such as tubules in horns and equine hooves, which enhance the material's toughness. Due to fiber orientation, fiber concentration, and the presence of features like tubules along a specific direction, toughness is typically found to be anisotropic ( Bertram and Gosline, 1986 ).

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Ashby diagram demonstrating toughness vs. modulus for different biological material

Reproduced with permission ( Wang et al., 2016b ). Copyright 2016, Elsevier.

Hydration-induced shape recovery

Keratin systems often function as protective layers which undergo significant deformation. Many of these systems are permanent and cannot remodel or self-heal through biological processes after experiencing considerable deformation, such as in the bighorn sheep horn ( Huang et al., 2019b ), feathers ( Liu et al., 2015b ; Sullivan et al., 2018 ), and pangolin scales ( Liu et al., 2016b ). A solution to this lack of regenerative capacity is keratin's ability to undergo hydration-assisted shape recovery. This phenomenon was discovered by Liu et al. (2015b) , who observed 98% shape recovery in compressed peacock tail feathers after seven cycles of deformation to over 90% strain ( Liu et al., 2015b ). After the keratin is deformed plastically, the recovery process involves water infiltrating the amorphous keratin matrix, causing swelling, which forces the deformed crystalline regions of the IFs to regain their initial shape by breaking and reforming hydrogen bonds ( Huang et al., 2019b ). Also, the feather shaft was shown to have hydration-assisted shape and strength recovery. The feather shaft was subjected to bending and then allowed to soak in water for 24 hr, and after one cycle, it was found to recover its strength by ∼80% ( Sullivan et al., 2018 ). The mechanism proposed by Sullivan et al. (2018) for the feather is shown in Figure 7 ( Sullivan et al., 2018 ). The Bighorn sheep horn was also shown to recover its shape by soaking in water after severe compression of 50% strain which was further assisted by the hollow tubules ( Huang et al., 2019b ). In a similar study by Liu et al. (2016b) , the pangolin scale was shown to have hydration-assisted strength recovery after indentation, which simulated penetration-induced injury by a predator. The self-healing was attributed to the swelling of the keratin-based material allowing for an increase in flexibility of keratin fibers to reorientate and straighten ( Liu et al., 2016b ).

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Hydration-induced mechanical reversiblilty is a common trait amongst keratinous systems

Reversible deformation of the feather shaft induced by hydration; top: restraightening of a deformed feather with hydration and recovery of its initial shape; bottom: sequence of events as the IF-amorphous matrix composite is first deformed and then hydrated.

Adapted with permission ( Quan et al., 2021 ). Copyright 2021, Nature.

The top sequence of Figure 7 shows the gradual restraightening of the feather shaft as it is hydrated. Plastic deformation causes permanent deformation of the amorphous matrix (bottom sequence), which is weaker than the IFs. The IFs undergo buckling on the compression side. Upon hydration, water molecules penetrate the amorphous matrix and cause swelling, which forces the crystalline IFs to straighten and realign. Upon drying, the matrix shrinks again, and the original configuration is established. These studies show that hydration actuates shape recovery in α -keratin and β -keratin, which is not surprising as both keratins have similar structures involving crystalline IFs embedded in an amorphous matrix.

Thermal properties

Another common function of keratin is to serve as a thermal insulating barrier in hair, wool, fur, and feathers, to name a few. Often the goal of these systems is to trap air pockets within the insulating layer. This method is very effective since air has an extremely low thermal conductivity of just ∼0.0264 Wm −1 K −1 ( Mao and Russell, 2007 ). As noted previously, the self-assembly process of natural keratinous materials has afforded some organisms with precisely controlled meso-, micro-, and nanostructures. For thermal insulation, this ability has been utilized to generate lightweight systems that trap significant amounts of air with minimal material. Note that keratin by itself has a low thermal conductivity of just 0.19 Wm −1 K −1 . However, when arranged into low-density wool, the combined thermal conductivity is reduced to 0.03 W m −1 K −1 ( Mao and Russell, 2007 ). Nature's ability to produce these intricate structures in abundance has made certain keratinous systems like feathers, wool, and fur some of humanity's most valuable thermal insulators to date. In humans, bipedalism concentrates exposure from the sun to the head, and this is exactly where capillarity is highest. The remainder of the body is only covered by vestigial hair, and this enables an increase in sweat glands, which enhances the ability of the body to regulate the temperature and has helped humans to develop an amazing ability to run for extended distances.

Bioinspired materials based on keratinous systems

Keratin is one of the most essential biopolymers found in nature, appearing in the integument of many vertebrates, as discussed in Section 1 . Keratinous materials are especially intriguing due to their hierarchical structures, which vary widely across organisms and are found in a broad range of morphologies that are tuned for their specific functions. To show that these configurations give rise to the high performance of natural keratinous materials and can be a source of bioinspiration, these naturally occurring geometries are replicated in engineered materials by simplifying integral designs and scaling them to more appropriate sizes for processing and mechanical testing. Additionally, many of these studies rely on numerical and analytical models to better understand the mechanical behavior and deformation mechanisms of these bioinspired systems. This section will review these efforts through a bioinspired lens, focusing on how keratin-based systems and their structures achieve diverse functions.

The many functions of keratinous materials, shown in Figure 8 , will lay the framework for reviewing their associated bioinspired materials. Table 4 highlights some common examples of systems for each function and their relevant structures.

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Keratin provides many functions in nature

In the following section, bioinspired designs based on keratinous systems will be broken down into the classifications shown in this figure.

Keratin biological systems, their principal functions, and related structures

Mechanical applications

Keratin-based materials are frequently utilized in nature as structural load-bearing components that provide protection and withstand high impact forces. Keratinous systems perform admirably under such diverse mechanical demands, even compared to some of the most advanced engineered materials ( Lee et al., 2011 ). One reason is that keratin's mechanical properties can be tuned by hydration, providing a stiff (∼10 GPa) load-bearing material when dry or a ductile rubbery material when fully hydrated (∼0.1 GPa) ( Bertram and Gosline, 1987 ; Collins et al., 1998 ; Huang et al., 2019a ; Zhang et al., 2007 ). Another reason is that keratin takes on the form of a wide range of structures with intricate geometrical features at multiple length scales that synergistically lead to high mechanical performance. This subsection will review keratinous systems with remarkable mechanical properties and instances where their structural features have been used as inspiration for synthetic materials.

One of the most common keratinous systems that has been studied for bioinspiration is the hoof wall of horses and bovines ( Bertram and Gosline, 1987 ; Douglas et al., 1996 ; Huang et al., 2019a ; Kasapi and Gosline, 1996 , 1997 , 1999 ; Li et al., 2010 ). Horse hooves hit the ground at a speed of ∼8m/s ( Parsons et al., 2011 ) and can experience impact forces of ∼16.1 N/kg (deceleration of ∼56 g) ( Lanovaz et al., 1998 ; Setterbo et al., 2009 ). The hoof wall is composed of dead keratinocyte cells that cannot repair themselves yet can survive many regular impacts. This characteristic has made the hoof wall a prime candidate for designing bioinspired materials with high impact resistance and energy absorption capabilities. The hoof wall has an intricate hierarchical structure, depicted in Figure 9 A, that has been shown to augment keratin's bulk properties. At the mesoscale, the hoof has hollow cavities (∼40 micrometers in diameter) surrounded by relatively stiff elliptical regions (with a major axis of ∼200 micrometers and a minor axis of ∼100 micrometers) that run parallel to the surface of the hoof wall ( Huang et al., 2019a ). These tubules are embedded in a lamellar matrix composed of stacked, microscale, pancake-shaped cells (keratinocytes). These two geometries work in concert to provide the hoof with high fracture control ( Bertram and Gosline, 1986 ; Kasapi and Gosline, 1997 , 1999 ) and impact toughness ( Huang et al., 2019a ; Kasapi and Gosline, 1996 ).

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Horse hooves have been a great source of inspiration for tough material designs with fracture control properties

(A) A schematic of the horse hoof's micro- and meso-structure showing reinforced tubules embedded in layers of pancake-shaped cells. These cells are filled with IFs. Reproduced with permission ( Kasapi and Gosline, 1999 ). Copyright 1999, Company of Biologists.

(B) Schematic showing different epoxy arrangements infiltrated PLA samples inspired by the hoof's layered structure.

(C) Crack propagation through flat layered samples before peak stress (top left), at peak stress (bottom left), during failure (top right), and after failure (bottom left).

(D) Failure pattern of zigzag layered samples.

(E) Schematic showing how cracks interact with a jagged layered structure.

(F) Force-extension curve (left) and energy absorption-extension curve (right) of samples with layered structures of different angles. Reproduced with permission ( Rice and Tan, 2019 ). Copyright 2019, Elsevier Ltd.

Rice and Tan ( Rice and Tan, 2019 ) drew inspiration from the lamellar structure found in the horse hoof's intertubular matrix to design improved composite materials. In hooves, the lamellar structure has shown strong retardation of fracture propagation by causing cracks to divert along the interlayer interfaces away from the living tissue at the hoof's interior ( Bertram and Gosline, 1986 ; Kasapi and Gosline, 1997 , 1999 ). To harness this fracture control mechanism for engineered composites, Rice and Tan ( Rice and Tan, 2019 ) manufactured a layered material with alternating soft (ductile) and stiff (brittle) regions, composed of 3D printed polylactic acid (PLA) layers infiltrated with epoxy or resin, as shown in Figure 9 B. Their goal was to demonstrate that this bioinspired structure could successfully be utilized in synthetic composites and explore the effects of layer thickness, layer angle, and notch location on crack propagation. Single-edge notched bending tests on monolithic samples of resin, epoxy, and PLA showed that cracks traveled directly through the material with negligible deflection. Similar results were found for samples that contained flat lamellae and thin PLA layers, as shown in Figure 9 C. The shear stress near the crack tip initiates debonding between the soft and hard layers; this gives rise to a crack-deflection mechanism similar to those found in hooves. Maximum shear stress develops at 45° to the original notch tip, while the lowest shear stress occurs at 90° to the notch. So, flat layers (layers oriented at a 90° angle to the notch like those in Figure 9 C) experience the least debonding and exhibit minimal crack deflection. However, these samples have the benefit of being very stiff and require high peak forces to failure. Figure 9 D shows how the introduction of angles into the lamellar structure can affect the crack path through the material. As the angle of the layers relative to the crack tip nears 45°, more shear stress builds up between the soft and hard layers causing the crack to deflect along the interface of the two materials. Figure 9 E compares the force-extension curves of samples with layer angles of 60°, 70°, and 90°. Layers at 60° begin to debond at very low forces, while layers at 90° do not exhibit any debonding. Lamellar structures oriented at 70° are an ideal compromise, providing some stiffness and resistance to fracture before absorbing energy by debonding along the zigzag interface. Figure 9 E also shows each model's energy absorption curves and indicates that after 14 mm of extension, the 70° model absorbs more energy than the traditional 90° model. One final factor that was found to be very important for this configuration is the layer thickness. When the ductile PLA layer was too thin, the crack fractured through it, and minimal deflection was observed. Higher peak forces and energy absorption were found for thicker ductile layers.

Several researchers have also explored the characteristic tubular structures found in hooves. Wang et al. (2020a) 3D printed simplified tubular arrangements based on bovine hooves ( Wang et al., 2020a ). The tubules were modeled as hollow hexagonal prisms with varying angles that are inspired by the different angles of the intertubular layers found in the hoof. Three different configurations, shown in Figure 10 A, were prepared for single-edge notched bending tests. The first model (G1) had no internal structure and was composed of bulk PLA. The second model (G2) had three rows of tubules, each offset from the previous row by 22.5°. The final model (G3) had the same structure as G2, but the tubules had a deflection of 15°. The introduction of tubules significantly improved the mechanical performance of the material with an increase of 39% in K IC and 55% in G IC from the G1 model to the G2 configuration. Figure 10 A shows the K IC and G IC results normalized by volume, which indicates the superiority of the G2 design. The samples with tubular elements had a confined fracture pattern, which was given credit for the enhanced toughness and energy absorption of the G2 and G3 models.

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Tubular structures in hooves have attracted significant attention for bioinspired designs

(A) Schematic of different tubular arrangements modeled after the hoof with tubules (yellow) represented as hexagonal prisms (left). Graph of normalized K IC  for each model (middle), and representative images of the damage zone for each model after testing (right). Reproduced with permission ( Wang et al., 2020a ). Copyright 2020, Elsevier B.V.

(B) Schematic of different models with increasing complexity culminating in double-phase tubules embedded in a layered structure (top). Images (middle) and optical micrographs (bottom) of the different samples after drop tower tests where the impact energy was 100KJ/m 2 . Open Access ( Huang, 2018 ).

Huang et al. (2018) combined reinforced tubular and lamellar structures to understand the impact-resistant synergy that these arrangements provide. Four different models were created using a multi-material 3D printer. These can be seen in Figure 10 B. Single-phase samples were made out of stiff and brittle VeroClear. A softer, more ductile polymer called TangoBlackPlus was used to print the black, interlayer regions on the models. Both of these phases are proprietary materials produced by Stratasys, Ltd. Each sample was impacted with 100 kJ/m 2 of energy, and the results are shown in Figure 10 B. The single-phase samples failed and fractured into many pieces, while the other three samples all remained intact; only the double-phase tubule reinforced sample prevented cracking from reaching the sample's corners. Optical microscopy images of the damaged samples are shown at the bottom of Figure 10 B, where the tubules' crack arresting capabilities can be observed ( Huang, 2018 ).

Ma et al. (2020) formed tubular structures inspired by the equine hoof wall's architecture to achieve outstanding crashworthiness. As shown in Figure 11 , they modified traditional square tubes by replacing the vertices with the unit geometrical structure. The conception of these structures was inspired by the tubular geometry present in the keratinous equine hoof wall. They also modified the side walls to corrugated plates, inspired by secondary epidermal lamella in an inner lamellar layer. The samples were manufactured using the aluminum alloy AA6061-0. Ma et al. (2020) demonstrated that the hoof-inspired geometry (hoof-wall inspired corrugated tube [HCT]) could significantly improve crashworthiness using compression tests and finite elemental analysis. The HCT provided a 269% increase in energy absorption and 124% increase in specific energy absorption over traditional square tubules in compression testing ( Ma et al., 2020 ).

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Designs containing structures inspired by the hoof wall have been fabricated to create materials with improved crashworthiness

The top two rows of images show the naturally occurring horse hoof, while the bottom row shows designs of increasing complexity that incorporate the tubular and lamellar microstructure of the keratinous hoof sheath. Reproduced with permission ( Ma et al., 2020 ). Copyright 2020, Elsevier Ltd.

Horns have a very similar structure to hooves with hollow tubular elements embedded in lamellar stacks of flat cells. However, the tubules in horns are perpendicular to loading at the impact zone and lack the reinforced region surrounding the tubules that is found in hooves. Figure 12 A shows SEM images of the tubular and lamellar structure of the bighorn sheep horn alongside 3D printed models of the horn, including a single-phase block of stiff VeroClear with and without an array of tubules and two-phase lamellar structures (the second phase being ductile TangoBlackPlus). Figure 12 B shows how the bioinspired models are compared to horn samples under compression. When samples were compressed with the loading axis parallel to the lamellae, they showed much lower strength. This behavior is due to delamination between the soft and hard phases, similar to the response found in horns. When samples were compressed perpendicular to the tubules, the hollow cavities collapse, leading to a slight decrease in stiffness and strength but an increase in plastic deformation and final compressive strain. Again, this performance mirrored that of real horns, suggesting that this structure could also have good energy absorption capacity under impact ( Huang, 2018 ).

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Bighorn sheep horns can endure tremendous impacts and have been the muse for several impact-resistant bioinspired designs

(A) The horn's structure (top) with SEM images of its tubular and layered structure. Schematics and images of bioinspired designs with unreinforced tubules embedded in a layered configuration. The layers relative to the tubules' orientation are the opposite of the hooves while the orientation of the tubules to the impact direction is also reversed.

(B) Stress-strain curves of the horn and bioinspired samples in different orientations.

(C) Images of failure mechanisms of bioinspired samples when compressed in different orientations with respect to print direction. Open Access ( Huang, 2018 ).

However, 3D printed polymer models of keratinous structures have been limited by their inability to capture these systems' full complexity and mechanical functionality. While the shape of the stress-strain curves of the printed samples and horn samples are similar, Figure 12 C shows that their failure mechanisms are quite different. For example, the 3D printed samples developed stress concentrations around the tubules leading to cracking when compressed perpendicular to the tubules. This behavior was not observed in the horn, which was able to distribute stress more uniformly ( Huang et al., 2017 ). Also, when horn samples were compressed parallel to the tubules, tubule buckling was observed. In the 3D printed samples, it was the lamellae that buckled rather than the tubules. These differences are likely due to disparities in material properties between printed and natural samples, the lack of lower-order hierarchical structure in the printed models, and processing restrictions that create weak interfaces and residual stress in 3D printed components. The print direction additionally influences the mechanical response. While 3D printing biomimetic structures have huge potential, this example underscores some of this technique's limitations ( Huang et al., 2017 ).

Huang et al. (2018) also tested the recoverability of compressed 3D printed samples inspired by bighorn sheep horns. Dynamic and quasi-static recovery tests on horn samples showed that, when exposed to water, keratinous materials can regain much of their initial shape after compression. In keratin, this process is highly dependent on hydration, which disrupts the hydrogen bonds within and between the macromolecular chains and allows them to be reformed in a recovered position once the load is released. A similar process can be achieved in synthetic polymers by raising the specimen's temperature over the glass transition temperature. After being compressed to 50% strain, the 3D printed samples were exposed to 62°C for 15 min. Similar to the horn results, damage from compression in the longitudinal and transverse directions was irrecoverable due to lamellae buckling and shear band formation. However, in the radial direction, much of the structure and the stress-strain curve was recovered in subsequent compression cycles, suggesting that keratinous materials can also provide a structural blueprint for shape recovery materials ( Huang, 2018 ).

Kassar et al. (2016) produced foam liner material for motorcycle helmets inspired by the microstructure of horns. Helmets and horns both have an outer structure that is mainly responsible for energy absorption during impact. Soft inner tissue that distributes the load increases the deceleration distance and thus protects the head. Following a similar principle, they designed foams with varying tubular porosity. As observed in horn structures, the tubules' porosity was varied from 0%, near the head, to 10% in the middle and ∼30% on the outer shell ( Kassar et al., 2016 ). This spatial change in porosity is a classic example of a gradient structure, one of the hallmarks of biological materials ( Liu et al., 2017 ). Figure 13 shows the bioinspired design. To assess the design, modified drop tower tests according to “United Nations Economic Commission for Europe Standard” for motorcycle helmets ECE 22.05 were performed using the foam manufactured by EPS material. The design was able to meet safety thresholds far below the limits stipulated by the ECE 22.05 motorbike helmet testing standard.

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Bighorn sheep horns absorb tremendous impacts in nature, so researchers envision helmets inspired by the horn's microstructure

(A) Visualization of the hierarchical structure with an emphasis on the microstructure of the bighorn sheep horn.

(B) Conception of a helmet with a gradient in tubular porosity between the interior and exterior.

(C) Cross section of the protective tubular region showing a variation in tubule size through the helmet's thickness. Reprinted with permission ( Kassar et al., 2016 ).

The above efforts have taken bioinspiration from microscale structural elements of hooves and horns. However, Sun et al. (2014) designed rear under-run protection devices (RUPDs) for heavy trucks inspired by the macroscale geometry of the sheep horn. The RUPD prevents the entry of small-scale vehicles under the rear end of the heavy truck. The design was analyzed using a finite element analysis. The authors concluded that, compared to the normal RUPD of the same thickness, the bioinspired design could provide better protection when rear-end accidents happen; this is due to its enhanced energy absorption and structural strength ( Sun et al., 2014 ). Zhang et al. (2008) took inspiration from buffalo hooves to design impellers for a paddy field. They studied the buffalo hooves' curvature that allows them to maneuver through the field with relative ease. The impeller designed with similar curvature has a 38% increase in pull force and was more efficient than standard blades ( Zhang et al., 2008 ).

Baleen is the filter-feeding system found in the oral cavity of baleen whales, some of the largest animals on the planet, and is composed of highly mineralized keratin. To withstand the forces associated with filter-feeding, some whales have evolved baleen with complex structures that provide remarkable fracture toughness. The baleen plates contain a tubular sandwich structure that can be seen in Figure 14 A. The tubular region has a structure that is reminiscent of hooves but has a much higher mineral content that arises from hydroxyapatite nanocrystals embedded among the keratin IFs. The sandwich structure, composed of a solid shell around the tubular zone, provides high flexural stiffness and strength relative to the material's weight. Much like the tubule lamellae found in hooves, the concentric layered arrangement around the hollow cavities serves to deflect cracks and increase fracture toughness. This structure is highly anisotropic. The differences that arise from different loading directions can be seen in Figure 14 B. Loading parallel to the tubules gives higher Elastic modulus but less ductility than the loading perpendicular to the tubules. This anisotropy has a profound effect on fracture toughness ( Wang et al., 2019 ).

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Whale baleen is a part of the filter-feeding apparatus of baleen whales and is able to withstand high stresses and impacts from fish that get sucked into the whale's mouth. Bioinspired models have shown that the structure of the baleen helps endow it with admirable properties

(A) Image of a cross section of whale baleen showing the tubule layer sandwiched between a solid shell of keratin.

(B) Stress-strain curves of the baleen in each orientation showing significant differences in response based on loading direction. Stress-strain curves of the bioinspired models, indicating the design's superiority with all of the features incorporated in tandem in model iv. Reproduced with permission ( Wang et al., 2019 ). Copyright 2018, WILEY-VCH Verlag GmbH & Co. KGaA.

Four 3D printed models were fabricated to investigate the role that each of these features plays in the baleen. The most complex model (model IV) printed using three different materials most closely represents baleen. The mineralized lamellae were simulated using a stiff polymer and the matrix using a ductile polymer, while a polymer of intermediate stiffness represented the unmineralized lamellae filaments. Each successive model adds a new design element. Model I contains just a sandwich structure of soft material between two stiff layers; model II adds the concentric filament structure; model III includes the hollow cavity at the center of tubules. Model IV combines all of these features with the stiff lamellar rings shown in yellow. The addition of filaments raised the samples' stiffness, while the hollow cavities slightly decreased the sample strength at strain rates of 0.28 s −1 and 10 −4 s −1 but increased it at strain rates of 10 −2 s −1 . The addition of the stiff lamellar rings unsurprisingly increased the models' stiffness and strength and led to significantly more strain-rate stiffening and strengthening. These phenomena were also observed in the natural baleen. Wang et al. (2019) concluded that model IV provides the best mechanical performance showing that the features found in keratinous whale baleen can be utilized as beneficial structural design elements ( Wang et al., 2019 ).

In summary, bioinspired research on mechanical keratinous tissue has focused on several features: tubules (as found in the hoof, horn, and baleen), lamellar structures (found in all keratinous materials), and macroscale geometry (like hoof curvature or horn shape). When composite materials incorporate tubules or lamellae, they find improved fracture toughness due to crack interactions at these structures' interfaces. Similarly, macroscale geometries are practical but largely unexplored avenues of inspiration for specific functions like impellers or bumpers.

Thermal insulation

Keratinous systems are some of nature's best insulation by virtue of their elaborate structures that trap air. Many synthetic fibers are more inherently resistant to heat transfer. However, with their hierarchy of air-trapping features, natural keratinous systems are still some of the most superb thermal insulators. The popular and unsurpassed down jackets use feathers. As a result, researchers have tried to recreate these natural insulators' configurations in engineered materials to harness their desirable thermal capabilities.

Some organisms, like polar bears and penguins, can thrive in the most extreme conditions on earth due to their keratinous thermal protection ( Jia et al., 2017 ; Metwally et al., 2019 ). Polar bear hairs consist of a hollow porous interior that provides superior thermal properties surrounded by a shell of aligned fibers, which supplies mechanical stability. SEM images of these hairs are shown in Figure 15 A. Individual hairs are approximately 200 micrometers in diameter, while the interior pores measure 15-20 micrometers across. The length scale of these pores is significant because it allows the hairs to trap substantial amounts of air, providing a thermal buffer between the bear's living tissue and the surrounding arctic temperatures that can reach as low as −45°C.

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Polar bears can survive in some of the harshest environments on earth, largely due to their warm fur. Bioinspired models based on porous hairs have been fabricated to harness the remarkable thermal properties exhibited by polar bear hair

(A) SEM images of polar bear hair radial (left) and longitudinal (right) cross sections.

(B) Design set up for freeze spinning system used to fabricate bioinspired polar bear hairs fibers.

(C) SEM images of bioinspired hair cross sections fabricated at different temperatures.

(D) Plot of average pore size vs. fiber strength in the bioinspired fibers.

(E) Plot of difference in heat between the top of fibers and bottom of fibers with varying average pore size when placed on a heated stage over a range of temperatures (−20°C - 80°C). Reproduced with permission ( Cui et al., 2018 ). Copyright 2018, WILEY-VCH Verlag GmbH & Co. KGaA.

Since 3D printing cannot manufacture architectures on the scale of micrometers, Cui et al. (2018) used freeze spinning to create bioinspired synthetic fibers that could mimic the polar bear hair. This process is similar to freeze-casting in that it harnesses directional ice crystal growth to create a porous lamellar structure within an aqueous solution. However, freeze spinning performs this technique within a stable, extruded liquid wire. Once the wire is frozen, the material is freeze-dried to preserve the intricate microstructure formed by the ice crystals, and the completed porous fiber can be woven into a textile ( Cui et al., 2018 ). This process is visualized in Figure 15 B.

As with freeze-casting, several parameters can be adjusted to control the production, such as solution viscosity, extrusion speed, and freezing temperature. For the latter, Cui et al. (2018) found that the temperature at which the ice crystals are formed can be used to control the pore size and orientation in the fiber, as shown in Figure 15 C. As the temperature is lowered from −40 C o to −196 C o , more ice crystals are formed, but the freezing process occurs quickly, giving the crystals less time to propagate through the solution. The result is more numerous pores that are smaller in size. When the fiber is frozen in liquid nitrogen, a random porous network is produced, but when crystals are formed at higher temperatures, the pores align in the crystal growth direction. The pores' alignment and size have a significant effect on the fibers' tensile properties, as seen in Figure 15 D. Aligned pores provide better strength and elongation than a random porous network ( Cui et al., 2018 ).

In comparison, fibers with larger pores tended to have higher strength but lower average elongation than fibers with smaller pores. Smaller pores, however, provide better thermal properties. This behavior was determined by heating fibers with different pore sizes on a stage and measuring the temperature on fibers' surfaces using IR images. These results are summarized in Figure 15 E. This biomimetic material also showed promising results for thermal cloaking and, when embedded with carbon nanotubes, electro heating ( Cui et al., 2018 ).

Feathers are among the most ubiquitous materials used as thermal insulators due to their extreme lightweight and durability. Different types of feathers are distinguished by their structure and location on the bird: contour (body feathers) and plume (down feathers). Down feathers are primarily responsible for thermal insulation, which is attributed to their hierarchical foam-based structure creating large surface areas for trapping heat. Some academics have posited that Eiderdown, in particular, is the most thermally insulating natural material in the world ( Kasturiya et al., 1999 ). Down benefits from an impressive strength-to-weight ratio ( Gao et al., 2007 ; Havenith, 2010 ), compressibility ( Gao et al., 2010 ), and compression recovery ( Martin, 1987 ), making it invaluable as bodily insulation in extreme environments. The first use of down jackets was seen in expeditions to Mount Everest in 1922 and by 1933 in down sleeping bags, which have become a staple of mountaineering in the harshest of climates. While this application of keratinous tissue is hardly bioinspiration, this review would be incomplete if it did not mention the pervasiveness of feathers in a vast range of textiles from common bedding to elite sub-zero clothing ( Fuller, 2015 ). Even before down became popular, other keratin sources such as wool and animal fur have played a dominant role in the human race's ability to inhabit some of the coldest regions on earth.

One of the driving enterprises of the industrial revolution was the production of textiles. With such vast commercial implications, research on manufacturing cheap, synthetic fabrics with properties similar to wool and fur has been evolving for centuries. Modern clothing is often a mix of natural materials such as wool or cotton and synthetic fibers like polyester. In some cases, natural fibers have been replaced entirely. Examples include synthetic cashmere, which is usually a combination of rayon, nylon, and polyester, and fleece, typically composed of PET. Ultimately, many of our modern textiles are bioinspired materials that are attempting to replicate the success of traditional but expensive, labor-intensive keratinous systems.

In summary, keratinous materials' thermal insulation revolves around hierarchical surface texture or internal pores that are meant to trap air pockets and create a buffer between the animal and its surroundings. Efforts to recreate these structures using synthetic materials have been quite successful, highlighting that this is a fruitful area of study.

Reversible adhesion

Reversible or non-destructive adhesion allows for repeated attachment and detachment cycles that do not damage the substrate. Nature employs a variety of reversible adhesive strategies: mechanical interlocking, friction, chemical bonding, dry adhesion (i.e., van der Waals), wet adhesion (i.e., capillary), and suction (i.e., pressure differential). Often, organisms will use a combination of the above attachment methods to adhere to surfaces successfully. These processes are strongly dependent on the environment (predominantly wet vs. dry and smooth vs. rough). Mechanical interlocking, friction, dry adhesion, and wet adhesion are strongly dependent on having nanostructured surfaces. The hierarchical nature of keratin lends itself well to forming nanostructured and intricate designs. While the field of reversible adhesion is extensive ( Arzt et al., 2003 ; Gorb, 2008 ), our focus here is on materials inspired by keratin-based systems to highlight the diverse functionality that keratin offers. We will focus on the mechanical attachment found in the feather vane and dry adhesion found in gecko setae and their respective bioinspired designs. Claws and talons use a more conventional design principle, a relatively large hook, and will not be treated here.

The feather vane is directionally permeable, which effectively helps it capture air for lift ( Alibardi, 2007 ; Sullivan et al., 2017a ). This mechanism is controlled by the branching barbs' geometry and stiffness and interconnecting barbule network, which ultimately forms the feather vane ( Alibardi, 2007 ; Sullivan et al., 2017a ). Barbs, which branch from the rachis, are further branched into barbules. The barbules have hooklets (hamuli) on their extremities, which fit into the neighboring barb's groove, creating a highly ordered lattice of interconnected adjacent barbs ( Figure 16 A). Having multiple hooklets increases both the adhesion and the probability that two neighboring barbs will stay connected. The interconnected network of the feather vane, provided by this adhesive mechanism between the barbs and barbules, is credited as the essential element that allows birds to achieve flight ( Sullivan et al., 2017a ).

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Progression of bioinspired designs based on the attachment mechanism found in the feather vane

(A) SEM micrograph of the feather vane showing a branched network of barbs, barbules, and hooklets.

(B) First hook and groove-inspired sample.

(C) Modified hook and groove structure with a closer match in stiffness to the actual feather vane.

(D) Advanced replication of the feather vane to incorporate membrane flaps for directional permeability.

(E) The first groove-only unidirectional sliding structure.

(F) Two-dimensional sliding structure, which shows textile-like behavior.

(G) Cubic sliding structure which provides tailored stiffness in three dimensions. Adapted with permission ( Sullivan et al., 2019 ). Copyright 2019, Elsevier.

Several 3D printed bioinspired designs based on the reversible adhesive mechanism of the feather vane have been developed by Sullivan et al. (2019) . These 3D printed structures not only serve to understand better the mechanisms operating in the feather but extend beyond the scope of the intended function in nature to suggest innovative solutions for deployable structures, next-generation chain mail, and smart foams. The initial interlocking barbule bioinspired design was intended to mimic the attachment mechanism by scaling the dimensions to an appropriate size for 3D printing and mechanical testing. The first design in Figure 16 B helped demonstrate the feather vane's adherence through hooks and grooves that slide along each other. This feature is similar to the mechanism found in Velcro but is more organized and directional. While the first design served as a simplified model, the 3D printed material used was much stiffer than the feather vane. It did not accurately mimic the feather vane's elasticity and ability to re-adhere. The second model attempted to reduce the elastic mismatch by printing with a more flexible material which helped to elucidate how the material properties and the hook's geometry can enable sufficient adhesion that is damage-tolerant and can re-adhere readily ( Figure 16 C). Development of subsequent designs based on the barb and barbule interaction further increased in complexity to represent the feather vane with the inclusion of flaps that act as one-way valves ( Figure 16 D). These models suggest that two existing modes allow for tailored air permeability: (1) membrane flaps allow air to flow through space between barbules dorsally but not ventrally, and (2) the sliding of hooks along the grooves offers expansion within the feather vane (when hooks are closer to the base of the groove the vane is tighter, i.e., less permeable than when the hooks are at the tip of the groove). The purpose of this effort was to offer a simplified visualization of the complex nature of reversible adhesion and directional permeability in the feather vane ( Sullivan et al., 2019 ).

The subsequent iterations of designs expanded beyond just mimicking the feather vane by extracting fundamental design principles to optimize the interplay of tailorable, expansive materials. The ideas involved removing the hooks, which were shown to be the weakest point of adhesion from previous designs ( Figures 16 A–16D), and creating an exclusively grooved-based structure that had stoppers at the end to prevent complete detachment ( Figure 16 E). In the first groove-based structure, sliding was only able to occur in one direction. In this direction, sliding enabled an increase in flexibility while the perpendicular direction remained rigid. The design was further altered to allow sliding in both directions, which led to textile-like behavior when stretched open ( Figure 16 F). Finally, a cubic structure was developed, which allowed for sliding and manipulating the modulus in all three dimensions ( Figure 16 G). This progression in development highlights the importance of bioinspired design as a creative process reaching beyond the limitation imposed on nature to develop innovative materials.

The gecko setae are a most striking example of reversible adhesion in nature. Over the past two decades, these keratinous nanopillars have stirred up a tremendous amount of scientific interest, leading to the publication of hundreds of research papers and enough articles to be the topic of their own review ( Boesel et al., 2010 ; Jeong and Suh, 2009 ; Li et al., 2019 ; Niewiarowski et al., 2016 ; Russell et al., 2019 ; Stark and Mitchell, 2019 ; Wang et al., 2020b ; Zhou et al., 2013a ). However, interest in the gecko stretches back through the past century. Piquantly, geckos' mysterious ability to climb vertical walls and even to hang upside down on ceilings was correctly interpreted in 1902 by Franz Weitlaner ( Kroner and Davis, 2015 ). Since then, great strides in bioinspiration have come with vast amounts of research, and many groups have succeeded in making reversible dry adhesives based on the gecko setae structure. Here, we will only broadly cover this burgeoning area of study.

The gecko's adhesive pads utilize van der Waals forces and, to a lesser extent, capillary forces generated by the hierarchical broom-like geometry of setal arrays ( Li et al., 2019 ; Russell et al., 2019 ). The setae found on the gecko adhesive pad are arranged on lamellae and branch into hundreds of individual spatula-shaped tips (typically referred to as spatulae), as shown in Figure 17 A. van der Waals forces require extremely close contact (<10 nm) to generate a significant force, and this is accomplished by the flexible, branched nanostructure of the gecko pad. As the setae divide into smaller subdivisions with higher aspect ratios, their effective elastic modulus decreases, allowing them to conform easily to smooth and rough surfaces ( Russell et al., 2019 ). In the aggregate, the spatulae generate significant adhesive forces in the normal direction and frictional forces in the lateral direction, both of which are vital to the locomotion of the gecko ( Boesel et al., 2010 ). Further, these fine subdivisions have the added benefit of confining crack propagation if a single seta begins to fail ( Li et al., 2019 ).

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Geckos use van der Waals forces generated by densely packed setal arrays on the feet to climb even the sheerest surfaces. Many researchers have attempted to replicate this structure to create reversible, dry adhesives

(A) SEM image of the branched gecko setal array. The inset image shows the split-fiber endings with tilted, spatula-shaped tips ( Rong et al., 2014 ).

(B) SEM image of synthetic gecko-inspired adhesive composed of polymer micropillars with densely packed carbon nanotubes glued to the end. Open Access ( Rong et al., 2014 ). Copyright 2013, the authors.

(C) SEM image of bioinspired, tilted micropillars composed of polyurethane that mimic the gecko setae's directional gripping strength. Reproduced with permission ( Murphy et al., 2009a ). Copyright 2009, Wiley-VCH Verlag GmbH & Co. KGaA.

(D) SEM images of three hierarchical tiers of mushroom-shaped pillars composed of polyurethane that mimic the hierarchical branched structure found in the gecko pad. Reproduced with permission ( Murphy et al., 2009b ). Copyright 2009, American Chemical Society.

There are several additional characteristics of the gecko pad that make them particularly alluring to researchers. One of the gecko pad's most enticing features is its controllable and reversible adhesion, allowing it to be reused and not leaving behind any residue on the locomotive surface. The setal spatula-shaped tips attach to a surface when the gecko pulls its toes downward and inward, creating a small pulling angle between the setae and the surface. To detach from a substrate, the gecko pushes its toes upward and outward; this has the dual effect of increasing the pulling angle past the critical detachment angle (∼30°), while squeezing the setae to increase their effective elastic modulus and decrease their conformability to the surface ( Li et al., 2019 ; Russell et al., 2019 ; Tian et al., 2006 ). This process can be actualized in a matter of milliseconds allowing for rapid, reversible adhesion. Other unique properties provided by the branched setal arrangements include self-cleaning characteristics and adhesion in challenging environments like underwater and on surfaces with different polarity and roughness ( Russell et al., 2019 ; Stark and Mitchell, 2019 ).

Several research groups have hypothesized that the self-clean capabilities of the gecko-pad arise directly from the nanostructure of the setal configurations. Hansen and Autumn (2005) suggest that the primary reason for self-cleaning is the energetic disequilibrium between the substrate and the setae, but state that other factors like locomotion, particle rolling, and particles wedging between the setae could play a role ( Hansen and Autumn, 2005 ). Follow-up studies have confirmed that each of these mechanisms improve self-cleaning ( Hu et al., 2012 ; Mengüç et al., 2014 ), particularly when particles bond more strongly to the setae than the substrate. Xu et al. (2015) postulated that the dynamic motion of gecko toes (referred to as digital hyperextension) allows geckos to tune the pull-off velocity of the setal arrays ( Xu et al., 2015 ). Since the adhesive force between dirt particles and the substrate is velocity-dependent, and the force between particles and the setae is largely velocity-independent, increasing the pull-off rate can dislodge bonded particles from the surface of the toe pads. This velocity-controlled self-cleaning technique was applied to synthetic biomimetic materials with great success, achieving an ∼80% chance of particle detachment at high velocities (>1000  μm s −1 ) compared to the 0-40% chance of detachment beneath this threshold.

Many different approaches ( Arzt et al., 2021 ) have been utilized to fabricate nanostructures capable of generating dry, reversible adhesion inspired by the gecko setae. These include soft lithography ( Crosby et al., 2005 ; Lamblet et al., 2007 ; Peressadko and Gorb, 2004 ), injection molding ( Gorb et al., 2007a , 2007b ; Gorb and Varenberg, 2007 ; Varenberg and Gorb, 2007 ), hot embossing ( Hu et al., 2014 ), photolithography ( Aksak et al., 2007 ; del Campo and Greiner, 2007 ; Davies et al., 2009 ; Northen and Turner, 2005 , 2006 ), plasma etching ( del Campo and Greiner, 2007 ; Jeong et al., 2009 ), electron beam lithography ( Geim et al., 2003 ), carbon nanotubes (an example of which is shown in Figure 17 B) ( Hu et al., 2013 ; Qu et al., 2008 ; Rong et al., 2014 ; Yurdumakan et al., 2005 ), nanodrawing ( Jeong et al., 2006 ), micro/nanomolding ( Glassmaker et al., 2004 ; Greiner et al., 2007 ; Mahdavi et al., 2008 ; Sitti and Fearing, 2003 ), dip-transferring ( Murphy et al, 2009a , 2009b ), two-photon lithography ( Hensel et al., 2018 ), nanoimprint lithography ( Raut et al., 2018 ), and many more. As research groups have aimed to mimic the gecko pad's intricate structure more closely, the complexity of their fabrication processes has increased. Early techniques focused on only manufacturing a dense network of nanopillars. However, these studies showed that other design parameters need to be considered to truly capture the gecko pad's functionality.

For example, the natural setae are tilted, which creates much larger shear forces in the gripping direction than in the non-gripping direction, effectively enhancing the gecko pad's reversible adhesion ( Boesel et al., 2010 ). Figure 17 C shows tilted polyurethane fibers fabricated via inclined exposure and dip coating to capture this parameter. Another essential variable for gecko-inspired adhesives is the shape of the tip of the nanofibers. Many different arrangements have been investigated, but mushroom-shaped tips have proven to be the most successful design ( Russell et al., 2019 ).

del Campo et al. (2007) compared biomimetic arrays with various pillar shapes and found the mushroom configuration to have a pull-off strength 30 times that of cylindrical pillars ( del Campo et al., 2007 ). Spatular tips provided an intermediate degree of adhesion, while concave tips and spherical tips were slightly better than flat cylindrical pillars. Spuskanyuk et al. (2008) hypothesized that part of this larger adhesive force is due to the fact that mushroom-shaped tips are less adversely affected by edge defects than flat or cylindrical pillars ( Spuskanyuk et al., 2008 ). Further, stress concentrations are reduced at the contact interface ( Li et al., 2019 ). Several other papers ( Aksak et al., 2014 ; Balijepalli et al, 2016 , 2017 ; Fleck et al., 2017 ; Gorb et al., 2007a ) have examined the mushroom shape both experimentally and numerically and concluded that it is one of the best pillar designs for adhesion. Fleck et al. (2017) and Balijepalli et al. (2017 , 2016) considered fibril detachment as a crack propagating along the pillar-substrate interface and found that mushroom-shaped pillars reduce the corner stress intensity of the contact zone, thus reducing the likelihood of detachment ( Fleck et al., 2017 ) ( Balijepalli et al, 2016 , 2017 ). Review papers on the subject ( Boesel et al., 2010 ; Li et al., 2019 ; Russell et al., 2019 ; Wang et al., 2020b ; Zhou et al., 2013a ) have also noted several other benefits of the mushroom shape, including improved adhesion enhancement via contact splitting and increased crack trapping compared to flat cylinders.

Figure 17 D shows a three-layered hierarchical arrangement of polyurethane fibers with mushroom-shaped tips. This arrangement was manufactured with soft-lithography and capillary molding. Fiber aspect ratio, fiber radius, hierarchical branching arrangements, and material selection are all important factors as well. Figure 17 shows some of the tradeoffs that come with different manufacturing processes. While the polyurethane tips in Figure 17 D have controlled tip geometry, their aspect ratio and fiber density are much lower than those of the carbon nanotube tipped design in Figure 17 B. Neither of these designs was able to incorporate the tilted structure shown in Figure 17 C. The gecko pad's ability to optimize all of these different parameters simultaneously provides just another example of why natural keratin can be so impressive relative to manufactured materials and how there is so much for engineers to learn from nature.

The applications for gecko-inspired, dry, reversible adhesives are seemingly endless. One of the most popular uses of this emerging technology is soft robotics ( Li et al., 2016 ). The reversible dry adhesion is ideal for (unsurprisingly) climbing and gripping. It has been utilized for numerous commercial devices (like Onrobot's soft gripper and GECOMER's pick-and-place robotic systems), as well as countless academic pursuits ( Arzt et al., 2021 ; Asbeck et al., 2009 ; Dadkhah et al., 2016 ; Estrada et al., 2016 ; Hawkes et al, 2015a , 2015b ; Henrey et al., 2014 ; Jiang et al, 2015 , 2017 ; Kalouche et al., 2014 ; Ko et al., 2017 ; Purtov et al., 2015 ; Seo and Sitti, 2013 ; Song and Sitti, 2014 ; Zhou et al., 2013b ). Furthermore, several products utilizing adhesive materials based on the gecko pad are now commercially available from Geckskin, nanoGriptech, and Gottlieb Binder GmbH.

By way of their tunable and hierarchical structure, keratinous materials have evolved diverse methods to achieve reversible adhesion. In the feather, this is accomplished through the mechanical interlocking of hook-shaped barbs and barbules, while the gecko pad adheres to surfaces with van der Waals forces generated by its branched setal arrangement. These features have been translated to scaled up to macroscopic engineered systems (as in the feather-inspired 3D prints) and biomimetic nano- and microscale structures (for the gecko setae).

Lightweight structures

In engineering applications, sandwich structures are used for their ultra-lightweight, energy absorption capabilities, and comparable mechanical strength relative to bulk materials. Sandwich structures can be tailored by controlling the properties of the face (outer cortex) and core (foamy center) and their geometry. Typically, sandwich structures are constructed with a high modulus face and a low modulus core to achieve a lightweight yet stiff material with rectangular cross-sections. Sandwich structures are not limited to engineered materials and are found in abundance in keratin-based systems, including beaks ( Seki et al., 2006 ), feathers ( Liu et al., 2015a ; Sullivan et al., 2017b ), quills ( Yang et al., 2013 ), baleen ( Wang et al., 2019 ), and spines. Unlike engineered materials, the faces and core of biological materials are frequently made of the same material but occur in distinct phases: the face being more compact while the core is more porous. Here, we will review how the lightweight yet mechanically robust, keratin-based sandwich structures implemented in porcupine quills and hedgehog spines serve as the basis of lightweight bioinspired designs. While sandwich structures are not limited to keratinous materials, this review highlights the structural and functional diversity found in keratin systems that lend themselves to developing bioinspired structures.

The porcupine quill is composed of α-keratin and is a lightweight yet buckling-resistant structure that undergoes significant compressive and flexural loads during its service as a protective mechanism. The sandwich structure of the porcupine quill consists of a thin-walled cylindrical cortex enclosing a closed-cell foam. Some porcupine quills contain an additional structural element that reinforces the foamy center, which is referred to as a stiffener. The stiffeners have been found to increase the compressive strength and buckling resistance of porcupine quills ( Yang et al., 2013 ). Inspired by the stiffeners present in the porcupine quill, Tee et al. (2021) developed several 3D-printed cylinders with varying infill structures from uniform to non-uniform designs to mimic the radial structures found in the porcupine quill. However, mechanical testing was not performed, and little information is known on the degree of reinforcement the stiffeners provide and how their structure can be tailored ( Tee et al., 2021 ).

Hedgehog spines are similarly structured to porcupine quills and contain reinforcing stiffeners, further classified as longitudinal stringers and transverse plates ( Vincent, 2002 ; Vincent and Owers, 1986 ). Despite their structural similarities, porcupine and hedgehog spines serve different functions. Hedgehog spines are adhered to within the skin and are primarily used as shock absorbers upon falling from great heights, while porcupine quills can readily detach from the body and serve as a defensive mechanism. Due to their high stiffness and capabilities for impact resistance, hedgehog spines are a suitable inspiration for developing lightweight yet mechanically robust bioinspired designs.

Drol et al. (2019) , using X-ray microcomputed tomography, were able to capture the key internal structural design elements found in hedgehog spines, which were then used to create computational model abstractions in ABAQUS and compared to analytical models to better understand the role that stringers and plates play in the spine's flexural performance. Ten models with increasing complexity were generated. The most basic level is a simple hollow cylinder (level 1) and builds up to most realistically represent the spine (level 10) with a complex arrangement of longitudinal stringers and periodically arranged branched transverse plates. The beam models were subjected to 3-point bending with a displacement-controlled boundary condition in which the bending stresses, the normalized bending stresses, and the Von Mises stress contours were quantified. The hollow tube, the simplest case, is reported to have the highest specific stiffness; however, the lack of stiffeners limits its ability to reduce buckling. The model with the next highest effective stiffness is model 10, the most complicated and representative model of the spine. Model 10 contains longitudinal stringers and branched transverse plates with the smallest spacing between the central plates and the longitudinal stringers and a more accurate curvature connection between the stringers instead of a blocked fillet. The build-in model 10 allows for removing material while maintaining stiffness, creating a lightweight yet stiff structure. The longitudinal stringers aid in increasing the bending stiffness by localizing material further away from the central axis, which effectively increases the second area moment. The transverse plates provide reinforcement and help distribute the applied load evenly, minimizing buckling and localized failure. Furthermore, this study provides insight into how the structural organization of keratin-based materials, such as the hedgehog spines, can be directly translated to synthetic designs to develop tailored stiff and lightweight structures. This study's findings have even inspired the development of novel football helmet liners to help reduce traumatic brain injuries. This example illustrates how bioinspired designs stimulate innovation ( Drol et al., 2019 ).

The feather shaft is another example of how keratin can be used to achieve a lightweight yet mechanically robust structure that is able to withstand aerodynamic loads during flight. This behavior is primarily attributed to the sandwich structure of the feather shaft. The feather shaft is composed of an outer shell of compact keratin that surrounds a medullary center made of foamy keratin. Liu et al. (2015a) investigated the hierarchical structure and mechanical properties of the peacock tail feather shaft under tension and compression. They determined that the presence of the foam center enhanced failure resistance by delaying splitting and buckling of the cortex shell and exhibits overall improved compressive stability ( Liu et al., 2015a ). While there has been a significant amount of work dedicated to understanding the structure and mechanical properties of the feather shaft, there have been limited attempts toward the development of bioinspired sandwich structures based on the feather shaft. We suggest that this is an area of study for future work.

Many keratinous materials manage to achieve good mechanical properties while limiting their mass. Often this is accomplished with a sandwich structure consisting of foam surrounded by a stiff exterior face. Since low density is a highly coveted trait in engineered materials, these natural keratinous systems have served as the basis for bioinspired designs aimed to capture high strength to weight ratios.

Structural color

Besides the outstanding mechanical, lightweight, and thermal properties of avian feathers, these keratinous materials are also known to display a diverse range of vibrant colors. This property is in part due to structural coloration, which arises from the interactions of light with a submicron array of varying morphologies which include multilayer structures (as seen in the iridescent throat patch of the hummingbird) ( Gruson et al., 2019 ), two-dimensional photonic crystals (as seen in peacock and mallard feathers) ( Freyer and Stavenga, 2020 ; Stavenga et al., 2017 ; Weiss and Kirchner, 2010 ; Zi et al., 2003 ), or spinodal-like channel structures ( Parnell et al., 2015 ) (as seen in the Eurasian Jay Garrulus glandarius ). These nanostructures self-assemble and can occur as a multi-layered structure of β-keratin and a pigment-based protein (e.g., melanin, carotenoids), as shown in Figure 18 A. The combination of structural color from the sub-micron array of keratin and the absorption from the pigment is referred to as color mixing. β-keratin has a low refractive index (∼1.5), but when implemented in a multi-layer structure, it allows for high reflectance and vivid coloration ( Burg and Parnell, 2018 ). The presence of pigments strongly contributes to the vibrant coloration due to their high refractive indices and broad absorption spanning the UV-visible range. Structural color in avian feathers can occur as iridescent or non-iridescent and is strongly dependent on the underlying structure and organization. Typically, long-range order is responsible for producing iridescence, while short-range order is non-iridescent ( Noh et al., 2010 ). Thus, structural color in avian feathers is highly tunable and thus a desirable candidate for bioinspiration.

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Structural color found in avian feathers and bioinspired analogs

(A) Violet-backed starling and TEM micrograph of the multi-layered structure of hollo melanosomes and a thin film of keratin.

(B) Structural color produced by SMNPs. (C) Micrograph detailing the arrangement of SMNPs as a thin film. Adapted with permission ( Xiao et al., 2015 ). Copyright 2015, American Chemical Society.

Despite the vast arrangement of keratin in combination with pigmentation and the subsequent multitude of colors with varying optical properties (iridescent vs. non-iridescent) found in bird feathers, there have been limited ventures at bioinspiration. The most prevalent study that draws inspiration from feathers is the development of structural color produced by self-assembly of synthetic melanin nanoparticles (SMNPs) inspired by the assembled melanosomes in avian feathers ( Xiao et al., 2015 ). Xiao et al. (2015) used a vertical evaporation-based self-assembly method to develop thin films of SMNPs with a wide range of colors (red, orange, yellow, and green) ( Figures 18 B and 18C). The coloration produced is attributed to the thickness of the thin film which can be controlled by the concentration and evaporation rate. Additionally, the morphology of the SMNPs influences the packing, and, therefore, the film thickness and coloration produced. In avian feathers, there exists a diverse range of melanosome geometries from spherical to oblong and hollow to filled. These morphologies can additionally tune the coloration produced, which is an exciting avenue for future work. The SMNPs have a broad absorption spectrum (high absorption at short wavelength and low absorption at long wavelengths) and a relatively high refractive index (∼1.4–1.6 at 589 nm) which was found to be responsible for the enhanced color saturation and purity. In addition to the desirable optical properties, SMNPs are biodegradable and inherently biocompatible, making them suitable candidates for various applications ( Xiao et al., 2015 ).

Structural coloration is not limited in nature to keratinous materials and is additionally found in chitin-based materials such as the morpho butterfly and the exoskeletons of beetles ( Fu et al., 2016 ). These chitin-based systems have been extensively studied and have led to the development of numerous bioinspired structural colors ( Chung et al., 2012 ; Steindorfer et al., 2012 ; Zhang and Chen, 2015 ). Despite their lack of prevalence in bioinspired structural colors, there are still many opportunities awaiting to be explored in the field of avian feathers. This review highlights the importance of keratinous structural colors found in avian feathers and the vast potentials for these systems to serve as bioinspired candidates.

Hydrophobic surfaces

Hydrophobic surfaces are essential in both the engineering and the biological worlds. As such, researchers have been attracted to how living creatures can repel water by manipulating the contact angle of water droplets on their surfaces. Certain organisms, like ducks, excrete hydrophobic oils that can be spread on the surfaces of their feathers to repel water. Others, like the famous lotus flower, utilize nanoscale roughness to decrease the contact area of water droplets on their surface, with the two-fold benefit of keeping the organism dry while cleaning dirt and debris of the substrate as water droplets runoff. This phenomenon, dubbed the “lotus effect,” has been observed in several keratinous systems as well, which is particularly intriguing because keratin itself is very water absorbent. It should be noted that, in nature, several hydrophobic strategies are often utilized in tandem. The duck feather, for example, also has significant surface roughness, which helps to repel water along with the oil excreted by its uropygial gland, while the lotus leaf's nanostructure is covered by a thin, hydrophobic wax film that helps prevent water from penetrating the epidermis. Thus, researchers have attempted to imitate the surface features found on hydrophobic keratinous systems to create synthetic, water-repellent materials.

Penguin feathers have not only superior thermal insulating properties but also remarkable anti-icing properties. Despite spending a significant amount of time in freezing temperatures and swimming underwater, ice crystals are not typically observed on penguins' feathers. The secret to the ice-phobicity of penguin feathers is in its rough micro and nanostructure, which traps air in grooves, preventing supercooled water droplets from adhering and coalescing. A schematic of this water repulsion mechanism can be seen in Figure 19 A. This trapped air is also postulated to provide a thermal barrier that reduces ice adhesion strength and heat transfer during icing. On the surface of the barbules and hamuli are grooves that are about 100 nanometers deep. These grooves are responsible for the surface roughness that creates the air pockets shown in Figure 19 A ( Wang et al., 2016d , 2016e ).

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The multiscale surface roughness and fine nanoscaled grooves on feathers help them repel water

(A) Schematic showing how the hamuli on penguin feathers trap air beneath water droplets creating an air cushion and minimizing the amount of material in contact with the water.

(B) Bioinspired polyamide nanofiber membrane fabricated via asymmetric electrode electrospinning.

(C) Chart of contact angle and adhesive force versus location on the polyamide membrane highlighting the effect of fiber density. Reproduced with permission ( Wang et al., 2016d , 2016e ). Copyright 2016, American Chemical Society (D) SEM image of cotton fiber with precipitated chitosan nanoribbons on the surface inspired by duck feathers.

(E) SEM image of polyester fibers with precipitated chitosan “nanoflowers” on the surface. Reproduced with permission ( Liu et al., 2008 ). Copyright 2008, IOP Publishing Ltd.

Inspired by the penguin feather, Wang et al. (2016) used asymmetric electrode electrospinning to weave an anti-icing polyamide nanofiber membrane, as shown in Figure 19 B ( Wang et al., 2016d , 2016e ). The radially arranged fibers mimic the barb tips' structure, while other fibers randomly overlap this arrangement, creating a regular 3D network similar to that found in the feather. The fibers are densely packed near the triangular electrode compared to the fibers near the curved electrode, as shown in Figure 19 b1, b2, and b3. This fiber arrangement creates a gradient in the chemical surface structure. In the region of densely packed fibers, the static contact angle of water droplets was ∼154° with a low adhesion force of ∼37 μN. In this region, droplets struggled to permeate the tightly bound membrane. The few droplets that were able to adhere to the fibers coalesced with other droplets and resulted in self-propelled jumping, i.e., the droplets fell off the membrane naturally before freezing. As the distance between the fibers increases, more droplets were able to penetrate the membrane. The static contact angle and adhesion force of the water droplets were measured at 105.1° and 102μN, respectively. Figure 19 C shows the gradual change in these values through the membrane's radius. Droplet coalescence and jumping did not occur when the fibers' distance was greater than the diameter of droplets. After 3-4 hr at -5°C, some frost and ice were found on the less densely packed fibers but not in the densely packed fibrous network. This result shows that by tuning the density of the overlapping nanofiber network, anti-icing properties similar to those found in keratinous penguin feathers can be achieved.

Waterfowl, ducks in particular, are so famous for their anti-wetting capabilities that the phrase “like water off a duck's back” has worked its way into our everyday lexicon. Until recently, it was generally thought that this extraordinary hydrophobicity arose from the low surface energy of preening oil excreted from glands at the base of their tail and spread over the feathers. However, recent studies ( Nishino et al., 1999 ) of preening oil on smooth surfaces have revealed that it is not that special after all and is less hydrophobic than several synthetic resins and oils. The feather's structure, coupled with preening oil, makes water run off of a duck's back so efficiently. Like penguin feathers, duck feathers have multiscale textures, with the same branched structure and micro-sized surface features covered with nanoscale grooves and protuberances. Liu et al. (2008) mimicked this structure by precipitating chitosan nanostructures on the surface of textile microfibers. They did so by dip-coating fibers in an acidic solution containing chitosan before placing them in an ammonia gas environment. The ammonia is absorbed by the film, making the solution basic and causing the cationic polyelectrolyte chitosan to precipitate in nanofeatures on the textile substrate's surface. On cotton fibers, the chitosan formed long ribbons ( Figure 19 D), while on polyester fibers, the chitosan shrank down to nanosized flower shapes ( Figure 17 E). The result is a hierarchical arrangement of surface irregularities where the fibers themselves compose the microscale roughness, while the chitosan precipitates form the nano roughness. Once the fibers are dried, they are treated with polysiloxane to lower the fibers' surface energy (similar to the preening oil found on natural duck feathers). With just the polysiloxane treatment, the contact angle of a water droplet was 118° for cotton and 100° for polyester. When combined with the chitosan surface roughness, these values rose to 152° and 148°, respectively, showing how the combination of a low surface energy film and surface roughness, similar to feathers, can lead to the development of superhydrophobic materials ( Liu et al., 2008 ).

The oberhautchen (thin outer layer) of many lizard and gecko skins is composed of β-keratin. Many geckos also have tiny keratin spinules upon this outer layer that serve to repel water and disrupt bacterial growth. Figure 20 A (i-iii) shows images of the scales of Strophurus williamsi , a species of arboreal geckos found in Australia. Generally, these spinules are 0.5–4 μm long packed closely together with over 400 spinules per 10 μm 2 . These spinules are mounted upon scales that have a honeycombed-shaped basal layer composed of intersecting ribs. The static contact angle of water droplets was similar to that of feathers, ranging between 151 and 155°. Interestingly, gecko skin accomplishes such impressive superhydrophobicity from its spinule density rather than finer roughness structuring like the channels found on insect hairs or the hamuli in penguin feathers. These hairs not only prevent water from building up on the skin of the gecko but also allow the skin to clean itself, removing harmful bacteria and contaminants as droplets coalesce and run off the gecko with even the slightest tilt or perturbation ( Watson et al., 2015 ).

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Much like the gecko pad, the outer layer of skin on the gecko has hydrophobic, self-cleaning properties due to its rough mesostructure, which researchers have attempted to replicate

(A) SEM images of natural gecko skin (i-iii) alongside SEM images of biomimetic polystyrene replicas made via biotemplating (iv-vi).

(B) Close-up SEM images of gecko spinules and the different measurements used to characterize them (left). Various biomimetic replicas, like the ones shown in A iv-vi, were prepared using several polymer solutions. The resultant spinule shapes are visualized (right) and compared to the natural spinules found on the gecko. Open access ( Green et al., 2017 ). Copyright 2017, the authors.

Green et al. (2017) developed a benchtop biotemplating apparatus to fabricate synthetic replicas of gecko skin spinules with comparable hydrophobicity to emulate their antibacterial properties. To do so, negative molds were generated by coating shed gecko skin, which was adhered to a glass slide by a thin layer of water, with commercially available PVS. The water also served to inflate the spinules to mimic their natural state better. This negative mold was then used to fabricate gecko skin replicas from several different polymer solutions targeted toward various applications. These included a synthetic polystyrene solution and natural biopolymer solutions of chitosan, silk fibroin, fused bilayers of chitosan and alginate polysaccharides, and blended α-keratin hair extract ( Green et al., 2017 ).

Each solution was successfully used to form a replica of the gecko skin nanostructure; several images of the natural shed gecko skin compared with the polystyrene replica are shown in Figure 20 A (iv-vi). Some of the solutions were able to more closely mimic the gecko spinules' dimensions, as visualized in Figure 20 B. The curing process had a significant effect on the ability of each solution to closely resemble the geometry of the natural gecko spinules. For example, the polystyrene solution hardens slowly due to organic solvent evaporation, which resulted in stiffer spinules with less curvature. The metrics for measuring curvature in the spinules are shown by images “d” and “e” in Figure 20 B. The chitosan-based replicas, on the other hand, closely mirrored the curvature of the nano tip, as well as the thickness and height of the natural spinules. The biomimetic samples were only slightly less hydrophobic than the natural gecko skin obtaining a contact angle of about 134°. The synthetic spinule arrays also revealed notable anti-bacterial properties. Confocal microscopy showed that the spinules effectively disrupted bacterial cultures grown on the replicas removing as much as 95% of bacteria from the surface after water treatment. Vucko et al. (2008) developed a similar procedure using epoxy molds of live geckos to observe their oberhautchen without needing to kill and prepare them for SEM examination. This approach is highly applicable to other organisms and other research fields since it can be non-destructively performed on living creatures while generating finely detailed replicas for observation or functional use ( Vucko et al., 2008 ).

Many keratinous materials that provide thermal insulation also protect organisms from getting wet because significant surface roughness benefits both areas. In the case of hydrophobicity, this roughness comes in many forms in nature, such as hamuli, nano grooves, or spinules, but all have the objective of reducing the area in contact with water droplets allowing them to run off the surface efficiently. Like research on thermal insulation and reversible adhesion, studies on bioinspired surface roughness to achieve anti-wetting properties have shown great success and are a promising research area for bioinspiration.

Keratin as a material for engineered systems

So far, we have seen how keratinous structures provide beneficial properties that can be used to inspire engineered designs. However, keratin itself has often been utilized as a material for various applications due to its unique intrinsic properties. Over the past few decades, many researchers have explored how to connect different technologies such as materials science, applied health sciences, and engineering. This section will discuss possibilities to use keratin for applications in the: (i) biomedical, (ii) composite, and (iii) reversible material realms.

Historically, keratin was one of the first polymers used by humans before the plastics revolution in the 20 th century. Keratin extracted from tortoise shells has been used to craft fine components, like hairbrushes, for hundreds of years, while baleen from whales was famously used to make corsets ( McKittrick et al., 2012 ; Wang et al., 2016b ). Hair (typically human or horsehair) has also had versatile applications ranging from paintbrushes to the torsional springs used in ancient Greek and Roman artillery. Researchers have recently explored natural macromolecules as candidates to perform biochemical, mechanical, and structural roles due to their appealing properties.

Keratin can be extracted from various sources (typically wool, poultry feathers, or hair) using multiple different techniques. Common extraction methods include oxidative and reductive extraction, steam explosion extraction, or ionic liquids and eutectic solvents ( Feroz et al., 2020 ; Shavandi et al., 2017 ). Studies involving oxidative technologies and reductive extraction were initially applied to animal horns and hooves but were also used to extract keratin from wool and human hair. Early studies on the properties of extracted keratin led to increased interest in exploring keratin for medical applications. Among the first innovations were keratin powders for cosmetics, fibers, composites, and coatings for drugs ( Beyer, 1907 ; Dale, 1932 ; Goldsmith, 1909 ; Rouse and Van Dyke, 2010 ).

Biomedical usage

Recently, there has been a significant increase in the number of biomedical studies related to using keratin-based biomaterials. This variety of applications includes biomedicine, natural polymer flocculants, bioelectronics, biolubricant formulations, and manufacturing bone scaffolds ( Roy et al., 2015 ). Keratin is widely used in biomedical applications due to its biocompatibility, lack of immune reaction upon transplant, good cellular interaction, and biodegradability ( Dickerson et al., 2013 ; Rouse and Van Dyke, 2010 ).

Asia has taken the lead in keratin biomaterials research since the first medical application of pyrolyzed human hair by a Chinese herbalist dates from the 16th century ( Zhen, 2005 ). In the modern age, scaffolds, hydrogels, powders, films, and fibers have been prepared, starting with early studies by Japanese scientists ( Ito et al., 1982 ; Noishiki et al., 1982 ) in 1982 on vascular graft production with hemostatic properties. Researchers have also shown that keratin can be effectively used for peripheral nerve regeneration, drug delivery, hydrogel formation, and films that promote wound healing ( Rouse and Van Dyke, 2010 ). For medical applications, keratin has shown interesting characteristics, but its potential has not yet been fully explored. For example, areas such as wound healing, bone regeneration, peripheral nerve repair, antimicrobial activity, hemostasis, and cell adhesion of amino acid sequences (due to the Arg-Gly-Asp and Leu-Asp-Val binding motifs) have led to increasing interest in keratin for medical applications. Although keratin-based biomaterials show wide promise, there can be significant costs associated with the extraction and processing of keratin and its post-processed mechanical characteristics. In 1983 and 1985, researchers from Japan and the UK, respectively, published papers speculating on the prospect of using keratin as the building block for new biomaterials ( Jarman and Light, 1985 ; Various Authors, 1993 ).

Also, keratin biomaterials derived from wool and human hair have been shown to possess cell-binding motifs, such as leucine-aspartic acid-valine (LDV) and glutamic acid-aspartic acid-serine (EDS) binding residues, which are capable of supporting cellular attachment. Together, these properties create a favorable three-dimensional matrix that allows for cellular infiltration, attachment, and proliferation. Thus, the conservation of biological activity within regenerated keratin biomaterials could prove advantageous for controlling specific biological functions in various tissue engineering applications ( Rouse and Van Dyke, 2010 ).

Reconstituted biopolymers often suffer from inferior mechanical properties, which can pose a challenge for processing and limit applications. This is especially true for biomaterials made from extracted keratin fibers, despite the stellar mechanical properties found in natural keratinous materials. Thus, many studies have targeted keratin films, focusing on the physical strength and flexibility of the films while maintaining their excellent biological activity ( Rouse and Van Dyke, 2010 ). The addition of other biopolymers such as chitosan or silk-fibroin improves the mechanical properties of keratin. The chitosan-keratin films also had beneficial anti-microbial properties and proved to be suitable substrates for cell cultures ( Lin et al, 2017 , 2018 ; Tanabe et al., 2002 ). For silk-fibroin and keratin films, studies have shown that the two molecules interact synergistically and provide unique properties not found in pure keratin or silk-fibroin films. For example, the polarity of keratin's amino acids causes silk-fibroin to rearrange from a random-coil to β-sheet configuration ( Lee and Ha, 1999 ; Vu et al., 2016 ). As a result of these unique interactions, the combined film is more biocompatible ( Lee, 2001 ; Lee et al., 1998 ) and biodegradable ( Vasconcelos et al., 2008 ) than its constituents.

Keratin has also been explored as a raw material for cell scaffolds and shows significant promise due to its ability to self-assemble into complex 3D shapes. A host of fabrication techniques from electrospinning ( Wang et al., 2016d , 2016e ), wet spinning ( Yue et al., 2018 ), photomask micropatterning ( Yue et al., 2018 ), and compression molding/particulate leaching ( Katoh et al., 2004b ) to freeze casting of aqueous keratin solutions ( Lin et al., 2019 ; Tachibana et al., 2002 ) have been used to create keratin scaffolds. These scaffolds have many advantages, including a stable homogeneous, interconnected, porous structure ( Lin et al., 2019 ; Tachibana et al., 2002 ), free cysteine residues that can be used to bind bioactive substances to the scaffold surface ( Kurimoto et al., 2003 ; Tachibana et al., 2005 ), and resorbability ( Peplow and Dias, 2004 ) that make it a suitable material for tissue engineering and drug delivery ( Lin et al, 2017 , 2018 , 2019 ; Srinivasan et al., 2010 ; Verma et al., 2008 ). These properties have also led to studies on keratin-based biomaterials for wound ( Konop et al., 2017 ; Lin et al., 2018 ; Than et al., 2012 ; Wang et al., 2016d , 2016e ) and burn dressings ( Poranki et al., 2014 ).

Composite films of keratin and synthetic polymers have also been fabricated to create films with even better mechanical properties. For example, poly (diallyl dimethylammonium chloride) and poly (acrylic acid) were blended with keratin extracted from wool to fabricate thick films based on the principle of poly ionic complexation. This was accomplished using a layer-by-layer self-assembly method ( Ducheyne et al., 2017 ; Katoh et al., 2004a ; Sionkowska et al., 2010 ). Keratin blends with poly(ethylene oxide) have also been explored for usage as scaffolds for cell growth, wound dressings, and drug delivery membranes, while keratin mixed with polyamide 6 has been envisaged as a practical material for biomedical devices, active water filtration, and textile fibers ( Zoccola et al., 2007 ).

Keratin's emerging role as a medical biomaterial revolves around many of the same aspects that make it a successful biological material. Its tunable properties and architecture make it viable for numerous different applications, while its abundance and natural origin make it appealing to researchers as an economical, sustainable, and biocompatible material. However, it is limited by the mechanical weakness of reconstituted keratin and the lack of cheap and scalable extraction techniques ( Shavandi et al., 2017 ).

Composite materials have steadily grown in popularity over the past decades due to their lightweight yet mechanically robust properties. However, these synthetic materials are traditionally produced from petroleum-based plastics, which are increasingly expensive and environmentally harmful. Many researchers aim to tackle this problem with biodegradable, renewably sourced composites made of biopolymer matrixes and natural fibers. Knowledge of the properties of available biodegradable polymers and natural fibers is essential for manufacturing a biodegradable composite ( Shrivastava and Dondapati, 2021 ).

Polymers reinforced with natural fibers, commonly named “bio-composites,” have started to be used industrially in the automotive and building sectors as well as the consumer goods industry. Green composites are a specific class of bio-composites where a bio-based polymer matrix such as a biodegradable polyurethane is reinforced by natural fibers such as keratin ( García et al., 2008 ; Quirino et al., 2014 ; Zia et al., 2008 ). Väisänen et al. (2016) describe natural fiber-polymer composites (NFPCs) as renewable and sustainable materials since they are composed of natural fibers embedded in a polymer matrix which may also be of biological origin (e.g., PLA) ( Väisänen et al., 2016 ).

Conzatti et al. (2013) reported on the valorization of keratin-based wastes, made of unserviceable poor quality raw wools from farm breeding, fiber byproducts from textile processing, and horns, nails, hair, and feathers from butchery. Zoccola et al. (2009) estimated that keratin wastes from breeding, butchery, and textile industry, made up of wool, hair, feathers, beaks, hooves, horns, and nails, have been estimated worldwide to be more than 5,000,000 tons/year ( Zoccola et al., 2009 ). With an increasing demand for sustainable materials, these protein byproducts are beginning to be regarded as renewable resources worthy of better exploitation ( Conzatti et al., 2013 ).

Extracted keratin has also gained popularity as a component for composite production as both a filler material and a fiber reinforcement. This interest is primarily driven by keratin's availability and environmental benefits (biodegradable, renewable, leftovers from other products) on top of their beneficial properties.

Donato et al. ( Donato and Mija, 2019 ) discussed the manufacturing of keratin-based composites with different polymers in detail. To form efficient keratin-polymer composites, it is essential to have good adhesion between the fiber and polymer matrix. Since keratin fibers have numerous hydrophilic surfaces, this can lead to weak mechanical properties of the overall composite material. As a result, coupling agents are sometimes required to boost interfacial adhesion. For example, Song et al. (2017) used functionalized cellulose nanocrystals to crosslink keratin fiber while also serving as reinforcement. This interfacial treatment resulted in marked improvements in tensile strength, elongation to failure, and toughness of such a composite. Further, the incorporation of cellulose nanocrystals reduced the keratin's water sensitivity which is a barrier for many in vivo applications ( Song et al., 2017 ). More approaches to coupling agents and keratin-polymer composites are discussed in detail by Shavandi and Ali ( Shavandi and Ali, 2019 ).

As attractive as synthetic polymers are, their use is coming under scrutiny due to the realization that petroleum reserves are finite and that oil prices are likely to rise steadily over the next few decades. Furthermore, with global environmental awareness at an all-time high, synthetic polymers have lost some of their luster. The synthetic fiber industry as it currently exists will ultimately decline and be replaced by an industry based on renewable feedstocks ( Fudge et al., 2010 ). Recent works on the mechanical properties of fibers isolated from hagfish slime suggest that these unique fibers may one day be replicated in a way that is environmentally sustainable and economically viable. These “slime threads” consist of bundles of 10 nm protein nanofibers known as IFs, which form part of the cytoskeleton in most animal cells ( Koch et al, 1994 , 1995 ). Keten et al. (2010) explored the nanoconfinement of β-sheet crystals in silk as a means to control stiffness, strength, and toughness. This study highlighted another feature that makes β-sheet crystals an attractive model: they self-assemble from soluble precursors into 10 nm filaments in aqueous buffers ( Keten et al., 2010 ). The key to the high strength and toughness of spider silk and hagfish threads are the β-sheet crystallites that simultaneously crosslink the protein molecules and arrange them into a structure in which “sacrificial bonds” increase the energy required to break the material ( Keten et al., 2010 ; Koch et al., 1994 ; Mostaert and Jarvis, 2007 ).

Pourjavaheri et al. (2018) developed a bio-composite from chicken feather waste and thermoplastic polyurethane. This composite material was fabricated via solvent-casting evaporation at eight different compositions. The thermo-mechanical properties of the composites were assessed using thermogravimetry, dynamic mechanical analysis, and stress-strain measurements with hysteresis loops. The results showed that keratin derived from a current waste product from the poultry industry could effectively and cheaply provide the thermo-mechanical properties required of composite materials ( Pourjavaheri et al., 2018 ). Similarly, Tran and Mututuvari (2016) developed composite materials made from keratin, cellulose, and chitosan combinations. They found that adding cellulose and chitosan improved the mechanical and thermal stability of the overall material but hindered the reformation of α-helices. Instead, when combined with these biopolymers, the keratin preferred the extended β-sheet morphology or amorphous configurations ( Tran and Mututuvari, 2016 ).

Reversible materials

Another attractive characteristic of keratin as raw material is its mechanical reversibility. This reversibility can be found in keratin due to the transition from α-keratin helices to β-keratin sheets. This transition has been observed as a result of stress along the longitudinal axis of the α-helix, as well as heat absorption or from a combination of the two ( Chapman, 1969a ; Haly and Snaith, 1970 ; Hearle, 2000 ; Mason, 1965 ; Miserez et al., 2009 ). Recently, Cera et al. (2021) have captured this reversible process using hydration as a trigger to fabricate 3D-printed, hierarchical shape-memory materials out of keratin extracted from animal hairs. Impressively, this material had a tensile strength and Young's modulus orders of magnitude higher than conventional water-triggered shape-memory materials ( Cera et al., 2021 ).

Fibrillar keratin was extracted from ground Angora wool using LiBr to induce a solid-liquid phase transition of the crystalline keratin and DTT to cleave the disulfide bonds in the hair matrix at 90 C o . The product was then filtered, cooled, centrifuged, and separated to obtain concentrated fibrillar keratin quantities. This extraction process is shown in Figure 21 A. When subject to shear stress and spatial constraint, the extracted keratin protofibrils self-organized into a nematic crystal phase. Adding NaH 2 PO 4 to the extracted keratin allows for tighter control of the nematic phase by introducing a charge screening effect which causes the keratin fibrils to interact more. This process makes the crystalized proteins stiffen and pack closer together with more alignment. The result is a shear-thinning, viscous, keratinous solution that is ideal for extrusion processing and whose properties can be tuned via the NaH 2 PO 4 concentration.

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Advances in 3D printing technology have recently made printing different biological materials more feasible

Cera et al. have utilized these advances to fabricate hydration-induced shape-memory components out of keratin.

(A) The keratin extraction process used to obtain printable, fibrillar keratin ( Cera et al., 2021 ).

(B) To obtain aligned fibrils, keratin fibers were fabricated using traditional wet-spinning. The resultant hierarchical structure is visualized here.

(C) Schematic of the atomic scale process for using water to lock and unlock the hydrogen bonds within α-helices or between the β-sheets. This mechanism endues the material with shape recovery properties.

(D) Images of the keratin printing process and final products (left); SEM image of the fine detail that can be obtained; birefringence images showing the alignment of the keratin fibers in the woven structure.

(E) Series of still images of the hydration-induced shape recovery of the printed samples composed of keratin, showing the prints returning to their initial form over a matter of seconds when submerged in water. Reproduced with permission ( Cera et al., 2021 ). Copyright 2020, the authors.

To maximize uncoiling when loaded and to improve tensile strength and strain-to-failure, α-helices were aligned using traditional wet-spinning. Spun fibers were exposed to hydrogen peroxide to restore the disulfide network of the keratin. Figure 21 B shows that the keratinous fibers (∼10 micrometers in diameter) maintained a hierarchical structure, with a core composed of fibrils that are approximately 50 nm in width. When stretched in the wet state, the α-helices unwind into β-keratin sheets. As the fibers dry while under a constant load, hydrogen bonds begin to form between the β-sheets, fixing them in place and making them metastable. In fact, when stretched to 80% strain and held in place for 10 min at room temperature, the fibers only shrunk back to 77% strain, showing the efficacy of these hydrogen bonds for locking the keratinous fiber into its new fixed shape. Upon rehydration, the hydrogen bonds are disrupted, and the fiber can return to its original shape. This process is visualized in Figure 21 C.

Owing to the shear-thinning properties of the keratinous solution, small diameter extrusion needles can be used to print different geometries with textural features on a scale of 50 micrometers. The keratinous material was printed into a hydrogel which served as support, as well as the coagulation bath. The keratin protofibrils aligned themselves along the print pathway, allowing finer control of the material's shape memory properties. Figure 21 D shows the 3D printing process to fabricate a flat star, ring, and flat strip. The middle image shows an SEM of the fine details that could be produced. The images on the right are birefringence images that show the common alignment of the keratin protofibrils. Once the keratinous material has been printed, it can be further manipulated into new shapes before the disulfide network is reformed by exposing the print to hydrogen peroxide. Figure 21 E shows a square print that was folded into an origami star shape before the disulfide network was reformed. Once the star shape is set with the hydrogen peroxide, water can be used to trigger shape recovery even when it has been deformed into a tube. In this case, it takes less than 2 min for the tube to recognizably transform back into the star origami arrangement, as seen in Figure 21 E ( Cera et al., 2021 ).


This review aims to establish a link between keratin as a fibrous biopolymer and as a material of engineering interest due to its wide-ranging functionality. Keratin fills many different niches in nature due to its inherent properties and its geometric tailorability on multiple length scales derived from its self-assembled hierarchical structure. We established the importance of each of these aspects by exploring keratin as a source of design inspiration alongside the keratin as a raw material for engineered systems.

Keratinous systems have been used to inspire materials with mechanical, thermal, reversible adhesive, lightweight, structural color, and hydrophobic characteristics. These bioinspired designs have not only been used to understand the success of biological materials better but have served also as a creative platform for researchers to extend natural design ideas beyond the limitations of nature, laying the groundwork for the next generation of functional materials. Keratin also has been used as filler or reinforcement in composites with an eye toward environmentally sustainable production and specific biomedical applications. Keratin's prolificity in the industrial world in wool and feathers alongside its beneficial material properties makes it a desirable constituent for expensive components like biomedical materials or fiber-reinforced composites.

Future directions

Keratin has a lot to offer to the scientific and engineering communities, but several obstacles need to be overcome to convert its propitious potential into reality. Here, we suggest several future directions to maximize the impact of keratinous materials on the engineering and scientific communities:

  • ○ As discussed in Sections 1 and 3 , 1 and 3 , keratin has a hierarchical structure that allows for tailorable material properties. When manufacturing bioinspired components, it can be challenging to find a material that matches the properties (i.e., Young's Modulus, strength, toughness, viscoelasticity, conductivity, density, and others) of natural keratin. This can make translations of natural keratinous designs to synthetic systems challenging. Recent developments in the 3D printing of keratin ( Cera et al., 2021 ) have the potential to eliminate this issue by allowing bioinspired designs to be printed using keratin.
  • ○ As shown in Figures 1 and ​ and2, 2 , keratin has an inimitable hierarchical structure that plays an important role in its extensive functionality, i.e., atomic-scale hydrogen bonds in the amino acids make keratin's properties highly tunable via moisture alongside the nanoscale α-helices, which allow for a phase transition at 20% strain while mesoscale features like lamellae, spinules, or spatulae, toughen, repel water, or adhere to surfaces, respectively. Engineers have struggled to replicate the multiscale ordered arrangements found in keratinous systems that help them be so multifunctional, and this remains a major challenge for the field going forward.
  • ○ While there has been significant research on various keratinous systems, there are other keratinous materials that have not yet been studied, particularly amongst reptiles and birds. Much of the research on keratin has revolved around its role in wool, hair, or human skin, which all possess the α-keratin. However, much less is known about β-keratin. Further, each keratinous system bears its unique structure optimized for its role in an organism. Exploring more keratinous systems will continue to reveal new design motifs and inspiration for engineered materials.
  • ○ Similarly, some keratinous systems have been explored, but few attempts have been made to replicate their structure in synthetic materials. These include pangolin scales, butterfly cocoons, nails, talons, claws, and beaks, amongst others.
  • ○ Harrington et al. (2016) eloquently state: “in the case of biological materials, a battery of selective pressures encountered over the evolutionary history of the organism influence the final product,” and as such biological materials are always multifunctional ( Harrington et al., 2016 ). However, engineers often replicate these materials with a singular objective in mind, ignoring the tremendous benefits of a multifunctional material. An exception is the gecko pad, where researchers have perused its reversible dry adhesion, self-cleaning capabilities, and toughness ( Boesel et al., 2010 ; Li et al., 2019 ). Taking a multifunctional approach to each bioinspired design could help to develop superior materials that can be used for numerous applications at once.
  • ○ A vast majority of the work on bioinspired keratinous materials has been done at the macro-, meso-, or micro-scale and is often scaled up for fabrication. An increased focus on generating these structures at their natural length scale could help recapture the original material's properties. Similarly, a broader thrust in exploring the nanoscale behavior of keratin could help develop hierarchical materials or unlock further functional mechanisms that larger-scale experiments have not revealed.
  • ○ Modeling is a beneficial way to understand the structure-property relationships, particularly for a complex biopolymer like keratin. Improved models would help to understand better the hierarchical synergies in keratin and which design parameters are most important for different functionalities.

Listed above are just some of the possibilities for future work on keratin as an engineering material. However, this list is not necessarily specific to keratin. Many other biopolymers like collagen, elastin, and chitin have similar wide-ranging usages in nature. Uncovering what niches each of these biopolymers can fill, how they succeed in so many different environments, and using them in engineered materials will provide a wealth of knowledge to the engineering community. All of this comes with the added benefit of biopolymers being renewable resources. With so many different utilities, understanding and replicating keratin-like structures has the potential to touch every corner of society.

Limitations of the study

This study is not a comprehensive review of all applications of keratin, particularly in the biomedical field. Further, certain popular research topics discussed above, such as gecko pads, are not reviewed comprehensively. For a deeper understanding of these areas, the authors refer readers to the cited review articles.


This research has been supported by funding from the National Science Foundation Mechanics of Materials and Structures program (Grant numbers 1926353 and 1926361). Discussions with Drs. Bin Wang, Tarah Sullivan, Yasuaki Seki, Wei Huang, Ekaterina Krutsik (Novitskaya), and Prof. Joanna McKittrick have been extremely helpful at reaching the conclusions presented here. We dedicate this review to Prof. McKittrick's memory.

Declaration of interests

The authors declare no competing interests.

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  1. Keratin: Protein, Structure, Benefits, Uses & Risks

    Keratin is a protein that helps form hair, nails and your skin's outer layer ( epidermis ). It helps support your skin, heal wounds and keep your nails and hair healthy. There are 54 kinds of keratin in your body. There are two types: Type I: Of the 54 kinds of keratins in your body, 28 of them are type I.

  2. Keratin

    Keratin ( / ˈkɛrətɪn / [1] [2]) is one of a family of structural fibrous proteins also known as scleroproteins. Alpha-keratin (α-keratin) is a type of keratin found in vertebrates. It is the key structural material making up scales, hair, nails, feathers, horns, claws, hooves, and the outer layer of skin among vertebrates.

  3. Keratin

    keratin, fibrous structural protein of hair, nails, horn, hoofs, wool, feathers, and of the epithelial cells in the outermost layers of the skin. Keratin serves important structural and protective functions, particularly in the epithelium.

  4. Keratin: Types, Structure, Benefits, Uses, and Safety

    Keratin is a protein in the cells on the surface of the skin. The fingernails, hair, and skin need keratin to grow, function, and stay healthy. Cosmetic treatments to improve hair and nail health are often enriched with keratin. Keratin also occurs naturally in some foods and can be taken in supplement form as well.

  5. Keratin: Structure, mechanical properties, occurrence in biological

    A ubiquitous biological material, keratin represents a group of insoluble, usually high-sulfur content and filament-forming proteins, constituting the bulk of epidermal appendages such as hair, nails, claws, turtle scutes, horns, whale baleen, beaks, and feathers.

  6. Structure and functions of keratin proteins in simple, stratified

    Historically, the term 'keratin' stood for all of the proteins extracted from skin modifications, such as horns, claws and hooves. Subsequently, it was realized that this keratin is actually a mixture of keratins, keratin filament-associated proteins and other proteins, such as enzymes.

  7. Brief introduction of keratin and its biological application ...

    1.1 Source of keratin. Keratin is the group of intermediary protein that shows in mammalian tissue configurations (hair, fur, nails, skin, wool, hooves, and horns) and birds (e.g., bird beaks and feathers) [1-3], which plays an important role in protecting the body (Fig. 1).It has a higher content in the epithelial tissue of higher vertebrates and mainly derived from the structural protein ...

  8. Biological importance and pharmaceutical significance of keratin: A

    Keratin is the most common structural protein in epithelial cells [20], and it is the most significant biopolymer in animals [21]. Furthermore, Keratin is one of the most rigid biological materials, with exceptional hardness and modulus despite being made entirely of polymeric elements and rarely containing minerals [22]. Keratinous materials ...

  9. Keratin: Structure, mechanical properties, occurrence in biological

    According to the Ashby map [13], shown in Fig. 1, keratin is among the toughest biological materials, possessing both high toughness and high modulus, although it is solely composed of polymeric constituents, and seldom contains minerals [14].

  10. The human keratins: biology and pathology

    The keratins are the typical intermediate filament proteins of epithelia, showing an outstanding degree of molecular diversity. Heteropolymeric filaments are formed by pairing of type I and type II molecules. In humans 54 functional keratin genes exist.

  11. A Review of Keratin-Based Biomaterials for Biomedical Applications

    The biological activity of keratin films was also increased by incorporating a cell adhesion peptide, Arg-Gly-Asp-Ser (RGDS), at the free cysteine residues of reduced keratin extracts. RGDS-carrying keratin films proved to be excellent substrates for mammalian cell growth, and this work again demonstrated the potential and versatility of ...

  12. Keratin-mediated hair growth and its underlying biological ...

    Keratin is a cytoskeletal protein that forms intermediate filaments within epithelial cells and participates in maintaining the strength of the cells 1.It is a major protein found within the hair ...

  13. What You Need to Know About Keratin Treatments

    Keratin is a type of protein that makes up the outer surface of your hair, skin and nails. "As you age, you lose keratin. It gets broken down or becomes damaged," explains Dr. Vij. "Overtime, factors like environmental triggers, UV chemicals and heat can break down keratin. Those factors basically cause the protein to change its natural shape."

  14. Keratin

    [ˈkɛrətɪn] Definition: Structural protein found in hair, nails, and outer layer of skin Table of Contents Keratin is a fibrous protein naturally present in hair, skin, and nails. In hair care, it serves as a protective and structural element, enhancing strength and smoothness while reducing frizz.

  15. Keratin as a Biopolymer

    Keratin has recently gained a lot of limelight among various proteins. Keratin is a renewable, sustainable, biocompatible, and biodegradable bioresource, and these characteristics make it a promising candidate for diverse applications. It is a fibrous protein mainly found in feathers, hair, wool, animal claws, and fingernails.

  16. Keratin

    Keratin constitutes the major component of the feather, hair, hooves, horns, and wool represents a group of biological material having high cysteine content (7-13%) as compared to other structural proteins. Keratin -based biomaterials have been investigated extensively over the past few decades due to their intrinsic biological properties and ...

  17. Keratin-rich foods and their benefits

    Keratin is a protein that helps maintain the structure of hair, nails, skin, and the lining of the internal organs. Certain nutrients support keratin production. ... Can 'biological race' explain ...

  18. Keratin

    Keratin is found in the human skin and the epidermal appendages of animals. It is a very viable and cost-effective protein for biomedical applications. Keratin-based biomaterials extracted from wool and human hair are biocompatible, biodegradable, nontoxic and very tunable.

  19. Preparation and applications of keratin biomaterials from natural

    Abstract Keratin is a kind of natural polymer that is abundant in feathers, wool, and hair. Being one of the natural biomolecules, keratin has excellent biological activity, biocompatibility, biodegradability, favorable material mechanical properties, and natural abundance, which exhibit significant biological and biomedical application potentials. At present, the strategies commonly used for ...

  20. Extraction and application of keratin from natural resources: a review

    Keratin belongs to the family of fibrous structural proteins called scleroproteins. It is the most abundant structural protein found in hair, nails, feathers, horns, claws of animals; along with collagen, it is the most important biopolymer encountered in animals.

  21. Keratin Structure, Function & Diseases

    A keratin protein is an intermediate filament used to provide structural integrity to the hair, skin, and nails. Proteins are made up of amino acids. Where is keratin produced? Keratin is...

  22. Engineering with keratin: A functional material and a source of

    Keratins are broadly classified as having either α- or β-ultrastructures (Figure 1).Typically, mammalian keratin is found in the α-keratin form, while avian and reptilian keratins are β-keratin types; however, one mammal, the pangolin, is known to have both α- and β-keratin domains in its scales (Wang et al., 2016c).Like all biological materials, both α- and β-keratinous materials form ...

  23. Engineering with keratin: A functional material and a source of

    Keratin is a ubiquitous biological polymer comprising the bulk of mammalian, avian, and reptilian epidermal appendages, including nails, hair, the outer layer of skin, feathers, beaks, horns, hooves, whale baleen, claws, scales, hagfish slime, and gecko pads ...