Abstract
Tissue regeneration is a process that recapitulates and restores organ
structure and function. Although previous studies have demonstrated
wound-induced hair neogenesis (WIHN) in laboratory mice (Mus), the
regeneration is limited to the center of the wound unlike those
observed in African spiny (Acomys) mice. Tissue mechanics have been
implicated as an integral part of tissue morphogenesis. Here, we use
the WIHN model to investigate the mechanical and molecular responses of
laboratory and African spiny mice, and report these models demonstrate
opposing trends in spatiotemporal morphogenetic field formation with
association to wound stiffness landscapes. Transcriptome analysis and
K14-Cre-Twist1 transgenic mice show the Twist1 pathway acts as a
mediator for both epidermal-dermal interactions and a competence factor
for periodic patterning, differing from those used in development. We
propose a Turing model based on tissue stiffness that supports a
two-scale tissue mechanics process: (1) establishing a morphogenetic
field within the wound bed (mm scale) and (2) symmetry breaking of the
epidermis and forming periodically arranged hair primordia within the
morphogenetic field (μm scale). Thus, we delineate distinct
chemo-mechanical events in building a Turing morphogenesis-competent
field during WIHN of laboratory and African spiny mice and identify its
evo-devo advantages with perspectives for regenerative medicine.
Subject terms: Biophysics, Cell biology, Developmental biology, Stem
cells, Skin diseases
__________________________________________________________________
How hair follicle regeneration arises readily in some species ie. spiny
rather than laboratory mice, is unclear. Here, authors compare them,
showing an optimal stiffness is needed for placode formation and the
difference in hair follicle regenerative behaviour after wounding is
linked to Twist1.
Introduction
The ultimate goal of regenerative medicine is to restore the function
and structure of the original tissue. Wound healing in adult humans and
mice generally undergoes re-epithelialization successfully yet fails to
develop further, resulting in a scar with excess collagen and an
absence of other skin appendages such as hair follicles. To facilitate
regenerative wound healing, we look into skin development to
recapitulate principles of hair follicular neogenesis.
Patterns form with the break of homogeneity and lead to the emergence
of new structure or arrangement^[58]1. In skin development, Turing
reaction-diffusion was shown to be involved in the periodic pattern
formation of feathers and hairs^[59]2,[60]3. Yet, before periodic
patterning occurs, a morphogenetic field competent for Turing mechanism
must take place; this should have proper cell density, the ability to
secret morphogens, and appropriate morphogen receptors^[61]4. Within
the morphogenetic field, FGFs, Wnt/β-catenin and Edar^[62]5–[63]7
signaling activate the epithelial cells to aggregate and form hair
placodes, which later interact with the dermal condensate (DC) and
invaginate into the dermis to form the foundation of a hair follicle.
This process is characterized by a series of cohesive molecular
signaling and also physical cellular events such as cell aggregation,
collective cell migration and proliferation. Wnt/β-catenin signaling
has been shown imperative to progress these cells into morphogenesis,
in which a series of other signaling molecules such as Lef1, Sox2,
Edar, Shh, MMP and Twist2 are also expressed^[64]8,[65]9.
Dynamic mechanical changes also occur during
morphogenesis^[66]10,[67]11. For epithelial cells to collectively
migrate during morphogenesis, there must be an emergence of a local
active stress acting at cell-cell or cell-matrix interfaces that
creates an anisotropic force field^[68]12–[69]14. Force generation by
myosin-II motors on actin filaments have been shown to drives cell and
tissue morphogenesis in drosophila embryonic development^[70]15. In
other words, in order for hair placode to form and invaginate, the
epithelial cells must overcome the physical barrier provided by the
dermal cells and the extracellular matrix (ECM) in order to invaginate
into the dermis. While previous studies uncovered mechanisms that can
turn on/off hair follicle development, the chemo-mechanical dynamics
that allows epithelial placode to form and invaginate into the dermis
is largely unknown. Previously, Oster, Murray and Harris constructed a
mathematical model that described the action of motile cells that could
produce stress on their environment and thereby produce heterogeneous
spatial patterns through mechanical means^[71]16. This basic theory
assumes there are two populations: a motile population of cells, n,
which can produce stress; and a tissue substratum, the ECM, which has
density. Critically, the ECM is treated as a viscoelastic material,
meaning that the ECM will deform subject to the traction forces
produced by the cells. Critically, if the forces are added and removed
quickly the ECM will relax back to its original shape. The convergence
of these morphological and molecular asymmetries lead to the formation
of DC^[72]17 and activation of β-catenin in the adjacent epidermal
cells to initiate feather bud gene expression^[73]18,[74]19.
In the avian skin development, the early formation of the morphogenetic
field is recognized as the feather tract field, and individual buds
form sequentially or simultaneously within the tract field, with some
species-based differences^[75]4,[76]18,[77]20. This implies there are
different ways to make competent morphogenetic fields^[78]21, while the
region outside of the tract field becomes the apteric region. Thus, in
skin development, periodic generation of skin appendages occur in two
steps: first the formation of morphogenetic field and then the periodic
patterning of cell collectives within the field.
Wound-induced hair follicle neogenesis (WIHN) is a regenerative outcome
of wound healing where fully function hair follicles develop de novo
from the center of large full thickness excisional wounds^[79]22. This
observation was originally described in rats, rabbits, sheep and even
humans, and investigated in depth in recent years^[80]23–[81]26. Cells
in wound bed may use different paths to reach the morphogenetic
competent state. We contemplate that to regenerate new hair follicles
in the wound bed of the adult skin, a morphogenetic field also has to
be established first, which then allows periodically arranged hair
germs to be generated. Since adult cells have different epigenetic
landscapes from embryonic cells, the generation of hair placodes in the
adult may not follow the exact pathway as in development. Investigating
the chemo-mechanical dynamics of the epithelial cells and the ECM
during hair follicular neogenesis would facilitate our understanding on
how to set up the morphogenetic field and initiate the signaling
events, which would have translational potential.
In search for the ultimate regeneration model, the African spiny mouse
(Acomys cahirinus) serves as an adequate model to study complete skin
regeneration. The spiny mouse has evolved to give away up to 70% of its
skin to its predator and still remarkably regenerates its entire skin
and appendages^[82]27–[83]29. Its skin is mechanically softer (20
times) than the laboratory mouse (Mus musculus) and much easier to
break (77 times less energy required)^[84]27. The gene expression
profiles of the spiny mouse wound show a dampened response of collagen,
MMP and inflammatory molecules after wounding, suggesting an
alternative microenvironment to enrich hair neogenesis^[85]30. On the
other hand, studies in laboratory mice have shown that traction, or
tension across the skin or wound, causes hypertrophic scar through FAK
signaling^[86]31, implying the significance of tensile state of the
connective tissue to tissue functioning^[87]32,[88]33. Nevertheless,
the initiating events and how these findings can translate into our
understanding in laboratory mice and human remain to be explored. Thus
we hypothesize that spatial tissue mechanics of the wound partake in
establishing the morphogenetic field for hair follicular neogenesis,
and there lies a chemo-mechanical signaling event that initiates the
symmetry breaking of the epidermis and leading to placode formation and
invagination. We show Twist 1 pathway plays a key role in modulating
tissue stiffness and facilitate hair formation. Furthermore, by
delineating the common and distinct features of laboratory and spiny
mouse during WIHN, we learn from evo-devo advantages to provide
perspective for future implications.
Results
Tissue mechanics set up morphogenetic field for wound-induced hair neogenesis
To examine the effects of tissue mechanics on wound induced hair
neogenesis, we first created 1 × 1 cm full thickness wounds on the
dorsal skin of 3-week-old C57Bl/6 mice, and observed new hair follicles
formed at the center of the wound on post-wound day 28 (PWD28,
Fig. [89]1a–c). To investigate the spatial stiffness distribution of
the wound bed, we used an atomic force microscopy (AFM)^[90]34 to
measure across the wound (Supplementary Fig. [91]S1a), and calculated
tissue stiffness from force–displacement curves (Supplementary
Fig. [92]S1b, c) using a modified Hertz model^[93]35,[94]36. We found
that the center of the PWD14 wound, where de novo hair follicles can be
observed, is significantly softer than the wound periphery (28.0 ± 1.1
vs 10.5 ± 0.6 kPa, Fig. [95]1d, e).
Fig. 1. Tissue mechanics set up morphogenetic field for wound-induced hair
follicle neogenesis.
[96]Fig. 1
[97]Open in a new tab
a A PWD28 C57Bl/6 mouse. b AP staining showing de novo hair follicle
formation at the center of the wound bed at PWD28. c Schematic diagram
showing the location of regenerated hair follicles (blue dots) in b. d
Stiffness heatmap overlaying the PWD14 wound. Colorimetric unit: kPa. e
Cross-sectional view of the PWD14 wound and distribution of wound
stiffness. Yellow arrow heads indicate the formation of hair placodes.
Heatmap and number indicate the spatial stiffness of the whole wound
bed. Unit: kPa. Green arrow indicates the range of the morphogenetic
field. f A Bleb-treated mouse on PWD28. g AP stain showing the number
of hair follicles increased significantly upon Bleb treatment. h
Schematic diagram of hair follicles (blue dots) in g. i Stiffness
heatmap of PWD14 Bleb-treated overlaying wound. j Dot plot showing
changes in HF upon Bleb treatment. n = 8 biologically independent
animals. Data are presented as mean values ± SEM. p = 0.003, unpaired
two-sided t-test. k The area of the wound bed under 15 kPa positively
correlates with the number of hair follicles. SHG of l PWD14 and m
PWD21 wounds. The color squares indicate the location of corresponding
enlarged photos from n PWD14 center, o PWD14 margin, p PWD21 center and
q PWD21 margin of the wound image. r Dot plot showing the number of
fibers per square unit. PWD14 C/M: p = 0.0323, PWD14/PWD21 C: p =
0.0495, PWD21 C/M: p < 0.0001. s Fiber thickness in respective wound
time and location. PWD14 C/M: p = 0.0454, PWD14/PWD21 C: p = 0.0020,
PWD21 C/M: p < 0.0461. r, s Data are presented as mean values ± SD.
n = 8 regions examined over four biologically independent animals.
One-way ANOVA, Tukey test. t Summary graph of wound stiffness on PWD14
and PWD21 with respect to wound location and Bleb treatment. n = 10
regions examined over five biologically independent animals per
location per condition. Data are presented as mean values ± SD.
*p = 0.0478; **p = 0.0061; ***p < 0.0001. One-way ANOVA, Tukey test. u
Illustration of tissue mechanics partake in setting up the
morphogenetic field for WIHN. Red line: wound margin. Green: soft,
morphogenetic field. Blue: hair placodes. Ctl: control; Bleb:
Blebbistatin; HF: hair follicle; sq: square; Lt: left; Rt: right;
Morph: morphogenetic. The images in a–i represent 8 out of 8
experiments performed. The images in l–q represent 4 out of 4
experiments performed.
To evaluate whether the observed tissue stiffness plays a role in hair
follicle neogenesis, we treated the wound daily from PWD10 to PWD16
with 100 μM of Blebbistatin (Fig. [98]1f), and observed an increase in
the number of hair neogenesis (Fig. [99]1g, h) and decrease in the
stiffness of the wound bed (Fig. [100]1i). Statistically, Blebbistatin
significantly increased the number of resultant hair follicles from
17.4 ± 2.1 to 34.5 ± 4.3 kPa (Fig. [101]1j, n = 8), and significantly
increased the area of the wound bed that is under 15 kPa from
2.38 ± 0.13 to 3.70 ± 0.26 mm^2 (Supplementary Fig. [102]S1d). To
correlate tissue stiffness with hair neogenesis, we quantified and
plotted the area of the wound under 15 kPa versus the number of hair
neogenesis observed in each wound, and found a strong correlation
(R^2 = 0.8082), suggesting stiffness may contribute to creating the
morphogenetic field for hair neogenesis (Fig. [103]1k).
We further explored the molecular constituents of the wound stiffness.
Collagen has been implicated as the main ECM of the wound^[104]37,
hence we used second harmonic generation (SHG) to visualize the amount
and organization of collagen fibrils in the wound (Fig. [105]1l–q).
Generally speaking, fibers in PWD14 wounds are thinner and less dense
than in PWD21 ones. Both the fiber density and thickness are
significantly higher in the wound margin than in the wound center in
PWD14 and PWD21 (Fig. [106]1r, s). Hence, the spatiotemporal
organization of collagen fibrils in the PWD14 and PWD21 wound beds
corresponds to its respective stiffness of the wound (Fig. [107]1t).
This implies the wound bed needs to be softer than 15 kPa for hair
neogenesis to occur (green bar, Fig. [108]1t). Blebbistatin treatment
not only softened the entire wound bed, but also more importantly,
significantly lowered the stiffness of intermediate margin-center
region of the wound (blue and purple lines, Fig. [109]1t), hence
setting up a larger morphogenetic field competent for hair neogenesis
(green bar, Fig. [110]1u). These results suggest tissue mechanics play
an important role in hair neogenesis, and the spatial organization of
collagen fibrils may be the main constituent of tissue stiffness in the
wound bed.
African spiny mice exhibit an optimal range of tissue stiffness for placode
formation
The African spiny mice are known to have robust ability in WIHN
(Fig. [111]2a). Unlike laboratory mice that show de novo hairs only in
the center of the wound bed, new hair follicles were observed across
the entire wound bed on PWD28 (Fig. [112]2b, c). To explore how these
events occur over time, we examined the spatiotemporal pattern of hair
placode emergence in spiny mice. Interestingly, we found an opposite
trend. In spiny mice, the hair placodes started to develop from the
periphery of the wound bed on PWD14 (Figs. [113]2e, e1–3). The center
of the wound bed did not form hair placodes until PWD21 (Fig. [114]2f).
How can we explain this opposite trend? Can tissue mechanics play a
role?
Fig. 2. African spiny mice exhibit an optimal range of tissue stiffness for
placode formation.
[115]Fig. 2
[116]Open in a new tab
a A spiny mouse on PWD28. b K15 wholemount immunostaining of a PWD28
spiny mouse wound. Red line: border of the wound bed. c Schematic
diagram of hair follicles (blue dot) in b. d The stiffness heatmap
overlaying the PWD14 spiny mouse wound. Colorimetric unit: kPa. e H&E
histology and stiffness heatmap of PWD14 and f PWD21 wounds. The color
and number indicate the stiffness of the wound at respective location.
Unit: kPa. (e1–e3) enlarged images of regions 1, 2, and 3 from e. g H&E
of the PWD28 wound. Yellow arrows indicate the formation of hair
placode. h SHG of PWD14 and PWD21 spiny mouse wounds. Red line: wound
border. i, IHC of collagen I and collagen III in spiny mouse at
different post-wound time. j Blebbistatin treatment significantly
reduced the resultant number of hair neogenesis. n = 5 biologically
independent animals. Data are presented as mean values ± SEM.
p < 0.0001, unpaired two-sided t-test. k Graph indicating the
respective wound stiffness of the specific wound location and time in
the laboratory and spiny mice. n = 10 regions examined over 5
biologically independent animals per location per condition. Data are
presented as mean values ± SD. ^###p < 0.0001 when compared between Mus
PWD14 and Acomys PWD14; *p = 0.0482 when compared between Mus PWD14 and
Mus/Bleb PWD14 center, p = 0.0218 when compared between Mus PWD14 and
Mus/Bleb PWD14 at the right of wound center, p = 0.0441 between Acomys
PWD14 and Acomys PWD21 at the left of wound center, p = 0.0381 when
compared between Acomys PWD14 and Acomys/Bleb PWD14 wound margin right,
p = 0.0421 at center-right, **p = 0.0089; ***p < 0.0001. One-way ANOVA,
Tukey test. l Illustration of morphogenetic field in PWD14 and PWD21
spiny mouse wound bed. Red: wound border. Green: morphogenetic field.
^###: p < 0.005. Ctl: control; Bleb: Blebbistatin; Lt: left; Rt: right;
Morph: morphogenetic. The images in a–g, i represent five out of five
experiments performed. The images in h represent 4 out of 4 experiments
performed.
We used an AFM to determine the spatiotemporal dynamics of tissue
stiffness during WIHN in spiny mice (Fig. [117]2d). In general, two
trends are similar to that of the laboratory mice: (1) the center of
the wound bed is softer than the wound periphery (Fig. [118]2d), and
(2) the overall stiffness of the wound increased from PWD14 to PWD21
(Fig. [119]2e, f). However, there are two major differences: (1) the
soft nature of the unwounded spiny mice skin and wound bed, and (2) the
periphery-to-center formation pattern of de novo hair primordia in the
spiny mice wounds. The unwounded spiny mouse skin and the stiffest
region of its wound bed, the wound margin, are still softer than
15 kPa, while the center of the wound bed is below 5 kPa on PWD14
(Fig. [120]2e). As a result, the hair primordia formation in the spiny
mouse wound followed a periphery to central trend; the hair placodes
only began to form where the wound stiffness was higher than 5 kPa
(Fig. [121]2f). Hence, the de novo hair follicles were more mature at
the wound periphery than at the wound center, and this morphological
feature is apparent on PWD28 (Fig. [122]2g). This result is best
explained by the presence of an optimal range between 5 and 15 kPa of
tissue stiffness for placode formation. To test this hypothesis, we
further softened the spiny mouse wound bed by Blebbistatin treatment
and showed that: (1) Blebbistatin treatment softened the wound bed and
significantly increased the wound area under 5 kPa (Supplementary
Fig. [123]S2a–c), and (2) the resultant number of hair neogenesis was
significantly reduced from 74.00 ± 6.14 to 12.40 ± 1.10 (Fig. [124]2j,
Supplementary Fig. [125]S2d). These results suggest that regions of
the wound bed that are either too stiff or too soft are not favorable
for new hair formation.
What is the molecular basis of these differences in tissue stiffness?
To investigate the structure and organization of collagen fibrils in
the spiny mouse wound as observed in the laboratory mice, we also used
SHG to visualize it. Interestingly, collagen fibrils were identified in
the unwounded spiny mouse skin but were almost undetectable within the
wound center in both PWD14 and PWD21 wounds (Fig. [126]2h). We analyzed
and compared the fiber thickness and fiber density of PWD14 and PWD21
spiny mouse wound margin (Supplementary Fig. [127]S2e, f), and showed
that the collagen fibrils in the spiny mouse wounds are significantly
thinner and less dense than that of the laboratory mice (Supplementary
Fig. [128]S2g, h). The spiny mouse skin has been reported to have high
levels of collagen III^[129]27; however, since type III are less
crystallined and generate little SHG signal^[130]38,[131]39, we further
used IHC to examine the expression of collagen I and III in the spiny
mouse wounds. We found that collagen III is the dominant collagen type
expressed in the early stages of spiny mouse wound healing
(Fig. [132]2i); nevertheless, this relationship is reversed later in
post 16-week wounds when collagen I was highly expressed in contrast to
collagen III (Supplementary Fig. [133]S2i). To summarize, we found that
there is a lower limit on the softness of the wound bed at 5 kPa in
order for hair neogenesis to occur, as the initial hair placodes can
only be observed in wound stiffness between 5 and 15 kPa in both
laboratory and spiny mice wounds (Fig. [134]2k). This implies that
5–15 kPa could be the optimal range for the wound to set up the
morphogenetic field for hair placode formation, and suggests why hair
neogenesis begins from the wound periphery and later to center in the
spiny mice (Fig. [135]2l).
RNA-seq analysis identifies epidermal Twist1 as an upstream regulator for the
formation of hair placodes in WIHN
To delineate the molecular mechanism underlining WIHN, specifically at
the morphogenetic field, we used a 5 mm punch biopsy to separate the
PWD14 epidermis into the regenerative wound center and the
non-regenerative wound margin, and performed RNA-seq analysis to look
for differentially expressed genes (DEG) between the two. PWD14 was
selected because our preliminary data showed that gene expression
related to hair neogenesis (Wnt5a, Lef1, Gli1, Fgf10, and Twist1)
peaked in the wound center of PWD14 epidermis (Supplementary
Fig. [136]3a, b). From this RNA-seq analysis, we identified 2780 DEG
(Fig. [137]3a). Among them, Wnt/β-catenin signaling (P = 1.49 × 10^−2),
Integrin signaling (1.84 × 10^−8), Stat3 pathway (1.01 × 10^−6),
inhibition of Matrix Metalloproteases (4.53 × 10^−7), EMT core genes
(4.23 × 10^−10), proliferation of epithelial cells (3.47 × 10^−17),
cell movement of epithelial cells (5.43 × 10^−13), and organization of
ECM are significantly enriched (Fig. [138]3b, c, Table [139]1).
Furthermore, we identify 20 significantly upregulated genes related to
hair placode formation (Table [140]2).
Fig. 3. Transcriptome analysis identifies Twist1 as an important
transcription factor during WIHN in laboratory mice.
[141]Fig. 3
[142]Open in a new tab
a Gene expression heatmap of PWD14 epidermis wound center vs wound
margin. b The significantly enriched pathways of DEG. c, Volcano plot
showing gene expression fold changes of representative DEG in MMP,
Twist1, Wnt/β-catenin related pathways, proliferation of epithelial
cells, cell movement of epithelial cells. Upregulation indicates high
expression in wound center. FC: fold change. C/M: wound center versus
margin. d Twist1 ranked 2nd by two-sided Fisher’s Exact Test p value in
the top ten differentially expressed upstream regulator. e IPA
identifies Twist1 as the top upstream regulator of the downstream DEG
(p = 1.1 × 10^−21). f Wholemount immunostaining of Twist1 at respective
location from the epidermal side of the PWD14 wound as illustrated. Red
line: wound border. Green line: morphogenetic zone. Blue dots: hair
placode. g IHC of Twist1 and Snai1 in PWD14 wounds containing the hair
placode. Dotted line demarcates the border of dermal epidermal
junction. Blue: Hoechst. Scale bar: 100 μm. a–d n = 2 biologically
independent experiments. The images in f, g represent 4 out of 4
biologically independent experiments performed.
Table 1.
The significantly enriched pathways in PWD14 epidermis wound center vs
margin RNA-seq analysis.
Enriched Pathway p-value Differentially expressed genes
Wnt/β-catenin signaling (X30) 1.49 × 10^−2 appl2, bmpr2, cdh5, csnk1d,
dkk3, dvl1, ep300, fzd2, gsk3b, hdac1, kremen2, lrp1, lrp6, ppard,
ppp2r1a, ppp2r2c, ppp2r3a, ppp2r5b, ptpa, sfrp1, sox15, sox18, src,
tgfb1, tgfb3, tle3, wnt16, wnt10a, wnt10b, wnt5a
Regulation of EMT Pathway (X28) 1.02 × 10^−2 dvl1, fzd2, hgf, lox,
map2k2, mmp2, mmp9, notch4, pard6b, pdgfrb, ralb, smad3, smurf1, snai1,
tgfb1, tgfb3, twist1, twist2, wnt10a, wnt10b, wnt5a, wnt9b
MMP genes (X1) 4.53 × 10^−7 adam12, hspg2, lrp1, mmp2, mmp3, mmp9,
mmp10, mmp11, mmp13, mmp14, mmp17, mmp19, mmp23b, sdc1, thbs2, timp1,
timp3
Organization of ECM (X84) 1.32 × 10^−23 adam12, adamts4, agrn, apbb2,
aplp1, bgn, bmp1, c6orf15, col16a1, col18a1, col1a1, col1a2, col27a1,
col3a1, col4a1, col4a2, col5a1, col5a2, col5a3, col6a1, col6a2, col6a3,
col6a4, col7a1, col8a1, ctsk, dcn, ddr1, egflam, elf3, emilin1, fbln5,
fbn1, fbn2, fn1, furin, ibsp, icam2, itga1, itga11, itga3, itga5,
itga9, itgb3, itgb6, jam2, kdr, lama4, lox, loxl1, mfap2, mmp1, mmp10,
mmp11, mmp13, mmp14, mmp19, mmp2, mmp3, mmp9, nid1, nid2, olfml2a,
olfml2b, pdgfra, pecam1, postn, prdx4, ptx3, pxdn, serpine1, sh3pxd2b,
sparc, spp1, timp1, tnc, tnf, vcam1, vcan, vtn, vwf
Proliferation of epithelial cells (X186) 3.47 × 10^−17 ahr, alms1,
areg, atm, bad, bcl11b, birc2, bnc1, brca1, calm1, casp3, casp8, ccnd3,
cd9, cdc25b, cdc73, cdkn1a, cdkn1b, cebpa, cers2, col8a1, creb3l3,
cryab, csf2rb, ctsv, cul3, cxcr2, dab2ip, eif4e, eng, ep300, epgn,
epha2, ercc1, ereg, esrra, ezh2, fbln5, fgfr2, fn1, frs2, fst, gata3,
glul, grn, gsk3b, has2, hbegf, heyl, hgf, hoxa5, hyal1, ifngr1, ift52,
ift74, ift80, igf1, il18, il22ra2, il24, il4r, il6r, inhba, inhbb,
itga1, itga3, itgb3, junb, kcnk2, kdr, klf10, klf5, klk3, klk6, klk8,
krt16, krt17, lgals7/lgals7b, lgr4, lmnb1, lrp6, maged1, map2k1,
map2k6, map2k7, mapk7, mapk8, mapk9, mapkapk2, marveld3, mfge8, mki67,
mmp14, mmp9, mt2, nab1, nab2, nfib, nfkbia, nme2, npm1, nr3c1, nrg1,
odc1, p2rx7, pkp3, postn, ppard, prlr, ptafr, pten, ptgs2, pthlh,
ptpn1, ptprk, rack1, rbl2, rela, relb, rgn, rida, rps6kb1, s1pr2,
sema4d, serpinf1, sfn, sfrp1, sh2b1, slc20a1, slc7a5, smad3, smad7,
snai2, socs1, socs3, sparc, spint2, spp1, stat5a, stmn1, tfrc, tgfb1,
tgfb3, tgm1, timeless, timp1, tnfaip6, tnfrsf11a, tnfrsf12a, tnfrsf1a,
tslp, twist1, twist2, uhrf1, vegfa, wnt10b, wnt16, wnt5a, yod1, zbtb16
[143]Open in a new tab
Comparison using two-sided Fisher’s Exact Test.
Table 2.
The 20 significantly upregulated genes related to hair placode
formation at epidermis wound center compared to wound margin on PWD14.
Gene Description Location Type(s)
cdh11 cadherin 11 Plasma Membrane other
meg3 maternally expressed 3 Other other
twist1 twist family bHLH transcription factor 1 Nucleus transcription
regulator
col1a1 collagen, type I, alpha 1 Extracellular Space other
col3a1 collagen, type III, alpha 1 Extracellular Space other
col6a2 collagen, type VI, alpha 2 Extracellular Space other
col6a3 collagen, type VI, alpha 3 Extracellular Space other
efemp2 EGF containing fibulin-like extracellular matrix protein 2
Extracellular Space other
fbn1 fibrillin 1 Extracellular Space other
fstl1 follistatin like 1 Extracellular Space other
lgals1 lectin, galactoside-binding, soluble, 1 Extracellular Space
other
timp2 TIMP metallopeptidase inhibitor 2 Extracellular Space other
cpxm1 carboxypeptidase X (M14 family), member 1 Extracellular Space
peptidase
dpysl3 dihydropyrimidinase like 3 Cytoplasm enzyme
ftl1 ferritin, light polypeptide Cytoplasm enzyme
pde5a phosphodiesterase 5A Cytoplasm enzyme
ppic peptidylprolyl isomerase C Cytoplasm enzyme
myh10 myosin, heavy chain 10, non-muscle Cytoplasm other
srpx2 sushi-repeat containing protein, X-linked 2 Cytoplasm other
igf2bp2 insulin like growth factor 2 mRNA binding protein 2 Cytoplasm
translation regulator
[144]Open in a new tab
To identify the potential upstream initiators of WIHN, we identified
114 transcription factors (TF) that are significantly upregulated at
the wound center, including those that are related to the Twist
(Twist1, Twist2, Snai1) and Wnt (Tcf23) pathways (Table [145]3). Among
them, Twist1 is the second most statistically significant upstream
regulator, which regulates many downstream DEG (P = 4.55 × 10^−11) as
indicated by the Upstream Analysis function in Ingenuity Pathway
Analysis (IPA) (Fig. [146]3d, e). The top significant transcription
factor is Nfkbia (3.28 × 10^−13) which is associated with the
inflammatory response after wounding (Fig. [147]3d). Wholemount
immunostaining also showed that epidermal Twist1 is highly expressed in
the PWD14 wound center and enriched in both hair placode and
inter-placode region more than in the margin (Fig. [148]3f,
Supplementary Fig. [149]3c). IHC staining of Twist1 and Snai1 (number 5
ranked TF) on PWD14 sections show both TF are expressed in the
epithelial hair placode (Fig. [150]3g). These findings point to ECM
remodeling, cell proliferation, cell movement, Stat3, and Wnt/β-catenin
as important signaling events during WIHN, and epidermal Twist1 appears
to be an important upstream regulator for hair placode formation.
Table 3.
The 114 significantly upregulated transcription factors at epidermis
wound center compared to wound margin on PWD14.
acap3 cers2 fam129b hr mier2 relb srebf1 wtip
aes churc1 fem1a id3 mllt1 rfx1 srebf2 zbtb42
arid3a cic fiz1 ier2 mnt rfxank srf zdhhc13
arid5a cited4 fosl1 ifi204 mxd1 rnf114 ssbp4 zfp219
asb1 creb3l3 fosl2 irf1 nab2 rnf25 tbx15 zfp369
asb6 csrnp1 foxc1 irf5 nfatc4 rnf4 tbx3 zfp444
atf4 ctbp2 foxp4 irf7 nfkbia sbno2 tcf23 zfp593
atf6b dnmt3l glis2 irx1 noct scand1 tcfl5 zkscan6
atxn7l3 e2f7 glmp jarid2 notch4 siah2 thap4 zxdc
barx2 eaf1 gpank1 junb pax9 smad3 trim16
bhlhe40 ehmt2 hdac1 klf10 phf1 smad7 tsc22d1
bnc1 elf3 helz2 lmo1 pitx1 snai1 tsc22d4
btg2 elk1 heyl lztr1 prrx1 sox15 twist1
carhsp1 elk3 hic2 maff rbpms sox18 twist2
cebpa esrra hopx maged1 rela sqstm1 ube2v1
[151]Open in a new tab
Based on p-value < 0.05.
Given the differences between laboratory and spiny mice during
WIHN^[152]29 (Figs. [153]1, [154]2), we also performed RNA-seq analysis
on spiny mouse wounds to identify key molecules for its hair primordia
formation. Since the entire spiny mouse wound bed is capable of
undergoing WIHN, we harvested the entire wound on PWD0, 14, 21, and 28
and separated them into epidermis and dermis to analyze the
quantitative changes of gene expression before, and during early, mid,
and late stages of WIHN (Fig. [155]4a). From the wound epidermis, the
proliferation of epithelial cells and cell movement associated genes
all peaked on PWD14, including Twist1 (Fig. [156]4b). In the dermis,
Twist1 and its related TF Zeb2 also peaked on PWD14, while Zeb1 and
Tgfb1 showed a different trend (Fig. [157]4c). Interestingly, many of
the ECM and MMP related genes along with other TFs such as Stat3 are
highly expressed on PWD21 and PWD28 (Fig. [158]4d), which may reflect
the softness of PWD14 spiny mouse wounds (Fig. [159]2k). IHC staining
of PWD14 spiny mouse wounds also show that Twist1, Zeb2, MMP9 and P-cad
are expressed around the hair placode, while E-cadherin is
downregulated at the hair placode. As suggested by its RNA-seq
analysis, Twist1, Zeb2 and MMP9 are also expressed in the dermis
(Fig. [160]4e). Lastly, using Harmine and GM6001, we show that
inhibiting Twist1 and pan-MMP activity in the spiny mouse wounds also
significantly reduced WIHN (Fig. [161]4f, g).
Fig. 4. Twist1 is also expressed and can modulate the outcome of WIHN in
spiny mice.
[162]Fig. 4
[163]Open in a new tab
a Gene expression heatmap of spiny mouse epidermis at different
post-wound days. b Proliferation of epithelial cells and cell movement
associated genes in the spiny mouse wound epidermis. c Twist1 related
transcription factors in the spiny mouse wound dermis. d ECM remodeling
in the spiny mouse wound dermis. e IHC of Twist1, Zeb2, MMP9, E-cad and
P-cad in PWD14 the regenerating hair placodes of the spiny mice. White
arrows point to the downregulation of E-cad at the tip of hair placode.
f The effects of Twist1 inhibitor Harmine and pan-MMP inhibitor GM6001
treatment on hair neogenesis in spiny mouse. Red: wound border; yellow
arrow: hair follicles. Observed on PWD35. g Dot graph of the number of
regenerated hair follicles (HF) observed in spiny mouse wounds under
different treatments. n = 6 biologically independent animals. Data are
presented as mean values ± SEM. *p = 0.0265; **p = 0.0027; unpaired
two-sided t-test. Ctl: control. The images in f represent six out of
six experiments performed. The images in e represent 4 out of 4
experiments performed.
It should be noted that while dermal Twist1 is highly expressed in
embryonic skin, it is absent in the E14 epidermal hair placode
(Supplementary Fig. [164]S3d). Identifying epidermal Twist1 in WIHN
suggests that the spatiotemporal dynamics of the tissue stiffness and
gene expression may play a more important role in the epithelial
placode formation in WIHN than in embryonic development.
Epidermal Twist1 regulates both epidermal and dermal cell behavior and tissue
stiffness towards hair primordia formation in WIHN
To verify the role of epidermal Twist1 in WIHN, we crossed the K14-Cre
mice with Twist1-loxP mice to generate the K14-Cre-Twist1 mutant mice
(Fig. [165]5a) that expressed little Twist1 in the epidermis
(Supplementary Fig. [166]S4a). The mutant mice showed a significant
decrease in hair neogenesis (Fig. [167]5b, c, n), and a significantly
stiffer wound bed than that of wild type mice (Fig. [168]5d, o,
Supplementary Fig. [169]S4b). We further used small molecule inhibitors
to suppress Twist1 (Harmine, Supplementary Fig. [170]S4a) and its
downstream MMP activities (GM6001), and both treatments significantly
decreased the number of hair neogenesis (Fig. [171]5e–j, n), stiffened
the wound center (Fig. [172]5o, Supplementary Fig. [173]S4b), and
reduced the wound area under 15 kPa (Supplementary Fig. [174]S4c).
Interestingly, in the perturbed wounds, the wound area under 15 kPa
showed a correlation with the resultant number of hair follicles
(R^2 = 0.7496, Supplementary Fig. [175]S4d), although the slope of the
trend line is much smaller than that of the control and Blebbistatin
treated samples (1.6828 vs 12.812, Supplementary Fig. [176]S4d and
Fig. [177]1k).
Fig. 5. Epidermal Twist1 regulates both epidermal and dermal cell behavior
and tissue stiffness towards hair primordia formation in WIHN.
[178]Fig. 5
[179]Open in a new tab
a A K14-Cre-Twist1 mouse with a PWD28 wound. b AP staining of a
K14-Cre-Twist1 PWD28 wound. c, Schematic diagram of b. d Stiffness
heatmap of PWD14 K14-Cre-Twist1 wound overlaying on the wound photo.
e–g AP staining of control (Ctl), and Harmine and GM6001 treated wounds
on PWD28. h–j Schematic diagrams of e–g, respectively. j Photo of PWD28
wound transfected with Twist1-overexpressing lentivirus. k bright field
and l fluorescence image of GFP-tagged-Twist1 overexpressing (oe) virus
in the PWD28 wound, colocalizing with hair follicles. m Schematic
diagram of k and l. n Dot plot showing the resultant hair follicle
number in wild type, K14-Cre-Twist1, Harmine-treated, GM6001-treated
and Twist1-overexpressing (oe) virus treated PWD28 wounds. n = 7
biologically independent animals. Data are presented as mean
values ± SEM. All comparisons made to WT. Twist1-oe: p = 0.0074,
K14-Cre-Twist1: p = 0.0008, Harmine: p < 0.0001, GM6001: 0.0044.
Unpaired two-sided t-test. o Changes in wound stiffness upon different
perturbations. K14-Cre-Twist1, GM6001 and Harmine treatments all
significantly increased the stiffness of the wound center region,
marked by asterisks. Green bar: 5–15 kPa morphogenetic range. n = 5
biologically independent animals. Data are presented as mean
values ± SEM. All comparisons made to WT of respective location on
wound. Center: K14-Cre-Twist1, p = 0.0006; GM6001, p = 0.0006; Harmine,
p = 0.0018. Center-left: K14-Cre-Twist1, p = 0.01115; GM6001,
p = 0.0137; Harmine, p = 0.0214. Center-right: K14-Cre-Twist1, p =
0.0083; GM6001, p = 0.006; Harmine, p = 0.0159. Unpaired two-sided
t-test. p qPCR analysis: ECM remodeling related-gene expression fold
change (FC) of wild type epidermis wound center vs K14-Cre-Twist1 wound
center. n = 3 biologically independent samples. Data are presented as
mean values ± SEM. All comparisons made to WT. Mmp2, p = 0.0483; Mmp9,
p = 0.0014; Mmp13, p = 0.0005; Itga1, p = 0.0071. Unpaired two-sided
t-test. q PWD14 wild type vs K14-Cre-Twist1 dermis wound center RNA-seq
analysis identifies significantly enriched pathway affected by
K14-Cre-Twist1 knockout. r Dermal condensate (DC) signature genes
downregulated in PWD14 K14-Cre-Twist1 dermis wound center. Red dotted
line: wound boarder. Yellow arrow: hair follicle. Harmine: Twist1
inhibitor. GM6001: pan-MMP inhibitor. Ctl: control. The images in a–m
represent 6 out of 7 experiments performed.
We further used lentivirus to transfect and overexpress Twist1 in the
wild-type mouse wounds on PWD10 and observed a significant increase in
the resultant hair follicle numbers on PWD28 (Fig. [180]5k–n).
Correspondingly, we re-analyzed public microarray data that compared
mouse strains with high and low WIHN capacity^[181]40, and found that
Twist1 expression levels are also significantly higher in the high
regenerative strain (C57BL/6 X FVB X SJL) than the low regenerative one
(C57BL/6, p = 0.004, Supplementary Fig. [182]S4e). These findings
verify that Twist1 and its downstream signals such as MMP play
important roles in controlling wound stiffness and hair follicle
neogenesis during WIHN (Fig. [183]5n, o).
To explore the signaling molecules perturbed by Twist1 knockdown during
WIHN, we compared the gene expressions of PWD14 wild type epidermis
wound center to that of K14-Cre-Twist1, and found downregulation in
many genes related to cell proliferation, Wnt/β-catenin signaling
(Supplementary Fig. [184]S4f, g) and ECM remodeling (Fig. [185]5p).
Previous studies have demonstrated that the epidermal placodes are
required for underlying dermal condensation and the ensuing hair
follicle development^[186]41,[187]42. To further explore whether
epidermal Twist1 affects DC fate acquisition during WIHN, we performed
RNA-seq analysis on PWD14 wild type dermis wound center, PWD14 wild
type dermis wound margin, and PWD14 K14-Cre-Twist1 dermis wound center.
There are 1623 DEG between PWD14 wild type dermis wound center vs
margin (Supplementary Fig. [188]S5a). Among them, DC signature genes
are significantly enriched (p = 1.06 × 10^−5), which suggests DC niche
formation in the PWD14 dermis wound center. In addition, we found 928
DEG between K14-Cre-Twist1 and wild type dermis wound center comparison
(Supplementary Fig. [189]S5b), and the significantly enriched pathways
include cell movement, ECM, inhibition of MMP, Wnt/β-catenin signaling
and integrin signaling (Fig. [190]5q). To further investigate the
contribution of epidermal Twist1 to dermal condensation, we overlapped
DEG from wild type dermis center vs margin (total 1623 genes) and those
from K14-Cre-Twist1 vs wild type dermis center genes (total 928 genes)
(Supplementary Fig. [191]S5c), and found 258 significantly overlapping
DEG (p = 1.37 × 10^−40). This indicates many DEG between wild type
dermis wound center and margin were perturbed by epidermal Twist1
knockout. Among these DEG, most of the significantly upregulated genes
were significantly downregulated by knocking out epidermal Twist1, and
vice versa (r = −0.61, p = 1.1 × 10^−27, Supplementary Fig. [192]S5d).
Similarly, we can identify 23 DC signature genes that are upregulated
in the wild type dermis wound center vs wound margin, which were also
downregulated by epidermal Twist1 knockout (Supplementary
Fig. [193]S5e). In total, there are 31 DC signature genes that are
downregulated in PWD14 K14-Cre-Twist1 dermis center versus wild type
(Fig. [194]5r). These results suggest that epidermal Twist1 plays an
essential role in regulating DC and following hair follicle
regeneration via an epidermal-dermal signaling interaction during WIHN.
Our findings here suggest tissue mechanics and epidermal Twist1 may
feed in Wnt/β-catenin based hair primordia formation pathway in WIHN.
This new “non-canonical” concept will be further discussed in
discussion, together with literature.
Turing-like mechanism explains an optimal wound stiffness range facilitates
new hair placode formation
From our previous mechanical analysis, we allude that an optimal range
of wound stiffness is required for hair neogenesis to occur. This also
suggests that in order for the hair placode to form and invaginate into
the dermis, the aggregated cells should be able to overcome the
physical barrier of the dermis. To quantify its mechanical properties,
we used the AFM to map multiple 100 × 100 μm squares in the wound
center to explore the apparent stiffness of the PWD14 hair placodes
versus surrounding wound bed on a micrometer scale (Fig. [195]6a,
left). The results show that the stiffness of hair placodes
(17.36 ± 0.34 kPa) is significantly higher than that of the wound
center (10.53 ± 0.58 kPa), but still much lower than the wound margin
(Fig. [196]6a, dot plot), implying that the activated epithelial
placode cells also undergo mechanical changes in addition to gene
expressions (Fig. [197]6a′). In parallel, we also found that the
average stiffness of E14 mouse embryonic skin to be 7.3 ± 0.6 kPa while
its hair placode is 9.6 ± 0.5 kPa (Supplementary Fig. [198]S6a, b),
which is comparable to the microenvironment of the wound center in
laboratory mice and the entire wound bed in spiny mice.
Fig. 6. Multiscale tissue mechanics set up morphogenetic field for WIHN.
[199]Fig. 6
[200]Open in a new tab
a AFM stiffness mapping demonstrates the stiffness of the hair placode.
This is based on experimental data. Top left: an AFM cantilever
scanning a PWD14 wound. Blue box demarcates the 100 × 100 μm scanning
area. Below: stiffness heatmap of a placode. Colorimetric unit: kPa.
Right panel: Dot plot showing the stiffness of placode with respect to
wound margin and center. Representative image of 3 out of 3
experiments. n = 9 regions examined over 3 independent biological
animals. Data are presented as mean values ± SD. p < 0.0001, unpaired
two-sided t-test. a′ Schematic diagram of placode stiffness and its
respective cross-sectional view. b Hypothetical model showing feedback
loop in a Turing system with an underlying measure of stiffness.
Simulations illustrating stationary distributions of u (activator, blue
background blocks) and E (stiffness, pick background blocks) with
increasing s (source coefficient, measures the strength of the E
source). The underlying domain is a square of side length 100 and the
color shows the value of S at each grid point, S(x, y). The ‘size’ of
the soft region is controlled by σ. From left to right and then the row
below, the values of s are 2, 4, 6, 8 and 10, and σ = 20. In all cases
random initial conditions from a uniform distribution of [0.5, 1.5] are
used. c Schematic drawing showing the opposite topology of
morphogenetic competent and non-competent region in the wound bed. d
Conceptual summary of the way we perceive the relationship between
tissue stiffness and morphogenetic field (Top). Summary based on data
from Figs. [201]1 and [202]2. Middle: It highlights the different
geographic distribution of the morphogenetic field (green) within a
wound bed (red frame), and also the periodic appearance of hair
primordia (orange) within the morphogenetic field (green). When the
wound stiffness is too low (blue), no placode can form. Bottom:
Stiffness of different wound beds predicate distribution of
morphogenetic field and placode formation.
Turing model has been proposed as the underlying mechanism of pattern
formation^[203]43. Here we look to construct a Turing system^[204]44 to
explain the differential placode formation pattern in laboratory and
spiny mice, which is linked to the underlying structure of the solution
region. Namely, the system should produce spots (hair placodes) within
a specific region of stiffness; if the solution region is too soft, or
too stiff, then the system does not pattern.
We consider three diffusible populations an activator,
[MATH: u :MATH]
, an inhibitor,
[MATH: v :MATH]
, and a measure of stiffness,
[MATH: E :MATH]
. Specifically,
[MATH: u :MATH]
,
[MATH: v :MATH]
and
[MATH: E :MATH]
are thought to be biochemical populations that are able to interact
with each other. The prototypical “Schnakenberg” Turing
kinetics^[205]43,[206]45,[207]46 exist between the populations
[MATH: u :MATH]
and
[MATH: v :MATH]
. The Schnakenberg kinetics are a general form of Turing kinetics,
whereby all dynamics can be connected to source parameters,
[MATH: α :MATH]
and
[MATH: β :MATH]
. Since we have no guidance on kinetics these are as good as any
kinetic type and swapping them for some more accurate kinetics should
not influence the resulting conclusions.
We adapt the Schnakenberg kinetics by modulating the inhibitor source
by the population
[MATH: E :MATH]
, which we take to be a stiffness measure of the field. Namely, a soft
field has low density ECM and, thus, (we assume) the soft field
produces more
[MATH: E :MATH]
, which, in turn produces more
[MATH: v :MATH]
. We consider a square spatial domain
[MATH: [−50,
50]×[−50, 50] :MATH]
, centered at zero, with Neumann boundary conditions and random initial
conditions. In terms of the mathematics we produce the following system
of interactions:
[MATH: dudt⏟Rateofchangeofu=∇2<
/mrow>u⏟Diffusionofu+α−u+
u2v<
mrow>⏟Schnakenberginteraction, :MATH]
1
[MATH: dvdt⏟Rateofchangeofv=Dv∇2<
/mrow>v⏟Diffusionofv+E−u2v⏟SchnakenberginteractionwithproductiondependentonE
, :MATH]
2
[MATH: dEdt⏟RateofchangeofE=DE∇2<
/mrow>E⏟DiffusionofE+sS(
x,y)⏟RegionofsofttissueactsasasourceforE−E⏟Edecaysovertime. :MATH]
3
The coefficients
[MATH: Dv
:MATH]
and
[MATH: DE
:MATH]
are positive constants, which measure how quickly the populations
spread. The source coefficient,
[MATH: s :MATH]
, measures the strength of the
[MATH: E :MATH]
source,
[MATH:
S(x,y)=exp−xσ2−yσ2. :MATH]
4
This means that the
[MATH: E :MATH]
source is a scaled Gaussian distribution centered at zero. The source
is highest in the center and decays towards the boundary. Physically,
this means that the tissue is softest in the center and stiffest on the
boundary. The ‘size’ of the soft region is controlled by
[MATH: σ :MATH]
, namely, increasing
[MATH: σ :MATH]
makes the source function ‘wider’ meaning that more of the tissue is
soft. An illustrative example of
[MATH: S :MATH]
is shown in Fig. [208]6B.
Simulations of increasing
[MATH: s :MATH]
can be seen in Fig. [209]6B. The top row (pink background) shows the
output
[MATH: u :MATH]
after a threshold (color change) has been applied, illustrates the
change in tissue stiffness. The bottom row (blue background) shows the
accompanying profile of
[MATH: E :MATH]
, reflects spot pattern formation. Namely, the spots in the blue
background illustrate the regions in which
[MATH: u>3.8 :MATH]
. Once again, as
[MATH: E :MATH]
decreases (from pink to green and blue) we expect the tissue beneath to
be softer. Thus, we see that as
[MATH: s :MATH]
increases left to right the center becomes lighter and lighter (pink
background blocks). Notably, as the center becomes too soft (the last 2
pink background blocks in the third row) we see that placodes stop
forming in this region (the last 2 blue background blocks in the bottom
row). Hence, we have produced a simulation in which a Turing pattern
has a feedback loop with an external field, which is considered to be a
measure of the underlying ECM stiffness. Critically, in order for spots
to form the media can be neither too stiff nor too soft.
Discussion
In summary, we show that multiscale tissue mechanics of the wound bed
partake in setting up the morphogenetic field for WIHN, and Twist1 is
an important chemo-mechanical regulator involved in initiating cellular
events that lead to placode formation and invagination through symmetry
breaking of the epidermis, ECM remodeling, collective migration and
epidermal-dermal crosstalk. As we try to recapitulate the developmental
process to facilitate regenerative wound healing, the mechanical
microenvironment of the tissue should also be considered. The
stiffening of the epithelial hair placode serves as an important
symmetry breaking point - cells invaginating into the soft dermis
(Fig. [210]6a). Identifying 5–15 kPa as the optimal stiffness range
(Fig. [211]2k) is important in setting up the morphogenetic field for
the hair placodes to overcome the physical barrier of the
microenvironment and invaginate into the dermis. Furthermore, different
spatiotemporal dynamics of wound stiffness in different species
predicate the distribution of morphogenetic field and placode formation
(Fig. [212]6c, d).
We demonstrate that there are two levels of symmetry breaking during
successful WIHN, in parallel to the developmental process. The first
level is the generation of morphogenesis competent field (green) from
the center (Fig. [213]6b, c). The second level is the generation of
periodically arranged hair germs forming (brown dots, Fig. [214]6b,
blue blocks) from the morphogenesis competent field. In the spiny mice,
the topology is reversed with the competence zone (green) on the
periphery, while the central field (pink) cannot form hairs
(Fig. [215]6c). By perturbing tissue stiffness, we can even generate a
concentric ring-shaped competent field, fulfilling the prediction of
the model (Fig. [216]6b, d). Additionally, the Turing mechanism can
also help explain the asymmetric field in the less uniform environment
(e.g., wound), development and growth^[217]47.
The key question is what factors are required to make a region
competent to undergo further periodic Turing patterning to generate
hair placodes in the adult skin. The multiscale tissue mechanics
perspective allows us to compare the similar and distinct pathways in
development and WIHN, and appreciate that laboratory and spiny mice
have evolved and manifested during regenerative wound healing, in
contrast to repair. The findings and concept, together with those in
recent WIHN studies are discussed in the following.
Macroscale symmetry breaking of tissue mechanics in the wound bed leads to
the emergence of morphogenetic field
By comparing the mechanical and molecular response of the laboratory
and spiny mouse during wound healing, we found that the hair placode
formation pattern is opposite in laboratory and spiny mice during WIHN
and this spatial difference is predicated by the wound bed with
stiffness between 5–15 kPa, which is optimal for hair neogenesis and
also the spatiotemporal expression of Twist1-related TFs.
We postulate that the spatiotemporal formation of the hair placodes is
a good indication of the morphogenetic field of the wound bed,
established partly by tissue mechanics. Previous studies have shown
that there is an optimal matrix stiffness for different cell behaviors
(e.g., cell migration, proliferation, differentiation), and very soft
matrix impedes focal adhesion formation and cell migration^[218]48. We
postulate that on PWD14 the wound center of the spiny mouse was still
too soft for epidermal cells to form the hair placode, a process that
requires epithelial cells to migrate and invaginate into the dermis,
hence it occurred later on PWD21 when the wound stiffness reached over
5 kPa. On the other hand, a wound bed stiffer than 15 kPa may be too
stiff for the epidermal hair placode cells to invaginate, as
demonstrated by the thick and dense collagen fibers in the PWD14
laboratory mouse wound margin (Fig. [219]1l–s). The high collagen III
expression in the spiny mouse wound in PWD14 and PWD21 (Fig. [220]2i),
in contrast to the very few detectable collagen fibrils by SHG
(Fig. [221]2h, Supplementary Fig. [222]S2e, f), suggest that collagen
III (not very crystallined and generates little SHG
signal^[223]38,[224]39) may be responsible for constituting the soft
wound bed that resembles the physical environment of the embryonic
skin, ideal for hair neogenesis. Recent studies also showed that ECM
stiffness and mechanical forces exerted from the epidermal cells
cohesively instruct tissue architectures and function^[225]49,[226]50.
Having a soft wound bed may also modulate the signaling of ECM
remodeling gene expression. The time-course RNA-seq analysis of the
spiny mouse wound showed an 1-week delay in the peak expression time of
Twist1-related genes (PWD14) and ECM, MMP, integrins and other TFs
(PWD21) in spiny mouse (Fig. [227]4b–d). We speculate that a suitable
tissue stiffness is also required for TF to enter the nucleus, as
supported by findings that force-induced nuclear deformation modulates
nuclear entry of TF^[228]51. Hinz has also proposed a candy wrap theory
to describe that certain level of mechanical force is required to
release active TGF-β from its latent form^[229]52. The spiny mouse
dermal cells also demonstrate fewer α-SMA-positive stress fibers upon
substrate stiffness increase^[230]53. The spiny mouse’s ability to keep
the wound bed soft during early stages of wound healing could delay the
nuclear entry and activation of the upstream TF and hence the
expression of certain ECM, MMP and integrins, and consequentially set
up the optimal molecular and mechanical wound bed as morphogenetic
field. Furthermore, it is worth noting that Twist1 expression is higher
in the dermis than epidermis (Fig. [231]4e) in spiny mouse around the
hair placode, which is more representative of the embryonic dermal
Twist1 expression during development (Supplementary Fig. [232]S3d).
These features could be the evolutionary advantages that the spiny mice
have evolved to promote regenerative wound healing and survival. These
similar yet distinct regulations of Twist1 and tissue mechanics between
laboratory and the spiny mice remain to be investigated.
Microscale symmetry breaking of tissue mechanics in the morphogenetic field
leads to the emergence of hair primordial
We have identified Twist1 as the key upstream chemo-mechanical
regulator to activate ECM remodeling, epithelial cell movement,
epithelial proliferation, Wnt/β-catenin signaling, and DC in the dermis
(Figs. [233]3–[234]5). Analysis from a previous microarray database
also showed that Twist1 expression is higher in the mice strain with
high regenerative capacity compared to a low capacity strain
(Supplementary Fig. [235]4e)^[236]40. Twist1 is shown to directly
regulate Cdh11, Grem1, Zeb1, Dkk3, Gli1, Fgfr1, Tbox18, Col6a2, and
Lamb1 (Supplementary Fig. [237]7) when we reanalyzed a database that
used H3K4me3 ChIP-seq to show the epigenetic reprogramming following
Twist1-mediated EMT in human epithelial cells^[238]54. Others have also
shown direct transcriptional binding of Twist1 on the Snai2
promoter^[239]55. This effect of Twist1 also corroborates with the
detected increase in the stiffness of the placode. At the same time,
Twist1 has also been shown to directly bind to the MMP promoter to
exert its transcriptional effect^[240]56. By inducing MMP activity at
the hair placodes and remodeling local ECM, we postulate this can lower
the physical barrier of the dermis to also facilitate placode
invagination. In this study, we used small molecule inhibitors Harmine
and GM6001 to inhibit Twist1 and pan-MMP activities, respectively.
Harmine targets the Twist1 pathways through its promotion of Twist1
protein degradation^[241]57 and is also capable of blocking the
activities of dual-specificity tyrosine phosphorylation-regulated
kinase family proteins and mitogen activated protein kinase^[242]58.
GM6001 is a potent reversible broad spectrum inhibitor of
zinc-containing proteases, including various MMPs (MMP-1, −2, −7, −8,
−9, −12, −13, −14, −16, and −26), disintegrin and metalloproteinase
domain-containing (ADAM) proteins ADAM9, ADAM10, ADAM12, and
ADAM17^[243]59. Although the results of the inhibitor treatment fell in
line with our hypothesis and other Twist1-functional perturbation
studies (lentivirus transfection and K14-Cre-Twist1 transgenic mice),
and there is no significant difference between the number of new hair
follicles observed in K14-Cre-Twist1 and Harmine treated wounds
(Fig. [244]5n), we acknowledge these inhibitors’ potential side effects
outside of Twist1 pathway activities.
Furthermore, in order for the epithelial cells to continue to
invaginate downward into the dermis, the cell number needs to increase,
which can be observed in the highly enriched epidermal proliferation
and movement genes in the gene set enrichment analysis (Fig. [245]3b).
Similar expression of Twist1-related genes at the hair placodes have
also been observed in our spiny mouse RNA-seq analysis (Fig. [246]4b),
others’ laboratory mice WIHN microarray database^[247]60 and human hair
follicle morphogenesis^[248]40. Our K14-Cre-Twist1 RNA-seq analyses
also imply that epidermal Twist1 plays a role in dermal-epidermal
interaction (Fig. [249]5q, r). The well-established morphogenesis
initiator of skin development, β-catenin, has been shown to directly
activate Twist1 expression in skull progenitor cells^[250]61.
Therefore, we reason that Twist1 is one of the chemo-mechanical
regulators that responds to β-catenin activation to induce symmetry
breaking of the morphogenetic field of the epidermis, which facilitates
dermal–epidermal interactions and initiates ECM remodeling, cell
proliferation and collective migration, leading to placode formation
and invagination during WIHN. Alternatively, Twist1 itself could also
play the role of a mechanosensor during mechanotransduction for the
wound induced hair follicle neogenesis^[251]62–[252]65.
WIHN studies identify the concept of noncanonical and canonical hair
primordia formation pathway
WIHN is a combination of local periodic patterning events and a global
influence that constitute the morphogenetic field. The canonical Wnt,
β-catenin and Shh^[253]66 have been identified as the critical
activators of WIHN. Ablation of Wnt in the wound epidermis via
inducible β-catenin deletion eliminates hair neogenesis, while
overexpression of Wnt in the wound epidermis enhances
it^[254]22,[255]67. On the other hand, non-canonical signaling also
regulates WIHN, some by interacting with the canonical signals. The δαT
immune cells secret Fgf9 to act on the neighboring myofibroblasts in
the wound, inducing them to secret Wnt2a ligand^[256]60. Double
stranded RNA, which is released during injury, activates Toll-like
receptor 3^[257]40 and its downstream effectors IL-6^[258]68 and Stat3
to promote hair neogenesis. This effect is achieved through the
induction of known hair morphogenetic molecules such as Edar, Wnt, and
Shh pathways.
Based on our earlier developmental studies of periodic formation of the
feather and hair germs, we noticed there are two waves of molecular
expression, which we name them restrictive and de novo mode,
respectively^[259]4. In short, restrictive mode molecules are present
before periodic patterning occurs, and are required for periodic
patterning process, while de novo molecules are the readout. Our data
showed Twist1 is expressed higher in the wound center (Fig. [260]3f),
it is not exclusively expressed in the nucleus of hair placode cells,
but also inter-follicular epidermis (Fig. [261]3f, g). These results
suggest that Twist1 belongs to the “restrictive mode” molecules, and
therefore they are present in the epidermis in both putative placode
and inter-placode regions. Our view is that Twist1 is initially broadly
expressed in the morphogenetic zone of the wound bed, and become
accentuated in the placode region and enter the nucleus when the sum of
all the upregulating factors for placode formation reaches a threshold.
This is supported by our experiments where either softening the wound
bed or overexpressing Twist1 in the wound enhanced WIHN. In spiny mice,
we have noted that epidermal Twist1 is expressed predominantly but not
exclusively in basal cells (Fig. [262]3f, g). While we postulate that
activation of Twist1 in the basal cells suggests a prelude for EMT,
recent discovery in the heterogeneity of wound epidermal cells^[263]69
eludes that a more comprehensive and single-cell resolution future
study is required to identify the molecular identities of these
Twist1-positive and placode forming cells.
In this study, we show that Twist1 plays a key role in epidermal and
dermal signaling during wound-induced hair primordia formation, which
is a different mechanism adopted during developmental process. Studies
have identified the binding sites for β-catenin on Twist1
promoter^[264]61, although in our K14-Cre-Twist1 gene expression
analysis, knocking out Twist1 also affected Wnt/β-catenin expression in
both the epidermis and dermis during WIHN. We postulate β-catenin may
be the initial activator of Twist1, and Twist1 can also loop back to
regulate canonical Wnt/β-catenin signaling. These findings provide
insights on the canonical/non-canonical molecular events during WIHN.
Furthermore, we also demonstrate that tissue mechanics, or the
stiffness of the wound bed, the ECM, and the stiffness of the cells
also partake during WIHN; the hair primordia formation is not exclusive
to molecular signaling events. The soft and easy-to-shed feature of the
African spiny mouse skin not only serves as an escape strategy from
predators^[265]27, but also fosters the optimal mechanical cue for hair
primordia formation during wound healing. Understanding the common and
distinct features of laboratory and spiny mice in response to wounding
shed light on evo-devo advantages and provide perspectives for future
implications.
Methods
Animal model
All animal work was performed according to the approved animal
protocol, guidelines and regulations for the care and use of laboratory
animals of University of Southern California (USC). Ethical approval
was obtained for all experiments performed. All mice were housed in
climate controlled indoor facilities in a temperature range between 21
and 26 °C with a 12:12-h controlled dark/light cycle. Humidity is
maintained at 30–70%. C57BL/6J mouse purchased from Jackson Lab was
used as the primary animal for this study. The K14-Cre-Twist1 mice were
bred by crossing the Twist1 conditional null (Twist1^+/+)^[266]70 and
Tg(KRT14-cre)1Amc/J (Jackson Lab) mice. The wild type Twist1^+/+ mice
were used as the control in transgenic mouse study. The African spiny
mouse, Acomys cahirinus, is a kind gift from Dr. Malcolm Maden at the
University of Florida and Dr. Ashley W Seifert of University of
Kentucky. A colony of captive-bred Acomys cahirinus was established at
USC, and all experiments were per formed with protocols approved by the
USC IACUC. We used both male and female 2-month-old spiny mice and
4-week-old C57Bl/6J mice for wound experiments unless otherwise
specified.
Wound-induced hair neogenesis assay
Mice were anesthetized using Ketamine–Xylazine (80 mg/kg; 5 mg/kg) and
analgesic Buprenorphine SR (0.5 mg/kg) was given by intraperitoneal
injection (IP) at the beginning of the procedure. A 1 × 1 cm square
full thickness wound was excised on the posterior dorsum of 4-week-old
mice (p28) using scissors, and let it heal by secondary intention. For
the spiny mice, 1.5 × 1.5 cm square full thickness wound was excised on
8 week-old mice. Additional DietGel Boost (ClearH2O) was placed on the
bottom of the cage during the first week post-operation. Mice of the
same sex from the same litter were housed together and provided with
half-dome shelter.
Alkaline phosphatase (ALP) stain
To detect newly forming dermal papillae, alkaline phosphatase staining
was performed as previously reported^[267]22. Briefly, full thickness
wounds were excised and epidermis separated from the dermis using 20 mM
EDTA. The dermis was fixed in acetone overnight at 4 °C, and washed in
PBS several times. The dermis was pre-incubated in ALP buffer (0.1 M
Tris-HCl, 0.1 M NaCl, 5 mM MgCl[2] and 0.1% Tween-20) for 30 min,
incubated with BCIP/NBT Color Development Substrate (Promega, Madison,
WI, USA) in ALP buffer at 37 °C until color development. The reaction
was stopped by washing with pH 8.0 Tris-EDTA and the tissue stored in
PBS with sodium azide.
Atomic force microscopy
AFM (NanoWizard 4a/CellHesion, JPK, Berlin, Germany) was setup for
contact mode indentation in PBS. The spring constants of all
cantilevers were calibrated via thermal noise method with correction
factor in liquid^[268]71,[269]72 prior to each measurement resulting in
values of 0.03 N/m. To allow for proper modeling of the data, a glass
bead (5 μm in diameter) was attached at the end of a tipless
rectangular cantilever (Arrow-TL1, NanoWorld, Neuchatel, Switzerland)
using 2-component epoxy (Gorilla Glue Epoxy Clear, Gorilla Glue
Company, Cincinnati, OH, USA). A force series identified a maximum
indentation force of 5 nN to show the most consistent results on test
samples. A constant rate of 1 μm/s was used for the entire approach and
retract sequence. Force-distance curves were collected and
post-processed using the JPK package software (Data Processing,
6.3.11). The force curves were analyzed using the Hertz model with a
spherical indentation^[270]35,[271]36
The force on the cantilever F(h) is given by:
[MATH: Fh=
Esample1−vsample24R3h3/2 :MATH]
where h is the depth of the indentation, E is the effective modulus of
a system tip-sample, v is the Poisson ratio for the sample, and R is
the radius of the AFM tip. The unit of Young’s modulus is calculated as
N/m^2, and expressed as pascal (Pa) or kilopascal (kPa). Poisson ratio
was set at 0.5 since the spherical tip was incompressible relative to
the sample. The temperature of the measurement was controlled at 32 °C
to mimic the surface temperate of mouse skin^[272]73.
To maintain the biomechanical force integrity of the dorsal wound, the
entire mouse skin organ was removed by creating an excision on the
ventral side midline, extending from the anterior neck region to the
posterior genital region, and then dissecting away the skin organ from
the underlying fascia. Normal skin and wound stiffness were immediately
measured after skin organ removal to prevent artifacts from tissue
decomposition. AFM measurements were positioned across the wound
starting from unwounded normal skin on one side and progressively
traveling through to the opposite side; the near wound edge, the near
wound bed, the wound center, the opposite wound bed, the opposite wound
edge, and the opposite normal skin. At least 5 indentation points were
taken for each region of interest.
Second harmonic generation imaging
The animals were anaesthetized using Isofluorane. Body temperature was
maintained with a homeothermic blanket system (Harvard Apparatus,
Holliston, MA, USA). The SHG images of animals were acquired using the
external detectors of an inverted Leica SP5 (Wetzlar, Germany)
multiphoton confocal fluorescence microscope powered by a Chameleon
Ultra-II MP laser at 860 nm and a 40x Zeiss water-immersion objective
(NA1.2). A Z-stack series of 3 μm per slice, 50 slices in total was
recorded during a time course starting at the cornified layer of the
epidermis and ending at 150 µm depth for each time point.
Heatmap of spatial stiffness of the wound
The interpolation of tissue stiffness was performed by using 3-D
meshgrid function of MATLAB (R2015b, Natick, MA, USA). After obtaining
a Young’s modulus (z) at a specific spatial location (x, y) in the
wound, a 3-dimensional matrix was defined. When the positions and
stiffness of all the measured spots were identified, we could
interpolate the stiffness of the positions in between to average the
stiffness of the nearest parameters using 3-D meshgrid function, by
defining (x, y) as meshgrid and (z) as griddata. In the end, the
heatmap was generated by defining the representative color of
stiffness.
Wound area calculation and stiffness analysis
The area of the wound is quantified by using the area measurement
function under ImageJ according to its user guide ImageJ/Fiji 1.46
(NIH, Bethesda, MD). The photo of the wound was taken with a scale bar,
hence the measured area in pixel unit can be converted to actual size.
To obtain the area of the wound under 15 kPa, we specifically adjusted
the (x, y, z) values according to each respective wound so the heatmap
was also actual size. Using Photoshop, we use the wand tool and set the
tolerance value to match that of 15 kPa on the scale bar. By using this
parameter, the wand tool could select the area of the wound along the
15 kPa line. This selected area is then saved and quantified using
ImageJ to obtain the final actual area.
Inhibitor treatments
Blebbistatin (Cayman, MI, USA), Harmine (Cayman, MI, USA) and GM6001
(Cayman, MI, USA) were dissolved in DMSO. 20 μl was applied once a day
directly to the wound surface starting at post-wound day 10 (PWD10) and
continuing until PWD16.
Histological preparations
The wound tissues were fixed in 4% PFA and dehydrated in a graded
alcohol series. The tissue was cleared in Xylene and embedded in
paraffin wax. 6 μm sections were cut on a microtome. H&E sections were
performed according to accepted protocol. Whole-mount tissues were
fixed in 4% PFA and then stored at 4 °C in PBS with NaAzide.
Paraffin section and whole-mount immunohistochemistry
Fixed tissues were permeabilized with methanol and blocked with 3%
H[2]O[2] for 30 min, and then serum blocked for 1 h. The primary
antibody was added and incubated over night at 4 °C with agitation. The
tissue was washed with TBST (Tris Buffered Saline Tween 20) and the
secondary antibody was added for 1 h at room temperature. The tissue
was washed with TBST and if utilized, a tertiary antibody was added for
1 h at room temperature. The tissue was washed and color was developed
using the AEC kit (Vector Laboratories, CA, USA) or fluorescence was
visualized with a fluorescence microscope. The whole-mount samples were
cleared in a series of Glycerol-PBS until 100% Glycerol for imaging.
The Twist1 (ab50887), Collagen I (ab34710) and Collagen III (ab7778)
antibodies are from Abcam (Cambridge, MA, USA), MMP9 (N2C1, GTX100458)
is from GeneTex (Irvine, CA, USA), Snai1 (13099-1-AP), P-cadherin
(13773-1-AP), E-cadherin (20874-1-AP) and Zeb2 (14026-1-AP) are from
Proteintech (Rosemont, IL, USA). The dilution ratio for section IHC was
1:50, 1:400 for wholemount immunostaining.
Harvesting wounds from laboratory mice for RNA extraction
The wound was harvested and a 3 mm diameter hole-punch biopsy was taken
from the geometric center of the wound, and the remaining wound tissue
was considered the margin. The epidermis and dermis were separated
manually under a dissecting microscope. The dissected dermis tissues
were immediately placed in liquid nitrogen for 30 s. The frozen tissues
were then disaggregated individually using a mortar and pestle, and
then collected into a 1.5 ml microtube. The dissected epidermis was
also collected into a 1.5 ml microtube.
Harvesting wounds from spiny mice for RNA extraction
The spiny skin (shaved) or wound was harvested and placed epidermal
side down in a 3 cm culture dish filled with a thin layer of 0.25%
Trypsin-EDTA (ThermoFisher, Waltham, MA, USA). The level of trypsin
should be just enough to cover the epidermis but not submerging the
tissue. The tissue is incubated at 4 °C for 6–12 h, rinsed in PBS and
the dermis-epidermis were separated manually under a dissecting
microscope. The epidermis was collected into a 1.5 ml microtube. The
dissected dermis was placed in liquid nitrogen for 30 s and
disaggregated using a mortar and pestle, and collected into a 1.5 ml
microtube.
RNA extraction and RNA-seq
The RNA was extracted using the RNeasy Mini Kit (QIAGEN, Hilden,
Germany). 1 µg of total RNA from each sample was used to construct an
RNA-seq library using TruSeq RNA sample preparation v2 kit (Illumina,
CA, USA). Sequencing (75 cycles single-end or paired-end reads) was
performed by USC Molecular Genomics Core using a NextSeq 500 sequencer
(Illumina, CA, USA).
RNA-seq and microarray analysis
The mouse mm10 reference genome, and RefSeq genome annotation
downloaded from the UCSC Genome Browser on 5 June, 2019 were used for
RNA-Seq analysis^[273]74. The alignment, quantification, normalization,
and differentially expression analysis were performed by STAR
2.6.1d^[274]75, htseq-count 0.6.0^[275]76, TMM^[276]77, and edgeR
3.26.8^[277]78, respectively. P value or False discovery rate < 0.05
was set as a threshold to identify differentially expressed genes
(DEG). The hierarchical clustering, Venn diagram, volcano plot, gene
expression profile, and scatter plot were carried out by Partek
Genomics Suite 7.18.0723 (Partek Inc. MO, USA). The pathway enriched
analysis based on Fisher’s exact test, and upstream analysis were
performed by Ingenuity Pathway Analysis (Content Version: 60467501,
Build: ing_beryl, Date: 11-20-2020; IPA, QIAGEN Inc. CA, USA). An HTMH
form Perl CGI program was performed for the statistical significance of
the overlap between two groups of genes
([278]http://nemates.org/MA/progs/overlap_stats.html). The gene set of
DC signature gene and EMT core genes were build based on Supplementary
Table [279]S1 of^[280]79 and upregulated genes in Supplementary
Table [281]S1 of^[282]80. Public microarray data under GEO database
([283]GSE50418) were re-analyzed using Partek Genomics Suite 7.18.0723
(Partek Inc. MO, USA).
Lentivirus production and transfection
The Twist1 overexpressing vector genome plasmid was cloned by inserting
Twist1 promoter into the lentiviral backbone:
5′LTR-cPPT-Ubq-eGFP-P2A-Twist1-WPRE-3′LTR. Twist1 promoter was
amplified from mouse genomic DNA, and the plasmid backbone was
purchased from Addgene (Watertown, MA, USA). The empty backbone without
Twist1 promoter insertion was used as control.
293T cells (ATCC^® CRL3216™) at 50–60% confluency were transfected with
10 μg vector genome plasmid, 10 μg of packaging construct ΔR8.2, and
2 μg envelope plasmid pCMV-VSVG using the calcium phosphate method.
10 mM sodium butyrate was added to fresh media 16 h post-transfection
and removed after 8 h. Virus-containing media was collected at 36 h
post-transfection, sterile filtered, and ultracentrifuged on a 20%
sucrose cushion at 110,863 g and 4 °C for 1.5 h before storing at
−20 °C for up to 30 days or −70 °C indefinitely.
The virus was applied to the wound on PWD10 to infect the tissue. The
efficiency of transfection can be visualized by detecting the eGFP
intensity under a fluorescent microscope, and later verified by frozen
section IHC.
RT-qPCR
RNA extraction was done with Zymo Research Direct-zol RNA Kits. Reverse
transcription was done using Superscript III First Strand Synthesis
kit. The RNA and cDNA concentrations were measured with the NanoDrop
2000 spectrophotometer and normalized between samples. Primers used for
qPCR are listed in Supplementary Table [284]S2. The Ct values were
measured using the Agilent Mx3000P qPCR system. The relative
quantification was done by pyQPCR Version 0.9 software.
Statistics
The Kolmogorov–Smirnov tests were conducted to test normal distributed
random samples. Two independent sample T-tests (two-tailed) were used
for comparing unpaired sample groups. For some datasets not equally and
normally distributed, Wilcoxon rank tests were conducted using IGOR Pro
or MATLAB to evaluate the statistically significant difference between
two samples. Chi-square test was conducted in MATLAB. The photographs
are representative samples of at least four replicates. Stiffness
measurements were reported as the averaged of at least four independent
samples. Hair follicle counts are reported as the average from at least
6 samples. Each bar on qPCR graph represents average and SE of three
independent samples. All data is presented as mean ± SD unless stated
otherwise. Results from student t-tests (two-tailed) with p < 0.05 was
considered significant. *, p < 0.05. **, p < 0.01. ***, p < 0.005.
Reporting summary
Further information on research design is available in the [285]Nature
Research Reporting Summary linked to this article.
Supplementary information
[286]Supplementary Information^ (1.7MB, pdf)
[287]Reporting Summary^ (348.7KB, pdf)
[288]Peer Review File^ (3.9MB, pdf)
Acknowledgements