Skip to main content
NCBI home page
Plastic and Reconstructive Surgery Global Open logoLink to Plastic and Reconstructive Surgery Global Open
. 2025 Aug 27;13(8):e7058. doi: 10.1097/GOX.0000000000007058

The Utility of De-epithelialization in Subcutaneous Skin Flap Transposition

Takuya Saito *,✉, Kenichiro Kawai †, Soh Nishimoto †, Hisako Ishise †, Koyo Nakajima †, Masao Kakibuchi †
PMCID: PMC12385113  PMID: 40881255

Abstract

Background:

To address soft tissue volume deficiencies following surgery or injury, plastic surgeons implant de-epithelialized flaps subcutaneously. During this procedure, residual epidermal components and skin appendages may inadvertently be implanted, potentially leading to postoperative complications. However, such occurrences are rare in clinical practice. To date, no studies have examined the long-term changes in subcutaneously transposed skin flaps.

Methods:

De-epithelialized and non–de-epithelialized flaps were implanted subcutaneously in rat models. Pathological analyses, Western blotting, and quantitative real-time polymerase chain reaction were performed to evaluate temporal changes.

Results:

In all cases of non–de-epithelialized flap implantation, subcutaneous masses were palpable at all time points. Pathological and molecular analyses revealed the formation of a single epidermal cyst at each site throughout the study period, with no significant changes in keratin 5 expression, a marker of the basal epidermal layer. In contrast, in de-epithelialized flap sites, subcutaneous masses observed at 1 month postoperatively gradually regressed over time. Clusters of small cysts that initially formed at 1 month postoperatively disappeared over time, coinciding with a reduction in keratin 5 expression. Furthermore, regardless of epithelialization status, the development of body hairs from subcutaneously implanted follicles progressively diminished.

Conclusions:

These findings suggest that in non–de-epithelialized flaps, residual epidermis persists and continues to differentiate, potentially leading to complications. In contrast, in de-epithelialized flaps, remaining skin appendages and epithelial components gradually regress, thereby reducing the risk of complications. The results underscore the clinical usage of de-epithelialization in subcutaneous skin flap transplantation.

Takeaways

Question: When a skin flap is implanted under the skin, de-epithelialization is performed to avoid complications caused by the epidermis. In this study, we examined the usefulness of de-epithelialization, which has not been previously studied.

Findings: De-epithelialized and non–de-epithelialized flaps were implanted subcutaneously in model rats, and changes over time of the flaps were examined. In the non–de-epithelialized group, epidermal differentiation continued subcutaneously and caused complications, whereas in the de-epithelialized group, epidermal differentiation did not occur and complications were clearly suppressed.

Meaning: Our study demonstrated the usefulness of de-epithelialization when a skin flap is transposed subcutaneously.

INTRODUCTION

Plastic surgeons routinely perform flap techniques for the reconstruction of the breast, head and neck, and other body regions. The flap technique involves transposing tissue with its own blood supply, distinguishing it from skin and fat grafts, which lack an intrinsic blood supply.1,2 This distinction allows flap techniques to provide more voluminous tissue than other methods.

To compensate for soft tissue volume loss after tumor resection or injury, the flap is implanted with its epidermis removed, and the process is called “de-epithelialization.”3 This procedure is performed to prevent the formation of cysts and infections, which may occur if the epidermis remains beneath the skin. Although de-epithelialization is widely practiced to reduce such complications, no studies have conclusively validated its efficacy.

During de-epithelialization, preserving the dermis rather than excising the entire skin layer is recommended to protect the subdermal vascular plexus and prevent volume loss.4 The remaining dermis also serves as structural support for suturing, as fat and muscle lack the necessary solidity. De-epithelialization is typically performed using scissors or a scalpel, and its completion is assessed visually. However, residual epidermal elements may be inadvertently left behind and placed under the skin. Additionally, epithelial components, such as hair follicles and stem cells from the lower bulge region of the follicular canal, are thought to persist despite the procedure.5

Singh et al6 reported a case in which breast skin necrosis led to the exposure of a de-epithelialized flap after breast reconstruction, revealing epithelialization centered around residual skin appendages. This suggests that skin appendages and stem cells remaining on the flap may contribute to postoperative complications, such as epidermal cyst formation even with de-epithelialization.

Moreover, because hair follicles extend into the fat layer, there is a possibility that residual hair follicles on the flap may continue functioning postoperatively, generating subcutaneous body hair even after de-epithelialization.7 This could lead to complications such as pilonidal sinus formation.8 However, such complications are rarely observed in clinical practice. If epidermal differentiation and hair follicular function are suppressed in the subcutaneous environment and no complications arise without de-epithelialization, the procedure may be unnecessary. The advantage of not requiring de-epithelialization is that volume loss is minimized.

Peer and Paddock9 studied skin grafts placed beneath the skin and reported cyst formation. They observed that hair follicles on de-epithelialized grafts were initially present but gradually developed into microscopic round cysts.9 Similarly, Dimitroulis and Slavin10 investigated full-thickness skin grafts placed into the craniomandibular joint of a rabbit model and found atrophy of sweat glands and hair follicles, along with cyst formation. However, these studies were conducted using graft models, not flap techniques.

Unlike skin grafts, skin flaps maintain an intact blood supply and include the deepest portions of hair follicles, such as the dermal papilla. To evaluate the usefulness of de-epithelialization, we examined time-dependent changes in keratinocytes and appendages in subcutaneously transposed skin flaps by comparing de-epithelialized and non–de-epithelialized flaps.

MATERIALS AND METHODS

Animals

Seven-week-old male F344/Jcl rats were purchased from CLEA Japan, Inc. (Tokyo, Japan). All animals were housed under standard conditions at Hyogo Medical University’s animal facility in accordance with the institution-approved guidelines.

Local Flap Technique in the Rat Model

Twelve rats were randomly assigned to 2 groups: the non–de-epithelialized group (group 1) and the de-epithelialized group (group 2). Four local flaps measuring 1 × 1.5 cm were created on the back of each rat (Fig. 1).

Fig. 1.

Fig. 1.

Open in a new tab

Four local flaps were created on the back of each rat, rotated 90 degrees, and sutured underneath the skin.

In group 1, the flaps were shaved but not de-epithelialized, except for the epidermis at the base of the flap, which was removed to facilitate complete wound closure (Fig. 2). In group 2, the epidermis of the flap was completely removed using scissors. The completion of de-epithelialization was assessed visually. All flaps were rotated 90 degrees and sutured subcutaneously (Fig. 3).

Fig. 2.

Fig. 2.

Open in a new tab

Intraoperative photographs showing the process of flap elevation. A, In group 1, the skin flap was shaved, and only the base of the skin flap was de-epithelialized. B, The wound was sutured.

Fig. 3.

Fig. 3.

Open in a new tab

Stepwise procedure. A, The flap was designed, and body hair was shaved. B, The local skin flap, including the fat layer, was elevated. C, The epidermis at the base of the skin flap was removed. D, A subcutaneous pocket was created, and the skin flap was rotated 90 degrees and inserted. E, The wound was closed with sutures.

Sample Collection and Analysis

At the end of the experiment, all animals were euthanized using carbon dioxide gas. Tissue samples were collected at 1, 3, and 6 months postoperatively to analyze time-dependent changes in local and pathological findings, including immunostaining with an antikeratin 5 antibody (Abcam, Cambridge, United Kingdom). Keratin 5 is expressed in the basal layer of the epidermis.11 Western blot analysis and quantitative real-time polymerase chain reaction (PCR) were performed to evaluate the differentiation potential of keratinocytes.

Western Blot Analysis

Keratin 5 expression levels were analyzed using the automated Abby instrument (ProteinSimple, San Jose, CA). Tissues were lysed with accessory sample buffer, and 3 µL of lysate was combined with a 1:60 dilution of primary anti-cytokeratin 5 antibody (Abcam) and analyzed according to the manufacturer’s instructions.

Quantitative Real-time PCR

Total RNA was extracted from each flap using ISOGEN (Nippon Gene Co., Ltd.) following the manufacturer’s instructions. Aliquots of 220 ng of total RNA were reverse-transcribed into complementary DNA using SuperScript III Reverse Transcriptase (Invitrogen, CA). Quantitative real-time PCR was conducted using SYBR Green PCR Master Mix (Applied Biosystems, CA) and the QuantStudio 12K Flex Real-Time PCR System (Thermo Fisher Scientific).

The following primers were used:

  • Rattus norvegicus keratin 5: 5′-ACACCCCTGAACCTGCAAAT-3′ (forward) and 5′-TCATTCTCAGCCGTGGTACG-3′ (reverse).

  • R. norvegicus cyclophilin A: 5′-GTCTGCTTCGAGCTGTTTGC-3′ (forward) and 5′-CCTTTCTCCCCAGTGCTCAG-3′ (reverse). Cyclophilin A was used as the internal control.

Statistical Analysis

All statistical analyses were performed using EZR (Saitama Medical Center, Jichi Medical University, Saitama, Japan), a graphical user interface for R (The R Foundation for Statistical Computing, Vienna, Austria). Specifically, EZR is a modified version of R Commander designed to incorporate statistical functions commonly used in biostatistics. Results are presented as means ± SD of n observations. Comparisons between more than 2 experimental groups were conducted using the Tukey–Kramer test. A P value of less than 0.05 was considered statistically significant.

RESULTS

Local Findings

  • Group 1: Subcutaneous masses were palpable at all 12 flap sites. Their average size was 1.5 cm at 1 and 3 months and 2 cm at 6 months, with no significant differences. Cysts containing body hair and foul-smelling serous fluid were observed at all time points. Hair within the cysts became progressively thinner over time (Fig. 4).

  • Group 2: Subcutaneous masses were palpable at all 12 sites. The average size decreased significantly from 1.375 cm (1 mo) to 0.75 cm (3 mo) and 0.7 cm (6 mo). At 1 month postoperatively, all 4 sites had formed cysts, and body hair was found inside. However, the body hair in the cyst was thinner and shorter compared with the other group, and no serous fluid was observed (Fig. 5A). At 3 months postoperatively, 1 of the 4 sites had formed a very small cyst and contained no obvious body hairs inside, whereas the others had only formed scar-like induration (Fig. 5B). At 6 months postoperatively, just a small lump was palpable except for a very small cyst in 1 site (Fig. 5C).

Fig. 4.

Fig. 4.

Open in a new tab

Local findings in group 1. A, At 1 month postoperatively, a cyst containing hard body hairs and foul serous fluid formed under the skin. B, At 3 months postoperatively, a cyst containing body hairs and serous fluid formed. However, the body hairs were thinner and mixed with wastes compared with those at 1 month postoperatively. C, At 6 months postoperatively, a cyst was formed, but the body hairs contained within were finer and contained a greater proportion of waste material.

Fig. 5.

Fig. 5.

Open in a new tab

Local findings in group 2. A, At 1 month postoperatively, a cyst containing body hairs, which were thinner and shorter than those in group 1, was observed. The cyst did not contain a serous component. B, At 3 months postoperatively, a cyst with only a small lumen was formed and contained no obvious body hairs. C, At 6 months postoperatively, only a small lump was palpable.

Pathological Findings

  • Group 1: One large cyst was detected in each site at 1 month postoperatively. The cyst wall was formed by stratified squamous epithelium, with keratinocyte denuclearization and keratinization toward the lumen, and unstructured waste material was found inside. The cyst wall included many skin appendages and was stained with anti-cytokeratin 5 antibody. Similar findings were seen at 3 and 6 months postoperatively (Fig. 6).

  • Group 2: The aggregate of cysts of various sizes and irregular shapes was observed in each site at 1 month postoperatively. The cyst walls consisted of squamous epithelium of similar thickness to that of the other group and were stained with anti-cytokeratin 5 antibody. No hair follicles were observed on the cyst wall, unlike in group 1(Figs. 7A, B). However, at 3 months postoperatively, the cysts forming the aggregate were smaller, their walls were thinner and not stained by immunostaining, and the sparsely stained areas corresponded to body hairs (Figs. 7C, D). At 6 months postoperatively, there was no obvious cyst, and it had been replaced by a granulation-like tissue composed of histiocytes and multinucleated giant cells. No obvious body hairs were observed, and the immunostaining areas were very sparse (Figs. 7E, F).

Fig. 6.

Fig. 6.

Open in a new tab

Pathological findings in group 1. At all postoperative time points, 1 cyst per site was observed, and the cyst wall formed by differentiated epidermis contained numerous hair follicles and was stained with anti-cytokeratin 5 antibodies. A, At 1 month postoperatively, HE staining was performed. B, At 1 month postoperatively, immunohistochemical staining was performed. C, At 3 months postoperatively, HE staining was performed. D, At 3 months postoperatively, immunohistochemical staining was used. E, At 6 months postoperatively, HE staining was performed. F, At 6 months postoperatively, immunohistochemical staining was performed. HE, hematoxylin and eosin.

Fig. 7.

Fig. 7.

Open in a new tab

Pathological findings in group 2. A, At 1 month postoperatively, HE staining showed a cluster of small, irregularly shaped cysts. There were no hair follicles, as seen in group 1. B, At 1 month postoperatively, immunohistochemical staining showed that the cyst wall was stained with anti-cytokeratin 5 antibody. C, At 3 months postoperatively, HE staining showed that the cysts that formed clusters were smaller than those at 1 month postoperatively. D, At 3 months postoperatively, immunohistochemical staining showed that the cyst wall was not stained with anti-cytokeratin 5 antibody. E, At 6 months postoperatively, HE staining showed that the cluster had been replaced by a granulation-like tissue composed of histiocytes and multinucleated giant cells. F, At 6 months postoperatively, immunohistochemical staining showed that the stained areas were very sparse.

Molecular Analysis

Western blot analysis revealed that the keratin 5 expression level was similar throughout the entire postoperative period in group 1. However, in group 2, it decreased significantly between the 1-month postoperative time point and after 3 months (Fig. 8). In group 2, a band of higher molecular weight than keratin 5 was observed, possibly due to glycosylation or other effects, but changes in the amount of the target protein over time were evident. Quantitative real-time PCR also showed a significant decrease in keratin 5 mRNA levels over time in group 2, whereas no obvious change was observed throughout the entire postoperative period in group 1 (Fig. 9).

Fig. 8.

Fig. 8.

Open in a new tab

Western blotting was performed to examine the expression of keratin 5 to determine whether epidermal differentiation occurs. In group 1, the expression level of keratin 5 remained the same for all postoperative periods. In group 2, expression was observed at the 1-month postoperative time point but not after 3 months.

Fig. 9.

Fig. 9.

Open in a new tab

Quantitative real-time PCR was performed to measure the amount of mRNA for keratin 5 with cyclophilin as the housekeeping gene. In group 1, no change over time was observed in the keratin 5 mRNA level. In group 2, there was a significant decrease in keratin 5 mRNA levels between the 1-month postoperative time point and after 3 months. Asterisk (*) indicate statistically significant differences P < 0.05.

DISCUSSION

It is generally believed that a fully differentiated epidermal layer in vitro requires culture under air-liquid interface conditions, as atmospheric exposure is crucial for normal keratinization.12 Teshima et al13 reported that epidermal layers in submerged cultures exhibited incomplete and abnormal differentiation; however, appropriate oxygen supply in the culture medium stimulated keratinocyte differentiation comparable to that observed under air-liquid interface conditions.

In group 1, despite complete subcutaneous embedding of the epidermis without atmospheric exposure, cysts filled with waste material formed. The keratinocytes forming the cyst walls exhibited progressive denucleation toward the lumen. The presence of numerous skin appendages in the cyst walls suggests that these cysts originated from the skin of the transposed flap. Keratin 5 expression persisted subcutaneously for up to 6 months postoperatively, as confirmed by Western blotting and quantitative real-time PCR. These findings suggest that the soft tissue covering the epidermis of the transposed flap provided an environment conducive to normal keratinization and differentiation, potentially due to sufficient oxygen supply. This implies that keratinocytes followed the complete differentiation cycle, including shedding, and continued expelling dermal waste over an extended period. The interfollicular epidermal stem cells in the epidermal basal layer may have contributed to sustained keratinocyte differentiation in this group.14,15

In group 2, cysts formed at 1 month postoperatively, likely originating from the epithelial components of residual hair follicles, as indicated by the absence of structural elements in the cyst walls except for small sweat gland–like structures. However, by 3 and 6 months, cyst formation was significantly reduced, with only 1 case per time point showing cyst presence. Western blotting and quantitative PCR demonstrated a time-dependent decline in keratin 5 expression. These findings suggest that in de-epithelialized flaps implanted subcutaneously, keratinocytes derived from deep hair follicles are eventually resorbed. Hair follicle stem cells (HFSCs), residing in the bulge region of hair follicles, are known for their role in epidermal regeneration and hair renewal.16–20 However, in this study, the remaining HFSCs did not induce keratinocyte differentiation or contribute to cyst formation subcutaneously during the long period of time. As noted in the Introduction section, Singh et al6 reported epithelial islands forming around skin appendages in exposed de-epithelialized flaps. This suggests that although HFSCs retain their regenerative potential, they are functionally suppressed when embedded subcutaneously.

In group 1, cyst formation was limited to a single large cyst per site, and no follicular enlargement was observed, unlike in group 2. This may be because waste products from follicles were expelled into the cyst, preventing accumulation within the follicles.

The development of hair follicles is regulated by signaling molecules secreted from HFSCs, which enhance dermal papilla cell activity and promote hair growth.21,22 In group 1, thick body hairs were observed within the cysts at 1 month postoperatively. However, hair accumulation was unexpectedly sparse, despite the retention of hair follicle structures at 6 months. Similarly, in group 2, fine body hairs were observed subcutaneously but disappeared by 6 months. These findings suggest that hair follicle function is progressively suppressed in subcutaneous environments, regardless of de-epithelialization. Possible mechanisms include lack of atmospheric exposure or ischemia due to tissue pressure, similar to the pathophysiology of pressure-induced alopecia. Further investigation into the mechanisms underlying subcutaneous suppression of hair follicle function could provide insights for alopecia treatment.

Despite the presence of subcutaneous body hair postoperatively, no cases of pilonidal sinus–like pathology were observed. Pilonidal sinus disease is thought to result from a combination of 3 factors: free hair invasion, embedding forces, and skin vulnerability in the gluteal region.23 This suggests that freely embedded hairs contribute to pilonidal sinus formation. However, in this study, body hairs remained encapsulated within cysts, likely preventing severe foreign body reactions.

If these findings are applied to humans, species differences must be considered. Rats and humans have similar skin thicknesses, but humans exhibit thinner keratin layers, thicker epidermis, and lower follicle density.24 In human skin, except for the palms and soles of the feet, the thickness of the epidermis is the same everywhere, but the density of skin appendages varies by sex, race, and body part. Given that cysts in group 2 appeared to originate from individual follicles, transient cyst formation may also occur in human de-epithelialized flaps, particularly when harvested from hair-dense regions. Further studies are needed to verify this possibility.

Our findings align with previous research by Peer and Paddock,9 who observed cyst formation in subcutaneously embedded epidermis using skin grafts. Unlike skin grafts, skin flaps retain a stable blood supply and include deeper follicular structures, such as the dermal papilla. This raised concerns about potential subcutaneous body hair development. However, our study demonstrates that subcutaneous conditions gradually suppress hair follicle activity. Although HFSCs in de-epithelialized flaps retain their native functions, keratin 5 expression monitoring indicates that keratinocyte differentiation does not occur subcutaneously over time.

The occasional cyst formation in group 2 at 3 and 6 months may be attributed to incomplete de-epithelialization. As in clinical practice, de-epithelialization in this experiment was assessed visually. Alternative methods, such as small dermatomes or vegetable peelers, were considered but deemed impractical for rats. Chemical peeling was also unsuitable due to potential subcutaneous inflammation. Future research should also focus on optimizing de-epithelialization techniques and minimizing oversight during the process.

Despite limitations such as species differences, small sample size, and challenges in achieving reliable de-epithelialization, this study suggests that failure to remove the epidermis in skin flap transposition may lead to complications. Proper de-epithelialization is crucial for reducing such risks.

CONCLUSIONS

When the epidermis is placed subcutaneously in skin flap transposition, complications may arise due to the continued differentiation of keratinocytes. In contrast, de-epithelialized flaps demonstrate a reduced risk of such complications, as the remaining epithelial components and skin appendages gradually regress over time. These findings underscore the clinical significance of de-epithelialization in subcutaneous skin flap transposition.

DISCLOSURE

The authors have no financial interest to declare in relation to the content of this article.

Footnotes

Published online 27 August 2025.

Disclosure statements are at the end of this article, following the correspondence information.

REFERENCES

  • 1.Serafin D, Georgiade NG. Microsurgical composite tissue transplantation. Ann Surg. 1978;187:620–628. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Lynch JB, Madden JJ, Jr, Franklin JD. Breast reconstruction following mastectomy for cancer. Ann Surg. 1978;187:490–501. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Cannady SB, Seth R, Fritz MA, et al. Total parotidectomy defect reconstruction using the buried free flap. Otolaryngol Head Neck Surg. 2010;143:637–643. [DOI] [PubMed] [Google Scholar]
  • 4.Laungani AT, Van Alphen N, Christner JA, et al. Three-dimensional CT angiography assessment of the impact of the dermis and the subdermal plexus in DIEP flap perfusion. J Plast Reconstr Aesthet Surg. 2015;68:525–530. [DOI] [PubMed] [Google Scholar]
  • 5.Schneider MR, Schmidt-Ullrich R, Paus R. The hair follicle as a dynamic miniorgan. Curr Biol. 2009;19:R132–R142. [DOI] [PubMed] [Google Scholar]
  • 6.Singh M, Carty M, Nuutila K, et al. Saved by de-epithelialization: DIEP flap dermal skin regeneration salvage after mastectomy skin flap loss. Plast Reconstr Surg Glob Open. 2015;3:e511. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7.Williams R, Westgate GE, Pawlus AD, et al. Age-related changes in female scalp dermal sheath and dermal fibroblasts: how the hair follicle environment impacts hair aging. J Invest Dermatol. 2021;141:1041–1051. [DOI] [PubMed] [Google Scholar]
  • 8.Feleshtynskyi Y, Balan I, Dyadyk O, et al. Morphological features of pilonidal cysts of the sacrococcygeal region. Pol Przegl Chir. 2021;94:20–27. [DOI] [PubMed] [Google Scholar]
  • 9.Peer L, Paddock R. Histologic studies on the fate of deeply implanted dermal grafts observations on sections of implants buried from one week to one year. Arch Surg. 1937;34:268. [Google Scholar]
  • 10.Dimitroulis G, Slavin J. Histological evaluation of full thickness skin as an interpositional graft in the rabbit craniomandibular joint. J Oral Maxillofac Surg. 2006;64:1075–1080. [DOI] [PubMed] [Google Scholar]
  • 11.Moll R, Divo M, Langbein L. The human keratins: biology and pathology. Histochem Cell Biol. 2008;129:705–733. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Seo A, Kitagawa N, Matsuura T, et al. Formation of keratinocyte multilayers on filters under airlifted or submerged culture conditions in medium containing calcium, ascorbic acid, and keratinocyte growth factor. Histochem Cell Biol. 2016;146:585–597. [DOI] [PubMed] [Google Scholar]
  • 13.Teshima H, Endo M, Furuyama Y, et al. Involvement of hypoxia-inducible factor activity in inevitable air-exposure treatment upon differentiation in a three-dimensional keratinocyte culture. FEBS J. 2023;290:2049–2063. [DOI] [PubMed] [Google Scholar]
  • 14.Watt FM. Epidermal stem cells: markers, patterning and the control of stem cell fate. Philos Trans R Soc Lond B Biol Sci. 1998;353:831–837. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Lowell S, Watt FM. Delta regulates keratinocyte spreading and motility independently of differentiation. Mech Dev. 2001;107:133–140. [DOI] [PubMed] [Google Scholar]
  • 16.Hoffman RM. The pluripotency of hair follicle stem cells. Cell Cycle. 2006;5:232–233. [DOI] [PubMed] [Google Scholar]
  • 17.Alonso L, Fuchs E. The hair cycle. J Cell Sci. 2006;119:391–393. [DOI] [PubMed] [Google Scholar]
  • 18.Garcin CL, Ansell DM, Headon DJ, et al. Hair follicle bulge stem cells appear dispensable for the acute phase of wound re-epithelialization. Stem Cells. 2016;34:1377–1385. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Takeo M, Lee W, Ito M. Wound healing and skin regeneration. Cold Spring Harb Perspect Med. 2015;5:a023267. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Taylor G, Lehrer MS, Jensen PJ, et al. Involvement of follicular stem cells in forming not only the follicle but also the epidermis. Cell. 2000;102:451–461. [DOI] [PubMed] [Google Scholar]
  • 21.Kim Y, Kim SB, Lee H, et al. AIMP1-derived peptide secreted from hair follicle stem cells promotes hair growth by activating dermal papilla cells. Int J Biol Sci. 2024;20:5764–5778. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Driskell RR, Clavel C, Rendl M, et al. Hair follicle dermal papilla cells at a glance. J Cell Sci. 2011;124:1179–1182. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Kanat BH, Sözen S. Disease that should be remembered: sacrococcygeal pilonidal sinus disease and short history. World J Clin Cases. 2015;3:876–879. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Jung EC, Maibach HI. Animal models for percutaneous absorption. J Appl Toxicol. 2015;35:1–10. [DOI] [PubMed] [Google Scholar]

Articles from Plastic and Reconstructive Surgery Global Open are provided here courtesy of Wolters Kluwer Health

RESOURCES