Leonard C. Sperling, MD

To the Editor:

Temporal triangular alopecia (TTA), a condition first described by Sabouraud¹ in 1905, is a circumscribed nonscarring form of alopecia. Also referred to as congenital triangular alopecia, TTA presents as a triangular or lancet-shaped area of hair loss involving the frontotemporal hairline. Temporal triangular alopecia is characterized histologically by a normal number of miniaturized hair follicles without notable inflammation.² Although the majority of cases arise between birth and 9 years of age,^3,4 rare cases of adult-onset TTA also have been reported.^5,6 Adult-onset cases can cause notable diagnostic confusion and inappropriate treatment, as reported in our patient.

A 25-year-old woman with a history of Hashimoto thyroiditis presented with hair loss affecting the right temporal scalp of 3 years' duration that was first noticed by her husband. The lesion was an asymptomatic, 6×8-cm, roughly lancet-shaped patch of alopecia located on the right temporal scalp, bordering on the frontal hairline (Figure 1). Centrally, the patch appeared almost hairless with a few retained terminal hairs. The frontal hairline was thinned but still present. There was no scaling or erythema, and fine vellus hairs and a few isolated terminal hairs covered the area. The corresponding skin on the contralateral temporal scalp showed normal hair density. The patient insisted that she had normal hair at the affected area until 22 years of age, and she denied a history of trauma or tight hairstyles. Initially diagnosed with alopecia areata by her primary care provider, the patient was treated with topical corticosteroids for 6 months without benefit. She was subsequently referred to a dermatologist who again offered a diagnosis of alopecia areata and treated the lesions with 2 intralesional corticosteroid injections without benefit. No biopsies of the affected area were performed, and the patient was given a trial of topical minoxidil.

The patient consulted a new primary care provider and was diagnosed with scarring alopecia. She was referred to our dermatology department for further treatment. An initial biopsy at the edge of the affected area was interpreted as normal, but after failing additional intralesional corticosteroid injections, she was referred to our hair clinic where another biopsy was performed in the central portion of the lesion. A 4-mm diameter punch biopsy specimen revealed a normal epidermis and dermis; however, in the lower dermis only a single terminal follicle was seen (Figure 2). Sections through the upper dermis (Figure 3) showed that the total number of hairs was normal or nearly normal with at least 22 follicles, but most were vellus and indeterminate hairs with only a single terminal hair. The dermal architecture was otherwise normal. Given the clinical and histologic findings, a diagnosis of TTA was made. Subsequent to the diagnosis, the patient did not pursue any additional treatment options and preferred to style her hair so that the area of TTA remained covered.

The differential diagnosis in adults presenting with a patch of localized alopecia includes alopecia areata, trichotillomania, pressure-induced alopecia, traction alopecia, lichen planopilaris, discoid lupus erythematosus, and rarely TTA. Temporal triangular alopecia is a fairly common, if underreported, nonscarring form of alopecia that mainly affects young children. A PubMed search of articles indexed for MEDLINE using the terms temporal triangular alopecia or congenital triangular alopecia or triangular alopecia documented only 76 cases of TTA including our own, with the majority of patients diagnosed before 9 years of age. Only 2 cases of adult-onset TTA have been reported,^5,6 possibly leading to misdiagnosis of adult patients who present with similar areas of hair loss. As with some prior cases of TTA,^5,7 our patient was misdiagnosed with alopecia areata and scarring alopecia, both treated unsuccessfully before a diagnosis of TTA was considered. Clues to the diagnosis included the location, the lack of change in size and shape, the lack of response to intralesional corticosteroids, and the presence of numerous vellus hairs on the surface. A biopsy of the visibly hairless zone was confirmatory. The normal or nearly normal number of miniaturized hairs in specimens of TTA suggest that topical minoxidil therapy (eg, 5% solution twice daily for at least 6 months) might be useful, but the authors have tried it on a few other patients with clinically typical TTA without discernible benefit. When lesions are small, excision provides a fast and permanent solution to the problem, albeit with the usual risks of minor surgery.

To the Editor:

Temporal triangular alopecia (TTA), a condition first described by Sabouraud¹ in 1905, is a circumscribed nonscarring form of alopecia. Also referred to as congenital triangular alopecia, TTA presents as a triangular or lancet-shaped area of hair loss involving the frontotemporal hairline. Temporal triangular alopecia is characterized histologically by a normal number of miniaturized hair follicles without notable inflammation.² Although the majority of cases arise between birth and 9 years of age,^3,4 rare cases of adult-onset TTA also have been reported.^5,6 Adult-onset cases can cause notable diagnostic confusion and inappropriate treatment, as reported in our patient.

A 25-year-old woman with a history of Hashimoto thyroiditis presented with hair loss affecting the right temporal scalp of 3 years' duration that was first noticed by her husband. The lesion was an asymptomatic, 6×8-cm, roughly lancet-shaped patch of alopecia located on the right temporal scalp, bordering on the frontal hairline (Figure 1). Centrally, the patch appeared almost hairless with a few retained terminal hairs. The frontal hairline was thinned but still present. There was no scaling or erythema, and fine vellus hairs and a few isolated terminal hairs covered the area. The corresponding skin on the contralateral temporal scalp showed normal hair density. The patient insisted that she had normal hair at the affected area until 22 years of age, and she denied a history of trauma or tight hairstyles. Initially diagnosed with alopecia areata by her primary care provider, the patient was treated with topical corticosteroids for 6 months without benefit. She was subsequently referred to a dermatologist who again offered a diagnosis of alopecia areata and treated the lesions with 2 intralesional corticosteroid injections without benefit. No biopsies of the affected area were performed, and the patient was given a trial of topical minoxidil.

The patient consulted a new primary care provider and was diagnosed with scarring alopecia. She was referred to our dermatology department for further treatment. An initial biopsy at the edge of the affected area was interpreted as normal, but after failing additional intralesional corticosteroid injections, she was referred to our hair clinic where another biopsy was performed in the central portion of the lesion. A 4-mm diameter punch biopsy specimen revealed a normal epidermis and dermis; however, in the lower dermis only a single terminal follicle was seen (Figure 2). Sections through the upper dermis (Figure 3) showed that the total number of hairs was normal or nearly normal with at least 22 follicles, but most were vellus and indeterminate hairs with only a single terminal hair. The dermal architecture was otherwise normal. Given the clinical and histologic findings, a diagnosis of TTA was made. Subsequent to the diagnosis, the patient did not pursue any additional treatment options and preferred to style her hair so that the area of TTA remained covered.

The differential diagnosis in adults presenting with a patch of localized alopecia includes alopecia areata, trichotillomania, pressure-induced alopecia, traction alopecia, lichen planopilaris, discoid lupus erythematosus, and rarely TTA. Temporal triangular alopecia is a fairly common, if underreported, nonscarring form of alopecia that mainly affects young children. A PubMed search of articles indexed for MEDLINE using the terms temporal triangular alopecia or congenital triangular alopecia or triangular alopecia documented only 76 cases of TTA including our own, with the majority of patients diagnosed before 9 years of age. Only 2 cases of adult-onset TTA have been reported,^5,6 possibly leading to misdiagnosis of adult patients who present with similar areas of hair loss. As with some prior cases of TTA,^5,7 our patient was misdiagnosed with alopecia areata and scarring alopecia, both treated unsuccessfully before a diagnosis of TTA was considered. Clues to the diagnosis included the location, the lack of change in size and shape, the lack of response to intralesional corticosteroids, and the presence of numerous vellus hairs on the surface. A biopsy of the visibly hairless zone was confirmatory. The normal or nearly normal number of miniaturized hairs in specimens of TTA suggest that topical minoxidil therapy (eg, 5% solution twice daily for at least 6 months) might be useful, but the authors have tried it on a few other patients with clinically typical TTA without discernible benefit. When lesions are small, excision provides a fast and permanent solution to the problem, albeit with the usual risks of minor surgery.

To the Editor:

Temporal triangular alopecia (TTA), a condition first described by Sabouraud¹ in 1905, is a circumscribed nonscarring form of alopecia. Also referred to as congenital triangular alopecia, TTA presents as a triangular or lancet-shaped area of hair loss involving the frontotemporal hairline. Temporal triangular alopecia is characterized histologically by a normal number of miniaturized hair follicles without notable inflammation.² Although the majority of cases arise between birth and 9 years of age,^3,4 rare cases of adult-onset TTA also have been reported.^5,6 Adult-onset cases can cause notable diagnostic confusion and inappropriate treatment, as reported in our patient.

A 25-year-old woman with a history of Hashimoto thyroiditis presented with hair loss affecting the right temporal scalp of 3 years' duration that was first noticed by her husband. The lesion was an asymptomatic, 6×8-cm, roughly lancet-shaped patch of alopecia located on the right temporal scalp, bordering on the frontal hairline (Figure 1). Centrally, the patch appeared almost hairless with a few retained terminal hairs. The frontal hairline was thinned but still present. There was no scaling or erythema, and fine vellus hairs and a few isolated terminal hairs covered the area. The corresponding skin on the contralateral temporal scalp showed normal hair density. The patient insisted that she had normal hair at the affected area until 22 years of age, and she denied a history of trauma or tight hairstyles. Initially diagnosed with alopecia areata by her primary care provider, the patient was treated with topical corticosteroids for 6 months without benefit. She was subsequently referred to a dermatologist who again offered a diagnosis of alopecia areata and treated the lesions with 2 intralesional corticosteroid injections without benefit. No biopsies of the affected area were performed, and the patient was given a trial of topical minoxidil.

The patient consulted a new primary care provider and was diagnosed with scarring alopecia. She was referred to our dermatology department for further treatment. An initial biopsy at the edge of the affected area was interpreted as normal, but after failing additional intralesional corticosteroid injections, she was referred to our hair clinic where another biopsy was performed in the central portion of the lesion. A 4-mm diameter punch biopsy specimen revealed a normal epidermis and dermis; however, in the lower dermis only a single terminal follicle was seen (Figure 2). Sections through the upper dermis (Figure 3) showed that the total number of hairs was normal or nearly normal with at least 22 follicles, but most were vellus and indeterminate hairs with only a single terminal hair. The dermal architecture was otherwise normal. Given the clinical and histologic findings, a diagnosis of TTA was made. Subsequent to the diagnosis, the patient did not pursue any additional treatment options and preferred to style her hair so that the area of TTA remained covered.

The differential diagnosis in adults presenting with a patch of localized alopecia includes alopecia areata, trichotillomania, pressure-induced alopecia, traction alopecia, lichen planopilaris, discoid lupus erythematosus, and rarely TTA. Temporal triangular alopecia is a fairly common, if underreported, nonscarring form of alopecia that mainly affects young children. A PubMed search of articles indexed for MEDLINE using the terms temporal triangular alopecia or congenital triangular alopecia or triangular alopecia documented only 76 cases of TTA including our own, with the majority of patients diagnosed before 9 years of age. Only 2 cases of adult-onset TTA have been reported,^5,6 possibly leading to misdiagnosis of adult patients who present with similar areas of hair loss. As with some prior cases of TTA,^5,7 our patient was misdiagnosed with alopecia areata and scarring alopecia, both treated unsuccessfully before a diagnosis of TTA was considered. Clues to the diagnosis included the location, the lack of change in size and shape, the lack of response to intralesional corticosteroids, and the presence of numerous vellus hairs on the surface. A biopsy of the visibly hairless zone was confirmatory. The normal or nearly normal number of miniaturized hairs in specimens of TTA suggest that topical minoxidil therapy (eg, 5% solution twice daily for at least 6 months) might be useful, but the authors have tried it on a few other patients with clinically typical TTA without discernible benefit. When lesions are small, excision provides a fast and permanent solution to the problem, albeit with the usual risks of minor surgery.

Case Report

A 44-year-old woman presented with a localized patch of hair loss on the frontal scalp of several month’s duration. She had been bitten by a tick at this site during the summer. Two months later a primary care provider prescribed triamcinolone cream because of intense itching at the bite site. The patient returned to her primary care provider 2 weeks later due to persistent itching, hyperpigmentation, and hair loss. At that time, the clinician probed the central portion of the lesion because of a concern for retained tick parts. A few weeks later, a dermatologist evaluated the patient and found a roughly circular zone of alopecia measuring approximately 5 cm in diameter that was just posterior to the left frontal hairline (Figure 1). In the center of the plaque there was a small eschar surrounded by a zone of hyperpigmentation, mild induration, and almost complete loss of terminal hairs. At the periphery of the lesion, hair density gradually increased and skin pigmentation normalized.

A punch biopsy was obtained from an indurated area of hyperpigmentation adjacent to the eschar. Both vertical and horizontal sections were obtained, revealing a relatively normal epidermis, a marked decrease in follicular structures with loss of sebaceous glands, and dense perifollicular lymphocytic inflammation with a few scattered eosinophils (Figures 2 and 3).

Historical Perspective

Tick bite alopecia was first described in the French literature in 1921¹ and in the English-language literature in 1955.² A few additional cases were subsequently reported.^3-5 In 2008, Castelli et al⁶ described the histologic and immunohistochemical features of 25 tick bite cases, a few of which resulted in alopecia. Other than these reports, little original information has been written about tick bite alopecia.

Clinical and Histologic Presentation

Tick bite alopecia is well described in the veterinary literature.^7-9 It is possible that the condition is underreported in humans because the cause is often obvious or the alopecia is never discovered. The typical presentation is a roughly oval zone of alopecia that develops 1 to 2 weeks after the removal of a tick from the scalp. Often there is a small central eschar representing the site of tick attachment and the surrounding scalp may appear scaly. In one report of 2 siblings, multiple oval zones of alopecia resembling the moth-eaten alopecia of syphilis were noted in both patients, but only a single attached tick was found.² In some reported cases, hair loss was only temporary, and at least partial if not complete regrowth of hair occurred.^3,4 Follow-up on most cases is not provided, but to our knowledge permanent alopecia has not been described.

Information about the histologic findings of tick bite alopecia is particularly limited. In a report by Heyl,³ biopsies were conducted in 2 patients, but the areas selected for biopsy were the sites of tick attachment. Centrally dense, acute, and chronic inflammation was seen, as well as marked tissue necrosis of the connective tissue and hair follicles. Peripheral to the attachment zone, tissue necrosis was not found, but telogen hairs with “crumpled up hair shafts” were present.³ The histologic findings presented by Castelli et al⁶ were based on a single case of tick bite alopecia; however, the specimen was a generous excisional biopsy, allowing for a panoramic histologic view of the lesion. In the center of the specimen, hair follicles were absent, but residual follicular streamers and follicular remnants were surrounded by lymphocytic inflammation. Sebaceous glands were conspicuously absent, but foci with naked hairs, fibrosis, and granulomatous inflammation were seen. Peripherally, the hair follicles were thinned and miniaturized with an increased number of catagen/telogen hairs. Some follicles showed lamellar fibroplasia and perifollicular chronic inflammation. The inflammatory infiltrate consisted predominantly of helper T cells with a smaller population of B lymphocytes and a few plasma cells.⁶ In 2016, Lynch et al⁵ described a single case of tick bite alopecia and noted pseudolymphomatous inflammation with germinal center formation associated with hair miniaturization and an elevated catagen/telogen count; focal follicular mucinosis also was noted.Our histologic findings are similar to those of Castelli et al,⁶ except that the inflammatory infiltrate was clearly B-cell dominant, with a suggestion of germinal center formation, as noted by Lynch et al.⁵ This inflammatory pattern often can be encountered in a chronic tick bite lesion. Destruction of follicles and associated sebaceous glands and their replacement by follicular scars indicate that at least in the central portion of the lesion some permanent hair loss occurs. The presence of catagen/telogen hairs and miniaturized follicles indicates the potential for at least partial regrowth.

Similar to other investigators who have described tick bite alopecia, we can only speculate as to the mechanism by which clinical alopecia occurs. Given the density of the inflammatory infiltrate and perifollicular inflammation, it seems reasonable to assume that inflammation either destroys hair follicles or precipitates the catagen/telogen phase, resulting in temporary hair loss. The inflammation itself may be due to the presence of tick parts or the antigens in their saliva (or both). The delay between tick attachment and the onset of alopecia can be attributed to the time it takes follicles to cycle into the catagen/telogen phase and shed the hair shaft.

Case Report

A 44-year-old woman presented with a localized patch of hair loss on the frontal scalp of several month’s duration. She had been bitten by a tick at this site during the summer. Two months later a primary care provider prescribed triamcinolone cream because of intense itching at the bite site. The patient returned to her primary care provider 2 weeks later due to persistent itching, hyperpigmentation, and hair loss. At that time, the clinician probed the central portion of the lesion because of a concern for retained tick parts. A few weeks later, a dermatologist evaluated the patient and found a roughly circular zone of alopecia measuring approximately 5 cm in diameter that was just posterior to the left frontal hairline (Figure 1). In the center of the plaque there was a small eschar surrounded by a zone of hyperpigmentation, mild induration, and almost complete loss of terminal hairs. At the periphery of the lesion, hair density gradually increased and skin pigmentation normalized.

A punch biopsy was obtained from an indurated area of hyperpigmentation adjacent to the eschar. Both vertical and horizontal sections were obtained, revealing a relatively normal epidermis, a marked decrease in follicular structures with loss of sebaceous glands, and dense perifollicular lymphocytic inflammation with a few scattered eosinophils (Figures 2 and 3).

Historical Perspective

Tick bite alopecia was first described in the French literature in 1921¹ and in the English-language literature in 1955.² A few additional cases were subsequently reported.^3-5 In 2008, Castelli et al⁶ described the histologic and immunohistochemical features of 25 tick bite cases, a few of which resulted in alopecia. Other than these reports, little original information has been written about tick bite alopecia.

Clinical and Histologic Presentation

Tick bite alopecia is well described in the veterinary literature.^7-9 It is possible that the condition is underreported in humans because the cause is often obvious or the alopecia is never discovered. The typical presentation is a roughly oval zone of alopecia that develops 1 to 2 weeks after the removal of a tick from the scalp. Often there is a small central eschar representing the site of tick attachment and the surrounding scalp may appear scaly. In one report of 2 siblings, multiple oval zones of alopecia resembling the moth-eaten alopecia of syphilis were noted in both patients, but only a single attached tick was found.² In some reported cases, hair loss was only temporary, and at least partial if not complete regrowth of hair occurred.^3,4 Follow-up on most cases is not provided, but to our knowledge permanent alopecia has not been described.

Information about the histologic findings of tick bite alopecia is particularly limited. In a report by Heyl,³ biopsies were conducted in 2 patients, but the areas selected for biopsy were the sites of tick attachment. Centrally dense, acute, and chronic inflammation was seen, as well as marked tissue necrosis of the connective tissue and hair follicles. Peripheral to the attachment zone, tissue necrosis was not found, but telogen hairs with “crumpled up hair shafts” were present.³ The histologic findings presented by Castelli et al⁶ were based on a single case of tick bite alopecia; however, the specimen was a generous excisional biopsy, allowing for a panoramic histologic view of the lesion. In the center of the specimen, hair follicles were absent, but residual follicular streamers and follicular remnants were surrounded by lymphocytic inflammation. Sebaceous glands were conspicuously absent, but foci with naked hairs, fibrosis, and granulomatous inflammation were seen. Peripherally, the hair follicles were thinned and miniaturized with an increased number of catagen/telogen hairs. Some follicles showed lamellar fibroplasia and perifollicular chronic inflammation. The inflammatory infiltrate consisted predominantly of helper T cells with a smaller population of B lymphocytes and a few plasma cells.⁶ In 2016, Lynch et al⁵ described a single case of tick bite alopecia and noted pseudolymphomatous inflammation with germinal center formation associated with hair miniaturization and an elevated catagen/telogen count; focal follicular mucinosis also was noted.Our histologic findings are similar to those of Castelli et al,⁶ except that the inflammatory infiltrate was clearly B-cell dominant, with a suggestion of germinal center formation, as noted by Lynch et al.⁵ This inflammatory pattern often can be encountered in a chronic tick bite lesion. Destruction of follicles and associated sebaceous glands and their replacement by follicular scars indicate that at least in the central portion of the lesion some permanent hair loss occurs. The presence of catagen/telogen hairs and miniaturized follicles indicates the potential for at least partial regrowth.

Similar to other investigators who have described tick bite alopecia, we can only speculate as to the mechanism by which clinical alopecia occurs. Given the density of the inflammatory infiltrate and perifollicular inflammation, it seems reasonable to assume that inflammation either destroys hair follicles or precipitates the catagen/telogen phase, resulting in temporary hair loss. The inflammation itself may be due to the presence of tick parts or the antigens in their saliva (or both). The delay between tick attachment and the onset of alopecia can be attributed to the time it takes follicles to cycle into the catagen/telogen phase and shed the hair shaft.

Case Report

A 44-year-old woman presented with a localized patch of hair loss on the frontal scalp of several month’s duration. She had been bitten by a tick at this site during the summer. Two months later a primary care provider prescribed triamcinolone cream because of intense itching at the bite site. The patient returned to her primary care provider 2 weeks later due to persistent itching, hyperpigmentation, and hair loss. At that time, the clinician probed the central portion of the lesion because of a concern for retained tick parts. A few weeks later, a dermatologist evaluated the patient and found a roughly circular zone of alopecia measuring approximately 5 cm in diameter that was just posterior to the left frontal hairline (Figure 1). In the center of the plaque there was a small eschar surrounded by a zone of hyperpigmentation, mild induration, and almost complete loss of terminal hairs. At the periphery of the lesion, hair density gradually increased and skin pigmentation normalized.

A punch biopsy was obtained from an indurated area of hyperpigmentation adjacent to the eschar. Both vertical and horizontal sections were obtained, revealing a relatively normal epidermis, a marked decrease in follicular structures with loss of sebaceous glands, and dense perifollicular lymphocytic inflammation with a few scattered eosinophils (Figures 2 and 3).

Historical Perspective

Tick bite alopecia was first described in the French literature in 1921¹ and in the English-language literature in 1955.² A few additional cases were subsequently reported.^3-5 In 2008, Castelli et al⁶ described the histologic and immunohistochemical features of 25 tick bite cases, a few of which resulted in alopecia. Other than these reports, little original information has been written about tick bite alopecia.

Clinical and Histologic Presentation

Tick bite alopecia is well described in the veterinary literature.^7-9 It is possible that the condition is underreported in humans because the cause is often obvious or the alopecia is never discovered. The typical presentation is a roughly oval zone of alopecia that develops 1 to 2 weeks after the removal of a tick from the scalp. Often there is a small central eschar representing the site of tick attachment and the surrounding scalp may appear scaly. In one report of 2 siblings, multiple oval zones of alopecia resembling the moth-eaten alopecia of syphilis were noted in both patients, but only a single attached tick was found.² In some reported cases, hair loss was only temporary, and at least partial if not complete regrowth of hair occurred.^3,4 Follow-up on most cases is not provided, but to our knowledge permanent alopecia has not been described.

Information about the histologic findings of tick bite alopecia is particularly limited. In a report by Heyl,³ biopsies were conducted in 2 patients, but the areas selected for biopsy were the sites of tick attachment. Centrally dense, acute, and chronic inflammation was seen, as well as marked tissue necrosis of the connective tissue and hair follicles. Peripheral to the attachment zone, tissue necrosis was not found, but telogen hairs with “crumpled up hair shafts” were present.³ The histologic findings presented by Castelli et al⁶ were based on a single case of tick bite alopecia; however, the specimen was a generous excisional biopsy, allowing for a panoramic histologic view of the lesion. In the center of the specimen, hair follicles were absent, but residual follicular streamers and follicular remnants were surrounded by lymphocytic inflammation. Sebaceous glands were conspicuously absent, but foci with naked hairs, fibrosis, and granulomatous inflammation were seen. Peripherally, the hair follicles were thinned and miniaturized with an increased number of catagen/telogen hairs. Some follicles showed lamellar fibroplasia and perifollicular chronic inflammation. The inflammatory infiltrate consisted predominantly of helper T cells with a smaller population of B lymphocytes and a few plasma cells.⁶ In 2016, Lynch et al⁵ described a single case of tick bite alopecia and noted pseudolymphomatous inflammation with germinal center formation associated with hair miniaturization and an elevated catagen/telogen count; focal follicular mucinosis also was noted.Our histologic findings are similar to those of Castelli et al,⁶ except that the inflammatory infiltrate was clearly B-cell dominant, with a suggestion of germinal center formation, as noted by Lynch et al.⁵ This inflammatory pattern often can be encountered in a chronic tick bite lesion. Destruction of follicles and associated sebaceous glands and their replacement by follicular scars indicate that at least in the central portion of the lesion some permanent hair loss occurs. The presence of catagen/telogen hairs and miniaturized follicles indicates the potential for at least partial regrowth.

Similar to other investigators who have described tick bite alopecia, we can only speculate as to the mechanism by which clinical alopecia occurs. Given the density of the inflammatory infiltrate and perifollicular inflammation, it seems reasonable to assume that inflammation either destroys hair follicles or precipitates the catagen/telogen phase, resulting in temporary hair loss. The inflammation itself may be due to the presence of tick parts or the antigens in their saliva (or both). The delay between tick attachment and the onset of alopecia can be attributed to the time it takes follicles to cycle into the catagen/telogen phase and shed the hair shaft.

Potassium hydroxide (KOH) preparations remain an important bedside test for prompt and accurate diagnosis of superficial fungal infections known as dermatophytoses. This tool has been used for at least 100 years, with early terminology referring to it as potash; for the last century, it has largely been a technique passed down as a skill from master technician to learning apprentice. The original pioneer of the KOH preparation remains a mystery.¹

Variations on techniques for performing the KOH preparation exist, and tips and tricks on the use of this test are a hot topic among dermatologists.² Although primary care and dermatology-specific publications espouse the importance of the KOH preparation,^3,4 it has unfortunately been identified and labeled as one of the forgotten diagnostic tools.⁵

It is incumbent on dermatologists to educate medical students and residents using a simple and specific method to ensure that this simple and effective technique, with sensitivity reported between 87% and 91% depending on the expertise of the examiner,⁶ remains part of the clinical armamentarium. One concern in the instruction of large groups of students and clinicians is the ready accessibility or availability of viable skin samples. This article describes a method of collecting and storing skin samples that will allow educators to train large groups of students on performing KOH preparations without having to repeatedly seek skin samples or patients with superficial skin infections. A detailed description of the pedagogy used to teach the preparation and interpretation of KOH slides to a large group of students also is reviewed.

Specimen Collection

The first step in teaching the KOH preparation to a large group is the collection of a suitable number of skin scrapings from patients with a superficial fungal skin infection (eg, tinea corporis, tinea versicolor). A common technique for obtaining skin samples is to use a no. 15 scalpel blade (Figure 1) to scrape the scale of the lesion at its scaly border once the area is moistened with an alcohol pad or soap and water.⁷ The moisture from the alcohol pad allows the scale to stick to the no. 15 blade, facilitating collection. Once a suitable amount of scale is collected, it is placed on a glass microscope slide by smearing the scale from the blade onto the slide. This process has been modified to facilitate a larger quantity of specimen as follows: dermatophyte-infected plaques with scale are rubbed with the no. 15 blade and the free scale drops into a standard urine specimen cup. This process is repeated multiple times from different sites to capture the displaced scale with the dermatophyte. We have found that as long as the specimen cups are sealed tightly and stored in a relatively dry and cool environment (room temperature), the samples can be used to construct KOH teaching slides for at least 3 years. We have not used them beyond 3 years but suspect that they would continue to be viable after this time.

Figure 1. Scraping at leading edge of lesion with a no. 15 scalpel blade after cleaning the area with an alcohol pad.

Preparation of Slides

Given that time for teaching often is limited, it is beneficial to fix many skin scrapings on a large number of glass slides prior to the session, which enables students to simply add KOH to the slides on the teaching day. To prepare the slides in advance, it is necessary to gather the following materials: a specimen cup with skin samples, glass slides, pickups or tweezers, a small pipette, a cup of water, protective gloves, and a pencil. After donning protective gloves, the pickups or tweezers are used to retrieve a few flakes of scale from the specimen cup and place them on the center of a glass slide. Using the pipette, 1 or 2 drops of water are added to the scale, and the slide is then allowed to dry. The slides are marked with the pencil to indicate the “up” side to prevent the students from applying KOH solution to the wrong side of the slide. The skin scale is fixed in place on the slide as the water evaporates and may be stored until needed for use in a standard slide box or folder.

Performing the KOH Preparation

On the day of teaching, it is helpful to engage the entire group of students with an introductory lecture on the purpose and use of the KOH preparation. Upon completion, students move to a workstation with all of the materials needed to prepare the slide. Additional items needed at this time are 10% KOH solution, coverslips, and a heating device (eg, lighter, Bunsen burner, match)(optional). Students are instructed to place 1 or 2 skin scales onto a glass slide or retrieve a slide with skin scales already fixed, and then add 1 drop of 10% KOH solution directly to the sample (Figure 2). Next, they should place a slide coverslip onto the KOH drop and skin sample using a side-to-side technique that will move the scale into a thin layer within the KOH solution and push away any excess solution to the periphery (Figure 3). Large amounts of excess KOH solution should be cleared away with a paper towel, lens paper, or tissue. The heat source can be used to gently heat the underside of the glass slide (Figure 4), but it often is sufficient to simply wait 3 to 5 minutes for the KOH solution to take effect. The heat accelerates the maceration of the scale and makes it easier to see the hyphae among the keratinocytes. Some physicians advocate the use of dimethyl sulfoxide in lieu of heating,⁸ but this solution may not be available in all primary care settings.

Figure 2. Adding 10% potassium hydroxide solution to skin samples.

Figure 3. Placing the slide coverslip.

Figure 4. Applying gentle heat to the potassium hydroxide preparations.

Microscopic Examination

Prior to examining the slides under the microscope, students may complete a self-guided tutorial (eg, digital or paper slide show) on the various features seen through the microscope that are indicative of dermatophytes, including branching hyphae and yeast buds. They also should be educated about the common appearance of artifacts that may resemble hyphae. Once the students have completed the tutorial, they may proceed to microscopic examination.

 While the students are viewing their slides under the microscope, we find it helpful to have at least 1 experienced faculty member for every group of 10 students. This instructor should encourage the students to lower the microscope condenser all the way to facilitate better observation. Students should start with low power (×4 or red band) and scan for areas that are rich in skin scale. Once a collection of scale is found, the student can switch to higher power (×10 or yellow band) and start scanning for hyphae. Students should be reminded to search for filamentous and branching tubes that are refractile. The term refractile may be confusing to some students, so we explain that shifting the focus up or down will show the hyphae to change in brightness and may reveal a greenish tint. Another helpful indicator to point out is the feature that hyphae will cross the border of epidermal skin cells, whereas artifacts will not  (Figure 5). Once the students have identified evidence of a dermatophyte infection, they must call the instructor to their station to verify the presence of hyphae or yeast buds, which helps confirm their understanding of the procedure. Once the  student accurately identifies these items, the session is complete.

Figure 5. Example of hyphum crossing epithelial cell borders. Figure 5. Example of hyphum crossing epithelial cell borders.

Comment

The use of a KOH preparation is a fast, simple, accurate, and cost-effective way to diagnose superficial fungal infections; however, because of insufficient familiarity with this tool, the technique often is replaced by initiation of empiric antifungal therapy in patients with suspected dermatophytosis. This empiric treatment has the potential to delay appropriate diagnosis and treatment (eg, in a patient with nummular dermatitis, which can clinically mimic tinea corporis). One way to encourage the use of the KOH preparation in the primary care and dermatologic setting is to educate large groups of next-generation physicians while in medical training. This article describes a teaching technique that allows for long-term storage of positive skin samples and a detailed description of the pedagogy used to train and educate a large group of students in a relatively short period of time.

All KOH preparations fall under the US federal government’s Clinical Laboratory Improvement Amendments and require proficiency testing.⁹ Although the teaching method presented here is designed for teaching medical students, it may be utilized to educate or refamiliarize experienced physicians with the procedure in an effort to improve proficiency in point-of-care testing programs used in many health care systems to comply with the Clinical Laboratories Improvement Amendments. Future analyses could assess whether the method described here improves provider performance on such proficiency measures and whether it ultimately helps ensure quality patient care.

Potassium hydroxide (KOH) preparations remain an important bedside test for prompt and accurate diagnosis of superficial fungal infections known as dermatophytoses. This tool has been used for at least 100 years, with early terminology referring to it as potash; for the last century, it has largely been a technique passed down as a skill from master technician to learning apprentice. The original pioneer of the KOH preparation remains a mystery.¹

Variations on techniques for performing the KOH preparation exist, and tips and tricks on the use of this test are a hot topic among dermatologists.² Although primary care and dermatology-specific publications espouse the importance of the KOH preparation,^3,4 it has unfortunately been identified and labeled as one of the forgotten diagnostic tools.⁵

It is incumbent on dermatologists to educate medical students and residents using a simple and specific method to ensure that this simple and effective technique, with sensitivity reported between 87% and 91% depending on the expertise of the examiner,⁶ remains part of the clinical armamentarium. One concern in the instruction of large groups of students and clinicians is the ready accessibility or availability of viable skin samples. This article describes a method of collecting and storing skin samples that will allow educators to train large groups of students on performing KOH preparations without having to repeatedly seek skin samples or patients with superficial skin infections. A detailed description of the pedagogy used to teach the preparation and interpretation of KOH slides to a large group of students also is reviewed.

Specimen Collection

The first step in teaching the KOH preparation to a large group is the collection of a suitable number of skin scrapings from patients with a superficial fungal skin infection (eg, tinea corporis, tinea versicolor). A common technique for obtaining skin samples is to use a no. 15 scalpel blade (Figure 1) to scrape the scale of the lesion at its scaly border once the area is moistened with an alcohol pad or soap and water.⁷ The moisture from the alcohol pad allows the scale to stick to the no. 15 blade, facilitating collection. Once a suitable amount of scale is collected, it is placed on a glass microscope slide by smearing the scale from the blade onto the slide. This process has been modified to facilitate a larger quantity of specimen as follows: dermatophyte-infected plaques with scale are rubbed with the no. 15 blade and the free scale drops into a standard urine specimen cup. This process is repeated multiple times from different sites to capture the displaced scale with the dermatophyte. We have found that as long as the specimen cups are sealed tightly and stored in a relatively dry and cool environment (room temperature), the samples can be used to construct KOH teaching slides for at least 3 years. We have not used them beyond 3 years but suspect that they would continue to be viable after this time.

Figure 1. Scraping at leading edge of lesion with a no. 15 scalpel blade after cleaning the area with an alcohol pad.

Preparation of Slides

Given that time for teaching often is limited, it is beneficial to fix many skin scrapings on a large number of glass slides prior to the session, which enables students to simply add KOH to the slides on the teaching day. To prepare the slides in advance, it is necessary to gather the following materials: a specimen cup with skin samples, glass slides, pickups or tweezers, a small pipette, a cup of water, protective gloves, and a pencil. After donning protective gloves, the pickups or tweezers are used to retrieve a few flakes of scale from the specimen cup and place them on the center of a glass slide. Using the pipette, 1 or 2 drops of water are added to the scale, and the slide is then allowed to dry. The slides are marked with the pencil to indicate the “up” side to prevent the students from applying KOH solution to the wrong side of the slide. The skin scale is fixed in place on the slide as the water evaporates and may be stored until needed for use in a standard slide box or folder.

Performing the KOH Preparation

On the day of teaching, it is helpful to engage the entire group of students with an introductory lecture on the purpose and use of the KOH preparation. Upon completion, students move to a workstation with all of the materials needed to prepare the slide. Additional items needed at this time are 10% KOH solution, coverslips, and a heating device (eg, lighter, Bunsen burner, match)(optional). Students are instructed to place 1 or 2 skin scales onto a glass slide or retrieve a slide with skin scales already fixed, and then add 1 drop of 10% KOH solution directly to the sample (Figure 2). Next, they should place a slide coverslip onto the KOH drop and skin sample using a side-to-side technique that will move the scale into a thin layer within the KOH solution and push away any excess solution to the periphery (Figure 3). Large amounts of excess KOH solution should be cleared away with a paper towel, lens paper, or tissue. The heat source can be used to gently heat the underside of the glass slide (Figure 4), but it often is sufficient to simply wait 3 to 5 minutes for the KOH solution to take effect. The heat accelerates the maceration of the scale and makes it easier to see the hyphae among the keratinocytes. Some physicians advocate the use of dimethyl sulfoxide in lieu of heating,⁸ but this solution may not be available in all primary care settings.

Figure 2. Adding 10% potassium hydroxide solution to skin samples.

Figure 3. Placing the slide coverslip.

Figure 4. Applying gentle heat to the potassium hydroxide preparations.

Microscopic Examination

Prior to examining the slides under the microscope, students may complete a self-guided tutorial (eg, digital or paper slide show) on the various features seen through the microscope that are indicative of dermatophytes, including branching hyphae and yeast buds. They also should be educated about the common appearance of artifacts that may resemble hyphae. Once the students have completed the tutorial, they may proceed to microscopic examination.

 While the students are viewing their slides under the microscope, we find it helpful to have at least 1 experienced faculty member for every group of 10 students. This instructor should encourage the students to lower the microscope condenser all the way to facilitate better observation. Students should start with low power (×4 or red band) and scan for areas that are rich in skin scale. Once a collection of scale is found, the student can switch to higher power (×10 or yellow band) and start scanning for hyphae. Students should be reminded to search for filamentous and branching tubes that are refractile. The term refractile may be confusing to some students, so we explain that shifting the focus up or down will show the hyphae to change in brightness and may reveal a greenish tint. Another helpful indicator to point out is the feature that hyphae will cross the border of epidermal skin cells, whereas artifacts will not  (Figure 5). Once the students have identified evidence of a dermatophyte infection, they must call the instructor to their station to verify the presence of hyphae or yeast buds, which helps confirm their understanding of the procedure. Once the  student accurately identifies these items, the session is complete.

Figure 5. Example of hyphum crossing epithelial cell borders. Figure 5. Example of hyphum crossing epithelial cell borders.

Comment

The use of a KOH preparation is a fast, simple, accurate, and cost-effective way to diagnose superficial fungal infections; however, because of insufficient familiarity with this tool, the technique often is replaced by initiation of empiric antifungal therapy in patients with suspected dermatophytosis. This empiric treatment has the potential to delay appropriate diagnosis and treatment (eg, in a patient with nummular dermatitis, which can clinically mimic tinea corporis). One way to encourage the use of the KOH preparation in the primary care and dermatologic setting is to educate large groups of next-generation physicians while in medical training. This article describes a teaching technique that allows for long-term storage of positive skin samples and a detailed description of the pedagogy used to train and educate a large group of students in a relatively short period of time.

All KOH preparations fall under the US federal government’s Clinical Laboratory Improvement Amendments and require proficiency testing.⁹ Although the teaching method presented here is designed for teaching medical students, it may be utilized to educate or refamiliarize experienced physicians with the procedure in an effort to improve proficiency in point-of-care testing programs used in many health care systems to comply with the Clinical Laboratories Improvement Amendments. Future analyses could assess whether the method described here improves provider performance on such proficiency measures and whether it ultimately helps ensure quality patient care.

Potassium hydroxide (KOH) preparations remain an important bedside test for prompt and accurate diagnosis of superficial fungal infections known as dermatophytoses. This tool has been used for at least 100 years, with early terminology referring to it as potash; for the last century, it has largely been a technique passed down as a skill from master technician to learning apprentice. The original pioneer of the KOH preparation remains a mystery.¹

Variations on techniques for performing the KOH preparation exist, and tips and tricks on the use of this test are a hot topic among dermatologists.² Although primary care and dermatology-specific publications espouse the importance of the KOH preparation,^3,4 it has unfortunately been identified and labeled as one of the forgotten diagnostic tools.⁵

It is incumbent on dermatologists to educate medical students and residents using a simple and specific method to ensure that this simple and effective technique, with sensitivity reported between 87% and 91% depending on the expertise of the examiner,⁶ remains part of the clinical armamentarium. One concern in the instruction of large groups of students and clinicians is the ready accessibility or availability of viable skin samples. This article describes a method of collecting and storing skin samples that will allow educators to train large groups of students on performing KOH preparations without having to repeatedly seek skin samples or patients with superficial skin infections. A detailed description of the pedagogy used to teach the preparation and interpretation of KOH slides to a large group of students also is reviewed.

Specimen Collection

The first step in teaching the KOH preparation to a large group is the collection of a suitable number of skin scrapings from patients with a superficial fungal skin infection (eg, tinea corporis, tinea versicolor). A common technique for obtaining skin samples is to use a no. 15 scalpel blade (Figure 1) to scrape the scale of the lesion at its scaly border once the area is moistened with an alcohol pad or soap and water.⁷ The moisture from the alcohol pad allows the scale to stick to the no. 15 blade, facilitating collection. Once a suitable amount of scale is collected, it is placed on a glass microscope slide by smearing the scale from the blade onto the slide. This process has been modified to facilitate a larger quantity of specimen as follows: dermatophyte-infected plaques with scale are rubbed with the no. 15 blade and the free scale drops into a standard urine specimen cup. This process is repeated multiple times from different sites to capture the displaced scale with the dermatophyte. We have found that as long as the specimen cups are sealed tightly and stored in a relatively dry and cool environment (room temperature), the samples can be used to construct KOH teaching slides for at least 3 years. We have not used them beyond 3 years but suspect that they would continue to be viable after this time.

Figure 1. Scraping at leading edge of lesion with a no. 15 scalpel blade after cleaning the area with an alcohol pad.

Preparation of Slides

Given that time for teaching often is limited, it is beneficial to fix many skin scrapings on a large number of glass slides prior to the session, which enables students to simply add KOH to the slides on the teaching day. To prepare the slides in advance, it is necessary to gather the following materials: a specimen cup with skin samples, glass slides, pickups or tweezers, a small pipette, a cup of water, protective gloves, and a pencil. After donning protective gloves, the pickups or tweezers are used to retrieve a few flakes of scale from the specimen cup and place them on the center of a glass slide. Using the pipette, 1 or 2 drops of water are added to the scale, and the slide is then allowed to dry. The slides are marked with the pencil to indicate the “up” side to prevent the students from applying KOH solution to the wrong side of the slide. The skin scale is fixed in place on the slide as the water evaporates and may be stored until needed for use in a standard slide box or folder.

Performing the KOH Preparation

On the day of teaching, it is helpful to engage the entire group of students with an introductory lecture on the purpose and use of the KOH preparation. Upon completion, students move to a workstation with all of the materials needed to prepare the slide. Additional items needed at this time are 10% KOH solution, coverslips, and a heating device (eg, lighter, Bunsen burner, match)(optional). Students are instructed to place 1 or 2 skin scales onto a glass slide or retrieve a slide with skin scales already fixed, and then add 1 drop of 10% KOH solution directly to the sample (Figure 2). Next, they should place a slide coverslip onto the KOH drop and skin sample using a side-to-side technique that will move the scale into a thin layer within the KOH solution and push away any excess solution to the periphery (Figure 3). Large amounts of excess KOH solution should be cleared away with a paper towel, lens paper, or tissue. The heat source can be used to gently heat the underside of the glass slide (Figure 4), but it often is sufficient to simply wait 3 to 5 minutes for the KOH solution to take effect. The heat accelerates the maceration of the scale and makes it easier to see the hyphae among the keratinocytes. Some physicians advocate the use of dimethyl sulfoxide in lieu of heating,⁸ but this solution may not be available in all primary care settings.

Figure 2. Adding 10% potassium hydroxide solution to skin samples.

Figure 3. Placing the slide coverslip.

Figure 4. Applying gentle heat to the potassium hydroxide preparations.

Microscopic Examination

Prior to examining the slides under the microscope, students may complete a self-guided tutorial (eg, digital or paper slide show) on the various features seen through the microscope that are indicative of dermatophytes, including branching hyphae and yeast buds. They also should be educated about the common appearance of artifacts that may resemble hyphae. Once the students have completed the tutorial, they may proceed to microscopic examination.

 While the students are viewing their slides under the microscope, we find it helpful to have at least 1 experienced faculty member for every group of 10 students. This instructor should encourage the students to lower the microscope condenser all the way to facilitate better observation. Students should start with low power (×4 or red band) and scan for areas that are rich in skin scale. Once a collection of scale is found, the student can switch to higher power (×10 or yellow band) and start scanning for hyphae. Students should be reminded to search for filamentous and branching tubes that are refractile. The term refractile may be confusing to some students, so we explain that shifting the focus up or down will show the hyphae to change in brightness and may reveal a greenish tint. Another helpful indicator to point out is the feature that hyphae will cross the border of epidermal skin cells, whereas artifacts will not  (Figure 5). Once the students have identified evidence of a dermatophyte infection, they must call the instructor to their station to verify the presence of hyphae or yeast buds, which helps confirm their understanding of the procedure. Once the  student accurately identifies these items, the session is complete.

Figure 5. Example of hyphum crossing epithelial cell borders. Figure 5. Example of hyphum crossing epithelial cell borders.

Comment

The use of a KOH preparation is a fast, simple, accurate, and cost-effective way to diagnose superficial fungal infections; however, because of insufficient familiarity with this tool, the technique often is replaced by initiation of empiric antifungal therapy in patients with suspected dermatophytosis. This empiric treatment has the potential to delay appropriate diagnosis and treatment (eg, in a patient with nummular dermatitis, which can clinically mimic tinea corporis). One way to encourage the use of the KOH preparation in the primary care and dermatologic setting is to educate large groups of next-generation physicians while in medical training. This article describes a teaching technique that allows for long-term storage of positive skin samples and a detailed description of the pedagogy used to train and educate a large group of students in a relatively short period of time.

All KOH preparations fall under the US federal government’s Clinical Laboratory Improvement Amendments and require proficiency testing.⁹ Although the teaching method presented here is designed for teaching medical students, it may be utilized to educate or refamiliarize experienced physicians with the procedure in an effort to improve proficiency in point-of-care testing programs used in many health care systems to comply with the Clinical Laboratories Improvement Amendments. Future analyses could assess whether the method described here improves provider performance on such proficiency measures and whether it ultimately helps ensure quality patient care.