A New Technique for Quantitative Analysis of Hair Loss in Mice Using Grayscale Analysis

Summary

Alopecia is a common form of hair loss which can occur in many different conditions, including as a side-effect of chemotherapy. We have developed a method to quantify hair loss in mice, utilizing a standard gel imager to perform a grayscale analysis, to facilitate study of promising new alopecia therapies.

Abstract

Alopecia is a common form of hair loss which can occur in many different conditions, including male-pattern hair loss, polycystic ovarian syndrome, and alopecia areata. Alopecia can also occur as a side effect of chemotherapy in cancer patients. In this study, our goal was to develop a consistent and reliable method to quantify hair loss in mice, which will allow investigators to accurately assess and compare new therapeutic approaches for these various forms of alopecia. The method utilizes a standard gel imager to obtain and process images of mice, measuring the light absorption, which occurs in rough proportion to the amount of black (or gray) hair on the mouse. Data that has been quantified in this fashion can then be analyzed using standard statistical techniques (i.e., ANOVA, T-test). This methodology was tested in mouse models of chemotherapy-induced alopecia, alopecia areata and alopecia from waxing. In this report, the detailed protocol is presented for performing these measurements, including validation data from C57BL/6 and C3H/HeJ strains of mice. This new technique offers a number of advantages, including relative simplicity of application, reliance on equipment which is readily available in most research laboratories, and applying an objective, quantitative assessment which is more robust than subjective evaluations. Improvements in quantification of hair growth in mice will improve study of alopecia models and facilitate evaluation of promising new therapies in preclinical studies.

Introduction

Alopecia (hair loss) can be a psychologically and emotionally distressing event with multiple causes. Male-pattern baldness is the most common cause of alopecia, affecting approximately two thirds of males by age 35¹. A similar pattern of hair loss can be observed in females with polycystic ovarian syndrome. In both of these disorders, the hair loss is androgen mediated. Alopecia can also occur as an autoimmune disease, alopecia areata, which affects 1.7% of the population².  Alopecia can occur as a side-effect of some medical treatments, such as chemotherapy³.  A high percentage (65-85%) of chemotherapy patients experience some degree of alopecia^4,5. The psychological consequences of hair loss have been well studied in the chemotherapy setting. Chemotherapy-induced alopecia can result in anxiety, depression, a negative body image, lowered self-esteem and a reduced sense of well-being^6,7. A high percentage (47-58%) of female cancer patients consider hair loss to be the most traumatic aspect of chemotherapy, and up to 8% decline treatment for fear of hair loss^4,6. There is also evidence in androgenic alopecia to support therapy to reduce psychological and even medical consequences of hair loss^8,9. Likewise, alopecia areata has been reported to have severe psychological consequences², and the patchy nature of the hair loss can lead to a more unpleasant cosmetic result than most other causes of hair loss.

While drugs with mild anti-androgenic effects (i.e., spironolactone) had been used with limited success as therapy for alopecia, the first effective medication for alopecia was minoxidil¹⁰. This antihypertensive has an observed side-effect of causing hair growth, and is now used as topical therapy for many forms of alopecia. However, responses are often incomplete, with some subjects showing only slowing of hair loss rather than actual regrowth¹⁰. Finasteride is a competitive antagonist to type II 5α-reductase which blocks conversion of testosterone to dihydrotestosterone, resulting in improvements in androgenic alopecia at the expense of partial systemic androgen blockade. Response rates with long-term (10 years) therapy are around 50%¹¹. Overall, despite considerable research in this area, there is still no adequate therapy for hair loss.

For decades, scientists and clinicians have examined methods of measuring scalp hair growth in clinical trials. With the development of drugs that treat alopecia, there has been a greater need for reliable, economical and minimally invasive means of measuring hair growth and, specifically, response to therapy. Image analysis technology for a precise quantification of hair density in patients with hair loss disorders yielded consistent and valid results in the past using a variety of techniques, including analysis of digitized images¹², image analysis of individual hairs and skin lesions¹³, and microscopic scanning to quantify hair mass in a defined scalp region¹⁴ .

Unfortunately, while the above methodologies have provided improved assessment of efficacy for hair growth-promoting interventions in clinical trials, these methods have not been applied to rodent studies in preclinical investigations. Our goal is to develop a consistent and reliable method to quantify hair loss in mice, which will allow investigators to more accurately assess and compare new therapeutic approaches for various forms of alopecia. We have developed a methodology using equipment readily available in most laboratories which will allow rapid and reliable quantification of hair density in mice with brown or black hair. This methodology has been tested in mouse models of chemotherapy-induced alopecia, alopecia areata, and alopecia from waxing. A detailed protocol is presented for performing these measurements, including validation data from C57BL/6 and C3H/HeJ strains of mice. As this technique relies on detecting light absorption from pigments in the hair shaft, it cannot be used to detect hair growth in white mice or albino mice.

Protocol

Ethics Statement: All studies involving animals must be approved by the IACUC of the institution (for the data which follows, protocols were approved by the Montefiore IACUC, protocol #11-6-240 and #13-7-100). Animals are provided light anesthesia as indicated for the sole purpose of keeping them still during photography, there are no painful procedures required for this protocol.

1. Acquisition of Photographs

1. Set focus and field of view for gel imager using paper with printed text. Verify uniformity of light source across photographed region. To help ensure uniformity, use a gel imager with a built in light source for reflective photography. Do not use a transillumination light source, as this would create a silhouette of the animal which is unsuitable for further grayscale analysis.
   NOTE: This will place the mouse slightly out of focus, which provides optical averaging across the region of interest (ROI) and will help reduce quantal errors for very small (i.e., <10 pixel) regions of interest. Larger regions of interest will not be affected by this minor focusing adjustment, nor will they be affected by differences in the size of the animal.
2. Anesthetize animals using ketamine (100 mg/ml)/xylazine (20 mg/ml) (2:1), as this provides rapid anesthesia effect and rapid recovery and is optimal for photographing multiple animals.
   NOTE: Anesthesia is confirmed when the animal is still enough to permit photography. As animals typically recover from this light anesthesia within 10-15 min, the veterinarian has not recommended use of eye ointments.
3. Place anesthetized animals on gel imager in vertical alignment (as close to parallel as possible).
   1. For dorsal photographs, place the animals in prone position with limbs extended.
   2. For ventral photographs, place animals in the supine position, taking care that the animals are not laterally rotated.
4. Place grayscale standard in photographed region.
5. Close access door. Important: Ambient light can introduce variations in exposure.
6. Set F-stop to an exposure which places the region of interest within the linear range of acquisition (reading F-stop).
   NOTE: Most systems will display where the image is saturated.
7. Take photograph.
8. Change F-stop to another exposure setting which places the region of interest within the linear range by increasing or decreasing the F-stop by 1, such that both the standard and the region of interest remain in a linear range of acquisition (reference F-stop).
9. Take photograph.
10. Once photography is completed, place animals on a warming table and monitor until they can maintain sternal recumbency. Return animals to their cages. Return the group of animals to the vivarium once all animals are fully recovered.

2. Quantification of Light Absorption

1. Mark regions of interest on the images of the animals using provided software for gel imager.
   1. For whole animal dorsal view, use a rectangular or oval image extending from the upper limbs to the lower limbs, extending laterally as much as possible such that no part of the box extends beyond the back of the animal as shown in Figure 1A.
      NOTE: One could also mark the area of interest using a free-hand drawing tool.
   2. For whole animal ventral view, use 2 rectangles: one covering the pelvic region and one covering the chest area as shown in Figure 1B.
   3. For a smaller region of interest, i.e., location of drug administration, mark as appropriate.
2. Mark region(s) of interest on grayscale absorption standard.
3. Record absorption from each of the marked regions of interest.

3. Analysis

Absorption levels obtained for the regions of interest may need to be normalized to the background standard for comparison between photographs. The relationship of exposure to absorption is log-log (i.e., the relationship is linear between log(exposure) and log(absorption)). Using this relationship, the absorption of the region of interest can be normalized to a standard background value, which will allow absorptions to be compared directly between different photographs, including those taken at different time points (i.e., serial measurements within a study protocol). The procedure for performing these corrections is detailed in optional steps 3.1 and 3.2.

1. (Optional) Plot curve of log (exposure) vs. log(absorption) using values obtained from grayscale absorption standard.
2. (Optional) Adjust for variations in the standard of each photograph as follows:
   1. Select F-stop for reading (as determined in step 1.6) and F-stop for reference (as determined in step 1.8).
   2. Calculate average absorption of the standard in all photographs at the reading F-stop. This average is the Reference Standard Value (RSV).
   3. Calculate difference in absorption between the reading and reference F-stop settings in each photograph for the standard (delta-S) and for each defined ROI (delta-ROI-1, delta-ROI-2 …..delta-ROI-X)
   4. Calculate the corrected absorption for each ROI as follows: Corrected absorption (ROI-X) = Absorption (ROI-X) at reading F-stop + (RSV - absorption of standard at reading F-stop) * (delta-ROI-X /delta-S)
3. Compile experimental data and perform data analysis using standard statistical techniques for continuous variables (i.e., ANOVA, T-test).

Results

This technique was validated using C3H/HeJ engrafted mice, the mouse model of alopecia areata¹⁶. These animals develop global hair loss which progresses gradually over time. However, the hair loss occurs in patches, and may vary from one mouse to the next, introducing considerable variability and hampering qualitative assessments. This model provided an opportunity to test correlations between measurements of optical density at different locations on the mouse as an assessment of validity.

Discussion

In this report a detailed description is provided of a new technique for quantifying hair loss in rodents. This technique utilizes a gel imager for image acquisition and analysis, equipment which is readily available in most laboratories. The measurements have been shown to be robust to minor variations in technique (Figures 4-5), and are well-correlated to the degree of visual hair loss in treatment models (Figure 6). These techniques have been successfully employed in ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
KODAK Gel Logic 100 Imaging System	Eastman Kodak Company, Rochester, NY, USA		Gel imager for obtaining and analyzing photographs, must have built in light source for reflective photography.
The Kodak/Tiffen Q-13 Gray Scale	Imatest		Greyscale standard
C57BL/6J Mice	Jackson Laboratories, Bar Harbor, Maine		Mice for representative study
C3H/HeJ engrafted mouse	Jackson Laboratories, Bar Harbor, Maine		Mice for representative study