Isolation and Characterization of Antler-Derived Multipotent Stem Cells

Abstract

Recent studies have reported that stem cells can be isolated from various tissues such as bone marrow, fatty tissue, umbilical cord blood, Wharton's jelly, and placenta. These types of stem cell studies have also arisen in veterinary medicine. Deer antlers show a seasonal regrowth of tissue, an unusual feature in mammals. Antler tissue therefore might offer a source of stem cells. To explore the possibility of stem cell populations within deer antlers, we isolated and successfully cultured antler-derived multipotent stem cells (MSCs). Antler MSCs were maintained in a growth medium, and the proliferation potential was measured via an assay called the cumulative population doubling level. Immunophenotyping and immunostaining revealed the intrinsic characteristic stem cell markers of antler MSCs. To confirm the ability to differentiate, we conducted osteogenic, adipogenic, and chondrogenic induction under the respective differentiation conditions. We discovered that antler MSCs have the ability to differentiate into multiple lineages. In conclusion, our results show that deer antler tissue may contain MSCs and therefore may be a potential source for veterinary regenerative therapeutics.

Introduction

Recently, stem cell research has been an interesting and ever-progressing field within cell biology. Research in stem cells is conducted with animal as well as human stem cells. Deer antlers constitute a rare example of a completely regenerating organ in mammals. Because of the distinctive regenerative ability of deer antlers, there have been many studies to determine the characteristics of these antlers. Nevertheless, little is known about antlers. It has been suggested that the unusual feature of an organ that undergoes annual regrowth might indicate the presence of a stem cell population. Some studies have reported that cells isolated and cultured from deer antlers may comprise a population of stem cells (5,20,31). However, these studies did not fully characterize the stem cells, and the number of available studies is not sufficient. Therefore, we sought to further elucidate the characteristics of antler stem cells.

Stem cell studies have been conducted using various animal tissues as well as human tissues. In small animals, many reports have shown that stem cells can be isolated and characterized from bone marrow, fatty tissue, umbilical cord matrix, umbilical cord blood, and amniotic membrane (11,14,30,33). Likewise, some reports also confirmed the possibility of stem cell populations from diverse tissues in large animals, such as horses and pigs (16,21,23,29). In deer, we thought that the antler might be a potential stem cell source due to its peculiar regenerative ability.

In this study, we successfully isolated and characterized antler tissue-derived multipotent stem cells (MSCs) and confirmed that the isolated cells are self-renewing and can differentiate into multiple different lineages. We also optimized the cell culture conditions for the antler MSCs by assaying cell viability against various cell culture medium conditions, including the concentration of fetal bovine serum (FBS) and growth factors. Using optimized culture conditions, antler MSCs displayed vigorous cell proliferation and normal chromosome numbers and morphology. For these results, several essential experiments were conducted such as long-term cell population assay, expression pattern of stem cell marker, cell surface markers by fluorescence-activated cell sorting (FACS) analysis, in-depth multilineage differentiation assay, and karyotype assay. In this respect, this study shows obvious differences from other previous studies (5,9,31). Finally, we suggest that deer antler tissue might be a valuable source of stem cells and could potentially be a useful source of regenerative therapeutics in veterinary science.

Materials and Methods

Tissue Collection From Antlers

All of the described experiments were approved by and followed the regulations of the Institute of Laboratory Animals Resources (SNU-120202-1, Seoul National University, Korea). All safety compliances were strictly observed and adhered to as outlined in the Policy and Regulation for Care and Use of Laboratory Animals (Institute of Laboratory Animal Resources, Seoul National University). The antler samples (n = 3) were obtained from different specimens. Therefore, there was no sample pooling in our study. All samples (n = 3) were obtained from 2-year-old farmed Sika deer males as previously described (18). Briefly, the deer were anesthetized with Hellabrunn mixture [125 mg xylazine (Rompun; Bayer Korea Co., Seoul, Korea) and 100 mg ketamine (Ketara; Yuhan Co., Seoul, Korea)], and then the antlers were shaved and cleaned to conduct biopsy. The antler tip was taken for a biopsy, which was conducted with sterilized surgical instruments. The section used in the biopsy was approximately less than 10 mm diameter and was collected from the specimens after approximately 45–60 days of the initiation of the antler. A crescent-shaped incision was made in the skin using a surgical blade, and then tissue collection was completed using a sterile biopsy needle (BD Biosciences, San Jose, CA, USA). To prevent contamination and damage to the tissues, all samples were immediately moved into sterile tubes (BD Biosciences), stored at 4°C and transported to the laboratory as quickly as possible.

Cell Isolation and Culture

Cell isolation and culture were performed as previously described with some modifications (18). In brief, the collected antler tissue was minced with a surgical blade (Swann-Morton, Sheffield, UK) and scissors (Miltex, York, PA, USA) under sterile conditions. The minced tissues were digested with an enzyme solution prepared with collagenase type II (2 mg/ml; Gibco BRL, Grand Island, NY, USA) at 37°C for approximately 3–4 h. The digested tissues were passed through a cell strainer (100-μM mesh size; BD Biosciences) to remove any debris. Then, the samples were washed in phosphate-buffered saline (PBS; Gibco BRL) for two to three times by centrifugation at 350 × g for 5 min. The cell pellet was then seeded in various types of culture media, including Dulbecco's modified Eagle's medium (DMEM), Eagle's minimum essential medium (EMEM), DMEM/F-12, and D-media (formula No. 78-5470EF; all from Gibco BRL). The cells were cultured in a humidified atmosphere containing 5% CO₂. The basal culture medium was changed three times each week, and the cells passaged after reaching 80–90% confluency. To confirm the effect of FBS (Gibco BRL), the doses of 20%, 10%, 5%, 3%, and 1% of FBS were tested via cumulative population doubling level (CPDL) with each medium condition tested. To confirm the effect of growth factors, bFGF (basic fibroblast growth factor, Abcam, Cambridge, MA, USA) and IGF-1 (insulin-like growth factor-1, Abcam) were added to each medium condition at various concentrations (100, 20, 10, and 0 ng/ml) and tested via CPDL.

CPDL Analysis

The estimated growth efficiency and proliferation potential of antler MSCs were determined by CPDL analysis using the formula: CPDL = ln(N_f / N_i)ln2, where N_i is the initial seeding cell number, N_f is the final harvest cell numbers, and ln is the natural log. Cells (5 × 10⁴) were plated in triplicate in a six-well culture plate (Nunc, Rochester, NY, USA) and subcultured 5–7 days later. The final cell numbers were counted, and 5 × 10⁴ cells were replated. To evaluate the cumulated doubling level, the population doubling for each passage was calculated and then added to the population doubling levels of the previous passages.

Karyotype Analysis

To detect any chromosomal abnormalities in the antler MSCs, karyotype analysis was conducted followed by the standard methods of Q band at passage 5. Briefly, antler MSCs were arrested with 0.1 μg/ml colchicine (Sigma-Aldrich, St. Louis, MO, USA) for 20 min. The cells were then suspended in a hypotonic solution (0.075 M KCl; Sigma-Aldrich) and incubated for 20 min at 37°C. The cells were pelleted at 335 × g (1,000 rpm) for 10 min and fixed by washing three times in methanol/glacial acetic acid (3:1; Sigma-Aldrich). Chromosome spreads were obtained by pipetting suspension drops onto clean glass and allowing them to air dry. Cells undergoing metaphase were captured with a CCD camera (Olympus, Tokyo, Japan), chromosomes were counted, and the banding pattern was analyzed.

Flow Cytometry Analysis of Surface Antigen Expression

Antler MSCs were analyzed by flow cytometry on a FACS Aria analyzer with FACS DIVA software (BD Biosciences). The cells were detached with a solution containing 0.25% Trypsin-EDTA (Gibco BRL) and then washed three times with PBS. After washing, the cells were fixed by ice-cold ethanol for 30 min. After three washes with PBS, the cells were stained with fluorescein isothiocyanate (FITC)-conjugated anti-human cluster of differentiation 31 (CD31), CD45, CD62P, CD105, CD133, human leukocyte antigen (HLA)-DR, and octamer-binding transcription factor 4 (Oct4; all from BD Biosciences). After three washes with PBS, the cells were resuspended in 500 μl PBS and analyzed with a FACS Aria.

Immunocytochemistry

For immunostaining, 1 × 10⁴ cells were plated in a four-chamber slide (Nunc) of each well. After 1–2 days, the cells were fixed in 4% paraformaldehyde (Sigma-Aldrich) for 20 min and were then permeabilized for 10 min at room temperature in 0.5% Triton X-100 (Sigma-Aldrich) diluted in PBS. After washing three times, the cells were blocked with normal goat serum (NGS; Zymed, San Francisco, CA, USA) overnight at 4°C. Cells were then incubated with primary antibodies overnight at 4°C. After washing three times, the cells were incubated with the secondary antibodies Alexa 488 and 594 (diluted 1:1,000; Molecular Probe, Inc., Eugene, OR, USA) for 1 h. Finally, for nuclear staining, DAPI (4′,6-diamidino-2-phenylindole; 1 mg/ml, Gibco BRL) was diluted 1:100 in PBS and applied to the samples for 15 min. Cell images were captured on a confocal microscope (Eclipse TE2000; Nikon, Tokyo, Japan). The primary antibodies used in this study for immunostaining purposes were mouse anti-ATP-binding cassette, subfamily G, member 2 (Abcg2; Santa Cruz Biotechnology, Santa Cruz, CA, USA), rabbit anti-nestin (Santa Cruz Biotechnology), rabbit anti-C-myc (Abcam), mouse anti-CD90 (BD Biosciences), rabbit anti-Cripto-1 (Abcam), mouse anti-CD9 (Abcam), rabbit anti-Nanog (Abcam), mouse anti-stage-specific embryonic antigen 1 (SSEA1; Abcam), rabbit anti-sex-determining region Y box 2 (Sox2; Chemicon, Temecula, CA, USA), mouse anti-l-alkaline phosphatase (AP; Santa Cruz Biotechnology), mouse anti-SSEA4 (Abcam), and rabbit anti-Oct4 (Abcam).

Osteogenic Differentiation

Osteogenic differentiation was conducted with osteogenic medium, composed of 50 μM ascorbic acid 2-phosphate, 100 nM dexamethasone, 10 mM β-glycerophosphate (all Sigma-Aldrich), and 10% FBS in DMEM. Additionally, DMEM with 10% FBS medium was used as a control. The medium in each experimental group was changed two times each week. After 3 weeks, Alizarin Red S staining was conducted to detect calcium deposition. For Alizarin Red S staining, the cells were rinsed with PBS and fixed with 70% ice-cold ethanol for 1 h at 4°C. Following three washes using distilled water, the samples were stained with 40 mM Alizarin Red S (pH 4.2; Sigma-Aldrich) for 10 min at room temperature. Nonspecific dye was removed by washing with distilled water five times. Alizarin Red S was solubilized in 100 mM cetylpyridinium chloride (Sigma-Aldrich) for 1 h. The release of the solubilized Alizarin Red S was measured at 570 nm using a spectrophotometer (24). For Von Kossa staining, the cells were soaked in 5% silver nitrate (Sigma-Aldrich) for 1 h with concomitant exposure to ultraviolet light (Philips, Eindhoven, Holland), followed by 5% sodium thiosulfate (Sigma-Aldrich) for 5 min, and then counterstained with Nuclear Red Stain (Sigma-Aldrich) for 5 min.

Adipogenic Differentiation

The adipogenic differentiation medium was composed of 500 μM 3-isobutyl-1-metyl-xanthine (IBMX), 1 μM dexamethasone, 60 μM indomethacin, and 5 μg/ml insulin (all Sigma-Aldrich). Once the cells reached 80–90% confluency, they were treated with the adipogenic differentiation medium for 3 weeks. To evaluate adipogenic differentiation, Oil Red O staining was conducted. The cells were fixed with 10% formalin (Sigma-Aldrich) for at least 1 h and rinsed with 60% isopropanol (Sigma-Aldrich) prior to incubation in freshly diluted Oil Red O (Sigma-Aldrich) for 10 min. Bound stain was solubilized using 100% isopropanol, and then the released stain was measured at 500 nm using a spectrophotometer (Beckman Coulter, Fullerton, CA, USA).

Chondrogenic Differentiation

Chondrogenic differentiation was performed as previously described with some modifications (36). Briefly, 2 × 10⁵ cells were added to a 15-ml polypropylene tube (BD Biosciences) and centrifuged to make a pellet. Chondrogenesis differentiation medium (Lonza, Basel, Switzerland) was added to the cell pellet at 37°C in a 5% CO₂ incubator for 3 weeks. The differentiation medium was replaced three times per week. After 3 weeks, the round pellets were embedded in paraffin and cut into 3-mm sections. For histological evaluation, the sections were stained with toluidine blue (Sigma-Aldrich). A 3-mm cell pellet slice mounted on a slide was deparaffinized and hydrated with distilled water. For toluidine blue staining, the slide was immersed in toluidine blue working solution for 1 min. Excess unbound stain was removed using several washes of distilled water. The slide was quickly dehydrated with sequential washes of 95% and absolute alcohol. For Alcian blue-PAS staining, the slide was stained with Alcian blue, 0.5% periodic acid, and Schiff's reagent (all Sigma-Aldrich). The slides were cleared in xylene (Sigma-Aldrich) and covered with Canada balsam (Sigma-Aldrich) and a coverslip (Marienfeld, Lauda-Konigshofen, Germany).

Statistical Analysis

The data were analyzed by two-way ANOVA and unpaired t test using the Bonferroni–Dunn method to correct for multiple comparisons to determine the p value. The results are expressed as the mean plus or minus the standard error. Statistically significant data are indicated by asterisks (***p < 0.001, **p < 0.01).

Results

Isolation and Culture of Antler MSCs

To establish cell culture conditions, we isolated and cultured antler cells with four different types of culture media: DMEM, MEM, D-Media, and DMEM/F-12. The different media were assayed for optimal cell growth and evaluated using CPDL analysis. The cultured cells were adherent to the plastic culture surfaces, and the morphology of the cells was very similar among all culture medium types tested (Fig. 1A–D). To detect the proliferation effect among the culture medium types, we performed CPDL and measured the cell population from passage 3 to passage 23. During this time, there was no significant difference among the different types of culture media on CPDL, and stable increases in the cell growth curves were observed under all conditions (Fig. 1E). There were no morphological changes observed in the cells until passage 23 (data not shown). Although not statistically significant, the cells cultured with DMEM displayed more vigorous cell proliferation compared with EMEM, D-Media, and DMEM/F-12 media (Fig. 1E). DMEM was henceforth used to culture antler MSCs in following experiments.

Figure 1.

Primary culture and effect of dose-dependent serum on the antler MSCs. (A–D) Phase contrast images of antler mesenchymal stem cells (MSCs) at passage 1 (P1). The cells were cultured with four different types of media (with 10% FBS): (A) Dulbecco's modified Eagle's medium (DMEM), (B) Eagle's minimum essential medium (EMEM), (C) D-Media, and (D) DMEM/F-12. Scale bar: 50 μm. (E) Cell growth curve of antler MSCs with four different types of media. CPDL was evaluated from P3 to P23.

Proliferation Assay with Serum and Growth Factors

The ability of cells to proliferate is generally associated with the dose of serum and growth factors in the media. In the case of serum conditions, many cultured cell lines are cultured with 10% serum (11,19,23,25,33,36). Accordingly, 10% serum was applied to the antler cells in the experiments described here (5,20,31). However, to determine the optimal serum conditions for cell growth, antler MSCs were also cultured with various serum concentrations. The cell growth curves were calculated via CPDL with 1%, 3%, 5%, 10%, or 20% FBS in the media. CPDL was conducted from passage 3 to passage 7. There proved to be a statistically significant difference when comparing the growth curves for those cells grown with 1% FBS at passages 3 and 4 with all other serum concentrations. After passage 5, both the 1% and 3% FBS conditions showed statistically significant differences in proliferation (Fig. 2A). However, we confirmed that FBS concentrations greater than 5% had no statistically significant effect on cell growth curves. The differences among cell populations according to serum concentrations were confirmed by phase contrast microscopy (Fig. 2B–F). We observed a marked difference in the cell population in the 1% and 3% FBS conditions compared with other serum concentrations. Therefore, we determined that the medium containing 5% serum was the best cell culture condition for deer antler MSCs.

Figure 2.

Proliferation effect of serum concentrations and growth factors on antler MSCs. (A) Cell growth curves of antler MSCs with various concentrations of FBS. CPDL was evaluated from passage 3 to passage 7 with five different concentrations of FBS (1%, 3%, 5%, 10%, and 20%). The experiments were conducted using DMEM. Compared with 5%, 10%, and 20% FBS, concentrations of either 1% or 3% FBS displayed a significant difference on CPDL. All analyses were performed in triplicate; the means ± standard deviations are plotted (**p < 0.01, ***p < 0.001). (B–F) Phase contrast images of antler MSCs with five different concentrations of FBS at passage 7. (B) 1% FBS, (C) 3% FBS, (D) 5% FBS, (E) 10% FBS, and (F) 20% FBS of antler MSCs. Scale bar: 100 μm. (G, H) Cell growth curves of antler MSCs with growth factors: (G) bFGF and (H) IGF-1. No significant difference was observed (W/O, without).

To determine the role of growth factors in optimizing the growth medium conditions, our previous study (24) showed that that bFGF and IGF had an important role in cell proliferation with human MSCs. Therefore, we evaluated the effect of bFGF and IGF-1 in the cell proliferation of antler MSCs. The growth factors bFGF and IGF-1 were separately assayed for cell proliferation from passage 3 to passage 7. Four different concentrations were tested: 100, 20, 10, and 0 ng/ml. We observed no significant differences among the resulting cell proliferation curves within both the bFGF and IGF-1 groups (Fig. 2G and H). We also confirmed that there were no differences in cell morphology among the various conditions of growth factors (data not shown). For these reasons, the basal culture condition of antler MSCs was maintained in DMEM (supplemented with 5% FBS) without any growth factors.

Analysis of Immunophenotype and Karyotype

To determine whether they showed the characteristics of stem cells, we conducted flow cytometry analysis of antler MSCs at passage 5 and used five CD markers (CD31, CD45, CD62p, CD105, and CD133), HLA-DR, and a stem cell marker (Oct4). In the FACS analysis, antler MSCs had positive expression patterns of CD105 (79%) and Oct4 (69.6%). CD105, also called SH2, is a well-known MSC marker, and Oct4 is also a specific stem cell marker (Fig. 3A). The other markers (CD31, CD45, CD62p, CD133, and HLA-DR) displayed nearly no expression. Therefore, the antler MSCs exhibited a stem cell phenotype.

Figure 3.

Flow cytometry and karyotype analysis. (A) FACS analysis was performed at passage 5. Values show the intensity of the indicated antigen. (B) Karyotype of antler MSCs at passage 5, showing a euploid number of chromosomes. CD31, cluster of differentiation 31; HLA-DR, human leukocyte antigen- DR; Oct4, octamer-binding transcription factor 4.

To confirm a normal chromosome count using our culturing procedure, karyotype analysis was conducted with antler MSCs. The cells were confirmed to contain 66 chromosomes, which is the expected normal karyotype of deer cells (Fig. 3B).

Immunostaining of Antler MSCs with Various Stem Cell Markers

To investigate the expression of stem cell markers in antler MSCs, we conducted immunostaining with several stem cell markers. In total, 12 stem cell markers were used for staining. Among the 12 markers tested, seven displayed positive expression. Furthermore, in the FACS analysis (Fig. 3A), Oct4 showed the positive expression pattern, which was highly stained in the nucleus (Fig. 4). Cells also maintained highly positive expression patterns of CD9, C-myc, and Sox2. These genes are known markers of embryonic stem cells (22) and germline stem cells (15). We also observed slightly positive expression for nestin, SSEA4, and Cripto-1. Nestin is a known marker for neural stem cells and follicle stem cells (1,34), while SSEA4 and Cripto-1 are embryonic stem cell markers (26). However, the cells showed negative expression patterns in Abcg2, CD90, AP, Nanog, and SSEA1, all of which are also known stem cell markers.

Figure 4.

Immunostaining of antler stem cells. Antler MSCs were immunostained with ATP-binding cassette, subfamily G, member 2 (Abcg2), nestin, CD9, Cripto-1, C-myc, CD90, sex-determining region Y box 2 (Sox2), alkaline phosphatase (AP), Nanog, stage-specific embryonic antigen 1 (SSEA1), SSEA4, and Oct4. DAPI (4′,6-diamidino-2-phenylindole) staining was used for nuclei detection. Scale bar: 100 μm.

Induction of Differentiation

To evaluate the differentiation ability of antler MSCs, three different types of differentiation assays were conducted. To test for osteogenic differentiation, the cells were cultured in osteogenic induction medium. The differentiation medium was replaced every 3 days for 3 weeks. To detect calcium deposition, Alizarin Red S and Von Kossa staining were conducted in both the osteogenic differentiation condition and the basal culture condition (Fig. 5). The basal culture condition was used as a control and was composed of DMEM with 5% FBS. Under the differentiation conditions, the cells were strongly positively stained with Alizarin Red S (Fig. 5A, B) and Von Kossa staining (Fig. 5E, G). Not surprisingly, under the basal culture condition, the cells were not stained with Alizarin Red S (Fig. 5C, D) and Von Kossa staining (Fig. 5F, H). To quantify the extent of cellular differentiation, Alizarin Red S-stained cells were eluted with 100 mM cetylpyridinium chloride, and the absorbance was measured. Compared with the basal culture condition, osteogenic differentiated cells showed approximately 18-fold greater values of Alizarin Red S staining (Fig. 5I).

Figure 5.

Osteogenic differentiation. (A–H) Osteogenic differentiation of antler MSCs. (A–D) Alizarin Red S and Von Kossa staining after 3 weeks of osteogenic induction and control condition. Osteogenic differentiated cells (A, B, E, and G) were grown in osteogenic induction medium. Negative control cells (C, D, F, and H) were grown in DMEM with 10% FBS. No staining with Alizarin Red S and Von Kossa staining was observed under the control conditions. Differentiated cells stained strongly with Alizarin Red S (A and B) and Von Kossa (E and G). Scale bar: 100 μm. (I) For quantification, Alizarin Red S-stained cells were solubilized with 100 mM cetylpyridinium chloride, and the absorbance was measured spectrophotometrically at 570 nm for 0.5 s. Compared with the negative control, differentiated cells showed approximately 18-fold greater absorbance values. All analyses were performed in triplicate, and the means ± standard deviations are plotted (***p < 0.001).

To confirm chondrogenic differentiation abilities, antler MSCs were seeded into 15-ml polypropylene tubes and centrifuged to form a pellet. The pellet was incubated at 37°C in a 5% CO₂ incubator with the chondrogenic induction medium. The medium was changed once every 3 days for 3 weeks. After differentiation, we examined the pellet formation in the bottom of the polypropylene tube. The pellet appeared to be ovoid and opaque (Fig. 6A, B). The pellet was then fixed and stained with toluidine blue (Fig. 6C) and Alcian blue-PAS (Fig. 6D), both of which exhibited positive staining patterns.

Figure 6.

Chondrogenic differentiation. (A–D) Chondrogenic differentiation of antler MSCs. (A) Pellet formation was observed at the bottom of the tube. The white arrow indicates the pellet. (B) The shape of chondrogenic pellet. (C) Toluidine blue and (D) Alcian blue-periodic acid and Schiff's reagent (PAS) staining for chondrogenic pellets. The stained tissue displayed a typical cartilaginous tissue phenotype. Scale bar: 100 μm.

To confirm adipogenic differentiation abilities, antler MSCs were treated with adipogenic induction medium that was changed once every 3 days for 3 weeks. After differentiation, the cells were stained with Oil Red O to detect fatty droplets. Under differentiation conditions, fatty droplets were observed via positive Oil Red O staining (Fig. 7A, B). The white arrows indicate the stained fatty droplets in Oil Red O (Fig. 7B). The basal culture medium was used as a negative control, which ultimately displayed negative staining (Fig. 7C, D). To quantify adipogenic differentiation, stained cells were eluted with 100% isopropanol, and the absorbance was measured. The differentiated cells showed approximately ninefold higher absorbance than that of the negative control (Fig. 7E).

Figure 7.

Adipogenic differentiation. (A–D) Adipogenic differentiation of antler MSCs. Oil Red O staining was conducted after 3 weeks of adipogenic induction. (A, B) Adipogenic differentiated cells were grown in adipogenic induction medium. Fatty droplets were strongly stained with Oil Red O. (C, D) Negative controls showed no staining with Oil Red O. White arrows indicate stained fat droplets. Scale bar: 100 μm. (E) For quantification, stained cells were solubilized with 100% isopropanol, and the absorbance was measured spectrophotometrically at 500 nm for 0.5 s. Compared with the negative control, differentiated cells showed approximately ninefold greater absorbance values. All analyses were performed in triplicate, and the means ± standard deviations are plotted (***p < 0.001).

Discussion

For decades, stem cell research has been an increasingly interesting issue with implications in human regenerative medicine. As stem cell research has developed, many tissues have been discovered to possess a stem cell population. In veterinary medicine, researchers have also sought stem cell populations in animals. Indeed, studies have not only successfully isolated and characterized stem cells from several animal tissues (21,23,29,30,33), but they have also identified forms of animal stem cell therapies for use in regenerative medicine (6,16,25).

In mammals, after individuals reach adulthood, the ability to regenerate damaged organs is commonly limited. However, deer antlers are very peculiar organs in that they are lost and regrown annually (17,28). Research has been conducted for decades, both in vivo and in vitro, to understand the mechanism and regulation of antler growth (2,3,32). Several studies previously reported the identification of possible stem cells from deer antlers (18,20,31). Though more detailed experiments are needed to establish antler stem cells, Rolf et al. (31) indicated that deer antlers contain a stem cell population via positive STRO-1 staining. These cells were able to differentiate into osteogenic, chondrogenic, and adipogenic lineages under the inductive media. However, to strictly confirm the identification of a stem cell population, it is essential to conduct a cell population assay until late passages, as well as a karyotype assay, immunophenotyping, stem cell marker expression patterns, and an in-depth multilineage differentiation assay with isolated cells.

In this study, we report the isolation and characterization of an antler-derived MSC population. Antler MSCs were cultured from antler tissues and tested to optimize the culture condition using various cultured medium formulations and concentrations of FBS and growth factors. We observed that the most abundant cell growth was measured using DMEM compared with EMEM, D-media, and DMEM/F-12. Regardless of culture media, antler MSCs proved to be adherent to plastic surfaces without undergoing any morphological changes. Many previous studies of antler cells suggested the use of DMEM for culture media, although there was no reported assay for optimal culture conditions (18,20,31). In testing for the ability of serum and growth factors to aid in culture viability, various concentrations were used in establishing culture conditions via a CPDL assay. Many studies reported that stem cells are usually cultured with 10% FBS or fetal calf serum (FCS). However, we observed that antler MSCs had a very similar cell growth population and morphology at concentrations greater than 5% FBS (Fig. 2).

Several studies have examined the effect of growth factors on antler growth, both in vivo and in vitro, using such factors as sex hormones and IGF-1 (3,32). However, their effects remain controversial, and more evidence is needed to establish the effect of growth factors on deer antler growth (3,18,27,32). Our previous research showed the effect of bFGF and IGF-1 in cell proliferation and self-renewal with human umbilical cord-derived MSCs (24). Both bFGF and IGF-1 are well-known and important regulators of self-renewal and differentiation in stem cell biology. Our results indicated that antler MSCs seemed entirely uninfluenced by bFGF and IGF-1 in regard to their cell growth population. We observed very similar growth proliferation profiles and cell morphology. Antler MSCs were confirmed to have vigorous cell proliferation capabilities without added growth factors. Therefore, we established and confirmed the optimal antler MSC culture conditions, which gave rise to robust cell proliferation.

The analysis of FACS showed the antler MSCs have the characteristic of immunophenotypes known as the stem cells (10). Additionally, antler MSCs have positive expression patterns of CD105 (79%) and Oct4 (69.6%). CD105, also known as endoglin, is used as a marker for MSCs, and Oct4 is also expressed in embryonic stem cells and adult stem cells (9). Negative expression profiles were confirmed using CD31 (0%), CD45 (0.1%), CD62p (0.1%), CD133 (0.3%), and HLA-DR (0.3%), which display a distinctive patterns of stem cell markers (10). CD133 is known as a surface marker for hematopoietic stem cells (35). In the immunostaining of antler MSCs, several positive expression patterns were observed with stem cell markers. However, unexpectedly, some stem cell markers, such as Abcg2, CD90, AP, Nanog, and SSEA1, were not expressed. Whether these results should be considered as typical characteristics of antler MSCs or as an artifact of the current limited supply of stem cell markers is unclear. We used the human host antibodies for FACS and immunostaining because of a lack of deer-specific antibodies. Due to the limitation of existing animal-specific antibodies, many stem cell studies have conducted all experiments with various host antibodies in the field of veterinary stem cell biology (20,23,29,36). To circumvent this limitation in the future, the development of deer host-specific antibodies is essential to confirm the identification of antler MSCs.

The distinctive features of stem cells are the ability to self-renew and the capability to differentiate. In the present study, antler MSCs were subjected to a differentiation assay for osteogenic, adipogenic, and chondrogenic lineages under culture conditions specific to each lineage. We confirmed the multilineage differentiation ability of antler MSCs.

In this study, we successfully isolated antler MSCs and confirmed the possibility of culturing them through several dozen passages. Stem cell characteristics were identified using FACS, immunostaining, and differentiation assays. Furthermore, antler MSCs showed no alterations in chromosome counts via karyotype analysis. Clinical cell therapy applications require a large number of stem cells, and to this end, antler MSCs proliferation was stable until passage 23. These results suggest that the isolated cells can potentially be used for cell therapy in veterinary medicine.

Until now, the clinical cell therapy applications of antler cells with large or wild animals have largely remained nonexistent. The existence of wild animals, including deer, is important for the ecological system. However, because of the growth of human populations and urban development, the risk of injury to wild animals has increased (4,8). Especially, deer–vehicle collision is frequent, which can induce damages such as bone fracture, blooding, and skin injury (13). In this aspect, antler MSCs can be used for therapeutic application for wild animal treatments.

However, some studies of horse stem cells have reported the clinical application of equine tissue-derived stem cells under various conditions (7,12,19), which remains a current issue in regenerative medicine.

In conclusion, we suggest that antler MSCs could be a potential source of stem cells and therefore could be useful in tissue engineering.