Abstract
The effects of mono-(2-ethylhexyl) phthalate (MEHP), an active metabolite of di-(2-ethylhexyl) phthalate (DEHP), on prepubertal guinea pig testes in vitro were investigated. The testes of 35-day-old guinea pigs were surgically excised. They were seeded in a defined medium containing antibiotics and administered MEHP at concentrations of 1, 10, and 100 nmol/ml, respectively. The control groups were administered a similar volume of corn oil vehicle. The tissues were incubated for 3, 6, and 9 h. The specimens were collected at 3, 6, and 9 h after treatment. They were fixed in 4%paraformaldehyde or 5% glutaraldehyde. For quantitation of the apoptotic spermatogenic cells, the terminal dUTP nick end-labeling (TUNEL) staining was performed by light microscopy. Detachment and displacement of spermatogenic cells, thin seminiferous epithelia, and Sertoli cell vacuolization were observed. Maximal testicular damage was recognized at 100 nmol/ml 9 h after MEHP treatment. The percentage (%) of apoptotic spermatogenic cells significantly increased at 3, 6, and 9 h after treatment, compared to the control groups. Because the loss of spermatogenic cells by MEHP treatment varies among species, the present study, using guinea pigs, was designed and conducted to obtain further information.
Mono-(2-ethylhexyl) phthalate (MEHP) is a well-characterized Sertoli cell toxicant, and is an active toxic metabolite of di-(2-ethylhexyl) phthalate (DEHP). DEHP is widely dispersed throughout the environment due to its increased use in the production of plastic products (Giam et al. 1978; Cole et al. 1981; Boekelheide 1993; Giammona et al. 2002). Sertoli cells are the direct target cell for MEHP, and the primary consequence of MEHP treatment in rodents showed a large increase in spermatogenic cell apoptosis (Richburg and Boekelheide 1996). Apoptosis of selected spermatogenic cells normally occurs in the testis, and is essential for the maintenance of spermatogenesis (Roosen-Runge 1973; Print and Loveland 2000). The increased of the incidence of spermatogenic cell apoptosis is often observed as a result of various forms of physically or chemically injured testes (Richburg 2000). Testicular damage is a relatively common finding in toxicology studies (Gray and Gangolli 1986). Recent studies revealed that MEHP induced testicular injury in mice and rats, and caused the impairment of male fertility (Iona et al. 2002; Suominen et al. 2003). Marked species differences in sensitivity to phthalate esters have been reported (Gray et al. 1982; Richburg et al. 2000). Even within one species (mouse), distinguishable differences were noticed among different strains (Koji 2001). Moreover, in most of the studies on phthalate esters, rats and mice have been used as an animal model. Regarding the MEHP effects on guinea pig testes, no work has been carried out during the last two decades. Therefore, the present study was conducted in vitro to investigate the effects of MEHP on testes in male guinea pigs during the prepubertal stage of growth.
MATERIALS AND METHODS
Animals
Twelve prepubertal male guinea pigs (28-day-old) were purchased from Charles River Laboratories, Japan (Crj). The body weight of the animals ranged from 280 to 300 g. They were acclimatized for 1 week prior to use. The guinea pigs were housed three per one plastic cage, maintained on a 12 h light/dark cycle at a constant temperature (70°F ±2°F) and humidity (35% to 70%), and provided water and rodents Pellets (Oriental Yeast, Japan) ad libitum. The handling of all animals was maintained in accordance with the National Institute of Health (NIH) Guidelines for the Care and Use of Laboratory Animals. The use of animals for this experiment was approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Tokyo, Japan.
CHEMICAL/DRUG
MEHP was purchased from Tokyo Kasei Kogyo, Japan (Figure 1). MEHP was dissolved in an appropriate volume of corn oil vehicle prior to administration. An apoptosis detection kit was purchased from Takara Chemical, Japan.
Experimental Design
The testicular tissue cultures obtained from guinea pigs were divided into three groups. Each group consisted of four animals. Within each group, they were divided into two subgroups. Group 1: corn oil (vehicle) control. Group 2: MEHP treatment at the concentration of 1, 10, and 100 nmol/ml, respectively.
Tissue Processing and Treatment
The animals were anesthetized with sodium pentobarbital (Nimbutal; 50 mg/kg body weight), and subsequently sacrificed by decapitation. The prepubic region of the animals were cleaned and disinfected with ethanol. The testes were surgically removed, and decapsulated in phosphate-buffered saline (PBS). The seminiferous tubules were treated with collagenase, washed three to five times with PBS, and placed in Dulbecco’s minimum essential medium (DMEM; Sigma Chemical, USA). After washing three to five times with DMEM, the tissues were carefully placed on DMEM with antibiotics containing 200-unit penicillin 100 IU ml−1, streptomycin 100 IU ml−1, gentamycin 40 mg ml−1, and fungizone 0.5 μg ml−1, and supplimented with 10% fetal bovine serum (Cansera International, Canada). Next, the seminiferous tubules were placed on 24 multiwell plate (Sumilon), and/or Millipore filter with reconstituted basement membrane extracts (Hadley et al. 1988). Simultaneously, they were also placed on a 100-cm2 culture dish (Fulcon). The culture was then treated with MEHP at the concentrations of 1, 10, and 100 nmol/ml, respectively. The control group received a similar volume of corn oil vehicle. Both MEHP and vehicle-treated samples were incubated at 32.5°C in a humidified atmosphere containing 95% air and 5% CO2 and was continued for 9 h.
Histopathological Preparation
At 3, 6, and 9 h after treatment, the tissue were collected, washed gently with PBS, and immediately fixed in 4% paraformaldehyde. The tissues were dehydrated in a series of graded ethanol, cleared in xylene, and embedded in paraffin. The paraffin blocks were cut at 5 μm in thickness. The sections were then stained with Meyer’s hematoxylin and eosin and/or periodic acid–Schiff (PAS)-hematoxylin staining for general histopathological examination.
Transmission Electron Microscopy
For transmission electron microscopy, the specimens were fixed in 5% glutaraldehyde–0.05 M cacodylate buffer (pH, 7.4) at 4°C for 2.5 h and then washed three or five times with the same buffer. They were postfixed in 1% osmium tetroxide (OsO4) for 1 h, dehydrated through a graded series of ethanol, and embedded in Araldite. Thin sections were cut at approximately 1 μm in thickness, stained with 1% toluidine blue, and observed using light microscopy. Ultrathin sections were cut, and stained with uranyl acetate and lead citrate, and examined with a JEM-1200 EX transmission electron microscope at 80 kV.
In Situ TUNEL Staining and Quantitation
In order to assess quantitatively the incidence of apoptotic spermatogenic cells after MEHP treatment, the in situ terminal deoxynucleotidyl transferase-mediated digoxigenin- dUTP nick end-labeling (TUNEL) technique was conducted in prepubertal male guinea pigs in vitro. This method provides a very sensitive indicator to detect breaks in DNA and often labels cell DNA early in the process of apoptosis before morphologic evidence of cell death can be seen. TUNEL was performed by using the in situ Apoptosis Detection Kit (Takara, Japan). Briefly, the tissue sections were deparaffinized and digested with 20 μg/ml proteinase K (Takara, Japan) at 37°C for 15 min. After washing three to five times with PBS, they were treated with terminal deoxynucleotidyl transferase (TdT) enzyme and Labeling Safe Buffer (Takara, Japan). The TdT reaction was conducted at 37°C for 90 min. After further washing three to five times with PBS, they were incubated with horseradish peroxidase (HRP)-goat anti-biotin (Takara, Japan) at 37°C for 30 min, reincubated with 0.05% diaminobenzidine (DAB) and 0.02% H2O2 for visualization of HRP sites, counterstained with methyl green, and observed using a light microscope. Tunel-positive spermatogenic cells in the seminiferous tubules were counted and tabulated.
Statistical Analysis
Only intact and round seminiferous tubules were picked up. The number of apoptotic spermatogenic cells was counted in 100 randomly selected round seminiferous tubules from each of three different groups. The data, calculated as a percentage of total, were expressed as the mean ± SEM. Significance between groups (p < .05) was determined by single-factor analysis of variance(ANOVA) with a Fisher’s least significant differences test comparison using Stat View software (Abacus Concepts, Berkeley, CA, USA).
RESULTS
Histopathological Findings
Light Microscopy
Histopathology was evaluated using cross-sections of seminiferous tubules stained with PAS-hematoxylin. At 3 h after MEHP treatment, the detachment and displacement of spermatogenic cells were frequently encountered compared to the control (Figure 2A, B ). From 6 to 12 h after MEHP treatment, progressive detachment and displacement of spermatogenic cells, thin seminiferous epithelia, disappearance of defined tubular lumen, and Sertoli cell vacuolization were distinctly recognized (Figure 2B ).
Transmission Electron Microscopy
The seminiferous tubules obtained from the petridish and/or Millipore filter membrane were examined using transmission electron microscopy. Most of the cells in the control group were normal in structure (Figure 3A ). Degenerative Sertoli cells with several large vacuoles were frequently encountered in the treated groups (Figure 3B ). No morphological difference was observed between the samples from the Petri dish and those from Millipore filter membrane.
Quantification of TUNEL-Positive Spermatogenic Cells
The percentage of apoptotic spermatogenic cells significantly increased throughout the treatment period. Furthermore, the maximal increase was observed at 100 nmol/ml in all treated groups, compared to the control groups (p < .05). At 9 h after treatment, the number of apoptotic spermatogenic cells increased about fourfold at 100 nmol/ml of MEHP (Figures 4A, B, C, D and 5).
DISCUSSION
Tissue degeneration is a common feature of developmental processes in vertebrates during embryonic and postnatal periods. At that time, excess or redundant cells are eventually removed by cell death. In adult life, cell death is also thought to play an important role in the regulation of tissue mass under both physiological and pathological conditions. Moreover, under experimentally manipulated conditions, such as cryptorchidism, ischemia-reperfusion, hypophysectomy, treatment with gonadotropin-releasing hormone (GnRH) antagonists, heating, and drug treatment, spermatogenic cell degeneration occurs during various stages of spermatogenesis (Sinha-Hikim et al. 1995; Cai 1997; Nandi et al. 1999).
MEHP is the active metabolite of phthalic acid ester DEHP. A number of studies revealed that phthalic acid esters reduced fertility and induced testicular atrophy in laboratory animals (Gray and Butterworth 1980; Gray and Beamand 1984; Albro 1987; Kasahara et al. 2002).
In most of the in vivo studies on phthalate esters, some major changes such as detachment and sloughing of spermatogenic cells, atrophy of seminiferous tubules, and loss of testicular, seminal vesicle, and prostate weights have been reported (Thomas and Thomas 1984; Gray and Gangolli 1986; Dalgaard et al. 2001). The present study revealed detachment of spermatogenic cells, disappearance of normal tubular lumen, thin tubular epithelia, and vacuolization of Sertoli cells. These findings are similar to those reported in other laboratory animals (Gray et al. 1982; Richburg and Boekelheide 1996).
Both in vivo and in vitro experiments have demonstrated that the Sertoli cell is the primary site of phthalate-induced testicular toxicity (Thomas et al. 1987; Li et al. 2000; Richburg et al. 2000). Sertoli cell alterations include vacuolization, inhibition of seminiferous tubule fluid formation, loss of mitochondrial succinate dehydrogenase activity, inhibition of transferrin secretion, and abnormalities in the glycolytic pathway. In addition, MEHP has been reported to specifically disrupt follicle stimulating hormone–induced increases in cyclic AMP in cultured rat Sertoli cells. These Sertoli cell alterations lead to a progressive degeneration of spermatocytes and spermatids, ultimately sloughed off into the tubular lumen (Boekelhiede 1993; Grasso et al. 1993). The experiments using Sertolispermatogenic cells coculture system showed that MEHP induced a concentration-dependent increase in the rate of spermatogenic cell detachment (Creasy et al. 1988). Our present findings are similar to the observations reported in rats (Creasy et al. 1988).
In order to study the consequences of phthalate-induced Sertoli cell toxicity, we investigated the influence of MEHP on spermatogenic cell apoptosis in young guinea pig testes. Apoptotic spermatogenic cells were observed in controls at 3 h. The incidence of spermatogenic cell apoptosis significantly increased at 3, 6, and 9 h after MEHP treatment. The maximal percentage of apoptotic spermatogenic cells was observed at the concentration of 100 nmol/ml MEHP 9 h after treatment. TUNEL-positive spermatogenic cells in the present study significantly increased (approximately fourfold) over baseline at 100 nmol/ml MEHP 9 h after treatment. This finding was also consistent with the findings in wild-type mice (Giammona et al. 2002). They reported that the incidence of TUNEL-positive spermatogenic cells in B6 wild-type and gld mice testes significantly increased over baseline at 6, 12, and 48 h after MEHP treatment, but declined to return to baseline until 96 h.
Spermatogenic cell apoptosis from birth to adult life is essential for spermatogenesis. Physiological apoptosis continuously occurs in rats to limit the size of the spermatogenic cell population to numbers that can be adequately supported (Richburg et al. 1999). Richburg and Boekelheide (1996) demonstrated a progressive increase in spermatogenic cell apoptosis from 6 to 12 h after MEHP treatment in rats, but a decrease at 3 h after treatment. The increase in the number of apoptotic spermatogenic cells reported in different laboratory animals after phthalate esters treatment supports our findings.
Although most of the studies on phthalate esters induced testicular toxicity were limited to rats and mice, marked species and strain differences in sensitivity were observed. Comparative studies with di-n-butylphthalate (DBP), a phthalic acid ester, revealed that rats and guinea pigs were severely affected, mice were less affected, and hamsters showed no testicular damage (Gray et al. 1982; Giammona et al. 2002).
Many hypotheses exist in the literature regarding the mechanism of phthalate esters–induced testicular toxicity. Apoptosis has been described to occur in cells after growth factor or hormone deficiency (Bardon et al. 1987; Kyprianou and Isaacs 1988; Tapanainen et al. 1993). Therefore, in an analogous manner, altered secretion of Sertoli cell factors as a result of MEHP treatment may trigger spermatogenic cell apoptosis.
MEHP could also be envisioned to interact directly with the apoptotic signaling pathway. The cellular mechanisms and signal transduction events which regulate apoptosis are poorly defined and currently the subject of intensive research. Moreover, the trigger for apoptosis might be integrated with other normal cellular signal transduction systems (Corcoran et al. 1994). MEHP disrupts the follicle stimulating hormone-linked signal transduction pathway in primary Sertoli cell cultures by a mechanism located in the cell membrane (Grasso et al. 1993). These findings suggest that MEHP interacts directly with components of the apoptotic signal transduction pathway. It may be concluded that MEHP induces testicular toxicity in guinea pigs in vitro, and the testicular tissue culture system is useful for screening the testicular toxicity of phthalate esters and other environmental persistent reproductive toxicants.
However, the exact mechanism of MEHP-induced testicular toxicity still remains unclear. Afurther study is now underway in our laboratory to clarify the mechanism of action of MEHP-induced testicular dysfunction in guinea pig.
Footnotes
Figures
This work was carried out in the Department of Veterinary Anatomy, The University of Tokyo, Japan. The authors thank Mr. I. Tsugiyama for his technical assistance and excellent animal care during this research. Cordial thanks are also extended to Dr. Mitsuharu Matsumoto (Kagoshima University, Japan) for his continuous encouragement, valuable suggestion, and critical reviewing of this manuscript. This work was supported in part by a grant-in-aid from the Japan Society for Promotion of Sciences (JSPS), and in part by a grant-in-aid from the Ministry of Health, Labour and Welfare, Japan.