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BOR - Papers in Press, published online ahead of print September 15, 2004.
Biol Reprod 2004, 10.1095/biolreprod.104.031278
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BIOLOGY OF REPRODUCTION 72, 86–96 (2005)
DOI: 10.1095/biolreprod.104.031278
© 2005 by the Society for the Study of Reproduction, Inc.

Effects of Acid Sphingomyelinase Deficiency on Male Germ Cell Development and Programmed Cell Death1

Marjut Otala2,3, Markku O. Pentikäinen4, Tiina Matikainen3,5, Laura Suomalainen3, Jukka K. Hakala4, Gloria I. Perez5, Mikko Tenhunen6, Krista Erkkilä3, Petri Kovanen4, Martti Parvinen7, and Leo Dunkel3,8

Program for Developmental and Reproductive Biology,3 Biomedicum Helsinki and Hospital for Children and Adolescents, University of Helsinki, FIN-00029 HUS, Helsinki, Finland Wihuri Research Institute,4 FIN-00140, Helsinki, Finland Vincent Center for Reproductive Biology,5 Vincent Obstetrics and Gynecology Service, Massachusetts General Hospital, and Department of Obstetrics, Gynecology, and Reproductive Biology, Harvard Medical School, Boston, Massachusetts 02114 Department of Oncology,6 Helsinki University Hospital, FIN-00029, Helsinki, Finland Department of Anatomy,7 University of Turku, FIN-20520 Turku, Finland Department of Pediatrics,8 University of Kuopio, FIN-70211, Kuopio, Finland


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Deficiency of acid sphingomyelinase (ASM), an enzyme responsible for producing a pro-apoptotic second messenger ceramide, has previously been shown to promote the survival of fetal mouse oocytes in vivo and to protect oocytes from chemotherapy-induced apoptosis in vitro. Here we investigated the effects of ASM deficiency on testicular germ cell development and on the ability of germ cells to undergo apoptosis. At the age of 20 weeks, ASM knock-out (ASMKO) sperm concentrations were comparable with wild-type (WT) sperm concentrations, whereas sperm motility was seriously affected. ASMKO testes contained significantly elevated levels of sphingomyelin at the age of 8 weeks as detected by high-performance, thin-layer chromatography. Electron microscopy revealed that the testes started to accumulate pathological vesicles in Sertoli cells and in the interstitium at the age of 21 days. Irradiation of WT and ASMKO mice did not elevate intratesticular ceramide levels at 16 h after irradiation. In situ end labeling of apoptotic cells also showed a similar degree of cell death in both groups. After a 21-day recovery period, the numbers of primary spermatocytes and spermatogonia at G2 as well as spermatids were essentially the same in the WT and ASMKO testes, as detected by flow cytometry. In serum-free cultures both ASMKO and WT germ cells showed a significant increase in the level of ceramide, as well as massive apoptosis. In conclusion, ASM is required for maintenance of normal sphingomyelin levels in the testis and for normal sperm motility, but not for testicular ceramide production or for the ability of the germ cells to undergo apoptosis.

apoptosis, spermatogenesis, stress, testis


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Acid sphingomyelinase (ASM) is a lysosomal enzyme that belongs to a family of sphingomyelinases that catabolize membrane sphingomyelin (SM) to ceramide and phosphorylcholine [1, 2]. Deficient ASM activity is the cause of Niemann-Pick disease (NPD) in which SM degradation is impaired, leading to elevated levels of SM and cholesterol [3]. Cells received from NPD patients as well as mice lacking a functional ASM gene have become useful tools when investigating ceramide-mediated signal transduction pathways.

Acid sphingomyelinase knock-out (ASMKO) mice appear normal at birth, begin to express symptoms for neurologic disease including ataxia and mild tremors at about 2–4 mo of age, and die at the age of 6–8 mo [4, 5]. Their fecundity is affected since reduced fertility can be observed before the onset of behavioral deficits [6]. Therefore, lipid accumulation may cause severe physiological defects in the testis and harm germ cell development or function. A lack of ASM may also lead to defective ceramide production. Ceramide, which is a lipid second messenger, has been found to be pro-apoptotic in several biological systems [7, 8]. Compelling evidence has been presented on sphingomyelinase activation and generation of ceramide as a response to ionizing radiation [9, 10]. Ceramide can be further metabolized to sphingosine, which again can be phosphorylated into sphingosine-1-phosphate (S1P). S1P counterbalances the effects of ceramide and thereby inhibits apoptosis in several cell types, including the male germ cells [11–14]. Thus, lack of ASM in the testis may lead to impaired production of ceramide and deficient germ cell apoptosis.

ASM deficiency renders many cell types completely or partially resistant to apoptosis [15–17]. It plays an important role in female germ cell death. In a recent study it was found that apoptosis was diminished in ASMKO mouse fetal ovaries, resulting in an increased number of primordial follicles and ovarian hyperplasia in neonatal mice [13]. ASMKO mouse oocytes were also able to resist chemotherapy-induced apoptosis in vitro. Most importantly, resistance to irradiation-induced apoptosis was achieved in the wild-type (WT) oocytes in vivo when they were protected by S1P [13].

Both female and male germ cells are susceptible to irradiation, and infertility may follow cancer treatments. We have previously found that irradiation damages the early developmental stages of mouse spermatogonia [18]. In the male, S1P is able to partially protect testicular germ cells of WT C57BL/6 mice against radiation-induced apoptosis in vivo [18] and to suppress human male germ cell apoptosis in vitro by 30% [14]. Male testicular tissue has also been shown to produce increased amounts of ceramide when germ cell apoptosis is induced in vitro [14]. Therefore, a pathway involving ceramide may be functional in male germ cell apoptosis. To study the SM pathway more closely in the male gonad, we investigated the role of ASM deficiency in the mouse testis 1) in vivo in germ cell development and physiological apoptosis at different developmental ages, 2) in vivo in irradiation-induced apoptosis, and 3) in vitro in serum deprivation–induced apoptosis.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Mice

The mouse colony was established from heterozygous (±) breeding pairs, which had a mixed 129/SV-C57BL/6 genetic background. The generation of the mice has originally been published by Horinouchi et al. [5]. A polymerase chain reaction genotyping method was used to maintain the breeding colony and to choose suitable mice for the experiments. WT and ASM homozygous-null (ASMKO) male litter mates were used for the experiments. Testis histology was analyzed at the age of 7 days (juvenile, no active spermatogenesis, main cell types spermatogonia, immature Sertoli cells, and immature Leydig cells); 21 days (juvenile, onset of spermatogenesis, appearance of different developmental stages of germ cells, mature Sertoli cells, and Leydig cells starting to proliferate) [19]; 8 wk (sexually mature, fully functional spermatogenesis); and 20 wk (neurological symptoms) males. In the in vivo and in vitro experiments, 8- to 10-wk-old young adult mice were used. The animals were kept in a 12-h light/dark cycle, and they received food and water ad libitum. All animal protocols were reviewed and approved by the Massachusetts General Hospital Institutional Animal Care and Use Committee, Boston, and by the Institutional Animal Care and Use Committee of the Wihuri Research Institute, Helsinki, Finland, and were performed in strict accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals.

Sperm Analysis

WT and ASMKO mice were killed at the age of 20 wk, and sperm were collected from cauda epididymis and capacitated in vitro for 2 h in human tubal fluid medium (Specialty Media, Phillisburg, NJ) supplemented with 0.5% BSA under oil. All samples were analyzed for sperm concentration and motion parameters by the HTM-IVOS semen analyzer (Version 10HTM-IVOS; Hamilton-Thorn Company, Beverly, MA). Setting parameters and the definition of measured sperm motion parameters were established by the Hamilton-Thorn Company (frames acquired = 30; frame rate = 60 Hz; straightness [STR] threshold = 80%; duration of the tracking time = 0.38 seconds). To measure both sperm concentration and motility, aliquots of semen samples (5 µl) were placed into a prewarmed (37°C) Makler counting chamber (Sefi-Medical Instruments, Haifa, Israel). A minimum of 200 sperm from at least four different fields was analyzed from each specimen.

Measurement of Ceramide and Sphingomyelin Levels

Levels of ceramide and SM were analyzed as described previously in detail [14]. Briefly, tissue from mouse testes ({approx}100 mg) was homogenized in 1030 µl of homogenization buffer (1% Triton X-100, 150 mM NaCl, 10 mM Tris pH 7.4, 1 mM EDTA, 0.2 mM PMSF, 0.5% NP-40, 1 µg/ml leupeptin). After centrifugation the supernatants were collected for protein concentration measurements by the DC protein assay (Bio-Rad Laboratories, Hercules, CA). The lipids were extracted by a method modified from Bligh and Dyer [20] as described [14] and analyzed by the high-performance, thin-layer chromatography method using dichloromethane/ methanol/glacial acetic acid (100:2:5, v/v/v) for ceramides and chloroform/ methanol/glacial acetic acid/water (100:60:20:5, v/v/v/v) for SM. The individual lipid classes were visualized and analyzed as described [14]. Ceramide and SM concentrations were normalized to the total cell protein, and the ceramide/SM weight ratio was calculated. The lipid standards used in this analysis were from Sigma (St. Louis, MO).

Electron Microscopy (EM)

Small pieces of testicular tissue were fixed in 2.5% glutaraldehyde in 0.1 mol/L phosphate buffer, dehydrated, and embedded in epoxy resin. The samples were sectioned at 50 nm with an ultramicrotome (Reichert Jung, Vienna, Austria) mounted on nickel grids and stained with uranyl acetate and lead citrate using the Leica EMstain apparatus (Leica, Vienna, Austria) according to the manufacturer's instructions. Observations were made with a JEOL JEM 1200 EX transmission EM (JEOL, Tokyo, Japan).

Irradiation of Mice

The mice received a dose of 0 or 0.5 Gy for the total body at the Department of Oncology, Helsinki University Hospital, Helsinki, Finland. The irradiation was performed as described previously [18] with the Varian Clinac 600C linear accelerator using a 6-MV photon beam, and the dose rate was 2 Gy/min. Sixteen hours after irradiation, one testis was snap-frozen for measurements of ceramide and sphingosine levels, and the other was used for stage-specific squash preparations. Testicular germ cell apoptosis can be detected in the rat 8 h after irradiation, and the highest numbers of apoptotic germ cells are found at approximately 16 h [21]. After 21 days each testis from another group of mice was weighed, and DNA flow cytometric analyses were performed. The time required for spermatogonia to develop into spermatocytes and spermatids is 21 days. In the case that mouse spermatogonia are damaged, as occurs normally as a consequence of irradiation, the number of spermatocytes and spermatids was reduced after the period of 21 days [18].

Nonradioactive In Situ End Labeling of DNA (ISEL)

Short segments of seminiferous tubules (approximately 1 mm in length) were transferred in 10–20 µl of PBS on silane-coated microscope slides. A cover slip was placed on each tubulus segment to squash the tubulus and produce a monolayer of cells around both ends of the segments. The samples were fixed and dehydrated as previously described [22–24]. DNA 3' end-labeling was performed as described [18, 25]. In brief, after rehydration the preparations were permeabilized by microwaving for 5 min in citrate buffer (10 mmol/L citrate, pH 6.0); incubated in terminal transferase reaction buffer (1 mol/L potassium cacodylate, 125 mmol/L Tris-HCl, and 1.25 mg/ml BSA, pH 6.6); and 3' end-labeled with Dig-dd-UTP (Roche) for 1 h at 37°C by the terminal transferase (Tdt; Roche) reaction. Dig-dd-UTP was located with a peroxidase-conjugated antidigoxigenin antibody (Anti-Digoxigenin-POD; Roche), and the antibody was detected with 0.05% diaminobenzidine substrate (Sigma). The samples were counterstained lightly with hematoxylin and then dehydrated and mounted.

DNA Flow Cytometry

The testes were decapsulated in phosphate-buffered saline on a Petri dish, and the seminiferous tubules were teased apart and prepared into 1 mm long segments under a transillumination stereomicroscope. The different developmental stages of the epithelial cycle are distinguishable by their varying capacity to absorb light, so that the more a seminiferous tubule segment has advanced-stage spermatids, the darker it seems under a preparation microscope. DNA flow cytometric analyses were performed as described previously, with some modifications [18, 26, 27]. The stage-specific 1 mm long single segments of seminiferous tubules were treated with a detergent (0.3% Nonidet P-40; BDH, Poole, England) in PBS containing 0.2% bovine serum albumin (Sigma) and ribonuclease A (5 µg/ml; Sigma) for 10–15 min at 4°C, after which a 10-sec vortex and incubation for 15 min at 37°C followed. Propidium iodide (25 µg/ml; Sigma) and diluted fluorescent particle solution (10 µl; 475 beads/µl; TrueCount Beads, Becton Dickinson, Mountain View, CA) were added to the samples. The samples were filtered and then analyzed by a FACSCalibur flow cytometer (Becton Dickinson) at 488 nm. A total of 5000 fluorescent impulses were counted excluding the beads and debris. CellQuest Pro software (Becton Dickinson) was used for calculating the number of nuclei in each peak of the DNA histograms. The number of cells in each population was converted to absolute numbers by using a multiplication factor that was derived from the ratio of added to counted numbers of standard beads. In the obtained histograms, step 1–12 spermatids form the 1C (haploid chromosome number) peak, and the 4C peak is comprised mainly of primary spermatocytes, excluding preleptotene spermatocytes [26]. We have previously shown that the radiation-induced reduction mainly takes place in mouse 1C (spermatids) and 4C (spermatocytes) cell populations, and it can be seen by flow cytometry 21 days after irradiation [18].

Tissue Culture

For the in vitro experiments, testicular tissue was cultured as described, with some modifications [14, 24, 25, 28]. The testes were decapsulated in phosphate-buffered saline on a Petri dish, and the seminiferous tubules were microdissected in tissue culture medium (nutrient mixture Ham F-10; Life Technologies, Paisley, UK) supplemented with 0.1% bovine albumin (Sigma) and 10 µg/ml gentamicin (Life Technologies). Segments of seminiferous tubules were cultured instead of isolated germ cells to maintain as physiological an environment as possible for the germ cells. The segments of seminiferous tubules were incubated at 34°C under serum-free conditions in a humified atmosphere containing 5% CO2. We first made a time series for which WT tissue was cultured for 0, 24, 48, 72, or 96 h to examine the rate of apoptosis induction in mouse germ cells under serum-free conditions. After culture the tissue was snap-frozen in liquid nitrogen and stored in –80°C for Southern blot analysis of fragmented DNA. We then chose 24- and 48-h cultures for the following in vitro experiment.

Segments of seminiferous tubules of WT and ASMKO mice were cultured for 0 (freshly snap-frozen), 24, or 48 h in the serum-free culture conditions. After culture the tubules were either frozen in liquid nitrogen or squashed stage specifically on microscope slides under cover slips as explained below. Southern blot analyses and in situ end labeling (ISEL) were performed, and ceramide and SM levels were measured.

Southern Blot Analysis of Apoptotic DNA Fragmentation

DNA was extracted from frozen tissue using an Apoptotic DNA Ladder Kit (Roche Molecular Biochemicals, Mannheim, Germany) as described previously in detail [14, 25]. Briefly, DNA was quantified and labeled with digoxigenin-dideoxy-UTP (Dig-dd-UTP; Roche) by the terminal transferase (Roche) reaction. The samples were then electrophoresed, blotted, and crosslinked onto nylon membranes. The attached 3' end-labeled DNA fragments were localized with alkaline phosphatase– conjugated antidigoxigenin antibody (Anti-Digoxigenin-AP, alkaline phosphatase conjugates; Roche), and the bound antibody was detected by the chemiluminescence reaction (Roche). X-ray films exposed to the luminescent membranes were digitized by a tabletop scanner (Hewlett Packard Scanjet 6300C) and analyzed with Gel plot 2 macro for Scion Image beta 4.0.2 (Scion Corporation, Frederick, MD) analysis software. The low molecular weight DNA fragments (<1.3 kb) were quantitated against the positive control sample provided by the manufacturer (Roche).

Data Presentation and Statistical Methods

Comparisons between WT and ASMKO lipid concentrations, testicular weights, DNA ladders, and sperm counts were performed by two-tailed Student t-test, and stage-specific comparisons were done by Mann-Whitney U test. Multiple comparisons between the lipids and testicular weights were done by ANOVA followed by two-tailed t-tests. P-values < 0.05 were considered significant. Results are expressed as mean ± SEM.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Physiological Germ Cell Death

Sperm concentrations were not statistically different between 20-wk-old WT and ASMKO mice (Table 1). However, the percentage of motile spermatozoa was severely reduced in the ASMKO mice (P < 0.001), and the defect in ASMKO sperm motility seemed even greater when progressive motility was measured (P < 0.001).


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  		TABLE 1. Effects of ASM deficiency on sperm concentration and motility at the age of 20 wk.*

ASMKO mouse testes had significantly elevated SM contents (27.68 ± 3.92 µg/mg protein) as compared with WT mice (6.47 ± 0.56 µg/mg protein) at the age of 8 wk (P < 0.001), but the ceramide levels were at a comparable level (Fig. 1). No significant differences could be detected between WT and ASMKO testicular weights at the ages of 7 days, 21 days, 8 wk, and 20 wk, although there was a tendency for slightly heavier testes in the sexually mature ASMKO mice (8 and 20 wks; Fig. 2).



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  		FIG. 1. Basal sphingomyelin and ceramide levels in WT and ASMKO testes at sexual maturity. Each bar represents the mean lipid concentrations obtained from independent experiments (number of experiments indicated by n) ± SEM. ***P < 0.001



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  		FIG. 2. Testicular weights of WT and ASMKO mice at the ages of 7 days, 21 days, 8 wk, and 20 wk. Each bar represents the mean of three to eight testes (number of testes indicated by n) from different animals ± SEM

Low molecular weight DNA fragmentation analysis (Southern blotting) revealed increased apoptosis in the ASMKO testes at the age of 21 days as compared with the ages of 7 days, 8 wk, and 20 wk (P < 0.05), indicating that the well-established physiological wave of germ cell apoptosis required for the development of functional spermatogenesis in rodents [29–34] also takes place in the ASMKO testes (Fig. 3A). Little germ cell apoptosis could be detected at the age of 7 days, and at the age of 8 wk the amount of apoptosis had again declined to a basal level (comparable with the level of apoptosis in WT mice of the same age; data not shown) and remained at this basal level also at the age of 20 wk. By histological examination of electron micrographs we were able to detect pathological vesicles from the age of 21 days onward in Sertoli cell cytoplasms and in the interstitium (Fig. 3B). Such vesicles were not observed in the WT tissue. At this age large numbers of apoptotic spermatocytes, typical for the normal physiological apoptotic wave in the rodent testis, also were found in the ASMKO testes (Fig. 3B). The testes from 8-wk-old sexually mature ASMKO mice withheld pathological vesicles within the seminiferous tubules in the Sertoli cells and outside the tubules in the interstitium (Fig. 3C). By the age of 20 wk the size of the vesicles had grown and they had become more numerous in the Sertoli cells as well as in the interstitium (Fig. 3D). In none of the ages were we able to observe pathological accumulation of vesicles in the spermatogonia, spermatocytes, or in round spermatids. The neck regions of elongated spermatids were also devoid of pathological vesicles.



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  		FIG. 3. Physiological apoptosis in ASMKO testicular tissue and testis morphology. A) Quantification of low molecular weight DNA. Each bar represents a mean of four independent experiments ± SEM. *P < 0.05. B) Pathological droplets (black arrows) in ASMKO Sertoli cell cytoplasm at the age of 21 days. The droplets were observed in the Sertoli cell cytoplasm between the germ cells and also close to Sertoli cell nuclei (black double arrow). Several apoptotic spermatocytes with condensing chromatin (open arrows) as well as some terminal-stage apoptotic cells (double open arrow) were observed. C) More pathological accumulation could be detected in the testes from 8-wk-old mice (black arrows) and it was located in the Sertoli cell cytoplasm (Sertoli cell nucleus indicated with black double arrow) as well as in the interstitium. D) Abundant droplets (black arrows) were found in the testes of 20-wk-old ASMKO mice. Original magnifications B–D and insets x2500

Germ Cell Susceptibility to Radiation-induced Cell Death

Sixteen hours after irradiation, SM levels in the WT mouse testes were low, whereas they were markedly elevated in the ASMKO testes (P < 0.001; Fig. 4). However, the level of SM in the irradiated WT and ASMKO mice did not differ from that of the corresponding nonirradiated mice (compare with Fig. 1). Surprisingly, ceramide levels were comparable with those in the nonirradiated testes in both groups (Fig. 4; compare with Fig. 1), and the level of ceramide was similar between the WT and ASMKO testes.



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  		FIG. 4. Sphingomyelin and ceramide levels in WT and ASMKO testes 16 h after irradiation. Each bar represents the mean lipid concentrations obtained from independent experiments (number of experiments indicated by n) ± SEM. ***P < 0.001

No statistically significant differences in testicular weights at 16 h after irradiation could be observed between the WT and ASMKO mice or in relation to mean testicular weights before irradiation (0 h; Fig. 5). Twenty-one days after irradiation, weights of the WT testes had decreased by approximately 20% as compared with the nonirradiated testes (P < 0.01), whereas no significant reduction in testicular weights could be observed in the ASMKO mice. Consequently, the ASMKO testes were heavier than the WT testes 21 days after irradiation (P < 0.05).



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  		FIG. 5. Testicular weights 21 days after irradiation. Each value represents the mean of 3–20 testes (number of testes indicated by n) from different animals ± SEM. **P < 0.01; *P < 0.05

Spermatogonia was the main cell type that died after irradiation, although apoptotic spermatocytes and occasional spermatids were also found (Fig. 6A). The level of apoptosis in stage-specific squash preparations for which ISEL of DNA fragmentation had been performed was similar between the WT and ASMKO mice in all of the stages II–V, VII–VIII, and IX–XI (Fig. 6B). In stages II–V and IX–XI, the mean number of apoptotic cells ranged between 20 and 30 apoptotic cells per 1 mm segment of the seminiferous tubulus. In stages VII–VIII, four or three apoptotic cells could be found on average in the WT and ASMKO tubulus segments, respectively.



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  		FIG. 6. Germ cell apoptosis 16 h after irradiation. A) Apoptotic germ cells in squash preparations from WT and ASMKO mice. Pictures from stages IX–XI are shown. PSC, Pachytene spermatocyte; SG, spermatogonium. Original magnification, upper panels x200 and lower panels x400. B) Number of ISEL-positive germ cells in stages II–V, VII–VIII, and IX–XI 16 h after irradiation. Each bar represents the mean value of apoptotic cells in three segments per stage per animal from three animals ± SEM

The long-term tolerance of the ASMKO mouse germ cells to irradiation, as compared with the WT mouse germ cell response, was measured by counting the number of spermatids (1C) and spermatocytes (4C) by flow cytometry in stages II–V, VII–VIII, and IX–XI 21 days after irradiation. No differences in cell numbers between the two groups in any of the stages investigated could be detected, although a statistically nonsignificant trend toward higher numbers of remaining cells in the ASMKO mouse testes was observed (Fig. 7).



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  		FIG. 7. Cell numbers 21 days after irradiation in 1C (spermatids) and 4C (spermatocytes) cell populations as detected by flow cytometry. Each value represents the mean number of cells in three testes from different animals ± SEM

Germ Cell Susceptibility to Tissue Culture–induced Cell Death

In our preliminary experiment we cultured WT mouse testis tissue under serum-free conditions for up to 96 h, after which we performed Southern blotting of apoptotic DNA. Apoptosis induction could be seen after culture for 24 h, and the ladder pattern became more prominent at 48 h. After a 72-h culture, the ladders started to become unclear. The pattern changed into a mere smear as necrosis took over after 96 h of culture (data not shown). We therefore chose to culture testis tissue from WT and ASMKO mice in vitro for 24 and 48 h for further experiments.

We first investigated the amounts of SM and ceramide in the tissue cultures. The level of SM remained much lower in the WT testes than in the ASMKO testes at all time points (0, 24, and 48 h) investigated (P < 0.001; Fig. 8). The level of ceramide increased significantly in both groups after culture for 24 and 48 h. After culture for 24 h, the level of ceramide had increased by approximately 76% (P < 0.05) in the WT tissue and by approximately 73% (P < 0.05) in the ASMKO tissue. After culture for 48 h the increase was 76% in the WT tissue (P < 0.05) and 77% in the ASMKO tissue (P < 0.01) as compared with the uncultured tissue (Fig. 8). The level of ceramide was comparable in the WT and ASMKO mice at all time points investigated. Moreover, in both groups, apoptosis increased markedly during the 48-h cultures (P < 0.001), but there were no differences in the amount of cell death between the WT and ASMKO germ cells in any of the time points investigated (Fig. 9).



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  		FIG. 8. Sphingomyelin and ceramide levels in WT and ASMKO testes after 24 and 48 h of culture. Each bar represents a mean of independent experiments (number of experiments indicated by n) ± SEM. *P < 0.05; **P <0.01; ***P < 0.001



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  		FIG. 9. Germ cell apoptosis in segments of seminiferous tubules incubated under serum-free conditions. A) Radiograph of tissue cultured for 0, 24, and 48 h. B) Low molecular weight DNA (<1.3 kb) quantification. Each bar represents a mean of four independent experiments ± SEM

To more closely examine germ cell apoptosis in the cultures, we used the ISEL technique to count apoptotic cells in stage-specific squash preparations. Although apoptotic cells of all cell types were identified, in vitro culture mainly seemed to affect spermatocytes and early spermatids and to a much lesser extent spermatogonia and late spermatids (Fig. 10A). We found clusters of high numbers of ISEL-positive cells as well as individual dying cells among large areas of healthy tissue. There was much variation in the number of apoptotic cells between the tubulus segments. At quantification, cells in the very early stages of apoptosis were not counted. No statistically significant differences could be found between the WT and ASMKO mice in the 0, 24, or 48 h time points in any of the stages, II–V, VII– VIII, or IX–XI (Fig. 10B). At 24 h in stages II–V and VII– VIII, there was a tendency toward greater numbers of apoptotic cells in the ASMKO tubulus segments, but this failed to reach statistical significance. Accordingly, when apoptotic DNA fragmentation in the cultured tissue was measured by Southern blotting (Fig. 9B), no statistical differences were found between the genotypes. At 0 h both WT and ASMKO testes had an average of four apoptotic cells per 1-mm tubulus segment in stages II–V and one apoptotic cell per 1-mm segment in stages VII–VIII. Stages IX–XI contained more apoptotic cells, the average number being approximately 16 in the WT and 19 in the ASMKO tubulus segments. In the cultured tissue several apoptotic cells could be found in all stages.



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  		FIG. 10. Germ cell apoptosis in vitro. A) Apoptotic cells in cultured WT and ASMKO testis tissue. PSC, Pachytene spermatocyte; ST, spermatid. Original magnification x400. B) Number of ISEL-positive germ cells in uncultured tissue (0 h) and in tissue cultured for 24 and 48 h. Each bar represents the mean of apoptotic cells in different tubulus segments (number of tubulus segments indicated by n) ± SEM


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study we investigated the effects of ASM deficiency on testicular germ cell development and apoptosis. To find out whether ASM deficiency disrupts sperm production, we first investigated ASMKO sperm quality. We found that sperm motility in the ASMKO mice was impaired, but that the sperm concentrations were not statistically different between WT and ASMKO mice. This suggests that control of germ cell numbers cannot be severely disturbed in the ASMKO testes.

Our result that ASMKO sperm motility is defective at the age of 5 mo is in accordance with the results of Butler et al. [6]. They found that in ASMKO sperm, the rapid motility was reduced and greater numbers of static sperm were observed than in WT sperm. The etiology behind the poor sperm quality was found to result from membrane lipid accumulation and a subsequent regulatory volume decrease defect in the developing ASMKO sperm [6]. Pathological lipid accumulation in sperm plasma membrane causes bending of the sperm to relieve increased surface area. Mutant sperm are nevertheless able to regain normal morphology after incubation in mild detergent, which demonstrates that the bending defects are a direct consequence of membrane lipid accumulation. Further, mitochondrial membrane lipid accumulation leads to decreased mitochondrial membrane potential and thus interferes with the energy-generating capacity of the sperm [6]. Both plasma membrane and mitochondrial membrane lipid accumulation most likely impair sperm motility, as normal flagellar motion requires straight sperm tail morphology as well as metabolically active mitochondria. Additionally, lipid accumulation in epididymal epithelial cells may result in impaired physiological and morphological maturation of spermatozoa [6]. Thus, poor sperm motility does not seem to be due to defects in ASM-mediated apoptosis.

After assessment of ASMKO mouse sperm quality, we continued to more closely investigate the role of the SM pathway in males in early stages of germ cell development and in the capacity of ASM-mutant cells to die by apoptosis, especially after pathological insults. Thus, our aim was to investigate the role of ASM deficiency in the mouse testis 1) in vivo in germ cell development and physiological apoptosis at different developmental ages, 2) in vivo in radiation-induced apoptosis, and 3) in vitro in serum deprivation–induced apoptosis. We observed that the ASMKO testes contained elevated SM levels and pathological droplets. Physiological apoptosis at the onset of sexual maturation and at sexual maturity were comparable with the WT mice. The WT and the ASMKO mice also had similar responses to apoptosis induced in vivo by irradiation and in vitro by serum deprivation.

An important feature of the sexually mature ASMKO mice is the strikingly elevated intratesticular SM content, indicating that ASM is essential for efficient SM degradation in the testes. In accordance with Butler et al. [6], we observed pathological vesicles in the ASMKO mouse Sertoli cells and the interstitium, but not in the testicular germ cells, and found that the accumulation advanced with age. We were able to detect incipient accumulation already at the age of 21 days. Lipid-laden foam cells are known to be a part of the etiology of ASM deficiency [6, 35, 36]. The regulatory volume decrease deficits may explain why the developing germ cells seem to lack intracytoplasmic pathological vesicles as observed by EM. The ASMKO Sertoli cells that express gross lipid accumulation may not be able to phagocytose residual bodies from mature spermatids [6], resulting in excess surface areas in the midpiece of the sperm tail in only late spermatids and spermatozoa.

It seems that although pathological vesicle accumulation in the Sertoli cells is an integral part of the ASMKO phenotype, its effects on Sertoli cell function are surprisingly small. Impaired Sertoli cell function would result in reduced sperm concentrations. We found that at the age of 5 mo ASMKO mouse sperm concentrations were not significantly reduced despite substantial pathological accumulation in the Sertoli cell cytoplasms. In accordance, Butler et al. [6] observed equivalent sperm concentrations in WT and ASMKO mice at the age of 6 mo. Thus, the Sertoli cells seem to be able to nurture the germ cells rather well regardless of the pathological intracytoplasmic vesicles.

We then investigated whether physiological germ cell apoptosis is disrupted in the ASMKO mouse testes at different ages. After birth mouse spermatogenesis first begins at the age of 2–5 wk by the sequential appearance of germ cells corresponding to each developmental stage of the seminiferous tubule. Around the age of 3 wk a physiological apoptotic wave eliminates large numbers of spermatogonia and spermatocytes [31]. This apoptosis is most likely required to maintain a proper number of maturing germ cells per Sertoli cell. If the early apoptotic wave is disturbed, as in mice defective of the pro-apoptotic bax gene [30] or mice expressing high levels of the antiapoptotic Bcl-xL or Bcl-2 proteins [31, 37], spermatogenesis is highly abnormal and sterility ensues. However, ASMKO mice did express increased germ cell apoptosis at the age of 3 wk as detected by large numbers of dying spermatocytes and increased low molecular weight DNA fragmentation. We also observed equivalent numbers of apoptotic cells in the sexually mature ASMKO mice as compared with the age-matched WTs.

Male and female germ cells seem to differ in regard to the role of ASM in the fetal gonads. Male germ cells most likely develop normally in the fetal period despite the ASM deficiency, as detected by normal testicular weights, morphology, and amount of apoptosis as at Day 7 postpartum. In striking contrast, ASM is essential for generating death signals in the fetal female germ line. Neonatal (Day 4 postpartum) ASMKO ovaries had an increased number of primordial follicles as well as significant ovarian hyperplasia, indicating defective normal apoptotic deletion of fetal oocytes [13]. In males ASM did not prove important to physiological apoptosis in the sexually mature mice. Accordingly, in the ASMKO females histomorphometric analysis revealed that the differences in the number of oocytes in neonatal ovaries was preserved at the age of 42 days when female mice achieve sexual maturity, indicating that the rate of apoptosis is preserved postnatally. However, ASM no doubt is important to induced germ cell apoptosis in the female [13].

The molecular mechanisms behind the different effects of ASM deficiency on male and female germ cells thus far remain to be elucidated. In the case of oocytes, much evidence has been presented on behalf of ceramide being the central death-inducing agent. Although measuring ceramide levels within oocytes has not been possible, an antibody directed against ceramide has provided evidence that it is generated in oocytes and has a central role in female germ cell apoptosis [38]. Although ceramide may be a key player in ovarian follicle apoptosis in all developmental stages (primordial to antral), its impact on male germ cell apoptosis is much less straightforward. Based on our previous study on the potential of S1P, a rheostat for ceramide—to partially protect the very early, but not the later developmental stages of spermatogonia—it is possible that different germ cell developmental stages may proceed into apoptosis through different routes. Additional evidence is provided by the cell cycle regulator protein p53, which seems to be involved in irradiation-induced death of the differentiating spermatogonia, but not in that of the more radioresistant stem spermatogonia [39]. Thus, the ASM-mediated pathway may merely be an alternative route to apoptosis in the male germ cells, which is very different from the central role of ASM in oocytes.

Responses of WT and ASMKO testes to irradiation were similar. We found no differences in the amount of apoptosis between the two groups at 16 h or at 21 days after irradiation, although at 21 days a trend for slightly larger cell populations was detected in the ASMKO testes, which could in part explain why the ASMKO mice on average seemed to have heavier testes than the WT mice after the recovery period. The observation that ceramide was not elevated in either group at 16 h is interesting. Some reports suggest that ceramide is generated rapidly after irradiation [9, 15, 40–42]. According to other researchers, elevation of ceramide levels may be a late response and ceramide may remain elevated for longer periods after irradiation [43–45]. The response seems cell type–specific. In testicular tissue the injured cells proceed into apoptosis gradually so that in the rat testis the highest numbers of dying cells can be found at approximately 16 h after irradiation and apoptotic cells can still be detected at 42 h [21]. Therefore, one could assume that the production of ceramide would remain elevated for several hours postirradiation. Since ceramide generation precedes the onset of apoptosis [14, 15], the optimum time to measure possible peak ceramide levels would be before the 16 h time point. Also, according to a previous study on mice, the number of abnormal spermatogonia reaches a peak 12 h after irradiation and the total number of spermatogonia then declines [46]. Thus, the time point of 16 h may be too late for the detection of peak ceramide levels. The number of apoptotic cells at any given time point is nevertheless quite minute in relation to the total cell population in the testicular tissue, and therefore the elevation of the ceramide levels may constantly remain undetectably low, although existent.

In our in vitro experiments we cultured WT and ASMKO testicular tissue under serum- and hormone-free conditions. In this in vitro model apoptosis has previously been effectively induced in human seminiferous tubules [24]. In human testicular tissue germ cells in the later phases of differentiation (i.e., spermatocytes and early spermatids) are most sensitive to withdrawal of survival factors [24], and this seemed to be the case also in the cultured mouse tubules. In our 24 and 48 h cultures, the level of ceramide became clearly elevated and massive germ cell apoptosis was observed in both the WT and ASMKO mice. However, there were no differences in the amount of ceramide or germ cell apoptosis between the two genotypes. Interestingly, it seems that the ASMKO germ cells possess a means of producing high levels of ceramide other than through the ASM enzyme. Accordingly, it has been previously shown that WT and ASMKO spermatozoa exhibit a similar degree of SM degradation in situ, and therefore have sphingomyelinase activity distinct from the ASM enzyme [6]. This may be true for the earlier stages of spermatogenic cells as well.

Accumulating evidence has been presented on the involvement of the mitochondria-dependent apoptotic pathway in male germ cell apoptosis induced by stress stimuli, such as deprivation of serum and hormones [47] or heat exposure [48]. An important event in the mitochondrial apoptosis pathway is the permeation of the mitochondrial outer membrane and the release of cytochrome c. Ceramide has been proposed to act on multiple sites in the mitochondria [49]. It has been shown to be able to inhibit the mitochondrial respiratory chain complex III [50] and to form stable pores in the mitochondrial membranes [51]. These pores are large enough to enable the release of cytochrome c [51] that allows the subsequent activation of caspases. Although the mechanism by which ceramide induces cell death in the mitochondrial apoptosis pathway still remains unclear, it most likely involves the release of cytochrome c [49].

Ceramide can be synthesized from SM not only by ASM but also by neutral sphingomyelinase (NSM) [52–55]. In the testis both ASM and NSM activities have been identified [56]. ASMKO mice possess physiologic levels of NSM activity despite the deficiency of the ASM enzyme [5], and therefore NSM may participate in ASMKO germ cell apoptosis. Another pathway to produce ceramide is de novo synthesis in which the enzyme ceramide synthase catalyses the formation of dihydroceramide, which is rapidly oxidized into ceramide [52–55]. Other factors may also contribute to pathological germ cell apoptosis observed in the ASMKO mice. Ceramide can be rapidly converted, for example, into sphingosine, and may therefore not be detectable after irradiation. Sphingosine again has been reported to enhance apoptosis [45]. Additionally, apoptosis pathways other than one dependent on ceramide accumulation may be functional in the male germ cells.

The aims of the present study (i.e., the role of ASM in male germ cell development and apoptosis) have not been investigated before. Our results shed light on the involvement of the SM pathway in male germ cell apoptosis. Taken together, ASM deficiency results in abnormally high intratesticular SM contents and defective sperm motility, but does not affect sperm concentrations. Responses to in vivo– and in vitro–induced apoptosis were quite similar in the WT and the ASMKO mouse testicular tissue. In vivo, no ceramide accumulation could be detected in either of the groups 16 h after irradiation. In vitro, the level of ceramide elevated significantly and to a comparable level in both groups during the 24- and 48-h cultures. These results suggest that the physiological effects of ASM deficiency on the testes are due to pathological lipid accumulation and that the ASM enzyme is not crucial for male germ cell apoptosis.


   ACKNOWLEDGMENTS
 
We thank J. Tilly and R. Kolesnick for providing the mice from the colony of the Vincent Center for Reproductive Biology, Massachusetts General Hospital/Harvard Medical School, Boston, Massachusetts, and J. Pru for assistance with the mice. We are grateful to V. Ahokas for skillful technical assistance. A. Ketola is thanked for help with the statistical analyses.


   FOOTNOTES
 
1 Supported by the Sigrid Jusélius Foundation (Finland), the Cancer Society of Finland, the Nona and Kullervo Väre Foundation (Finland), Helsinki Biomedical Graduate School (Finland), the Finnish Foundation for Pediatric Research, the Finnish Cultural Foundation, and the Research and Science Foundation of Farmos (Finland). Back

2 Correspondence: Marjut Otala, Program for Developmental and Reproductive Biology (5th Floor, Room B529b), Biomedicum Helsinki, University of Helsinki, P.O. Box 700, FIN-00029 HUS, Helsinki, Finland. FAX: 35 894 717 1947; marjut.otala{at}hus.fi Back

Received: 25 April 2004.

First decision: 10 June 2004.

Accepted: 27 August 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Levade T, Jaffrezou JP. Signalling sphingomyelinases: which, where, how and why?. Biochim Biophys Acta 1999 1438:1-17[Medline]
  2. Marchesini N, Hannun YA. Acid and neutral sphingomyelinases: roles and mechanisms of regulation. Biochem Cell Biol 2004 82:27-44[CrossRef][Medline]
  3. Kolodny EH. Niemann-Pick disease. Curr Opin Hematol 2000 7:48-52[CrossRef][Medline]
  4. Otterbach B, Stoffel W. Acid sphingomyelinase-deficient mice mimic the neurovisceral form of human lysosomal storage disease (Niemann-Pick disease). Cell 1995 81:1053-1061[CrossRef][Medline]
  5. Horinouchi K, Erlich S, Perl DP, Ferlinz K, Bisgaier CL, Sandhoff K, Desnick RJ, Stewart CL, Schuchman EH. Acid sphingomyelinase deficient mice: a model of types A and B Niemann-Pick disease. Nat Genet 1995 10:288-293[CrossRef][Medline]
  6. Butler A, He X, Gordon RE, Wu HS, Gatt S, Schuchman EH. Reproductive pathology and sperm physiology in acid sphingomyelinase-deficient mice. Am J Pathol 2002 161:1061-1075[Abstract/Free Full Text]
  7. Kolesnick RN, Kronke M. Regulation of ceramide production and apoptosis. Annu Rev Physiol 1998 60:643-665[CrossRef][Medline]
  8. Kolesnick R, Fuks Z. Radiation and ceramide-induced apoptosis. Oncogene 2003 22:5897-5906[CrossRef][Medline]
  9. Haimovitz-Friedman A, Kan CC, Ehleiter D, Persaud RS, McLoughlin M, Fuks Z, Kolesnick RN. Ionizing radiation acts on cellular membranes to generate ceramide and initiate apoptosis. J Exp Med 1994 180:525-535[Abstract/Free Full Text]
  10. Verheij M, Bose R, Lin XH, Yao B, Jarvis WD, Grant S, Birrer MJ, Szabo E, Zon LI, Kyriakis JM, Haimovitz-Friedman A, Fuks Z, Kolesnick RN. Requirement for ceramide-initiated SAPK/JNK signalling in stress-induced apoptosis. Nature 1996 380:75-79[CrossRef][Medline]
  11. Spiegel S, Cuvillier O, Edsall LC, Kohama T, Menzeleev R, Olah Z, Olivera A, Pirianov G, Thomas DM, Tu Z, Van Brocklyn JR, Wang F. Sphingosine-1-phosphate in cell growth and cell death. Ann N Y Acad Sci 1998 845:11-18[CrossRef][Medline]
  12. Olivera A, Kohama T, Edsall L, Nava V, Cuvillier O, Poulton S, Spiegel S. Sphingosine kinase expression increases intracellular sphingosine-1-phosphate and promotes cell growth and survival. J Cell Biol 1999 147:545-558[Abstract/Free Full Text]
  13. Morita Y, Perez GI, Paris F, Miranda SR, Ehleiter D, Haimovitz-Friedman A, Fuks Z, Xie Z, Reed JC, Schuchman EH, Kolesnick RN, Tilly JL. Oocyte apoptosis is suppressed by disruption of the acid sphingomyelinase gene or by sphingosine-1-phosphate therapy. Nat Med 2000 6:1109-1114[CrossRef][Medline]
  14. Suomalainen L, Hakala JK, Pentikainen V, Otala M, Erkkila K, Pentikainen MO, Dunkel L. Sphingosine-1-phosphate in inhibition of male germ cell apoptosis in the human testis. J Clin Endocrinol Metab 2003 88:5572-5579[Abstract/Free Full Text]
  15. Santana P, Pena LA, Haimovitz-Friedman A, Martin S, Green D, McLoughlin M, Cordon-Cardo C, Schuchman EH, Fuks Z, Kolesnick R. Acid sphingomyelinase–deficient human lymphoblasts and mice are defective in radiation-induced apoptosis. Cell 1996 86:189-199[CrossRef][Medline]
  16. Pena LA, Fuks Z, Kolesnick RN. Radiation-induced apoptosis of endothelial cells in the murine central nervous system: protection by fibroblast growth factor and sphingomyelinase deficiency. Cancer Res 2000 60:321-327[Abstract/Free Full Text]
  17. Lozano J, Menendez S, Morales A, Ehleiter D, Liao WC, Wagman R, Haimovitz-Friedman A, Fuks Z, Kolesnick R. Cell autonomous apoptosis defects in acid sphingomyelinase knockout fibroblasts. J Biol Chem 2001 276:442-448[Abstract/Free Full Text]
  18. Otala M, Suomalainen L, Pentikainen MO, Kovanen P, Tenhunen M, Erkkila K, Toppari J, Dunkel L. Protection from radiation-induced male germ cell loss by sphingosine-1-phosphate. Biol Reprod 2004 70:759-767[Abstract/Free Full Text]
  19. Vergouwen RP, Jacobs SG, Huiskamp R, Davids JA, de Rooij DG. Proliferative activity of gonocytes, Sertoli cells and interstitial cells during testicular development in mice. J Reprod Fertil 1991 93:233-243
  20. Bligh EG, Dyer WJ. A rapid method for total lipid extraction and purification. Can J Biochem Physiol 1959 37:911-917
  21. Henriksen K, Kulmala J, Toppari J, Mehrotra K, Parvinen M. Stage-specific apoptosis in the rat seminiferous epithelium: quantification of irradiation effects. J Androl 1996 17:394-402[Abstract/Free Full Text]
  22. Henriksen K, Hakovirta H, Parvinen M. Testosterone inhibits and induces apoptosis in rat seminiferous tubules in a stage-specific manner: in situ quantification in squash preparations after administration of ethane dimethane sulfonate. Endocrinology 1995 136:3285-3291[Abstract]
  23. Parvinen M, Hecht NB. Identification of living spermatogenic cells of the mouse by transillumination-phase contrast microscopic technique for "in situ" analyses of DNA polymerase activities. Histochemistry 1981 71:567-579[CrossRef][Medline]
  24. Erkkila K, Henriksen K, Hirvonen V, Rannikko S, Salo J, Parvinen M, Dunkel L. Testosterone regulates apoptosis in adult human seminiferous tubules in vitro. J Clin Endocrinol Metab 1997 82:2314-2321[Abstract/Free Full Text]
  25. Pentikainen V, Suomalainen L, Erkkila K, Martelin E, Parvinen M, Pentikainen MO, Dunkel L. Nuclear factor-kappa B activation in human testicular apoptosis. Am J Pathol 2002 160:205-218[Abstract/Free Full Text]
  26. Toppari J, Bishop PC, Parker JW, diZerega GS. DNA flow cytometric analysis of mouse seminiferous epithelium. Cytometry 1988 9:456-462[CrossRef][Medline]
  27. Kangasniemi M, Veromaa T, Kulmala J, Kaipia A, Parvinen M, Toppari J. DNA flow cytometry of defined stages of rat seminiferous epithelium: effects of 3 Gy of high-energy X-irradiation. J Androl 1990 11:312-317[Abstract/Free Full Text]
  28. Erkkila K, Suomalainen L, Wikstrom M, Parvinen M, Dunkel L. Chemical anoxia delays germ cell apoptosis in the human testis. Biol Reprod 2003 69:617-626[Abstract/Free Full Text]
  29. Russell LD, Alger LE, Nequin LG. Hormonal control of pubertal spermatogenesis. Endocrinology 1987 120:1615-1632[Abstract/Free Full Text]
  30. Knudson CM, Tung KS, Tourtellotte WG, Brown GA, Korsmeyer SJ. Bax-deficient mice with lymphoid hyperplasia and male germ cell death. Science 1995 270:96-99[Abstract/Free Full Text]
  31. Rodriguez I, Ody C, Araki K, Garcia I, Vassalli P. An early and massive wave of germinal cell apoptosis is required for the development of functional spermatogenesis. Embo J 1997 16:2262-2270[CrossRef][Medline]
  32. Russell LD, Chiarini-Garcia H, Korsmeyer SJ, Knudson CM. Bax-dependent spermatogonia apoptosis is required for testicular development and spermatogenesis. Biol Reprod 2002 66:950-958[Abstract/Free Full Text]
  33. Jeyaraj DA, Grossman G, Petrusz P. Dynamics of testicular germ cell apoptosis in normal mice and transgenic mice overexpressing rat androgen-binding protein. Reprod Biol Endocrinol 2003 1:48[CrossRef][Medline]
  34. Jahnukainen K, Chrysis D, Hou M, Parvinen M, Eksborg S, Soder O. Increased apoptosis occurring during the first wave of spermatogenesis is stage-specific and primarily affects midpachytene spermatocytes in the rat testis. Biol Reprod 2004 70:290-296[Abstract/Free Full Text]
  35. Miranda SR, Erlich S, Visser JW, Gatt S, Dagan A, Friedrich VL Jr, Schuchman EH. Bone marrow transplantation in acid sphingomyelinase–deficient mice: engraftment and cell migration into the brain as a function of radiation, age, and phenotype. Blood 1997 90:444-452[Abstract/Free Full Text]
  36. Erlich S, Miranda SR, Visser JW, Dagan A, Gatt S, Schuchman EH. Fluorescence-based selection of gene-corrected hematopoietic stem and progenitor cells from acid sphingomyelinase–deficient mice: implications for Niemann-Pick disease gene therapy and the development of improved stem cell gene transfer procedures. Blood 1999 93:80-86[Abstract/Free Full Text]
  37. Furuchi T, Masuko K, Nishimune Y, Obinata M, Matsui Y. Inhibition of testicular germ cell apoptosis and differentiation in mice misexpressing Bcl-2 in spermatogonia. Development 1996 122:1703-1709[Abstract]
  38. Tilly JL, Kolesnick RN. Sphingolipids, apoptosis, cancer treatments and the ovary: investigating a crime against female fertility. Biochim Biophys Acta 2002 1585:135-138[Medline]
  39. Hasegawa M, Zhang Y, Niibe H, Terry NH, Meistrich ML. Resistance of differentiating spermatogonia to radiation-induced apoptosis and loss in p53-deficient mice. Radiat Res 1998 149:263-270[Medline]
  40. Chmura SJ, Nodzenski E, Beckett MA, Kufe DW, Quintans J, Weichselbaum RR. Loss of ceramide production confers resistance to radiation-induced apoptosis. Cancer Res 1997 57:1270-1275[Abstract/Free Full Text]
  41. Herr I, Wilhelm D, Bohler T, Angel P, Debatin KM. Activation of CD95 (APO-1/Fas) signaling by ceramide mediates cancer therapy– induced apoptosis. Embo J 1997 16:6200-6208[CrossRef][Medline]
  42. Tepper AD, Cock JG, de Vries E, Borst J, van Blitterswijk WJ. CD95/ Fas-induced ceramide formation proceeds with slow kinetics and is not blocked by caspase-3/CPP32 inhibition. J Biol Chem 1997 272:24308-24312[Abstract/Free Full Text]
  43. Bose R, Verheij M, Haimovitz-Friedman A, Scotto K, Fuks Z, Kolesnick R. Ceramide synthase mediates daunorubicin-induced apoptosis: an alternative mechanism for generating death signals. Cell 1995 82:405-414[CrossRef][Medline]
  44. Liao WC, Haimovitz-Friedman A, Persaud RS, McLoughlin M, Ehleiter D, Zhang N, Gatei M, Lavin M, Kolesnick R, Fuks Z. Ataxia telangiectasia–mutated gene product inhibits DNA damage–induced apoptosis via ceramide synthase. J Biol Chem 1999 274:17908-17917[Abstract/Free Full Text]
  45. Nava VE, Cuvillier O, Edsall LC, Kimura K, Milstien S, Gelmann EP, Spiegel S. Sphingosine enhances apoptosis of radiation-resistant prostate cancer cells. Cancer Res 2000 60:4468-4474[Abstract/Free Full Text]
  46. Hasegawa M, Wilson G, Russell LD, Meistrich ML. Radiation-induced cell death in the mouse testis: relationship to apoptosis. Radiat Res 1997 147:457-467[Medline]
  47. Erkkila K, Pentikainen V, Wikstrom M, Parvinen M, Dunkel L. Partial oxygen pressure and mitochondrial permeability transition affect germ cell apoptosis in the human testis. J Clin Endocrinol Metab 1999 84:4253-4259[Abstract/Free Full Text]
  48. Sinha Hikim AP, Lue Y, Diaz-Romero M, Yen PH, Wang C, Swerdloff RS. Deciphering the pathways of germ cell apoptosis in the testis. J Steroid Biochem Mol Biol 2003 85:175-182[CrossRef][Medline]
  49. Birbes H, Bawab SE, Obeid LM, Hannun YA. Mitochondria and ceramide: intertwined roles in regulation of apoptosis. Adv Enzyme Regul 2002 42:113-129[CrossRef][Medline]
  50. Gudz TI, Tserng KY, Hoppel CL. Direct inhibition of mitochondrial respiratory chain complex III by cell-permeable ceramide. J Biol Chem 1997 272:24154-24158[Abstract/Free Full Text]
  51. Siskind LJ, Colombini M. The lipids C2- and C16-ceramide form large stable channels. Implications for apoptosis. J Biol Chem 2000 275:38640-38644[Abstract/Free Full Text]
  52. Spiegel S, Foster D, Kolesnick R. Signal transduction through lipid second messengers. Curr Opin Cell Biol 1996 8:159-167[CrossRef][Medline]
  53. Mathias S, Pena LA, Kolesnick RN. Signal transduction of stress via ceramide. Biochem J 1998 335:465-480
  54. Hannun YA, Luberto C, Argraves KM. Enzymes of sphingolipid metabolism: from modular to integrative signaling. Biochemistry 2001 40:4893-4903[CrossRef][Medline]
  55. Ohanian J, Ohanian V. Sphingolipids in mammalian cell signaling. Cell Mol Life Sci 2001 58:2053-2068[CrossRef][Medline]
  56. Spence MW, Burgess JK, Sperker ER. Neutral and acid sphingomyelinases: somatotopographical distribution in human brain and distribution in rat organs. A possible relationship with the dopamine system. Brain Res 1979 168:543-551[CrossRef][Medline]



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