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Regular Article |
a Developmental Biology Program, Institute of Biotechnology, University of Helsinki, FIN-00014 Helsinki, Finland
b The Institute of Biogenesis Research, Department of Anatomy and Reproductive Biology, University of Hawaii Medical School, Honolulu, Hawaii 96822
c Department of Comparative Medicine
d Department of Molecular and Medical Genetics, Oregon Health & Science University, Portland, Oregon 97201-3098
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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2 and GnRH, that were previously mapped to that region and the functions of which in spermatogenesis are well established were not altered in DMS. As a consequence, positional cloning of the DMS locus will be essential to identify new molecules potentially involved in arrest at M-I. Furthermore, mice carrying this genetic trait might be useful for studies of assisted reproductive technologies and male contraceptives.
assisted reproductive technology, apoptosis, meiosis, spermatogenesis, testis
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Spermatogenesis is complex: it requires hormonal regulation originating in the central nervous system, as well as a large number of well-orchestrated molecular mechanisms controlling cellular differentiation in the seminiferous tubules of the testis. In mammals, spermatogenesis is a lifelong process that can be classified into three phases: 1) proliferation, 2) meiosis, and 3) spermiogenesis [6]. During the proliferation phase, spermatogonia stem cells give rise to both stem cells and committed precursors, the latter of which continue to differentiate into spermatocytes. The ratio between self-renewal and differentiation of the spermatogonia stem cells is regulated by glial cell line-derived neurotrophic factor (GDNF) [7]. After the proliferative phase, spermatocytes enter meiosis, a complex process sequentially leading to haploid round spermatids, elongated spermatids, and spermatozoa. Characteristic of meiosis are the molecular mechanisms related to checkpoint control [8]. Checkpoint control genes were first identified in yeast and Drosophila from mutants showing meiotic arrest and apoptosis [9]. Mammalian homologues of those checkpoint control genes that were inactivated by targeted mutagenesis revealed similar phenotypes [8]. Male sterility due to meiotic arrest, during pachytene stage or metaphase I, has been identified in a number of genetic models in the mouse, predominantly by targeted gene knockout, but also by standard transgenesis (for review see Venables and Cooke [10]). The present study relates to a line of transgenic mice in which transgene insertion caused a dominant trait of male sterility. Characterization of this phenotype revealed a spermatogenesis defect, which was due to spermatocytes undergoing meiotic arrest and apoptosis.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Expression plasmid pVgRXR (Invitrogen, Carlsbad, CA) was restricted with SfiI, a 6.9-kilobase (kb) DNA fragment isolated and used to generate transgenic mice by microinjection. B6D2F1 mice were used as the embryo donor strain for transgene injections, and the protocols employed followed standard procedures described by Hogan et al. [11].
Histology
Postnatal testes and epididymis from DMS mice and nontransgenic littermates were freshly dissected, fixed in 4% paraformaldehyde for 2–24 h depending on size, and processed in paraffin. Sections were cut at 5-µm thickness and stained with hematoxylin and eosin.
TUNEL Labeling
After deparaffinization of slides, apoptotic cells were detected with the ApopTag in situ Apoptosis Detection Kit (Intergen, Purchase, NY). The kit was used according to the manufacturer's instruction. Sections were then subjected to nuclear counterstaining with methyl green according to the manufacturer's instructions (Vector Laboratories, Inc., Burlingame, CA).
Immunohistochemistry
Deparaffinized testis sections were incubated overnight with rat monoclonal antibody GATA-1 at a dilution of 1:50 [7] (Santa Cruz Biotechnology, Santa Cruz, CA) at 4°C, washed three times for 5 min in PBS, and incubated with biotinylated anti-rat immunoglobulin G secondary antibody (Vector Laboratories) at 1:500 for 30 min at room temperature. After washes in PBS, the sections were treated by standard labeling procedures according to the manufacturer's instructions (Vector Laboratories). After immunohistochemical staining, nuclear counterstaining was performed with methyl green according to the manufacturer's instruction (Vector Laboratories).
Radioactive In Situ Hybridization
Radioactive in situ hybridization was performed as previously described [12]. Antisense and sense RNA probes from rat 2-kb full-length GDNF family receptor (GFR)2 cDNA, mouse 1-kb epithelial microtubule-associated protein (E-MAP)-115 cDNA, and mouse 3ß-hydroxysteroid dehydrogenase isomerase (HSD) cDNA described previously [13–15] were synthesized using appropriate RNA polymerases and 35S-labeled UTP. The hybridization temperature was 52°C, and autoradiography slides were exposed at 4°C for 2 wk. Histologic counterstaining was performed with hematoxylin. The slides were photographed with an Olympus Provis microscope equipped with a CCR camera (Photometrics Ltd., Harpenden, Herts, UK). Photograph editing was done using PhotoShop 4.0 (Adobe Systems, San Jose, CA). Darkfield images were inverted, artificially stained in red, and combined with brightfield images.
Fluorescence In Situ Hybridization
Primary fibroblast cells from line DMS were treated with 20 ng/ml colcemid for 2 h followed by hypotonic treatment for 10 min using 0.075 M KCl. Cells were then fixed with modified Carnoy fixative (3:1, methanol:glacial acetic acid) for 10 min and processed as described [16]. Chromosome spreads were made by dropping cell suspension onto a clean microscope slide. The slides were stained with Hoechst 33258 (Molecular Probes, Eugene, OR) (1 µg/ml) and exposed to UV light (302 nm) for 30 min. The probe, pVgRXR, was nick-translated using digoxygenin-11-dUTP (Boehringer-Mannheim, 
Roche Diagnostics, Nutley, NY). One slide of cell line DMS, denatured in 70% formamide, and 90 ng of labeled probe (also denatured) were hybridized overnight. The next day, the slide was postwashed in 50% formamide (15 min at 43°C) followed by a 2x saline-sodium citrate (SSC) wash (8 min at 37°C) and then detected with anti-digoxygenin-rhodamine. The process was repeated the following week using the same protocol but detecting with anti-digoxygenin-fluorescein isothiocyanate (FITC) instead of rhodamine. The image analysis for acquisition, display, and quantification of hybridization signals was performed with Cytovision System (Applied Imaging, Santa Clara, CA).
Southern Blot Hybridization Analysis
Southern blot hybridization was performed according to standard methods as previously described [17]. Wild-type and DMS mouse genomic DNAs were digested with various restriction enzymes including BamHI, BclI, BglII, BstEII, EcoRI, and HindIII, and 10 µg per lane was separated on an agarose gel. The hybridization of the DNA was done with [-32P]dCTP-labeled rat GFR2 full-length cDNA or mouse 206-bp GnRH cDNA probes. Hybridization was performed at 42°C for 16 h in a solution containing 50% formamide, 6x SSPE, 5x Denhardt, 0.5% SDS, and 100 µg/ml denatured salmon sperm DNA. After hybridization, the filter was washed five times, 1 min each, at room temperature, in 2x SSC, 0.1% SDS, and twice for 15 min at 65°C in 0.2x SSC, 0.1% SDS. The filter was exposed to a Fuji phosphoimager screen o/n and processed in a FujiBAS phosphoimager (Fuji, Stamford, CT).
Total RNA Isolation and Northern Blotting
Total RNAs were isolated with TRIZOL Reagent kit (Gibco Life Technology, Gaithersburg, MD) according to the manufacturer's protocol, and 30 µg of total RNA was used in each lane. Northern blotting was carried out according to standard protocol [17]. The probes of GFR2 and GnRH cDNA (described previously) labeled with [32P]dCTP were used in hybridization as in Southern blot analysis. The filters were exposed overnight in a FujiBAS phosphoimager screen, and the screen was developed and analyzed in the FujiBAS phosphoimager.
Flow Cytometry
Karyotypes of seminiferous epithelia were analyzed by FACScan flow cytometry (Becton Dickinson, Franklin Lakes, NJ) after ethidium bromide or propidium iodide labeling of the nuclei (CellFIT Cell-Cycle Analysis version 2.0.2). Mouse spleen cells served as diploid cell controls.
Intracytoplasmic Sperm Injection and Round Spermatid Injection
Intracytoplasmic sperm injection (ICSI) and round spermatid injection (ROSI) were performed according to standard procedures [18, 19] with some modification. The donor spermatozoa were obtained from testis, cauda epididymis, and round spermatids isolated from the testis of DMS mutant male mice at age 11 mo. Oocytes were obtained from donor B6D2F1 females at approximately 12 wk of age. Surrogate mothers of developing embryos were CD1 females approximately 4 mo of age. Two-cell embryos developed from ICSI and ROSI oocytes were transferred into the oviduct of surrogate females (CD1) that had mated with vasectomized males of the same strain during the previous night. The females were allowed to deliver and raise their young. Females without visible signs of pregnancy were killed on Day 19.5 dpc (days postcoitum) and examined for the presence or absence of fetuses and implantation sites in the uteri.
In detail, a sperm mass was collected from the cauda epididymis and placed at the bottom of a 1.5-ml plastic centrifuge tube containing 200 µl of CZB medium (37°C) (Specialty Media, Phillipsburg, NJ). This allowed the spermatozoa to disperse into the medium. We found that the spermatozoa were immotile. Spermatozoa viability was then further examined using a live-dead sperm-staining kit (FertiLight; Molecular Probes). A single spermatozoon with normal head and tail (less than 1% of the entire sperm population) was selected and injected into an oocyte. A drop of sperm suspension was mixed (1:2) with 10% polyvinyl pyrolidone (PVP) (w/v) in Hepes-CZB; then, the head of a spermatozoon was separated from the tail by applying a few piezo pulses to the head-tail junction [20]. Only the head was injected into each oocyte. The operations were carried out at room temperature (25°–27°C). Sperm injections were completed within 1 h after the sperm suspension was mixed in PVP medium (Medi-Cult, Hopkinton, MA). Sperm-injected oocytes were placed in 50-µl droplets of CZB under mineral oil and incubated at 37°C in a humidified atmosphere of 5% CO2 in air.
For ROSI, seminiferous tubules placed in Hepes-CZB were cut into small pieces using a pair of fine scissors and pipetted vigorously with a small-bore pipette to release spermatozoa and spermatogenic cells into the medium. Round spermatids, characterized by their small size as well as a centrally located, distinct nucleolus, were individually picked up in a large-bore pipette and transferred to 10% PVP in Hepes-CZB. The nucleus of each spermatid was injected into a single oocyte. A group of 15 spermatid-injected oocytes was activated by culturing them for 5–6 h in Ca2+-free CZB containing 10 mM SrCl2 [21]. Those oocytes with two distinct pronuclei and a second polar body were considered normally fertilized. Normally fertilized ICSI and ROSI oocytes were further cultured in CZB until they reached the two-cell stage.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Microinjection of a 6.9-kb fragment of transgene DNA encoding the ecdyson-regulated expression cassette pVgRXR resulted in five founder animals, three females and two males, which were fertile and bred normally. Screening for the presence of the transgene was performed by polymerase chain reaction analysis of mouse genomic DNA obtained from tail samples (data not shown). In the process of establishing transgenic lines from the different founders, it was found that F1 generation males from female founder 101 were sterile. Female transgenic offspring was fertile, but males failed to produce offspring, suggesting a dominant inheritance of male sterility (DMS). In fact, transgenic males mated and produced vaginal plugs in females, but these matings did not result in any signs of pregnancies. Gross anatomical analysis of reproductive organs from DMS mice, including ovary, uterus, seminal vesicle, epididymis, and penis, did not reveal abnormalities; even the testis were not noticeably smaller than those in nontransgenic littermates.
Histologic Analysis
Normal spermatogenesis in nontransgenic littermates at the age of 6 wk gave rise to a number of round and elongated spermatids at the end of the first wave (Fig. 1A). In contrast, the mutant littermates' testes showed only a small number of spermatids but a large number of meiotic metaphase I figures, suggesting a block of progression from metaphase to anaphase during the first meiotic division (Fig. 1, B and C). Premeiotic pachytene spermatocytes and metaphase I spermatocytes had extensive darkly stained chromatin and eosinophilic cytoplasm (Fig. 1C), which are histologic characteristics of degenerating germ cells. Progressive degeneration in the epithelia of seminiferous tubules from DMS mice was not severe; phenotypically, the differences observed between 16-wk-old adult mouse testis and 6-wk-old mouse testis were minimal (Fig. 1, D–F). In DMS testis, the greatest amount of degeneration was found in M-I spermatocytes, but degeneration was also found to a much lesser extent at the spermatid stage (Fig. 1G). Consistent with the arrest of spermatogenesis observed in DMS testis was the pronounced absence of spermatozoa in the epididymal duct. Instead of mature sperm, round cells, possibly arrested spermatogenic cells and round spermatids, were present in the duct, perhaps sloughed off from the seminiferous tubules (Fig. 1, H and I).
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In Situ Assay for Apoptosis
Arrest during meiotic progression is usually followed by programmed cell death, according to Ashley [8]. To assess the fate of the differentiating germ cells in DMS testis, specific staining for cells undergoing programmed cell death was evaluated by TUNEL labeling. Apoptotic cells, although rarely found in wild-type testis (Fig. 2A), were present in abundance among spermatocytes mostly in stage XII seminiferous tubules in DMS testis (Fig. 2B). Most TUNEL-positive cells were either metaphase I or premeiotic pachytene spermatocytes (Fig. 2, B and C). Cell types other than germ cells were occasionally TUNEL positive, suggesting that apoptosis occurred after alterations of the tubular microenvironment (Fig. 2C). Apoptotic germ cells could also be found in the epididymal duct of the DMS mouse line (Fig. 2D), suggesting that some degenerating seminiferous epithelial cells became discharged from the tubules and migrated to the epididymal duct.
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Gene Expression in DMS Testis
To assess whether meiosis metaphase I arrest in DMS spermatocytes was cell autonomous or caused by malfunctioning and dying somatic cells in the tubules, Sertoli cell-specific and Leydig cell-specific gene expression patterns were evaluated. In seminiferous tubules, GATA1 protein is expressed specifically in Sertoli cell nuclei, where it assumes a segmental distribution pattern. Immunohistochemical analysis of Sertoli cells from DMS mice and control littermates revealed a normal distribution pattern of GATA1 (Fig. 3, A and B). Sertoli cell function is further characterized by the 115-kDa epithelial microtubule-associated protein (E-MAP-115), a structural component required for the normal manchette formation between Sertoli cells and spermatids [14]. In situ hybridization studies showed that E-MAP 115 was expressed in both wild-type and DMS testis (Fig. 3, E and F). Reduced E-MAP-115 signal density in DMS mice is likely due to the reduced number of spermatids. Leydig cells specifically express 3ß-hydroxysteroid dehydrogenase isomerase (3ß-HSD) [15], which controls the steroidogenic pathways in the gonads, as well as in the adrenal cortex; it is a key enzyme in regulating the formation of 
4-3-ketosteroids from 5-3-ß-hydroxysteroids. The expression pattern of 3ß-HSD in Leydig cells in DMS mice (Fig. 3D) was similar to the pattern in wild-type mice (Fig. 3C).
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Mapping Transgene Locus by Fluorescence In Situ Hybridization
A critical objective was to identify the chromosomal locus of transgene insertion. Meeting this objective was an essential step in identifying candidate genes that might be affected by the transgene insertion. Toward this goal, fluorescence in situ hybridization (FISH) was performed using a cDNA probe specific for the transgene. The FISH probe used from the transgene pVgRXR included the VP-16/ecdyson receptor region, which does not contain sequences with extensive homology to any known mouse locus. FISH analysis was performed on primary fibroblasts derived from the DMS mouse line, which were mitotically arrested at metaphase with colcemid. Hybridization was performed with a digoxygenin-labeled probe as described [16]. Detection with FITC showed unequivocally a fluorescent signal on one copy of chromosome 14, at cytogenetic offset D3-E1. The signal is consistent with a hemizygous transgene insertion locus; its intensity suggests that tandem arrays of several copies of the transgene had inserted (Fig. 4).
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Southern, Northern, and In Situ Analyses of Candidate Genes
Southern blot analysis of genomic DNA, restriction-digested with various enzymes, using a GFR2 cDNA probe showed that the patterns between DMS and wild-type animals were identical (Fig. 5A). Northern analysis using RNA from several tissues isolated from DMS mice and nontransgenic control littermates was performed. Using this analysis, GFR2 expression was identified in the brain of both wild-type animals and DMS transgenic mice. No detectable difference in the magnitude of the signal was observed (Fig. 5B). Expression of GFR2 in the testis was analyzed using in situ hybridization and was localized to both round and elongated spermatids from wild-type testis (Fig. 5C). In DMS testis, the GFR2 signal was found to be localized on the occasionally formed round spermatids, but the overall expression level was substantially diminished (Fig. 5D). This diminished expression may be a consequence of the reduced number of cells present. Another candidate gene, GnRH, which was previously mapped to chromosome 14, also did not show polymorphisms in either Southern or Northern analysis (data not shown).
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Flow Cytometry
Karyotype analysis of seminiferous epithelial cells from DMS mice and control littermates was performed using flow cytometry (Fig. 6, A and B). Consistent with histologic analysis of spermatocytes showing metaphase I arrest, the number of haploid cells (1n) in the DMS mouse testis was dramatically reduced to approximately half of the number of haploid cells in the wild-type mouse testis. In contrast, the number of cells with 2n and 4n DNA content was comparable in testis from DMS mice and control wild-type littermates.
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Progeny Yielded From Intracytoplasmic Round Spermatids Injection but Not Spermatozoa
Two series of experiments were performed. Spermatozoa and spermatids of one male were used for both ICSI and ROSI on the same day. The data from those experiments are summarized in Table 1.
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Intracytoplasmic sperm injection We found that cauda epididymal spermatozoa were invariably immotile; less than 1% had normal heads and tails. All of the spermatozoa were diagnosed "dead" by the live/dead sperm staining, implying that their plasma membranes had been damaged extensively. Even though the epididymal spermatozoa were dead in a conventional sense, about half of the sperm-injected oocytes were fertilized, and most of them were developed to two-cell stage. However, neither of two surrogate mothers that received these embryos became pregnant. No embryos and no implantation sites were seen in the uteri of surrogate mothers on 19.5 dpc, indicating that the transferred two-cell embryos either did not reach the blastocyst stage or, if they did, did not implant.
Round spermatid injection In contrast, two surrogate females became pregnant after transfer of ROSI-derived two-cell embryos. One female delivered one pup (one male), and the other delivered five pups (two males, three females).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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2 and GnRH, were not shown to be implicated in the DMS mouse line. The DMS male mice did mate normally and produce copulation plugs at normal frequency. This indicated that they did not have any neurologic or behavioral problems related to infertility. Male infertility phenotypes in men and in mouse models are occasionally associated with neurologic syndromes such as, for example, in the mutant weaver [22]. Neurologic manifestations can also be implicated with reduced fertility because of a lower mating frequency, as observed in the estrogen receptor gene knockout [23].
All transgene-positive male mice were sterile. Examination of the spermatogenesis defects in DMS testis revealed metaphase arrest of meiosis I accompanied by large numbers of apoptotic spermatocytes. We initially set out to investigate the cellular origin of the mutation causing spermatocyte arrest by checking gene expression in somatic cells. Expression of GATA1 and 3ß-HSD in Sertoli cells and Leydig cells of DMS testis, respectively, was indistinguishable from that in control testis, suggesting that somatic cell types were not overtly affected. However, as the mutated locus is not known, it is currently impossible to determine if the phenotype is due to aberrant signaling by somatic cells or if it is germ cell autonomous. Either of these scenarios could be implicated. Abnormal Sertoli cell or Leydig cell function has been associated with a substantial number of cases of sterility in men [24] and with mouse models deficient for Bclw [25] and protein C inhibitor [26].
Programmed cell death of male germ cells is a fairly common cause of sterility. As spermatocytes become committed to meiosis, they are subjected to checkpoint controls that engage a multitude of genes to monitor the status and fidelity of the reduction division; failure to pass checkpoint control results in apoptosis [8, 9]. The mouse model carrying a targeted mutation in the meiotic checkpoint control gene ataxia-telangiectasia, ATM [27], supports this model. Checkpoint control-mediated meiotic arrest—at early zygotene to midlate pachytene stages—and subsequent apoptosis are common when extensive regions of unpaired DNA are caused by transgene insertion or radiation damage. However, this mechanism is unlikely in DMS mice because the total size of the transgene insertion is on the order of around 100 kb, substantially less than the sizes reported for insertions of BAC or YAC transgenes which did not affect male fertility. Examples of large segments of unpaired chromosomes from tandem arrays of the transgene DNA and translocations causing meiotic arrest have previously been reported [2]. The sterility phenotype in DMS is sex specific as it affects spermatogenesis but not oogenesis. Mapping of the locus of transgene insertion revealed a locus on chromosome 14; this indicates that although the phenotype is sex specific, genotypically there is no sex linkage. Autosomal mutations specifically affecting spermatogenesis have been described for a number of loci [10].
Male sterility has been frequently found in mice carrying targeted mutations at loci that are ubiquitously expressed, including the protooncogene c-abl [28], the transcription factor Egr4 [29], and the basigin locus, which encodes a protein that belongs to the immunoglobulin superfamily [30]. Targeted gene knockout of DNA for the mismatch repair genes MLH1 and PMS2 [31], the heat shock protein HSP 70-2 [32], and the RNA-binding protein, TLS [33] had similar consequences. In addition, heterologous expression of certain transgenes has been shown to cause male sterility phenotypes; those transgenes include the v-mos oncogene, but not the c-mos oncogene, which caused meiotic arrest and anomalies of spindles due to altered patterns of -, ß-, and 
-tubulins and phosphoproteins [34].
We have established that the DMS mutation is inherited in a dominant fashion. This dominant trait may be compatible with the following two models. 1) Transgene insertion may lead to the ectopic expression or silencing of a gene that interferes with meiosis I. 2) Alternatively, the insertion may truncate a gene causing a dominant-negative mutant. The second scenario would most likely apply if the molecules involved functioned as dimers or multimers. As we do not yet have detailed information about the locus of transgene insertion, it is currently not possible to establish if either of the these models, or an alternative model, best describes the DMS mouse mutant. However, a common aspect to dominant mutations is variable penetrance of the phenotypic consequences. Incomplete penetrance of meiotic arrest causing sterility has been reported for the Egr4 gene knockout in mice. In this study, spermatocytes showed metaphase I arrest and apoptosis, except for a small number of cells that evaded checkpoint control and developed to spermatids and spermatozoa. This arrest and apoptosis resulted in severe oligozoospermia. The detailed mechanisms that account for incomplete penetrance of meiotic arrest in the absence of the zinc-finger transcription factor, Egr4, as demonstrated by Tourtellotte et al. [29], have yet to be shown. In the DMS mutant line, we showed a further example of sterility inherited with incomplete penetrance of spermatogenesis. Because the spermatogenesis was incomplete, a few round spermatids and spermatozoa could be collected. This provided the opportunity to evaluate these cells in assisted reproductive technologies.
Assisted fertilization techniques such as ICSI have become routine in human infertility clinics to overcome male infertility due to severe oligospermia and asthenospermia [35]. In the mouse, sperm precursors such as spermatids and spermatocytes have been used as substitutes for spermatozoa using modified protocols [17, 36]. Surprisingly, in our attempts to use spermatozoa and round spermatids from DMS transgenic mice, in ICSI and ROSI, respectively, we found that the latter but not the former approach was successful in rescuing male progeny. It is very likely that the spermatogenic cells of DMS transgenic mice die shortly after their transformation from spermatids to spermatozoa. It is possible that elongated spermatids and recently transformed spermatozoa are still alive and competent to participate in fertilization and development.
Molecular cloning of the locus of transgene insertion will eventually elucidate the molecular mechanism underlying this mutation and, as it is a dominant trait, permit the evaluation of strategies for male contraceptives. Toward that goal, the locus of transgene insertion was identified on chromosome 14 D3-E1. Insertional mutations caused by transgenes can be accompanied by large chromosomal rearrangements, severely hampering attempts for their molecular identification and characterization. The availability in the near future of DNA sequences covering the entire mouse genome should help accelerate the molecular analysis of transgene insertions, as small query sequences will readily permit the establishment of a contiguous map of a locus.
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ACKNOWLEDGMENTS |
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2 probe, Dr. A.H. Payne (University of Michigan) for the 3ß-HSD probe, Dr. M. Komada and Dr. P. Soriano (Fred Hutchinson Cancer Researcher Center, Seattle, WA) for the plasmid pBS-EMAP, and Dr. Sergio Ojeda (Primate Center, Oregon Health Sciences University) for providing the GnRH probe. We also thank Anne Scullin for editing the manuscript. |
FOOTNOTES |
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1 This work was supported in part by the Oregon Cancer Institute. 
2 Correspondence: Manfred Baetscher, Department of Comparative Medicine, Oregon Health & Science University, 3181 SW Sam Jackson Park Rd., Portland, OR 97201-3098. FAX: 503 494 4338; baetsche{at}ohsu.edu
3 Current address: Department of Psychiatry & Behavioral Sciences, University of Washington Medical Center, Seattle, WA 98195
Accepted: October 24, 2001.
Received: August 29, 2001.
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, and 
, respectively