Molecular Human Reproduction, Vol. 6, No. 3, 219-225,
March 2000
© 2000 European Society of Human Reproduction and Embryology
Testis and spermatozoa |
Expression of protamine-1 and -2 mRNA during human spermiogenesis
1 Institut für Veterinäranatomie, Frankfurter Strasse 98, D-35392 Giessen, 2 Institut für Pathologie, Langhansstrasse 10, D-35392 Giessen, and 3 Institut für Anatomie und Zellbiologie, Grosse Steinstrasse 52, D-06097 Halle, Germany
Abstract
During spermiogenesis, the histone-to-protamine replacement causes the compaction of the spermatid chromatin. The genes for protamines, PRM-1 and PRM-2, are transcribed in round and elongating spermatids. The transcripts are stored in a translationally-repressed state by the binding of protein repressors before being translated in elongating and elongated spermatids. RNA extracts from homogenized whole testis samples supply only average data, and cell-specific and stage-specific expression cannot be addressed. Therefore, we used UV-laser-assisted cell-picking (UV-LACP) to select spermatids of defined differentiation steps. Subsequent reverse transcription–polymerase chain reaction (RT–PCR) with intron-spanning primer pairs allowed the detection of DNA-free and pseudogene-free PRM-1 and PRM-2 cDNA. Additional in-situ hybridization with digoxygenin-labelled cRNA probes exhibited PRM-1 and PRM-2 mRNA from step 1/2 spermatids to step 4 spermatids, but not in elongated spermatids. RT–PCR revealed amplicons for PRM-1 and PRM-2 in all spermatids except step 3 round spermatids. Applying proteinase K digestion, PRM-1 and PRM-2 transcripts were also detected in step 3 spermatids indicating that protein repressors may bind to both PRM-1 and PRM-2 mRNA in step 3 round spermatids. These data demonstrate that the combination of UV-LACP and non-radioactive in-situ hybridization appear to be a suitable approach for the study of cell-specific and stage-specific gene expression during spermiogenesis.
human testis/in-situ hybridization/laser-assisted cell-picking/protamines/spermiogenesis
Introduction
In round spermatids, both histones and non-histone proteins are replaced by transition proteins. In elongating spermatids, transition proteins are removed from the condensing chromatin and are replaced by protamines constituting the nuclear proteins of elongated spermatids and mature spermatozoa (Hecht, 1989a
,b, 1990a,Hecht, b; Oliva and Dixon, 1991; Dadoune, 1995; Siffroi et al., 1999; Steger, 1999). In man, the histone-to-protamine exchange is only ~85% complete (Tanphaichitr et al., 1978; Gatewood et al., 1987; Prigent et al., 1996).
Protamine-bound DNA is the most tightly compacted DNA (Pogany et al., 1981; Ward and Coffey, 1991) causing cessation of transcription in elongating spermatids. Thus major modifications in both nuclear and cytoplasmic structures continue throughout spermiogenesis; stringent temporal and stage-specific gene expression via transcriptional and translational control mechanisms is of fundamental importance to ensure complete differentiation of round spermatids into mature spermatozoa. In transgenic mice, premature expression of protamine-1 causes precocious nuclear condensation and arrests spermatid differentiation (Lee et al., 1995).
Specific sequences in the 3'-untranslated region (3'-UTR) of PRM-1 mRNA cause delayed translation of both a human growth hormone reporter mRNA (Braun et al., 1989) and a human green fluorescent protein reporter mRNA (Schmidt et al., 1999). Since transgenes lacking these sequences showed no delayed translation (Braun, 1990; Fajardo et al., 1997), these sequences appear responsible for masking PRM-1 mRNA. In round spermatids, most of the mRNA is stored as translationally inactive ribonucleoprotein (RNP) particles, involving protein repressors binding to specific sequences located in the 3'-UTR or to the 160–180 residues counting poly-A tail of the transcripts (Stern et al., 1983; Hecht, 1989a,b, 1990a, Hecht, b; Morales et al., 1991; Dadoune, 1995; Kleene, 1996; Steger, 1999). Translation subsequently takes place in elongating and elongated spermatids after mRNA undergo a partial poly-A shortening by deadenylation. Before or during translation, mRNA for both protamines are subjected to a shortening process, namely from 0.62 to 0.45 kb for PRM-1 mRNA and from 0.9 to 0.7 kb for PRM-2 mRNA (Domenjoud et al., 1991). It has been shown that 85–95% of PRM-1 and PRM-2 mRNA with poly-A tails of 160–180 residues are translationally inactive, whereas 80–95% of PRM-1 and PRM-2 mRNA with poly-A tails of 30 residues are translationally active (Kleene et al., 1984, Heidaran and Kistler, 1987; Kleene, 1989, 1993).
Recently, protamine-2 has been recommended as spermatogenic cell marker for molecular diagnosis of spermatogenesis in non-obstructive azoospermia providing valuable information about the existence, the stage of differentiation and the physiology of spermatids and spermatozoa (Lee et al., 1998). Although protamine-1 and -2 proteins are known to be present from step 4 elongating spermatids to step 8 elongated spermatids (Roux et al., 1987, 1988; LeLannic et al., 1993; Lescoat et al., 1993; Prigent et al., 1996; Siffroi et al., 1999) and transcripts for PRM-1 and PRM-2 have been observed in human spermatids (Wykes et al., 1995, 1997; Saunders et al., 1996; Siffroi et al., 1999), mRNA expression has not been assigned to the various steps of spermatid differentiation.
In the present study, the cell-specific and stage-specific expression of both PRM-1 and PRM-2 mRNA were investigated to gain further insight into the regulation of gene expression in haploid spermatids during normal spermiogenesis. Non-radioactive in-situ hybridization as well as UV-laser-assisted cell-picking (UV-LACP) followed by reverse transcription–polymerase chain reaction (RT–PCR) analysis, recently demonstrated as appropriate methods for the study of testicular tissue (Steger et al., 1998, Pauls et al., 1999), were applied and compared regarding the sensitivity of both methods.
Materials and methods
Testicular tissue
Extraction of RNA was performed on four testes from two orchidectomized men with prostatic carcinoma, aged 52 and 72 years. The patients were not treated with any drugs prior to orchidectomy. For UV-LACP, two testes from a 63 year old man with epididymitis and two testes from a 63 year old man with prostatic carcinoma were snap-frozen and stored in liquid nitrogen. For in-situ hybridization, 20 testicular biopsy specimens from 10 men (aged 30–45 years; mean 37.2 years) with obstructive azoospermia were fixed by immersion in Bouin's fixative and embedded in paraffin using standard techniques. All patients revealed normal endocrine values and histologically normal spermatogenesis (score 
8, according to Holstein and Schirren, 1983).
UV-LACP, first strand cDNA synthesis, and RT–PCR
For UV-LACP, 5 µm cryostat sections were mounted onto glass slides, stained with haematoxylin for 45 s, and immersed in absolute ethanol. Spermatids from defined stages of the seminiferous epithelial cycle were collected using the UV-laser MicroBeam system (Palm GmbH, Wolfratshausen, Germany) and the inverted Axiovert 135 microscope (Carl Zeiss, Oberkochen, Germany). The isolated cell profiles were harvested by a sterile injection needle mounted on a digitally controlled micromanipulator (Palm GmbH), transferred into a reaction tube containing 10 µl of first strand buffer (52 mmol/l Tris–HCl, 78 mmol/l KCl, 3.1 mmol/l MgCl2, pH 8.3), snap-frozen, and stored at –80°C until further processing.
Prior to cDNA synthesis, reaction tubes with picked cell profiles were incubated at 70°C for 10 min and then rapidly cooled on ice-water. For proteinase K (Sigma, Deisenhofen, Germany) digestion, the picked cell profiles were treated with either 100 µg/ml proteinase K or 400 µg/ml proteinase K to release RNA from RNA-binding proteins. After incubation at 53°C for 30 min, samples were incubated at 99°C for 10 min to destroy proteinase K.
First strand cDNA synthesis was performed using the GeneAmp RNA–PCR Kit, according to the manufacturer's instructions (Perkin Elmer, Foster City, CA, USA). 2 µl each of MgCl2 (25 mmol/l) and 10x PCR buffer II (100 mmol/l Tris–HCl, 500 mmol/l KCl, pH 8.3), 1 µl each of dNTPs (5 mmol/l each), random hexamers (50 µmol/l), and MuLV reverse transcriptase (50 IU), and 0.5 µl of RNase inhibitor (10 IU) were added to 10 µl of first strand buffer containing the picked cell profiles, and incubated at 20°C for 10 min, at 43°C for 75 min, and at 99°C for 5 min.
Using the GeneAmp RNA–PCR Kit, according to the manufacturer's instructions (Perkin Elmer), RT–PCR conditions were 1x 2 min 45 s 94°C/60x 45 s 94°C, 45 s 58°C, 45 s 72°C/1x 7 min 72°C/4°C. 8 µl of cDNA were added to 4 µl 10x PCR buffer II, 3 µl MgCl2 (25 mmol/l), 8 µl dNTPs (5 mmol/l each), 0.5 µl AmpliTaqGoldTM, 1.5 µl of each primer (10 µmol/l), and ultrapure H2O to a final volume of 50 µl. The DNA sequences for human PRM-1 and PRM-2 (Domenjoud et al., 1990) were generated using the primer pairs listed in Table I.
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The primer pairs employed in this study flanked the single intron of PRM-1 DNA and PRM-2 DNA to preclude genomic DNA amplification within the expected product length of 153 bp (PRM-1 cDNA) and 294 bp (PRM-2 cDNA). To exclude genomic DNA or pseudogene amplification, genomic DNA was isolated from similar testicular tissue with the QIAmp Blood Kit (Qiagen, Hilden, Germany). PCR analysis was carried out with the same primer pairs applied in RT–PCR. Amplification products were visualized by agarose gel electrophoresis and ethidium bromide staining. For each RT–PCR, controls were performed by omitting reverse transcriptase or by using primary spermatocytes instead of spermatids.
Cloning of human PRM-1 and PRM-2 cDNA, and digoxigenin-labelled cRNA probes for PRM-1 and PRM-2 mRNA
Total RNA was extracted from testicular tissue with Trizol Reagent, according to the manufacturer's protocol (Life Technologies, Eggenstein, Germany). First strand cDNA synthesis was performed using the Superscript II Kit, according to the manufacturer's instructions (Life Technologies).
The cDNA clones for human PRM-1 and PRM-2 (Domenjoud et al., 1990) were generated using RT–PCR (1x 3 min 95°C/35x 1 min 95°C, 1 min 69°C, 2 min 72°C/1x 10 min 72°C/4°C) with the primer pairs listed in Table I
.
A 156 bp RT–PCR product of the human PRM-1 cDNA and a 294 bp RT–PCR product of the human PRM-2 cDNA were subcloned in pGEM-T (Promega, Heidelberg, Germany). The plasmids were transformed in the XL1-Blue Escerichia coli strain (Stratagene, Heidelberg, Germany) and extracted by column purification, according to the manufacturer's instruction (Qiagen).
In-vitro transcription of digoxygenin (DIG)-labelled cRNA was performed using the RNA–DIG Labelling Mix (Boehringer Mannheim, Mannheim, Germany) and RNA-polymerases T7 and SP6. Prior to cRNA synthesis, the vectors containing the PRM-1 and PRM-2 inserts had been digested with NcoI or NotI (New England Biolabs, Schwalbach, Germany) for the production of sense cRNA (NcoI) or antisense cRNA (NotI). After phenol extraction, the dried pellet was reconstituted in 100 µl RNase-free water. The concentration of the DIG–RNAs was estimated by a semiquantitative dot blot test (Jackson, 1991).
In-situ hybridization
In-situ hybridization was performed as previously reported (Steger et al., 1998). Briefly, 7 µm sections from Bouin-fixed and paraffin-embedded tissue samples were mounted on slides coated with aminopropyltriethoxysilane (Sigma). Deparaffinized and rehydrated tissue sections were digested with proteinase K (20 µg/ml 1x PBS) for 30 min at 37°C, post-fixed in 4% paraformaldehyde for 10 min and prehybridized in 20% glycerol for 30 min. Sections were then incubated with the DIG-labelled sense and antisense cRNA probes. Both PRM-1 cRNA and PRM-2 cRNA were used at a dilution of 1:100 in hybridization-buffer containing 50% deionized formamide, 10% dextran sulphate, 2x sodium chloride/sodium citrate (SSC), 1x Denhardt's solution, 10 µg/ml salmon sperm DNA (Sigma) and 10 µg/ml yeast t-RNA (Sigma). Hybridization was performed overnight at 37°C in a humid chamber containing 50% formamide in 2x SSC.
Post-hybridization washes were performed, according to a previously described method (Lewis and Wells, 1992). Tissue samples were incubated overnight at 4°C with an anti-DIG Fab-antibody conjugated to alkaline phosphatase (Boehringer Mannheim). Staining was visualized with NitroBlue Tetrazolium/5-bromo-4-chloro-3-indolyl phosphate (NBT/BCIP) (KPL, Gaithersburg, MD, USA) in a humid chamber and protected from light. Finally, sections were mounted in Glycergel (Dako, Hamburg, Germany). For each test, control incubations were performed using DIG-labelled cRNA sense probes.
Quantification and statistical analysis
In sections subjected to in-situ hybridization, the ratio of labelled to unlabelled spermatids was determined in 10 seminiferous tubules from each biopsy. For stage-specific quantification (Clermont, 1963), this ratio was determined in 10 seminiferous tubules for each stage of the seminiferous epithelial cycle. Data were analysed using Student's t-test. P < 0.05 was considered to be statistically significant.
Results
In-situ hybridization visualized a stage-specific expression of PRM-1 and PRM-2 mRNA (Table II; Figure 1). In stage I/II of the seminiferous epithelial cycle containing two generations of spermatids, namely steps 1/2 round spermatids and step 7/8 elongated spermatids, 62 and 53% of step 1/2 round spermatids revealed a weak signal for PRM-1 and PRM-2 respectively, whereas step 7/8 elongated spermatids were completely negative for both PRM-1 and PRM-2 (Figures 1 A, B). 80.5% (PRM-1) and 76% (PRM-2) of step 3 round spermatids (stage III; Figures 1 C, D) and 62% (PRM-1) and 68.5% (PRM-2) of step 4 elongating spermatids (stage IV; Figures 1 E, F) exhibited a strong signal for both PRM-1 and PRM-2. Step 5 elongated spermatids (stage V; Figures 1G,H) were again completely devoid of both PRM-1 and PRM-2 mRNA. Summarized, ~68% and ~66% of steps 1–4 spermatids revealed signals for PRM-1 and PRM-2 mRNA respectively. Differences between steps 1/2, step 3, and step 4 spermatids were not significant. Control incubations with DIG-labelled PRM-1 and PRM-2 cRNA sense probes were completely negative (data not shown).
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Employing UV-LACP, cell profiles of 10–20 spermatids of a defined step of differentiation were cut out of the seminiferous epithelium (Figures 2A–D
). Subsequent RT–PCR analysis of these cell profiles revealed transcripts of both PRM-1 and PRM-2 in all steps of spermatid differentiation. However, we noticed considerable variation among different UV-LACP processed cell profiles in the efficiency of RT–PCR amplification (Figure 3A). With the exception of step 3 round spermatids, the average success of RT–PCR amplification of PRM-1 and PRM-2 transcripts was 53 and 81% respectively (Table II). In steps 1/2 round spermatids, revealing a weak staining by in-situ hybridization, 46% of RT–PCR samples (PRM-1) and 73% of RT–PCR samples (PRM-2) displayed positive signals. In step 4 elongating spermatids, revealing a strong staining by in-situ hybridization, 50% of RT–PCR samples (PRM-1) and 100% of RT–PCR samples (PRM-2) exhibited positive signals. In elongated spermatids, which were completely negative by in-situ hybridization, 71% (step 5/6) and 46% (step 7/8) of RT–PCR samples were positive for PRM-1, while 75% (step 5/6) and 75% (step 7/8) of RT–PCR samples were positive for PRM-2. In step 3 round spermatids, which revealed a strong staining by in-situ hybridization, only 8% (PRM-1) and 25% (PRM-2) of RT–PCR samples exhibited positive signals. However, in step 3 round spermatids, positive signals for both PRM-1 and PRM-2 cDNA could be obtained and, therefore, the discrepancy between in-situ hybridization and RT–PCR analysis could be removed when cell profiles were digested with proteinase K prior to first strand cDNA synthesis and RT–PCR amplification (Table II; Figure 3B). Using 100 µg/ml proteinase K and 400 µg/ml proteinase K, 25 and 100% of RT–PCR showed positive signals for PRM-1, while 37.5% and 50% of RT–PCR showed positive signals for PRM-2 respectively. Primary spermatocytes which are known to be devoid of protamines, revealed no positive signals and, therefore, served as negative control (Figures 3A,B).
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Discussion
The human genes PRM-1, PRM-2, and TNP-2 exist as a linear array on chromosome 16p13.3 (Domenjoud et al., 1990, Schlüter et al., 1992) being regulated as a single genetic unit (Choudhary et al., 1995). Applying radioactive in-situ hybridization, PRM-2 mRNA was shown to be the most abundant transcript in human testis. PRM-1 and TNP-2 mRNA were present in approximately equal quantities (Wykes et al., 1995). These results are in contrast with those of others suggesting that TNP-2 mRNA is expressed at a very low level in human testis (Schlueter et al., 1992, 1993). The human TNP-2 gene differs from that of other mammalian species, that have been investigated so far, by the absence of the conserved 5'GCCATCAC3' nucleotide sequence in the 3'-UTR (Schlueter et al., 1992, 1993). The difficulty in demonstrating TNP-2 transcripts in human testis may be due to insufficient storage of TNP-2 mRNA as RNP particles. Using Northern blot analysis and fluorescence in-situ hybridization, PRM-2 mRNA was again the most abundant transcript in human testis. Comparatively, transcripts for PRM-1 and TNP-2 were present at a level of ~50% and ~3% of that of PRM-2 mRNA (Choudhary et al., 1995). Applying non-radioactive in-situ hybridization, TNP-2 mRNA cannot be demonstrated in human testis (Steger et al., 1998).
In this study, we investigated the expression of both PRM-1 and PRM-2 mRNA during normal spermatogenesis. In contrast to previous studies (Choudhary et al., 1995; Wykes et al., 1995, 1997; Saunders et al., 1996), we were able to demonstrate cell-specific and stage-specific expression applying non-radioactive in-situ hybridization and UV-LACP followed by RT–PCR analysis.
PRM-1 and PRM-2 mRNA was demonstrated in the cytoplasm from steps 1/2 round spermatids to step 4 elongating spermatids. While both mRNA revealed only weak hybridization signals in step 1/2 spermatids, strong staining was observed in step 3 and step 4 spermatids. Therefore, translation of the corresponding proteins, P1 and P2, known to be present in the nucleus from step 4 to step 8 spermatids (Roux et al., 1987, 1988; LeLannic et al., 1993; Lescoat et al., 1993; Prigent et al., 1996; Siffroi et al., 1999), appears to be delayed.
In haploid spermatids, translational arrest is due to the storage of mRNA as RNP particles (Stern et al., 1983; Penttilä et al., 1995; Kleene, 1996) in chromatoid bodies which can frequently be observed in round spermatids (Biggiogera et al., 1990; Moussa et al., 1994). The translational arrest is caused by the binding of protein repressors to the 3'-UTR or the poly-A tail of mRNA. Numerous mouse mRNA-associated proteins have already been identified (Grange et al., 1987; Kwon and Hecht, 1991, 1993; Berger et al., 1992; Murray et al., 1992; Kwon et al., 1993, Fajardo et al., 1994; Kleene et al., 1994; Gu et al., 1995; Schumacher et al., 1995a,b; Lee et al., 1996).
Step 3 round spermatids exhibited a strong signal using in-situ hybridization, but in contrast displayed nearly no signal applying the more sensitive RT–PCR technique. However, amplification of both PRM-1 and PRM-2 cDNA could be obtained when cell profiles were subjected to proteinase K treatment prior to first strand cDNA synthesis and RT–PCR amplification. This is in line with data obtained by in-situ hybridization where paraffin sections had also been subjected to proteinase K digestion prior to hybridization of the DIG-labelled cRNA probe. These data suggest a tight binding of protein repressors preventing the transcripts of PRM-1 and PRM-2 from being translated in step 3 round spermatids.
For step 1/2 and step 4 spermatids, data obtained by UV-LACP RT–PCR were in line with those obtained by in-situ hybridization. RT–PCR producing no amplification products may be explained by the selection of spermatids being devoid of transcripts for PRM-1 and PRM-2. During normal spermiogenesis, only 68 and 66% of round and elongating spermatids revealed hybridization signals for PRM-1 and PRM-2 mRNA respectively.
RT–PCR positive signals in step 5/6 and step 7/8 elongated spermatids being completely negative using in-situ hybridization may be caused by remnants of untranslated mRNA, which can only be detected by the more sensitive RT–PCR technique. Both PRM-1 and PRM-2 mRNA have been demonstrated even in epididymal and ejaculated human spermatozoa (Pessot et al., 1989; Kumar et al., 1993; Miller et al., 1994; Miller, 1997; Wykes et al., 1997).
Applying non-radioactive in-situ hybridization, both protamines exhibited an identical localization with protamine-2 showing less intensive signals than protamine-1. However, quantitative evaluation revealed no significant differences in the distribution of PRM-1 and PRM-2 mRNA. These results are in contrast with previous studies (Choudhary et al., 1995; Wykes et al., 1995) which demonstrated that, in human testis, PRM-1 mRNA is present at a level of ~50% of that of PRM-2 mRNA. These data correspond with the results obtained by RT–PCR following UV-LACP. Here, the average success of RT–PCR amplification (except step 3 round spermatids) of PRM-1 and PRM-2 transcripts was 53 and 81% respectively.
In conclusion, our data demonstrate that the combination of non-radioactive in-situ hybridization and UV-LACP RT–PCR is a suitable approach for the study of cell and stage-specific gene expression during spermiogenesis. Furthermore, the inclusion of proteases more specific than proteinase K prior to first strand cDNA synthesis may contribute to our understanding of the timing and functional interaction of protein repressor molecules to mRNA transcripts.
Acknowledgments
We are grateful to Professor Dr L.Hertle, Department of Urology of the University, Münster, for providing the biopsies. The skilful technical assistance of C. Fröhlich and A. Hax is gratefully acknowledged. We like to thank PD Dr R.M.Bohle, Professor Dr W.Kummer and Professor Dr W.Seeger, project 21 of SFB 547, for constant support during the course of the study. Funding of this research programme was provided by DFG grant STE 892/1–1.
Notes
4 To whom correspondence should be addressed 
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Submitted on September 17, 1999; accepted on December 6, 1999.
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