Modification of chromosomal architecture in human spermatozoa with large vacuoles

Authors


Correspondence:

Anne Perdrix, Reproductive Biology Laboratory – CECOS, EA 4308 “Spermatogenesis and male gamete quality”, Rouen University Hospital, 1 rue de Germont, 76031 Rouen Cedex, France.

E-mail: anne.perdrix@chu-rouen.fr

Summary

Human normal spermatozoa present a specific chromatin organization, illustrated particularly by the non-random chromosome positioning. Spermatozoa with large vacuoles, described using motile sperm organelle morphology organization (MSOME), are associated with nuclear alterations, such as abnormal chromatin condensation and aneuploidy. To question a probable association between large nuclear vacuoles and chromatin disorganization, we evaluated chromosomes X, Y and 18 topography in normal spermatozoa (NS) compared with spermatozoa with large vacuoles (SLV). After centrifugation on a gradient density system, 229 NS (spermatozoa presenting a normal nuclear shape and a vacuole area <6.5% of head area) from 10 normal semen samples and 221 SLV (spermatozoa presenting a vacuole area >13% of head area) from 10 semen samples with teratozoospermia were selected using MSOME. A three-colour FISH was carried out using α-satellite centromeric probes for chromosomes X, Y and 18. For each chromosome, longitudinal and spatial positioning of centromeres was analysed. Distribution of each chromosome was non-random in NS and in SLV, whatever the methodology used. Using longitudinal positioning, distribution of chromosome 18 and chromosome Y centromeres did not differ significantly between SLV and NS. On the contrary, chromosome X centromeres were more frequently positioned in the posterior region of sperm nucleus in SLV (= 0.01). Considering spatial positioning, distributions differed significantly between SN and SLV for chromosome Y (= 0.02) and chromosome 18 (< 10−4) and marginally for chromosome X (= 0.08). Our study concluded to a modification in chromosomes X, Y and 18 centromere topography between NS and SLV, representing a novel and supplementary evidence to argue chromatin disorganization in SLV.

Introduction

Spermatozoa are characterized by a unique chromatin organization, based on the replacement during spermiogenesis of histones by protamines; as a result, sperm chromatin is coiled into toroids (Ward, 1993), instead of nucleosome organization observed in somatic cells (Ward & Coffey, 1991), allowing an extreme compaction of sperm DNA. A supplementary level of complex nuclear organization is represented by chromosome topology. Indeed, as observed in somatic cells (Foster & Bridger, 2005; Cremer & Cremer, 2010), individual chromosomes occupy distinct territories (Haaf & Ward, 1995; Zalensky et al., 1995) and the preferential longitudinal positioning has been established for 11 chromosomes in human spermatozoa (Zalensky & Zalenskaya, 2007).

The condensed, insoluble and highly organized nature of sperm chromatin acts to protect genetic integrity during transport of the paternal genome through the male and female reproductive tracts. It also ensures that paternal DNA is delivered in the form that sterically allows the proper fusion of two gametic genomes and enables the developing embryo to correctly express the genetic information (Erenpreiss et al., 2006). Sperm nucleus seems consequently crucial for fertilization and embryo development. Moreover, accumulating evidence suggests that disturbances in the organization of the genomic material in sperm nuclei are negatively correlated with the fertility potential of spermatozoa (Agarwal & Said, 2003). Several sperm nuclear alterations have been associated with infertility: sperm DNA damage (Barratt et al., 2010), sperm chromatin immaturity (Oliva, 2006), sperm aneuploidy (Tempest & Griffin, 2004). Role of sperm chromosome topography in male infertility have also been questioned (Finch et al., 2008; Ioannou et al., 2011), and the specific architecture of sperm chromatine seemed to be an integral part of epigenetic mechanism in embryogenesis (Cremer et al., 2004; Claussen, 2005).

As a consequence, in Assisted Reproduction Technology (ART), sperm nucleus appeared more and more determinant to improve pregnancy rates and became the centre of interest of several studies (Lewis, 2007; Barratt et al., 2010). Among sperm nuclear investigations, Bartoov et al. (2001) proposed an observation system, in real time and at a high magnification of up to ×6600 called the Motile Sperm Organelle Morphology Examination (MSOME), using an inverted microscope equipped with Nomarski interferential contrast optics. Normal spermatozoa have been defined, and new abnormalities, as nuclear vacuoles, have been described (Bartoov et al., 2002). Large vacuoles have been associated with sperm nuclear alterations (i) DNA damage (Franco et al., 2008; Garolla et al., 2008; Oliveira et al., 2010; de Almeida Ferreira Braga et al., 2011), (ii) aneuploidy (Garolla et al., 2008; Perdrix et al., 2011), and (iii) chromatin immaturity (Boitrelle et al., 2011; Cassuto et al., 2011; Franco et al., 2012; Perdrix et al., 2011). The presence of large vacuoles in sperm heads has been suspected to be deleterious on ART outcome (Berkovitz et al., 2006a; Vanderzwalmen et al., 2008; Cassuto et al., 2009; de Almeida Ferreira Braga et al., 2011; Oliveira et al., 2011). To clarify vacuole impact on embryo development, the chromatin assessment in spermatozoa with large vacuoles became of major interest (Boitrelle et al., 2011).

The purpose of this study was to precise the chromatin alterations associated with the presence of large vacuoles in sperm head. We used chromosome positioning to assess chromatin integrity, comparing chromosome topology between spermatozoa with large vacuoles and normal spermatozoa.

Materials and methods

Patients

A total of 20 men were included in this study. All the participants gave their agreement to provide semen samples for the study. Ten men, with age ranged from 26 to 41 years, presented normal semen parameters [according to World Health Organization (WHO) criteria; WHO, 2010] and were considered as controls. Ten other men, with age ranged from 28 to 43 years, presented altered semen parameters, with, at least, teratozoospermia (normal forms <50%, according David's modified classification) (Auger et al., 2001). Karyotypes in peripheral blood lymphocytes were analyzed for patients with altered sperm parameters and were normal for every patient.

Cell preparation

Semen samples were collected after 3–5 days of sexual abstinence, and liquefied for 20 min at 37 °C. After performing semen analysis according to WHO (2010) guidelines for each sample, 1 mL of native spermatozoa was centrifugated on a gradient density system (PureSperm 100; JCD, La Mulatière, France). A droplet of the motile fraction obtained was placed into a glass box (Willco Sterile, GWST-100; Biosoft International, Amsterdam, The Netherlands) under sterile paraffin oil (Ovoil 100; Vitrolife, Göteborg, Sweden) to perform high magnification observations. For each patient, spermatozoa morphology was carried out at high magnification (MSOME) (×6600) using an inverted microscope equipped with Nomarski differential interference contrast optics (Leica DMI 6000B; Leica, Solms, Germany). Twenty-five spermatozoa were randomly photographed (Leica Application Suite version 3.4.0; Leica) and morphology assessment was conducted using Leica IM 1000 software. Length, width and surface of head, number as well as area of vacuoles were also assessed with this software (Saïdi et al., 2008).

Then, motile spermatozoa selection was performed at ×6600 magnification using a micropipette injection (ICSI Micropipets MIC-50-30; Humagen, Charlottesville, VA, USA) and a micromanipulator system. Two types of spermatozoa were isolated from the two types of semen samples (i) normal spermatozoa (NS), representing spermatozoa without any vacuoles, or with a unique small vacuole (vacuole area <6.4% of head area), from the 10 samples with normal semen parameters, (ii) spermatozoa with large vacuoles (SLV), representing spermatozoa with a vacuole area >13% of sperm head area, from the 10 abnormal semen samples (Saïdi et al., 2008; Perdrix et al., 2012). For each type of spermatozoa 20–50 spermatozoa per patient were selected and put down on glass slides (Cytoslides; Thermo Fisher Scientific, Runcorn, Cheshire, UK) in a spotted area. The number of isolated spermatozoa was limited by the selection duration. A maximum time of 2 hours has been used for this technique to avoid possible effects of time selection on the incidence of vacuolated spermatozoa (Peer et al., 2007). After air-drying, sperm cells were fixed with methanol (Chelli et al., 2010; Perdrix et al., 2011). The slides were stored in a freezer at −20 °C until their use.

Slides were washed in phosphate-buffered saline (PBS) (Biomérieux, Marcy l'Etoile, France), then in 2 × SSC (standard saline citrate) (Sigma-Aldrich, Saint-Quentin Fallavier, France). Sperm cells were briefly decondensed using a solution of 25 mm DTT (Dithiothreitol) (Sigma Chemical CO, Saint Louis, MO, USA), 1 m TrisHcl pH 9.5 at room temperature. Treatment with DTT has been proved to preserve sperm nucleus structure (Hazzouri et al., 2000), not only sperm shape (Celik-Ozenci et al., 2003) but also sperm chromocenter (Gurevitch et al., 2001). Slides were washed in 2 × SSC, then in PBS, and dehydrated in a 70–100% ethanol series and air-dried.

FISH

A three-colour FISH was carried out using α-satellite centromeric probes for chromosome X in green (CEP X Spectrum Green; Abbott, Rungis, France), chromosome Y in yellow (CEP Y Sat III Spectrum Orange, CEP Y Sat III Spectrum Green; Abbott) and chromosome 18 in red (CEP 18 Spectrum Orange; Abbott). The yellow colour was obtained by mixing an equal volume of CEP Y labelled with Spectrum Orange and CEP Y labelled with Spectrum Green (Milazzo et al., 2006; Garolla et al., 2008; Wiland et al., 2008; Perdrix et al., 2011).

Briefly, probes and hybridization solution were removed from the freezer (−20 °C) and allowed to warm up to room temperature. The hybridization mixture contained each centromeric probe filled up with hybridization solution to a final volume of 9 μL. Nine microlitre of the hybridization mixture were placed on the spotted area of the slide, and a 20 mm × 20 mm coverslip was carefully applied. Both sperm cells and probes were simultaneously denatured at 73 °C for 5 min on heated plate (Hybaid; Omnigene, Teddington, UK) and hybridized overnight at 37 °C.

After hybridization, coverslips were removed. Slides were washed once in a solution of SSC 0.4X/NP40 0.3% (Ipegal; Sigma Chemical CO) for 2 min at 73 °C and once in a solution of SSC 2X/NP40 0.1% at room temperature for 1 min. After dehydration in a 70–100% ethanol series, slides were counterstained with a 4′,6-diamidino-2-phenylindole (DAPI) (Counterstain 1, Adgenix, Voisin le Bretonneux, France) solution diluted in antifade mounting medium (Antifade; MP-QBio-Gene, Illkirch, France) and coverslips were mounted. A highly concentrated DAPI solution (1 μg/mL) was used to detect more easily the few number of MSOME selected spermatozoa fixed on glass side.

Microscopy

Spermatozoa were examined with an epifluorescence microscope (Nikon Microphot-FXA; Nikon Instruments, Champigny sur Marne, France) at a magnification of ×630. A triple band-pass filter set (FITC/rhodamine/DAPI) was used. Images were acquired with a CCD camera (MetaSystems, Altlussheim, Germany). The positioning of the chromosome centromeres in the sperm cells was analysed using ISIS (In Situ Imaging System) MetaSystems software (v. 5.0; Altlussheim, Germany). Green, yellow and red signals detected chromosomes X, Y and 18, respectively. Two fluorescent spots of comparable size and intensity, separated by at least one spot diameter, were considered as two copies of the corresponding chromosome. Spermatozoa with diffuse fluorescent signals and overlapping nuclei were classified as ambiguous and were not included in the count. Chromosome relative positions were analyzed in a minimum of 200 normal spermatozoa and 200 spermatozoa with large vacuoles. Aneuploidy frequency was defined by the sum of frequencies of 1818XX, 1818YY and 1818XY diploid spermatozoa, XX, YY and 1818 disomic spermatozoa as well as XY hyperhaploid spermatozoa.

Chromosome positioning

Centromere positions of chromosomes X, Y and 18 were analysed using 2 methodologies: linear positioning and positioning in 2 dimensions (2D).

Linear positioning

Sperm nucleus was divided into three equal parts along its long axis, from the acrosome to the point of sperm tail attachment, defining anterior part (a), median part (m) and posterior part (p). The frequency of centromeres for each chromosome positioned in the anterior, median and posterior part of the sperm head was calculated (Fig. 1).

Figure 1.

Determination of linear and spatial positioning of chromosome centromeres in human sperm nuclei. (a) Example of FISH centromere signals in sperm cells. The nuclear counterstain is DAPI (blue), the chromosome X centromeric probe is green and the chromosome 18 centromeric probe is red. Assessment was performed at ×630 magnification. (b and c) Schematic view of the sperm nucleus presenting a red signal for chromosome 18 centromere and a green signal for chromosome X centromere. (b) Illustration of linear positioning: sperm nucleus was divided in three equal parts: anterior (a), median (m) and posterior (p); FISH centromere signals were localized within each part. (c) Illustration of spatial positioning: using the methodology proposed by Zalenskaya & Zalensky (2004): each FISH signal was localized using three geometrical parameters: L (length of the long sperm head axis), H (elevation over the long axis) and D (distance from the tail attachment point).

2D positioning

Centromeres were localized along the short and long sperm axis to evaluate if chromosomes were situated deep inside the nucleus (central localization) or close to the sperm membrane (peripheral localization), close to the acrosome (anterior localization) or near the tail (posterior localization). We used the graphic model described by Zalenskaya & Zalensky (2004). Several geometrical parameters such as length of long axis (L: from tail attachment point to acrosome); distance from FISH centromere signal to tail attachment point (D); distance from FISH centromere signal and long axis (H) were assessed. Values of D/L and H/L were calculated, corresponding to the chromosome centromere coordinates along the long and the short axis, respectively, of the sperm head. The highest D/L values close to 1.0 marked the position of centromeres close to an acrosome area. The lowest H/L values close to 0.0 marked the central position of the centromeres within the sperm nucleus; the highest H/L values close to 0.5 marked position more peripherally (Fig. 1).

Statistical analysis

Semen and MSOME parameters were compared between patients and controls using Mann–Whitney U-test. Vacuole distributions obtained with MSOME analysis were compared using Chi-squared test. Aneuploidy frequencies were compared between normal spermatozoa and large vacuoles spermatozoa using Chi-squared test.

For linear positioning, Chi-squared test was used to determine whether the distribution of chromosome centromeres between anterior, median and posterior parts of the sperm nuclei were statistically different from random distribution. Then, chromosome positionings were compared between normal spermatozoa and spermatozoa with large vacuoles using Chi-squared test.

For 2D positioning, Kolgomorov–Smirnov test was used to compare chromosome distribution to random distribution. Mean H/L and D/L values were compared between the two types of spermatozoa using the t-test or Mann–Whitney U-test. Then, to assess chromosome topography in 2D, using individual values in opposition with mean values, a table was created with three columns representing the D/L values varying from 0.0 to 1.0 and spaced by 0.33, and two lines for H/L values from 0.0 to 0.5 and spaced by 0.25. In the 6 table-cells, centromere distribution of each chromosome was compared between normal spermatozoa and spermatozoa with large vacuoles using the chi-squared test. A value of < 0.05 was considered to be significant.

Results

Studied population

Semen parameters and MSOME analysis are summarized in Table 1. Teratozoospermia was observed in all patient semen sample; combined with asthenozoospermia (progressive motility <32%) for five patients, and with oligoasthenozoospermia (sperm concentration <15 × 106/mL or <39 × 106/ejaculate, and progressive motility <32%) for three patients. Sperm cell count (= 0.0005), progressive motility (= 0.0003) and normal forms (< 10−4) were significantly decreased in patients compared with controls.

Table 1. Conventional semen parameters, MSOME parameters and aneuploidy rates in MSOME selected spermatozoa considered as normal or with large vacuoles, in 10 patients and ten controls. A value of < 0.05 was considered to be significant
Semen parametersMSOME parametersSelected spermatozoa
Sperm count (106/mL)Progressive motility (%)Normal Morphology (%)Sperm head length (μm)Sperm head width (μm)Sperm head size (μm2;)Sperm vacuoles mean numberMean vacuoles area (%)Vacuole localizationNumber of SLVNumber of NSAneuploidy rate (%)
AntMedPost
% n % n % n
  1. Ant: anterior; Med: median; MSOME: Motile Sperm Organelle Morphology Examination; n: number of vacuoles observed; NS: normal spermatozoa; Post: posterior; SEM: standard error of the mean; SLV: spermatozoa with large vacuoles. a= 0.0005. b= 0.0003. c< 10−4. d= 0.57. e= 0.001. f= 0.06. g= 0.16. h< 10−4. i= 0.09.

Patients
 P11010334.53.312.41.68.233.31359237.7329 3.5
 P24525485.23.113.91.710.948.82148.8210027 0.0
 P3220105.43.214.22.316.643.92554.4311.8129 13.8
 P45830265312.11.38.551.51739.4139.1317 17.6
 P52040215.53.114.21.710.426.21157.12416.7714 35.7
 P61630424.83.213.31.98.258.32837.5180023 13.0
 P71635235.23.213.91.91755.32638.3186.4315 13.3
 P8820155.42.913.81.812.254.32543.5202.2121 14.3
 P9520184.9312.51.81546.72148.9222.2122 9.1
 P1019251653.2132.211.548.12642.6239.3524 16.7
 Mean ± SEM19.9 ± 5.5a25.5 ± 2.7b25.2 ± 3.9c5.1 ± 0.1d3.1 ± 0.1e13.3 ± 0.2f1.8 ± 0.1g11.8 ± 1.1h46.6 ± 3.1 46.9 ± 2.5 5.5 ± 1.7 22.1 ± 1.7 13.7 ± 3.0i
Controls
 C15035625.13.515.51.349.4384.4276.32 224.5
 C24845764.83.413.52.23.432.71827.3154022 3212.5
 C324355353.213.81.7638.11661.92600 219.5
 C46540614.93.414.124.535.31858.8305.93 2516.0
 C59035695.23.414.41.95.436.21759.6284.32 210.0
 C63035515.43.415.41.6412.2580.5337.33 911.1
 C76045545.13.314.71.47.822.9871.4255.72 2010.0
 C8185405053.313.61.37.524.2875.82500 185.5
 C9165456353.213.71.22.235.51158.1186.52 395.1
 C103840524.83.213.41.47.926.5967.6235.92 229.1
 Mean ± SEM75.5 ± 17.7a39.5 ± 1.4b59.1 ± 2.7c5.1 ± 0.1d3.3 ± 0.1e14.2 ± 0.2f1.6 ± 0.1g5.2 ± 0.6h27.3 ± 3.2 64.5 ± 5.1 8.1 ± 3.6  22.9 ± 2.58.3 ± 1.4i

Using MSOME analysis, despite a sperm head width increased in patients (= 0.001), sperm head size did not differ significantly between patients and controls (= 0.06). Vacuole area was larger in patients than in controls (< 10−4), but vacuole number was similar (= 0.16). Vacuoles were preferentially localized in the anterior and median parts of sperm heads, (91.8% in controls, 93.5% in patients). Vacuole distributions differed significantly between controls and patients (< 10−4), with more vacuoles located in the anterior part of sperm heads in patients.

There was no significant difference in aneuploidy rates in SLV from patients (13.7%) compared with SN from controls (8.3%) (= 0.09) (Table 1).

Chromosome positioning

Linear positioning

Results of linear positioning assessment are presented in Table 2. Centromere distributions were not random for each chromosome, in normal spermatozoa and in spermatozoa with large vacuoles. Most of centromeres were located in the median part of sperm head (range of means: 44.3–64.3%), whatever the chromosome and whatever the type of spermatozoa. Topographies of chromosome 18 and chromosome Y centromeres did not differ significantly between normal spermatozoa and spermatozoa with large vacuoles. On the contrary, chromosome X distribution was significantly different in normal spermatozoa compared with spermatozoa with large vacuoles (= 0.01); indeed, chromosome X centromeres were more frequently posterior in spermatozoa with large vacuoles.

Table 2. Linear positioning of chromosome X, Y and 18 centromeres, distributed between the anterior, median and posterior parts of sperm head, in normal spermatozoa and spermatozoa with large vacuoles. A value of < 0.05 was considered to be significant
ChromosomeChromosome distribution in normal spermatozoa %(n)Chromosome distribution in spermatozoa with large vacuoles %(n)Comparison of chromosome distributions between normal spermatozoa and spermatozoa with large vacuoles (pb)
Anterior localizationMedian localizationPosterior localizationComparison to random distribution (pa)Anterior localizationMedian localizationPosterior localizationComparison to random distribution (pa)
  1. n: number of centromeres observed.

  2. a

    Chromosome distributions compared with an expected random distribution of 33.3/33.3/33.3 using the chi-squared test.

  3. b

    Comparison of chromosome distributions between normal spermatozoa and spermatozoa with large vacuoles using the chi-squared test.

X 35 (41)60.7 (71)4.3 (5)<10−442.5 (42)44.3 (47)13.2 (14)0.0020.01
Y27.7 (31)64.3 (72) 8 (9)<10−428.7 (35)57.4 (70)13.9 (17)10−40.09
1822.6 (48)60.4 (128)17 (36)<10−4 23 (48)52.6 (110)24.4 (51)<10−30.14

2D positioning

Results of centromere distribution evaluated using H/L and D/L parameters are presented in Table 3. Figure 2 provides an illustration of the centromere distribution in 2D, in the virtual matrix used for Chi-squared test. Centromere distributions were not random for each chromosome, in normal spermatozoa and in spermatozoa with large vacuoles. Considering mean values (i) comparing D/L values using T-test, chromosomes 18, X and Y appeared substantially more anterior in normal spermatozoa vs. spermatozoa with large vacuoles and (ii) comparing H/L values using Mann–Whitney U-test, chromosomes 18 and X appeared more peripheral in normal spermatozoa vs. spermatozoa with large vacuoles, but no difference was observed for chromosome Y. Considering every individual centromere values (i) distributions in 2D of chromosome 18 and chromosome Y centromeres differed significantly between normal spermatozoa and spermatozoa with large vacuoles (< 10−3 and = 0.04 respectively) and (ii) for chromosome X, 2D distribution was marginally different between normal spermatozoa compared with spermatozoa with large vacuoles (= 0.08).

Table 3. Positioning in two dimensions of chromosome X, Y and 18 centromeres, using the graphic model proposed by Zalenskaya & Zalensky (2004), in normal spermatozoa and spermatozoa with large vacuoles. A value of < 0.05 was considered to be significant
Centromere co-ordinateChromosome XChromosome YChromosome 18
Normal spermatozoa (= 116)Spermatozoa with large vacuoles (= 103)Comparison between normal spermatozoa and spermatozoa with large vacuoles (pb)Normal spermatozoa (= 112)Spermatozoa with large vacuoles (= 122)Comparison between normal spermatozoa and spermatozoa with large vacuoles (pb)Normal spermatozoa (= 212)Spermatozoa with large vacuoles (= 209)Comparison between normal spermatozoa and spermatozoa with large vacuoles (pb)
Mean ± SEComparison to random distribution (pa)Mean ± SEComparison to random distribution (pa)Mean ± SEComparison to random distribution (pa)Mean ± SEComparison to random distribution (pa)Mean ± SEComparison to random distribution (pa)Mean ± SEComparison to random distribution (pa)
  1. D/L: chromosome centromere co-ordinate along the long axis of the sperm head; H/L: chromosome centromere co-ordinate along the short axis of the sperm head; n: Number of centromeres observed.

  2. a

    Chromosome 2D distributions compared with an expected random distribution using Kolgomorov–Smirnov test.

  3. b

    Comparison of D/L and H/L mean values obtained for chromosome X, Y and 18 centromeres in normal spermatozoa and spermatozoa with large vacuoles using t-test or Mann–Whitney U-test.

D/L0.64 ± 0.01<10−40.59 ± 0.01<10−40.030.60 ± 0.01<10−40.56 ± 0.01<10−40.040.56 ± 0.01<10−40.53 ± 0.01<10−40.05
H/L0.16 ± 0.01<10−40.12 ± 0.01<10−40.030.16 ± 0.01<10−40.14 ± 0.01<10−40.30.18 ± 0.01<10−40.14 ± 0.01<10−4<10−4
Figure 2.

Schematic representation of 2D-positioning assessment for chromosome X, Y and 18 centromeres, in normal spermatozoa and spermatozoa with large vacuoles. Centromeres were localized in a plan using their coordinates among the long sperm head axis (D/L) and the short sperm head axis (H/L). Areas of centromere distribution were virtually divided into six (for chromosome 18 and Y) or four (for chromosome X) table-cells, in order to compare chromosome distributions between normal spermatozoa and spermatozoa with large vacuoles, using the chi-squared test within each table-cell. Normal spermatozoa are represented by black points and spermatozoa with large vacuoles by black triangles. D/L: chromosome centromere coordinate along the long axis of the sperm head. H/L: chromosome centromere coordinate along the short axis of the sperm head.

Haploid spermatozoa population

To evaluate aneuploidy influence on chromosome topography, we performed chromosome positioning assessment in the restricted haploid spermatozoa population.

With linear positioning assessment, a non-random distribution in normal haploid spermatozoa (chr X: < 10−4; chr Y: < 10−4; chr 18: < 10−4) and in haploid spermatozoa with large vacuoles (chr X: = 0.02; chr Y: = 0.001; chr 18: = 0.002) was confirmed. As observed in the whole spermatozoa population, comparisons of centromere distributions between normal haploid spermatozoa and haploid spermatozoa with large vacuoles demonstrated no significant difference for chromosome Y (= 0.59) and chromosome 18 (= 0.1); chromosome X topography remained significantly modified between normal haploid spermatozoa and haploid spermatozoa with large vacuoles (= 0.0001).

With 2D positioning, the analysis of chromosome centromeres distribution, restricted to haploid spermatozoa, confirmed the data obtained for the whole spermatozoa population: we demonstrated (i) a non-random topography in normal haploid spermatozoa (chr X: < 10−4; chr Y: < 10−4; chr 18: < 10−4) and in haploid spermatozoa with large vacuoles (chr X:< 10−4; chr Y: < 10−4; chr 18: < 10−4) and (ii) a modified topography in haploid spermatozoa with large vacuoles for chromosome Y (< 10−3), chromosome 18 (< 10−3) and not for chromosome X (= 0.14).

Discussion

To evaluate the impact of sperm vacuoles on chromosome positioning, our study was based on (i) a MSOME selection of defined spermatozoa considered normal or with large vacuoles, (ii) an evaluation of chromosome topography for each type of spermatozoa, and (iii) a comparison of the chromosome distributions between normal spermatozoa and spermatozoa with large vacuoles.

In our MSOME selection methodology, spermatozoa with large vacuoles were isolated when their vacuole area was superior to 13%. In the absence of consensus on ‘large’ vacuole definition, varying from more than 25% (Boitrelle et al., 2011) or 50% of sperm head area (Franco et al., 2008, 2012; Mauri et al., 2010; Oliveira et al., 2010; Akl et al., 2011; Silva et al., 2012) to a diameter of more than 1.5 μm and visible at 400× magnification (Watanabe et al., 2011), this cut-off has been proposed to define abnormal sperm vacuole area, using a strict objective MSOME analysis (Saïdi et al., 2008; Perdrix et al., 2012); this value represented the smallest mean vacuole area observed only in abnormal semen samples. Isolation of spermatozoa with large vacuoles was performed using spermatozoa of selected infertile patients presenting semen alterations, whereas normal spermatozoa were selected from spermatozoa presenting normal semen parameters. Indeed, vacuole area seemed to be correlated with semen alterations (Cassuto et al., 2012; Perdrix et al., 2012); in particular, vacuoles have been supposed to increase when percentage of spermatozoa with normal form decreases (Sermondade et al., 2007; Saïdi et al., 2008; Oliveira et al., 2009; Perdrix et al., 2011, 2012). The number of MSOME selected spermatozoa was limited not only because of time consuming of the technique but also to avoid the potential effects of incubation time on sperm vacuole occurrence (Peer et al., 2007). Consequently, our study was limited to 200 normal spermatozoa and 200 spermatozoa with large vacuoles; this sample size corresponded to previously reported data focused on individual MSOME selected spermatozoa sample size varying from 100 (Garolla et al., 2008) to 350 (Franco et al., 2008), 360 (Cassuto et al., 2012), 400 (Perdrix et al., 2011) and 450 (Boitrelle et al., 2011) for each type of selected spermatozoa.

Assessment of centromere topography was performed using two methodologies. The first one, named linear positioning and based on a simple localization of the FISH signals among the long axis of the sperm head, represented the most instinctive and the most common method used in literature (Luetjens et al., 1999; Hazzouri et al., 2000; Terada et al., 2000; Sbracia et al., 2002; Zalenskaya & Zalensky, 2004; Foster et al., 2005; Mudrak et al., 2005; Olszewska et al., 2008; Wiland et al., 2008). Using this methodology, we observed a preferential localization of chromosome X, Y and 18 centromeres in the median part, in vacuolated and not vacuolated spermatozoa, as described by Olszewska et al. (2008) (medial located 18, X and Y centromeres: 55–65%) or Wiland et al. (2008) (medial located X and Y centromeres: 58–68%). On the contrary, other authors reported a preferential anterior position for chromosome X centromere (Luetjens et al., 1999; Hazzouri et al., 2000; Zalenskaya & Zalensky, 2004) or chromosome Y centromere (Sbracia et al., 2002), and a preferential posterior position for chromosome 18 centromere (Sbracia et al., 2002). Confronted with these contradictions, Olszewska et al. (2008) pointed the imperfections of this method (influence of FISH probes used, of studied sub-regions number in sperm head, of borderline FISH signals) and concluded to the importance of applying a ‘more objective’ model with normalized coordinates. Indeed, the simplicity of the linear positioning allowed to assess several spermatozoa: from 100 spermatozoa per chromosome (Mudrak et al., 2005) to 1000 spermatozoa per chromosome (Sbracia et al., 2002), with most of publications based on an analysis of 200 spermatozoa per chromosome (Luetjens et al., 1999; Zalenskaya & Zalensky, 2004; Olszewska et al., 2008; Wiland et al., 2008). However, this methodology presents some limits: linear positioning, by definition, only provided centromere localization information among one dimension, considering the long axis of sperm head. Three-dimensional analysis was performed to observe chromosome distribution in sperm head, but such studies required a confocal laser scanning microscope (Hazzouri et al., 2000), and the number of assessed spermatozoa was limited (about 30 spermatozoa per chromosome (Hazzouri et al., 2000). Zalenskaya & Zalensky (2004) proposed a semi-quantitative method describing chromosome positioning in two dimensions, and based on a precise measurement of chromosome centromere coordinates along the long and the short axis of sperm head. This method allowed a positioning assessment of a large number of spermatozoa close to 200 spermatozoa per chromosome (Zalenskaya & Zalensky, 2004; Olszewska et al., 2008), and provided information about chromosome topography not only from the acrosome to the tail but also from the periphery to the centre of sperm head. This is the reason why this method has been chosen in our study. To interpret data obtained from multiple measurements of hundreds of spermatozoa, Zalenskaya & Zalensky (2004) used mean values; this statistical strategy, instinctive and very useful, suffered anyway from a little lost of information because of the heterogeneity of the values. Consequently, we proposed in our study, to take into account each centromere of chromosome, individually, defined by both long axis coordinate and short axis coordinate into the sperm head.

Our first results confirmed a non-random localization of chromosome X, Y and 18 centromeres in human sperm head (Haaf & Ward, 1995; Tilgen et al., 2001; Zalenskaya & Zalensky, 2004; Manvelyan et al., 2008), independently of the presence or the absence of vacuoles. Secondly, our study demonstrated the following (i) with linear positioning method, a significant difference for chromosome X distribution between normal spermatozoa and vacuolated spermatozoa, and not for chromosomes 18 and Y and (ii) using 2D method, a marginal difference for chromosome X, and a significant difference for chromosome 18 and Y distributions, between normal spermatozoa and vacuolated spermatozoa. Seemingly contradictory, these results were complementary: linear positioning assessment proved a modification in chromosome distributions among the long axis of sperm head between the two types of spermatozoa, whereas 2D positioning added information about chromosome distributions among the short axis. In our study, X chromosomes varied essentially in the anteroposterior distributions between vacuolated and not vacuolated spermatozoa; on the contrary, chromosomes 18 and Y varied predominantly in their positions in depth. The complementarity of linear and 2D-positioning results have already been demonstrated as following: Olszewska et al. (2008) concluded to an absence of difference in the longitudinal localization of chromosomes 15, 18 and X between control males and infertile patients with increased level of aneuploidy evaluated using linear positioning method, but demonstrated a significant difference using spatial positioning method.

Considering that our study was performed on particular spermatozoa, selected from patients with specific semen profile, it appeared necessary to discuss a confusion bias, potentially implicated in chromosome topography modifications between normal spermatozoa and spermatozoa with large vacuoles. Firstly, semen alterations have been suspected to be associated with sex chromosome position changes (Finch et al., 2008), and abnormal sperm morphology has been specifically supposed to be related with an abnormal nuclear organization (Gurevitch et al., 2001); in our study, nuclear disorganization cannot be dissociated from altered spermatogenesis. However, a recent publication seemed to minimize the impact of altered spermiogenesis on chromosome topography (Ioannou et al., 2011). Secondly, oligozoospermia (Rives et al., 1999; Vegetti et al., 2000; Calogero et al., 2001; Tempest & Griffin, 2004), asthenozoospermia (Vegetti et al., 2000; Tempest & Griffin, 2004) and teratozoospermia (Calogero et al., 2001; Harkonen et al., 2001; Machev et al., 2005) have been related to sperm aneuploidy. Confusingly, sperm aneuploidy has been associated with modification of chromosome localization in sperm heads: Olszewska et al. (2008) demonstrated disturbances in the centromere area in sperm nuclei of infertile patients with an increased level of aneuploidy compared with fertile males; in sperm cells of infertile patients with an increased level of aneuploidy, ‘some differences in the preferential longitudinal centromere positioning between sperm nuclei with hyperhaploidic karyotype (n + 1) vs. normal karyotype (n)’ were observed. Consequently, impact of aneuploidy in our results must be considered (i) aneuploidy level was not significantly increased in vacuolated spermatozoa from OAT patients compared with non-vacuolated spermatozoa from controls and (ii) exclusion of aneuploid spermatozoa did not modified our results, confirming the independency of chromosome topography changes with respect to aneuploidy, as previously observed by Sbracia et al. (2002).

The association between vacuoles and changes in chromosomes positioning observed in our study should be reinforced considering reported common points between these two semen characteristics. Both sperm head vacuoles (Zamboni, 1987; Mundy et al., 1994) and chromosome topology modifications in sperm nucleus (Finch et al., 2008; Wiland et al., 2008) have been described to be the result of a global spermiogenesis alteration. Furthermore, sperm head vacuoles and chromosome topography seemed to have concordant impact on embryonic development: nuclear architecture has been supposed to influence the early embryonic development (Haaf & Ward, 1995; Foster et al., 2005), but also later stages of zygote development; in parallel, sperm vacuoles have been associated with a poorer embryonic quality (Berkovitz et al., 2006b; Vanderzwalmen et al., 2008; Cassuto et al., 2009) essentially because of an effect on late embryonic development (Berkovitz et al., 2006a; Hazout et al., 2006; Vanderzwalmen et al., 2008). Both sperm head vacuoles (Figueira Rde et al., 2011) and chromosome topography (Luetjens et al., 1999; Terada et al., 2000; Zalensky & Zalenskaya, 2007; Olszewska et al., 2008) have been implicated in sex-chromosome aneuploidy in ICSI-derived embryos.

In conclusion, chromosome architecture was modified in spermatozoa with large vacuoles compared with normal spermatozoa. Considering chromosome positioning as a reflect of chromatin integrity, our results emphasize the hypothesis of an association between nuclear vacuoles and sperm chromatin alterations. Consequently, MSOME analysis could be considered more informative than a strict morphological assessment of spermatozoa to improve male infertility diagnosis and treatment. However, further studies are necessary to confirm our preliminary results and to determine mechanism implicated in the association between chromosome disorganization and sperm vacuoles.

Acknowledgements

Anne Perdrix and Albanne Travers performed experiments. Nathalie Rives designed the research study. Anne Perdrix and Florian Clatot analysed the data. Louis Sibert, Valérie Mitchell, Fanny Jumeau and Bertrand Macé recruited patients, participated in design of the study and revised the manuscript critically. Anne Perdrix and Nathalie Rives wrote the article.

The authors wish to thank Julien Dutheil, staff member at Max-Planck Institute for Terrestrial Microbiology, junior researcher in the Institute of Evolutionary Sciences of Montpellier (ISE-M, UMR 5554), who performed statistical analysis.

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