Signification of DNA integrity in sperm of Palaemon serratus (Pennant 1777): Kinetic responses and reproduction impairment

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Signification of DNA integrity in sperm of Palaemon serratus (Pennant 1777): Kinetic responses and reproduction impairment Alexandre Errauda, Marc Bonnardb, Olivier Geffardc, Romain Coulauda, Agnès Poreta, Aurélie Duflota, Joëlle Forget-Leraya, Alain Geffardb, Benoit Xuereba,∗ a

Normandie Univ, UNIHAVRE, UMR-I 02 SEBIO, FR CNRS 3730 SCALE, 76600, Le Havre, France Université Reims Champagne Ardenne, UMR-I 02 SEBIO, 51100, Reims, France c IRSTEA, UR MALY Laboratoire d’écotoxicologie, centre de Lyon-Villeurbanne, F-69616, Villeurbanne, France b

A R T I C LE I N FO

A B S T R A C T

Keywords: Comet assay Crustacean Palaemon serratus Spermatozoa Reproduction success

The study of the effects of contamination on sperm quality not only provides an early, specific and integrative response to the fraction of bioavailable pollutants, but also has been shown to predict the potential of this fraction to modify an organism's capacity to reproduce. In addition, fertility damage in invertebrates has been addressed as a major problem that may pose a threat to the maintenance of populations. In this context, the present study proposes a methodology based on the measurement of sperm DNA integrity to evaluate the impact of paternal damaged DNA on the reproductive success of Palaemon serratus. A preliminary methodological optimization step was carried out to assess the kinetics of response of spermatozoa as well as the sensitivity of the spermatozoa according to their location in the genital tract. Spermatozoa appeared to be sensitive to a short in vivo exposure to the direct acting agent methyl methanesulfonate (i.e. MMS; 2 days), with a persistence of damage even after a 30 days' recovery in a clean environment, suggesting a probable lack of DNA repair machinery. Moreover, our results revealed no difference in the level of DNA damage in mature spermatozoa whatever the exposure in spermatophore located in the terminal ampulla or in the proximal and distal part of the vas deferens. Finally, a significant decrease in the percentage of naturally bred prawns has been observed at the highest concentration of MMS (i.e. 100 μM). Nevertheless, no reproduction impairment (i.e. fertilization rate and early embryo development) following a paternal exposure has been shown in spite of very high levels of sperm DNA damage. In regard to the literature, this result raises questions concerning the kinetics of expression of genotoxic damage on progeny in the Palaemon model and future work will be led in this way.

1. Introduction

invertebrates (Akcha et al., 2004; Lacaze et al., 2010; Lewis and Galloway, 2008, 2009). Although this approach could itself be proposed as an end goal allowing to detect damage from the molecular to the individual level, it does not necessarily account for an impact at higher biological levels (i.e. populations, communities). Indeed, beyond the diagnostic of effects on the health of individuals, it now appears to be more important to understand how these sub-individual effects can be translated into individual and population dynamics, particularly in functionally important species, in order to promote the predictive and also ecological relevance of these indicators (Adams et al., 1989; Baird et al., 2007; Jha, 2008). In this way, the possibility to link biomarker responses to fitness impairment, such as effects on growth, maintenance and reproduction, has been proposed by several groups of authors (Allen and Moore, 2004; Amiard and Amiard-Triquet, 2008). The assessment of the impact of a genotoxic exposure on the

A large diversity of anthropogenic contaminants is constantly discharged into the environment. Aquatic areas are considered to be the ultimate destination of this multi-contamination comprised of up to one third of compounds described as potentially genotoxic for living organisms (Claxton et al., 1998; Ohe et al., 2004). Consequently, the assessment and the maintenance of aquatic ecosystem quality has increased the need to develop robust, integrative early-warning tools as biomarkers to establish cause-effect relationships between genotoxic exposure and health impairment in wildlife (Oost et al., 2003). Nowadays, there are a number of laboratory and field studies demonstrating that an exposure to environmental contaminants induces DNA damage in somatic and germ cells, be it in fish species (Baumgartner et al., 2009b; Santos et al., 2013; Devaux et al., 2011, 2015) or macro-

∗

Corresponding author. E-mail address: [email protected] (B. Xuereb).

https://doi.org/10.1016/j.marenvres.2019.01.005 Received 1 October 2018; Received in revised form 9 January 2019; Accepted 11 January 2019 0141-1136/ © 2019 Elsevier Ltd. All rights reserved.

Please cite this article as: Erraud, A., Marine Environmental Research, https://doi.org/10.1016/j.marenvres.2019.01.005

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reproduction process and its potential repercussions on fitness in wildlife populations can be legitimately addressed as major issues in ecotoxicology (Baumgartner et al., 2009a). Indeed, whether DNA damage in somatic cells can cause mutations resulting in multiple cell proliferation and cancer affecting individual health, damaged-DNA in germ line cells may contribute to hereditary defects through the deleterious mutation transmission leading to teratogenic effects, a recruitment decrease (Mitchelmore and Chipman, 1998) and, in the worst case scenario, an elevated extinction risk for sensitive species (Diekmann et al., 2004a, b). Contrary to oocytes, which contain the cellular machinery to prevent and repair DNA against environmentally induced damage, sperm cells are generally considered to reduce and lose this capacity progressively during spermatogenesis (Aitken et al., 2004). Incidentally, human health research has clearly demonstrated that the male reproductive system is a major target for environmental chemicals and damaged sperm DNA has been adversely involved in the increase of birth defects and childhood diseases (e.g. Aitken and De Iuliis, 2007; Talbot and Chacon, 1981). Concerning wildlife, few studies have investigated this issue. Despite that, clear correlations between spermatozoa DNA damage and impairments in recruitment success (i.e. increase of malformations and mortality within progeny) have been reported in fish (Baumgartner et al., 2009b; Devaux et al., 2011, 2015; Santos et al., 2013), as well as in invertebrates (Lacaze et al., 2011b; Lewis and Galloway, 2009). Consequently, the monitoring of sperm DNA damage in sentinel species presents a great interest in the perspective of an environmental survey due to (1) its potential to integrate the effects of a large range of pollutants and (2) its apparent signification in terms of impact on population recruitment. It is therefore important to improve our knowledge relative to the genotoxicity on spermatic cells and its repercussions on reproduction and development success in key species. Notably, special attention should be paid to groups of crustaceans. Indeed, these organisms adopt a vast array of sperm designs (i.e. cellular morphologies and functionalities depending on the fertilization strategy; Braga et al., 2013) which are very different from the ent-aquasperm commonly found in vertebrates and mollusks, being the most commonly used organisms in biomonitoring. Although crustaceans have received less attention, they have proved relevant for the quality assessment of water bodies, especially in fresh water (e.g. Chaumot et al., 2015). Furthermore, some recent reports highlighted the importance of considering the impact of chemicals on male fertility in crustaceans as a serious concern (review in Lewis and Ford, 2012; Yang et al., 2008). Among marine crustaceans, Palaemon genus have been commonly used as a relevant sentinel species for assessing the health conditions of coastal systems (Bocquene et al., 1995; Frasco et al., 2008; Key et al., 2006). These species are logistically interesting since they can be sampled throughout the year and are relatively easy to identify, to manipulate and to maintain in a laboratory. The Palaemon genus is widespread and common in the coastal and estuarine waters of Western Europe, where they are often found in high density. These prawns are important trophic components for many fish and other crustacean species, including commercially-valuable ones, and play a major part in the detritus breakdown process as primary and secondary consumers (Anderson, 1985). Measurement of sperm DNA damage by Comet assay has been previously optimized in the coastal species Palaemon serratus (Erraud et al., 2017, 2018), highlighting the potential of this tool as a biomarker for exposure to genotoxic insults in marine crustaceans. In order to improve the interpretation of this biomarker, the present study aimed (1) to describe the kinetic occurrence and persistence of DNA damage in spermatozoa submitted to genotoxic stress and (2) to assess the consequences of this damage on the reproduction success parameters of prawns (i.e. fertilization and embryo-development). To achieve this, sperm DNA damage was measured after different times of in vivo exposure of prawns to a concentration gradient of the direct alkylating compound methyl methanesulfonate (MMS), but also after different times and conditions of depuration by the transfer of prawns

into a healthy medium. Next, the effects of sperm DNA damage on the fertilization rate and the embryonic development were explored after cross-breeding of exposed males with unexposed females. 2. Materials and methods 2.1. Sampling and stabulation of prawns Within the common prawn populations from the French coast of the English Chanel, the major peak of reproductive activity occurs from December to March (Campillo, 1979). During this period, the prawns are located in the deeper waters of the subtidal zone. In order to study this species during the reproductive period, adult specimens of P. serratus (i.e. 52.13 ± 11.78 of body size) were collected by a professional fisherman of Le Havre (DYFLO) using specific traps in the 2-nautical mile zone between Octeville-sur-mer and Cauville-sur-mer (Normandy, France). Prawns were brought to the laboratory in 30 L-plastic containers supplied with artificial seawater (ASW; salinity of 33, pH 8.2 and approximate temperatures of the environment). ASW was obtained by dissolution of TETRA®Sea salt (i.e. salt used in marine aquarium maintenance) at a concentration of 40 g.L−1. Sexually mature females and males were immediately separated on the basis of their secondary sexual characters as described in Erraud et al. (2017) and kept in 30 Lplastic containers, under oxygenation until the beginning of the experiment the next morning. 2.2. Kinetic of response of P. serratus spermatozoa For each experiment, prawns were kept in 2 L beakers supplied with 1 L of artificial sea water, under a photoperiod of 12 h light and 12 h dark cycle. Mortality was followed with the daily removal of dead/ moribund prawns during the experiment. Every beaker was provided with additional aeration to maintain optimum oxygenation conditions (i.e. upper to 8 mg.L−1 of dissolved O2), and the water was daily monitored in each control beaker for ammonium (NH4+) and nitrite/ nitrate (NO2-/NO3-) levels which were consistently below detectable levels. Prawns were fed daily ad libitum with pellet B-Peanaeus Grower RCE 1 (Le Gouessant®) according to manufacturer's recommendations during this experiment period. 2.2.1. Sperm cells’ sensitivity after an in vivo exposure to MMS Adult P. serratus specimens (N = 240) were exposed for 2, 4, 7 and 14 days to 0, 4, 20 and 100 μM of MMS to study the dose-/time-response relationships of sperm DNA damage. MMS stock solutions were prepared in ASW (salinity of 33) at concentrations ranging from 4 to 100 mM. Exposure media were obtained by adding 1 mL of the appropriate MMS stock solution to 1 L of uncontaminated ASW. Three replicates of 5 prawns (n = 15 per time/dose conditions) were placed in glass 2 L-beakers filled with 1 L of each test solution and media were renewed daily. At the end of the exposure period, surviving prawns from each condition were sacrificed and spermatozoa were immediately collected for an individual assessment of DNA damage (see section 2.4). 2.2.2. Depuration kinetics of encapsulated spermatozoa in the spermatophore during the exposure Prawns (N = 120) were exposed for 2 days to a range of MMS (i.e. 0, 4, 20 and 100 μM) with 30 specimens per concentration in semi-static conditions (as described in section 2.2.1). At the end of the 2 day-exposure period, 40 prawns were sacrificed (nT0 = 10 per MMS concentration) to assess the initial post-exposure level of sperm DNA damage. The remaining 80 prawns (i.e. 20 per MMS concentration) were transferred into an uncontaminated medium (i.e. 5 specimens in 1 L of ASW). Forty prawns were sacrificed after 15 and 30 days of depuration (nT15 and nT30 = 10 per MMS concentration). The sperm samples were obtained and analyzed as described in section 2.4. 2

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4, 20 and 100 μM) with 30 specimens per concentration in semi-static conditions (as described in section 2.2.1). At the end of the exposure, 40 prawns (n = 10 per MMS concentration) were sacrificed to determine the level of DNA damage at T0. Among the remaining 80 prawns, 60 specimens (i.e. 15 per MMS concentration) were individually distributed in an uncontaminated medium (i.e. 1 specimen per 2 L-beaker supplied with 1 L of ASW) which was slowly aerated to maintain optimum oxygenation conditions (see section 2.1). The remaining 20 prawns (n = 5 per MMS condition) were stocked in ASW (i.e. 5 per 2 Lbeaker) for a renewal in case of mortality during the experiment. Then, 60 females in stage 5 of ovarian maturation were individually placed with males (n = 15 per initial MMS concentrations) for mating in a maximum period of 30 days. Reproduction required the molt of the females before the deposition of spermatophores on the female abdomen. Mortality, molt and oviposition of females were controlled daily. After fertilization, each ovigerous female was marked with a tag for an individual follow-up and placed in a 30 L-tank with constant filtration and aeration. The fertilization rate was assessed for each couple after 24–48 h post-mating. For that, an egg grasp of the first pair of periopods (i.e. at least 100 eggs per female) was carefully extracted using fine forceps and observed using a binocular magnifier (MZ75, Leica Microsystems). Fig. 2A and B shows normal embryo segmentation after 24–48 h post fertilization. The percentage of abnormalities was calculated after counting the number of aborted eyes, undeveloped embryos and undifferentiated embryos in which cells have degenerated with an impaired membrane (Fig. 2C–F). Teratogenicity was monitored after 45 days at the 8th stage of development (Fig. 1). At this stage, the nearly-ready-to-hatch embryo is characterized by the appearance of Ommatidia (Om) around the eyes. In the cephalothoracic region, the antennules, antennae and mandibles are more developed. The abdominal region is organized into five segments, and the latter becomes longer. The cephalothoracic carapace (CC) is formed and covers the heart, and the cephalothoracic appendages (CA). The transparency of the carapace makes it possible to recognize yolk granules in the middle intestine. Some of the different types of malformations in embryos are illustrated in Fig. 2G–J.

2.2.3. Depuration kinetics of the cells of the spermatic line exposed in the genital tract Male prawns (N = 120) were exposed for 2 days to a range of MMS (0, 4, 20 and 100 μM) with 30 specimens per concentration in semistatic conditions (as described in section 2.2.1). At the end of the 2 dayexposure period, spermatophores of all prawns from each condition tested were immediately extracted and 40 sperm suspensions (nT0 = 10 per condition) with cellular survival > 85% were randomly selected to determine the levels of DNA damage at T0. Just after the spermatophore extraction, the 40 prawns were transferred to an uncontaminated medium (i.e. 5 specimens in 1 L of ASW). Spermatophore extractions were de novo performed for all survival specimens again after 15 and 30 days of rearing in the uncontaminated medium. Fifteen days between each extraction were needed while awaiting the formation of a new spermatophore in the terminal ampullae. Forty sperm suspensions (nT15 and nT30 = 10 per conditions) displaying cellular survival > 85% were randomly selected for DNA integrity analysis. The sperm samples were obtained and analyzed as described in section 2.4. 2.3. Effect of sperm DNA damage on fertilization success and early embryo development For this experiment, some basic elements of Palaemonidae prawns’ reproduction are required for a global understanding of this process. During the period before reproduction, oogenesis begins with ovaries that gradually increase in volume related to the thermal conditions. Thus, 5 stages of ovarian development can be easily identified according to the color and the place taken by the ovaries in the cephalothorax: from stage 1 (i.e. filiform ovaries, not visible) to stage 5 (i.e. ovaries that take up the whole cephalothorax and are dark brown) (Richard, 1978). It is important to note that several inter-molt cycles are necessary to bring the ovaries to stage 5. Once the females have mature ovaries (i.e. December–January period), females molt, thus sending a signal of availability for males. Briefly, males come to deposit, after or during mating, a pair of spermatophores on the thelycum of the female. In a period of 18–24 h after mating, the female releases her eggs at the third pair of periopods (i.e. gonopores), taking advantage of a decalcification of the vulval opercula, and mixes everything with the pleopods, allowing fertilization. Then, the eggs are incubated, attached to pleopodal setae until advanced larval stages. The embryonic development is constituted of 9 stages (Müller et al., 2004, Fig. 1). The time of embryonic development is temperature dependent. According to the works of Richard (1978), 56 days are needed to achieve complete development at a temperature of 15 °C (i.e. the laboratory temperature used in this experiment). So, to assess reproduction impairment following paternal exposure, male prawns (N = 120) were exposed for 2 days to a range of MMS (0,

2.4. Procedure of the sperm DNA damage analysis The measurement of DNA damage in the prawns’ spermatozoa was performed according to the methodology described in Erraud et al. (2017). Beforehand, male specimens were weighed and the total cephalothoracic length (LCT) was measured. Spermatophores were extracted by a gentle pressure between the fifth pair of pereiopods allowing the expulsion of spermatophores from the terminal ampullae. Then, spermatophores were transferred into a 1.5 mL-microtube, weighed, immersed with 600 μL of artificial seawater (adjusted to the Fig. 1. Morphological pattern of the embryonic developmental stages of P. serratus: (1) spawning egg: (2) Cleavage; (3) Germinal disc; (4) embryonized Nauplius, (5–6) initial post-nauplius; (7) mid postnauplius; (8) final post-nauplius; (9) pre-hatching embryo. (An) Antennulae, (AS) abdominal segment, (At) antennae, (Bl) blastomeres, (CC) cephalothoracic carapace, (CA) cephalothoracic appendages, (Ch) chorion, (CP) caudal papilla, (GD) germinal disc, (Ey) eye, (OL) optical lobes, (Om) ommatidia, (PnA) post-naupliar appendages, (Te) telson, (YM) yolk mass. Source A. Erraud.

3

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Fig. 2. Malformations of P. serratus embryos, 24–48 h after fertilization (left picture) and 45 days old embryos developed (right picture) in non-contaminated water at 15 °C, (A)–(B): example of normal embryos of Palaemon serratus observed under binocular magnifiers x8. (A) complete formation of 4 quadrants, (B) stage 32 nucleis (C)–(F): example of malformed embryos of Palaemon serratus (C) unfertilizated oocyte, (D) aborted eyes, (E) undeveloped embryos, (F) undifferentiated embryos in which cells have degenerated with impaired membranes, (G) aberrant cleavages and edema, (H) nonspecific malformations, (I) enlarged embryos and impaired membranes and (J) abnormalities at the eyes stage dead. Source A. Erraud.

hemolymphatic osmolality of P. serratus at i.e. 950–1000 mOsmol.kg−1) and ripped by pipetting up and down until the entire laceration of spermatophores. For each specimen tested in the present study, the mortality of sperm-suspension was assessed by a membrane permeability test after mixing cells 1:1 (v/v) with an isotonic Trypan-blue dye solution (i.e. 0.4% w/v). Cellular mortality was performed in KOVA® slides, using a photonic microscope (x400). Mortality measurements were performed on the spermatozoa suspensions of 20 males. To avoid the measurement of DNA damage due to cytotoxic events leading to an over-estimation, a minimum of ten different males displaying a mortality rate (< 15%) were selected for the assessment of DNA damage using Comet assay (n = 10). Prior to the Comet assay, superfrosted microscope slides were first covered with a normal melting point agarose-type I (0.8% w/v) in Ca2+ - Mg2+ free PBS (i.e. 10 mM, pH 7.4) and dried overnight at ambient temperature. After collection of the spermatozoa, 60 μL of the cell suspension (i.e. 106 cells.mL−1) were equally mixed with 60 μL of 1% low-melting-point agarose-type VII in Ca2+ - Mg2+ free PBS (i.e. 10 mM, pH 7.4, 37 °C - 0.5% final agarose concentration), and immediately deposited onto the coated slides and finally covered with a 24 × 60-mm coverslip. From this point, all steps of the Comet assay protocol were performed in the dark with inactinic light to prevent additional DNA damage. Slides were cooled for 10 min at 4 °C on ice for agarose solidification. After removal of the coverslip, slides were placed in a freshly prepared lysing solution (i.e. 2.5 M NaCl, 100 mM Na2EDTA, 10 mM Tris, 1% Triton X-100 and 10% DMSO added immediately before use, pH 10) at 4 °C in the dark for 1 h. After cell lysis, slides were gently placed in a horizontal electrophoresis chamber filled with freshly prepared alkaline solution (i.e. 300 mM NaOH, 1 mM Na2EDTA, pH > 13). DNA was then allowed to unwind for 15 min. DNA Electrophoresis was performed in a standard Comet Assay Tank (i.e. 20 slides; 1050 mL of alkaline solutions; 24 cm × 27 cm x 7 cm (W x L x H)) under 0.83 V cm−1 for 24 min. After electrophoresis, slides were washed in a neutralization buffer (i.e. 0.4 M Tris–HCl, pH 7.5) at 4 °C for 20 min. The last step consisted of a dehydration of slides in absolute ethanol for 15 min. DNA embedded in the slide was stained with 30 μL of a DAPI solution at a concentration of 2 μg mL−1. Slides were blindly observed using an epifluorescence-reversed microscope (Eclipse TE2000-U, Nikon®). The slide scoring was assessed by visual scoring according to the method of Collins (2004) which was demonstrated as trustworthy as image analysis (e.g. Azqueta et al., 2011), and proved to be appropriate to detect DNA damage in sperm from P. serratus (Erraud et al., 2017). A minimum of 150 spermatozoa per slide was counted and

classified into five categories of Comet according to the degree of DNA damage, from 0 (i.e. no tail) to 4 (i.e. almost all DNA in the tail indicating highly damaged DNA). The DNA damage level was expressed in arbitrary units. This arbitrary unit graduated from 0 to 400 was calculated by applying the following formula:

AU =

(0×NO) + (1 × N1) + (2 × N2) + (3 × N3) + (4 × N4) × 100 ∑ comet

with N indicating the number of cells counted on the microscopic slide in each class of DNA damage. 2.5. Statistical analysis Statistical analyses were performed with the R software (R Development Core Team, 2016) using RStudio environment (R Studio Team, 2016). All results are expressed as mean ± standard deviation. For experimental data of viability, DNA damage, fertilization rate and the percentage of embryo abnormalities, normality was assessed by a Shapiro-Wilk test and homoscedasticity of variance by the Bartlett test. Depending on the normality of data, two different statistical approaches were performed: i) non-parametric approach with Kruskal & Wallis rank sum tests followed by Nemenyi post-hoc tests (using PMCMR Rpackage; Pohlert, 2014) if significant differences (p-value < 0,05) and ii) linear modelling approach with ANOVA tests followed by post hoc comparison with Student's t tests in case of significant differences (pvalue < 0,05). 3. Results 3.1. Dose- and time-response kinetic of P. serratus sperm DNA damaging toward MMS in vivo exposure Fig. 3 represents the level of DNA damage measured in spermatozoa of P. serratus prawns after 2, 4, 7 and 14 days of in vivo exposure to 0, 4, 20, 100 μM of MMS. No significant effect of MMS on survival of prawns was observed, with 26.7, 33.3, 23.3 and 26.7% of mortality after 14 days of exposure for exposure to 0, 4, 20 and 100 μM, respectively. Otherwise, all deaths resulted from cannibalism between congeners occurring after the molt. The viability rate of spermatozoa was superior to 85% at all tested conditions despite the severe damage levels observed at the highest concentration (e.g. spermatic viabilities of 93,1 ± 5.1, 90.7 ± 3.4, 91.2 ± 2.9 and 90.6 ± 4.1% after 2, 4, 7 and 14 days of exposure to 100 μM). No significant difference of 4

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Fig. 3. Kinetic induction of sperm DNA damages measured using Comet assay from Palaemon serratus in-vivo exposed (i.e. 2, 4, 7 and 14 days, 12 °C) to a gradient of one model genotoxic stressors (i.e. MMS). Results are shown in boxplot (i.e. the median, the first and the third quartiles, the non-outliers range and the outliers). The mean (red point) was added to boxplots to provide as much information as possible of data set (n = 15). * Denotes statistically significant difference from control. (*p < 0.05; **p < 0.01 and ***p < 0.001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

3.1.1. Remanence of sperm DNA damage by passive depuration Fig. 4 represents the level of spermatozoa DNA damage measured at the end of the 2 day-exposure to MMS (i.e. T0), as well as after 14 and 30 days of depuration by transfer of prawns into clean artificial seawater (i.e. T14 and T30). Slight prawn mortality was observed at the outcome of the experiment, independently of the initial MMS concentration. Only 9 of the initial 120 prawns died due to cannibalism. No effect of the MMS treatment or the experiment duration on the viability rate of spermatozoa were observed (Kruskal & Wallis rank sum test, p > 0.05) with similar mean values of 91.5 ± 3.6%, 89.2 ± 5.2%, 94.0 ± 4.5% at T0, T14 and T30, respectively. The levels of DNA

viability was observed between all pair conditions in this experiment (ANOVA, p = 0.748). Constant levels of sperm DNA damage were observed in the control groups during the 14 days of experiment (i.e. 60.5 ± 11.0, 65.5 ± 9.3, 64.6 ± 13.0 and 55.8 ± 12 AU after 2, 4, 7 and 14 days; ANOVA, p = 0.963). All conditions resulted in a significant increase in DNA damage compared to their respective control (t – test, p < 0.001). On the other hand, the sperm DNA damage of the treated groups displayed positive dose response relationships (Spearman correlation test; p < 10−16; ρ = 0.962, 0.967, 0.968 and 0.967 for 2, 4, 7 and 14 days, respectively).

Fig. 4. Remanance of sperm DNA damages of Palaemon serratus measured using Comet assay after an in vivo exposure to a gradient of one model genotoxic stressors (i.e. MMS 12 °C) during their presence in the terminal ampulae and after a depuration phase (i.e. 14 and 30 days). Results are shown in boxplot (i.e. the median, the first and the third quartiles, the non-outliers range and the outliers). The mean (red point) was added to boxplots to provide as much information as possible of data set (n = 15). * Denotes statistically significant difference from control. (*p < 0.05; **p < 0.01 and ***p < 0.001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.) 5

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treatment or the depuration duration (Kruskal & Wallis rank sum test, p > 0.05), with mean values of 91.5 ± 3.6, 91.3 ± 3.8, 90.0 ± 4.7% for T0, T15 and T30, respectively. The constant level of DNA damage in the control group (i.e. 59.7 ± 12.5, 70.5 ± 11.2 and 65.8 ± 13.4 AU after T0, T15 and T30; ANOVA, p = 0.179) and the levels of damage recorded post-exposure in treated groups (i.e. 59.7 ± 12.5, 109.9 ± 15.2, 221.2 ± 9.6 and 316.8 ± 1.9 AU for 0, 4, 20 and 100 μM of MMS, at T0) were very close to the ones observed during both of the two previous experiments. As for the passive depuration, spermatozoa from the initially treated groups displayed no decrease in DNA damage after the 2 extraction/reformation cycles of spermatophores; but quite the opposite, a significant increase was observed from T0 to T15 (Nemenyi post-hoc test, p = 0.0017, p = 0.0003 and p = 8.2 10−5 for 4, 20 and 100 μM respectively), followed by a stabilization from T15 to T30 (Nemenyi post-hoc test, p = 0.7511, p = 0.9936 and p = 0.7367, for 4, 20 and 100 μM respectively). For example, the level of sperm DNA damage in prawns exposed at 100 μM reached 398.0 ± 1.3 and 396.9 ± 2.1 AU. These results are identical to those obtained during passive depuration and will require new experiments to understand this phenomenon.

damage recorded post-exposure (i.e. 59.7 ± 12.5, 109.9 ± 15.2, 221.2 ± 9.6 and 316.8 ± 1.9 AU for 0, 4, 20 and 100 μM of MMS, at T0) were very close to the ones measured during the previous experiment (see section 3.1) in the same respective conditions. The level of the control group's damage remained constant throughout the experiment (i.e. 59.7 ± 12.5, 69.0 ± 13.9 and 58.3 ± 9.0 AU at T0, T14 and T30, respectively; ANOVA, p = 0.143). After the transfer of prawns to a clean medium, the spermatozoa of treated groups displayed no decrease in DNA damage. On the contrary, a significant increase was measured in the group of prawns treated at the concentration of 4 μM of MMS, from T0 to T30, with a sperm DNA damage level of 109.9 ± 15.2, 140.1 ± 24.7 and 198.4 ± 13.5 AU, respectively (Nemenyi post-hoc test p = 0.0004). For the groups treated at 20 and 100 μM, a significant increase in the level of DNA damage was observed at T14 (Nemenyi post-hoc test, p = 0.0024 and p = 0.0004, respectively) followed by stabilization at T30 (Nemenyi post-hoc test, p = 0.071 and p = 0.991, respectively). This increase could be attributed to a phenomenon of inertia of the genotoxic impregnation induced by a peak of exposure to MMS. In any case, this observation further supports the relevance of these biomarkers demonstrating a repercussion of the damage over time. Future experiments can be conducted to understand this phenomenon. At T14 and T30, DNA damage reached 310.2 ± 13.7 and 315.9 ± 6.5 AU in the groups treated at 20 μM, and maximum values of 392.3 ± 4.8 and 392.9 ± 2.3 AU, at 100 μM.

3.2. Impact on fertilization and embryonic development The analyses of spermatozoa viability and DNA damage after the initial exposure of 2 days to the MMS gradient produced results very similar to the ones obtained during the two previous experiments in the respective conditions (i.e. 90.4 ± 2.6, 90.4 ± 1.9, 88.9 ± 2.6 and 91.5 ± 4.0% of viability, and 58.7 ± 6.0, 106.9 ± 14.1, 216.1 ± 10.7 and 322.3 ± 5.0 AU, for 0, 4, 20 and 100 μM, respectively). Table 1 illustrates the repeatability of the sperm DNA damage observed during the three different experiments, despite the fact that these exposures were performed on independent pools of specimen sampling during two different years (i.e. Kinetic response in 2016, and remanence of sperm DNA damage and reproduction impairment in 2107). It can be noted that the levels observed into the control conditions were similar (Wilkoxon test, p > 0.05, mean variation coefficient 0.1%). Concerning the treatment conditions, the results recorded

3.1.2. Remanence of sperm DNA damage by active depuration Fig. 5 represents the level of spermatozoa DNA damage measured at the end of the 2 day-exposure to MMS (i.e. T0), then after a forced depuration by the transfer of prawns into clean artificial seawater associated with 2 fifteen-day cycles of spermatophore extraction/reformation (i.e. T15 and T30). Prawn mortalities of 26.7, 10.0, 13.3 and 13.3% (i.e. 8, 3, 4 and 4 dead specimens) were observed for control, 4, 20 and 100 μM, respectively, during the experiment. The majority of deaths occurred either immediately after a few days after extraction or, to a minor extent, by cannibalism post-molting. No effect of the initial MMS treatment on prawn survival was characterized. The viability rate of spermatozoa remained rigorously close regardless of the initial MMS

Fig. 5. Remanence of sperm DNA damages of Palaemon serratus measured using Comet assay after an in vivo exposure to a gradient of one model genotoxic stressors (i.e. MMS 12 °C) during their presence in the spermatophore and after a depuration phase (i.e. 15 and 30 days). Results are shown in boxplot (i.e. the median, the first and the third quartiles, the non-outliers range and the outliers). The mean (red point) was added to boxplots to provide as much information as possible of data set (n = 15). * Denotes statistically significant difference from control. (*p < 0.05; **p < 0.01 and ***p < 0.001). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.) 6

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Table 1 Repeatability of sperm DNA integrity measured by the Comet assay in Palaemon serratus exposed 2 days-in vivo to a gradient of a well-known genotoxicant (i.e. MMS). This table is a compilation of the data obtained during 3 independent experiments (i.e. kinetics of response; reproduction impairment; recovery). The results are expressed in mean ± standard deviation. (a) and (b) denotes significant difference between conditions. Experimentations

Kinetic response Recovery Reproduction impairment

MMS concentrations (μM) 0

4

20

100

60.5 ± 11.1 (a) 58.2 ± 12.8 (a) 58.7 ± 5.9 (a)

90.6 ± 13.3 (a) 109.9 ± 15.2 (b) 106.9 ± 14.1 (b)

181.2 ± 21.4 (a) 221.1 ± 9.6 (b) 216.1 ± 10.7 (b)

287.6 ± 11.4 (a) 317.1 ± 1.7 (b) 322.3 ± 5.7 (b)

during the first experiment (i.e. Kinetic response) were significantly lower (Wilkoxon test, p < 0.05). However, the levels of DNA damage stayed closed as illustrated by the low variation coeffficients (i.e. 9.6, 9.9 and 5.4% respectively for the treatments with 4, 20 and 100 μM of MMS). The mating step (i.e. couples of pre-exposed male and healthy females, individually kept in uncontaminated seawater until spawning) lasted 5.6 ± 4.7, 6.6 ± 6.7, 7.0 ± 6.8 and 12.2 ± 9.6 days in means for 0, 4, 20 and 100 μM within the 30 day-experiment duration. No significant effect of the initial MMS treatment was observed (Kruskal & Wallis rank sum test, p = 0.3486). During this period, mortality was observed among the males (i.e. 30, 25, 45 and 35% of the 20 males pretreated with 0, 4, 20 or 100 μM of MMS) and the females (i.e. 30% for each group of males pre-treated). Females needing no pre-exposure, the dead ones were immediately replaced. In fine, 12, 10, 9 and 11 couples were followed for each condition of male pre-treatment. Only, 11, 8, 7 and 8 of these couples led to a spawning, and 1, 2, 2 and 3 females displayed no fertilized oocytes. Within the fertilized females (n = 11, 8, 7 and 8), the rate of dividing embryos after 24–48 h was not affected by the pre-treatment of males with MMS (i.e. 95.2 ± 3.8, 93.8 ± 1.9, 94.2 ± 6.1 and 93.5 ± 2.7% for 0, 4, 20 and 100 μM; Kruskal & Wallis rank sum test, p = 0.533; Fig. 6). Lastly, only 6, 4, 4 and 6 specimens among these fertilized females reached completion of the 45 days of embryonic development (the rest of the females lost their eggs during the 45 days of incubation). No significant effect of a paternal damaged DNA on the embryonic development was observed (i.e. 0.7 ± 0.2, 0.8 ± 0.1, 0.8 ± 0.1 and 1.0 ± 0.4% of abnormal embryos in females mating with pre-exposed males to 0, 4, 20 and 100 μM; (Kruskal & Wallis rank sum test, p = 0.543; Fig. 7).

4. Discussion 4.1. Kinetic responses of the sperm DNA damage measurement in P. serratus Considering that DNA integrity results from both genotoxic pressure and DNA repair, the level of DNA damage depends on (1) the dose and time of exposure to a genotoxic element, (2) the sensitivity of cells depending on their maturation stage and (3) the DNA repair capabilities. Consequently, the understanding of each of these three aspects constitutes an essential prerequisite to rigorously conduct our study. Due to the lack of knowledge relative to genotoxic effects on spermatozoa of palaemonids, a preliminary phase consisted in describing the kinetic response of the studied biomarker in terms of replicability, sensitivity and effective period. More globally, this knowledge is needed for the interpretation of Comet assay on palaemonid sperm, in the perspective of a deployment as a biomarker in biomonitoring surveys. The first experiment of MMS exposure demonstrated that mature spermatozoa of the marine crustacean P. serratus are quickly affected by the direct acting genotoxicant. Indeed, MMS led to significant effects from the lowest tested time and dose (i.e. 2 days of in vivo exposure at 4 μM). The severity of damage was positively correlated with the time and dose of exposure to reach a maximum value of 288 ± 11 A.U. (on a scale of 400). Despite these extreme values, no evidence of spermatozoa mortality was highlighted, the viability rate remaining higher than 93% at 100 μM. Although, it is quite difficult to make a rigorous inter-study comparison due to differences related to the time of MMS exposure, the sensitivity of the biological model, the comet assay protocol or the parameter used for expressing DNA damage, the effects of in vivo exposure to MMS have already been demonstrated in several aquatic invertebrates. In the same range of concentrations of MMS, Lacaze et al. (2010) showed in the amphipod Gammarus fossarum, significant increases in sperm DNA damage from 5 days of exposure at

Fig. 6. Fertilization rate of prawns Palaemon serratus after a 2 day-in vivo exposure of males to a gradient of a well-known genotoxicant (i.e. MMS). Results are shown in boxplot (i.e. the median, the first and the third quartiles, the non-outliers range and the outliers). The mean (red point) was added to boxplots to provide as much information as possible of data set (n = 5). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.) 7

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Fig. 7. Percentage of embryo abnormalities in the pre-hatching stage of prawns Palaemon serratus previously fertilized with sperm suspension from 2 day-in vivo exposed male prawns to a gradient of a well-known genotoxicant (i.e. MMS) and after 45 days of development on the abdomen on the females at 15 °C. Results are shown in boxplot (i.e. the median, the first and the third quartiles, the non-outliers range and the outliers). The mean (red point) was added to boxplots to provide as much information as possible of data set (n = 5). (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

4 μM, and recorded high DNA damage with around 75% of tail DNA at the highest tested concentration of 100 μM. On another concentration scale, the in vivo exposure of the mussel Mytilus edulis for 72 h to 47.2 mM of MMS induced less DNA damage, with 20% of tail DNA measured in sperm (Lewis and Galloway, 2009). In the present study, spermatozoa of the common prawn P. serratus appeared to be a sensitive cellular type with regard to the genotoxic aggression by MMS. Of course, a controlled exposure to MMS remains a genotoxic model and does not transcribe the environmental complexity. This has, however, provided the prerequisites necessary to calmly pursue the experimental approach. In view of the results obtained in this first step, further investigations were performed on the basis of a 2 day-exposure since this short period appeared adequate to study 1) realistic environmental damage levels at the concentration of 4 and 20 μM (Erraud et al., 2018) and 2) an extreme situation at 100 μM. All of these exposures, conducted according to an identical procedure on differents pools of organisms, attested the large degree of repeatability of the biological response (Table 1). Compared with somatic cells and oocytes, spermatozoa are in most cases considered to be lacking in the ability to repair DNA damage (Aitken et al., 2004). This can be explained by a reduced cellular body and a condensed nucleus, which result in a cellular metabolism limited to functions crucial for fertilization success. However, contrary to the majority of taxa currently used in ecotoxicology, the Palaemonids have spermatozoa provided with a large, decondensed nucleus (Braga et al., 2013), which justifies taking an interest in DNA remanence of mature spermatozoa. Otherwise, different spermatogenetic stages, displaying variable degrees of repair ability, could be present in the genital tract (i.e. testis and/or vas deferens). In addition, these cells in the tract (i.e. mature or not) are not embedded in the spermatophore matrix and could consequently be submitted to more marked genotoxic pressure in comparison with the spermatozoa stocked in ejaculatory bulbs. In the

present work, we have thus investigated the effective period of DNA damage in spermatozoa of P. serratus, in trying to integrate the different aspects previously cited. The aim was to ensure that the alterations measured after MMS-exposure were representative of those transmitted during fertilization, even in the case where a delay is observed between the end of exposure and mating. So, when prawns were only placed in a healthy medium after MMS exposure (i.e. without forced spermatophore renewal), no decrease of the DNA damage levels was observed even after 30 days of recovery, whatever the MMS treatment initially applied (Fig. 4). This result would suggest that the mature spermatozoa embedded in spermatophore were incapable of any DNA repair ability. Our observations are in agreement with those reported in the amphipod G. fossarum after 4 days of recovery post MMS-exposure (i.e. 5 days) (Lacaze et al., 2011a). Conversely, Lewis & Galloway (2009) observed a partial reversibility (i.e. 37%) of DNA damage in spermatozoa of the mussel Mytilus edulis after only 72 h post exposure. However, it is notable in the present study that the level of damage remained constant for 30 days, without cellular viability being altered. That suggests, in natura, the spermatozoa of prawns could integrate the genotoxic information for more than one month. In the second place, when the formation of new spermatophores was forced by manual extraction just after MMS exposure, then after 15 days of recovery in a healthy medium, no decrease in the DNA damage levels was observed either (whatever the initial MMS treatment). The levels of damage were similar to those recorded in the previous experiment of passive recovery. These results showed that the spermatophore matrix of palaemonids does not protect against genotoxic insults. They also suggested that the germ cells in genital tracts were sensitive to the genotoxic insult of MMS and led to permanent DNA damage as measured in mature spermatozoa. These germ cells would appear as deficient in DNA repair as mature spermatozoa. Lacaze et al. (2011a) reported in their study that the differences of sensitivity between the different stages of 8

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an in vivo exposure of MMS where spermatozoa present maximum levels. However, it is important to note that females began to present molting difficulties after 20 days in the reproduction period in a 2Lbeaker, with an increase in new appendix malformation and a bad calcification of pleopods and periopods, which stay trapped in the older exoskeleton. This observation was already noted by Patrois (1974) and has been allocated to dietary deficiencies. Although the food was adapted for Peneidae aquaculture, it would have been preferable to use food which the common prawn naturally eats in the sea. During this experiment, in control conditions as with males previously exposed to MMS concentrations, almost all eggs were fertilized (i.e. fertilization rate ranged from 93.5 to 95.1%) and the number of embryos recorded was the same. This lack of contamination effect on the fertilization was already noted in several species such as the marine molluscs Mytilus edulis and Crassostrea gigas (Zhou et al., 2006) and the polychaete Arenicola marina (Lewis and Galloway, 2009) as well as in the freshwater crustacean Gammarus fossarum (Lacaze et al., 2011b) and certain fish species (Devaux et al., 2011, 2015; Santos et al., 2013, 2014; Zhou et al., 2006). This lack of effect is not unexpected since in the majority of species, zygotic gene transcription is activated from the blastula stage until the end of the embryogenesis. Indeed, oocytes contain a stock of maternal mRNAs which take charge of embryogenesis until the blastula stage. In many species, the blastula stage is attained during the first 24 h post fertilization which is not the case for P. serratus prawns in our laboratory conditions, with a stage comprised between the complete formation of four quadrants (i.e. blastomeres) and the 32 nucleis stage before the completion of the segmentation (i.e. blastula). After 45 days of embryo development, only a few females attained the end of development with 54.5%, 50%, 57.1% for control, 4, and 20 μM of MMS, respectively. However, 75% of females reached the end of development for 100 μM of MMS (i.e. 6 females of the 8 remaining). The majority of egg loss occurred independently of the different conditions during the fluttering of pleopods. This observation was already noted in Macrobrachium nobili and attributed as a consequent abrasion of eggs during the ventilator movements of the pleopods and an erosion of individual territories (Balasundaram and Pandian, 1982). The results obtained in the present study are the first reporting of no significant effect on the embryonic development after fertilization with damaged sperm DNA (Table 2). Indeed, Lewis and Galloway (2009) reported teratogenic effects following paternal exposure to MMS, after only 24 h of embryonic development, in Mytilus edulis and Arenicola marina. The authors observed significant effects from 10 to 30% of Tail DNA, respectively for the two species. Although these levels of DNA damage may appear to be relatively average, the MMS concentrations tested to reach them were in a range of 163–472 times higher than those tested in this experiment, for in vivo exposure times of 24 and 72 h. In exposure conditions closer to those used in our study, Lacaze et al. (2011b) investigated, in Gammarus fossarum, the occurrence of prehatch embryo-larval abnormalities after 21 days of development, after 5-day paternal exposure to a MMS gradient. The authors detected a significant effect (i.e. 30% of defects) for an exposure dose of 20 μM. According to their model, they estimated an effective threshold of 20%

spermatogenesis in G. fossarum could exist depending on whether cells have been exposed during their differentiation or in a mature stage. In fact, in the present case, it could be suspected that all cells analyzed using Comet assay were exposed to MMS at a late stage of spermatogenetic maturation. Indeed, Campillo. (1979) observed within the P. serratus population of Roscoff (Brittany; France), during the reproduction period (i.e. from December), that the vas deferens of males was full of mature spermatozoa. At the same time, in crustaceans, it is acknowledged that the main steps of spermatogenesis occur before the cells reach the test lumen, even if some cellular properties seem to be established later (i.e. in vas deferens or post mating), such as capacitation (Subramoniam, 2016). Further studies aiming to compare the repair capabilities of different spermatogenetic stages, could thus consist in performing the same experiment of active depuration during the reproductive latency period. The vas deferens being empty during this period (Campillo, 1979), the reconstitution of spermatophores after forced extractions would involve the production of new spermatozoa from earlier spermatogenetic stages. In fine, this first study of the kinetic response has made it possible to validate the experimental condition needed to investigate the consequences of an altered paternal DNA on reproductive success. From a more environmental point of view, these results suggest that a male prawn, which was exposed to a genotoxic pressure (even episodic), keeps the scar within its spermatoza for more than one month, and this altered sperm may be used during several matings during the reproductive period as shown by Campillo. (1979) or by Balasundaram and Pandian (1982). 4.2. Impact on offspring (fertilization and embryonic development) In the present study, the impact of damaged sperm DNA on the reproduction process was assessed by natural reproduction in laboratory conditions. In control conditions, 92% of couples naturally bred (i.e. 11 couples out of the 12 remaining), highlighting a good methodological approach for the reproduction process in laboratory conditions. In exposed conditions to MMS concentrations, the number of natural breeding-couples significantly decreased reaching 80%, 78% and 73% for 4 μM, 20 μM and 100 μM, respectively. To our knowledge, this work is the first report of reproduction performed in laboratory conditions with Palaemon serratus and more globally in Palaemon genus, which makes it particularly difficult to interpret these results. However, we can hypothesize that this decrease in the reproduction process may be attributed to some poor global physiological conditions of male prawns. Indeed, the reproduction process requires a significant amount of energy and good physiological conditions. During the mating of Palaemon genus, male pleopods beat vigorously while the female turns over. The male genital pores do not reach the female sternum so it is a complex method of reproduction demanding a good synchronization of the male's pleopods of to knock its spermatophore against the female sternum (Chow et al., 1982). Moreover, even if the prawns' global physiological conditions cannot be determined from these experiments and require further investigations, it is easy to suppose that other cells indispensable to the prawns' metabolism did not escape unscathed after

Table 2 Bibliography relative to the effect of paternal DNA damaged by in vivo laboratory or natural exposure on the development of offspring in aquatic species.+ = Effect; - = No effect; N/A = Not available). Type of development

Species

Fertilization

Embryonic development

Larval development

Direct Indirect Indirect Indirect Indirect Indirect Indirect

Gammarus fossarum Arenicola marina Mytilus edulis Chondrostomas nasus Gasterosteus aculeatus Salmo trutta Salvelinus alpinus

– – – – – – –

+ + + N/A N/A – –

N/A N/A + + + N/A

9

Reference Lacaze et al. (2011b) Lewis & Galloway (2009) Devaux et al. (2015) Santos et al., (2013) (2014) Devaux et al. (2011) present study

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of Tail DNA. In fish, which have been the focus of most studies in recent years, the trend is not quite the same. Devaux et al., (2011), in brown trout (Salmo trutta) males and of Arctic char (Salvelinus alpinus) exposed to MMS by intraperitoneal injection (i.e. 50 mg/kg dry weight) a very slight increase in embryonic abnormalities hatching was recorded. Their observations were finally quite comparable to the results obtained in the present study (except its significance), since the anomaly rates went from 0.08 ± 0.17 for controls to 0.61 ± 0.64 for brown trout treated, and from 0.85 ± 0.65% to 1.36 ± 0.89% in Arctic charr. On the other hand, if damage to the paternal DNA did not influence either the fertilization or the survival rate of the embryos up to the hatching stage, it subsequently led to a wide range of abnormalities in the larvae. For example, the authors report a frequency of 20% skeletal abnormalities in larvae from trout treated with MMS against 2% in controls. Similarly, they observe 5 times more cumulative mortality after two months of growth, in the treated lots. Santos et al. (2013, 2014) also demonstrated a significant impact of paternal exposure to MMS on posthatching larval development of the Gasterosteus aculeatus stickleback. In contrast, the authors do not refer to embryonic pre-hatch development. To finish in a more environmental context, Devaux et al. (2015) artificially fertilized oocytes of healthy female fish Chondrostoma nasus, with male seeds taken from polluted sites. The authors observed a positive correlation between damage to sperm DNA and the occurrence of post-hatch larval anomalies, but do not refer to pre-hatch development. Some hypotheses can be made to explain our results. The first hypothesis is that the damaged DNA of sperm can be repaired inside the egg cytoplasm by repair mechanisms just between the entry of spermatozoa into the cytoplasm of the eggs and the beginning of the phase S (Kopeika et al., 2004). This hypothesis has already been proposed by Ciereszko et al. (2005) after an exposure to UV radiation and hydrogen peroxide of sperm of sea lamprey (Petromyzon marinus). This group of authors observed a significant increase in DNA damage after sperm exposure but the repair of oxidative DNA damage by the oocyte after fertilization. This observation was already shown in mice to investigate their capacity to repair damaged spermatozoa and has been attributed to depend on several factors: the type of DNA damage; the percentage of the sperm DNA affected and the quality of the oocyte (Ashwood-Smrth and Edwards, 1996; Matsuda and Tobari, 1988; Sakkas et al., 1998; Sakkas and Alvarez, 2010). Ahmadi and Ng (1999) showed that the mouse oocyte has the capacity to repair DNA damage of sperm but only when it is damaged less than 8%. In our studies, this hypothesis is difficult to apply because of the maximum level of DNA damage measured (AU ≈ 400), indicating that the totality of spermatozoa was severely affected. The second hypothesis is that the embryo abnormalities occurring after fertilization with damaged sperm DNA are not perceptible with simply a visual determination. For that, in future works, the impact of a damaged sperm DNA could be studied by a more detailed detection, for example studying chromosome aberrations or the level of damaged DNA in progeny. The last hypothesis is that parental DNA expression is not equivalent between species. Indeed, all the studies presented previously agree that DNA damage does not alter the fertilization capacity of spermatozoa but leads to significant abnormalities of development. However, the dynamics of occurrence of deleterious effects during development seem completely different depending on the species (Table 2). In addition, development should not be considered with respect to the hatching or parturition event but rather on the basis of transitions in the embryonic, larval and juvenile phases. It can thus be considered for a direct-development organism, such as the gammare that hatches as a juvenile, in which the larval development takes place in the egg, unlike the fish species presented above, in the mussel or prawn. Palaemonidae hatch as larvae and will undergo one or more metamorphoses to reach the juvenile state. In the case of our shrimp study, developmental alterations were sought during the embryonic development phase. Therefore, it is perfectly conceivable that the effects of a damaged paternal DNA may be expressed later during larval development as was the case in trout (Devaux et al., 2011). In mice,

recent studies demonstrated that more than 1000 genes in progeny can be reduced to a total silence and this phenomenon differs according to the stage of development. In the embryo, maternal alleles are expressed but in the adult, these are the paternal alleles and this, whatever the sex of the animal. It can be assumed that the footprint is a consequence of a parental conflict over the allocation of resources to the offspring (Gregg et al., 2010). This seems all the more relevant as the organisms that have exhibited the earliest damage are short-lived organisms (i.e. Mytilus edulis or Arenicola marina). In Palaemon serratus, the reproductive cycle is relatively long and depends on the temperature. Indeed, during this experimentation 45 days at 15 °C were necessary to obtain embryo stage 8. To reach the juvenile stage, the embryos should have passed through an additional stage of embryogenesis followed by an outbreak stage Zoe I, and still undergo a succession of metamorphoses. In fine, the predictive character of the application of the Comet on spermatozoa of Palaemon sp. could not be established. Although the impact of DNA damage on offspring may be suspected, we are unable to describe an effective threshold. Future investigations should be conducted to assess whether deleterious effects can be observed at later stages of development (i.e. larval and juvenile). 5. Conclusion The present study addressed for the first time the possibility of using the Comet assay to assess spermatozoa integrity in a palaemonid prawn and more globally in marine crustaceans in order to diagnose reproduction impairment in progeny. The procedure described here makes it possible (i) to emphasize the sensitivity of Palaemon sp spermatozoa after a short-term exposure to a fairly low concentration of the direct genotoxicant MMS, (ii) to demonstrate the high repeatability of the sperm DNA integrity, (iii) to ensure the level of paternal DNA damage transmitted by natural reproduction after active and passive depuration experiments. Our results showed that mature spermatozoa embedded in spermatophore can be quickly affected by a range of concentrations of MMS. Moreover, no repair capability in the decondensed nucleus of spermatozoa of prawns was shown. This methodological optimization was an inescapable first step to assess the reproduction impairment of progeny following a paternal exposure to a direct acting agent (MMS). Our results do not provide correlations between the induction of DNA strand breaks in sperm and consequences on the fertilization rate and abnormalities in embryos. However, an effect on the success of reproduction was highlighted and further research could be led to explain this phenomenon. To our knowledge, this is the first report of an absence effect of a paternal genotoxicant exposure on embryo-development abnormalities. Obviously, further work could also be addressed to improve our understanding of consequences of DNA damage in sperm on the reproductive potential of male prawns on the last stage of development ranging from hatching rate to delay in juvenile growth. Acknowledgement This study was supported by project ECOTONES funded by the program Seine-Aval V (Public Interest Groups Seine-Aval), the Research Federation CNRS 3730 SCALE and the Normandie Region. The authors also thank Mrs. D. Hallidy and Mrs. A. Le Saux for proof-reading the English. References Adams, S.M., Shepard, K.L., Greeley, M.S., Jimenez, B.D., Ryon, M.G., Shugart, L.R., McCarthy, J.F., Hinton, D.E., 1989. The use of bioindicators for assessing the effects of pollutant stress on fish. Mar. Environ. Res. 28, 459–464. Ahmadi, A., Ng, S.C., 1999. Fertilizing ability of DNA‐damaged spermatozoa. J. Exp. Zool. 284 (6), 696–704. Aitken, R., De Iuliis, G., 2007. Origins and consequences of DNA damage in male germ cells. Reprod. Biomed. Online 14, 727–733.

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