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Amphibian sperm DNA fragmentation Kate Pollock

Bachelor of Applied Science (Animal Studies)

Bachelor of Applied Science (Honours Class 1)

A thesis submitted for the degree of Master of Philosophy at

The University of Queensland in 2015

School of Agriculture and Food Sciences

i

Abstract

There are extremely limited studies that have investigated DNA fragmentation in

amphibians and even less information exists pertaining to DNA fragmentation in the spermatozoa.

A DNA fragmentation assay applied and validated for amphibian spermatozoa would not only

provide the first evidence of this phenomenon in a new major taxon, but also prove a valuable tool

for developing improvements in ART used in the captive propagation of threatened and endangered

species. Consequently, the fundamental aim of this project was the development and validation of

the sperm chromatin dispersion (SCD) test for amphibian spermatozoa with the purpose of

investigating the dynamic behaviour of amphibian sperm DNA in response to cryopreservation.

The SCD test was first applied to African clawed frog (Xenopus laevis) as an amphibian

sperm model using a species-specific modified lysing solution for protein depletion. Sperm DNA

fragmentation (SDF) was assessed immediately following activation of sperm motility (T0) and

again after one hour and 24 hours of incubation in order to produce a range of spermatozoa with

differing levels of DNA damage. The SCD procedure resulted in the production of three nuclear

morphotypes; amphibian sperm morphotype 1 (ASM-1) and ASM-2 showed no evidence of DNA

damage, whereas ASM-3 spermatozoa were highly fragmented with large halos of dispersed DNA

fragments and a reduced nuclear core. Levels of SDF revealed by the SCD test were highly

correlated with results produced using in situ nick hybridisation with DNA-specific molecular

probes (ISNT) and the double comet assay (r = 0.9613). The alkaline step of the double-comet

assay revealed the extensive presence of structural single-stranded DNA breaks (SSB) and provided

grounds for the suggestion that alkali labile sites (ALS) are a constitutive feature of African clawed

frog sperm chromatin.

SDF is a dynamic process and has been shown to increase with incubation at room

temperature as well as following cryopreservation. However, the mechanisms of DNA cryoinjury

and the resulting effect on the potential reproductive output of ART remain open to interpretation.

Consequently, dynamic changes to the basal level of DNA fragmentation were examined in fresh

African clawed frog spermatozoa in comparison with the post-thaw integrity of cryopreserved (-80

°C) and frozen (-20 °C) sperm DNA. SDF was examined initially at T0 and over incubation of up to

60 minutes. The difference in SDF was only marginal for fresh, frozen (-20 °C) and cryopreserved

(-80 °C) spermatozoa examined at the onset of the experiment immediately upon thawing, however,

a significant increase (p = 0.004) in SDF, albeit of a low magnitude, was observed in activated

ii

cryopreserved (-80 °C) spermatozoa following incubation. Although African clawed frog

spermatozoa appeared to be highly resistant to cryopreservation-induced DNA damage,

cryopreserved-thawed spermatozoa yielded a significantly lower (p = 0.0065) fertilisation rate than

fresh spermatozoa.

In order to provide further insight into cryoinjury of amphibian sperm chromatin, the

dynamic loss of DNA integrity in fresh and cryopreserved cane toad (Bufo marinus) spermatozoa

was correlated with changes in sperm viability and motility. Analysis of SDF was conducted at T0

immediately following activation and/or thawing and again following T3 and T6 hours of

incubation at room temperature. Sperm motility was activated by decreasing the osmolality of the

sperm extender to < 50 mOsmol kg-1 with dH20. Sperm viability was assessed using a dual emission

technique that incorporated the nucleic stains SYBR-14 and propidium iodide. Spermatozoa were

classified as either live (green fluorescence) or dead (red fluorescence). Dead spermatozoa were

further classified into two categories; dead-compact, where spermatozoa retained a normal

elongated, rod-like morphology and dead-swollen, where spermatozoa appeared enlarged with less

defined morphology. Assessment of SDF at the population level in fresh spermatozoa, as well as the

dynamics of sperm chromatin longevity, revealed that cane toad sperm DNA was highly stable

throughout the duration of the incubation period. Results demonstrated a surprisingly poor recovery

of both motility and viability in cane toad spermatozoa following freeze-thawing and a significant

increase in the proportion of spermatozoa showing nuclear swelling (p < 0.0001). The incidence of

dead-swollen spermatozoa observed in the cane toad was interpreted to represent the transition from

ASM-1 to ASM-2 described in African clawed frog spermatozoa and lent support to the hypothesis

that ALS may exist in the sperm chromatin of both amphibians.

i

Declaration by author

This thesis is composed of my original work, and contains no material previously published

or written by another person except where due reference has been made in the text. I have clearly

stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical

assistance, survey design, data analysis, significant technical procedures, professional editorial

advice, and any other original research work used or reported in my thesis. The content of my thesis

is the result of work I have carried out since the commencement of my research higher degree

candidature and does not include a substantial part of work that has been submitted to qualify for

the award of any other degree or diploma in any university or other tertiary institution. I have

clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

I acknowledge that an electronic copy of my thesis must be lodged with the University

Library and, subject to the policy and procedures of The University of Queensland, the thesis be

made available for research and study in accordance with the Copyright Act 1968 unless a period of

embargo has been approved by the Dean of the Graduate School.

I acknowledge that copyright of all material contained in my thesis resides with the

copyright holder(s) of that material. Where appropriate I have obtained copyright permission from

the copyright holder to reproduce material in this thesis.

ii

Publications during candidature

Pollock, K., Gosálvez, J., Arroyo, F., López-Fernández, C., Guille, M., Noble, A., and Johnston,

S.D. (2015). Validation of the sperm chromatin dispersion (SCD) test in the Amphibian Xenopus

laevis using in situ nick translation and comet assay. Reproduction, Fertility and Development 27,

1168-1174 – Incorporated as Chapter 2.

Publications included in this thesis




1168-1174 – Incorporated as Chapter 2.

Contributor Statement of contribution

Ms Kate Pollock (Candidate) Designed the experiments (70%)

Conducted the experiments (50%)

Analysed and presented the data (70%)

Wrote and edited the paper (70%)

Prof Jaime Gosálvez Designed the experiments (20%)



Ms Francisca (Paqui ) Arroyo Conducted the experiments (40%)

Advised on experimental design (30%)

Dr Carmen López-Fernández Advised on experimental design (60%)

Prof Matt Guille Advised on experimental design (10%)

Dr Anna Noble Conducted the experiments (10%)

Assoc Prof Steve Johnston Designed the experiments (10%)



iii

Contributions by others to the thesis

Assoc Prof Steve Johnston (University of Queensland, School of Agriculture and Food

Sciences; UQ-SAFS) and Prof Jaime Gosálvez (Autonomous University of Madrid, Genetics

Department; AUM) assisted with the conception of hypotheses presented in this thesis,

experimental design and manuscript revisions. Dr Richard Wilson (University of Queensland,

Mathematics Department) assisted with the statistical analyses and interpretation of the research

data reported in Chapters 3 and 4. Ms Francisca (Paqui) Arroyo (AUM) and Dr Anna Noble

(European Xenopus Resource Centre, Portsmouth, UK; EXRC) provided technical advice and

assistance with experiments presented in Chapter 2 and/or 3. Assoc Prof Steve Johnston, Dr Tamara

Keeley, Mr Tianyi Feng and Ms Alyce Swinbourne (all of UQ-SAFS) assisted practically with the

logistics of experiments reported in Chapter 4. Prof Matt Guille (EXRC), Mr Edward Qualischefski

and Mr Scott Kershaw (both of UQ-SAFS) provided animals for experiments in Chapters 3 and 4

respectively.

Statement of parts of the thesis submitted to qualify for the award of

another degree

Nil

iv

Acknowledgements

Firstly, I would like to express my deepest gratitude to my principle supervisor, Assoc Prof

Steve Johnston for providing me the opportunity to undertake this research. Thank you, Steve for

your guidance, patience and encouragement without which I would not have been able to see this

project through to completion. My heartfelt thanks for your emotional support at times when I lost

direction and for your significant editorial advice. My sincere thanks also go to Prof Jaime Gosálvez

and Dr Carmen López-Fernández and their team at the Autonomous University of Madrid, Genetics

Department for patiently explaining and guiding me through the assay procedures and for

laboratory assistance (BFU-2013-44290-R). Thank you for welcoming me into your lab and

supporting my transition to a lifestyle in Madrid.

I am indebted to Ms Francisca (Paqui) Arroyo and Dr Tamara Keeley for their significant

contributions to the experimental work. Thank you, Paqui for devoting much of your personal time

to travel with me to Portsmouth, UK to assist with experimental work and for your patient technical

advice. Special thanks go to Dr Tamara Keeley for spending many hours slaving away over a

fluorescent microscope to assist with data collection and for willingly standing by me to work well

into the night. Your “be prepared for anything” snack bag approach to experimental work will

certainly stay with me. My sincere thanks also go to Mr Tianyi Feng and Ms Alyce Swinbourne for

taking time away from their own academic pursuits to assist with experimental work. I also wish to

thank Prof Matt Guille and Dr Anna Noble of the European Xenopus Research Centre, Portsmouth,

UK for allowing me access to their facilities and animals. I am truly thankful to Dr Richard Wilson

for taking the time to explain statistical concepts and for willingly agreeing to devote time to

performing complex statistical analyses despite an already full workload.

Finally, I would like to express my warmest appreciation and gratitude to my family; my

parents Ian and Christine Pollock, brother Matthew Pollock and fiancé James Lewis. Without your

love and support through both the high and low times, I could not have achieved this milestone.

Thank you for your words of encouragement, hugs and many cups of tea. You carry this success

with me.

v

Keywords

amphibian spermatozoa, sperm DNA fragmentation, sperm chromatin dispersion, SCD,

cryopreservation, Xenopus laevis, Bufo marinus

Australian and New Zealand Standard Research Classifications

(ANZSRC)

ANZSRC code: 060802, Animal Cell and Molecular Biology, 50%

ANZSRC code: 060803, Animal Developmental and Reproductive Biology, 50%

Fields of Research (FoR) Classification

FoR code: 0601, Biochemistry and Cell Biology, 50%

FoR code: 0608, Zoology, 50%

vi

Table of contents

Abstract i

Declaration by author i

Publications during candidature ii

Publications included in this thesis ii

Contributions by others to the thesis iii

Statement of parts of the thesis submitted to qualify for the award of another degree iii

Acknowledgements iv

Keywords v

Australian and New Zealand Standard Research Classifications (ANZSRC) v

Fields of Research (FoR) Classification v

Table of contents vi

List of tables x

List of figures xii

List of abbreviations xiv

Glossary xvi

An explanation of the thesis format xviii

CHAPTER 1: Introduction and review of the literature: Sperm chromatin in the Amphibia 1

1.1 Abstract 2

1.2 Introduction 2

1.3 The amphibian sperm cell 4

1.3.1 Acrosome 4

1.3.2 Nucleus 5

1.3.3 Sperm tail and undulating membrane 6

1.4 DNA packaging in mammalian and lower vertebrate sperm nuclei 7

1.4.1 Chromatin remodelling in mammalian spermatozoa 7

1.4.2 Amphibian sperm nuclear basic proteins (SNBPs) 9

1.4.3 Sperm chromatin structure 12

1.5 Aetiology of sperm DNA damage 14

1.5.1 Defective spermiogenesis 16

1.5.2 Reactive oxygen species (ROS) and oxidative stress 17

1.5.3 Abortive apoptosis 18

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1.6 Iatrogenic induced sperm DNA damage and sperm DNA fragmentation dynamics 19

1.7 The effect of cryopreservation on DNA integrity 21

1.7.1 Oxidative stress as a mechanism of DNA cryoinjury 21

1.7.2 Osmotic stress and activation of sperm motility 22

1.7.3 Fertilisation success following cryopreservation 23

1.7.4 The influence of seasonal breeding on DNA integrity 24

1.8 Review of technologies used to assess DNA integrity 25

1.8.1 Sperm chromatin structure assay (SCSA) 27

1.8.2 Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) 28

1.8.3 Single-cell gel electrophoresis (SCGE) or comet assay 29

1.8.4 Sperm chromatin dispersion (SCD) test 31

1.9 Conclusion 32

1.10 Thesis aims and structure 33

CHAPTER 2: Validation of the sperm chromatin dispersion (SCD) test in the Amphibian Xenopus

laevis using in situ nick translation and comet assay 35

2.1 Abstract 36

2.2 Introduction 36

2.3 Materials and methods 38

2.3.1 Animals, sperm recovery and experimental design 38

2.3.2 Sperm chromatin dispersion test 39

2.3.3 In situ nick translation (ISNT) 39

2.3.4 Comet assay 40

2.3.5 Statistical analysis 41

2.4 Results 41

2.4.1 Sperm morphology after the SCD test 41

2.4.2 In situ DNA labelling of DNA strand breaks 41

2.4.3 Double-comet assay 43

2.5 Discussion 46

CHAPTER 3: Dynamics of sperm DNA fragmentation in Xenopus laevis using the sperm chromatin

dispersion (SCD) test; the effect of activation and cryopreservation. 50

3.1 Abstract 51

3.2 Introduction 51

viii



3.3.2 Sperm freezing (-20 °C) and cryopreservation (-80 °C) 55

3.3.3 In vitro fertilisation (IVF) 56



3.4 Results 57

3.4.1 Dynamics of sperm DNA fragmentation 57

3.4.2 Regression analysis of SDF after incubation for fresh, frozen (-20 °C) and

cryopreserved (-80 °C) samples 58

3.4.3 Effect of SDF on fertilisation success after cryopreservation at -80 °C 59

3.5 Discussion 60

CHAPTER 4: Effect of cryopreservation on the dynamic behaviour of sperm DNA fragmentation,

motility and viability in cane toad (Bufo marinus) spermatozoa 64

4.1 Abstract 65

4.2 Introduction 65



4.3.2 Assessment of sperm motility and viability 70



4.4 Results 71

4.4.1 Sperm morphology 71

4.4.2 Dynamics of DNA fragmentation in fresh spermatozoa 72

4.4.3 Dynamics of motility in fresh spermatozoa 74

4.4.4 Dynamics of viability in fresh spermatozoa 75

4.4.5 Dynamics of DNA fragmentation in cryopreserved spermatozoa 77

4.4.6 Dynamics of motility in cryopreserved spermatozoa 79

4.4.7 Dynamics of viability in cryopreserved spermatozoa 79

4.5 Discussion 79

CHAPTER 5: General discussion 84

5.1 Introduction 84

ix

5.2 Validation of the sperm chromatin dispersion (SCD) test in amphibian spermatozoa 84

5.3 Dynamics of sperm DNA fragmentation in fresh amphibian spermatozoa 86

5.4 Activation of motility and sperm DNA fragmentation 87

5.5 Effect of cryopreservation on the DNA integrity of amphibian spermatozoa 88

5.6 Effect of cryopreservation on sperm motility and viability 89

5.7 Limitations of the thesis 90

5.8 Summary of new and significant findings and recommendations for future research 91

CHAPTER 6: References 93

x

List of tables

Chapter 1

Table 1.1 Amino acid composition of amphibian sperm nuclear basic proteins 10

Table 1.2 Summary of the tests used to analyse the integrity of sperm chromatin,

including a description of the assay principle, detection methods and their

main strengths and weaknesses (Tavalaee et al. 2008; Sharma and

Agarwal 2011) 26

Chapter 2

Table 2.1 Percentage of SDF as assessed by the SCD in fresh activated African

clawed frog spermatozoa in three frogs incubated over 24 h at 25 °C 45

Table 2.2 Percentage of SDF as assessed by the double-comet assay in fresh

activated African clawed frog spermatozoa in three frogs incubated over

24 h at 25 °C 45

Chapter 3

Table 3.1 Regression analysis of SDF in African clawed frog incubated at room

temperature for 60 minutes examined fresh as well as following freezing (-

20 °C) and cryopreservation (-80 °C) 59

Table 3.2 Fertilisation success resulting from an IVF procedure using fresh and

cryopreserved-thawed (-80 °C) African clawed frog spermatozoa 60

Chapter 4

Table 4.1 Mean percentage and regression analysis of SDF, viability and motility in

fresh cane toad spermatozoa incubated at 22 °C for 6 hours 74

xi

Table 4.2 Mean percentage and regression analysis of SDF, viability and motility in

cryopreserved-thawed cane toad spermatozoa incubated at 22 °C for 6

hours 78

xii

List of figures

Chapter 1

Figure 1.1 Diagrammatic representation comparing Anuran spermatozoa of the

families (a) Dendrobatidae (Garda et al. 2002) and (b) Pseudinae (Garda et

al. 2004) 5

Figure 1.2 Scanning electron microscopy micrographs of the sperm heads and acetic

acid-urea polyacrylamide gel electrophoresis (AU-PAGE) analysis of the

SNBPs of three amphibian species 13

Figure 1.3 A two-step hypothesis to explain the aetiology of DNA fragmentation in

spermatozoa (Aitken and De Iuliis 2011) 15

Figure 1.4 Fluorescence photomicrographs of DNA fragmentation in African clawed

frog (modified from Pollock et al. 2015) spermatozoa evidenced by

different assay techniques 30

Chapter 2

Figure 2.1 African clawed frog sperm nuclear morphotypes after the SCD test 42

Figure 2.2 Evidence of DNA fragmentation in African clawed frog spermatozoa

following the double (neutral + alkaline) comet assay at T0 immediately

following activation 44

Figure 2.3 Correlation between the percentage of SDF identified in three male

African clawed frogs immediately following activation as estimated by

SCD test (ASM-3) and double-comet assay (ACM-2 + ACM-3 + ACM-

2+3) 46

xiii

Chapter 3

Figure 3.1 A representation of the experimental design detailing the division and

treatment of a single testis for an individual African clawed frog 54

Figure 3.2 Logistic model plots representing the dynamics of SDF in the African

clawed frog after 60 minutes of incubation at 25 °C 58

Chapter 4

Figure 4.1 Representation of the experimental design detailing the division and

treatment of testes for an individual cane toad 69

Figure 4.2 Cane toad sperm nuclear morphotypes after the SCD test 72

Figure 4.3 Logistic model plots representing the dynamics of SDF in cane toad

spermatozoa over 6 hours of incubation at 22 °C 73

Figure 4.4 Logistic model plots representing the dynamics of sperm motility in cane

toad spermatozoa over 6 hours of incubation at 22 °C 75

Figure 4.5 Logistic model plots representing the dynamics of sperm viability in cane

toad spermatozoa over 6 hours of incubation at 22 °C 76

Figure 4.6 Logistic model plots representing the dynamics of chromatin swelling

(dead-swollen spermatozoa) in cane toad spermatozoa over 6 hours of

incubation at 22 °C 77

xiv

List of abbreviations

ACM amphibian comet morphotype

ACT activated spermatozoa

AEC animal ethics committee

ALS alkali labile sites

ART assisted reproductive techniques

ASM amphibian sperm morphotype

ATP adenosine triphosphate

CCD charge-coupled device

Cy3 cyanine3 N-hydroxysuccinimide esters

DFI DNA fragmentation index

dH20 distilled water

DMSO dimethyl sulphoxide

DNA deoxyribonucleic acid

DSB double-stranded DNA breaks

DTT dithiothreitol

dUTP deoxyuridine triphosphate

EDTA ethylenediaminetetraacetic acid

EXRC European Xenopus Resource Centre

FBS foetal bovine serum

FITC fluorescein isothiocyanate

g gram

h hour

H histone

H1V histone 1 variant

hCG human chorionic gonadotropin

HDS high DNA stainability

INACT inactivated spermatozoa

ISNT in situ nick translation

IU international unit

IUCN International Union for Conservation of Nature

IVF in vitro fertilization

kg kilogram

xv

LN2 liquid nitrogen

m minute

M molar

MBS modified Barth’s saline

mg milli gram

min minute

mL milli litre

mm milli metre

mmol milli molar

mOsmol milli osmole

PBS phosphate-buffered saline

pH hydrogen ion concentration, -log 10

ROS reactive oxygen species

SAR simplified amphibian ringer

SCD sperm chromatin dispersion test

SCSA sperm chromatin structure assay

SDF sperm DNA fragmentation

SEM standard error of mean

SNBPs sperm nuclear basic proteins

SPSS statistical package for the social sciences

SSB single-stranded DNA breaks

T time

TBE tris–borate–EDTA buffer

TdT terminal deoxynucleotidyl transferase

TEM transmission electron microscopy

TP transition protein

TUNEL terminal dUTP nick-end labelling

μL micro litre

V volt

v/v volume:volume ratio

w/v weight:volume ratio

xvi

Glossary

Covalent disulphide bond – Strong inter- and intra-molecular bonds that arise from amino acid

(cysteine) residues between protamines to stabilise the sperm chromatin structure.

Denaturation – Separation of a double-stranded DNA helix into two single strands, which occurs

when the hydrogen bonds between the strands are broken by application of a high

temperature or strong acid.

Histone hyperacetylation – Removal of the positive charge of the histones by acetylation decreases

the interaction of the histones with the negatively charged phosphodiester backbone of the

DNA helix, causing relaxation of the condensed chromatin structure.

Osmolality – A measure of solute concentration (osmoles) per kilogram of solvent (mOsmol kg-1).

This differs from osmolarity which is a measure of solute concentration per litre of solvent

(mOsmol L-1).

Phosphodiester backbone – Alternating sugar and phosphate groups that form the structural

framework of a negatively charged single strand of DNA. The molecular double-helix shape

of double-stranded DNA is formed by two linear phosphodiester backbones that run

opposite each other and twist together in a helical shape.

Recombinational repair – Exchange of nucleotide sequences between two molecules of DNA to

repair breaks that occur on both strands i.e. double-stranded DNA breaks.

Spermatogenesis – Process by which spermatozoa are produced by way of mitosis and meiosis.

Initial cells called spermatogonia yield primary spermatocytes by mitosis, which divide to

produce two secondary spermatocytes (Meiosis I). Each secondary spermatocyte divides

into two spermatids (Meiosis II) that mature during spermiogenesis.

Spermiogenesis – Final stage of spermatogenesis during which the spermatids mature to become

motile spermatozoa. This process is typically divided into four stages characterised by

condensation of the sperm chromatin, formation of the acrosomal cap and tail and removal

of excess residual cytoplasm.

xvii

Transcription – Process by which a DNA sequence is read and copied by the enzyme RNA

polymerase to produce a complementary RNA strand (messenger RNA). This process is

important for the synthesis and regulation of proteins which perform important functions

within the spermatozoon.

Translation – Process by which messenger RNA produced during transcription is decoded to

produce specific combinations of amino acids which are linked together to form a

polypeptide chain. These chains are then folded to form proteins that constitute the structure

of the DNA molecule.

xviii

An explanation of the thesis format

This thesis has been formatted so that relevant papers published (Chapter 2), submitted

(Chapter 1) or prepared for publication (Chapters 3 and 4) during the candidature have been

presented as thesis chapters. Chapter 2 (published as Pollock et al. 2015) was accepted for

publication prior to submission of Chapter 1 as a review article. No studies had examined chromatin

damage in amphibian spermatozoa prior to the commencement of this candidature. Consequently, in

order to present the most up to date information pertaining to amphibian sperm DNA fragmentation

in Chapter 1, references to Pollock et al. (2015), were included. All references have been removed

from the individual chapters and presented in the general reference list at the end of the thesis.

1

CHAPTER 1: Introduction and review of the literature: Sperm

chromatin in the Amphibia

Pollock, K. 1, Gosálvez, J. 2, López-Fernández, C. 2, and Johnston, S.D.1

1School of Agriculture and Food Science, The University of Queensland, Gatton, Australia 4343. 2Department of Biology, Genetics Unit, The Autonomous University of Madrid, Madrid, Spain

20849.

Presented as submitted to the Journal of Herpetology

2

1.1 Abstract

The focus of amphibian conservation has recently shifted towards the establishment of a

genetic resource bank that in association with assisted reproductive technologies (ART) could be

used as a means of securing anuran biodiversity at risk of extinction (Kouba and Vance 2009).

Gamete cryopreservation is one such technology that provides a mechanism and network for

sustaining the genetic viability of small fragmented populations and is thus a fundamental aspect of

endangered species conservation. However, the process of gamete cryopreservation is not without

adverse effects and increasing importance has been placed on understanding the effect of the freeze-

thaw process on sperm DNA integrity. Despite extensive literature detailing sperm chromatin

organisation and DNA fragmentation in mammalian species, current knowledge pertaining to

amphibians is extremely limited. Consequently, this review will gather current information on

amphibian sperm chromatin organisation, as well as explore the etiology of DNA fragmentation in

vertebrate spermatozoa more generally, with a specific interest on the effect of cryopreservation.

We maintain that a better understanding of sperm chromatin structure and behaviour will lead to

improved methods of amphibian sperm preservation.

1.2 Introduction

Amphibian conservation continues to receive considerable attention due to the large number

of species threatened with extinction (Kouba and Vance 2009). According to the most recent update

of the International Union for Conservation of Nature (IUCN) Red List in 2008, an estimated one-

third of nearly 6000 amphibian species worldwide are experiencing rapid population declines, while

as many as 159 species may already be extinct; 38 species are known to be extinct, one is extinct in

the wild (only to be found in captivity) and at least 120 species have not been seen in recent years

(IUCN 2008). In Australia alone, 48 of 223 species (nearly 22%) are experiencing population

declines (IUCN 2008). With nearly half (42%) of the remaining amphibian species worldwide

continuing to decline in population size, such an unprecedented escalation in the number of

threatened species is of dire concern. Although habitat loss remains the most significant threat to

populations in the wild, other factors such as environmental contaminants, pathogens (particularly

the fungal disease chytridiomycosis) and climate change, are all contributing to the global

amphibian extinction crisis (Kouba and Vance 2009; Hayes et al. 2010). The unique anatomy and

physiology of amphibians leaves them particularly sensitive to fluctuations in environmental

conditions. Their highly porous skin coupled with a dependence on water for breeding means they

3

often react adversely to changes in the environment before that of higher order vertebrates (Kouba

and Vance 2009; Burlibasa and Gavrila 2011). Accordingly, amphibians have been described as a

sentinel species for modelling the effects of climate change and environmental damage (Kloas and

Lutz 2006; Kouba and Vance 2009; Burlibasa and Gavrila 2011; Valencia et al. 2011).

The focus of amphibian conservation has recently shifted towards the establishment of a

genetic resource bank that in association with assisted reproductive technologies (ART) could be

used as a means of securing anuran biodiversity at risk of extinction, as well as providing a

mechanism and network for sustaining the genetic viability of small fragmented existing

populations (Kouba and Vance 2009). For example, in vitro fertilisation (IVF) in amphibians is

becoming an increasingly useful protocol as part of species propagation used to support the genetic

management and reproductive output of amphibian breeding programs (Kouba and Vance 2009).

As such, gamete cryopreservation is a fundamental aspect of endangered species conservation and

successful protocols have been established for a number of amphibian species (Browne et al. 1998;

Michael and Jones 2004; Sargent and Mohun 2005).

Understanding the effect of sperm cryopreservation and climate change phenomena on

breeding capacity, specifically gamete DNA stability, is of critical importance for the conservation

of threatened and endangered amphibians and their environment. The integrity of sperm DNA is

now recognised as a potential indicator of fertility and is essential for successful syngamy and

embryonic development (D’Occhio et al. 2007; Gosálvez et al. 2015). There have been extremely

limited studies that have investigated DNA fragmentation in amphibians, and the majority of these

have focused on the effects of environmental toxicity on somatic cells (Wang and Jia 2009; Marselli

et al. 2010; Valencia et al. 2011). To date, only two studies exist reporting DNA fragmentation in

amphibian spermatozoa (in African clawed frog; Xenopus laevis; Pollock et al. 2015 and in Western

clawed frog; Xenopus tropicalis; Morrow et al. 2014; poster presented at the World Congress of

Reproductive Biology).

Given the vast differences in the protein composition and structure of spermatozoa across

different vertebrate taxa, it is not surprising that species-specific protocols are required to accurately

assess DNA fragmentation (Gosálvez et al. 2011a). Hence, development of a DNA fragmentation

assay for a novel species, such as the amphibian, requires an understanding of basic sperm structure

and chromatin composition in order to ensure a targeted and validated approach. Although there is a

reasonable body of literature on amphibian sperm morphology as it relates to phylogenic

4

relationships (Ausio 1995; Kwon and Lee 1995; Van der Horst et al. 1995), few of these studies

have described amphibian sperm chromatin structure (Bols and Kasinsky 1971; Kasinsky et al.

1985; Takamune et al. 1991; Yokota et al. 1991; Ausio et al. 2007). Consequently, this review will

aim to summarise the information available on amphibian sperm chromatin organisation, as well as

explore the etiology of DNA fragmentation in vertebrate spermatozoa more broadly, with a specific

interest on the effect of cryopreservation and activation of sperm motility. This review will then

conclude with a comparative analysis of the various techniques used to assess DNA damage and

those most apt to investigate amphibian sperm DNA.

1.3 The amphibian sperm cell

The ultrastructural components of the amphibian sperm cell have been well described (see

Swan et al. 1980; Selmi et al. 1997; Scheltinga et al. 2003). The majority of studies have focused

on spermiogenesis or cytological descriptions of spermatozoa but the predominant interest has been

on the taxonomic classification of the extant amphibian orders; for example, recent studies (e.g.

Ausio 1995; Kwon and Lee 1995; Van der Horst et al. 1995) have used apomorphies of sperm

ultrastructure to investigate the phylogenetic relationships of the amphibians. The three amphibian

orders (Urodela, Anura and Apoda) each possess a unique morphology, substantially different to

that of the classical head-midpiece-tail configuration of eutherian spermatozoa. While considerable

variation in sperm ultrastructure has been documented within families (Garda et al. 2004), basic

components of amphibian spermatozoa remain constant.

1.3.1 Acrosome

The acrosome complex is located at the anterior region of the sperm head and consists of

two conical structures that cover the nucleus; the acrosome vesicle and the subacrosomal cone

(Figure 1.1). The subacrosomal cone separates the nucleus from the acrosome vesicle (Costa et al.

2004). In some spermatozoa, a well-defined nuclear space (the endonuclear canal) exists between

the subacrosomal cone and the nucleus and may result from the chromatin compaction and

associated shrinking of the nucleus during spermiogenesis (Amaral et al. 2000; Garda et al. 2002).

The elongated and pointed acrosome observed in most Anura (genera Hyla, Bufo and Alytes;

Seshachar 1943) is considered an adaptation for the penetration of the gelatinous layer surrounding

the oocyte (Costa et al. 2004). Urodele spermatozoa possess an apical acrosomal barb that protrudes

from the head axis (Selmi et al. 1997); a similar structure is absent in both Anura (Kwon and Lee

5

1995) and Apoda (Van der Horst et al. 1995). The acrosome contains hydrolytic enzymes required

for the penetration of the glycoprotein-based vitelline envelope of the oocyte during fertilisation.

Figure 1.1. Diagrammatic representation comparing Anuran spermatozoa of the families (a) Dendrobatidae (From Garda et al. 2002) and (b) Pseudinae (From Garda et al. 2004).

1.3.2 Nucleus

The amphibian sperm nucleus varies greatly in size and appearance. It is a bulb-shaped

structure in the Apoda and pointed in most Anura (Seshachar 1943). The shape of the sperm head of

individual species is a result of nuclear condensation and shaping during spermiogenesis; the

progressive reduction in the volume of the nucleus is presumably due to a loss of nucleoplasm as

6

the chromatin condenses (Garda et al. 2002). This reduction is considerable in Apoda where the

nucleus is observed to condense by 80 – 95% (Seshachar 1943). The resulting nucleus is typically

either cylindrical or tapers at the anterior end. Unique to urodele spermatozoa is the presence of a

nuclear ridge; a closely packed bundle of parallel tubules that flanks the length of the nucleus

(Selmi et al. 1997). Positioned below the posterior portion of the nucleus is the nuclear fossa; it is a

deep cavity filled with electron-dense material that contains the proximal and distal centrioles

(Selmi et al. 1997). The nuclear fossa is surrounded peripherally by abundant mitochondria

responsible for converting oxygen and nutrients into adenosine triphosphate (ATP) required for the

metabolic processes of the spermatozoon, as well as providing energy for motility (Selmi et al.

1997; Costa et al. 2004). The mitochondria are characterised by concentric cristae that typically

form a spiral aggregation or mitochondrial collar around the base of the axoneme that originates

from the distal centriole (Seshachar 1943; Selmi et al. 1997; Scheltinga et al. 2003; see Figure 1.1).

1.3.3 Sperm tail and undulating membrane

The sperm tail consists of the axoneme and axial sheath. The axoneme is formed from the

distal centriole during spermiogenesis and the two structures are continuous in the mature

spermatozoa. The axoneme contains the microtubule arrangement typical of vertebrates with a

central pair of microtubules surrounded by a circular array of nine outer pairs (Duellman and Trueb

1986; Scheltinga et al. 2003; Costa et al. 2004). Notable differences occur in the tail structure of

anurans (see Figure 1.1). While urodele sperm characteristically have only a single flagellum,

anuran spermatozoa may have one or two tail filaments. The tail filament consists of either a simple

flagellum (as in the genera Rana, Hyla, Pelobates and Xenopus) or a flagellum with an undulating

membrane (genera Alytes, Bufo, Bombina and Discoglossus; Seshachar 1943; Koch and Lambert

1990). The undulating membrane is comprised of an accessory fibre often referred to as the axial

fibre that runs parallel to the axoneme and is united to it by a thin membrane. This membrane

contains dense material that appears to run longitudinally along the axis of the sperm tail.

Cinemicrographic analysis shows that the longer, flexuous axoneme moves actively to propel the

spermatozoa, while the axial fibre follows it passively with a lower amplitude of movement. The

functional significance of the undulating membrane is unlikely to directly aid propulsion, but rather,

may be to provide rigidity for the sperm tail without incurring an energy deficit large enough to

necessitate a long mid-piece to fuel propulsion, thus reducing the energy requirements of the

spermatozoon (Swan et al. 1980). This is supported by the observation that the axial fibre in the

undulating membrane of the newt (genus Notophthalmus; Swan et al. 1980) sperm tail appears to be

7

composed of material with keratinous-like properties, conferring an elastic rigidity of the axial

fibre.

1.4 DNA packaging in mammalian and lower vertebrate sperm nuclei

During mammalian spermiogenesis, the DNA molecule undergoes a series of structural

changes that allow the chromatin to be remodelled into a highly compacted form. Condensation of

the chromatin not only helps to streamline the mature spermatozoon by reducing the cell volume,

but also serves a protective role, reducing the susceptibility of the DNA to physical damage,

mutagens and environmental stressors, so that normal fertilisation, chromatin decondensation can

syngamy occur (Love and Kenney 1998; Gavrila and Mircea 2001; Ausio et al. 2007). The

reorganisation and packaging of the chromatin is accompanied by a marked change in molecular

composition facilitated by the sequential replacement of sperm nuclear basic proteins (SNBPs)

during the Golgi phase of spermiogenesis (Oko et al. 1996). These proteins are characterised as

belonging to discrete groups with variation in physical and chemical composition and include

histones, transition proteins, protamines and protamine-like proteins that comprise a structural

intermediate between histones and protamines (Bloch 1969). Although sperm nuclear condensation

appears to be an essentially conserved process, the protein complements present in the sperm

chromatin vary considerably between species and thus, the specific patterns and mechanisms by

which the chromatin is packaged are also likely to be highly divergent (Manochantr et al. 2005;

Enciso et al. 2011). There is, however, unanimous agreement that the preliminary chromatin

organisation of early-stage spermatids resembles the nucleosomal structure of mitotic somatic cells

(Birstein 1982). Although it is widely accepted that the sperm chromatin structure of eutherian

mammals is basically similar across species (Birstein 1982), little comparative information exists,

especially with respect to amphibians. Therefore, this review will primarily focus on DNA

packaging in mammalian sperm but with reference to amphibians where this information is

available.

1.4.1 Chromatin remodelling in mammalian spermatozoa

The first level of chromatin packing in amphibians does not differ from that in mammals

(Birstein 1982). During the initial stages of spermatogenesis, sperm DNA is closely associated with

histones and is organised into nucleosomal core particles similar to somatic cells (D’Occhio et al.

2007). At this stage, the nucleosome is still comprised of an octamer of core histones around which

8

the chromatin is wrapped forming a solenoid structure (Sharma and Agarwal 2011). The more

histones retained in the chromatin, the less tightly the chromatin will be compacted (Sharma and

Agarwal 2011). Chromatin remodelling is facilitated by loosening of the chromatin by histone

hyperacetylation and the action of DNA topoisomerase II, which relieves the torsional stress

brought about by the nucleosomal structure by producing temporary nicks in the sperm DNA

(Gavrila and Mircea 2001; Sharma and Agarwal 2011).

The first detectable structural change in the condensed state of chromatin in mammalian

sperm coincides with the deposition of lysine-rich transition proteins in place of the histones. This

results in a loss of the nucleosomal structure of the chromatin coupled with the cessation of

transcription (Oko et al. 1996). In the Syrian hamster (Mesocricetus auratus; Brewer et al. 2002),

two transition proteins (TP) have been implicated in the process of sperm chromatin condensation;

TP1 and TP2. The preferential binding of TP2 to CG sequences indicates this protein is primarily

responsible for impairing transcription, while TP1 promotes the repair and/or ligation of single-

stranded DNA breaks induced during the removal of histones by binding to the exposed bases

(Caron et al. 2001; Brewer et al. 2002). In grasshopper spermatozoa (Pyrgomorpha conica; Cerna

et al. 2008), temporary DNA nicking also appears to be synconised with the transient presence of

triplex DNA motives. These motives (also known as H-DNA), occur when two pyrimidine

nucleotide strands share a common purine strand, allowing single-stranded DNA to hybridise with

complimentary strands; these arrangements may contribute to recombinational repair of double-

stranded DNA breaks during the histone-protamine transition (Cerna et al. 2008).

The most significant level of chromatin compaction is attained following the synthesis and

deposition of protamines in place of the transition proteins during spermatid elongation of late

spermiogenesis (Brewer et al. 2002). Protamines are highly charged arginine-rich proteins that are

thought to facilitate condensation by neutralising the negative charge along the phosphodiester

backbone of the DNA at the time of binding (Brewer et al. 2002). The primary function of

protamines is to ensure that the sperm genome remains transcriptionally inert until it can be

deposited inside an oocyte and reactivated during fertilisation (Brewer et al. 2002). In eutherian

mammalian sperm nuclei, the chromatin is further stabilised by inter- and intra-molecular covalent

disulphide bonds that arise from amino acid (cysteine) residues between the protamines, resulting in

a rigid para-crystalline-like structure (Gavrila and Mircea 2001; Cortés-Gutiérrez et al. 2009).

Almost all metatherian (excluding some Planigale spp.) and prototherian mammals examined thus

far do not appear to contain any cysteine residues in their sperm protamine and therefore lack the

9

capacity for disulphide bonding to stabilise the sperm chromatin (Cortés-Gutiérrez et al. 2014b;

Johnston et al. 2015). While generally uncommon, cysteine residues have been reported in the

histone variants of the grey short-tailed opossum (Monodelphis domestica; Johnston et al. 2015), as

well as in some lower vertebrates (Saperas et al. 1992; Rocchini et al. 1996; Zhang et al. 1999),

indicating the possibility of histone-mediated disulphide bonds in these species. Hence, alternative

stabilising mechanisms are likely to function in these species but much further investigation is

required.

1.4.2 Amphibian sperm nuclear basic proteins (SNBPs)

Electrophoretic, cytochemical and amino acid analysis has demonstrated that SNBPs vary

considerably among amphibians. This protein diversity is well illustrated in anurans, where a

number of genera, including Xenopus, Bufo, Hyla and Rana possess SNBPs significantly different

from each other (Bols and Kasinsky 1971; Kasinsky et al. 1985; see Table 1.1). Amino acid

sequence analysis of SNBPs identified that mature Japanese common toad sperm (Bufo japonicus;

Takamune et al. 1991) are composed exclusively of two protamines rich in arginine, while the

mature spermatozoa of African clawed frog (Yokota et al. 1991; Ausio et al. 2007; Figure 1.2b)

contain six different SNBPs in addition to four types of core histones. In contrast, protamines are

totally absent in the sperm of Rana spp. such as the American bullfrog (Rana catesbeiana; Itoh et

al. 1997; Ausio et al. 2007; Figure 1.2a), Indian bullfrog (Rana tigerina; Manochantr et al. 2005)

and northern leopard frog (Rana pipiens; Alder and Gorovsky 1975) and consequently, the

nucleosomal structure is composed of characteristically protamine-deficient chromatin; only core

histones and the lysine-rich variant (H1V) of histone H1 are present.

10

Table 1.1. Amino acid composition of amphibian sperm nuclear basic proteins.

Japanese common toad1

(Bufo japonicas)

African clawed frog2

(Xenopus laevis)

American bullfrog3

(Rana catesbeiana)

P PL H

Lys 5.4 5.4 19.1

His 9.6 1.9 1.7

Arg 42.1 28.0 6.9

Asx 1.7 4.4 4.7

Thr 6.9 9.7 5.2

Ser 5.5 9.8 6.7

Glx 5.0 4.8 7.0

Gly 0.8 8.4 7.8

Ala 4.9 11.5 14.0

Cys 0.3 0.4 <0.1

Val 5.2 3.2 6.5

Met 0.0 1.6 0.5

Ile <0.1 1.8 3.8

Leu <0.1 3.6 6.2

Phe 0.0 1.0 1.2

Tyr 2.8 4.2 2.3

Trp 0.0 0.0 -

Pro 9.5 2.1 6.2

P – protamine type chromatin composition; PL – protamine-like chromatin composition; H – histone type chromatin composition; 1Data from Takamune et al. 1991; 2Data from Yokota et al. 1991; 3Data from Itoh et al. 1997.

The lack of transition proteins and presumably the reduced capacity for DNA repair

available during spermiogenesis in some lower vertebrates does not appear to greatly influence the

basal level of single-strand breaks observed during DNA fragmentation analysis (Caron et al.

2001). Fish (Tinca tinca; López-Fernández et al. 2009 and zebrafish; Danio rerio; Gosálvez et al.

2014) spermatozoa lacking transition proteins generally exhibit a low level of sperm DNA

fragmentation (not larger than 5%) at the time of activation. This trend is not dissimilar from that

observed in mammals, such as the ram (López-Fernández et al. 2008a) and koala (Phascolarctos

cinereus; Zee et al. 2009), suggesting that another mechanism of chromatin stabilisation and/or

DNA single-strand break repair may be at work in lower vertebrate spermatozoa. Protamine-like

11

proteins are present instead of transition proteins in the spermatozoa of some amphibians (Bufo and

Rana spp. and African clawed frog; Risley et al. 1986 and Gavrila and Mircea 2001 respectively).

These highly basic proteins have been described as an evolutionary precursor to protamines and are

highly heterogeneous, but typically have a high combined arginine-lysine content of at least 35 –

50%, albeit not as high as in protamines (Kasinsky et al. 1999; Lewis and Ausio et al. 2002; Ausio

et al. 2007). Since protamine-like proteins usually coexist with histones in the mature amphibian

spermatozoon, it is possible that protamine-like proteins may play a role in the maintenance of the

condensed chromatin structure (Itoh et al. 1997). Lewis and Ausio (2002) suggest that the higher

arginine content of protamine-like proteins leads to increased charge density and greatly enhances

the compaction of the DNA and stabilisation of the sperm nucleus. Indeed, a high percentage

composition of arginine in the SNBPs of koala (Gosálvez et al. 2011b) and African clawed frog

(Yokota et al. 1991; Ausio et al. 2007) spermatozoa (61.7% and 41.2% respectively), may

compensate for the absence of disulphide bonding to stabilise the sperm chromatin and thereby

explain the resistance to DNA fragmentation observed in these two species (Johnston et al. 2012b

and Pollock et al. 2015, respectively). Confirmation of a reduced arginine content of the SNBPs in

species that lack the capacity for disulphide bond formation and that are also known to exhibit rapid

sperm DNA fragmentation dynamics (for example in fish; López-Fernández et al. 2009; Gosálvez

et al. 2014) would lend support to this theory.

The protein composition of the sperm nucleus is an important consideration when

developing species-specific protocols for DNA fragmentation analysis, such as in the amphibian.

This is particularly the case for DNA fragmentation assays such as the sperm chromatin dispersion

(SCD) test and comet assay where visualisation of a differential chromatin dispersion pattern is

dependent on sufficient protein removal (Gosálvez et al. 2011a; Cortés-Gutiérrez et al. 2014b). As

the sperm cells of each species are likely to contain different protamine residues, it is extremely

important to modify the strength of the lysis solution used for protein depletion before an accurate

assessment of DNA fragmentation can be determined (Gosálvez et al. 2011a; Cortés-Gutiérrez et

al. 2014b). Furthermore, the presence or absence of protamines/cysteine residues and associated

disulphide bonds to stabilise the chromatin is also likely to influence the basal level of DNA

damage as well as the longevity of the spermatozoa when used in vitro. Significantly higher levels

of fragmented DNA in response to oxidative stress compared to eutherian mammals have been

observed in marsupial species (Eastern grey kangaroo; Macropus giganteus and common wombat;

Vombatus ursinus; Enciso et al. 2011) as well as in fish (López-Fernández et al. 2009; Gosálvez et

al. 2014); all these subeutherian species lack disulphide bonds. In contrast, African clawed frog

12

spermatozoa appeared to be fairly resistant to DNA fragmentation (Pollock et al. 2015). Although

cysteine residues have been reported in the protamine-like proteins and H1 histones of some bivalve

molluscs (Zhang et al. 1999), anemones (Rocchini et al. 1996) and tunicates (Saperas et al. 1992;

Zhang et al. 1999), Yokota (et al. 1991) claim that none of the SNBPs of African clawed frog

spermatozoa contained cysteine, despite amino acid sequence analysis revealing molar percentages

up to 0.8% in seven of the eight proteins examined. While it is possible that the negligible levels of

cysteine residues observed in African clawed frog spermatozoa may contribute to the stabilisation

of the sperm chromatin, it seems more likely to be a function of the high arginine content of the

SNBPs (up to 41.2%; Yokota et al. 1991) in this species. Interestingly, DNA fragmentation was

only revealed in common dunnart (Sminthopsis murina; Johnston et al. 2015) spermatozoa

following exposure to aggressive protein lysis and a high concentration of a disulphide bond-

reducing agent (1.0 M β-mercaptoethanol), despite possessing cysteine-free protamines; this

phenomenon prompted the authors to suggest that the cysteine residues associated with some

histones/histone variants, as seen in some lower vertebrates (Saperas et al. 1992; Rocchini et al.

1996; Zhang et al. 1999), could potentially establish disulphide bonds in species that retain high

levels of histones in their mature spermatozoa. Yet, the sperm chromatin of zebrafish characterised

solely by somatic histones and a nucleosomal core structure (Ausio et al. 2014), is among the most

susceptible to rapid DNA fragmentation. Clearly, the relationship between SNBP composition and

the associated stability of the sperm chromatin requires further examination.

1.4.3 Sperm chromatin structure

Various models have been proposed to explain the tertiary structure of mammalian sperm

DNA. An early model proposed by Balhorn (1982) suggests that protamines bind to DNA by lying

lengthwise inside the minor groove. The DNA strands of the sperm nucleus are packaged side by

side in a linear array so that the DNA-protamine complex of one strand fits into the major groove of

a neighbouring DNA strand. The positively charged arginine residues of the protamine molecule

neutralise the phosphodiester backbone of the DNA double helix so that the DNA-protamine

complexes of neighbouring strands can bind together by van der Waals forces; a form of

intermolecular attachment arising from the attractive and repulsive forces created by the correlated

positive-negative polarisation of the neighbouring protamine molecules (Ward 1993). Based on this

model, Ward (1993) presented the “sperm doughnut loop domain model”. In this model, protamines

function by binding to the DNA molecule in a non-specific manner and coiling the chromatin into

toroidal, or ring-like, ‘doughnuts’ (Ward 1993; Bewer et al. 2002). Each toroid corresponds to a

13

single DNA loop domain attached to the sperm nuclear matrix. Evidence of this toroidal form of

chromatin organisation in amphibian spermatozoa is limited. Selmi et al. (1997) observed a

concentric lamellar organisation of the chromatin fibres in the spermatozoa of the alpine newt

(Triturus alpestris). However, it would appear unlikely that the doughnut loop model applies to

amphibian species that fully or partially retain histones in the chromatin of mature spermatozoa.

Nuclease digestion and electron micrography of American bullfrog (Itoh et al. 1997; Ausio et al.

2007) spermatozoa known to be completely lacking protamines, has revealed that the chromatin

consists of irregularly-spaced nucleosomes reminiscent of the ‘beads on a string’ arrangement first

described in amphibian erythrocytes by Olins and Olins (1973) and similar to that of somatic cells

(see Figure 1.2d).

Figure 1.2. Scanning electron microscopy micrographs of the sperm heads and acetic acid-urea polyacrylamide gel electrophoresis (AU-PAGE) analysis of the SNBPs of three amphibian species: (a) American bullfrog (Rana catesbeiana); (b) African clawed frog (Xenopus laevis); (c) Cane toad (Bufo marinus). Note that mature spermatozoa of the American bullfrog are composed entirely of histones (H1-4), while African clawed frog and Cane toad spermatozoa respectively contain histones with protamine-like proteins (or sperm-specific proteins; SP2-6) or protamine (P). AUA – axoneme-undulating membrane-axial rod; H – head; MP – midpiece; S – single flagellum with axoneme; (d) Illustrated cross-sections of chromatin fibres comprised of histones (H) or protamines/protamine-like proteins (P/PL; From Ausio et al. 2007).

14

Despite a nucleosomal chromatin structure in the spermatozoa of some amphibians, the

chromatin can still undergo complete condensation to the same degree as in mammalian

spermatozoa, at least as visualised by transmission electron microscopy, suggesting that another

mechanism of chromatin packaging may function in these species (Manochantr et al. 2005; Ausio et

al. 2014). Factors such as a high nucleosome repeat length and histone H1 content in addition to

post-translational core histone modifications may function to promote chromatin condensation in

species that retain a nucleosomal structure (Ausio et al. 2014). Lewis and Ausio (2002) propose a

model of chromatin organisation in protamine-deficient species whereby protamine-like protein 1

(PL-1) neutralises the charge of a length of linker DNA between the nucleosomes, thus allowing the

DNA to wrap a further two times around the nucleosome creating a highly compacted structure.

Saperas et al. (2006) reported a novel chromatin structure in striped red mullet (Mullus surmuletus)

spermatozoa where compaction is mediated by protamine-like molecules; in this species the

chromatin consists of irregular parallel DNA bundles (see Figure 1.2d), reminiscent of the toroidal

chromatin organisation typically associated with protamines. Similarly, rooster (Gallus gallus

domesticas; Ausio et al. 2014) spermatozoa are composed of cysteine-free protamines, where the

chromatin appears as elongated fibres or rods that tend to align with the long axis of the nuclei. A

combination of both toroids and rods have been observed in mouse (Mus musculus) spermatozoa

(Ausio et al. 2014).

1.5 Aetiology of sperm DNA damage

When evaluating the DNA fragmentation dynamics of a target species, it is first necessary to

differentiate between inherent constitutive DNA modifications that are observed immediately

following ejaculation/activation, mostly associated with the normal chromatin remodelling process

and inducible or iatrogenic modifications that arise as a function of incubation time following

ejaculation/activation (Alvarez and Gosálvez 2012). Manipulation of spermatozoa in vitro for ART

exposes the spermatozoa to exogenous stressors that can influence the longevity of sperm DNA in a

given species (Gosálvez et al. 2011b). The aetiology of sperm DNA fragmentation is multi-factorial

and three major mechanisms have been proposed for the endogenous formation of DNA damage in

the spermatozoon; (1) defective chromatin remodelling during spermiogenesis resulting in

unresolved strand breaks, (2) excessive generation of reactive oxygen species (ROS) which

ultimately lead to oxidative stress and (3) abortive apoptosis (Sharma and Agarwal 2011). While

damage to the DNA could arise due to any combination of these mechanisms, they also appear to be

intrinsically related. Aitken and De Iuliis (2011) proposed a two-step hypothesis to explain the

15

relationship between these factors (Figure 1.3): The disruption of spermiogenesis as a consequence

of oxidative stress within the testes results in the production of spermatozoa with poorly packaged

chromatin that are themselves more vulnerable to oxidative attack. As the defective spermatozoa

default to the apoptotic pathway, the sperm mitochondria generate ROS that in turn attack the sperm

chromatin, leading to extensive DNA damage.

Figure 1.3. A two-step hypothesis to explain the aetiology of DNA fragmentation in spermatozoa. Step (1) The disruption of spermiogenesis as a result of oxidative stress within the testes brought about via an array of exogenous factors, results in the production of spermatozoa with poorly packaged chromatin that are in turn more vulnerable to oxidative attack. Step (2) As the defective spermatozoa default to the apoptotic pathway, the sperm mitochondria generate ROS which in turn attack the sperm chromatin, leading to extensive DNA damage (Aitken and De Iuliis 2011).

16

1.5.1 Defective spermiogenesis

Spermiogenesis is a key event in the occurrence of DNA damage in the male germ line.

During spermiogenesis the chromatin undergoes extensive remodelling involving physiological

DNA strand breaks. McPherson and Longo (1993) suggested that the programmed formation and

subsequent ligation of transient DNA nicks is required to facilitate protamination during chromatin

packing. These nicks are produced by the endogenous nuclease activity of topoisomerase II and are

considered necessary to relieve the torsional stress associated with the nucleosomal structure and

aid chromatin rearrangement during the displacement of histones by protamines; this allows the

chromatin to unwind and condense into a toroidal form (Sakkas and Alvarez 2010). Ligation of

DNA breaks is essential for preserving the integrity of the primary DNA structure. It has, therefore,

been proposed that differentiating spermatids must be equipped with enzyme activity capable of

repairing cleavage sites (Cabrero et al. 2007). Single-stranded breaks in elongating mammalian

spermatids are typically repaired by end-joining repair processes coupled with the progressive

nuclear accumulation of TP1 (Caron et al. 2001; Grégoire et al. 2011). DNA integrity is normally

fully restored before the spermatozoa are released from the germinal epithelium during

spermiogenesis (Sharma and Agarwal 2011). The lack of transition proteins available during

amphibian (i.e. non-mammalian; Ausio et al. 2007) spermiogenesis suggests that this mechanism of

DNA nick ligation is somewhat altered in this and other lower vertebrate taxa. While abnormal

chromatin condensation as well as reduced fertility have been observed in TP1-deficient mice (Yu

et al. 2000), the lack of transition proteins in the grasshopper (Eyprepocnemis plorans; Cabrero et

al. 2007) does not appear to hinder DNA repair, as evidenced by the detection of the DNA-repair

protein Ku70. Thus, the end joining repair pathways typical of mammalian sperm also appear to

function in lower vertebrates in the absence of transition proteins, suggesting that the participation

of transition proteins may simply enhance reliability of the DNA repair mechanism but are not

essential to its function (Grégoire et al. 2011).

Chromatin remodelling is an inherent sensitive step during spermiogenesis whereby

disruption of this process may result in spermatozoa with chromatin packaging abnormalities or

unresolved single and double-stranded DNA breaks. The presence of endogenous DNA nicks in

ejaculated spermatozoa may indicate incomplete maturation during spermiogenesis and are likely to

make the affected spermatozoa more susceptible to post-testicular assault by ROS and denaturation

(Sakkas and Alvarez 2010). Indeed, ‘structural’ single-stranded DNA breaks persist in the mature

spermatozoa of a number of species (Cortés-Gutiérrez et al. 2007; Johnston et al. 2009; López-

17

Fernández et al. 2009; Zee et al. 2009; Pollock et al. 2015). These breaks arise from alkali labile

sites (ALS); highly sensitive DNA motifs (regions of specific repetitive sequences), such as

apurinic or apirimidinic sites that produce single-stranded DNA following incubation in alkaline

solutions (Cortés-Gutiérrez et al. 2007). Cortés-Gutiérrez et al. (2009) proposed that ALS are

constitutive structural characteristics of the DNA and are likely to be universally present in the

sperm nucleus of all mammalian species. Assessing the presence of ALS is interpreted as an

alternative method of DNA packaging and may function in addition to and/or in the absence of

disulphide bonding to produce the highly condensed state of the chromatin (Cortés-Gutiérrez et al.

2009). These sites are believed to facilitate extreme chromatin compaction in the absence of

disulphide bond-mediated protamination in the koala (Zee et al. 2009), short-beaked echidna

(Tachyglossus aculeatus; Johnston et al. 2009), fish (López-Fernández et al. 2009) and African

clawed frog (Pollock et al. 2015). Pollock et al. (2015) identified extensive structural single-

stranded DNA breaks in African clawed frog spermatozoa; differentiation between single and

double-stranded DNA breaks via the double comet assay revealed that at least 91% of the sperm

population exhibited constitutive rather than true DNA damage immediately following sperm

activation.

1.5.2 Reactive oxygen species (ROS) and oxidative stress

Oxidative stress is one of the most significant factors contributing to sperm DNA damage

following ejaculation, although the exact mechanisms of its action remain unresolved. ROS are

highly active oxidising agents such as free radicals and hydrogen peroxide (H2O2; Tavalaee et al.

2008). The generation of low levels of ROS is a normal physiological process required for the

signalling events that control sperm capacitation, the acrosome reaction, hyperactivation and sperm-

oocyte fusion (Said et al. 2012). A primary source of ROS in the seminal plasma is derived from the

spermatozoa themselves; for example, the degree of ROS production in fish is controlled by an

antioxidant system composed of enzymatic and non-enzymatic components that act as ROS

scavengers (Tavalaee et al. 2008; Martínez-Páramo et al. 2012). Excessive generation of ROS by

defective spermatozoa, especially immature spermatozoa carrying surplus residual cytoplasm and

by seminal leukocytes leads to an imbalance in the antioxidant capacity of the seminal plasma and

the onset of oxidative stress (Said et al. 2012). Spermatozoa are particularly susceptible to oxidative

stress due to the high concentration of unsaturated fatty acids in the plasma membrane, a low

concentration of scavenging enzymes in the cytoplasm and a limited capacity for repair (Martínez-

Páramo et al. 2012). ROS attack the unsaturated fatty acids in the plasma membrane, damaging the

18

permeability and fluidity of the membrane as well as initiating a lipid peroxidation cascade, an

important mechanism of action of oxidative stress that can lead to chromatin cross-linking, base

changes and DNA strand breaks (Martínez-Páramo et al. 2012; Said et al. 2012).

Numerous studies have reported the detrimental effect of oxidative stress on sperm function

and DNA integrity (Cabrita et al. 2011; Enciso et al. 2011). In addition to the antioxidant capacity

of the seminal plasma, efficient chromatin condensation is essential for the protection of sperm

DNA from oxidative attack. It has been suggested that oxidative stress in the testes results in the

production of spermatozoa with poorly remodelled chromatin that are in turn more susceptible to

oxidative attack by ROS inherent in the seminal plasma (Aitken and De Iuliis 2012). Enciso et al.

(2011) suggests that adequate oxidation of thiols for chromatin condensation during epididymal

transit is necessary to ensure efficient protamination and disulphide bond formation required to

stabilise and protect eutherian spermatozoa against oxidative attack and DNA damage. It is,

therefore probable, that amphibian spermatozoa lacking protamines, and consequently disulphide

cross-linking (ie. Rana spp.; Risley et al. 1986), would be particularly sensitive to oxidative stress.

Similarly, marsupial spermatozoa (koala, Eastern grey kangaroo and common wombat; Enciso et al.

2011) inherently lacking disulphide bonds are much more sensitive to oxidative attack than those of

eutherian species, demonstrating a significant increase in single-stranded DNA breaks following

exposure to H2O2 and imposed oxidative stress. Little information exists describing the effects of

ROS on amphibian spermatozoa. Exposure of Mongolian toad (Bufo raddei; Feng et al. 2011)

spermatozoa to 4-Nonylphenol (an estrogen-like organic pollutant) and H2O2 has been shown to

significantly impair sperm motility and the integrity of the plasma membrane (as assessed by

morphology criteria); the effect of ROS on amphibian sperm DNA has yet to be explored.

1.5.3 Abortive apoptosis

Apoptosis is a mode of physiologically programmed cell death and is fundamental for

normal spermatogenesis (Yu et al. 2011). During spermatogenesis, the testis is characterised by

very high rates of cell proliferation and differentiation. An overabundance of cell proliferation

disrupts the supportive capacity of the Sertoli cells and necessitates the elimination of excess germ

cells by apoptosis (Sakkas and Alvarez 2010). This process is based on a genetic mechanism that

induces a series of cellular, morphological and biochemical alterations leading the cell to suicide

and become phagocytosed without eliciting an inflammatory response (Said et al. 2012). However,

this mechanism may not always operate efficiently post-meiotically. As male germ cells develop

19

into highly differentiated and transcriptionally inert spermatozoa, they lose their capacity to drive

the process of apoptosis to completion (Aitken and De Iuliis 2011; Sharma and Agarwal 2011).

Instead, a proportion of differentiating germ cells are thought to undergo a restricted ‘abortive’ form

of this process leading to defective remodelling and DNA cleavage, but retaining the capacity to

develop into mature spermatozoa with the potential for fertilisation (Barratt et al. 2010). The

presence of mature spermatozoa in the ejaculate expressing distinct markers of apoptosis-related

damage, such as DNA strand breaks, therefore, suggests that an abortive apoptosis may have taken

place (Yu et al. 2011).

Despite extensive evidence demonstrating the presence of DNA fragmentation in mature

spermatozoa, the significance of apoptosis as a primary cause remains unresolved. Apoptosis

appears to be intrinsically linked to ROS generation and oxidative stress. Aitken and De Iuliis

(2011) speculate that mature spermatozoa default to an apoptotic pathway in response to oxidative

stress; this in turn induces DNA damage through an oxidative attack mediated by mitochondrial

ROS generation. Furthermore, Sakkas and Alvarez (2010) have proposed that apoptosis is

responsible for the process of stripping the cytoplasm in the final stages of sperm maturation.

Consequently, abortive apoptosis may be in part responsible for the release of immature

spermatozoa carrying surplus residual cytoplasm and the resulting excessive generation of ROS in

the seminal plasma and the onset of further oxidative stress (Said et al. 2012).

1.6 Iatrogenic induced sperm DNA damage and sperm DNA fragmentation dynamics

Semen parameters such as motility and vitality are usually evaluated only once after sperm

collection. Similarly, sperm DNA fragmentation has typically been represented as a single value at

the time of assessment, giving rise to a belief in the static nature of this parameter (López-

Fernández et al. 2009). Based on the assumption that sperm DNA is unstable when maintained in a

para-biological environment, such as the conditions used to store semen following ejaculation or

which mimic the female reproductive tract, sperm DNA quality is likely to change during the useful

lifespan of a sperm sample (Gosálvez et al. 2011a). It is, therefore, probable that measurements of

DNA damage performed at the time of ejaculation/activation and at the time of processing for ART

are likely to differ due to the time lapse between both periods. Recent studies have demonstrated

that sperm DNA fragmentation is in fact a highly dynamic process, increasing from the time of

ejaculation up until fertilisation of the oocyte (López-Fernández et al. 2007; 2008ab). The

conditions of sperm storage particularly influence sperm DNA longevity and a certain amount of

20

iatrogenic DNA damage is to be expected once the spermatozoa are incubated ex vivo for ART

(Gosálvez et al. 2011a).

Comparative analysis of the dynamics of sperm DNA fragmentation in different species

highlights the varying resistance of sperm chromatin to external stressors. A progressive loss of

sperm DNA quality has been observed in a number of species, including but not limited to, the ram

(López-Fernández et al. 2008a), stallion (López-Fernández et al. 2007), boar (Pérez-Llano et al.

2010), donkey (Equus asinus; Cortés-Gutiérrez et al. 2008a), rhinoceros (Diceros bicornis,

Rhinoceros unicornis, Ceratotherium simum; Portas et al. 2009), koala (Zee et al. 2009; Johnston et

al. 2012b), short-beaked echidna (Johnston et al. 2009), fish (López-Fernández et al. 2009;

Gosálvez et al. 2014) and recently, in the amphibian (Pollock et al. 2015). There is considerable

inter-species, as well as inter-individual variation in the basal level of damaged DNA and the rate at

which the DNA fragments. This is not surprising given the large variation in the protein

composition of sperm DNA of different species. For example, in fish (López-Fernández et al. 2009;

Gosálvez et al. 2014) an increase in fragmented DNA was triggered only a few minutes after

activation in freshwater, while in the boar (Pérez-Llano et al. 2010) spermatozoa were incubated at

37 °C in a species-specific semen extender for days before an increase in sperm DNA fragmentation

was observed. The rates of DNA damage reported in fish are the fastest measured thus far and may

be due in part to osmotic stress imposed on the spermatozoa following activation and/or a

chromatin structure lacking disulphide bonds (López-Fernández et al. 2009; Gosálvez et al. 2014).

Interestingly, the African clawed frog (Pollock et al. 2015) that utilises the same strategy of

osmotically induced activation as in fish and which also lacks the capacity for disulphide bonding in

the sperm chromatin, exhibited a much slower rate of DNA damage; the level of sperm DNA

fragmentation as determined by the SCD test ranged from 8.7 – 20.3% following incubation for one

hour compared to 45 – 90% in fish (López-Fernández et al. 2009). The relative stability of the

chromatin in African clawed frog spermatozoa is comparable to that seen in the koala (Johnston et

al. 2012b) where a remarkably low level of DNA damage was observed in chilled preserved

spermatozoa even after 16 days of storage at 4 °C. These observations give support to the

probability other mechanisms may be functioning to stabilise the sperm chromatin of the African

clawed frog and other disulphide bond-deficient species and that this is perhaps related to a high

arginine content of the SNBPS. The highly variable rate at which DNA quality decays highlights

the importance of considering inherent differences to the sperm chromatin structure when applying

a DNA fragmentation assay to a novel species.

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1.7 The effect of cryopreservation on DNA integrity

The effect of cryopreservation on reproductive potential, specifically sperm DNA stability,

is of critical importance for improving ART for the conservation of endangered amphibians. An

absolute requirement for the use of frozen-thawed sperm for ART is that the cryopreservation

process must not induce alterations that would impede progeny viability and development (Labbe et

al. 2001). Despite many refinements in methodology and successful protocols established for a

number of amphibian species (Beesley et al. 1998; Browne et al. 2002a; Michael and Jones 2004;

Sargent and Mohun 2005) the procedure of cryopreservation is not without substantial adverse

effects. Sperm cryopreservation promotes cellular cryodamage that can compromise sperm quality

in terms of motility, vitality, membrane stability and ultimately DNA integrity, and can lead to

impaired fertilisation potential and in mammals, impaired embryonic development (Martínez-

Páramo et al. 2012). However, it has still not been clearly established whether there exists a direct

relationship between cryopreservation and sperm DNA damage. Gosálvez et al. (2010), suggest that

the incidence of increased single- and/or double-stranded DNA breaks following cryopreservation

is not a direct consequence of the freezing process, but rather, that modifications to the chromatin-

SNBP interaction as a result of freeze-thawing predisposes the sperm DNA to fragmentation that

manifests, not immediately, but over time following thawing. In most mammalian species,

including but not limited to the koala (Johnston et al. 2012a), stallion (López-Fernández et al.

2007), ram (López-Fernández et al. 2008a) and Asian elephant (Elephas maximus; Imrat et al.

2012), the basal level of sperm DNA damage observed immediately after thawing does not differ

significantly from that observed in fresh spermatozoa. However, the chromatin stability tends to

deteriorate more rapidly following thawing and incubation and can substantially limit the longevity

of the DNA.

1.7.1 Oxidative stress as a mechanism of DNA cryoinjury

There is strong evidence that the sperm cryopreservation procedure is associated with

increased generation of ROS and consequently, oxidative stress may be a significant mechanism of

sperm cryoinjury (Wang et al. 1997; Baumber et al. 2003; Martínez-Páramo et al. 2012).

Preparation of spermatozoa for cryopreservation involves dilution of the seminal plasma, the

predominant source of antioxidant protection, predisposing cryopreserved sperm to oxidative stress

(Baumber et al. 2003). Recent investigations have demonstrated that the addition of antioxidant

supplements to human semen can protect sperm DNA from oxidative injury during

22

cryopreservation and subsequent thawing (Thomson et al. 2009; Branco et al. 2010). Similarly,

significant decreases were observed in levels of sperm DNA fragmentation in the stallion (Baumber

et al. 2003), ram (Peris et al. 2007) and gilthead sea bream (Sparus aurata; Cabrita et al. 2011) that

had been treated with antioxidant-supplemented extenders prior to cryopreservation. In contrast,

Lahnsteiner et al. (2011) reported that supplementation of the cryopreservation extenders with

enzymatic and non-enzymatic antioxidants achieved only minor positive effects in brook trout

(Salvelinus fontinalis) and rainbow trout (Oncorhynchus mykiss) and concluded that oxidative

damage seems to play no role in cryoinjury. Interestingly, rainbow trout sperm exhibit a marked

resistance to cryopreservation-induced DNA damage, despite possessing protamines devoid of

cysteine residues responsible for the formation of disulphide bonds to stabilise and protect the

chromatin from oxidative attack (Labbe et al. 2001). The vastly different species-specific responses

to sperm cryopreservation and proposed mechanisms of cryoinjury remain open to interpretation, so

that making predictions about the effect of cryopreservation on the sperm of endangered

amphibians is difficult.

1.7.2 Osmotic stress and activation of sperm motility

It is possible that the dramatic changes in the osmotic environment associated with the

addition and removal of cryoprotectants contribute to the incidence of spermatozoa with fragmented

DNA post-thaw. During cryopreservation, the formation of extracellular ice crystals forms a hyper-

tonic environment resulting in the outflow of water across the plasma membranes of the

spermatozoa and a decrease in cellular volume; the removal of cryoprotectant upon thawing restores

the osmotic environment, causing a further influx of water and further damage to the sperm cell

(Johnston et al. 2012a). Although a number of studies have demonstrated the damaging effects of

aniso-osmotic media on spermatozoa in terms of plasma membrane integrity and motility (Blasse et

al. 2010), few have examined the effect of osmotic stress on sperm DNA. Johnston et al. (2012a)

recorded significant increases in the percentage of koala spermatozoa with fragmented DNA

following exposure to both hypo-osmotic and hyper-osmotic media. Furthermore, the authors

reported a significant correlation in DNA fragmentation between cryopreserved-thawed

spermatozoa and those exposed to a hypo-osmotic media. Similarly, hyper-tonic medium has been

shown to induce DNA fragmentation and apoptosis in fish epithelial cells (Hashimoto et al. 1998).

These observations differ from findings in rainbow trout where the sperm nucleus was shown to be

highly resistant to osmotic shock and maintained DNA stability after exposure to hyper-osmotic

cryoprotectant (Labbe et al. 2001). It is likely that species-specific differences to the structure of the

23

spermatozoa, particularly plasma membrane composition, determine their capacity to withstand

changes in osmolality during cryopreservation.

The need to activate motility in amphibian spermatozoa prior to use with ART imposes

further osmotic stress and is a significant factor impacting the longevity of both fresh and

cryopreserved-thawed spermatozoa. Spermatozoa from externally fertilising species, such as

amphibians and fish, remain inactive in vivo due to the isotonic environment of the testes (Kouba et

al. 2009; Browne et al. 2015). Activation of sperm motility in fresh water aquatic breeding anurans

occurs by spawning into hypo-tonic fresh/pond water with a low osmolality. Wolf and Hendrick

(1971) noted that the maximum rate of fertilisation in the African clawed frog was restricted to only

10 – 15 minutes following activation of fresh spermatozoa, while spermatozoa stored in the

inactivated state at 0 °C in isotonic DeBoers solution retained the capacity to fertilise for up to 24

hours. Remarkably, cane toad (Browne et al. 2001) spermatozoa showed evidence of fertilisation

after ten days of cold storage (at 0 °C) in simplified amphibian ringer solution (SAR). Although

delaying activation of sperm motility may prolong the fertilising capability of amphibian

spermatozoa, the progression of SDF during in vitro incubation is likely to impair embryo survival

(Gosálvez et al. 2014). For example, oocyte fertilisation rate in zebrafish was not dramatically

affected by using spermatozoa that had been subjected to controlled DNA damage, however,

embryo viability was significantly reduced (Gosálvez et al. 2014). Similarly, Pérez-Cerezales et al.

(2010) reported that single- and double-stranded DNA breaks and oxidised bases in cryopreserved

rainbow trout spermatozoa induced embryo developmental failures during gene expression and

resulted in a high abortion rate. These observations highlight the variable fertility of amphibian

spermatozoa handled ex vivo and reinforce the idea of using the spermatozoa for ART as quickly as

possible following activation.

1.7.3 Fertilisation success following cryopreservation

While few amphibian studies have used cryopreserved gametes for ART, the general

assumption is that fresh spermatozoa have a greater fertilisation capacity than frozen-thawed

spermatozoa. Interestingly, the spermatozoa of anuran amphibians do not appear to undergo cold-

shock comparable to mammalian spermatozoa when rapidly exposed to low temperatures; in fact,

many artificial fertilisation protocols recommend storage at 0 – 4 °C to prolong sperm viability

(Hollinger and Corton 1980; Mansour et al. 2009). Numerous amphibian studies have achieved

consistently high fertilisation rates using chilled fresh sperm samples, while cryopreserved

24

amphibian spermatozoa have achieved vastly mixed results (Browne et al. 1998; Buchholtz et al.

2004; Sargent and Mohun 2005). For example, in the African clawed frog, evidence of fertilisation

using fresh spermatozoa was observed in 80 – 93% of oocytes (Wolf and Hedrick 1971 and

Hollinger and Corton 1980, respectively), while Sargent and Mohun (2005) reported that the

number of oocytes fertilised using cryopreserved spermatozoa (23 – 81%) was never as great as in a

control fertilisation. The remarkable variation in fertilisation rates achieved following

cryopreservation highlights the importance of considering different freeze-thaw, as well as artificial

fertilisation protocols when comparing the results of different studies. It would be particularly

interesting to explore the relationship between amphibian sperm DNA damage resulting from

cryoinjury and associated embryo development; this may help to explain those situations where

artificial fertilisation failed to achieve viable embryos, despite the successful recovery of sperm

motility post-thaw.

Although the impact of cryopreservation on amphibian sperm DNA remains to be explored

in greater detail, it is well documented that the process of cryopreservation induces the impairment

of other parameters of sperm quality likely to impact fertilisation capacity, such as motility,

viability and plasma membrane integrity. The transition of intracellular water to ice during the

freezing process causes detrimental biophysical changes to the spermatozoa, particularly to the

plasma membrane and mitochondria (Martínez-Páramo et al. 2012). The mitochondrial collar is

integral to progressive movement of anuran spermatozoa and is particularly susceptible to

rupture/loss as a result of cryoinjury; Kouba and Vance (2009) noted the absence of the

mitochondrial collar in 90% of cryopreserved-thawed Bufo spp. spermatozoa. Labbe et al. (2001)

reported a very low increase in the proportion of rainbow trout spermatozoa with altered DNA after

cryopreservation compared to the 20 fold increase in the proportion of spermatozoa reported as

having damaged membranes and mitochondria (DeBaulny et al. 1997). Similarly, a marked

decrease in motility and viability was observed in African clawed frog spermatozoa immediately

post-thaw (Sargent and Mohun 2005; Mansour et al. 2009).

1.7.4 The influence of seasonal breeding on DNA integrity

Recent studies have suggested that susceptibility of sperm chromatin to cryopreservation

may vary throughout the year and that initial evaluation of fresh spermatozoa may not always

provide an accurate indication of the fertilising potential after freezing and thawing (Costanzo et al.

1998; Pérez-Cerezales et al. 2010). One of the factors affecting cryopreservation success is the

25

timing of sperm collection during the reproductive season. In seasonal breeding fish, the quality of

the spermatozoa fluctuates throughout the spawning season and increased levels of DNA damage

have been observed in frozen-thawed rainbow trout spermatozoa collected outside the peak

spawning period (Pérez-Cerezales et al. 2010). This increase in DNA damage could be related to

the ageing of spermatozoa stored in the ducts and the presence of immature, apoptotic or necrotic

sperm cells (Pérez-Cerezales et al. 2010). Costanzo et al. (1998) examined freeze tolerance in Rana

spp. and reported an increased susceptibility to cryoinjury in the freeze-tolerant wood frog (Rana

sylvatica) when exposed to sporadic, but naturally occurring episodes of subzero temperatures

shortly prior to the spawning period. Consequently, species-specific breeding seasons may

influence the quality of frozen-thawed spermatozoa and prove an important consideration when

collecting and cryopreserving amphibian sperm with the intention of utilising ART.

1.8 Review of technologies used to assess DNA integrity

A series of assays have been designed to evaluate the integrity of sperm DNA, including

tests for the assessment of DNA dependent proteins and for the detection of DNA breaks. Originally

developed and validated for investigating DNA damage in somatic cells, these methods have been

adapted to enable the reagents involved to access the more highly compacted DNA of spermatozoa

(Sharma and Agarwal 2011). Sperm DNA fragmentation can be assessed directly using

methodologies that employ enzymatic reactions to incorporate fluorescent labels to single- or

double-stranded DNA breaks, such as terminal deoxynucleotidyl transferase-mediated dUTP nick

end labeling (TUNEL) and in situ nick translation (ISNT), or using indirect methods that assess the

susceptibility of the sperm chromatin to DNA denaturation, including the single-cell gel

electrophoresis assay (comet assay), SCD test and sperm chromatin structure assay (SCSA;

Gosálvez et al. 2014). Table 1.2 highlights the primary advantages and disadvantages of each test.

The application of these assays has been driven mainly by the need for ART for commercial

breeding, endangered species conservation and treatment of human infertility. As DNA integrity is

typically poorly correlated with classical parameters of semen quality, this important component of

sperm viability would remain cryptic without the means to identify potential damage (López-

Fernández et al. 2008b).

26

Table 1.2. Summary of the tests used to analyse the integrity of sperm chromatin, including a description of the assay principle, detection methods and their main strengths and weaknesses (Tavalaee et al. 2008; Sharma and Agarwal 2011).

Assay Assay principle Detection Strengths Weaknesses

SCSA Susceptibility of DNA to heat or acid denaturation

Flow cytometry

Rapid, large number of spermatozoa counted by flow cytometry, highly sensitive and reproducible, objective

Special equipment required (flow cytometer)

TUNEL

Single and double-stranded DNA breaks

Fluorescence microscopy/ flow cytometry

Simple and fast, highly sensitive, large number of spermatozoa counted by flow cytometry

Variable protocols, special equipment required (flow cytometer)

COMET

Single and double-stranded DNA breaks (alkaline) or double-stranded DNA breaks only (neutral)

Fluorescence microscopy

Simple and inexpensive, highly sensitive

Labour intensive, subjective, variable protocols, requires imaging software and experienced technician

SCD

DNA dispersion halo

Fluorescence microscopy

Cheap and simple to perform

Subjective, possible fluorochrome fading

The predictive value of the assays in determining the fertilising capacity of spermatozoa is

of primary interest and appears to vary depending on whether actual DNA damage such as strand

breaks or susceptibility of the DNA to imposed denaturation (potential damage) is being measured

(Marchesi et al. 2007). The lack of standardised protocols can make it difficult to determine

whether observed differences in the extent of chromosomal abnormalities or DNA damage are real

(related to biology) or due to differences in testing methods and this should be taken into

consideration when comparing data across laboratories. Sperm preparation technique is paramount

and must be appropriately controlled to limit the potential for discrepancies in DNA integrity

between the examined sperm population and the neat semen (Marchesi et al. 2007). Furthermore,

inter-species application is likely to cause additional variation due to the species-specific protamine

compositions of the sperm chromatin, highlighting the importance of validating the assay when

applying a protocol to a novel species (Gosálvez et al. 2011a). A key consideration of any test is the

number of cells counted; counting higher numbers of spermatozoa allows for greater accuracy and

thereby any assay based on a low cell count will yield significantly wide confidence intervals

(WHO 2010).

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1.8.1 Sperm chromatin structure assay (SCSA)

The SCSA is one of the most widely utilised tests of sperm DNA damage and remains the

test for which most data are available, albeit the majority pertain to human spermatozoa. Developed

over 30 years ago, the SCSA has been used to measure over 100 000 animal and human sperm

samples (Evenson 2011). The SCSA identifies spermatozoa with abnormal chromatin packaging by

measuring the susceptibility of DNA to heat or acid induced denaturation in situ (Evenson et al.

1999). This test involves treating raw semen with a low pH buffer that denatures the DNA at sites

where strand breaks occur, inducing further damage to poorly packaged chromatin (Marchesi et al.

2007). The sample is then stained with acridine orange and processed by flow cytometry. Acridine

orange is a metachromatic dye that intercalates with the native (double-stranded) DNA structure to

fluoresce green; association with single-stranded DNA causes the dye to bind externally and emit a

red fluorescence (Marchesi et al. 2007; Tavalaee et al. 2008). Barratt et al. (2010) suggest that the

emitted red fluorescence may not be limited to pre-existing DNA damage and that sperm with

inherently lower structural chromatin stability may be more vulnerable to acid induced denaturation

and therefore, more likely to yield a false positive result. This is evidenced by Dias et al. (2006)

observation that caput stallion spermatozoa with condensed, but un-stabilised chromatin were more

sensitive to acid induced DNA denaturation than mature cauda spermatozoa. Additionally, the

SCSA does not account for inherent species-specific differences in chromatin structure that may

render the DNA more susceptible or resistant to acid denaturation, such as the presence of

protamine-mediated disulphide bonds that act to stabilise the sperm chromatin and may thereby

restrict access to the DNA molecule. In this respect, the SCSA differs from the comet assay and

SCD test that include a pre-treatment step employing a reducing agent to specifically loosen the

disulphide bonds protecting the chromatin (Cortés-Gutiérrez et al. 2014b).

The results of the SCSA are expressed as the DNA Fragmentation Index (DFI), representing

the sperm fraction with single stranded breaks and High DNA Stainability (HDS), characterising

immature spermatozoa with increased accessibility to the acridine orange dye due mainly to

dysfunctional chromatin condensation (Tavalaee et al. 2008; Evenson 2011). An advantage of the

SCSA, shared also by the TUNEL assay is the large number of spermatozoa assessable by flow

cytometry; this provides greater accuracy and makes this technique statistically robust, objective

and highly reproducible (Barratt et al. 2010). However, this assay is restricted to laboratories that

have a flow cytometer and highly skilled technicians (Agarwal and Said 2004).

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1.8.2 Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL)

The TUNEL assay was first described by Gorczyca et al. (1993) for the detection of DNA

strand breaks in human spermatozoa. The TUNEL assay detects both single and double-stranded

DNA breaks or ‘nicks’ by tagging the free 3’-OH end of the affected chain with labelled nucleotides

(dUTP) in an enzymatic reaction catalysed by terminal deoxynucleotidyl transferase (TdT) (Chohan

et al. 2006; Marchesi et al. 2007). The incorporation of dUTP is quantified by fluorescence

microscopy or flow cytometry and results are expressed as the percentage of spermatozoa that are

TUNEL positive; spermatozoa with normal DNA will have only background fluorescence, while

the more dUTP that is incorporated, the greater the number of DNA lesions (Evenson and Wixon

2006; Marchesi et al. 2007).

The TUNEL assay is a reproducible and inexpensive technique for assessing DNA

fragmentation in spermatozoa and results of studies on human spermatozoa have been shown to

correlate well with the SCSA (Aravindan et al. 1997; Chohan et al. 2006). However, the assay has

been criticised due to its many variations in methodology, limiting the value of results compared

across different laboratories (Evenson and Wixon 2006; Muratori et al. 2010ab). Most TUNEL

protocols use the fixative formaldehyde to prepare sperm for dUTP labelling. Muratori et al.

(2010b) identified that DNA fragmentation in the same semen sample was greatly affected by the

concentration and length of incubation in the fixative and suggested that formaldehyde may have a

damaging action on the sperm chromatin, particularly in spermatozoa with poor or absent

protamination, such as in many amphibians. However, it is also possible that the addition of

formaldehyde may simply have improved the efficiency of DNA labelling and thus the detection of

more underlying DNA damage. Mitchell et al. (2011) suggests that the conventional TUNEL assay

is in fact not sensitive enough and that the highly compacted nature of the sperm chromatin impedes

incorporation of dUTP labelling. Treatment with the disulphide bond reducing agent dithiothreitol

(DTT) to relax the chromatin prior to fixation appears to enhance the responsiveness of the TUNEL

assay as well as the strength of the signals produced. However, this modification is contradictory to

the accepted belief that TUNEL detects ‘real’ and not imposed DNA damage and further highlights

the need for a standardised protocol.

29

1.8.3 Single-cell gel electrophoresis (SCGE) or comet assay

The comet assay or single-cell gel electrophoresis enables the analysis of DNA damage in

an individual cell following lysis of the nuclear proteins binding the DNA (Barratt et al. 2010).

Ostling and Johanson (1984) first used the technique under neutral buffer conditions to visualise the

migration of double-stranded DNA loops from irradiated somatic cells in the form of a tail

unwinding from the relaxed nucleus. Singh et al. (1988) subsequently modified the assay to

incorporate alkaline electrophoresis buffers to deproteinize the nuclei and increase sensitivity to

detect both single and double-stranded DNA breaks without distinguishing between the two

(Agarwal and Said 2004). Electrophoresis causes DNA fragments to migrate away from the central

DNA core. Double-stranded DNA that contains no lesions will remain in the comet head, whereas

shorter fragments of nicked single and/or double-stranded DNA migrate to produce a tear drop-

shaped comet tail (Zini and Libman 2006; Marchesi et al. 2007; Figure 1.4b and c). DNA breaks are

evaluated by measuring the number of spermatozoa with migration comets, as well as the length of

the comet tail and/or the percentage of DNA present in the tail (Cotelle and Férard 1999; Agarwal

and Said 2004).

Enciso (2009) further proposed a two dimensional comet assay to achieve a simultaneous

characterisation of single and double-stranded DNA breaks, allowing a more precise and

comprehensive analysis of DNA damage in the individual spermatozoon. The aptly named two-

tailed or double-comet assay is a modification of the original neutral comet assay, whereby a second

(alkaline) electrophoresis step is performed on the same slide after an offset in orientation of 90°;

this enables the visualisation of comets on both the X (double-stranded DNA) and Y-axis (single-

stranded DNA) for a single spermatozoon (Enciso 2009; see Figure 1.4c). The ability to

differentiate between both single and double-stranded DNA breaks, raises the potential to relate

impaired fertility to the type of DNA damage present; double-stranded DNA breaks are generally

considered the most genotoxic forms of DNA damage with the greatest influence on fertility (Zee et

al. 2009).

While the comet assay is simple, fast to perform and produces results comparable to the

SCSA and TUNEL assays (Aravindan et al. 1997), the labour-intensive process of analysis requires

a skilled technician to accurately interpret the results with a level of reproducibility, limiting its use

in the clinical ART setting (Simon and Lewis 2011). Additional confusion comes from the lack of

standardisation of the parameters used to calculate the percentage of spermatozoa with damaged

30

DNA; reliance on fluorescence microscopy not only limits the number of spermatozoa able to be

analysed, there is also the potential for subjectivity to bias results without the use of image analysis

software (Fairbairn et al. 1995). The comet assay is currently primarily used for genotoxic studies

on human and animal somatic cells (including amphibian erythrocytes and spermatozoa; Simon and

Lewis 2011 and Pollock et al. 2015 respectively) and is frequently used to validate the type of DNA

damage (whether single or double-stranded) identified by the SCD test (López-Fernández et al.

2009; Zee et al. 2009; Pollock et al. 2015).

Figure 1.4. Fluorescence photomicrographs of DNA fragmentation in African clawed frog (modified from Pollock et al. 2015) spermatozoa evidenced by different assay techniques; (a) SCD test showing sperm nuclei with coiled cores and small, compact halos of dispersed chromatin representing non-fragmented DNA. Arrow indicates spermatozoon with a large, stellar halo of dispersed chromatin characteristic of fragmented DNA; (b and c) Comet morphologies revealed following the double (neutral + alkaline) comet assay; (b) sperm nucleus with a defined nuclear core and compact comet tail representing structural single-stranded DNA breaks; (c) comets comprised of both single- and double-stranded DNA breaks indicating extensive DNA damage in a single sperm nucleus.

31

1.8.4 Sperm chromatin dispersion (SCD) test

The SCD test is a relatively new addition to the assortment of techniques used to assess

sperm DNA integrity. This procedure measures the susceptibility of sperm DNA to fragmentation

and is based on the principle that controlled denaturation and removal of nuclear proteins results in

nucleoids with a central core and a peripheral halo of chromatin dispersion (Fernández et al. 2003;

Figure 1.4a). The halo corresponds to the spreading of relaxed nuclear DNA loops attached to

protein residues and in damaged spermatozoa, fragments of broken DNA (Gosálvez et al. 2011a).

In the original Halosperm® protocol developed for humans, spermatozoa are incubated in hydrogen

chloride for acid denaturation prior to treatment with a lysing solution to remove the nuclear

membranes and proteins (Fernández et al. 2003). Using this technique, spermatozoa with

fragmented DNA show restricted loop dispersion with very limited or absent haloes, while

spermatozoa with undamaged and properly condensed chromatin produce large halos of relaxed

DNA loops (Fernández et al. 2003). Quantifying the size and the incidence of the halos by

fluorescence or bright-field microscopy provides an index of the amount of DNA damage present.

Enciso (2006) first identified the need to consider inter-specific differences in chromatin

structure and/or organisation in sperm cells when applying the SCD to non-human spermatozoa;

using the Halosperm® protocol, undamaged boar sperm appeared highly resistant to releasing the

compacted DNA loops, making discrimination between halo sizes difficult. Subsequently, the

Sperm-Sus-Halomax® was developed as a variant of the SCD test, where the acid treatment was

removed and a species-specific modified lysing solution used for protein depletion; in this version

of the SCD test, spermatozoa with fragmented DNA produced large halos of chromatin dispersion,

while those with non-fragmented DNA showed very limited relaxation of the DNA loops (Enciso

2006). The SCD test has since been adapted to the unique chromatin structure of a range of

domestic and wildlife species, including the short-beaked echidna (Johnston et al. 2009), koala

(Johnston et al. 2007), stallion (López-Fernández et al. 2007 and donkey; Cortés-Gutiérrez et al.

2008a), boar (López-Fernández et al. 2008b), rhinoceros spp. (Portas et al. 2009), fish (López-

Fernández et al. 2009; Gosálvez et al. 2014) and most recently in the African clawed frog (Pollock

et al. 2015).

Due to its recent development, limited data exists comparing results obtained using the SCD

with other tests, however, strong correlations have been shown with the SCSA (Fernández et al.

2005; Muriel et al. 2006), as well as both the SCSA and TUNEL (Chohan et al. 2006). The SCD

32

test is inexpensive and can be conducted over a short period of time without the need for complex

and expensive laboratory equipment (Gosálvez et al. 2011a). Fluorochrome fading may limit the

repeatability of the assay over time, however, the SCD can also be performed with brightfield

staining methods. Manual assessment of DNA fragmentation restricts the number of spermatozoa

able to be counted and may impede the accuracy of the SCD test compared to assays that involve

flow cytometry. While the SCD test is simple to perform, adequate training and routine proficiency

testing is recommended to reduce the potential for intra-observer and inter-observer variability in

scoring of DNA fragmentation (Chohan et al. 2006). As part of the development of the SCD for

each novel species, the SCD is routinely validated using ISNT and comet assay protocols.

1.9 Conclusion

This review gathered current information on amphibian sperm chromatin structure and

behaviour in light of sperm preservation. Understanding the effect of the freeze-thawing procedure

on sperm DNA stability is of critical importance for the conservation of threatened and endangered

amphibians. Despite extensive literature detailing sperm chromatin organisation and DNA

fragmentation in mammalian species, similar knowledge pertaining to amphibians is extremely

sparse. The highly diverse protein composition of amphibian and lower vertebrate spermatozoa

more generally reinforces the importance of developing species-specific DNA fragmentation

assays. The relationship between chromatin structure and the associated stability of the sperm

chromatin is complex and requires further examination, particularly with respect to the

presence/absence of cysteine residues and the arginine content of the SNBPs, as well as constitutive

DNA modifications such as ALS. Comparative analysis of sperm DNA fragmentation dynamics in

different species highlights the varying resistance of sperm chromatin to external stressors and is an

important consideration when manipulating amphibian spermatozoa in vitro for ART. Continued

research into the application of DNA fragmentation assays to amphibian spermatozoa, particularly

with respect to cryoinjury, would not only provide additional evidence of this phenomenon in a new

major taxon, but has the potential to improve assisted reproductive technologies used for the

conservation of threatened and endangered amphibian species. Although significant advancements

have been made in the area of amphibian ART, there is an urgent need to investigate the impact of

cryopreservation on the longevity of amphibian sperm DNA integrity. Although it has still not been

clearly established whether there exists a direct relationship between cryopreservation and sperm

DNA damage, there is strong evidence to suggest that the freeze-thaw procedure can substantially

limit the structural integrity of the DNA over time. Additionally, investigation into the relationship

33

between sperm DNA fragmentation in amphibian spermatozoa and the resulting impact on

fertilisation rate and embryo development will provide insight into the mechanisms leading to

reproductive failure and may in turn prove useful for improving the reproductive output of

amphibian ART.

1.10 Thesis aims and structure


amphibians and only a single study has specifically examined DNA fragmentation in the

spermatozoa (Morrow et al. 2014; poster presented at the World Congress of Reproductive

Biology). A DNA fragmentation assay applied and validated for amphibian spermatozoa would not

only provide additional evidence of this phenomenon in a new major taxon, but also prove a

valuable tool for developing improvements in ART used in the captive propagation of threatened

and endangered species. Consequently, the fundamental aim of this project was the development

and validation of the SCD test for amphibian spermatozoa with the purpose of examining the post-

thaw survival of cryopreserved spermatozoa. This objective was addressed through the following

chapters.

Chapter 1: Sperm chromatin in the Amphibia – presented as submitted to the Journal of

Herpetology

Development of a species-specific protocol for DNA fragmentation assessment in a novel

species, such as the amphibian, requires an understanding of the basic structure and protein

composition of the spermatozoa. This review aimed to summarise the information available on

amphibian sperm chromatin organisation, as well as explore the etiology of DNA fragmentation.

Prior to the commencement of this candidature, no studies had examined chromatin damage in

amphibian spermatozoa. This review was submitted for publication prior to thesis submission and

consequently contains references to data presented in Chapter 2, in order to present the most up to

date information pertaining to amphibian sperm DNA fragmentation assessment.

Chapter 2: Validation of the sperm chromatin dispersion (SCD) test in Xenopus laevis – presented

as published in Reproduction, Fertility and Development 27, 1168-1174.

34

The first step towards using the SCD procedure as an investigative tool of amphibian sperm

biology is the co-validation of the technique up against other methods of assessing DNA damage.

This experiment applied the SCD test to an amphibian sperm model (African clawed frog) using a

species-specific lysing solution for protein depletion. Results of the SCD test were then validated

using in situ nick hybridisation with DNA-specific molecular probes (ISNT) and the double comet

assay. The double comet assay characterised both single- and double-stranded DNA breaks

simultaneously in the same spermatozoon. Sperm microgel slides were prepared for ISNT and the

double comet assay using the same species-specific lysing solution for protein depletion as in the

SCD test. While an additional study has since investigated DNA fragmentation in amphibian

spermatozoa prior to the submission of this thesis (Morrow et al. 2014; poster presented at the

World Congress of Reproductive Biology), experiments presented in Chapter 2 provided the first

evidence of amphibian sperm DNA damage.

Chapters 3 and 4: Dynamics of sperm DNA fragmentation in Xenopus laevis and Bufo marinus –

presented as prepared for submission to Reproduction, Fertility and Development

These investigations applied the SCD test to cryopreserved-thawed amphibian spermatozoa

in an attempt to provide insight into cryoinjury of the sperm chromatin and the resulting effect on

the potential reproductive output of ART. This was achieved by examining the dynamic changes to

the basal level of DNA fragmentation in fresh African clawed frog (Chapter 3) and cane toad

(Chapter 4) spermatozoa in comparison with the post-thaw integrity of cryopreserved sperm DNA.

Freezing additional samples of African clawed frog and cane toad spermatozoa without

cryoprotectant provided further insight into the mechanisms of cryoinjury. An IVF procedure using

cryopreserved African clawed frog spermatozoa was used to assess the extent to which freeze-

thawing was correlated with a decline in fertilisation rate. Chapter 4 also examined the relationship

between cryoinjury and other parameters of sperm quality, such as motility and viability. SYBR-14

and propidium iodide are fluorescent membrane-permeant nucleic acid stains that were used in

conjunction with fluorescence microscopy to differentiate live and dead spermatozoa.

35

CHAPTER 2: Validation of the sperm chromatin dispersion (SCD)

test in the Amphibian Xenopus laevis using in situ nick translation and

comet assay

Pollock, K.1, Gosálvez, J.2, Arroyo, F.2, López-Fernández, C.2, Guille, M.3, Noble, A.3, and

Johnston, S.D.1

1School of Agriculture and Food Science, The University of Queensland, Gatton, Australia 4343. 2Department of Biology, Genetics Unit, The Autonomous University of Madrid, Madrid, Spain

20849. 3European Xenopus Resource Centre, The University of Portsmouth, Portsmouth, United Kingdom

PO1 2DY.

Presented as published in Reproduction, Fertility and Development 27, 1168-1174.

36

2.1 Abstract

The integrity of sperm DNA is becoming increasingly recognised as an important parameter

of semen quality, but there are no published reports of this procedure for any amphibian. The

primary aim of this study was to apply a modified sperm chromatin dispersion (SCD) test

(Halomax) to an amphibian sperm model (African clawed frog; Xenopus laevis) and to validate the

assay against in situ nick translation (ISNT) and the double-comet assay procedure. Inactivated

spermatozoa were collected from fresh testes (n = 3). Sperm DNA fragmentation (SDF) for each

sperm sample was conducted immediately following activation (T0) and again after one hour (T1)

and 24 h (T24) of incubation at room temperature in order to produce a range of spermatozoa with

differing levels of DNA damage. The SCD procedure resulted in the production of three nuclear

morphotypes; amphibian sperm morphotype 1 (ASM-1) and ASM-2 showed no evidence of DNA

damage, whereas ASM-3 spermatozoa were highly fragmented with large halos of dispersed DNA

fragments and a reduced nuclear core. ISNT confirmed that ASM-3 nuclei contained damaged

DNA. There was a significant correlation (r = 0.9613) between the levels of ASM-3 detected by the

SCD test and SDF revealed by the double-comet assay.

2.2 Introduction

The integrity of sperm DNA is becoming increasingly recognised as an important parameter

of semen quality and essential for embryonic development and ultimately reproductive success

(D’Occhio et al. 2007). During spermiogenesis, the DNA molecule undergoes a series of structural

changes that allow the chromatin to be remodelled into a highly compacted form. This tight

configuration is susceptible to disruption by sperm DNA fragmentation (SDF). SDF is regarded as

damage to the topology and function of the chromatin complex via single- or double-stranded DNA

breaks that ultimately lead to impaired fertility (López-Fernández et al. 2009). SDF may include

both inherent constitutive DNA modifications that are observed immediately following sperm

emission, mostly associated with the normal chromatin remodelling process, and inducible or

iatrogenic modifications to the DNA that arise as a function of incubation time in vitro and which

are a result of external factors such as the conditions of sperm storage during processing for assisted

reproductive technologies (ART; Gosálvez et al. 2011a; Alvarez and Gosálvez 2012).

As DNA integrity is poorly correlated with standard parameters of semen quality, a means

to identify existing or potential damage to the inheritable genetic material is a necessary component

37

of ART (López-Fernández et al. 2008b). The sperm chromatin dispersion (SCD) test is based on the

principle that controlled denaturation and removal of nuclear proteins using a lysing solution results

in nucleoids with a central core and a peripheral halo from the dispersion of damaged chromatin

(Fernández et al. 2003). The halo corresponds to relaxed DNA loops attached to protein residues

and in damaged spermatozoa the halo is larger, more diffuse and formed by discrete DNA

fragments (Gosálvez et al. 2011a). Quantifying the size and the incidence of these haloes by

fluorescence or brightfield microscopy provides an index of the amount of DNA damage present in

the sample. Given the vast differences in the protein composition and structure of sperm DNA

across the different vertebrate taxa, it is not surprising that species-specific protocols are required to

ensure a targeted and validated approach to DNA fragmentation assessment (Gosálvez et al.

2011a).

Originally developed for assessing DNA fragmentation in human spermatozoa (Halosperm;

Fernández et al. 2003), the SCD test (as the Halomax) has now been successfully adapted to the

unique chromatin structure of a range of domestic and wildlife species including the koala

(Phascolarctos cinereus; Johnston et al. 2007), short-beaked echidna (Tachyglossus aculeatus;

Johnston et al. 2009), stallion (López-Fernández et al. 2007), donkey (Equus asinus; Cortés-

Gutiérrez et al. 2008a), boar (López-Fernández et al. 2008b), rhinoceros (Diceros bicornis,

Rhinoceros unicornis and Ceratotherium simum; Portas et al. 2009) and bony fish (Tinca tinca;

López-Fernández et al. 2009); the SCD test was validated in each of these species separately and

employs a species-specific modified lysing solution for protein depletion.


amphibians, with the majority of this work focussed on the effects of environmental toxicity on

somatic cells (Wang and Jia 2009; Maselli et al. 2010; Valencia et al. 2011); there have been none

that have specifically examined DNA fragmentation in amphibian spermatozoa. A DNA

fragmentation assay applied and validated for amphibian spermatozoa would be a useful tool for not

only comprehending the importance of this phenomenon in a new major taxon, but also for

understanding the impact of changing environmental factors and disease on amphibian

spermatogenesis and for developing improvements for use of the spermatozoa in ART, used in the

captive propagation of threatened and endangered species. For example, in vitro fertilisation (IVF)

in amphibians is becoming an increasingly useful protocol as part of species propagation used to

support the genetic management and reproductive output of amphibian breeding programs (Kouba

and Vance 2009).

38

Recent studies have demonstrated that SDF is, in fact, a highly dynamic process and

typically increases from the time of ejaculation or thawing following cryopreservation up until the

time of fertilisation (Gosálvez et al. 2011b). This dynamic loss of DNA integrity is particularly

relevant for externally fertilising vertebrates such as fish (López-Fernández et al. 2009), where

spawning in fresh or seawater osmotically initiates sperm activation in order for external

fertilisation to occur. It is likely that once activation has commenced there could also be a

consequent rapid loss of DNA integrity; this phenomenon has never been examined but is highly

relevant when developing and refining protocols for sperm conservation. The first step towards

using the SCD procedure as an investigative tool of amphibian sperm biology is the co-validation of

the technique with other methods of assessing DNA damage. Therefore, the aim of this study was to

apply the SCD test to an amphibian sperm model (African clawed frog) and then to validate these

results against other procedures for assessing DNA fragmentation, i.e. DNA breakage labelling and

the comet assay. In situ nick translation (ISNT) involves the direct incorporation of labelled

nucleotides to the free ends of either single- or double-stranded DNA breaks. The comet assay uses

a lysing solution to release the nuclear proteins binding the DNA, followed by electrophoresis,

which causes DNA fragments to migrate away from the central DNA core.

2.3 Materials and methods

2.3.1 Animals, sperm recovery and experimental design

For this study, three male wild-type African clawed frogs were sourced from a breeding

facility maintained at the European Xenopus Resource Centre (EXRC, Portsmouth, United

Kingdom) with controlled conditions for housing, photoperiod, temperature, water quality and

feeding. All animals were healthy, with a history of reproductive success. Sperm samples were

recovered according to EXRC protocol (EXRC 2012). Briefly, animals were euthanased by an

overdose of Tricaine–MS222 (Sigma Aldrich, Gillingham, UK) and the whole testes dissected into

1.0 M modified Barth’s saline (MBS) at room temperature. A portion of the testis was macerated by

gentle application of an Eppendorf pestle in 200 µL of chilled (4 °C) 0.1 M MBS to achieve sperm

activation and attain a final concentration of ~10 × 106 spermatozoa mL–1. Identification of sperm

motility under 400× magnification confirmed activation. Analysis of SDF for each sperm sample

was conducted immediately following activation (T0) and again after one hour (T1) and 24 h (T24)

of incubation at room temperature. The restricted incubation period of 1 h reflected the rapid

39

decline in the fertilisation capacity and motility of African clawed frog spermatozoa following

activation (Wolf and Hedrick 1971; Sargent and Mohun 2005).

2.3.2 Sperm chromatin dispersion test

The extent of DNA fragmentation in each sample was assessed with a prototype kit

(Halomax Proto-Xenopus) adapted from the Halomax kit (Halotech SL, Madrid, Spain) using the

following method. Aliquots of 50 µL of low-melting-point agarose in Eppendorf Safe Lock tubes

(Eppendorf, Hamburg, Germany) were suspended on a float in a water bath at 90 – 100 °C for five

minutes to melt the agarose. The temperature of the agarose was then equilibrated to 35 °C in a

second water bath before 25 µL of the sperm suspension (spermatozoa in 0.1 M MBS) was added

and gently mixed. Aliquots (10 µL) of the spermatozoa–agarose mixture were pipetted onto pre-

treated glass slides (provided in the Halomax kit) and covered with a glass coverslip (18 × 18 mm).

The slides were then placed on a chilled metallic plate (4 °C) and left to solidify into a thin microgel

for five minutes in a refrigerator at 4 °C. The coverslips were gently removed and the slides

promptly introduced (maintained at a horizontal position) into a plastic bath of lysing solution

provided in the Halomax kit for five minutes to deproteinise the sperm chromatin. The slides were

removed from the lysing solution and washed in a bath containing dH2O for five minutes and then

sequentially dehydrated in increasing concentrations of ethanol (70, 90 and 100%) for two minutes

each and finally left to air-dry. For visualisation under fluorescence microscopy the slides were

mounted with equal parts SYBR Green 1 (Thermo Fisher Scientific Inc., Munich, Germany) and

Vectashield mounting medium (Vector Laboratories, Burlingame, California, USA) to inhibit

fluorochrome fading. A total of 300 spermatozoa were counted per sample using a Leica DMRB

epifluorescence microscope (Leica Microsystems, Barcelona, Spain) equipped with single-band

fluorescence block filters for green (fluorescein isothiocyanate (FITC) equivalent) and red

(Cyanine3 NHS ester (Cy3) equivalent) fluorescence. The percentage of spermatozoa with

fragmented DNA was calculated.

2.3.3 In situ nick translation (ISNT)

For ISNT, sperm microgel slides were prepared at T0 as described previously for the SCD

test and then subsequently processed for ISNT (López-Fernández et al. 2009). After protein lysis,

the slides were washed four times in phosphate-buffered saline (PBS) for five minutes and then

incubated in reaction buffer for DNA polymerase I (10 mM TRIS-HCl, 5 mM MgCl2 and 7.5 mM

40

Dithiothreitol (DTT); pH 7.5) for five minutes. Next, 100 µL of reaction buffer containing 25 units

of DNA polymerase I and a nucleotide mix including digoxigenin-11-dUTP were pipetted onto the

slide, covered with a glass coverslip (18 × 18 mm) and incubated in a moist chamber at 37 °C for 30

min. The slides were then washed in Tris–borate–EDTA (TBE) buffer (89 mM boric acid and 2.5

mM EDTA, Tris-base; pH 8.3) and finally dehydrated in sequential ethanol baths (70, 90 and

100%) and air-dried. The incorporated nucleotide was detected by incubation (30 min) with anti-

digoxigenin-FITC (Roche, Madrid, Spain). Spermatozoa were visualised using propidium iodide (2

µg mL–1) in Vectashield mounting medium. All images were captured with a charge-coupled device

(CCD) camera (Leica DFC350 FX; Leica Microsystems).

2.3.4 Comet assay

In the present study, a two-directional or ‘double’ comet assay was used to characterise both

double-stranded breaks (DSB) and single-stranded breaks (SSB) simultaneously in the same

spermatozoon (Enciso et al. 2009). This approach enabled the visualisation of comets that are run

perpendicular to each other (DSB; horizontal direction and SSB; vertical direction). Sperm microgel

slides were prepared as described previously for the SCD test above. For the initial neutral

electrophoresis step, the slide was treated with the lysis solution provided in the Halomax kit for

five minutes to deproteinise the sperm chromatin and then removed and washed in 1 × TBE buffer

solution (0.089 M Tris, 0.089 M boric acid and 0.002 M EDTA) for 5 minutes. The slides were then

electrophoresed (8 min; 20 V) in 1 × TBE buffer to allow the DNA fragments resulting from DSB

to migrate away from the sperm nucleus towards the anode during application of the electrical

current. At the completion of electrophoresis, the slide was removed and placed in 0.9% NaCl

solution for two minutes. The slide was then transferred to a chilled (4 °C) alkaline solution (0.03 M

NaOH and 1 M NaCl; pH 12.5–13) for two and a half minutes to cleave the alkali-labile sites of the

DNA. The slide was introduced into a fresh alkaline buffer (0.03 M NaOH; pH 12.5–13) at room

temperature for the second electrophoresis (four min; 20 V) having been repositioned 90° clockwise

from its original position during the first electrophoresis step. This resulted in the migration of

DNA fragments arising from SSB towards the anode while the DNA fragments from DSB remained

in their original positions brought about by the first electrophoresis. Finally, the slide was washed

with neutralising buffer (0.4 M Tris-HCL; pH 7.5) for five minutes, followed by two minutes in 1 ×

TBE buffer and subsequent dehydration in a series of 70, 90 and 100% ethanol baths for two

minutes each. Comets of fragmented sperm DNA were visualised using GelRed (Biotium,

Hayward, California, USA).

41

2.3.5 Statistical analysis

A Pearson’s correlation was performed between the percentage of SDF revealed by the SCD

test and comet assay using the StatPlus:mac statistical analysis program for Mac OS Version 2009

(see www.analysis.com/en/).

2.4 Results

2.4.1 Sperm morphology after the SCD test

Sperm samples processed with the SCD test for the visualisation of DNA fragmentation

revealed three primary sperm morphotypes. Amphibian sperm morphotype type 1 (ASM-1)

maintained the normal slightly coiled filiform shape of the sperm head, showing only a small, dense

halo of chromatin dispersion, which corresponded to spermatozoa with intact DNA (Figure 2.1a).

ASM-2 was similar to ASM-1 in terms of chromatin dispersion, but the original morphology of the

nucleus had swollen or decondensed into a more spherical appearance (Figure 2.1b); while there

was clearly some structural change to the nucleus, the DNA of these nuclei was typically not

fragmented. At T0, immediately following activation, most of the spermatozoa displayed an

elongated coiled core, but after 1 h of incubation the proportion of spermatozoa with rounded

morphologies increased (compare Figures 2.1a and b). In contrast to ASM-1 and ASM-2, ASM-3

displayed a large, stellar halo of dispersed chromatin with multiple fragments of DNA (Figure

2.1a). Spermatozoa containing damaged DNA typically showed less fluorescence than non-

fragmented DNA as the DNA fragments were more widely dispersed in the microgel; i.e. the

nuclear core decreased in size as the DNA halo expanded. In some cases, the halo became

progressively difficult to visualise as the chromatin dispersed further from the nuclear core; the

nuclear core also became progressively smaller in size with greater DNA damage (Figure 2.1b).

2.4.2 In situ DNA labelling of DNA strand breaks

ISNT was employed to confirm whether the different halo morphologies of chromatin

dispersion identified by the SCD test occurred in conjunction with DNA strand breaks. In situ

labelling involved the direct incorporation of labelled nucleotides to the free 3’-OH ends resulting

from SSB or DSB. After DNA labelling, all sperm nuclei were positively labelled with green

fluorescence, but with varying intensities (Figure 2.1c).

http://www.analysis.com/en/

42

Figure 2.1. African clawed frog sperm nuclear morphotypes after the SCD test. Arrows indicate spermatozoa with large, stellar halos of dispersed chromatin corresponding to fragmented DNA (ASM-3). (a) Spermatozoa observed at T0 with coiled cores and small, compact halos of dispersed chromatin representing non-fragmented DNA (ASM-1). The spermatozoon containing fragmented DNA has retained a defined core. (b) A higher proportion of spermatozoa with round and slightly swollen cores was observed following 1 h of incubation at room temperature (ASM-2: arrow). Note the diminished core and halo of the fragmented spermatozoon. (c) In situ nick translation showing the direct incorporation of labelled nucleotides (green signal) in all African clawed frog spermatozoa processed with the SCD test including a single spermatozoon with fragmented DNA (arrow).

43

2.4.3 Double-comet assay

The double-comet assay revealed three distinct comet morphologies indicating different

degrees of severity in DNA damage (Figure 2.2). The first comet morphology (ACM-1) was

revealed only after the alkaline electrophoresis and manifested as comets with a consistent nuclear

core and tail size indicative of SSB (Figures 2.2a and b); these comets were observed in almost all

spermatozoa and were interpreted as DNA motifs that may be transformed into SSB after treatment

with alkaline solutions. The second comet morphology (ACM-2) was characterised by the

migration of DNA fragments in the same direction as structural SSB; however, the comets

possessed notably longer comet tails and a much reduced nuclear core (Figure 2.2e). We propose

that these SSB comets are likely to contain more single-stranded DNA breaks than what would

normally be regarded as structural SSB. The third comet morphology (ACM-3) was only revealed

following application of the neutral electrophoresis whereby DNA fragments migrated in the

direction perpendicular to that of SSB comets, producing comets of DNA fragments arising from

DSB (Figures 2.2c and d). ACM-2 and ACM-3 were regarded as spermatozoa with fragmented

DNA and were often observed to occur concurrently in a single spermatozoon (ACM-2+3). The

incidence of fragmented sperm nuclei revealed by the SCD test (ASM-3) and double-comet assay

(ACM-2, ACM-3 and ACM-2+3) is reported in Tables 2.1 and 2.2, respectively. A correlation

analysis to compare the results of estimates of the total SDF from the SCD (ASM-3) and the

double-comet assay (ACM-2 + ACM-3 + ACM-2+3), revealed a strong positive correlation (r =

0.9613; Figure 2.3), although the double-comet assay typically gave slightly higher values of total

SDF than the SCD test.

44

Figure 2.2. Evidence of DNA fragmentation in African clawed frog spermatozoa following the double (neutral + alkaline) comet assay at T0 immediately following activation; (a, b) sperm nuclei with large nuclear cores and compact comet tails representing structural SSB only (ACM-1); (c) sperm nucleus with extensive DNA damage indicated by the presence of comets comprised of both SSB and DSB (ACM-2+3); (d) sperm nucleus showing a comet comprised of structural SSB and DSB and (e) sperm nucleus showing a high degree of SSB as evidenced by a long comet tail and much reduced nuclear core (ACM-2).

45

Table 2.1. Percentage of SDF as assessed by the SCD in fresh activated African clawed frog spermatozoa in three frogs incubated over 24 h at 25 °C.

Male Time (T) ASM-1 ASM-2 ASM-3 Total SDF

(ASM-3)

1 T0 85.7 12.0 2.3 2.3

T1 68.7 16.0 15.3 15.3

T24 12.0 41.0 47.0 47.0

2 T0 92.7 2.0 5.3 5.3

T1 57.7 22.0 20.3 20.3

T24 4.0 35.0 61.0 61.0

3 T0 97.7 1.0 1.3 1.3

T1 68.3 23.0 8.7 8.7

T24 8.7 57.0 34.3 34.3

Table 2.2. Percentage of SDF as assessed by the double-comet assay in fresh activated African clawed frog spermatozoa in three frogs incubated over 24 h at 25 °C.

Male Time (T) ACM-1 ACM-2 ACM-3 ACM-2+3 Total SDF

1 T0 91.2 0.9 0.6 7.3 8.8

T1 82.1 2.3 7.3 8.3 17.9

T24 40.0 4.0 2.0 54.0 60.0

2 T0 91.0 1.0 2.0 6.0 9.0

T1 64.7 15.0 13.0 7.3 35.3

T24 20.0 7.0 5.0 68.0 80.0

3 T0 92.4 0.6 5.0 2.0 7.6

T1 83.1 2.3 7.3 7.3 16.9

T24 40.0 4.0 2.0 54.0 60.0

46

Figure 2.3. Correlation between the percentage of SDF identified in three male African clawed frogs immediately following activation as estimated by SCD test (ASM-3) and double-comet assay (ACM-2 + ACM-3 + ACM-2+3).

2.5 Discussion

The present study has demonstrated a strong positive correlation between the levels of SDF

revealed by the SCD test (ASM-3) and the double-comet assay (ACM-2 + ACM-3 + ACM-2+3).

While differentiation between single- and double-stranded DNA breaks was possible using the

double-comet assay, rarely did SDF, as revealed by SSB and DSB, occur separately but occurred

concurrently. ISNT was used to further confirm that ASM-3 morphology was associated with SDF

damage. We are therefore confident that the SCD procedure validated in this study can now

subsequently be used to investigate the behaviour of amphibian sperm DNA in response to a range

of endogenous and exogenous stressors. We have already seen in the present study that SDF

increases with incubation at room temperature, but the effect of cryopreservation and subsequent

incubation is likely to be even more pronounced.

The SCD procedure resulted in three distinctive morphotypes. ASM-3 was the result of

massive chromatin dispersal away from the nucleus and is clearly associated with severely damaged

DNA; this was also evidenced by the fact that the nucleoid core had coincidently reduced with

increased levels of DNA dispersal. Equally, the morphology of ASM-1 was also relatively

straightforward to interpret, with the sperm nucleus retaining its general morphology, but only

47

showing evidence of a small halo of DNA migration; this phenomenon is indicative of an ordered

chromatin relaxing following denaturation and protein removal. However, the morphology of ASM-

2 is somewhat more difficult to explain, as these sperm nuclei showed no evidence of massive DNA

migration or reduction of the nuclear core, but a characteristic change in shape of the nucleus from a

spiral elongated form to rounded head morphology; interestingly a shift in frequency from ASM-1

to ASM-2 was also observed with in vitro room-temperature storage, indicating that this change is

likely to be a consequence of the activated spermatozoa being subject to induced iatrogenic damage.

The mechanism leading to the loss of the original nuclear morphology may be similar to the

typical chromatin swelling or relaxation that has been observed in other species devoid of

disulphide bonds in their protamines. In these species it is possible that the nuclear basic proteins of

the spermatozoon render the DNA more susceptible to protein depletion (Zee et al. 2009). The

chromatin-swelling phenomenon observed here is similar to that described in the koala (Johnston et

al. 2006; 2007; 2012a), common wombat (Vombatus ursinus; Johnston et al. 2006) and fish

(López-Fernández et al. 2009). Chromatin swelling in African clawed frog spermatozoa without

DNA fragmentation may be due to the extensive presence of alkali labile sites (ALS), highly

sensitive DNA motifs (possible regions of specific repetitive sequences; Cortés-Gutiérrez et al.

2009) or baseline DNA lesions that are produced when the spermatozoa are processed in vitro;

these DNA motifs become ‘hot-spots’ producing stretches of single-stranded DNA following

alkaline denaturation (Cortés-Gutiérrez et al. 2009; 2014a). This phenomenon would also explain

the pervasiveness of structural SSB revealed in sperm nuclei by the alkaline step of the double-

comet assay and suggests that ALS are a constitutive feature of the African clawed frog sperm

genome. Perhaps this is why all spermatozoa, including those exhibiting small compact chromatin

halos of undamaged DNA, expressed positive in situ nick translation labelling, given that the DNA

nucleotide is incorporated into the free ends of single- or double-stranded DNA breaks. These sites

are proposed as an alternative DNA configuration that functions in addition to protamine-mediated

disulphide bonds to produce the highly condensed state of the chromatin during spermiogenesis.

Cortés-Gutiérrez et al. (2009) suggest that ALS are likely to be universally present in the mature

spermatozoa of all mammalian spermatozoa and, accordingly, structural SSB have been observed in

several mammalian species (Cortés-Gutiérrez et al. 2008b; 2009; Enciso et al. 2009). However,

they persist more extensively in disulphide-bond-deficient species such as the koala (Zee et al.

2009), short-beaked echidna (Johnston et al. 2009) and fish (López-Fernández et al. 2009). Thus,

the prevalence of ALS in the mature spermatozoa of African clawed frog is likely to be related to

48

the absence of sperm-specific protamines in the chromatin, the cysteine residues of which are

essential for the formation of disulphide bonds (Yokota et al. 1991; Ausio et al. 2007).

Interestingly, in the present study, the absence of protamines–cysteine residues and

associated disulphide bonds in African clawed frog spermatozoa did not appear to make the affected

spermatozoa more susceptible to DNA fragmentation in vitro. An underlying assumption of the

study was that DNA fragmentation would proceed in a rapid manner consistent with that of other

protamine-deficient species. However, in the three males examined, the SDF did not exceed 61% in

activated sperm samples over the duration of the 24 hour incubation period. This phenomenon is in

direct contrast to that observed in fish, where an increase in SDF occurs only a few minutes after

sperm activation (López-Fernández et al. 2009). The rapid rate of DNA fragmentation observed in

fish may be due to the combination of osmotic stress imposed on the spermatozoa following

activation in addition to a chromatin structure lacking disulphide bonds (López-Fernández et al.

2009). It is, therefore, interesting to note that the African clawed frog, which utilises the same

strategy of external fertilisation and osmotically induced activation and possesses a sperm

chromatin structure devoid of disulphide bonds, does not exhibit a similarly rapid decay of DNA

quality. This may indicate that another mechanism of chromatin stabilisation functions in African

clawed frog spermatozoa and highlights the importance of considering inter-species differences in

sperm chromatin structure when applying the SCD test to a novel species.

This study supports the hypothesis that the SCD test can be used to successfully identify

DNA damage in African clawed frog spermatozoa and produces results concordant with other

standard DNA fragmentation assays. The unique situation with African clawed frog sperm nuclear

basic proteins reinforces the importance of developing species-specific DNA fragmentation assays.

This is particularly the case for the SCD test where visualisation of a differential chromatin

dispersion pattern and, thus, an accurate assessment of DNA damage, is dependent on sufficient

protein removal and exposure of the DNA molecule. As the first amphibian protocol for the

assessment of SDF, this protocol has the potential to benefit assisted breeding programs using ART

to support amphibian conservation efforts. The ability to identify males whose sperm DNA is

compromised or particularly sensitive to fragmentation following activation will optimise mate

pairing, minimise wastage of female gametes and may aid the reproductive output of ART. Given

that gamete cryopreservation is a fundamental aspect of ART, it would be prudent to assess the

impact of freeze–thawing on the structural integrity of amphibian sperm DNA. Despite successful

protocols established for several amphibian species (Beesley et al. 1998; Browne et al. 1998;

49

2002ab; Michael and Jones 2004; Sargent and Mohun 2005), the process of cryopreservation

inherently promotes cellular damage that can compromise the stability of the sperm chromatin.

Furthermore, the information presented in this study has the potential to provide an insight

into the mechanisms of amphibian decline in the wild. Given their unique anatomy, amphibians are

particularly sensitive to fluctuations in environmental conditions; their exposed and permeable skin

coupled with a dependence on water for breeding means that they often react adversely to changes

in the environment before higher-order vertebrates (Burlibasa and Gavrila 2011). Additionally, the

potential for a high rate of mutagenic change or impaired spermatogenesis in the testes means that

amphibian sperm chromatin structural integrity (including sperm chromatin quality) could be a

useful index of environmental toxicity on the reproductive system; this study has provided the

technical basis and protocols by which the integrity of amphibian sperm chromatin might be

evaluated as such an index.

50

CHAPTER 3: Dynamics of sperm DNA fragmentation in Xenopus

laevis using the sperm chromatin dispersion (SCD) test; the effect of

activation and cryopreservation.

Pollock, K.1, Gosálvez, J.2, López-Fernández, C.2, Arroyo, F.2, Wilson, R.3, Guille, M.4, Noble, A.4

and Johnston, S.D.1

1School of Animal Studies, The University of Queensland, Gatton, Australia 4343. 2Department of Biology, Genetics Unit, The Autonomous University of Madrid, Madrid, Spain

20849. 3Department of Mathematics, The University of Queensland, St Lucia, Australia 4072. 4European Xenopus Resource Centre, The University of Portsmouth, Portsmouth, United Kingdom

PO1 2DY.

Presented as prepared for submission to Reproduction, Fertility and Development

51

3.1 Abstract

Sperm cryopreservation is a fundamental component of amphibian assisted reproductive

technology (ART) but this process promotes cellular damage that can potentially compromise

sperm DNA integrity. Sperm DNA fragmentation (SDF) is a dynamic process and has been shown

to increase with incubation at room temperature as well as following cryopreservation. The present

investigation was aimed at examining the dynamic changes to the basal level of DNA fragmentation

in fresh African clawed frog (Xenopus laevis) spermatozoa in comparison with the post-thaw

integrity of cryopreserved (-80 °C) and frozen (-20 °C) sperm DNA. Inactivated spermatozoa were

collected from fresh testicular macerates (n = 7); a portion of the testes was macerated in chilled (4

°C) 0.1 M MBS to achieve sperm activation. Analysis of SDF using the sperm chromatin dispersion

(SCD) test was conducted for each sperm sample at time 0 (T0) immediately following activation

and/or thawing and again following a further 10 (T10), 30 (T30) and 60 minutes (T60) of

incubation at room temperature. An IVF procedure was conducted to compare the fertilisation rates

of fresh and cryopreserved spermatozoa; the sperm suspensions were activated with 250 µL of

chilled (4 °C) 0.1 M MBS and added to a petri dish containing approximately 200 stripped oocytes.

The difference in SDF was only marginal for fresh, frozen (-20 °C) and cryopreserved (-80 °C)

spermatozoa examined at the onset of the experiment immediately upon thawing, however, a

significant increase (p = 0.004) in SDF, albeit of a low magnitude, was observed in activated

cryopreserved (-80 °C) following incubation. These results suggest that the sperm nucleus of

African clawed frog appear to be highly resistant to cryopreservation-induced DNA damage.

Cryopreserved-thawed spermatozoa yielded a significantly lower (p = 0.0065) fertilisation rate than

fresh spermatozoa. The information presented in this study has relevance to assisted breeding

programs using cryopreserved sperm to support endangered amphibian conservation.

3.2 Introduction

The focus of amphibian conservation has recently shifted towards establishing a genetic

resource bank coupled with assisted reproductive technologies (ART) as a means to secure

biodiversity at the risk of extinction, as well as help to sustain the genetic viability of extant

populations (Kouba and Vance 2009). As such, gamete cryopreservation is a fundamental aspect of

amphibian conservation. The effect of cryopreservation on breeding capacity, specifically sperm

DNA stability, is of critical importance for improving ART for the conservation of threatened and

endangered amphibians. A requirement for the use of cryopreserved-thawed sperm for ART is that

52

the cryopreservation process must not induce alterations that would impede progeny viability and

development (Labbe et al. 2001). Despite many refinements in methodology and successful

protocols established for a number of amphibian species (Beesley et al. 1998; Browne et al. 1998;

2002a; Michael and Jones 2004; Sargent and Mohun 2005), the procedure of sperm

cryopreservation is not without adverse effects. Cryopreservation promotes cellular damage that can

compromise sperm quality in terms of motility, vitality, membrane stability and DNA integrity

(Martínez-Páramo et al. 2012). The prevalence of chromatin damage in mammalian spermatozoa is

known to result in impaired fertilisation, early embryonic loss and ultimately, diminished

reproductive success (Martínez-Páramo et al. 2012).

As DNA integrity is poorly correlated with classical parameters of semen quality, an

important component of sperm viability would remain cryptic without the means to identify subtle

defects or potential damage (López-Fernández et al. 2008b). Assessment of DNA integrity,

therefore, provides the best indication of seriously damaged spermatozoa. The sperm chromatin

dispersion (SCD) test is a DNA fragmentation assay based on the differential response of

fragmented and intact spermatozoa nuclei to controlled denaturation and deproteinisation treatments

and has been previously adapted to the chromatin structure of African clawed frog spermatozoa and

validated using in situ nick translation (ISNT) and the double comet assay (Pollock et al. 2015). The

SCD test (Halosperm®; Fernández et al. 2003) is currently used extensively in human assisted

reproduction laboratories and has been successfully adapted (as the Halomax®; HalotechSL,

Madrid, Spain) for use in many mammalian and lower vertebrate species such as the koala

(Phascolarctos cinereus; Johnston et al. 2007), short-beaked echidna (Tachyglossus aculeatus;

Johnston et al. 2009), boar (Pérez-Llano et al. 2006), ram (López-Fernández et al. 2008a), stallion

(López-Fernández et al. 2007), donkey (Equus asinus; Cortés-Gutiérrez et al. 2008a), rhinoceros

(Diceros bicornis, Rhinoceros unicornis and Ceratotherium simum; Portas et al. 2009) and fish

(López-Fernández et al. 2009; Gosálvez et al. 2014); the SCD test employs a species-specific lysing

solution for protein depletion and was validated in each of these species separately.

Numerous studies have attributed damage to the chromatin structure of spermatozoa due to

the process of freezing and subsequent thawing. Semen processing such as dilution, cooling,

addition of cryoprotectants and thaw temperature, can impact the degree of DNA fragmentation

observed. An increase in sperm DNA fragmentation (SDF) subsequent to cryopreservation has been

reported in a range of wildlife and commercial species, including rainbow trout (Oncorhynchus

mykiss; Labbe et al. 2001), stallion (Baumber et al. 2003), chicken (Mericanel della Brianza;

53

Gliozzi et al. 2011), elephant (Elephas maximus; Imrat et al. 2012), koala (Johnston et al. 2012b)

and most recently in Western clawed frog (Xenopus tropicalis; Morrow et al. 2014). The

application of a DNA fragmentation assay to cryopreserved-thawed amphibian spermatozoa will

provide an insight into cryoinjury of the sperm chromatin and the resulting effect on the potential

reproductive efficiency of ART. Consequently, the present investigation was aimed at examining

the dynamic changes to the basal level of DNA fragmentation in fresh amphibian spermatozoa in

comparison with the post-thaw integrity of cryopreserved (-80 °C) and conventionally frozen (-20

°C) sperm DNA. An in vitro fertilisation (IVF) procedure was also used to assess the extent to

which cryopreservation was correlated with a decline in fertilisation rate.



Seven male wild-type African clawed frogs (Xenopus laevis) were sourced from a breeding

facility maintained at the European Xenopus Resource Centre (EXRC; Portsmouth, United

Kingdom) under controlled conditions for housing, photoperiod, temperature, water quality and

feeding. All animals were healthy with a history of reproductive success. A summary of the

experimental design is depicted in Figure 3.1. In accordance with the EXRC protocol (EXRC

2012), sperm samples were recovered from the whole testes of euthanased animals (Tricaine/MS-

222; Sigma Aldrich, Dorset, England, UK) and dissected within 1.0 M MBS (Modified Barth’s

Saline) at room temperature and cut into thirds. A portion of the testis was macerated by gentle

application of an Eppendorf pestle in 200 µL of chilled (4 °C) 0.1 M MBS to achieve sperm

activation and attain a final concentration of approximately 10 x 106 spermatozoa mL-1.

Identification of sperm motility under x 400 magnification confirmed activation. Another segment

of the testis was macerated in a fresh aliquot (200 µL) of 1.0 M MBS to maintain inactivation. The

third portion of testis was macerated in 250 µL of Leibovitz L-15 (0.15 M NaCl) tissue culture

medium (Sigma Aldrich, Dorset, England, UK) in preparation for cryopreservation. Analysis of

SDF for each sperm sample was conducted at time 0 (T0) immediately following maceration and

again following 10 (T10), 30 (T30) and 60 minutes (T60) of incubation at room temperature. The

incubation period was restricted due to the rapid decline in the fertilisation capacity and motility of

African clawed frog spermatozoa following activation (Wolf and Hedrick 1971; Sargent and Mohun

2005). Sargent and Mohun (2005) reported a half-life of sperm motility ranging from less than two

54

minutes to 21 minutes in the African clawed frog and similar results have been observed in

cryopreserved-thawed spermatozoa (Mansour et al. 2009).

Figure 3.1. A representation of the experimental design detailing the division and treatment of a single testis for an individual African clawed frog; T(x) – time of incubation in minutes; SCD – sperm chromatin dispersion test; ACT – activated spermatozoa; INACT – inactivated spermatozoa; IVF – in vitro fertilisation.

55

3.3.2 Sperm freezing (-20 °C) and cryopreservation (-80 °C)

The need for a constant supply of liquid nitrogen is a significant constraint on semen

collection and cryopreservation in the field for the assessment of SDF (Faszer et al. 2006).

Alternatives to conventional liquid nitrogen controlled rate freezers that do not require large

volumes of cryogen have demonstrated good recovery rates, however, typically require expensive

specialist equipment (Faszer et al. 2006), while commercial portable freezers have a considerably

smaller temperature range. Consequently, in order to determine the effect of freezing at a higher

temperature on SDF and facilitate ease of sample processing in the field, 20 µL aliquots of both

inactivated and activated spermatozoa were frozen without any additional cryoprotectant in

Eppendorf Safe Lock Tubes® (Eppendorf, Hamburg, Germany) at -20 °C in a conventional freezer

immediately following maceration (T0) and again following 10 (T10), 30 (T30) and 60 minutes

(T60) of incubation at room temperature. After at least 24 hours, frozen aliquots were thawed by

hand (approximately 30 seconds) and processed with the SCD test immediately following thawing

(T0).

Aliquots of inactivated spermatozoa were also cryopreserved at T0 according to the EXRC

protocol (Noble and Jafkins 2010) which was modified from that developed by Sargent and Mohun

(2005) for the African clawed frog. Briefly, a portion of the testis was macerated by gentle

application of an Eppendorf pestle in an Eppendorf Safe Lock Tube® (Eppendorf, Hamburg,

Germany) containing 250 µL of Leibovitz L-15 (0.15 M NaCl) tissue culture medium (Sigma

Aldrich, Dorset, England, UK) supplemented with 10% (v/v) foetal bovine serum (FBS) and 2.0

mM L-glutamine. The sperm preparation was cooled to 4 °C and diluted 1:1 with chilled (4 °C)

cryoprotectant (0.4 M sucrose, 10 mM NaHCO3, 2 mM pentoxyfylline, egg yolk and dH2O; Noble

and Jafkins 2010). Egg yolk was included in the cryoprotectant as it has been shown to significantly

improve viability of African clawed frog spermatozoa after freeze-thawing (Sargent and Mohun

2005). Optimal results have been obtained using a slow-freeze regime (unpublished data; Noble and

Jafkins 2010), whereby aliquots (250 µL) in thin-walled Eppendorf tubes were placed in a

polystyrene box (measuring approximately 20 x 15 cm with 3 cm thick walls), closed with a layer

of aluminium foil and stored in a -80 °C freezer as per EXRC protocol (Noble and Jafkins 2010).

Cryopreserved aliquots were thawed by hand (approximately 30 seconds) after a minimum of 24

hours of frozen storage. Aliquots were divided in half and one of each half was diluted 1:1 with 125

µL of chilled (4 °C) 0.1 M MBS to achieve sperm activation. Analysis of SDF was conducted

56

immediately following thawing (T0) and again after 10 (T10), 30 (T30) and 60 minutes (T60) of

incubation at room temperature.

3.3.3 In vitro fertilisation (IVF)

To confirm the fertilising capacity of each male and examine the effect of cryopreservation

on reproductive output, aliquots of fresh and cryopreserved-thawed (-80 °C) spermatozoa were used

for an IVF procedure with fresh oocytes retrieved from gravid female African clawed frogs. Egg

laying was induced in three females of proved fertilisation success using subcutaneous injections of

human chorionic gonadotropin (Chorulon 450 – 600 IU hCG), according to EXRC protocol (EXRC

2012). The sperm suspensions were activated with 250 µL of chilled (4 °C) 0.1 M MBS and added

to a petri dish (100 mm x 15 mm) containing approximately 200 stripped oocytes and gently

agitated to ensure thorough mixing and a single layer of oocytes. After 10 minutes, the oocytes were

flooded with chilled (4 °C) 0.1 M MBS and stored in a refrigerator at 4 °C for at least 24 hours. The

degree of fertilisation success was determined by calculating the proportion of oocytes that showed

evidence of nuclear cleavage after 24 – 48 hours; a target fertilisation rate of > 95% is typical

(EXRC 2012).


The Xenopus-Halomax® kit adapted from the Sperm-Halomax® kit (HalotechSL, Madrid,

Spain) was used to assess the extent of DNA fragmentation in each sample and has been described

previously (Pollock et al. 2015). Briefly, 50 µL aliquots of low point melting agarose in Eppendorf

Safe Lock Tubes® (Eppendorf, Hamburg, Germany) were suspended in a water bath at 90 – 100 °C

for five minutes and then equilibrated to 35 °C in a second water bath. The extended sperm

suspension (10 µL) was added to the agarose and gently mixed, before 10 µL aliquots were pipetted

onto pre-treated glass slides (provided in the Sperm-Halomax® kit) and covered with a glass

coverslip (18 x 18 mm). The slides were then left to solidify on a chilled metallic plate in a

refrigerator (4 °C) for five minutes. After the coverslips were removed, the slides were promptly

introduced (maintained at a horizontal position) into a lysing solution (10 mL) containing 2 M

NaCl, 0.5% SDS, 0.01 Triton X, 0.2 M Tris-HCL and 0.02 M EDTA; pH 7 (provided in the Sperm-

Halomax® kit and adjusted for amphibian spermatozoa) for five minutes. The slides were removed

from the lysing solution and washed in dH2O for five minutes and then dehydrated in sequential

ethanol baths (70, 90 and 100%) for two minutes each and finally left to air-dry. Spermatozoa were

57

mounted with equal parts SYBR Green 1 (Sigma Aldrich, Madrid, Spain) and Vectashield

mounting medium (Vector Laboratories, Burlingame, California, USA) and observed using a Leica

DMRB epifluorescence microscope (Leica Microsystems, Barcelona, Spain) equipped with single-

band fluorescence block filters for green (fluorescein isothiocyanate (FITC) equivalent) and red

(Cyanine3 NHS ester (Cy3) equivalent) fluorescence. A total of 300 spermatozoa were counted per

sample and the percentage of spermatozoa with fragmented DNA calculated. All images were

captured with a CCD camera (Leica DFC350 FX, Leica Microsystems, Barcelona, Spain.


Statistical analyses were performed using the Statistical Package for the Social Sciences

(SPSS, Inc., Chicago, USA). The relationship between SDF and the different storage methodologies

was determined using a linear logistic regression model. An F-test was used for the comparison of

fertilisation success. Differences were considered significant with P-values less than 0.05.

3.4 Results

3.4.1 Dynamics of sperm DNA fragmentation

In general, the basal level of SDF observed at the onset of the experiment (T0) was low,

with values not exceeding 2% for inactivated and activated spermatozoa of fresh samples, as well as

frozen (-20 °C) and cryopreserved samples immediately upon thawing (Figure 3.2). While the level

of SDF fluctuated slightly throughout the incubation period for fresh and frozen (-20 °C) samples,

values obtained after 60 minutes of incubation did not exceed 3% in almost all cases, with the

notable exception of Male 1 where the SDF reached 5% and 11% for fresh inactivated and activated

spermatozoa respectively. Almost all cryopreserved spermatozoa showed a marked increase in

DNA fragmentation after 30 minutes of incubation, reaching a mean (± SEM) SDF of 4.4 ± 1.4%

and 5.7 ± 1.0% at the completion of the incubation period for inactivated and activated

spermatozoa, respectively. The spermatozoa of male 3 appeared particularly robust to the stresses of

cryopreservation and showed almost no change in SDF for inactivated spermatozoa and only a

marginal increase of less than 3% for activated spermatozoa.

58

Figure 3.2. Logistic model plots representing the dynamics of SDF in the African clawed frog after 60 minutes of incubation at 25 °C. SDF – sperm DNA fragmentation.

3.4.2 Regression analysis of SDF after incubation for fresh, frozen (-20 °C) and cryopreserved (-80

°C) samples

A regression analysis was conducted to examine the relationship between SDF and the

different storage methodologies (Table 3.1). There was increased variability in the initial levels of

SDF observed between the different treatments. However, this variability was not consistent across

all treatments; the initial levels of SDF for inactivated (p = 0.2505) and activated (p = 0.7948)

cryopreserved spermatozoa were not significantly different. Consequently, this variability is more

likely to reflect experimental method variation rather than real differences between animals. While

59

variation in the rate of DNA fragmentation accounted for a significant proportion of the overall

variation in SDF, the difference between the upper and lower confidence limits was only marginal,

indicating that the actual change in SDF was small relative to 0%. The rates of increase in SDF for

fresh and frozen (-20 °C) samples were negligible and did not change significantly from 0% (p >

0.05) for the duration of the incubation period, irrespective of whether the spermatozoa were

inactivated or in the activated state. An exception was male 1, which showed a significant increase

in SDF for both inactivated (p < 0.0001) and activated (p = 0.0240) fresh spermatozoa. A

significant increase in SDF was observed in cryopreserved samples (p < 0.006) with the exception

of male 3 whose SDF remained low (p = 0.3975).

Table 3.1. Regression analysis of SDF in African clawed frog incubated at room temperature for 60 minutes examined fresh as well as following freezing (-20 °C) and cryopreservation (-80 °C).

Source of variation

Initial SDF at T0

(β0: intercept)

Rate of increase in SDF

(β1: slope)

Fresh INACT < 0.0001* > 0.5 (except male 1; < 0.0001*)

ACT 0.0010* > 0.5 (except male 1; 0.0240*)

Frozen (-20 °C) INACT 0.0005* > 0.5

ACT 0.0111* > 0.5

Cryo (-80 °C) INACT 0.2505 < 0.006* (except male 3; 0.3975)

ACT 0.7948 < 0.006*

SDF – sperm DNA fragmentation; T0 – time of incubation in minutes at onset of experiment; INACT – inactivated spermatozoa; ACT – activated spermatozoa motility; *denotes significant p-value.

3.4.3 Effect of SDF on fertilisation success after cryopreservation at -80 °C

Aliquots of fresh and cryopreserved-thawed (-80 °C) sperm samples were used for IVF with

fresh oocytes retrieved from gravid female African clawed frogs. The degree of fertilisation success

was determined by calculating the proportion of oocytes that showed evidence of cleavage after 24

– 48 hours. SDF at the time of IVF was similar for fresh and cryopreserved-thawed samples.

Nevertheless, cryopreserved-thawed sperm samples yielded a significantly lower rate of fertilisation

success (p = 0.0065; Table 3.2).

60

Table 3.2. Fertilisation success resulting from an IVF procedure using fresh and cryopreserved-thawed (-80 °C) African clawed frog spermatozoa.

Fresh Cryopreserved (-80 °C) p-value

Mean (± SEM) no. oocytes fertilised (%) 80.4 ± 4.7 32.3 ± 14.9 0.0065*

Mean (± SEM) SDF (T0) 0.4 ± 0.2 0.3 ± 0.1 0.1342

SEM – standard error of mean; SDF – sperm DNA fragmentation; T – time of incubation in minutes at onset of experiment; *denotes significant p-value.

3.5 Discussion

The observations of the present study support the dynamic nature of DNA fragmentation

(Gosálvez et al. 2011b), however, the loss of DNA quality manifests slowly in the spermatozoa of

the African clawed frog. Although there are limited dynamic studies of SDF against which to base

our comparisons, the basal level of SDF in fresh African clawed frog spermatozoa (0 – 2%) was

similar to that of previously examined species, albeit with values at the lower end of the observed

ranges (Johnston et al. 2007; López-Fernández et al. 2007; Cortés-Gutiérrez et al. 2008a; López-

Fernández et al. 2009; Portas et al. 2009). Although there was some evidence of individual

variation (see male 1 – Figure 3.2), the sperm DNA of African clawed frog was remarkably stable

over the 60 minute period of incubation at room temperature; this chromatin stability was apparent

for both inactivated and activated spermatozoa. This finding is surprising, given that activation of

motility in spermatozoa from anuran amphibians, which occurs by deposition into hypo-tonic

fresh/pond water (low osmolality of < 50 mOsmol kg-1), poses a significant osmotic challenge to

the spermatozoa (Browne and Zippel 2007; Kouba et al. 2009). In fact, Wolf and Hendrick (1971)

noted that the maximum rate of fertilisation in the African clawed frog was restricted to only 10 –

15 minutes following activation, while spermatozoa stored at 0 °C in isotonic DeBoers solution

retained the capacity to fertilise for up to 24 hours. Consequently, an underlying assumption of the

study was that DNA fragmentation would proceed in a rapid manner consistent with that of other

externally fertilising species that require osmotically induced activation of the spermatozoa.

However, in almost all cases, the SDF did not exceed 3% for inactivated or activated sperm samples

after 60 minutes of incubation. The significant increase in SDF seen for Male 1 is a notable

exception and may reflect an inherent sensitivity of the sperm chromatin. This finding is interesting

and provides insight into the degree of inter-individual variation that may exist at the population

level.

61

The remarkable stability of the sperm chromatin in the African clawed frog is in direct

contrast to the situation observed in fish (López-Fernández et al. 2009; Gosálvez et al. 2014) where

an increase in SDF occurs only a few minutes after sperm activation. The rapid rate of DNA

fragmentation following activation observed in fish spermatozoa may be facilitated by a chromatin

structure lacking disulphide bonds (López-Fernández et al. 2009). It is generally accepted that the

presence or absence of protamines/cysteine residues and associated disulphide bonds to stabilise the

sperm chromatin is likely to affect the dynamics by which DNA fragments (Cortés-Gutiérrez et al.

2014b). Significantly higher levels of fragmented DNA in response to oxidative stress compared to

eutherian mammals has been also observed in the koala, common wombat (Vombatus ursinus) and

Eastern grey kangaroo (Macropus giganteus), all of which lack disulphide bonding in their sperm

chromatin (Enciso et al. 2011). The observation that African clawed frog spermatozoa do not

exhibit a similarly rapid decay of DNA quality, despite also possessing a chromatin structure devoid

of disulphide bonds (Yokota et al. 1991; Ausio et al. 2007) is interesting and may suggest that

another mechanism of chromatin stabilisation functions in this species. This may be related to the

high arginine content of the protamine-like proteins present in African clawed frog spermatozoa

(Yokota et al. 1991; Ausio et al. 2007), which are proposed to increase the charge density and

greatly enhance the compaction of sperm DNA (Lewis and Ausio 2002).

The results of the present study have shown that freezing African clawed frog spermatozoa

at -20 °C without any additional cryoprotectant does not have an appreciable effect in terms of

DNA fragmentation. Similarly, vitrification (ultra-rapid freezing) of spermatozoa without the use of

cryoprotectants has achieved post-thaw recovery of DNA integrity comparable to fresh spermatozoa

in the rabbit (Rosato and Iaffaldano 2013) and human (Isachenko et al. 2004ab). This observation

has practical implications for the assessment of SDF in the field. As the loss of DNA quality in

frozen sperm samples proceeded in a manner consistent with fresh spermatozoa, it is possible to use

spermatozoa frozen in this manner to obtain an index of SDF comparable to fresh sperm samples.

This will facilitate more convenient sperm collection in the field with the use of a commercial

freezer, without the need for large quantities of liquid nitrogen. It is possible that the stability of the

sperm chromatin after 60 minutes of incubation reflects that the spermatozoa were frozen without

cryoprotectant and therefore, avoided dramatic changes in the osmotic environment and associated

oxidative stress arising from the production of ROS such as free radicals (Gosálvez et al. 2011a).

A number of cryopreservation protocols have been developed for African clawed frog

spermatozoa and have yielded mixed success. Sperm cryopreservation protocols that involve

62

penetrating cryoprotectants developed for other amphibian species (Beesley et al. 1998; Browne et

al. 1998; Costanzo et al. 1998) have not been successful in cryopreserving the spermatozoa of the

African clawed frog (Bucholz 2004; Sargent and Mohun 2005). While Mansour et al. (2009)

recorded positive results using DMSO, the EXRC protocol (Noble and Jafkins 2010) involves the

non-penetrating cryoprotectant sucrose shown to protect African clawed frog spermatozoa from

cryoinjury (Sargent and Mohun 2005). Preliminary studies conducted by Mansour et al. (2009),

demonstrated that processing Xenopus spp. spermatozoa at 4 °C prolonged its viability; similar

results have been recorded for the cane toad (Bufo marinus; Browne et al. 2002b). Consequently,

sperm preparations and cryoprotectant medium were chilled to 4 °C prior to freezing.

African clawed frog spermatozoa appear to be highly resistant to cryopreservation-induced

DNA damage (particularly Male 3 – see Figure 3.3). Although the cryopreserved (-80 oC) sperm

samples showed a significant increase in SDF after 60 minutes of incubation, the proportion of

sperm with SDF was still low, being less than 10%. Similarly, spermatozoa of numerous mammals

including dog (Prinosilova et al. 2012) and stallion (López-Fernández et al. 2007) retain a normal

chromatin structure immediately after thawing (Gosálvez et al. 2011b). Although Morrow et al.

(2014) observed a slightly higher SDF of 17.65% in Western clawed frog spermatozoa following

cryopreservation, it is unclear whether these results were obtained immediately after thawing or

following a period of incubation, making inferences about the possible causes of this variability

difficult. Interestingly, koala (Johnston et al. 2012b) and rainbow trout (Labbe et al. 2001)

spermatozoa also exhibited a marked resistance to cryopreservation-induced DNA damage, despite,

like the African clawed frog, lacking the capacity for disulphide bonding to stabilise and protect the

chromatin from in vitro denaturation (Yokota et al. 1991; Ausio et al. 2007). While the integrity of

the sperm chromatin remained fairly stable during the first minutes post-thawing, other studies of

amphibian sperm cryopreservation have reported a marked decrease occurring at this time in other

parameters of sperm quality such as motility and vitality (Sargent and Mohun 2005; Mansour et al.

2009). The transition of intracellular water to ice during the freezing process causes detrimental

biophysical changes to the spermatozoa, particularly to the plasma membrane and mitochondria

(Martínez-Páramo et al. 2012). Thus, the increase in SDF seen for cryopreserved spermatozoa may

be due in part to oxidative stress arising from the production of reactive oxygen species such as free

radicals, by the membrane-compromised and dying sperm population caused by lethal intracellular

ice formation (Gosálvez et al. 2011a).

63

The general assumption is that fresh spermatozoa stored at room temperature or chilled at 4

°C have a greater fertilisation capacity than cryopreserved-thawed spermatozoa. Accordingly,

cryopreserved-thawed African clawed frog spermatozoa yielded a significantly lower fertilisation

rate than fresh spermatozoa, despite exhibiting a similar level of SDF at the time of IVF. The

proportion of oocytes that showed evidence of cleavage after 24 – 48 hours of incubation with fresh

sperm samples (mean ± SEM; 80.4 ± 4.7%) was consistent with other results reported for the

African clawed frog; evidence of fertilisation using chilled fresh spermatozoa was observed in 80%

to 93% of oocytes by Wolf and Hedrick (1971) and Hollinger and Corton (1980) respectively, while

the European Xenopus Resource Centre (EXRC 2012) report that a target fertilisation rate of > 95%

is typical. Conversely, fertilisation protocols using cryopreserved amphibian spermatozoa have

achieved mixed success. Sargent and Mohun (2005) reported that the number of oocytes fertilised

using cryopreserved African clawed frog spermatozoa (23% to 81%) was never as great as in a

control fertilisation using fresh spermatozoa of the same sample. In the present study, the proportion

of oocytes fertilised using cryopreserved spermatozoa (mean ± SEM; 32.3 ± 14.9%) was similar to

results reported by Sargent and Mohun (2005), albeit at the lower end of the observed range. In the

can toad (Browne et al. 1998), values ranging from < 20% up to 81.4% have been recorded

depending on the concentration of the cryoprotectant used.

Although the impact of cryopreservation on amphibian sperm DNA remains to be explored

in greater detail, it is well documented that the process of freeze-thawing induces the impairment of

other parameters of sperm quality such as motility, viability and plasma membrane integrity. Labbe

et al. (2001) reported a very low increase in the proportion of rainbow trout spermatozoa with

altered DNA after cryopreservation compared to the 20 fold increase in the proportion of

spermatozoa reported as having damaged membranes and mitochondria (DeBaulny et al. 1997).

These findings support that DNA damage is only a minor component of cryoinjury and that

impaired fertilisation capacity is more likely related to poor recovery of motility (mitochondrial

damage), viability and plasma membrane integrity. In summary, the sperm DNA of the African

clawed frog is remarkably stable post-activation as well as post-thawing. It would be prudent for

future studies to incubate sperm samples for a longer period in order to reveal the true dynamics of

DNA fragmentation in this species. This would provide more information about the DNA quality of

individual males and is applicable to the use of assisted reproductive technologies for the captive

breeding of threatened and endangered species. Furthermore, freezing sperm samples at -20 °C may

prove a useful tool for assessing sperm DNA fragmentation in the field.

64

CHAPTER 4: Effect of cryopreservation on the dynamic behaviour of

sperm DNA fragmentation, motility and viability in cane toad (Bufo

marinus) spermatozoa

Pollock, K.1, Gosálvez, J.2, López-Fernández, C.2, Keeley, T.1, Feng, T.1, Wilson, R.3 and Johnston,

S.D.1,

1School of Animal Studies, The University of Queensland, Gatton, Australia 4343. 2Department of Biology, Genetics Unit, The Autonomous University of Madrid, Madrid, Spain

20849. 3Department of Mathematics, The University of Queensland, St Lucia, Australia 4072.

Presented as prepared for submission to Reproduction, Fertility and Development

65

4.1 Abstract

DNA fragmentation of amphibian spermatozoa is a dynamic process and has been shown to

increase with incubation at room temperature as well as following cryopreservation. However, the

mechanisms of DNA cryoinjury and the resulting effect on the potential reproductive output of

ART remain open to interpretation. The present investigation aimed to provide further insight into

cryoinjury of the sperm chromatin by correlating the dynamic loss of DNA integrity in fresh and

cryopreserved cane toad (Bufo marinus) spermatozoa with other important parameters of sperm

quality. Spermatozoa were cryopreserved in simplified amphibian ringer (SAR) without

cryoprotectant or in 15% (v/v) dimethyl sulfoxide (DMSO). Analysis of sperm DNA fragmentation

(SDF) using the sperm chromatin dispersion (SCD) test was conducted at time 0 (T0) immediately

following activation and/or thawing and again following T3 and T6 hours of incubation at room

temperature. Nucleic acid stains SYBR-14 and propidium iodide were used to assess sperm viability

at these time points in addition to an assessment of sperm motility. Sperm chromatin appeared to be

highly stable following thawing and only showed a marked increase in SDF when frozen without

cryoprotectant (SAR). Total sperm motility was substantially lower than that reported previously in

this species. A significant increase in the proportion of spermatozoa showing nuclear swelling was

observed only in spermatozoa cryopreserved in SAR (p < 0.0001). The findings presented in this

study support that DNA fragmentation is only a minor component of cryoinjury and that poor

recovery of motility and viability are likely to have a greater effect on the useful lifespan of cane

toad spermatozoa post-thaw.

4.2 Introduction

The integrity of sperm DNA is now widely accepted as an important parameter of semen

quality and is essential for successful syngamy and embryonic development (D’Occhio et al. 2007).

A series of assays have been designed to evaluate the integrity of sperm chromatin, including tests

for the assessment of DNA dependent proteins and for the detection of DNA breaks. Originally

developed and validated for investigating DNA damage in somatic cells, these technologies have

been adapted to enable the reagents involved to access the more highly compacted DNA of

spermatozoa (Sharma and Agarwal 2011). The SCD test is a relatively new addition to the

assortment of technologies used to assess sperm DNA integrity and is based on the principle that

controlled denaturation and lysis of chromatin protein results in a differential DNA dispersion

pattern indicative of the severity and quantity of DNA breakage (Fernández et al. 2003). In the last

66

decade, the SCD test (Halosperm®; Fernández et al. 2003) has been successfully adapted (as the

Halomax®; HalotechSL, Madrid, Spain) to the unique chromatin structure of mammalian (Pérez-

Llano et al. 2006; Johnston et al. 2007; López-Fernández et al. 2007; Cortés-Gutiérrez et al. 2008a;

López-Fernández et al. 2008a; Johnston et al. 2009; Portas et al. 2009) and more recently to lower

order vertebrate (López-Fernández et al. 2009; Gosálvez et al. 2014; Morrow et al. 2014; Pollock et

al. 2015) spermatozoa, using a species-specific lysing solution for protein depletion. The

application of DNA fragmentation assays has been driven mainly by the need for assisted

reproductive technologies (ART) for the treatment of human infertility, commercial breeding and

endangered species conservation.

The worldwide decline in amphibian numbers has fuelled demand for the continued

development and refinement of ART for amphibian conservation. An estimated one-third of nearly

6000 amphibian species worldwide are experiencing rapid population declines (IUCN 2008).

Together with ART, captive insurance colonies are invaluable for protecting species at risk of

extinction by supporting the genetic viability of small, fragmented existing populations through the

reintroduction of tadpoles back into the wild. A noteworthy example of the successful application of

ART to an endangered amphibian is the production and subsequent release of nearly 2000

Wyoming toad (Bufo baxteri; Browne et al. 2006) tadpoles produced using in-vitro fertilisation.

Significant advancements have been made in the area of amphibian ART, specifically with regards

to artificial fertilisation, hormonal induction of gametes and short- to long-term storage of

spermatozoa (Browne and Zippel 2007). Sperm cryopreservation in particular has the potential to

maintain genetic diversity into perpetuity and is a fundamental component of an ART strategy

(Browne and Zippel 2007).

Understanding the impact of cryopreservation on the structural integrity of amphibian sperm

DNA is of critical importance for improving ART for the conservation of threatened and

endangered species. Pollock et al. (2015) have already shown that DNA fragmentation in

amphibian spermatozoa increases with incubation at room temperature and the effect of

cryopreservation and subsequent incubation is even more pronounced (See Chapter 3). Dilution of

the seminal plasma with cryoprotectants in preparation for cryopreservation predisposes

spermatozoa to oxidative and osmotic stress and can potentially contribute to the incidence of

spermatozoa with fragmented DNA post-thaw. While the mechanisms of DNA cryoinjury remain

open to interpretation, an increase in damaged DNA subsequent to the process of freezing and

thawing has been reported in a range of species including rainbow trout (Oncorhynchus mykiss;

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Labbe et al. 2001), stallion (Baumber et al. 2003), chicken (Mericanel della Brianza; Gliozzi et al.

2011), Asian elephant (Elephas maximus; Imrat et al. 2012) and koala (Phascolarctos cinereus;

Johnston et al. 2012b). Only a single study has examined the incidence of cryopreservation-induced

chromatin damage in amphibian spermatozoa (in Western clawed frog; Xenopus tropicalis; Morrow

et al. 2014; poster presented at the World Congress of Reproductive Biology). This study not only

reported an increase in sperm DNA fragmentation following cryopreservation, but also a higher

incidence of developmental defects in embryos produced using cryopreserved compared to fresh

spermatozoa (Morrow et al. 2014). In an attempt to provide further insight into cryoinjury of the

sperm chromatin and the resulting effect on the potential reproductive output of ART, the present

investigation aimed to correlate the dynamic loss of DNA integrity in fresh and cryopreserved

amphibian spermatozoa with other important parameters of sperm quality. The cane toad served as

a reproductive model for endangered amphibians in this study, as there was already an established

protocol for the cryopreservation of cane toad spermatozoa (Browne et al. 1998). Additionally, this

study aimed to identify an index of amphibian SDF at the population level by examining individual

variation in a large cohort of males. For experiments presented in Chapter 3 using African clawed

frog spermatozoa, the incubation period was too short and posed a significant limitation with

respect to the analysis of SDF dynamics and sperm chromatin longevity in this species.

Consequently, the experiments presented in this Chapter aimed to improve the previous

experimental design by employing a longer incubation period.



Adult male cane toads (n = 25; AEC Permit # AE04263) were sourced from a local supplier

(Peter Douche, Mareeba, Queensland, Australia). Males were euthanased by pithing with a metal

rod and the testes dissected within 2 mL 1:1 simplified amphibian ringer (SAR; composition in

mM: NaCl 113, CaCl2 1, KCl 2.0, NaHCO3 3.6; 202 mOsmol kg-1; pH 7.2) at room temperature (22

°C) in a petri dish. The isotonic osmolality of the SAR prevented activation of sperm motility. The

SAR was supplemented with dissolved gentamicin sulphate salt (Sigma-Aldrich, New South Wales,

Australia) with a final concentration equivalent to 100 µg/mL-1. This dose was based on the results

of Viveiros et al. (2010) who reported a decrease in bacterial growth and improved longevity in

chilled freshwater fish (Brycon orbignyanus) semen treated with 100 µg/mL-1 gentamicin. There are

68

currently no recommendations for optimal concentrations of antibiotics in amphibian spermatozoa

media.

A split-sample experimental approach was adopted to test the effects of activation of sperm

motility and cryopreservation (with and without cryoprotectant) on the integrity of cane toad sperm

DNA. This approach involved evenly distributing testis segments from each male across the three

experimental treatments; a summary of the experimental design is depicted in Figure 4.1. Both

testes from each individual (n = 25 males) were divided into three equal segments and one segment

from each testis macerated in a 1.5 mL Eppendorf tube containing 1 mL of room temperature 1:1

SAR to maintain inactivation of sperm motility. Sperm concentration was quantified for each

suspension using a Makler Chamber (Emgrid Australia, South Australia, Australia) and adjusted to

attain a final concentration of approximately ~10 × 106 spermatozoa mL–1. To examine variation in

endogenous levels of SDF, the SCD test was run at time 0 (T0), immediately following testis

maceration for all 25 males. Another two testis segments, one from each testis, were macerated in a

1.5 mL Eppendorf tube containing 1 mL of room temperature 1:4 SAR (diluted with dH20; 40

mOsmol kg-1) to activate sperm motility. Identification of sperm motility under x 400 magnification

confirmed activation. A dynamic assessment of sperm viability and motility was then carried out at

T0, T3 and T6 hours. From the initial population of 25 cane toads, six males with optimal sperm

concentration (≥ 100 × 106 spermatozoa mL–1; Browne and Zippel 2007) were selected. For these

males, SDF was assessed at T0 immediately following testis maceration for both inactivated (1:1

SAR) and activated (1:4 SAR) sperm suspensions, as well as after T3 and T6 hours of incubation at

room temperature. A dynamic assessment of sperm viability and motility (in activated samples

only) was also carried out at these time points.

The remaining two testis segments were prepared for cryopreservation according to the

protocol developed by Browne et al. (1998) for cane toad spermatozoa. Testis segments were

macerated in a 1.5 mL Eppendorf tube containing 1 mL of chilled (4 °C) 15% (v/v) dimethyl

sulfoxide (DMSO; Sigma-Aldrich, New South Wales, Australia) made up with 1:1 SAR (containing

100 µg/mL-1 gentamicin sulphate salt; Sigma-Aldrich, New South Wales, Australia) and

supplemented with 10% (w/v) sucrose. Sperm suspensions were loaded into 250 µL Cassou straws

(IMV Technologies, Maple Grove, Minnesota, USA) and sealed with moistened PVC powder. In

order to compare the effect of cryopreservation with and without cryoprotectant on sperm longevity,

straws of 250 µL were also frozen for sperm suspensions in 1:1 SAR. Straws were chilled on ice

before being cooled in a single batch in a programmable controlled rate cooling chamber (Freeze

69

Control CL-863, Cryologics Pty Ltd., Mulgrave, Victoria, Australia) using a slow cooling rate (see

Browne et al. 1998 for cooling protocol) and were subsequently stored in LN2. Following a period

of cold storage, the straws were thawed in a water bath at 25 °C for 1 minute before dilution with

room temperature dH20 to activate sperm motility employing a 1:4 and 1:40 dilution rate for straws

containing SAR and DMSO respectively. A dynamic assessment of SDF, sperm motility and

viability was then carried out at T0 immediately following thawing and again following incubation

for T3 and T6.

Figure 4.1. Representation of the experimental design detailing the division and treatment of testes for an individual cane toad; SAR – simplified amphibian ringer; DMSO – dimethyl sulfoxide; T – time of incubation in hours; SCD – sperm chromatin dispersion test; ACT – activated spermatozoa; INACT – inactivated spermatozoa.

70

4.3.2 Assessment of sperm motility and viability

Sperm suspensions were assessed for percentage of sperm motility immediately following

activation. Whole sperm suspensions were thoroughly homogenised and an aliquot of 8 µL loaded

onto a glass slide and examined at x 400 magnification using a phase-contrast microscope (CH-2

Olympus, Japan). Spermatozoa were classified as either motile or immotile. Motile spermatozoa

included all spermatozoa showing either progressive or non-progressive motility. Progressively

motile spermatozoa moved linearly regardless of speed, while non-progressively motile

spermatozoa showed no forward displacement (Christensen 2002; WHO 2010). Two replicate

counts of 100 spermatozoa were assessed for each sample and the percentage of total motility was

determined.

Sperm viability was assessed using a dual emission technique incorporating the nucleic

stains SYBR-14 and propidium iodide (LIVE/DEAD® Sperm Viability Kit Imaging Protocol,

Molecular Probes, ThermoFisher Scientific, Waltham, Massachusetts, USA). SYBR-14 permeates

intact plasma membranes and produces a green fluorescence and propidium iodide permeates

damaged plasma membranes and produces a red fluorescence. Sperm suspensions were

homogenised and an aliquot of 20 µL combined with 1 µL SYBR-14 and incubated in the dark for

two minutes. Following the addition of 1 µL propidium iodide and incubation for an additional two

minutes, the aliquot was mixed and 8 µL deposited on a glass slide and examined at x 400

magnification using an epifluorescence microscope (BA310 Epi-LED Motic, Barcelona, Spain).

Spermatozoa were classified as either live (green fluorescence) or dead (red fluorescence). Those

spermatozoa found to exhibit dual fluorescence were presumed to represent moribund or slightly

damaged spermatozoa that were beginning to show signs of impaired membrane integrity and were

classified as dead (Garner and Johnson 1995). Dead spermatozoa were further classified into two

categories; dead-compact, where spermatozoa retained normal elongated, rod-like morphology and

dead-swollen, where spermatozoa appeared enlarged with less defined morphology. Two replicate

counts of 100 spermatozoa were assessed for each sample.


The extent of DNA fragmentation in cane toad spermatozoa was assessed using an

amphibian prototype kit (Xenopus-Halomax®; Halotech DNA SL, Madrid, Spain) adapted from the

Sperm-Halomax® kit applied previously to the African clawed frog (Xenopus laevis; Pollock et al.

71

2015). Briefly, 0.5 mL Eppendorf tubes containing 500 µL of low melting point agarose were

suspended on a float in a beaker of water and heated in a microwave oven for five minutes. The

liquefied agarose was then equilibrated to 35 °C using a heated stage. An aliquot of the extended

sperm suspension (25 µL) was gently mixed into 50 µL agarose and 10 µL of this mixture deposited

on a pre-treated glass slide (provided in the Xenopus-Halomax® kit) and covered with a glass

coverslip (22 x 22 mm). The slides were then chilled for five minutes on a glass place in a

refrigerator at 4 °C. Once the agarose had solidified, the coverslip was removed and the slide placed

(maintained at a horizontal position) on a plastic support in a petri dish. Lysing solution (enough to

fill the viewing field) containing 2 M NaCl, 0.5% SDS, 0.01 Triton X, 0.2 M Tris-HCL and 0.02 M

EDTA; pH 7 (provided in the Xenopus-Halomax® kit) was added to the slide before the petri dish

was covered and left to incubate for five minutes. Excess lysing solution was removed by gently

flushing the slide with dH20. The slide was then dehydrated in sequential ethanol washes (70, 90

and 100%) for two minutes each and finally left to air-dry. Spermatozoa were mounted with equal

parts of fluorochromes fluoRed® and fluoGreen® (Halotech DNA SL, Madrid, Spain) prepared in

mounting medium (provided in the fluorochrome kit) and observed using an epifluorescence

microscope (BA310 Epi-LED Motic, Barcelona, Spain). Black and white images were captured

using 12-bit format with a CCD (Leica DFC350 FX, Leica Microsystems) and stored as Tiff black

and white 12-bit format. Two replicate counts of 100 spermatozoa were analysed per sample and

the percentage of spermatozoa with fragmented DNA calculated and the average taken as the result.


Statistical analyses were performed using the Statistical Package for the Social Sciences

(SPSS, Inc., Chicago, USA). The relationships between SDF, motility and viability in fresh and

cryopreserved spermatozoa were determined using a linear logistic regression model. Differences

were considered significant with P-values less than 0.05.

4.4 Results

4.4.1 Sperm morphology

The SCD test revealed three primary sperm morphotypes indicative of varying degrees of

DNA fragmentation. Amphibian sperm morphotype 1 (ASM-1) corresponded to spermatozoa with

intact DNA and was characterised by a small, dense halo of chromatin dispersion surrounding an

72

elongated and tapered nuclear core (Figure 4.2ab). The chromatin dispersion halo of ASM-2 was

similar to that seen in ASM-1, but the original morphology of the nucleus had swollen or

decondensed into a more spherical appearance; while there was clearly some structural change to

the nucleus, the DNA of these nuclei was typically not fragmented. ASM-3 represented

spermatozoa that contained damaged DNA and displayed a large, stellar halo of dispersed

chromatin with multiple fragments of DNA (Figure 4.2ac). This morphotype typically showed less

fluorescence than ASM-1 or ASM-2 as the DNA fragments were more widely dispersed in the

microgel. In spermatozoa with severely damaged DNA, the nuclear core appeared diminished in

size and the halo was difficult to visualise due to the extensive dispersion of the chromatin away

from the nuclear core.

Figure 4.2. Cane toad sperm nuclear morphotypes after the SCD test. (a) Spermatozoa observed at T6 with varying degrees of DNA damage. Arrow indicates spermatozoon beginning to show evidence of DNA fragmentation. (b) Spermatozoon with an elongated core and small, compact halo of dispersed chromatin representing non-fragmented DNA (ASM-1). (c) Spermatozoon with diminished nuclear core and large, stellar halo of dispersed chromatin corresponding to fragmented DNA (ASM-3).

4.4.2 Dynamics of DNA fragmentation in fresh spermatozoa

The basal level of SDF observed at T0 in 25 freshly collected cane toad testicular

homogenates was consistently low, with values ranging from 0 – 5% for both inactivated (mean ±

73

SEM; 2.1 ± 0.3%; Table 4.1) and activated spermatozoa (mean ± SEM; 2.8 ± 0.3%). While there

was significant variation in SDF at T0 between cane toads in each group (p = 0.0002), there was

little difference between the overall levels of SDF observed between inactivated and activated

spermatozoa (p = 0.5178; Figure 4.3). The rate of increase in SDF was negligible with little

difference in levels of SDF observed at T6 between inactivated and activated spermatozoa (p =

0.5094). Values obtained after 6 hours of incubation did not exceed an upper value of 6.5% and

9.5% for inactivated and activated spermatozoa respectively.

Figure 4.3. Logistic model plots representing the dynamics of SDF in cane toad spermatozoa over 6 hours of incubation at 22 °C. Frsh In – fresh, inactivated spermatozoa; Frsh Ac – fresh, activated spermatozoa; Sar Ac – spermatozoa cryopreserved in simplified amphibian ringer; DMSO – spermatozoa cryopreserved in dimethyl sulfoxide; SDF – sperm DNA fragmentation.

74

Table 4.1. Mean percentage and regression analysis of SDF, viability and motility in fresh cane toad spermatozoa incubated at 22 °C for 6 hours.

Time of incubation (hours) Source of variation between cane toads

T0 T3 T6 (β0: intercept) (β1: slope)

Mean SDF ± SEM (%) Initial SDF at T0

(%)

Rate of increase in

SDF (%)

INACT 2.1 ± 0.3 3.6 ± 0.4 4.3 ± 0.3 0.0002* 0.1658

ACT 2.8 ± 0.3 3.8 ± 0.5 5.0 ± 0.5 0.0002* 0.0883

Source of variation between INACT and ACT groups: 0.5178 0.5094

Mean total motility ± SEM (%) Initial motility at

T0 (%)

Rate of decrease in

motility (%)

ACT 35.5 ± 1.5 19.0 ± 1.2 9.5 ± 1.2 < 0.0001* < 0.0001*

Mean total viability ± SEM (%) Initial viability at

T0 (%)

Rate of decrease in

viability (%)

INACT 64.6 ± 2.0 61.4 ± 1.8 57.1 ± 1.6 < 0.0001* 0.0064*

ACT 65.6 ± 2.1 63.7 ± 1.9 59.3 ± 1.7 < 0.0001* 0.0008*

Source of variation between INACT and ACT groups: 0.0011* 0.1244

Mean dead-swollen ± SEM (%) Initial dead-

swollen at T0 (%)

Rate of increase in

dead-swollen (%)

INACT 15.6 ± 1.8 24.1 ± 1.8 35.4 ± 2.0 < 0.0001* < 0.0001*

ACT 17.1 ± 2.4 20.4 ± 2.2 27.6 ± 2.7 < 0.0001* < 0.0001*

Source of variation between INACT and ACT groups: < 0.0001* < 0.0001*

SDF – sperm DNA fragmentation; T(x) – time of incubation in hours; ACT – activated spermatozoa; INACT – inactivated spermatozoa; dead-swollen – dead-swollen spermatozoa; SEM – standard error of mean; *denotes significant p-value.

4.4.3 Dynamics of motility in fresh spermatozoa

Total motility in fresh cane toad spermatozoa at T0 varied considerably between the 25

males (p < 0.0001; Figure 4.4), and ranged from 26 – 53% (mean ± SEM; 35.5 ± 1.5%; Table 4.1).

Sperm motility declined significantly over the incubation period (p < 0.0001), and had dropped to <

10% in 65% of the males by T6 (mean ± SEM; 9.5 ± 1.2%). The lowest recorded percentage of

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motility at T6 was for Male 10, being < 1%. The highest percentage of sperm motility observed for

an individual at T6 was 20.5% (in Male 9).

Figure 4.4. Logistic model plots representing the dynamics of sperm motility in cane toad spermatozoa over 6 hours of incubation at 22 °C. Frsh In – fresh, inactivated spermatozoa; Frsh Ac – fresh, activated spermatozoa; Sar Ac – spermatozoa cryopreserved in simplified amphibian ringer; DMSO – spermatozoa cryopreserved in dimethyl sulfoxide.

4.4.4 Dynamics of viability in fresh spermatozoa

The initial percentage of sperm viability (live sperm) observed at T0 varied significantly

between the 25 cane toads for both inactivated and activated spermatozoa (p < 0.0001; Figure 4.5).

Values observed at T0 ranged from 42 – 83% and 52.5 – 86% for inactivated and activated

spermatozoa respectively. While there was a significant difference between the percentage of viable

spermatozoa observed at T0 for the two groups (p = 0.0011), the actual difference between the

mean (± SEM) percentage viability for inactivated (64.6 ± 2.0%) and activated (65.6 ± 2.1%)

spermatozoa was negligible (Table 4.1). Viability declined steadily for all spermatozoa irrespective

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of whether the spermatozoa were inactivated or in the activated state (p = 0.1244). After 6 hours of

incubation, mean values were significantly lower than values observed at T0 for both inactivated (p

= 0.0064) and activated (p = 0.0008) spermatozoa.

There was significant variation in the percentage of dead-swollen spermatozoa observed at

T0 (p < 0.0001), with values ranging from 5 – 29% and 4 – 39% for inactivated and activated

spermatozoa respectively (Figure 4.6). However, the actual difference between the mean (± SEM)

percentage of dead-swollen spermatozoa for inactivated (15.6 ± 1.8%) and activated (17.1 ± 2.4%)

samples was negligible (Table 4.1). The proportion of dead-swollen spermatozoa increased

significantly over the incubation period for all samples, however was higher for inactivated

compared to activated samples at T6 (p < 0.0001).

Figure 4.5. Logistic model plots representing the dynamics of sperm viability in cane toad spermatozoa over 6 hours of incubation at 22 °C. Frsh In – fresh, inactivated spermatozoa; Frsh Ac – fresh, activated spermatozoa; Sar Ac – spermatozoa cryopreserved in simplified amphibian ringer; DMSO – spermatozoa cryopreserved in dimethyl sulfoxide.

77

Figure 4.6. Logistic model plots representing the dynamics of chromatin swelling (dead-swollen spermatozoa) in cane toad spermatozoa over 6 hours of incubation at 22 °C. Frsh In – fresh, inactivated spermatozoa; Frsh Ac – fresh, activated spermatozoa; Sar Ac – spermatozoa cryopreserved in simplified amphibian ringer; DMSO – spermatozoa cryopreserved in dimethyl sulfoxide.

4.4.5 Dynamics of DNA fragmentation in cryopreserved spermatozoa

The basal level of SDF observed at T0 immediately following thawing was similar to that

seen prior to cryopreservation for spermatozoa cryopreserved in either DMSO (mean ± SEM; 4.2 ±

0.6%; Figure 4.3) or SAR (mean ± SEM; 5.1 ± 0.5%; p = 0.9966). A negligible increase in SDF

was observed at T6 for spermatozoa cryopreserved in DMSO (p = 0.6644), reaching a maximum of

17.5% in Male 6 (mean ± SEM; 9.4 ± 1.9%). The spermatozoa of Male 10 appeared particularly

robust to cryopreservation stress (DMSO) and showed almost no change in SDF, reaching a

maximum of only 3.5% at T6. Almost all spermatozoa frozen in SAR showed a marked increase in

SDF from T3 onwards (p = 0.0224; Table 4.2), with the exception of Male 10 that showed a much

lower level of SDF (9%) compared to the mean (± SEM; 29.6 ± 10.3%). At T6, the level of SDF in

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this male had increased to a level comparable with other males, although was at the lower end of the

observed range (72 – 97%). A notable anomaly was Male 5, where the level of SDF at T6 was only

29%.

Table 4.2. Mean percentage and regression analysis of SDF, viability and motility in cryopreserved-thawed cane toad spermatozoa incubated at 22 °C for 6 hours.

Time of incubation (hours) Source of variation between cane toads

T0 T3 T6 (β0: intercept) (β1: slope)

Mean SDF ± SEM (%) Initial SDF at T0

(%)

Rate of increase in

SDF (%)

DMSO 4.2 ± 0.6 6.5 ± 1.5 9.4 ± 1.9 0.2650 0.6644

SAR 5.1 ± 0.5 29.6 ± 10.3 75.9 ± 10.1 0.2359 0.0224*

Source of variation between DMSO and SAR groups: 0.9966 < 0.0001*

Mean total motility ± SEM (%) Initial motility at

T0 (%)

Rate of decrease in

motility (%)

DMSO 2.8 ± 0.8 0.0 0.0 N/A N/A

SAR 0.0 0.0 0.0

Mean total viability ± SEM (%) Initial viability at

T0 (%)

Rate of decrease in

viability (%)

DMSO 19.6 ± 3.0 10.0 ± 2.1 10.2 ± 2.5 < 0.0001* < 0.0001*

SAR 6.8 ± 1.0 8.7 ± 1.0 5.2 ± 1.5 0.0974 0.0479*

Source of variation between DMSO and SAR groups: < 0.0001* 0.0005*

Mean dead-swollen ± SEM (%) Initial dead-

swollen at T0 (%)

Rate of increase in

dead-swollen (%)

DMSO 2.7 ± 1.4 1.7 ± 1.3 4.3 ± 1.9 < 0.0001* 0.0004*

SAR 69.8 ± 4.9 75.5 ± 3.2 85.0 ± 3.2 < 0.0001* 0.0298*

Source of variation between DMSO and SAR groups: < 0.0001* < 0.0001*

SDF – sperm DNA fragmentation; T(x) – time of incubation in hours; DMSO – spermatozoa cryopreserved in dimethyl sulfoxide; SAR – spermatozoa cryopreserved in simplified amphibian ringer; dead-swollen – dead-swollen spermatozoa; SEM – standard error of mean; *denotes significant p-value.

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4.4.6 Dynamics of motility in cryopreserved spermatozoa

Following thawing and activation at T0, spermatozoa cryopreserved in DMSO showed only

very low levels of motility, no greater than 5% (mean ± SEM; 2.8 ± 0.8%; Figure 4.4). All

spermatozoa frozen in SAR without cryoprotectant were immotile at T0 after thawing and

activation.

4.4.7 Dynamics of viability in cryopreserved spermatozoa

At T0 immediately following thawing, viability of spermatozoa cryopreserved in DMSO

varied significantly between males (p < 0.0001), with values ranging from 12.5 – 30% (mean ±

SEM; 19.6 ± 3.0%). Sperm viability was significantly lower in samples frozen in SAR without

cryoprotectant (mean ± SEM; 6.8 ± 1.0%; p < 0.0001) and showed less variation between males

(range of 2 – 9%; p = 0.0974; Figure 4.5). Viability of spermatozoa cryopreserved in DMSO had

declined significantly by T3 (< 0.0001; Table 4.2), however then appeared to plateau and remain

stable for the remainder of the incubation period, with a maximum value of 21% for Male 5 (mean

± SEM; 10.2 ± 2.5%). Spermatozoa frozen in SAR also showed a significant decrease in viability at

T6 (p = 0.0298), reaching a maximum value of 10% for Male 5 (mean ± SEM; 5.2 ± 1.5%).

There was significant variation between toads in the percentage of dead-swollen

spermatozoa observed at T0 following thawing (p < 0.0001), with values ranging from 49 – 80%

and 0.5 – 9.5% for spermatozoa frozen in SAR and DMSO respectively. There was a highly

significant increase (p < 0.0001) in the percentage of dead-swollen spermatozoa at T0 for samples

frozen in SAR (mean ± SEM; 69.8 ± 4.9%; Figure 4.6) compared to cryopreservation in DMSO.

The percentage of dead-swollen spermatozoa had increased significantly by T6 (p = 0.0298),

reaching a maximum value of 93% (mean ± SEM; 85.0 ± 3.2%) for spermatozoa frozen in SAR.

While sperm samples cryopreserved in DMSO also showed a significant increase in the percentage

of dead-swollen spermatozoa at T6 (p = 0.0004), values observed at this time were still low, with no

dead-swollen spermatozoa observed for Male 5 (mean ± SEM; 4.3 ± 1.95).

4.5 Discussion

The results of this study have demonstrated that the amphibian SCD test (Xenopus-

Halomax®; Halotech DNA SL, Madrid, Spain; Pollock et al. 2015) can successfully identify

80

fragmented DNA in cane toad spermatozoa. Chromatin dispersion patterns revealed by the SCD test

were comparable to the sperm nuclear morphotypes described in African clawed frog (Pollock et al.

2015) spermatozoa and enabled the differential classification of intact and damaged sperm nuclei.

Assessment of SDF at the population level in fresh spermatozoa, as well as the dynamics of sperm

chromatin longevity, revealed that cane toad sperm DNA was highly stable throughout the duration

of the incubation period and has provided an index against which future dynamic studies of

amphibian SDF can be compared. It is generally accepted that the presence or absence of

protamines/cysteine residues and associated disulphide bonds to stabilise the sperm chromatin is

likely to influence the susceptibility of DNA to fragmentation in vitro (Cortés-Gutiérrez et al.

2014b). While the sperm nuclear basic protein (SNBP) composition of cane toad spermatozoa is

characterised by arginine-rich protamines (Ausio et al. 2007), the presence of cysteine residues has

not been confirmed. The conspecific Japanese common toad (Bufo japonicus; Takamune et al.

1991) has only low levels of cysteine residues in the sperm protamines. The chromatin stability

observed in cane toad spermatozoa may also therefore reflect a high arginine content of the SNBPs,

a feature proposed to increase charge density and greatly enhance the compaction of the DNA

(Lewis and Ausio 2002). Interestingly, African clawed frog (Pollock et al. 2015) and common

dunnart (Sminthopsis murina; Johnston et al. 2015) spermatozoa also displayed a remarkable level

of chromatin stability despite lacking cysteine-bearing protamines (Yokota et al. 1991; Ausio et al.

2007; Johnston et al. 2015). Clearly, the interaction between SNBP composition and the resulting

stability of the sperm chromatin is complex, making predictions about the expected rate of damage

in a given species difficult.

Activation of sperm motility did not appear to have a significant effect on either the stability

of the sperm chromatin or sperm viability observed in cane toad spermatozoa. In amphibians with

external fertilization, the spermatozoa remain inactive in vivo due to the isotonic environment of the

testes, only becoming motile following spermiation/spawning into hypo-tonic fresh water with an

osmolality of < 50 mOsmol kg-1 (Browne and Zippel 2007; Kouba et al. 2009). The need to activate

motility in amphibian spermatozoa prior to use with ART, imposes high physiological stress to

maintain the osmotic balance of the spermatozoa and it is a significant factor impacting the

longevity of both fresh and cryopreserved-thawed spermatozoa (Browne et al. 2015); as the energy

reserves of the spermatozoa are exhausted, osmoregulation fails resulting in damage to the plasma

membrane and ultimately cell death (Browne et al. 2015). Therefore, it would be reasonable to

assume that the progressive loss of DNA integrity and sperm viability would proceed more rapidly

in species that require osmotically-induced activation compared to internally fertilizing species. One

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would at least expect differences when comparing parameters of sperm quality in activated and

inactivated spermatozoa. Indeed, this is certainly the case for fish (Tinca tinca; López-Fernández et

al. 2009 and zebrafish; Danio rerio; Gosálvez et al. 2014), where an increase in SDF occurred only

a few minutes after sperm activation. However, in the 25 males examined in this study, the level of

SDF did not exceed 9.5% for inactivated or activated spermatozoa after 6 hours of incubation.

These findings are comparable with results reported in African clawed frog (Pollock et al. 2015).

Consequently, it does not appear that osmotic stress associated with the activation of sperm motility

is a significant causal factor of chromatin damage in cane toad spermatozoa. The accelerated loss of

the normal morphology observed for inactivated spermatozoa in this study is interesting and

suggests that a protective mechanism may function in cane toad spermatozoa that is triggered only

following osmotically-induced activation, perhaps as an adaptation to the often harsh environmental

conditions where cane toads are found.

In the current experiment, the degree of sperm motility observed at T0 in fresh samples was

lower than previously reported for the cane toad (Browne et al. 1998; Fitzsimmons et al. 2007).

Activation of motility in spermatozoa from anuran amphibians occurs by deposition into hypo-tonic

fresh/pond water with a low osmolality of < 50 mOsmol kg-1 (Browne and Zippel 2007; Kouba et

al. 2009). Despite ensuring that the osmolality of the diluted SAR (40 mOsmol kg-1) was within the

appropriate range to sufficiently activate and prolong the duration of sperm motility in this species,

total motility did not exceed 53%. The lack of standardised protocols for the handling and activation

of amphibian spermatozoa should be taken into consideration when comparing data across

laboratories. The composition and osmolality of the activation media particularly influence the

degree of sperm motility achieved; Fitzsimmons et al. (2007) found a strong interaction between

osmolality and addition of the energy substrate theophylline to the SAR that can impact the

percentage of motile cane toad spermatozoa. It is possible that the addition of antibiotic to the SAR

may have affected the percentage of sperm motility observed in this study. When amphibian sperm

suspensions are collected via testes extraction and maceration, bacterial contamination occurs from

the surrounding fluids and tissues (Silla et al. 2014). Excessive bacterial growth can generate

harmful metabolic by-products such as reactive oxygen species (ROS), which in turn leads to the

onset of oxidative stress that can lead to chromatin cross-linking, base changes and DNA strand

breaks (Martínez-Páramo et al. 2012; Said et al. 2012). It is, therefore important to control the

proliferation of bacteria within sperm suspensions, particularly where samples will be chilled-stored

for short periods prior to being utilised for IVF. However, there is some evidence in fish

(Christensen and Tiersch 1996), stallion (Jasko et al. 1993; Aurich and Spergser 2007) and

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amphibian species (Germano et al. 2013; Silla et al. 2014), that supplementation of sperm extenders

with broad-spectrum antibiotics such as penicillin, streptomycin and gentamicin can negatively

impact sperm function in terms of motility and viability, possibly as a result of species-specific

toxicity. With only two studies to date reporting the use of antibiotics with amphibian spermatozoa

(Germano et al. 2013; Silla et al. 2014), optimal concentrations have yet to be identified.

The spermatozoa of the cane toad appear to be highly resistant to cryopreservation-induced

DNA damage (particularly Male 10). Not surprisingly, there was little difference between values of

SDF obtained at the onset of incubation for fresh and frozen-thawed samples (either DMSO or

SAR). Similar findings have been reported in stallion (López-Fernández et al. 2007), ram (López-

Fernández et al. 2008a), dog (Prinosilova et al. 2012), koala (Johnston et al. 2012b) and rainbow

trout (Labbe et al. 2001). Although the chromatin structure may appear unaltered by the

cryopreservation process immediately upon thawing, the varying resistance of sperm DNA to

stressors associated with post-thaw incubation ex vivo, particularly influences the longevity of

sperm DNA quality (Gosálvez et al. 2011a). The formation of intracellular ice during the freezing

process promotes detrimental changes to the spermatozoa, particularly to the plasma membrane and

mitochondria (Martínez-Páramo et al. 2012). ROS produced by the membrane-compromised and

dying sperm population have been shown to facilitate oxidative stress that can potentially increase

the level of SDF over time (López-Fernández et al. 2007; Gosálvez et al. 2011a). These effects are

likely to be exacerbated in spermatozoa frozen without cryoprotectant, such as DMSO; DMSO is a

membrane-permeable solute that acts by displacing intracellular water, limiting the degree of

damage to the spermatozoa through ice crystallisation and cell shrinkage (Santo et al. 2011). The

significant increase in SDF observed at T6 for samples frozen in SAR only was therefore, not

surprising. Furthermore, there is considerable inter-species, as well as inter-individual variation in

the rate at which the sperm DNA fragments following thawing. For example, López-Fernández et

al. (2008a) reported aggressive DNA damage in ram semen samples with levels of SDF greater than

50% observed within 5 hours of thawing. Although cryopreserved cane toad spermatozoa in the

present study showed a marked increase in SDF after 6 hours of incubation, the proportion of sperm

with SDF was still low, being less than 17.5% (for spermatozoa cryopreserved in DMSO). This

level of SDF is comparable to that reported in Western clawed frog spermatozoa following

cryopreservation (17.65%), although it is not clear whether this result was obtained immediately

after thawing or following a period of incubation (Morrow et al. 2014). This finding is also similar

to that seen in the koala (Johnston et al. 2012b) where levels of damaged DNA plateaued after 8

hours of incubation and were typically less than 10% even after 48 hours. Unfortunately, the

83

incubation period in this study was too short to show the full dynamics of DNA fragmentation in

cane toad spermatozoa, making predictions about the longevity of the sperm chromatin problematic.

While the integrity of cane toad sperm chromatin remained fairly stable during the first

hours post-thawing, it is well documented that the process of cryopreservation induces the

impairment of other parameters of sperm quality in amphibian spermatozoa such as motility,

viability, plasma membrane integrity and fertilisation rate (Sargent and Mohun 2005; Mansour et al.

2009). This was particularly evident for Male 10 where the spermatozoa appeared particularly

robust against cryopreservation, yet showed a very poor recovery of motility and being consistent

with what was found in other males. The results of the present study have clearly demonstrated a

very poor recovery of motility and viability in cane toad spermatozoa following freeze-thawing.

Although it was not surprising that spermatozoa frozen without cryoprotectant showed no motility

after activation, spermatozoa cryopreserved in DMSO showed only a marginal increase in motility

at T0. This may be a result of toxicity caused by the addition of antibiotic to the SAR, given that the

degree of sperm motility observed in fresh samples was also lower than previously reported for the

cane toad (Browne et al. 1998; Fitzsimmons et al. 2007). The marked increase in the incidence of

abnormal sperm nuclear morphology (e.g. dead-swollen morphotypes) for spermatozoa frozen

without cryoprotectant is interesting and may be associated with chromatin swelling. Chromatin

swelling or relaxation has also been noted in membrane-compromised spermatozoa of the koala

(Johnston et al. 2006) and common wombat (Vombatus ursinus; Johnston et al. 2006) following

thawing as well as in fresh African clawed frog (Pollock et al. 2015) spermatozoa. In these species,

SNBPs devoid of disulphide bonds in addition to the extensive presence of ALS may render the

DNA more susceptible to denaturation and swelling (Zee et al. 2009). It is possible that the

incidence of dead-swollen spermatozoa observed in the cane toad in the present study is consistent

with the transition from ASM-1 to ASM-2 described in African clawed frog (Pollock et al. 2015)

and therefore, that dead-swollen spermatozoa do not necessarily contain fragmented DNA. The

findings presented in this study support that DNA fragmentation is only a minor component of

cryoinjury and that poor recovery of motility and viability (plasma membrane integrity) are likely to

have a greater effect on the useful lifespan of cane toad spermatozoa post-thaw. Information

presented in this study will also be applicable to threatened and endangered amphibians.

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CHAPTER 5: General discussion

5.1 Introduction

The fundamental aim of this project was the development and validation of the SCD test for

amphibian spermatozoa with the purpose of examining the post-thaw longevity of cryopreserved

spermatozoa. Co-validation of the SCD procedure with other techniques of assessing DNA damage

confirmed that the SCD test was able to produce differential chromatin dispersion patterns for the

accurate assessment of SDF. The studies presented in this thesis provided the first evidence of SDF

in amphibian spermatozoa, in addition to novel insights into the chromosomal organisation of

African clawed frog (Xenopus laevis) sperm DNA. Findings suggested that alkali-labile sites

(revealed by the double-comet assay as extensive single-stranded DNA breaks) were a constitutive

feature of African clawed frog sperm chromatin and that double-stranded DNA breaks represented

true DNA damage. Subsequent investigations applied the SCD test to cryopreserved-thawed

amphibian spermatozoa and provided insight into cryoinjury of the sperm chromatin and the

resulting effect on the potential application of ART. This was achieved by examining the dynamic

changes to the basal level of DNA fragmentation in fresh African clawed frog (Chapter 3) and cane

toad (Bufo marinus; Chapter 4) spermatozoa in comparison with the post-thaw integrity of

cryopreserved sperm DNA. Additional samples of African clawed frog and cane toad spermatozoa

were frozen without cryoprotectant to provide further insight into the mechanisms of cryoinjury. An

IVF procedure using cryopreserved African clawed frog spermatozoa was implemented to assess

the extent to which freeze-thawing and the associated DNA damage was correlated with a decline in

fertilisation rate. Chapter 4 also examined the relationship between cryoinjury and additional

parameters of sperm quality. The sperm DNA of both the African clawed frog and cane toad was

revealed to be highly stable, although this varied between animals. Assessment of SDF at the

population level in Chapter 4 revealed considerable inter-individual variation and provided an index

against which future dynamic studies of amphibian SDF can be compared. Further studies are

required to explore the full extent of amphibian sperm chromatin longevity.

5.2 Validation of the sperm chromatin dispersion (SCD) test in amphibian spermatozoa

This thesis aimed to apply the SCD test to the African clawed frog to enable visualisation of

DNA fragmentation in the first amphibian sperm model. There were previously no such studies that

had examined DNA fragmentation in amphibian spermatozoa. A species-specific modified lysing

85

solution (Xenopus-Halomax®; Halotech DNA SL, Madrid, Spain; Pollock et al. 2015) was used for

protein depletion to ensure a targeted and validated approach to DNA fragmentation assessment in

this new taxon. The SCD test is based on the principle that controlled denaturation and removal of

nuclear proteins using a lysing solution results in nucleoids with a central core and a peripheral halo

from the dispersion of damaged chromatin (Fernández et al. 2003). Quantifying the size and the

incidence of these haloes by fluorescence microscopy provided an index of the amount of DNA

damage present in each sample.

The SCD procedure applied to African clawed frog spermatozoa in Chapter 2 revealed three

distinctive amphibian sperm morphotypes. ASM-3 was the result of massive chromatin dispersal

away from the nucleus and was clearly associated with severely damaged DNA. The morphology of

the ASM-1 sperm nucleus retained its general shape and showed evidence of only a small halo of

DNA migration, while ASM-2 sperm nuclei showed a characteristic change in shape of the nucleus

from a spiral elongated form to a rounded head morphology. The loss of the original nuclear

morphology in ASM-2 was interpreted as chromatin swelling or relaxation that has been observed

in other species devoid of disulphide bonds in their protamines (Johnston et al. 2006; 2007; López-

Fernández et al. 2009; Johnston et al. 2012b). In these species, it is possible that the nuclear basic

proteins of the spermatozoon render the DNA more susceptible to protein depletion (Zee et al.

2009). Chromatin swelling in African clawed frog spermatozoa without DNA fragmentation may be

due to the extensive presence of alkali labile sites (ALS). Indeed, the pervasiveness of structural

SSB revealed in sperm nuclei by the alkaline step of the double-comet assay suggests that ALS are

a constitutive feature of African clawed frog sperm chromatin. This theory was further supported by

the observation that all spermatozoa, including those exhibiting small compact chromatin halos of

undamaged DNA, expressed positive in situ nick translation labelling, given that the DNA

nucleotide is incorporated into the free ends of single- or double-stranded DNA breaks. The

prevalence of ALS in the mature spermatozoa of the African clawed frog may be related to the

absence of sperm-specific protamines in the chromatin, the cysteine residues of which are essential

for the formation of disulphide bonds (Yokota et al. 1991; Ausio et al. 2007).

The results of the investigation presented in Chapter 4 demonstrated that the amphibian

SCD test (Xenopus-Halomax®; Halotech DNA SL, Madrid, Spain; Pollock et al. 2015) developed

for the African clawed frog (Chapter 2) can successfully identify fragmented DNA in cane toad

spermatozoa. Chromatin dispersion patterns revealed by the SCD test were comparable to the sperm

morphotypes described in the African clawed frog (Pollock et al. 2015) and confirmed that the

86

Xenopus-Halomax® assay was appropriate for the differential classification of intact and damaged

sperm nuclei in cane toad, the third amphibian species to date in which sperm DNA fragmentation

has been reported.

5.3 Dynamics of sperm DNA fragmentation in fresh amphibian spermatozoa

Sperm DNA fragmentation is a highly dynamic process and a certain amount of iatrogenic

DNA damage is to be expected once the spermatozoa are incubated ex vivo for ART (Gosálvez et

al. 2011a). A progressive loss of sperm DNA quality has been observed in a number of species,

including but not limited to, the ram (López-Fernández et al. 2008a), stallion (López-Fernández et

al. 2007), boar (Pérez-Llano et al. 2010), donkey (Equus asinus; Cortés-Gutiérrez et al. 2008a),

rhinoceros (Diceros bicornis, Rhinoceros unicornis, Ceratotherium simum; Portas et al. 2009),

koala (Phascolarctos cinereus; Zee et al. 2009; Johnston et al. 2012b), short-beaked echidna

(Tachyglossus aculeatus; Johnston et al. 2009) and fish (Tinca tinca; López-Fernández et al. 2009

and zebrafish; Danio rerio; Gosálvez et al. 2014). Comparative analysis of the dynamics of sperm

DNA fragmentation in different species highlights the varying resistance of sperm chromatin to

external stressors. Not surprisingly, there is considerable inter-species, as well as inter-individual

variation in the basal level of damaged DNA and the rate at which the DNA fragments. The large

variation in the protein composition of sperm DNA of different species is proposed to play a pivotal

role in determining the longevity of sperm DNA quality.

In this thesis, the dynamics of DNA fragmentation in fresh spermatozoa were examined in

two amphibian species with differing SNBP compositions: African clawed frog with protamine-like

proteins and no cysteine residues in the spermatozoa (Yokota et al. 1991; Ausio et al. 2007;

Chapters 2 and 3) and cane toad (Chapter 4) with protamines (Ausio et al. 2007) and perhaps low

levels of cysteine residues in the spermatozoa (similar to Japanese common toad; Bufo japonicus;

Takamune et al. 1991). An underlying assumption explored in Chapters 2 and 3 was that the

absence of protamines–cysteine residues and associated disulphide bonds in African clawed frog

spermatozoa would make the sperm chromatin more susceptible to DNA fragmentation in vitro.

Surprisingly, the rate of SDF did not proceed in a rapid manner consistent with that of other

protamine-deficient species, but rather appeared highly stable after incubation. In fact, this

observation is in direct contrast to that observed in fish (López-Fernández et al. 2009; Gosálvez et

al. 2014) spermatozoa which also possess a chromatin structure lacking disulphide bonds, yet

exhibited an increase in SDF after only a few minutes. Findings presented in Chapter 4 revealed

87

that cane toad sperm chromatin was also highly stable. While the SNBP composition of cane toad

spermatozoa is characterised by arginine-rich protamines (Ausio et al. 2007), the presence of

cysteine residues has not been confirmed. The remarkable chromatin stability reported in both

African clawed frog and cane toad spermatozoa suggests that the high arginine content of the

SNBPs, which is proposed to greatly enhance the compaction of sperm DNA, may play a more

pivotal role in maintaining the sperm DNA integrity of these two species than the capacity for

disulphide bonding (Lewis and Ausio 2002). The findings presented in this thesis have provided

new insight into the mechanisms of chromatin stabilisation that may function in amphibians.

5.4 Activation of motility and sperm DNA fragmentation

In amphibians with external fertilization, the spermatozoa remain inactive in vivo due to the

isotonic environment of the testes, only becoming motile following spawning into hypo-tonic

fresh/pond water with a low osmolality (Browne and Zippel 2007; Kouba et al. 2009). The need to

activate motility in amphibian spermatozoa prior to use with ART imposes high physiological stress

to maintain the osmotic balance of the spermatozoa and is a significant factor impacting the

longevity of both fresh and cryopreserved-thawed spermatozoa (Browne et al. 2015). As the energy

reserves of the spermatozoa are exhausted, osmoregulation fails, resulting in damage to the plasma

membrane and ultimately cell death (Browne et al. 2015). An underlying hypothesis of experiments

in Chapters 3 and 4 was that the progressive loss of DNA integrity would proceed more rapidly in

activated compared to inactivated spermatozoa. In fact, for the first assessment of amphibian sperm

DNA fragmentation dynamics (in African clawed frog; Chapter 3), the incubation period was

restricted to only 60 minutes due to the rapid decline in fertilisation capacity and sperm motility

previously reported for this species following activation (Wolf and Hedrick 1971; Sargent and

Mohun 2005; Mansour et al. 2009). However, the remarkable stability of the sperm chromatin

reported in both the African clawed frog (Chapter 3) and cane toad (Chapter 4), occurred

irrespective of whether the spermatozoa were in the inactivated or activated state. To date, only two

other dynamic studies of SDF have been conducted in externally fertilising species requiring

osmotically-induced activation of sperm motility (López-Fernández et al. 2009; Gosálvez et al.

2014); these studies reported a rapid increase in SDF within minutes of sperm activation. From the

findings presented in this thesis it is possible to conclude that osmotic stress associated with the

activation of sperm motility does not appear to contribute significantly to DNA damage in

amphibian spermatozoa.

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5.5 Effect of cryopreservation on the DNA integrity of amphibian spermatozoa

Despite many refinements in methodology and successful protocols established for a

number of amphibian species (Beesley et al. 1998; Browne et al. 1998; 2002ab; Michael and Jones

2004; Sargent and Mohun 2005), the procedure of sperm cryopreservation is not without adverse

effects. Dilution of the seminal plasma with cryoprotectants in preparation for cryopreservation

predisposes spermatozoa to oxidative and osmotic stress and can potentially contribute to the

incidence of spermatozoa with fragmented DNA post-thaw. Sperm suspensions examined in

Chapters 3 and 4 were cryopreserved according to species-specific optimal protocols developed for

the African clawed frog (Sargent and Mohun 2005; Noble and Jafkins 2010) and cane toad (Browne

et al. 1998). These investigations allowed comparisons between the basal level of SDF in fresh

amphibian spermatozoa and the post-thaw integrity of cryopreserved spermatozoa. Additionally,

Chapter 3 examined the post-thaw SDF of African clawed frog spermatozoa frozen at in a

conventional freezer without cryoprotectant. In this study, an IVF procedure was also used to assess

the extent to which cryopreservation was correlated with a decline in fertilisation rate.

Not surprisingly, there was little difference between values of SDF obtained at the onset of

incubation for fresh and frozen-thawed samples for both the African clawed frog and cane toad.

However, there was evidence of individual variation; for example, the spermatozoa of Male 3

(Chapter 3) and Male 10 (Chapter 4) were particularly resistant to cryopreservation-induced DNA

damage compared to other males. Furthermore, freezing African clawed frog spermatozoa at -20 °C

without any additional cryoprotectant did not have an appreciable effect in terms of DNA

fragmentation. These findings suggest that it is possible to use spermatozoa frozen in this

conventional manner to obtain an index of SDF comparable to fresh sperm samples. Not only did

the sperm chromatin structure in both species appear unaltered by the cryopreservation process

immediately upon thawing, but also appeared highly resistant to the stressors associated with

incubation ex vivo. Although cryopreserved spermatozoa in both species showed a significant

increase in SDF after incubation, the proportion of spermatozoa with SDF was still low compared to

results reported in some other species (López-Fernández et al. 2008a; Imrat et al. 2012). Rather, the

post-thaw DNA fragmentation dynamics reported in this thesis for African clawed frog and cane

toad spermatozoa most closely resembled observations in the koala (Johnston et al. 2012b) where

the spermatozoa also exhibited a marked resistance to cryopreservation-induced DNA damage and

only low levels of SDF were reported, even after incubation for 48 hours. Only one other study has

examined the incidence of cryopreservation-induced chromatin damage in amphibian spermatozoa

89

(in Western clawed frog; Xenopus tropicalis; Morrow et al. 2014; poster presented at the World

Congress of Reproductive Biology); although this study reported a slightly higher SDF following

cryopreservation, it is unclear whether these results were obtained immediately after thawing or

following a period of incubation, making inferences about the possible causes of this variability

difficult. In Chapter 3, cryopreserved-thawed African clawed frog spermatozoa yielded a

significantly lower fertilisation rate compared to fresh spermatozoa, despite exhibiting a similar

level of SDF at the time of IVF. Morrow et al. (2014) also reported a higher incidence of

developmental defects in embryos produced using cryopreserved compared to fresh spermatozoa.

These findings suggest that DNA damage was only a minor component of cryoinjury in African

clawed frog spermatozoa and that the impaired fertilisation capacity was perhaps more likely

related to poor recovery of motility, viability and plasma membrane integrity. This prompted the

analysis of additional parameters of sperm quality in cane toad spermatozoa in Chapter 4.

5.6 Effect of cryopreservation on sperm motility and viability

In an attempt to provide further insight into cryoinjury of the sperm chromatin and the

resulting effect on the potential reproductive output of ART, the investigation presented in Chapter

4 aimed to correlate the dynamic loss of DNA integrity in fresh and cryopreserved cane toad

spermatozoa with other important parameters of sperm quality. Spermatozoa were classified as

either motile or immotile. Motile spermatozoa included all spermatozoa showing either progressive

or non-progressive motility. Sperm viability was assessed using a dual emission technique that

incorporated the nucleic stains SYBR-14 and propidium iodide; SYBR-14 permeated intact plasma

membranes and produced a green fluorescence and propidium iodide permeated damaged plasma

membranes and produced a red fluorescence. It is well documented that cryopreservation impairs

motility, viability, plasma membrane integrity and fertilisation rate in amphibian spermatozoa

(Sargent and Mohun 2005; Mansour et al. 2009). Accordingly, the results presented in Chapter 4

clearly demonstrated a very poor recovery of motility and viability in cane toad spermatozoa

following cryopreservation. Not surprisingly, spermatozoa frozen without cryoprotectant showed no

motility after activation and a marked increase in the incidence of abnormal nuclear morphology

(dead-swollen). Chromatin swelling or relaxation has also been noted in membrane-compromised

spermatozoa of the koala (Johnston et al. 2006) and common wombat (Vombatus ursinus; Johnston

et al. 2006) following thawing and may be associated with the presence of ALS. It is possible that

the incidence of dead-swollen spermatozoa observed in the cane toad coincided with the transition

from ASM-1 to ASM-2 described in the African clawed frog in Chapter 2. It is also worth noting

90

that the addition of antibiotic to the sperm extenders may have affected the percentage of sperm

motility observed before and after freezing. There is some evidence in amphibians (Germano et al.

2013; Silla et al. 2014), that supplementation of sperm extenders with broad-spectrum antibiotics

such as penicillin, streptomycin and gentamicin can negatively impact sperm function in terms of

motility and viability. While it is possible the values presented in Chapter 4 may not accurately

reflect sperm motility and viability in the cane toad, the relative change in these parameters

following freezing and thawing is nonetheless interesting.

5.7 Limitations of the thesis

An initial hypothesis of the thesis was that DNA fragmentation in amphibians would

proceed in a rapid manner consistent with that of other externally fertilising species that require

osmotically-induced activation. This assumption was based on evidence provided by the only two

dynamic studies of DNA fragmentation in lower vertebrate species (López-Fernández et al. 2009;

Gosálvez et al. 2014), where a significant increase in SDF occurred within minutes of sperm

activation. In these species, the absence of protamines-cysteine residues and associated disulphide

bonds was thought to make the sperm chromatin more susceptible to DNA fragmentation in vitro, a

feature also common to African clawed frog SNBPs. The incubation period for experiments

presented in Chapter 3 was initially restricted to only 60 minutes due to the reported rapid decline in

fertilisation capacity and motility for African clawed frog spermatozoa following activation (Wolf

and Hedrick 1971; Sargent and Mohun 2005). Sargent and Mohun (2005) reported a half-life of

sperm motility ranging from less than two minutes to 21 minutes in the African clawed frog and

similar results have been observed in cryopreserved-thawed spermatozoa (Mansour et al. 2009).

Surprisingly, the sperm chromatin of the African clawed frog appeared highly stable and very little

change in SDF was recorded after incubation. This posed a significant limitation with respect to the

analysis of sperm DNA fragmentation dynamics in this species.

For experiments presented in Chapter 4 using cane toad spermatozoa, the incubation period

was increased to 72 hours. When amphibian sperm suspensions are collected via testes extraction

and maceration, bacterial contamination occurs from the surrounding fluids and tissues and is likely

to become exacerbated over time (Silla et al. 2014). Excessive bacterial growth can generate

harmful metabolic by-products such as ROS which in turn leads to the onset of oxidative stress that

can cause DNA strand breaks (Martínez-Páramo et al. 2012; Said et al. 2012). For this reason, the

extender and cryoprotectant solutions were supplemented with gentamicin to give a final

91

concentration equivalent to 100 µg/mL-1. This dose was based on the results of Viveiros et al.

(2010) who reported a decrease in bacterial growth and improved longevity in chilled freshwater

fish (Brycon orbignyanus) semen. Despite this, significant bacterial contamination was still

observed in the sperm suspensions of the current study after incubation for 24 hours. As a

consequence, only data up to T6 could be considered for analysis as the bacteria observed at T24

would likely have accelerated the production of ROS in the sperm samples and in turn falsely

inflated the level of SDF; hence, while a progressive loss of motility and viability was recorded for

cane toad spermatozoa in Chapter 4, the dynamics of SDF were still not able to be fully explored,

particularly for spermatozoa examined fresh and following cryopreservation in DMSO. Although

the information presented in this thesis provided valuable insight into the behavior of the sperm

chromatin in response to cryoinjury, the restricted incubation periods made predictions about the

ultimate longevity of amphibian sperm chromatin problematic.

5.8 Summary of new and significant findings and recommendations for future research

This thesis has led to an improved understanding of amphibian sperm chromatin structure

and the relationship between cryoinjury and sperm quality.

New and significant findings include:

1. Development of the Xenopus-Halomax® assay to identify DNA damage in both African

clawed frog and cane toad spermatozoa

2. Evidence that SSB and DSB occurred simultaneously in African clawed frog spermatozoa

3. Extensive SSB, interpreted as alkali-labile sites, were a constitutive feature of African

clawed frog sperm chromatin

4. Double-stranded DNA breaks represent true DNA damage in amphibian spermatozoa

5. SDF in amphibian spermatozoa does not progress at a rate consistent with other lower

vertebrates studied to date

6. Freezing African clawed frog spermatozoa at -20 °C can be used as an alternative to

cryopreservation to obtain an index of SDF comparable to fresh spermatozoa

7. The incidence of chromatin swelling in amphibian spermatozoa is not related to SDF

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Recommendations for future research include:

1. Investigation into species-specific toxicity of amphibian spermatozoa in response to

antibiotics and identification of optimal concentrations to prevent contaminate growth

2. Further dynamic studies of SDF in amphibian spermatozoa employing longer incubation

periods will reveal the maximum longevity of amphibian sperm chromatin

3. Further investigation into the relationship between SDF in amphibian spermatozoa and the

resulting impact on fertilisation rate and embryo development/normality

93

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