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Genome integrity maintenance during spermatogonial development

Zheng, Y.

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Yi Zheng

Genome integrity maintenance during spermatogonial development PhD Thesis, University of Amsterdam, The Netherlands © Yi Zheng 2018, Amsterdam All rights reserved. No parts of this dissertation may be reproduced, stored in a retrieval system of any nature, or transmitted in any form or by any means without written permission from the author. This thesis describes research performed in the Reproductive Biology Laboratory of the Center for Reproductive Medicine, Academic Medical Center, University of Amsterdam, The Netherlands. ISBN: 978-94-6332-307-9 Cover: SMC5/6 molecule by Dideke Emma Verver Printing: GVO drukkers & vormgevers B.V.


ACADEMISCH PROEFSCHRIFT

ter verkrijging van de graad van doctor

aan de Universiteit van Amsterdam

op gezag van de Rector Magnificus

prof. dr. ir. K.I.J. Maex

ten overstaan van een door het College voor Promoties ingestelde commissie,

in het openbaar te verdedigen in de Agnietenkapel

op donderdag 15 februari 2018, te 14.00 uur

door

Yi Zheng geboren te Sichuan, China

Promotiecommissie Promotor: Prof. dr. S. Repping AMC-UvA

Co-promotor: Dr. G. Hamer AMC-UvA

Overige leden: Dr. ir. W.M. Baarends Erasmus Universiteit Rotterdam

Prof. dr. N. Zelcer AMC-UvA

Prof. dr. C.J.F. van Noorden AMC-UvA

Dr. N.A.P. Franken AMC-UvA

Prof. dr. D.G. de Rooij Universiteit Utrecht

Faculteit der Geneeskunde

Table of contents

Chapte 1 7

General introduction and outline of the thesis

Chapter 2 21

Non-SMC element 2 (NSMCE2) of the SMC5/6 complex helps to resolve topological stress Verver DE#, Zheng Y#, Speijer D, Hoebe R, Dekker HL, Repping S, Stap J, Hamer G #equal contribution International Journal of Molecular Sciences. 2016 Oct 26;17(11). pii: E1782

Chapter 3 49

Trivial role for NSMCE2 during in vitro proliferation and differentiation of male germline

stem cells Zheng Y, Jongejan A, Mulder CL, Mastenbroek S, Repping S, Wang Y, Li J, Hamer G

Reproduction. 2017 Sep;154(3):81-95

Chapter 4 77 On the increasing sensitivity of differentiating spermatogonia to DNA damage Zheng Y, Jongejan A, Mulder CL, van Daalen SKM, Mastenbroek S, Hwang G, Jordan P, Repping S,

Hamer G

Submitted

Chapter 5 107

Spermatogonial stem cell autotransplantation and germline genomic editing: a future cure for spermatogenic failure and prevention of transmission of genomic diseases Mulder CL#, Zheng Y#, Jan SZ, Struijk RB, Repping S, Hamer G*, van Pelt AM #equal contribution, *corresponding author

Human Reproduction Update. 2016 Sep;22(5):561-73

Chapter 6 139

General discussion and implications for future research

Chapter 7 155

Summary Samenvatting

Acknowledgements 160

PhD portfolio 162 About the author 163

List of publications 164

Chapter 1

General introduction and outline of the thesis

8 Chapter 1

Background Spermatogenic failure

An estimated 10-15% of couples suffer from subfertility [1, 2], defined as the inability to

conceive after one year of unprotected intercourse [3, 4]. Although the most important factor

that affects human fertility is female age, in about half of these couples reduced semen

quality is commonly observed [3, 5]. Reduced semen quality can be characterized by low

sperm counts (oligozoospermia), low sperm motility (asthenozoospermia), low number of

morphologically normal sperm (teratozoospermia) or the most extreme clinical presentation-

a complete absence of sperm in the semen (azoospermia) [6]. Azoospermia can be

subdivided into obstructive and non-obstructive azoospermia [2]. In the case of obstructive

azoospermia, the process of spermatogenesis is most often not affected, but the

spermatozoa cannot reach the semen due to a physical obstruction. In the case of non-

obstructive azoospermia, the lack of sperm in the semen is caused by severely decreased or

absent sperm production in the testis, often referred to as spermatogenic failure. Despite the

clinical importance, very little is known about the etiology of spermatogenic failure. There are

only a few established causes for spermatogenic failure, including DNA damage caused by

chemo- or radiotherapy [7], structural or numerical chromosomal abnormalities [5] and Y-

chromosome deletions [8]. Nonetheless, the etiology of spermatogenic failure remains

unknown in most cases. It is presumed that genetic mutations lie at the base of many cases

of spermatogenic failure [9, 10]. Yet, no direct treatment options for spermatogenic failure are

currently available to allow these men to achieve genetic parenthood. The only option to date

is the use of testicular sperm extraction (TESE) in combination with intra-cytoplasmic sperm

injection (ICSI). The drawback is however that the chance of finding spermatozoa upon

TESE in men with non-obstructive azoospermia is roughly 50% and that ICSI implies ovarian

hyperstimulation of the unaffected female partner as well as fertilization and culture of the

resulting embryos in vitro. If indeed spermatogenic failure is genetic in origin, this would

require a precisely patient-specific targeted therapeutic approach, or germline genome

modification to restore the genome into its original ‘fertile’ state. This is currently not yet

feasible.

Spermatogenesis and spermatogonial stem cells (SSCs) Spermatogenesis is an intricate developmental process ultimately leading to the

continuous production of spermatozoa. The whole process comprises three consecutive

developmental stages: the spermatogonial stage (mitotic proliferation and differentiation), the

spermatocyte stage (meiosis) and the spermatid stage (spermiogenesis) [11]. Specifically,

General introduction 9

spermatogenesis initiates from type A spermatogonia that undergo multiple mitotic divisions

and then differentiate into intermediate and type B spermatogonia. Type B spermatogonia

will then divide to form pre-leptotene spermatocytes that replicate their DNA and enter

meiosis. The spermatocytes will subsequently undergo two consecutive meiotic divisions

(meiosis I and II) to generate round spermatids which then further develop into elongating

spermatids and eventually mature sperm.

The type A spermatogonia can be divided into undifferentiated and differentiating

spermatogonia. The undifferentiated spermatogonia proliferate freely and maintain

spermatogonial density in the testis. In contrast, the differentiating spermatogonia are

irreversibly committed towards meiosis and their divisions are strictly regulated. An important

subset of the undifferentiated spermatogonia are the spermatogonial stem cells (SSCs).

These cells can be defined by their ability to generate and maintain donor-derived

spermatogenesis when transplanted into infertile recipient testes [12]. To maintain lifelong

male fertility, a perfect balance between SSC self-renewal and differentiation is essential.

Too much self-renewal may lead to tumor-like germ cell clusters, while excessive

differentiation will lead to germ cell depletion [13]. Despite the apparent importance of this

balance, knowledge regarding the molecular mechanisms underlying SSC self-renewal and

differentiation remains limited [11].

The spermatogonial response to DNA damage DNA damage, for instance caused by irradiation or chemotherapy, often results in germ

cell apoptosis. Many cancer patients undergoing chemo- or radiotherapy are therefore

confronted with reduced fertility [14-16]. Furthermore, DNA damage in germ cells that is not

correctly repaired can lead to genetic mutations or chromosomal aberrations that can be

transmitted to the offspring. For this reason it is thought that germ cells hold a unique

response to DNA damage. Indeed, they are generally much more prone to undergo

apoptosis in response to DNA damage than somatic cells [17, 18]. Even among the different

types of spermatogonia differences in radiosensitivity exist. Differentiating spermatogonia are

more radiosensitive and inclined to undergo apoptosis in response to irradiation than the

undifferentiated spermatogonial population [19]. Even between the undifferentiated

spermatogonia differences exist, with the self-renewing SSCs being the most resistant to

DNA damage [20-22]. It seems that, while differentiating spermatogonia with DNA damage

are readily eliminated, preservation of SSCs, and thus long-term male fertility, to some extent

prevails over the risk of mutated progeny. The mechanisms that determine these differential

responses of spermatogonial subtypes to DNA lesions remain largely unknown.

10 Chapter 1

Studies aimed at investigating the role of specific genes in the DNA damage response

of SSCs are hampered by the fact that many of these genes are embryonically lethal and

thus conventional knockout (KO) strategies are unlikely to work. In addition, reliable

conditional KO systems are not available for the earliest stages of spermatogenesis due to

the lack of suitable promoters.

Chromatin dynamics and the structural maintenance of chromosomes (SMC) 5/6 complex

Spermatogenesis, including spermatogonial differentiation, is associated with a

continuous and drastic transformation of chromatin structure and function. Failure to maintain

correct spatio-temporal organization of chromatin can lead to genomic instability, which often

results in germ cell apoptosis or, when all spermatogenic checkpoints fail, transmission of the

genomic abnormalities to the offspring [11, 23]. Thus, regulation of chromatin composition

and function and maintenance of genome integrity are of paramount importance to the

progression of spermatogenesis and safe reproduction.

Genomic integrity maintenance and other chromatin-based processes, e.g. DNA

replication, transcription and cellular differentiation, are for a large part orchestrated by SMC

protein complexes: SMC1/3 (cohesin), SMC2/4 (condensin) and SMC5/6. Of these the

SMC5/6 complex is composed of SMC5 and SMC6 and several non-SMC elements

(NSMCE1-4 in mammals, Figure 1). Together, these components can form a ring-like

structure able to hold two double-stranded DNA molecules together [24]. Of the NSMCEs,

NSMCE2 specifically associates with SMC5, where it displays an E3 small ubiquitin-related

modifier (SUMO) ligase activity that is involved in DNA damage repair [25-27].

Figure 1: The structure of the SMC5/6 protein complex. SMC5 and

SMC6 proteins form a ring-like structure together with several NSMCEs.

NSMCE2 specifically associates with SMC5. Image by Dideke Emma

Verver [24].


In the mouse and human testis, SMC5/6 has been described to be involved in several

meiotic processes including chromosome segregation, homologous chromosome synapsis

and meiotic sex chromosome inactivation [28, 29]. Interestingly, protein staining for SMC6

has recently been found to coincide with spermatogonial differentiation [29]. However, the

specific function of SMC5/6 in differentiating spermatogonia is currently not known. It may be

present in germ cells to prevent dangerous and error-prone recombination events in highly

repetitive genomic sequences such as the regions that surround centromeres [28-33].

Alternatively, the SMC5/6 complex may be involved in the maintenance of replication fork

stability and the prevention of replication-induced DNA damage [34-36]. A recent paper also

showed NSMCE2 to be expressed in mouse male germ cells from spermatogonia to round

spermatids [37]. Nevertheless, whether the SMC5/6 complex plays a role in spermatogonial

response to DNA damage has not been further investigated.

SSC culture SSCs account for only 0.02-0.03% of all germ cells [38]. Given the sparsity of SSCs in

the testis, SSC culture has become an established tool to expand and study this relatively

rare cell population in vitro. A breakthrough in SSC culture was accomplished in 2003, when

Shinohara’s group first reported a long-term culture system for mouse SSCs [39]. In this

culture system, primary undifferentiated spermatogonia, termed male germline stem (GS)

cells, are able to propagate in vitro for years without losing SSC properties [40]. Since then,

successful long-term SSC cultures from rats [41], hamsters [42], rabbits [43] and tree shrew

[44] have been achieved. SSCs from these species do not only propagate in vitro for a long

time but can also be used to generate transgenic offspring [45]. Based on the culture system

for rodent SSCs, we were the first to establish a culture system for adult [46] and prepubertal

human SSCs [47]. However, current cultures of human SSCs are not optimal and remain to

be improved. Most importantly, the developmental capacity of cultured human SSCs, i.e. the

capacity to initiate and maintain spermatogenesis, still needs to be demonstrated.

SSC transplantation In 1994, Brinster and colleagues developed a method of SSC transplantation by

injecting donor germ cells into the efferent duct or rete testis of a recipient testis [12, 48].

After transplantation, only SSCs are assumed to relocate to the stem cell niche at the basal

membrane of seminiferous tubules, from where they can reinitiate and maintain

spermatogenesis. It is well acknowledged that only SSCs have the capacity of producing

12 Chapter 1

donor-derived mature spermatozoa, whilst other more advanced germ cells will most likely

degenerate or further differentiate and disappear after a certain period of time.

Transplantation has therefore become the golden assay to determine the stem cell capacity

of spermatogonia. Later, this technique has successfully been used in rodent species other

than mice (e.g. rats), in domestic animals (e.g. boars, bulls, goats, sheep, cats, dogs) and

even non-human primates [49]. In 2012, Hermann et al. [50] first reported that autologous or

allogeneic transplantation of SSCs into the testis of recipient rhesus monkeys could

reestablish spermatogenesis and produce donor-derived sperm that were functional and able

to develop to embryos after fertilization.

Human SSC transplantation is currently regarded as a future treatment option for

prepubertal boys who have lost their germ cells due to the gonadotoxic side-effects of cancer

treatment such as chemo- or radiotherapy [51]. In order to preserve fertility in these

prepubertal boys, a testicular biopsy can be obtained before the initiation of chemo- or

radiotherapy. This testicular biopsy, or cultured and expanded cells from the biopsy, can be

cryopreserved. When the patient is cured from cancer and reaches adulthood, the cells can

be auto-transplanted into the testis where they will then hopefully initiate and maintain

spermatogenesis and restore fertility. This method is currently still under development in our

laboratory and the first auto-transplantation is expected to be conducted within five years

from now on. Because the efficiency of SSC transplantation highly relies on the number of

transplanted SSCs [52], development of culture systems to expand human SSCs is crucial.

In addition, the (epi)genomic stability of the cultured SSCs, the efficiency of the

transplantation technique in humans, and the safety of the patients and offspring need to be

guaranteed before clinical application of SSC transplantation can be considered [53].

Genome modification of SSCs The establishment of stable long-term culture systems to expand SSCs in vitro lays the

groundwork for the possibility to genetically modify SSCs. SSCs in culture can be subjected

to genetic manipulation, using similar protocols as developed for embryonic stem (ES) cells.

More specifically, SSCs can be transfected with plasmid vectors by prevailing methods such

as calcium phosphate precipitation, lipofection or electroporation, albeit very inefficiently [54].

To improve the efficiency, SSCs are typically transduced with viral-based vectors, such as

adenovirus, adeno-associated virus (AAV), retrovirus or lentivirus. Of these, lentiviral

transduction is advantageous in that transgenes can integrate into the genome and will thus

be stably expressed in SSCs [55]. In this way, genetically modified animal models, including

mice, rats and recently tree shrew, have been produced [44, 56, 57]. Nevertheless, as


retrovirus and lentivirus can integrate into the genome of the recipient, they are not suitable

for clinical use in the human.

Traditionally, genome editing has been achieved via homologous recombination in ES

cells, which is inefficient and time-consuming. Over the last decade, novel tools using

engineered nucleases to generate site-specific double-strand breaks (DSBs) have

revolutionized the field of genome editing. Of these, the novel CRISPR-Cas9 technique

(Figure 2) is unprecedentedly simple and efficient. The emergence of CRISPR-Cas9 greatly

facilitates genetic manipulation of SSCs, and to date both targeted gene KO and gene

correction have been achieved in rodent SSCs [58-60]. In the future, the combination of SSC

culture and CRISPR-Cas9 is expected to enable germline modification of all mammalian

species including humans.

Figure 2: A schematic illustration of the type II SpCas9 system. This most

commonly used CRISPR-Cas9 system is

composed of a single-guide RNA (sgRNA)

which contains a specific 20-nt sequence to

bind to the complementary genomic DNA

(gDNA) sequence. The 20-bp gDNA

sequence must precede 5’-NGG, the

protospacer-adjacent motif (PAM). In

conjunction with the endonuclease Cas9, a

DSB can occur at ~3-bp upstream of the

PAM. The triggered DSB is typically repaired

by non-homologous end joining (NHEJ),

which is error-prone and causes

insertions/deletions (indels) flanking the DSB site, possibly resulting in a frame-shift and therefore a

pre-mature stop codon. Alternatively, when a homologous DNA sequence is provided as a repair

template, homology-directed repair (HDR) can occur and precise genomic editing can be achieved.

Since SSCs are capable of transmitting genetic information from one generation to the

next, they are an ideal target for genetic manipulation to produce transgenic animal models

for biomedical research and animal production. The use of genetically modified SSCs could

also form a direct treatment option for men with spermatogenic failure in whom the disease is

caused by a genetic abnormality. In addition, the potential use of genome modification in

human SSCs opens up the possibility to prevent the paternal transmission of any genetic

14 Chapter 1

abnormality to offspring. Genome modification of human SSCs is in this way in essence a

potential alternative for existing methods aimed at preventing the transmission of genetic

abnormalities to offspring such as prenatal and preimplantation genetic diagnosis (PGD) and

an alternative for the possible future application of genome editing of human embryos. The

potential clinical application of germline genome editing, be it in SSCs or in embryos, is a

controversial and highly debated subject [61, 62].

Aim and outline of the thesis The specific aim of this thesis was to unravel the mechanisms that determine and

regulate the dynamic response to DNA damage during spermatogonial development, with a

specific focus on the role of the SMC5/6 complex. The development of SSC culture, together

with the development of CRISPR-Cas9 as described above, now for the first time open up

the possibility to study this using spermatogonia-specific genome modification. In the current

thesis, we combined an established culture system for spermatogonial proliferation and

differentiation with the CRISPR-Cas9 system to knock out genes of interest in the response

to DNA damage. Furthermore, we additionally addressed the broad and potentially large

clinical prospects and implications of using CRISPR-Cas9 to treat spermatogenic failure or to

prevent the transmission of genetic abnormalities to human offspring by transplantation of

genetically modified human SSCs.

In Chapter 2 we describe the use of CRISPR-Cas9 and a human osteosarcoma cell

line (U2OS) to investigate the molecular mechanisms by which the SMC5/6 complex

functions in genomic integrity maintenance.

In Chapter 3 we report an optimized protocol to generate genetically modified mouse

male germline stem cell lines using CRISPR-Cas9. Using this protocol, we generated a male

germline stem cell line devoid of NSMCE2, a subunit of the SMC5/6 complex, and then

interrogated the role of NSMCE2 in spermatogonial proliferation and differentiation.

In Chapter 4 we analyze the transcriptomes of irradiated and non-irradiated

undifferentiated and differentiating mouse male germline stem cells to gain insights into the

differential DNA damage responses of undifferentiated and differentiating spermatogonia.

In Chapter 5 we review the state of the art with respect to SSC transplantation and

genomic editing using CRISPR-Cas9, followed by envisioning the clinical prospects of SSC

transplantation, with or without genomic editing, to restore male fertility or prevent

transmission of genomic disorders.


In Chapter 6 we give an overview of the findings presented in this thesis and discuss

the implications of our results for future research.

In Chapter 7 we summarize the results presented in this thesis.

16 Chapter 1

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Chapter 2

Non-SMC element 2 (NSMCE2) of the SMC5/6 complex helps to resolve topological stress

Dideke E. Verver#

Yi Zheng#

Dave Speijer

Ron Hoebe

Henk L. Dekker

Sjoerd Repping

Jan Stap

Geert Hamer

#equal contribution

International Journal of Molecular Sciences

2016 Oct 26;17(11). pii: E1782

22 Chapter 2

Abstract The structural maintenance of chromosomes (SMC) protein complexes shape and

regulate the structure and dynamics of chromatin, thereby controlling many chromosome-

based processes such as cell cycle progression, differentiation, gene transcription and DNA

repair. The SMC5/6 complex is previously described to promote DNA double-strand breaks

(DSBs) repair by sister chromatid recombination, and found to be essential for resolving

recombination intermediates during meiotic recombination. Moreover, in budding yeast,

SMC5/6 provides structural organization and topological stress relief during replication in

mitotically dividing cells. Despite the essential nature of the SMC5/6 complex, the versatile

mechanisms by which SMC5/6 functions and its molecular regulation in mammalian cells

remain poorly understood. By using a human osteosarcoma cell line (U2OS), we show that

after the CRISPR-Cas9-mediated removal of the SMC5/6 subunit NSMCE2, treatment with

the topoisomerase II inhibitor etoposide triggered an increased sensitivity in cells lacking

NSMCE2. In contrast, NSMCE2 appeared not essential for a proper DNA damage response

or cell survival after DSB induction by ionizing irradiation (IR). Interestingly, by way of

immunoprecipitations (IPs) and mass spectrometry, we found that the SMC5/6 complex

physically interacts with the DNA topoisomerase II α (TOP2A). We therefore propose that the

SMC5/6 complex functions in resolving TOP2A-mediated DSB-repair intermediates

generated during replication.

Keywords Structural Maintenance of Chromosomes 5/6 complex (SMC5/6); Non-SMC Element 2

(NSMCE2); Topoisomerase II α (TOP2A); DNA double-strand breaks (DSBs); Ionizing

Radiation (IR); CRISPR-Cas9

NSMCE2 helps to resolve topological stress 23

Introduction

The structural maintenance of chromosome (SMC) protein complexes shape and

determine chromatin structure and function and are therefore implicated in many, if not all,

fundamental chromosome-based processes. The three SMC protein complexes, cohesin,

condensin and SMC5/6, all consist of two SMC subunits and several non-SMC elements, the

NSMCEs. The resulting ring-like complexes possess the capacity to hold two DNA double-

strands together, and are therefore able to physically shape the DNA in specific chromatin

structures [1, 2]. By doing so, SMC complexes control chromosome segregation, DNA repair,

transcription and replication, among other processes [1, 3-5]. Of the three SMC complexes,

the SMC5/6 complex has been most directly and exclusively described to be involved in DNA

damage repair and genomic integrity maintenance [6-8].

In mammals, SMC6 is highly expressed in the testis [9, 10] and we have recently found

SMC6 to be involved in crucial processes during mouse and human spermatogenesis,

including spermatogonial differentiation and meiosis [9, 10]. In various organisms, ranging

from yeast to humans, SMC5/6 is involved in numerous meiotic processes [11] such as

chromosome segregation [9, 10, 12, 13], homologous chromosome synapsis [9, 12-16] and

meiotic sex chromosome inactivation [10, 13]. Co-localization studies have suggested that

SMC5/6 prevents dangerous and error-prone homologous recombination (HR) in highly

repetitive, densely packed DNA regions such as the rDNA and pericentromeric

heterochromatin [10, 13, 15-18]. SMC5/6 seems to be involved in double-strand break (DSB)

repair as it is enriched at DSB sites in budding yeast and C. elegans [12, 14, 16]; it localizes

side by side with RAD51 in budding yeast and humans [9, 12, 16] and its deletion results in

an increase in RAD51 foci and chromosome fragmentation in C. elegans [14]. Furthermore,

Smc5/6 has been found to play a role in the resolution of meiotic recombination

intermediates and mutations of Smc5, Smc6 or the SUMO ligase domain of Nse2 lead to the

accumulation of toxic joint molecules in yeast and C. elegans [12, 15, 16, 19-22].

In budding and fission yeast the Smc5/6 complex is essential for the maintenance of

replication fork stability, the prevention of joint molecules and the resolution of such joint

molecules that would otherwise lead to mitotic failure (reviewed in [23-25]). In mice, ablation

of SMC6 results in embryonic lethality, whilst a mutation in its ATP hydrolysis motif only

generates a mild phenotype [26]. NSMCE2 has also been shown to be essential for mouse

development and it can suppress cancer and aging by limiting recombination and facilitating

chromosome segregation [27]. In line with these studies, a recent paper describes that

24 Chapter 2

depletion of SMC5 in mouse embryonic stem cells led to accumulation of cells in G2 and

subsequent mitotic failure and apoptosis [28].

From this increasing amount of data, it has become overwhelmingly clear that SMC5/6

is essential for maintaining genomic integrity by a variety of means. However, the exact roles

of the SMC5/6 complex in mammalian especially human cells remain poorly understood. By

using a commonly used human osteosarcoma cell line (U2OS), we extended our knowledge

regarding the roles of SMC5/6 in human genome integrity maintenance.

Materials and methods Cells and culture

U2OS cells were cultured at 37 °C and 10% CO2 in Dulbecco’s modified Eagle’s

medium (DMEM; (high glucose, pyruvate, L-glutamine); Thermo Fisher Scientific, Waltham,

MA, USA) supplemented with 10% Fetal Calf Serum (FCS), penicillin (100 U/mL) and

streptomycin (100 U/mL).

Design of single-guide RNAs (sgRNAs) and construction of the CRISPR-Cas9 plasmids

An online CRISPR-Cas9 Design Tool provided by Zhang’s lab (http://crispr.mit.edu)

was exploited to identify the targeting sequences. The 20-nt targeting sequences preceding

5′-NGG (the protospacer-adjacent motif, PAM), locating in early and conserved coding exons

among transcript variants, and with least predicted off-target sites in human genome were

selected. To target SMC6 and NSMCE2 in U2OS cells, two sgRNAs were designed for each

gene: 5′-GGTGACGAAGACGAATGTAA-3′ (in exon 3) and 5′-ATGCTTGGACCTTTTAAGTT-

3′ (in exon 4) for SMC6, 5′-TTCCAAGCCTGTATCAACTC -3′ and 5′-

AGCCTGTATCAACTCTGGTA-3′ (both in exon 3) for NSMCE2. The corresponding sense

and antisense strands of oligos were purchased from Sigma-Aldrich (St. Louis, MO, USA).

The CRISPR plasmids pSpCas9(BB)-2A-GFP (pX458) were obtained from Addgene

(Addgene plasmid 48138). The pX458 plasmids were digested with FastDigest BbsI

(Fermentas, Waltham, MA, USA). The oligos were then annealed and cloned into digested

pX458 according to the protocol described by Ran et al. [29].

Transfection of U2OS cells with CRISPR plasmids The constructed pX458 plasmids were transfected into U2OS cells with the 4D-

Nucleofector System (Lonza, Basel, Switzerland). For each nucleofection reaction,

approximately 500 ng plasmids were transfected into 100,000 U2OS cells using program

CM-104 and the SE Cell Line 4D-Nucleofector X Kit S (Lonza), according to the


manufacturer’s instructions. One day after nucleofection, the top 5%-10% GFP+ U2OS cells

were separated by FACS with a BD FACSAria cell sorter. In order to derive a stable

NSMCE2-deficient cell line from single U2OS cells, FACS was conducted to deposit single

GFP+ cells into 96-well plates (one cell per well).

Surveyor assay for verification of genome editing Genomic DNA of transfected U2OS cells was extracted with QIAamp DNA Mini Kit

(Qiagen, Hilden, Germany), according to the protocol provided by the manufacturer. The

genomic region (417 bp) containing both mutation sites in exon 3 of NSMCE2 was amplified

by PCR with Herculase II fusion polymerase (Agilent Technologies, Santa Clara, CA, USA).

The forward and reverse primers used for PCR were 5′-AATTTCAAGATGCCAGGACGT-3′

and 5′-GGATCTTCAAATCTTTGCCCAT-3′, respectively. PCR products were purified by

QIAquick PCR Purification Kit (Qiagen). Genomic modifications in the amplified region were

then detected with Surveyor Mutation Detection Kit (Integrated DNA Technologies, Coralville,

IA, USA), according to the manufacturer’s instructions. After Surveyor nuclease digestion,

the PCR products were run on a 2% agarose gel with ethidium bromide (EB) for visualization.

The insertion/deletion (indel) occurrence was estimated with the formula described by Ran et

al. [29].

Sequencing analysis of the targeting site in NSMCE2 Genomic DNA was extracted from each single cell-derived colonies, and the region

flanking the targeting site was amplified by PCR with Easy-A High-Fidelity PCR Cloning

Enzyme (Agilent Technologies). The primers used were identical to those for the Surveyor

assay. After purification, the PCR products were cloned into TOPO TA cloning vectors

(Thermo Fisher Scientific). The ligated vectors were transformed into One Shot TOP10

Chemically Competent E. coli (Thermo Fisher Scientific). For each reaction, forty colonies

were randomly picked and subjected to Sanger-sequencing to analyze mutations from all

alleles.

Off-target analysis in the NSMCE2-devoid cell line Genomic DNA was extracted from WT and NSMCE2 null U2OS cells, respectively. To

gain an overview of off-target effects in the established NSMCE2 null cell line, the 10 top-

ranking potential off-target sites provided by the online CRISPR Design Tool

(http://crispr.mit.edu) were analyzed. The selected sites included those preceding 5′-NAG,

the alternative PAM. The genomic regions flanking each potential off-target sites were

amplified by PCR with Herculase II fusion polymerase (Agilent Technologies). The selected

26 Chapter 2

off-target sites and the corresponding PCR primers are depicted in Table S1. Purified PCR

products were Sanger-sequenced for analysis of off-target effects.

Live cell microscopy Cells were plated in multi-chambered cover-glass slides (Labtek II, Nunc) in a density

of 1000 cells/cm2 and cultured overnight before starting imaging. Both U2OS WT and

NSMCE2 null cells were imaged at the same time, using a IRBE inverted phase contrast

microscope and a N Plan Apo L 40×/0.55 Ph2 objective. Images were captured every 10 min

for a total of 170 h, at 37 °C in an atmosphere containing 10% CO2. Medium was refreshed

under the microscope. Cell cycle duration was determined by calculating the time between

two cell divisions. Generations were aligned, in which we chose the third generation as the

one in which the medium was refreshed, based on the WT cells.

Distribution of cells over the cell cycle phase To determine the distribution of cells over the different cell cycle phases, colcemid

(KaryoMAX Colcemid Solution; Thermo Fisher Scientific) was added to the culture medium

to a final concentration of 0.1 µg/mL and DNA histograms were made after incubation. Cells

were detached by 0.05% trypsin/EDTA (Thermo Fisher Scientific), pelleted in serum-

containing medium and washed in PBS (Phosphate Buffered Saline). Cells were fixed and

stored in 100% EtOH at 4 °C. On the day of FACS analysis, cells were pelleted by

centrifugation and all EtOH was removed. Cells were resuspended in PBS and RNAse A

(final 1 mg/mL; Roche, Basel, Switzerland) was added. After vortexing, propidium iodide (PI;

final 25 µg/mL; Sigma-Aldrich) was added and the cell suspension was thoroughly vortexed

again. Cells were incubated for 15 min at 37 °C, after which the cell suspension was

transferred through a 21 G needle twice, in order to disrupt cell aggregates. DNA content

was analyzed on a LSR Fortessa FACS analyzer (BD Biosciences, San Jose, CA, USA)

using DivaTM acquisition and analysis software. Figures were constructed using FlowJo X

software.

Ionizing irradiation (IR) Exponentially growing cells were exposed to IR emitted by a 137Cs source (95% β-

emission). For immunocytochemistry, cells received 1 Gy of IR. For clonogenic assays, cells

received 0-8 Gy of IR.


Immunocytochemistry (ICC) For ICC, cells were plated on glass coverslides overnight, after which they were fixed in

4% formaldehyde/PBS for 10 min at room temperature (RT). In the case of IR treatment, IR-

treated cells (and their non-IR counterparts) were fixed at varying time points, ranging from 0

min to 6 h post IR. In the case of etoposide treatment, cells were incubated with 3 µM

etoposide for 1 h at 37 °C/10% CO2. Cells were fixed at varying time points after removal of

etoposide, ranging from 0 min to 6 h post etoposide treatment. After fixation, cells were

permeabilized for 10 min at RT in PBST (0.25% Tween-20/PBS). Next, non-specific

adhesion sites were blocked for 45 min at RT in PBST containing 1% bovine serum albumin

(BSA). To visualize SMC6 and γH2AX, slides were incubated for 2 h at RT in primary

antibodies guinea pig (GP) anti-SMC6 (1/200; custom made, peptide:

KRPRQEELEDFDKDGDEDE) and mouse anti-γH2AX (1/10,000; 05-636, Merck Millipore,

Darmstadt, Germany), diluted in 1% BSA/PBST. After incubation in corresponding secondary

antibodies (Goat-anti-GP Alexa488, Goat-anti-Mouse Alexa555, respectively; all diluted

1/1000 in 1% BSA/PBST), slides were washed and counterstained with DAPI and mounted

in Prolong Gold. Between all steps, except blocking and primary antibody incubation, 3 × 5

min washes in PBS were performed.

Widefield fluorescence microscopy images were acquired at RT using a Plan Fluotar

100×/1.30 oil objective on a Leica DM5000B widefield microscope equipped with a Leica

DFC365 FX CCD camera. Images were analyzed using Leica Application Suite Advanced

Fluorescence software. Figures were constructed using Adobe Photoshop CS5 version 12.0.

Confocal images for subsequent quantification were acquired at RT using a Leica TCS SP8

SMD confocal microscope equipped with a 63×/1.40 HC Plan Apo oil CS2 objective (Leica,

Wetzlar, Germany). For excitation of DAPI, a 405 nm UV Diode was used and for excitation

of other fluochromes, the VIS Argon 470–670 nm White Light Laser (WLL) was used.

Fluorescent signal was detected by PMTs and a HyD detector, and acquisition of the image

(stacks) was performed using LAS AF X software.

Quantitative imaging In order to enable identical staining and acquisition conditions for all samples, four time

points were chosen for quantitative imaging, 0 min, 30 min, 3 h and 6 h post IR or etoposide

treatment. Confocal image stacks were acquired using the following settings: resonant scan

= on; galvo flow and bidirectional X = off; line average = 4; acquisition = between lines; field

of view = 792 × 792 (zoom = 5.0); Z-step size = 0.20 µm; stack size = 8 µm total (42 steps).

One pixel = 47 × 47 × 200 nm (XYZ).

28 Chapter 2

Images were deconvolved using Huygens Essential software, with a maximum number

of iterations of 40, and a SNR setting of 12 (green channel) or 10 (red channel). By visual

inspection, cells with at least two clear SMC6 foci were identified for further analysis with

MatLab software. Using MatLab software, we isolated the γH2AX foci with a minimal size of

50 pixels (0.022 µm3) present in the nucleus. Next, the separate SMC6 foci were isolated

(minimal size 10 pixels), and the amount of γH2AX foci overlapping with a SMC6 focus was

determined. Statistical significance was determined by applying the Student’s t-test (two-

tailed, unpaired).

Clonogenic assay

Clonogenic assays were performed as described previously [30]. Four hours after

plating, cells were exposed to 0-8 Gy of IR or incubated for 1 h at 37 °C in 0-30 μM etoposide.

In each experiment, each dose was administered to 2 different cell densities. Experiments

were repeated at least 3 times. Survival capacity was calculated relative to the non-treated

control condition. Statistical significance was determined by applying the Student’s t-test

(one-tailed, paired).

Protein isolation Cells were detached, washed in PBS and pelleted by centrifugation. These cell pellets

were either snapfrozen in liquid nitrogen and stored or lysed directly. Cells were lysed in

RIPA buffer containing 1× PIC and 1× PhosSTOP (Roche) for 1 h on ice. The lysate was

centrifuged (16,000 rcf, 10 min, 4 °C) to clarify the extract.

Western blot analysis Western blot analysis of cell lysates was performed as previously described [10], using

the primary antibodies: SMC6 GP (1/200; custom made), SMC5 (1/1000; A300-236A, Bethyl

Laboratories, Montgomery, TX, USA), NSMCE2 (1/500; NBP1-76263, Novus Biologicals,

Littleton, CO, USA), TOP2A (1/1,000; TG2011-1, TopoGEN, Buena Vista, CO, USA) and β-

actin (1/5000; A1978, Sigma-Aldrich).

Immunoprecipitation (IP) IP was performed on lysed cells, using Dynabeads Protein A (Thermo Fisher Scientific).

Per IP, 1 × 106 cells and 50 µL dynabeads were used. Dynabeads were resuspended in AB

Binding & Washing buffer containing 2 µL anti-SMC6 GP (custom made) or anti-SMC5

(A300-236A, Bethyl Laboratories) antibody and incubated for 30 min with rotation at RT.


Using a magnet, the supernatant was removed, and the beads were washed by

resuspension in AB Binding & Washing buffer. After the removal of the buffer, the beads

were incubated in the cell lysate for 30 min with rotation at RT, after which the supernatant

was collected and the beads were washed. Elution of the precipitated antigen was achieved

after resuspension of the beads in RIPA buffer containing 1× PIC and 1× PhosSTOP,

addition of LDS Sample Buffer and Sample Reducing Agent and heating of the sample for 10

min at 70 °C. In preparation for Western blot analysis, the supernatant was also

supplemented with LDS Sample Buffer and Sample Reducing Agent and heated for 10 min

at 70 °C.

Mass spectrometry

Abcam rabbit anti-SMC6 (ab18039, Abcam, Cambridge, UK) was used for IP followed

by mass spectrometry of the several bands detected by the antibody in U2OS cells, following

the protocol described above. A total of approximately 7 × 106 cells, 100 µL dynabeads and 5

µL SMC6 Abcam antibody were used. All immunoprecipitated material was loaded on a

single lane of a 4%-12% bis-tris gradient gel (Thermo Fisher Scientific). After running, the gel

was washed 3 times 10 min in H2O to remove SDS, and subsequently stained in a colloidal

coomassie solution (PageBlue Protein Staining Solution; 24620, Thermo Fisher Scientific) for

1 h at RT, after which the excess of staining was washed away with H2O. Gels were stored in

1% acetic acid/H2O at 4 °C. Protein bands of interest were excised, alkylated and subjected

to tryptic digestion according to standard protocols. Further mass spectrometry analysis was

performed as described previously [31].

Results CRISPR-Cas9-mediated targeting of the SMC5/6 complex

In order to investigate the role of the SMC5/6 complex during different cellular

processes such as DNA repair, we used the novel CRISPR-Cas9 system to generate cells

lacking a fully functional SMC5/6 complex. U2OS cells were transfected with constructed

CRISPR plasmids (pX458) to target SMC6 or NSMCE2. One day after transfection, GFP+

cells harboring CRISPR plasmids were sorted by fluorescence-activated cell sorting (FACS)

and then cultured for five days (Figure 1A). To verify the occurrence of genome mutations in

the sorted cell fractions, a Surveyor assay was performed based on PCR amplicons of the

genomic DNA region around the targeting sites. Targeting of NSMCE2, but not SMC6,

yielded fragments with the expected sizes after Surveyor nuclease digestion, indicating

successful genome editing. The insertion/deletion (indel) occurrence brought by two

30 Chapter 2

individual single-guide RNAs (sgRNAs) targeting NSMCE2 was 17.2% and 16.6%,

respectively (Figure 1B). To derive a monoclonal knockout cell line, FACS was conducted to

deposit single GFP+ cells into 96-well plates. Single cells were then expanded for one to two

months. Consistent with the results of Surveyor assay, all single cell-derived colonies

appeared wild type for SMC6 after Sanger sequencing. In addition, for NSMCE2, we did not

achieve complete NSMCE2-knockout after propagation. However, one monoclonal cell

population (NSMCE2-1B) showed only one remaining wild type NSMCE2 allele, which was

effectively mutated after a second round of transfection and single cell sorting using the

NSMCE2-1B clone, resulting in the generation of a complete NSMCE2 null cell line

(NSMCE2-1R, Figure 1C). Both Sanger sequencing and Western blot analysis showed the

full null mutations in NSMCE2-1R cells (Figure 1C, D). Subsequently, by sequencing the 10

top-ranking potential off-target sites in the established NSMCE2 null cell line (Table S1), no

off-target alterations were detected.

Figure 1: CRISPR-Cas9-mediated targeting of NSMCE2. (A) Left panel, transfected U2OS cells

(GFP+) under bright field and fluorescence; right panel, FACS enrichment of GFP+ cells. Bar = 50 µm.

(B) Expected cleavage bands (approximately 304 and 113 bp, arrowheads) generated by Surveyor

nuclease digestion. For negative control (−) transfection of the pX458 plasmids without NSMCE2

sgRNA was performed. (C) Sequencing analysis for characterization of the CRISPR-Cas9-induced

frameshift mutations. Red letters represent the 20-nt targeting sequences, while blue letters refer to

the protospacer-adjacent motif (PAM). (D) Western blot analysis of the NSMCE2 protein in the final

NSMCE2 null and WT cells. β-Actin was used as a loading control.


Characterization of NSMCE2 null cells Morphologically, NSMCE2 null cells generally resemble WT cells, although NSMCE2

null cells clearly show more vacuoles, indicating increased cellular stress in the absence of

NSMCE2 (Figure 2A). In addition, time-lapse imaging revealed a significant 1.37-fold

increase in the cell cycle duration of NSMCE2 null cells (Figure 2B). When investigating the

distribution of cells among different cell cycle phases, the DNA histogram of NSMCE2 null

cells showed a recurring increase of approximately 10% in G0-1 phase compared to WT

(Figure 2C). To investigate whether all of the NSMCE2 null cells participate in the cell cycle,

we treated WT and NSMCE2 null cells with the M-phase blocking agent colcemid [32].

Although both WT and NSMCE2 null cells showed a rapid depletion of G0-1 cells after

colcemid treatment (Figure 2D, E), which is in accordance with the rapid cycling nature of

U2OS cells, there were always ~10% more NSMCE2 null cells remaining in G0-1, and even

after 96 h, a clear subpopulation of 16% remained (Figure 2D, E), indicating that these cells

do not participate in the cell cycle. Protein levels of SMC5 and SMC6 were not evidently

affected by the absence of NSMCE2 (Figure 2F).

32 Chapter 2

Figure 2: Analysis of NSMCE2 null cell growth characteristics. (A) Phase contrast images of WT

and NSMCE2 null cells. The latter shows a large number of vacuoles in the cytoplasm (arrow). Bar =

20 µm. (B) Average cell cycle duration of WT and NSMCE2 null cells over multiple generations

observed by live cell imaging. Data are presented as mean ± standard error of mean (SEM), n = 3. (C)

Cell cycle phase distribution analysis of WT and NSMCE2 null cells by DNA histograms shows a 10%

increase NSMCE2 null cells in G0-1. (D) G0-1 phase depletion by colcemid treatment. WT and NSMCE2

null cells were treated with colcemid for 0–96 h. In NSMCE2 null cells, a fraction of cells remained in

G0-1 even after 96 h. (E) Quantification for depletion of G0-1 cells with the time of colcemid treatment.

Data are presented as mean ± SEM, n = 3. (F) Western blot analysis of SMC5 and SMC6 proteins in

WT and NSMCE2 null cells. β-Actin was used as a loading control.

Irradiation-induced SMC6 foci formation occurs independent of NSMCE2 Because NSMCE2 is reported to be essential for the SMC5/6 function in the repair of

DNA DSBs [33-37], we performed immunocytochemical stainings (ICC) for SMC6 on WT and

NSMCE2 null cells at different time points after exposure to 1 Gy of ionizing irradiation (IR)

(Figure 3A). Indeed, both in WT and NSMCE2 null cells, IR induced SMC6 foci that co-

localize with DSBs (marked by γH2AX foci) and that gradually decrease post irradiation. This

expression pattern of SMC6 was similar for both WT and NSMCE2 null cells. After we

generated and applied a MatLab image analysis script that objectively isolates and quantifies

the number of SMC6 and γH2AX foci in each cell, we found no difference in average number

of γH2AX foci per nucleus at the chosen time points, indicating that both cell lines process

DSBs in a similar fashion (Figure 3B). Importantly, not all γH2AX foci were represented by a

SMC6 focus (Figure 3A). We therefore also determined the percentage of γH2AX foci

positive for SMC6. We found that roughly 50% of the DSB sites were positive for SMC6 in

both cell lines, indicating that the accumulation of SMC6 to sites of DSB damage is equally

efficient in WT and NSMCE2 null cells (Figure 3C). More etoposide-induced double-strand break (DSB) formation without NSMCE2

Next we interrogated whether the absence of NSMCE2 would influence the repair of

etoposide-induced DNA damage. The cells were exposed to etoposide, a cytotoxic agent

that acts by forming a complex with the DNA and the topoisomerase II enzyme [38, 39]. In

normal conditions, type II topoisomerase releases superhelical stress and untangles

chromosomes by creating transient DSBs, through which an unbroken DNA helix is

transferred before ligation of the break, thereby averting incidents such as stalled replication

forks or replication-induced joint molecules caused by DNA supercoiling [40]. Because


etoposide prevents re-ligation of the DNA strands after transient DSBs, etoposide treatment

will eventually lead to an increase of permanent DSBs [38, 39]. Indeed, when exposing the

WT and NSMCE2 null cells to etoposide, DSBs (marked by γH2AX) became readily

discernible (Figure 4A). However, in contrast to IR, the number of etoposide-induced DSBs

increased significantly in the absence of NSMCE2 (p < 0.02 after 30 min and p < 0.03 after 3

h, Figure 4B). Similar to IR, not all etoposide-induced DSB foci contained SMC6, and the

percentage of DSB foci containing SMC6 showed no significant difference between WT and

NSMCE2 null cells after etoposide treatment (Figure 4C).

Figure 3: Ionizing irradiation (IR)-induced double-strand break (DSB) foci formation in the absence of NSMCE2. (A) WT and NSMCE2 null cells were subjected to 1 Gy of IR, fixed at different

time points post IR (0 min: immediately after IR) and stained for γH2AX (a marker for DNA damage,

red) and SMC6 (green). SMC6 localized to sites of DNA damage in both WT and NSMCE2 null cells.

Bar = 5 μm. (B) Quantification of the average number of γH2AX foci in each cell. Data are presented

as mean ± SEM, n = 3. (C) Quantification of the average number of γH2AX foci that overlap with a

SMC6 focus. Data are presented as mean ± SEM, n = 3.

34 Chapter 2

Figure 4: Etoposide-induced DSB foci formation in the absence of NSMCE2. (A) WT and

NSMCE2 null cells were subjected to 3 µM etoposide for 1 h, fixed at different time points post

treatment (0 min: immediately after etoposide treatment) and stained for γH2AX (a marker for DNA

damage, red) and SMC6 (green). SMC6 localized to sites of DNA damage in both WT and NSMCE2

null cells. Bar = 5 μm. (B) Quantification of the average number of γH2AX foci in each cell.

Significantly more γH2AX foci were formed in NSMCE2 null cells. Data are presented as mean ± SEM,

n = 3. (C) Quantification of the average number of γH2AX foci that overlap with a SMC6 focus. Data

are presented as mean ± SEM, n = 3.

Absence of NSMCE2 affects survival upon etoposide-induced DSBs To measure the role of NSMCE2 in survival upon IR- or etoposide-induced DSBs, we

subjected both NSMCE2 null and WT cells to a clonogenic assay [30]. Firstly, the plating

efficiency, i.e., the percentage of plated single cells that develop into a cell colony of at least

50 cells determined in control conditions, was over three-fold lower in NSMCE2 null than in


WT cells (average of 25% compared to 75%, respectively) (Figure 5A). Interestingly, the

relative survival when cells were exposed to increasing doses of IR (0-8 Gy) only showed a

small difference between the WT and NSMCE2 null cells, with the latter being slightly more

sensitive (not statistically different though, Figure 5B). However, when exposed to 1 h of

increasing doses of etoposide, NSMCE2 null cells showed a clearly reduced viability

compared to WT cells (p < 0.05 in the case of 30 µM, Figure 5C).

Figure 5: Absence of NSMCE2 affects survival upon etoposide-induced DSBs. (A) Plating

efficiency of WT and NSMCE2 null cells during clonogenic assays. The survival capacity of plated

cells under non-challenged conditions was reduced in NSMCE2 null cells compared to WT. Data are

presented as mean ± SEM, n = 8. (B) Clonogenic assay after increasing doses of IR. NSMCE2 null

cells seemed to be slightly more sensitive to IR than WT cells (p > 0.05). Data are presented as mean

± SEM, n = 3. (C) Clonogenic assay after increasing doses of etoposide. NSMCE2 null cells were

significantly more sensitive to etoposide than WT cells. Data are presented as mean ± SEM, n = 3.

36 Chapter 2

The SMC5/6 complex physically interacts with topoisomerase II α (TOP2A) To validate that the SMC5/6 complex is indeed involved in topoisomerase II-mediated

relief of topological stress, we performed immunoprecipitations (IPs) using anti-SMC5 and

SMC6 antibodies. Both SMC5 and SMC6 clearly co-immunoprecipitated with each other, and

TOP2A clearly co-immunoprecipitated with SMC5 (Figure 6A), suggesting that the three

proteins are physically linked. To unequivocally establish the physical interaction among

these proteins, we conducted an additional IP with another antibody Abcam rabbit anti-SMC6,

followed by mass spectrometry of the bands that could represent SMC5/6 and TOP2A

(Figure 6B). We found that the band we presumed to represent full length SMC6 indeed

contained the SMC6 protein. Not unexpectedly, because SMC5 and SMC6 are physically

linked in the SMC5/6 complex, and SMC5 and SMC6 have equal sizes, we additionally

identified the SMC5 protein to be present at the same height (Figure 6B). Interestingly, the

lower running band (Figure 6B), which has been suggested to represent SMC6 lacking

posttranslational modifications [9, 10, 41], was convincingly identified as SEC23IP, a protein

previously shown to be involved in spermiogenesis [42]. Finally, we investigated the largest

band (approximately 175 kDa) that could potentially represent TOP2A and that was also

pulled down in this SMC6 IP (Figure 6B). Of the 23 identified peptides, 3 are homologous

between TOP2A and TOP2B, and 20 are unique to TOP2A. Because no specific peptides for

TOP2B were identified we conclude that this band indeed represented the protein TOP2A

(Data S1-S3).

Figure 6: The SMC5/6 complex physically interacts with TOP2A. (A)

Immunoprecipitations (IPs) and Western blot

analysis of SMC5 and SMC6. Both SMC5 and

SMC6 co-immunoprecipitated with each other.

Additionally, TOP2A co-immunoprecipitated with

SMC5. (B) For IP followed by mass spectrometry,

Abcam rabbit anti-SMC6 was used. SMC6-IP material was loaded on a 4%-12% bis-tris gradient gel

and stained with coomassie blue. Arrows indicate the bands that were isolated for mass spectrometry

analysis (green: potential SMC5/6 proteins; orange: potential TOP2A protein).

Discussion To study the SMC5/6 complex in an experimental setup, we used the novel CRISPR-

Cas9 system to target the SMC6 and NSMCE2 genes that encode the SMC6 and NSMCE2

subunits of the SMC5/6 complex, respectively. For NSMCE2, we did not get a complete


knockout after the first round of transfection and monoclonal isolation. The results were not

unexpected, given that U2OS is a cell line with chromosome counts in the hypertriploid range,

and that generating a true knockout is technically challenging since it requires disruption of

all functional copies of the gene. Consequently, a second round of gene targeting and single

cell expansion was performed. Eventually we established a bona fide cell line devoid of

NSMCE2. In the case of SMC6, however, we were unable to generate any CRISPR-modified

cells. Previous papers show that ablation of SMC6 resulted in embryonic lethality in mice [26]

and that conditional knockout of SMC5 in mouse embryonic stem cells induced apoptosis

[28]. Therefore, the failure to generate SMC6-deprived clones in our studies most likely

reflects that SMC6 is also essential for cell survival in humans.

To minimize the CRISPR-Cas9-induced off-target effects, we selected sgRNAs with the

highest specificity to coding exons of SMC6 and NSMCE2. Later, we analyzed the 10 top-

ranking potential off-target sites in the established NSMCE2 null cell line by sequencing, and

detected no off-target mutations. The results are in line with recent papers showing low

frequency of CRISPR-Cas9-induced off-target alterations in human cells [43, 44]. Hence,

although the possibility of off-target effects in modified cells could not be thoroughly excluded,

the differential phenotypes between NSMCE2 null and WT cells are thought to authentically

mirror gene functions.

We found clear differences in growth characteristics of the NSMCE2 null cells

compared to WT cells. The mutated cells have a prolonged cell cycle, and a clear larger

portion of the cells arrest in the G0-1 phase. Because this latter fraction is still equally present

after an extensive time in culture and multiple passages, it must be continuously

supplemented by cells exiting the cell cycle. Considering the differences in phenotypes, i.e.,

the presence of the G0-1-arrested cells, the overall slower growth rate and the reduced plating

efficiency of NSMCE2 null cells, it is plausible that the absence of NSMCE2 is not

immediately lethal to the majority of the cells, but poses growth challenges in normal culture

conditions that will ultimately arrest the cells.

These data are in line with a recent study performed in human breast cancer cells

(MCF-7), in which the depletion of NSMCE2 by RNA interference (RNAi) caused a slower

cell growth and increased percentage of G1 phase cells (70% vs. 55%-59% in control) [45].

Interestingly, depletion of NSMCE2 led to reduced levels of E2F1 protein and its downstream

target genes that are required for G1-S transition. This decreased growth rate was rescued

by ectopic expression of Flag-NSMCE2 but not its SUMO ligase-inactive mutant, suggesting

that the SUMO ligase activity of NSMCE2 ensures proper G1-S transition in these human

cancer cells [45].

38 Chapter 2

Although NSMCE2 is frequently linked to the DNA repair function of the SMC5/6

complex [33-37, 46, 47], the survival capacity of NSMCE2 null cells is only slightly affected

after increasing doses of IR. Likewise, IR-induced DSBs marked by γH2AX appear with

similar kinetic properties in WT and NSMCE2 null cells. We therefore conclude that NSMCE2

is not crucial for the repair of IR-induced DSBs. Nevertheless, the recruitment of SMC6 to

DSBs early post irradiation suggests that the SMC5/6 complex is involved in the early repair

of DSBs. In addition, NSMCE2 null cells did not display a differential survival rate in response

to cisplatin, a cytostatic agent causing DNA adducts and crosslinks that are generally

repaired by nucleotide excision repair (NER, Figure S1).

In contrast to the effects of IR or cisplatin, exposure to increasing doses of the

topoisomerase inhibitor etoposide does impair the survival capacity of NSMCE2 null cells. In

normal conditions, type II topoisomerase releases superhelical stress and untangles

chromosomes by creating transient DSBs, through which an unbroken DNA helix is

transferred before ligation of the break, in order to unwind the DNA double helix to prevent

supercoiling [40, 48]. Etoposide acts as a cytotoxic agent by forming a complex with the DNA

and the topoisomerase II enzyme, thereby preventing re-ligation of the DNA strands after

transient DSBs. Thus, etoposide treatment will ultimately lead to an increase of permanent

DSBs [38, 39]. The increased sensitivity to etoposide of NSMCE2 null cells is intriguing,

especially in the light of the interaction between SMC5/6 and TOP2A found in this study.

Interplay between TOP2A and the SMC5/6 complex has been suggested by several studies

in mouse and yeast [13, 49-52], but physical interaction between them is not confirmed.

Recently, it has been reported that Smc5/6 immunoprecipitated with the type II

topoisomerase in budding yeast [53]. Here, we for the first time unequivocally demonstrate

that TOP2A is indeed associated with the SMC5/6 complex in human cells. The absence of a

detectable band for TOP2A in the SMC6 guinea pig (GP) IP is most likely due to the lower

efficiency of the SMC6 GP antibody, which is supported by the observation that the SMC5 IP

generated more SMC6 protein than the SMC6 GP IP.

Topological tension is conceived when DNA molecules become supercoiled, for

instance preceding an advancing replication fork during chromosome duplication. This

positive supercoiling has to be removed in order for the replication fork to proceed. One

mechanism to avoid accumulation of supercoiled DNA ahead of the fork is to allow the

replication fork to proceed in a rotating manner following the turn of the helix. This will indeed

prevent the accumulation of supercoils ahead of the fork, but will simultaneously induce the

formation of sister chromatid intertwinings (SCI) behind the fork. Another way to release the

supercoiling is through topoisomerases, the enzymes creating single-strand nicks and


double-strand breaks. In this case TOP1 and TOP2 nick the DNA double helix ahead of the

fork, thereby allowing the unwinding of the supercoiled helix and the release of topological

tension. Because topological tension increases with the length of the chromosome,

topoisomerases are supposed to be more important to replication of longer chromosomes.

Indeed, budding yeast cells lacking functional topoisomerase I show a length-dependent

delay in replication [54]. In line with the observed interaction between SMC5/6 and TOP2A,

the association of budding yeast Smc5/6 with chromosomes during S-phase is also linearly

correlated with chromosome length, indicating that Smc5/6 somehow measures

chromosome length, probably by sensing topological tension [54, 55]. Moreover, Smc5/6

seems to also play a role in topological strain release, since budding yeast cells lacking

functional Smc6 or Nse2 show a delay in replication similar to Top2 mutants [54, 55]. Our

own data, showing that inhibition of topoisomerase activity has a more profound effect on

cells harboring an impaired SMC5/6 complex, together with physical interaction between the

SMC5/6 complex and TOP2A, further corroborate the presumed co-operation between

TOP2A and SMC5/6 at replication forks.

Considering the two mechanisms of tension release, SMC5/6 could function both

before and after the replication fork. Ahead of the fork, SMC5/6 could be responsible for the

correct repair of the TOP2A-induced DSBs. When the ligase function of TOP2A is inhibited

by etoposide, re-initiation of replication might rely more on SMC5/6, which will be challenged

when NSMCE2 deletion impairs SMC5/6 function. On the other side of the fork, SMC5/6

might be required to stabilize the SCIs, as proposed previously [54]. In this model, SMC5/6

associates to SCIs, thereby fixating them and allowing fork rotation, and reducing topological

tension. In parallel, budding yeast Smc5/6 is also involved in the actual resolving

recombination intermediates in order to prevent toxic chromosome structures [56]. In addition,

budding yeast Top2 is also essential for the removal of SCIs that would otherwise lead to

segregation errors during the subsequent M-phase [57-59]. Since both induction and removal

of SCIs involves transferring one DNA double helix through another via transient formation

and repair of a DSB [48], it is likely that SMC5/6 is also working together with TOP2A at the

level of SCIs.

Furthermore, in budding yeast, Top2 activity relies on sumoylation and failure to

sumoylate Top2 disrupts the ability of Top2 to separate replicated chromosomes [60]. Of

note, human TOP2 is also found conjugated to SUMO [61]. However, RANBP2 seems to be

the major SUMO E3 ligase for TOP2A in mice [62], and a mutation compromising NSMCE2

sumoylation activity does not affect murine lifespan [27]. Nevertheless, we cannot rule out

that SMC5/6 function at the replication fork might involve NSMCE2-mediated sumoylation of

TOP2A, which could explain the interaction between SMC5/6 and TOP2A and the effects

seen in NSMCE2 null cells.

40 Chapter 2

Our findings, demonstrating a physical interaction between SMC5/6 and TOP2A and

an increased sensitivity of NSMCE2 null cells to etoposide, suggest that the SMC5/6

complex helps to resolve topological stress. Since this physical interaction is even present in

cells that are not challenged by IR or cytotoxic agents, SMC5/6 and TOP2A seem already to

function together during S-phase under normal non-challenged circumstances. In this

respect, it is plausible that the fraction of NSMCE2 null cells arresting in G0-1 phase is

actually a representation of cells with stalled or collapsed replication forks very early in the

replication process, which occurs naturally, yet cannot be resolved properly due to the lack of

NSMCE2. While in WT cells these replication forks are normally repaired and restarted by

SMC5/6, the absence of NSMCE2 first induces a delay in early replication progression,

explaining the increased cell cycle duration of NSMCE2 null cells. Subsequently, residual

repair defects may trigger cells to eventually arrest in G0. In this respect, several studies in

yeast have suggested that Smc5/6-mutated cells will undergo cell division despite the

presence of chromosomal abnormalities caused by defective DNA repair mechanisms [33,

49, 52, 63, 64]. However, over time, the amount of chromosomal abnormalities within a cell

will accumulate, eventually leading to cell cycle arrest.

In conclusion, in the light of current literature and the data we present here, we propose

that the SMC5/6 complex functions in resolving TOP2A-mediated recombination

intermediates endogenously generated early during DNA replication in human cells.

Acknowledgments We thank Berend Hooibrink and Daisy Picavet of the core facility Cellular Imaging of

the Academic Medical Center (AMC) for assistance with and use of their equipment for

FACS analysis and confocal microscopy. Furthermore, we thank Klaas Franken, Hans

Rodermond and Bregje van Oorschot for assistance with and use of their 137Cs source for IR.

Finally, we thank Philip W. Jordan for generously providing the TOP2A antibody and for

fruitful discussions. This study has been supported by an AMC Fellowship, the People

Programme (Marie Curie Actions) of the European Union’s Seventh Framework Programme

(CIG 293765) to Geert Hamer and the China Scholarship Counsel (CSC) number

201306300081 to Yi Zheng.

Author contributions Dideke E. Verver, Yi Zheng, Jan Stap and Geert Hamer conceived and designed the

experiments. Dideke E. Verver, Yi Zheng, Dave Speijer, Ron Hoebe and Henk L. Dekker

performed the experiments. Dideke E. Verver, Yi Zheng, Jan Stap and Geert Hamer


analyzed the data. Dideke E. Verver, Yi Zheng, Sjoerd Repping and Geert Hamer wrote the

manuscript.

Conflicts of interest The authors declare no conflict of interest.

Abbreviations SMC Structural Maintenance of Chromosomes

NSMCE Non-SMC Element

DSB DNA double-strand break

IR Ionizing Radiation

TOP Topoisomerase

SCI Sister chromatid intertwining

42 Chapter 2

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Supplementary materials

Table S1: The selected off-target sites and the corresponding PCR primers.

Potential off-target

sites

Genomic loci

(GRCh38/hg38)

Mis

matches PCR primers

ACCCAAATCTGTATC

AACTC AGG

62065416-62065438,

chromosome 11 1, 2, 7, 8

F: GTCCCTCCATCTTGGTGCCT

R: GGGTCTTTGCTGCCTGTGA

CTCTTATCCTGTATC

AACTC AAG

43274942-43274964,


F: CACTGACAACAGGCATGAAAT

R: CTGGAGACTGAGGCAGGAGA

TTTCTATTCTGTATC

AACTC AAG

153197613-153197635,


F: ATTGGGCCTTATGAACTGATTC

R: TGGTCTACGCAGGGTAAGGATA

TGCCAAGGCTGTAT

GAACTC AGG

110987269-110987291,

chromosome 2 (in gene) 2, 8, 15

F: ACTGCTTGGAACAGTGAACATG

R: GCTGAGACTGATGAGCGATAAA

TCCAAAGTCTGTATC

AACTT TAG

2809500-2809522,

chromosome 8

2, 4, 8,

20

F: CAGCCAAACCATATCATTCTGT

R: TGTTTTCATGTTTGTGGCAGTG

TTCCAAGACTATATC

AACTA AAG

114757228-114757250,

chromosome 11 8, 11, 20

F: GTATTGCAGCAAGCCATTACC

R: AAGAATCTGCTCTGGAGGGAG

TTCCTGGCCTGTATC

AACAC AAG

52436381-52436403,

chromosome 12 5, 6, 19

F: ACGGAACCAGGTGAAGGAA

R: TTGGCACTTGGAGCGGTAG

TTTGAAGTCTGTATC

ATCTC AAG

104356169-104356191,

chromosome 5

3, 4, 8,

17

F: CACTCTTACTTTGTTCCCCACA

R: CCTCACTTGCCTTGCCTATT

TACTAAACATGTATC

AACTC AGG

31343808-31343830,

chromosome X 2, 4, 7, 9

F: GCTGGCGGGTGCAATTAGT

R: AGAGCAAGACCCTGACCCTAA

GCCCAAGCCAGTAT

CACCTC AAG

128369750-128369772,

chromosome 5

1, 2, 10,

17

F: CGGAAAGTGGGAGTAAGAAATC

R: GCTCTAATCACTGGCTATGCTAT

48 Chapter 2

Figure S1: Clonogenic assay after increasing

doses of cisplatin. No clear difference in

sensitivity to cisplatin could be detected between

WT and NSMCE2 null cells.

Data S1-S3 can be found at www.mdpi.com/1422-0067/17/11/1782/s1.

Chapter 3

Trivial role for NSMCE2 during in vitro proliferation and differentiation of male germline stem cells

Yi Zheng

Aldo Jongejan

Callista L. Mulder

Sebastiaan Mastenbroek

Sjoerd Repping

Yinghua Wang

Jinsong Li

Geert Hamer

Reproduction

2017 Sep;154(3):81-95

50 Chapter 3

Abstract Spermatogenesis, starting with spermatogonial differentiation, is characterized by

ongoing and dramatic alterations in composition and function of chromatin. Failure to

maintain proper chromatin dynamics during spermatogenesis may lead to mutations,

chromosomal aberrations or aneuploidies. When transmitted to the offspring, these can

cause infertility or congenital malformations. The structural maintenance of chromosomes

(SMC) 5/6 protein complex has recently been described to function in chromatin modeling

and genomic integrity maintenance during spermatogonial differentiation and meiosis.

Among the subunits of the SMC5/6 complex, non-SMC element 2 (NSMCE2) is an important

small ubiquitin-related modifier (SUMO) ligase. NSMCE2 has been reported to be essential

for mouse development, prevention of cancer and aging in adult mice and topological stress

relief in human somatic cells. By using in vitro cultured primary mouse spermatogonial stem

cells (SSCs), referred to as male germline stem (GS) cells, we investigated the function of

NSMCE2 during spermatogonial proliferation and differentiation. We first optimized a

protocol to generate genetically modified GS cell lines using CRISPR-Cas9 and generated

an Nsmce2-/- GS cell line. Using this Nsmce2-/- GS cell line, we found that NSMCE2 was

dispensable for proliferation, differentiation and topological stress relief in mouse GS cells.

Moreover, RNA sequencing analysis demonstrated that the transcriptome was only minimally

affected by the absence of NSMCE2. Only differential expression of Sgsm1 appeared highly

significant, but with SGSM1 protein levels being unaffected without NSMCE2. Hence, despite

the essential roles of NSMCE2 in somatic cells, chromatin integrity maintenance seems

differentially regulated in the germline.

NSMCE2 in male germline stem cells 51

Introduction

The process of spermatogenesis is characterized by ongoing and dramatic alterations

in composition and function of chromatin. Chromatin is the supra-molecular complex,

consisting of DNA and proteins, which packages, shapes and orchestrates the genome and

safeguards genomic stability. Incorrect spatio-temporal organization of chromatin can initiate

germ cell apoptosis, leading to spermatogenic arrest and male infertility [1]. If all

spermatogenic arrest mechanisms fail, incorrect chromatin architecture can even cause

chromosomal aberrations or aneuploidies, leading to congenital abnormalities in the offspring

[2].

Numerous chromatin-based processes, such as cell cycle progression, cellular

differentiation and genomic integrity maintenance, are to a large extent modulated by the

structural maintenance of chromosomes (SMC) protein complexes: SMC1/3 (cohesin),

SMC2/4 (condensin) and SMC5/6. Besides SMC5 and SMC6, the SMC5/6 complex contains

several non-SMC elements (NSMCEs), including NSMCE2. Together with these NSMCEs,

the SMC5/6 complex has a ring-like structure large enough to hold two double-stranded DNA

molecules together. This property of the SMC5/6 complex is pivotal for recombination-

mediated DNA damage repair [3-5] and resolving replication-induced topological stress in

yeast [6-8]. Of the four NSMCEs in mammals, NSMCE2 specifically associates with SMC5,

where it exhibits a C-terminal SP-RING domain with an E3 small ubiquitin-related modifier

(SUMO) ligase activity. This SUMO ligase activity of NSMCE2 is required for the DNA

damage repair activity of the SMC5/6 complex [9-11].

Using various model organisms, the function of the SMC5/6 complex during meiosis

has been extensively investigated and reviewed [12]. Meiotic processes involving the

SMC5/6 complex include chromosome segregation [13-17], homologous chromosome

synapsis [13-15, 18-20] and meiotic sex chromosome inactivation [14, 16]. Very likely, the

SMC5/6 complex prevents dangerous and error-prone recombination events in highly

repetitive, densely packed genomic regions such as the rDNA and pericentromeric

heterochromatin [14, 16, 19-22]. Aberrant recombination between such repetitive sequences

would otherwise interfere with the tightly regulated process of meiotic recombination, for

instance, by causing intra-chromosomal recombination events. Indeed, in yeast and C.

elegans, mutations in Smc5, Smc6 or the SUMO ligase domain of Nsmce2 lead to the

accumulation of toxic joint molecules caused by failure to resolve meiotic recombination

intermediates [13, 19, 20, 23-26].

In mitotically dividing yeast, the SMC5/6 complex is essential for the maintenance of

replication fork stability and the prevention of replication-induced topological stress [6-8].

Consistently, depletion of Smc5 in mouse embryonic stem (ES) cells leads to accumulation

52 Chapter 3

of cells in G2 and subsequent mitotic failure and apoptosis [27]. Using a human

osteosarcoma cell line (U2OS), we have recently shown that SMC6 interacts with DNA

topoisomerase II α (TOP2A) [28]. TOP2A is a topoisomerase that prevents supercoiling of

replicating DNA [29, 30]. Moreover, we have shown that the CRISPR-Cas9-mediated

removal of NSMCE2 in these cells led to increased sensitivity to the topoisomerase inhibitor

etoposide [28].

In the mouse and human, the SMC6 protein is most highly expressed in the testis

where it appears to be involved in spermatogonial differentiation [15, 16] and meiosis [14-16].

Likewise, also NSMCE2 has been identified to be expressed in developing mouse male germ

cells, from spermatogonia to round spermatids [31]. Recently, a study using a conditional

knock-out (KO) mouse model showed essential roles for SMC5/6 during meiotic

chromosome segregation [17]. However, suitable KO models to study the SMC5/6 complex

in mitotically dividing spermatogonia are currently not available. In mice, complete ablation of

SMC6 or NSMCE2 results in embryonic lethality [31, 32]. Hence, the role of SMC5/6 or

NSMCE2 in spermatogonia remains largely unknown.

Male germline stem (GS) cells, initially termed by Shinohara’s group, refer to the

cultured spermatogonial stem cells (SSCs) able to propagate in vitro for over 2 years without

losing SSC properties [33]. In the current study, we first optimized a protocol to generate

genetically modified mouse GS cell lines using CRISPR-Cas9. By applying this optimized

protocol, we generated an Nsmce2-/- GS cell line. Using this GS cell line, we studied the role

of Nsmce2 during in vitro spermatogonial proliferation and differentiation, gene expression

and the spermatogonial response to topological stress.

Materials and methods Animal use and care

Neonatal (4-5 d.p.p) DBA/2J (Charles River) male mice were used for GS cell isolation.

To acquire neonatal testis materials, donor mice were first anesthetized by 4% isoflurane

total body anesthesia followed by killing by decapitation and inactivation of the brain. Testes

were collected and the tunica albuginea was removed. Testicular tissues were cryopreserved

in supplemented MEM (Gibco, Thermo Fisher Scientific) containing 20% fetal bovine serum

(FBS) and 8% DMSO in a Coolcell freezing device and stored in liquid nitrogen (-196°C) for

future GS cell isolation. All animal procedures were in accordance with and approved by the

animal ethical committee of the Academic Medical Center, University of Amsterdam.


Design of Nsmce2- single-guide (sgRNA) and construction of CRISPR-Cas9 plasmids The online Optimized CRISPR Design Platform (http://crispr.mit.edu/) was utilized to

design Nsmce2-sgRNA. 5’-ACCCGTTACATATCCTTCAG-3’, followed by the protospacer-

adjacent motif (PAM) TGG (in exon 2) was selected as the target site. The corresponding

forward and reverse strand oligonucleotide was synthesized by Sigma-Aldrich, and then

annealed and cloned into the commercial linearized vector GeneArt CRISPR Nuclease

Vector with OFP Reporter (Thermo Fisher Scientific), following the protocol provided by the

manufacturer. The correct double-strand oligonucleotide insertion was confirmed by Sanger

sequencing after transformation and plasmid extraction.

GS cell culture and differentiation A mouse GS cell line was established following a previously published protocol [34].

Briefly, germ cells were isolated from the cryopreserved testes of 4-5 d.p.p DBA/2J male

mice by a two-step enzymatic dissociation. After overnight incubation on gelatin-coated wells,

the floating and loosely attached cells were collected and cultured in the complete GS cell

medium composed of StemPro-34 SFM medium (Thermo Fisher Scientific), StemPro-34

Supplement (Thermo Fisher Scientific), 1% FBS, 10 ng/ml recombinant human GDNF

(Peprotech), 10 ng/ml recombinant human bFGF (Peprotech), as well as other 17

components as previously reported [34, 35]. The cells were refreshed every 2-3 days and

passaged every 5-7 days at a ratio of 1: 4-6. From the third passage, the cells were

transferred to inactivated primary mouse embryonic fibroblast (MEF) feeder cells that had

been treated with 10 µg/ml mitomycin-C (Sigma-Aldrich) for 2-3 hours at 37 °C. The cells

were maintained at 37°C in an atmosphere of 5% CO2 in air. After ~1 month, the growth of

GS cells became stable. GS cells were cultured on MEFs unless otherwise stated. For

feeder-free culture, GS cells were seeded on wells pre-coated with laminin (20 µg/ml, Sigma-

Aldrich). For retinoic acid (RA)-induced differentiation of GS cells, the feeder-free culture was

adopted, and GS cells were exposed to medium containing 2µM all-trans-RA (Sigma-Aldrich)

for 3 days. In control groups, vehicle (0.1% ethanol) was added to the medium.

GS cell electroporation The constructed CRISPR plasmids targeting Nsmce2 were delivered to low-passage

GS cells (˂P10) by Neon electroporator (Thermo Fisher Scientific), following the

manufacturer’s guidance. The program used for electroporation was voltage 1100, width

20ms and pulse 2. Two days after electroporation, OFP+ cells were sorted by fluorescence-

activated cell sorter (FACS, BD Biosciences) and cultured on MEFs for recovery.

54 Chapter 3

Surveyor assay One week after FACS sorting, the genomic DNA of GS cells was extracted, and the

genomic region around the target site was amplified by PCR with the forward primer 5’-

GATGATGGCACAGTGCTTGG-3’ and the reverse primer 5’-

GGCAGTTCTGAGTGGAGGATTA C-3’. Herculase II fusion polymerase (Agilent

Technologies) was used for high-fidelity PCR amplification. PCR products were purified,

denatured and re-annealed to generate DNA heteroduplexes, followed by the Surveyor

nuclease (Integrated DNA Technologies) digestion, according to the protocol provided by the

manufacturer. The Surveyor nuclease digestion products were run and visualized on agarose

gels. The incidence of insertion/deletion (indel) was calculated using a previously described

formula [36].

GS cell clonal isolation and expansion One week after FACS sorting, the recovered GS cells were dissociated by accutase

(Thermo Fisher Scientific) and filtered through a 50µm mesh to remove cell aggregates. The

single GS cells were plated on 6-well plates pre-coated with laminin, at a density of 2,000-

4,000 cells/well. One week after plating, single GS cell-derived patches were detached with

0.5mM EDTA, manually picked under the microscope, and each cell patch was transferred to

one well of a 96-well plate pre-coated with MEFs. After ~1 month, the expanded colonies

were dissociated by accutase and transferred to individual wells of a 48-well plate with MEFs

and to larger wells thereafter. Since the clonal expansion proceeded to 6-well plates, the cell

culture was carried out routinely.

Genotyping monoclonal GS cell lines Before genotyping, a subpopulation of monoclonal GS cell lines was cultured on

laminin for several passages to thoroughly eliminate the mixed MEFs. Then, the genomic

DNA was extracted from each monoclonal GS cell line, and the region around the target site

was amplified by PCR with the uniform primers for the Surveyor assay, but with a different

high-fidelity polymerase (Easy-A high-fidelity PCR cloning enzyme, Agilent Technologies).

The purified PCR products were cloned into T-Vector pMD19 (TaKaRa) following the

manufacturer’s guidance. After transformation and overnight incubation, twenty colonies for

each reaction were picked at random and sequenced.


Off-target analysis The 10 top-ranking potential off-target loci, provided by the online Optimized CRISPR

Design Platform (http://crispr.mit.edu/), were analyzed. In brief, the genomic DNA was first

extracted from monoclonal GS cell lines, and the regions flanking each potential off-target

site were PCR-amplified with the Herculase II fusion polymerase (Agilent Technologies) and

the primers shown in Table 1. The purified PCR products were then sequenced for off-target

analyses. Table 1: The selected potential off-target sites and the corresponding PCR primers for

sequencing. Potential off-target sites Genomic loci

(GRCm38/mm10) Mis-matches

PCR primers

ACCTGTTACATATCCTTCGG

AGG

60636826-60636848,

chromosome 12, - 4, 19

F: AAGTTAGTTTAGGGCACAAAGG

R: AGTCCACCAGGTTAGAAAAGC

TCCTGTTACGTATCCTTCAG

CAG

4770918-4770940,

chromosome 1, + 1, 4, 10

F: GCTCCCTGGCTTTCTCATT

R: CTTGCCCGTGTCCTCTACTA

ACACTGTACATATCCTTCAG

CAG

22477599-22477621,

chromosome 18, - 3, 5, 6

F: TGGTCCAAAATTCGCTGTAA

R: GTGGCATCAGGGCAAACA

AAGCTTTCCATATCCTTCAG

TGG

107576559-107576581,

chromosome 6, + 2, 3, 5, 8

F: CCTGGAACTAACTCTAGGGATG

R: CTGGATGTTTCTTAATGGGACT

AGGCGTCTCATATCCTTCAG

TGG

148326544-148326566,

chromosome 2, - 2, 3, 7, 8

F: ATTTCCAGCAGAGTCCCACT

R: GACCCCAAGGCCATTATTC

TTCAGTTAGATATCCTTCAG

TGG

91837662-91837684,

chromosome 11, - 1, 2, 4, 9

F: CCTAAGCTGCTGCCTAAAAG

R: CACATGACATTCTGATCTTGCA

GCCAGTTCCACATCCTTCAG

CAG

34570031-34570053,

chromosome 2, + 1, 4, 8, 11

F: TGCTCAAGGAGGAGGAAACT

R: GGAAGGCTGGAAGTGGTGT

ACATGTTTCAGATCCTTCAG

AAG

75040008-75040030,

chromosome 3, - 3, 4, 8, 11

F: GTGGCAAATCTGGGTGGA

R: TCTGAGGGTAGGCTGTGAGG

ACTTCTTACAAATCCTTCAG

TGG

16346304-16346326,

chromosome 14, + 3, 4, 5, 11

F: TGGCAGTGATGAGAAAACGA

R: GCTCTGAGGATGGAATGGGT

GCCAGTTCCATTTCCTTCAG

TAG

123031365-123031387,

chromosome 10, + 1, 4, 8, 12

F: AGCTGGCAGTCTATGAGTCAAT

R: TGCCCAGTGCGATTTGAT

Western blot analysis

Before Western blot analysis, a subpopulation of Nsmce2+/+ and Nsmce2-/- GS cells

were cultured on laminin for several passages to thoroughly eliminate the mixed MEFs. Then,

the protein was isolated and Western blot analysis was conducted as previously reported [16,

28], with the LI-COR Odyssey imaging system (Biosciences). The primary antibodies used

were rabbit anti-NSMCE2 (1: 200; provided by Oscar Fernandez-Capetillo), rabbit anti-SMC5

56 Chapter 3

(1: 500; A300-236A, Bethyl Laboratories), guinea pig anti-SMC6 (1: 200; custom made,

peptide: KRPRQEELEDFDKDGDEDE), mouse anti-PLZF (1: 100; D-9, Santa Cruz

Biotechnology), mouse anti-OCT4 (1: 100; C-10, Santa Cruz Biotechnology), rabbit anti-

STRA8 (1: 1,000; ab49602, Abcam), rabbit anti-SGSM1 (1: 1,000; ab171943, Abcam),

mouse anti-β-actin (1: 5,000; A1978, Sigma-Aldrich) and rabbit anti-GAPDH (1: 400; FL-335,

Santa Cruz Biotechnology).

Cell cycle analysis Cell cycle analysis based on DNA content was performed as previously described [28].

DNA content was analyzed with the FACS analyzer (BD Biosciences) and the figures were

constructed via the FlowJo software. Data were presented as the mean ± standard error of

mean (S.E.M.). Differences between groups were assessed using the Student’s t-test. A

difference was considered significant when p˂0.05.

EdU assay The cell proliferation assay was performed using a Click-iT EdU Alexa Fluor 488

imaging kit (Thermo Fisher Scientific), following the protocol provided by the manufacturer. In

brief, GS cells were grown on laminin-coated glass coverslips in 24-well plates. On the day of

the treatment, cells were incubated with 10µM EdU diluted in complete medium for 2 h at

37 °C, followed by fixation in 4% paraformaldehyde (PFA) for 10 min. After permeabilization,

cells were incubated with the reaction cocktail for 30 min at room temperature (RT), and then

counterstained with DAPI for 5 min. Cells were mounted on glass slides with the Prolong

Gold anti-fade mountant (Thermo Fisher Scientific) and later subjected to visualization under

the microscope. For quantification of EdU+ cells, at least 300 cells were analyzed in each

group. Data were presented as the mean ± S.E.M. Differences between groups were

assessed using the Student’s t-test. A difference was considered significant when p˂0.05.

Immunocytochemistry GS cells were grown on laminin-coated glass coverslips in 24-well plates for all

immunocytochemical experiments. In case of etoposide treatment, GS cells were incubated

with 10µM etoposide for 3 h at 37 °C, then fixed at different time points, i.e. 0 h (immediately

post treatment), 1, 3 and 5 h after treatment respectively, in 4% PFA for 10 min. Cells were

permeabilized in phosphate-buffered saline (PBS) with 0.1% triton-X for 15 min, followed by

1 h of blocking in PBS with 1% bovine serum albumin (BSA) and 0.25% Tween20. Primary


antibodies were applied to cells at 4°C overnight. The primary antibodies used were mouse

anti-PLZF (1: 50; D-9, Santa Cruz Biotechnology), mouse anti-OCT4 (1: 50; C-10, Santa

Cruz Biotechnology), rabbit anti-LIN28A (1: 1,000; ab46020, Abcam), rabbit anti-ID4 (1: 100;

M106, CalBioreagents), mouse anti-ɣ-H2AX (1: 20,000; 05-636, Merck Millipore) and guinea

pig anti-SMC6 (1: 400; custom made). Omission of the primary antibodies and replacement

with mouse, rabbit and guinea pig isotype IgGs were used as negative controls. After

washing with PBS on the next day, the cells were incubated with the corresponding

secondary antibodies (Alexa Fluor 488 or 555, 1: 1,000; Thermo Fisher Scientific) for 1h at

RT. After counterstaining with DAPI for 5 min at RT, the cells were mounted on glass slides

with the Prolong Gold anti-fade mountant (Thermo Fisher Scientific) and later subjected to

visualization under the microscope. For quantification of ɣ-H2AX+ cells (˃5 ɣ-H2AX foci/cell),

at least 50 cells were analyzed in each group. Data were presented as the mean ± S.E.M.

Differences between groups were assessed using the Student’s t-test. A difference was

considered significant when p˂0.05.

Microscopy Fluorescence microscopy images were acquired at RT using a Leica DM5000B

microscope equipped with a Leica DFC365 FX CCD camera. Images were analyzed using

Leica Application Suite Advanced Fluorescence (LAS AF) software. The presented figures

were constructed using Adobe Photoshop CS6.

RNA sequencing (RNA-seq) Total RNA was extracted from Nsmce2-/- and Nsmce2+/+ GS cells, respectively, using

PureLink RNA Micro Kit (Thermo Fisher Scientific). Biological triplicates from different

passages were prepared. Total RNA was sent to BGI Tech Solutions (Hong Kong, China) for

library construction and sequencing (Illumina HiSeq 4000, paired end 100bp). Reads were

subjected to quality control and aligned to the UCSC mm10 (GRCm38.p4) genome using

HISAT2 (v2.0.4) [37]. Counts were obtained using HTSeq (v0.6.1) [38] using the UCSC

mm10 GTF. Statistical analyses were performed using the edgeR [39] and limma [40] R

(v3.2.2)/Bioconductor (v3.0) packages. All genes with no counts in any of the samples were

removed (15,633 genes), whilst genes with more than 1 count-per-million reads (CPM) in at

least 2 of the samples were kept (31,435 genes). Count data were transformed to log2-counts

per million (logCPM), normalized by applying the trimmed mean of M-values method [39] and

precision weighted using voom [41]. Differential expression was assessed using an empirical

Bayes moderated t-test within limma’s linear model framework including the precision

58 Chapter 3

weights estimated by voom. Resulting P values were corrected for multiple testing using the

Benjamini-Hochberg false discovery rate. Corrected P values of ˂0.05 were considered as

statistically significant. Genes were re-annotated using biomaRt (v2.26.1) using the Ensembl

genome databases (v85). The resulting DEGs (adj.P˂0.05) and entire RNA-seq data are

shown in Table 2 and Supplementary Table 1 (see section on supplementary data given at

the end of this article) respectively. Gene Set Enrichment Analysis (GSEA) software [42, 43]

was used to analyze differentially expressed gene sets, and the GSEA results are shown in

Supplementary Table 2.

Table 2: The DEGs obtained by analysis of RNA-seq data from Nsmce2-/- and Nsmce2+/+ GS

cells.

Gene symbol Description logFC adj.P.Val (˂0.05) Mx1 MX dynamin-like GTPase 1 2.647288067 0.046661415

Skor1 SKI family transcriptional corepressor 1 0.715634301 0.046661415

Myog myogenin 4.244660863 0.044188003

Gbgt1 globoside alpha-1,3-N-

acetylgalactosaminyltransferase 1

1.234137519 0.046661415

Sgsm1 small G protein signaling modulator 1 -1.795957187 0.003750701

Zfp358 zinc finger protein 358 -1.292106048 0.046661415

Smim22 small integral membrane protein 22 2.450729769 0.046661415

Gm9 predicted gene 9 6.259824968 0.046661415

Gm30332 predicted gene 30332 -6.010552517 0.003750701

Accession numbers All sequence data have been submitted to NCBI (SRA) and will be available under the

accession number: ID PRJNA379902.

Results Construction of CRISPR-Cas9 plasmids and gene targeting of Nsmce2 in GS cells

To perform loss of function study for Nsmce2, we first designed a sgRNA to target exon

2 of the mouse Nsmce2 locus (Figure 1A). The target site is located in an early and

conserved coding sequence of the two Nsmce2 transcript variants. Frameshift mutations at

this site will eliminate NSMCE2 function, including its only known activity, the E3 SUMO

ligase through its C-terminal SP-RING domain [9-11]. To enable recognition and fluorescent

sorting of transfected cells, we then constructed CRISPR-Cas9 plasmids containing this

sgRNA and an orange fluorescent protein (OFP) reporter (Figure 1B).

To establish a GS cell line, we isolated germ cells from the testes of 4-5 d.p.p DBA/2J

male mice and cultured the isolated germ cells following an established protocol published by


Shinohara’s group [34]. After 3-4 weeks, the GS cells were transferred to inactivated MEFs

for subculture and they formed the distinctive grape-like colonies (Figure 1C), consistent with

former reports [34, 44].

GS cells have been shown to be extremely refractory to commonly used transfection

methods such as calcium phosphate precipitation and lipofection [45]. Nevertheless, novel

electroporation devices can be harnessed to transfect spermatogonia with moderate

efficiency [46-50]. To this end, we utilized a Neon electroporator to deliver CRISPR vectors

into early passage GS cells. Two days after electroporation, OFP+ cells (Figure 1D) were

enriched by FACS (Figure 1E) and transferred to MEF feeder cells for recovery. One week

after FACS sorting, we conducted a Surveyor assay [36] based on the sorted cells. The

target site-specific PCR amplicons were digested by Surveyor nucleases, yielding fragments

with expected sizes indicative of indel mutations, demonstrating the occurrence of gene

editing (Figure 1F).

Figure 1: CRISPR-Cas9-mediated targeting of Nsmce2 in GS cells. (A) The design of a sgRNA

targeting exon 2 of the mouse Nsmce2 locus. Blue letters represent the 20-bp target, whereas red

letters refer to the PAM NGG. (B) A schematic overview of the constructed CRISPR-Cas9 plasmids

targeting Nsmce2. (C) The grape-like GS cell colonies when cultured on MEFs. Bar = 100µm. (D) GS

cells on laminin 1 day after electroporation. Bar = 100µm. (E) FACS enrichment of OFP+ cells

harboring CRISPR-Cas9 plasmids. (F) Surveyor assay-induced cleavage bands (approximately 300

and 218bp respectively, arrowheads). Negative control (-) represents the group without transfection.

Generation of Nsmce2-/- GS cell lines

Next, we derived monoclonal Nsmce2-/- GS cell lines. Initially, we exploited FACS to

deposit single GS cells into individual wells of one 96-well plate (1 cell/well) pre-coated with

MEF feeder cells. However, after 2 weeks of culture, we did not observe any GS cell clones.

60 Chapter 3

To overcome this hurdle, we determined to pick single cell-derived patches. Specifically, one

week after FACS sorting, the sorted and recovered GS cells were plated on 6-well plates

pre-coated with laminin, at a low density (2,000-4,000 cells/well) (Figure 2A). The wells were

pre-coated with laminin instead of MEFs because GS cells can attach rapidly to laminin,

thereby circumventing the problem of polyclonal formation when single GS cells are plated at

a low density. By contrast, we observed that GS cells attached slowly to MEFs and that

many single cells readily aggregated before attachment. Previous reports have shown that

MEFs as feeders are dispensable and GS cells can also be cultured on laminin for a long

time [51-53]. One week after plating, approximately 1-10% of single GS cells formed cell

patches comprising 4-6 cells (Figure 2B). They were then detached with EDTA, manually

picked under the microscope and each cell patch was transferred to one well of a 96-well

plate pre-coated with MEFs for further clonal expansion. After ~1 month, the expanded

colonies (Figure 2C) were dissociated and transferred to individual wells of a 48-well plate

coated with MEFs and to larger wells thereafter. Eventually, the clonal expansion proceeded

to 6-well plates. The whole process of the clonal expansion took ~2.5 months. In total, we

derived 7 GS cell lines from 48 picked single-cell patches. PCR amplification of the genomic

DNA region around the target site followed by TA cloning and Sanger sequencing revealed

that 2 of them carried gene modifications at the Nsmce2 locus. Unfortunately, one Nsmce2-/-

GS cell line failed to expand, probably due to premature passage when cells remained at a

small number. Fortunately, the other Nsmce2-/- GS cell line, harboring bi-allelic frameshift

mutations (Figure 2D), was successfully expanded. Western blot analysis, using a

transfected and single cell-derived Nsmce2+/+ GS cell line as a positive control, confirmed the

eradication of the corresponding NSMCE2 protein (Figure 2E). Protein levels of SMC5 and

SMC6 were not influenced by removal of NSMCE2 (Figure 2E). We validated these results

using lysates from Nsmce2+/+ and Nsmce2-/- MEFs (described in [31]) (Figure 2E). Finally, we

sequenced the 10 top-ranking potential off-target sites (Table 1) in the established Nsmce2-/-

cell line and detected no off-target mutations. This Nsmce2-/- GS cell line, together with the

control Nsmce2+/+ GS cell line used in Figure 2E, were used for all further experiments.

Removal of NSMCE2 does not influence the spermatogonial cell cycle or proliferation

Morphologically, Nsmce2-/- GS cells were indistinguishable from their Nsmce2+/+

counterparts, and both formed the characteristic grape-like colonies on MEFs (Figure 3A),

distinct from the ES-like multipotent GS (mGS) cell colonies. Because we have previously

demonstrated a prolonged cell cycle of Nsmce2-null U2OS cells [28], we investigated


whether the removal of NSMCE2 also alters the proliferation/cell cycle time of GS cells. The

two single cell-derived GS cell lines, Nsmce2+/+ and Nsmce2-/- respectively, did not have

significantly different cell doubling time (Figure 3B). Consistently, cell cycle analysis by DNA

histogram showed no difference between the two cell populations with respect to the ratios of

cells in different cell cycle phases (Figure 3C). To further investigate the role of NSMCE2 in

controlling GS cell proliferation, we performed an EdU incorporation assay. The proportion of

cells that incorporate the thymidine analog EdU (a measurement for DNA synthesis) did not

differ between Nsmce2+/+ and Nsmce2-/- GS cells (Figure 3D). Furthermore, both cell lines

could normally propagate in vitro for more than 35 passages, without significant changes in

cellular morphology or cell doubling time. Overall, the results suggest that the removal of

NSMCE2 does not influence in vitro spermatogonial proliferation.

Figure 2: Generation of Nsmce2-/- GS cell lines. (A) Single GS cells immediately after plating on

laminin. Bar = 100µm. (B) Single cell-derived patches 1 week after seeding. Bar = 100µm. (C) A single

GS cell-derived colony after 1 month of culture in a well of 96-well plate coated with MEFs. Bar =

100µm. (D) Sanger-sequencing analysis of the Nsmce2-/- GS cell line. (E) Immunoblotting of NSMCE2,

SMC5 and SMC6 in the established Nsmce2+/+ and Nsmce2-/- GS cell lines, and in the control

Nsmce2+/+ and Nsmce2-/- MEFs. β-Actin is used as a loading control.

62 Chapter 3

Figure 3: The deprivation of NSMCE2 does not influence the self-renewal of GS cells. (A)

Representative images of Nsmce2+/+ and Nsmce2-/- GS cells on MEFs. Bar = 100µm. (B) The doubling

time of Nsmce2+/+ and Nsmce2-/- GS cells. Data are presented as the mean ± S.E.M., n=3. (C) Cell

cycle analysis of Nsmce2+/+ and Nsmce2-/- GS cells showing the percentages of cells in different cell

cycle phases (G0-1, S and G2-M). Data are presented as the mean ± S.E.M., n=3. (D) Representative

images (the upper part) and quantification (the lower part) of Nsmce2+/+ and Nsmce2-/- GS cells that

incorporate EdU. Cells were counterstained with DAPI to visualize nuclei. Bar = 20µm. Data are

presented as the mean ± S.E.M., n=3.

Nsmce2-/- GS cells express typical markers of undifferentiated spermatogonia and can be induced to differentiation

To validate their undifferentiated spermatogonial identity, we performed

immunocytochemistry on Nsmce2-/- and Nsmce2+/+ GS cells. Both cell populations showed

staining for SSC/progenitor cell markers PLZF, LIN28A, OCT4 and ID4 (Figure 4A),

indicating their undifferentiated state. Negative controls, i.e. omission of primary antibodies or


replacement with isotype IgGs, did not yield any staining (Figure 4A). Because our previous

studies have shown that SMC6 marks spermatogonial differentiation [16], we next

investigated whether NSMCE2 plays a role during spermatogonial differentiation. To this end,

we induced spermatogonial differentiation by adding RA to the culture medium according to

previous papers [54, 55]. In line with previous papers [51, 54], when cultured on laminin, both

Nsmce2+/+ and Nsmce2-/- GS cells started to exhibit the typical rhomboid morphology with

long pseudopod-like extensions that is characteristic for undifferentiated spermatogonia.

Treatment with RA for 3 days made them, as expected for differentiating spermatogonia [54],

gradually lose this phenotype and become more round (Figure 4B), indicative of their

differentiation. Moreover, Western blot analysis showed that exposure to RA considerably

reduced the protein levels of PLZF and OCT4, whereas that of differentiation marker STRA8

[56, 57] was markedly increased in both cell lines (Figure 4C). Overall, these results

demonstrate that Nsmce2-/- GS cells have normal undifferentiated spermatogonial

characteristics and can be induced to differentiation normally.

64 Chapter 3

Figure 4: Nsmce2-/- GS cells express typical markers of undifferentiated spermatogonia and

can be induced to differentiation. (A) The staining of SSC/progenitor cell markers in GS cells on

laminin. Bar = 10µm. (B) Representative images of GS cells (on laminin) with/without RA treatment.

Asterisks (*) indicate the long pseudopod-like extension. Bar = 25µm. (C) Western blot analysis of

PLZF, OCT4 and STRA8 proteins in GS cells with/without RA-induced differentiation. GAPDH and β-

actin are used as loading controls.

Etoposide-induced DNA damage repair occurs independently of NSMCE2 in GS cells Because we have recently shown that SMC6 physically interacts with TOP2A and that

NSMCE2 is implicated in the response to etoposide-induced topological stress and in

subsequent DNA damage repair in U2OS cells [28], we investigated whether NSMCE2

functions similarly in GS cells. To this end, we exposed Nsmce2+/+ and Nsmce2-/- GS cells to

etoposide and quantified DNA damage formation and repair marked by ɣ-H2AX (Figure 5A

and B). Albeit at a low number, ɣ-H2AX staining could be discerned in a small fraction of

cells prior to etoposide treatment. Exposure to etoposide for 3 h triggered an increase of cells

displaying ɣ-H2AX staining, indicative of increased DNA damage. However, the absence of

NSMCE2 did not lead to more ɣ-H2AX+ cells after etoposide treatment (Figure 5A and B).

Also the decrease of ɣ-H2AX+ cells, indicative of DNA repair, followed similar dynamics

between Nsmce2+/+ and Nsmce2-/- GS cells. Moreover, in contrast to U2OS cells [28], ɣ-

H2AX did not co-localize with SMC6 in GS cells, regardless of etoposide treatment (Figure

5C). Negative controls, i.e. omission of primary antibodies or replacement with isotype IgGs,

did not yield any staining (Figure 5A and C). The above data suggest that NSMCE2 is not

involved in the response to etoposide-induced topological stress in GS cells. Deprivation of NSMCE2 results in significant downregulation of Sgsm1 at the RNA but not protein level

To gain a broader perspective on the overall molecular effects of NSMCE2 removal, we

conducted a RNA-seq experiment to compare the transcriptomes of Nsmce2-/- and

Nsmce2+/+ GS cells. According to the RNA-seq data of biological triplicates from different

passages (Figure 6A), inactivation of NSMCE2 generated only 9 differentially expressed

genes (DEGs, adj.P˂0.05). Of these 9 DEGs, 6 genes showed upregulation and 3 genes

showed downregulation (Table 2 and Supplementary Table 1). These RNA-seq data thus

suggest that removal of NSMCE2 has only very minimal effects on the spermatogonial

transcriptome. Notably, one known gene, Sgsm1, showed a very significant downregulation

without NSMCE2 (Table 2) and appeared to be highly expressed in human spermatogonia


(Human Protein Atlas, v15 [58]). Nevertheless, Western blot analysis showed comparable

protein expression of SGSM1 in Nsmce2-/- and Nsmce2+/+ GS cells (Figure 6B). Subsequent

GSEA indicated alterations of gene sets such as H3K27Me3 or H3K4Me2 related to histone

methylation and cancer development (Supplementary Table 2). Nonetheless, none of the

genes within these gene sets were differentially expressed (adj.P˂0.05).

Figure 5: Etoposide-induced DNA damage formation. (A) Representative images of ɣ-H2AX

staining in Nsmce2+/+ and Nsmce2-/- GS cells after etoposide treatment. GS cells were seeded on

laminin prior to etoposide treatment. Control, without etoposide treatment; 0h, immediately post

treatment. Bar = 10µm. (B) Percentage of ɣ-H2AX+ cells after etoposide treatment. Control, without

etoposide treatment; 0h, immediately post treatment. Data are presented as the mean ± S.E.M., n=3.

(C) No co-localization of ɣ-H2AX with SMC6. Bar = 5µm.

66 Chapter 3

Figure 6: The overall effects of NSMCE2 removal on the transcriptome of GS cells. (A) The

multidimensional scaling (MDS) plot (left panel) and heat map (right panel) illustrating the minimal

transcriptome variation between the WT and KO cells. KO, Nsmce2-/- GS cells; WT, Nsmce2+/+ GS

cells. (B) Western blot analysis of SGSM1 in Nsmce2+/+ and Nsmce2-/- GS cells. β-Actin is used as a

loading control.

Discussion SSCs, a subpopulation of undifferentiated spermatogonia, are best characterized by

their capability of self-renewal to maintain sufficient numbers as well as differentiation to

mature spermatozoa, thereby maintaining life-long male fertility. SSCs hold great value in

reproductive medicine. They can reestablish spermatogenesis following transplantation into

recipient testes, and thus they can be harnessed to restore fertility of, for instance, childhood

cancer survivors who lose their germ cells due to chemotherapy or radiotherapy [59].

Moreover, in combination with genomic modification, SSCs could theoretically be employed

to cure spermatogenic failure with known genetic causes or prevent inheritance of genomic

diseases [59]. For rodent SSCs, CRISPR-Cas9 has been applied successfully for precise

genomic modification and even to correct a genetic disease in the offspring [48-50]. Apart

from clinical applications, SSCs are also of substantial utility in biomedical research, e.g.

manipulation of SSCs provides an advantageous avenue to generate various animal models

with specific genotypes and phenotypes [60].

In spite of this, the establishment of genetically modified SSC lines has been inefficient

so far, primarily due to low transfection efficiency, difficulty in monoclonal isolation and

expansion of SSCs in vitro. These technical difficulties impede the generation of transgenic

animal models and the research on genes underlying spermatogenesis. Similar to other

primary stem cells, SSCs are very refractory to prevailing transfection approaches such as

calcium phosphate precipitation and lipofection [45]. Previous studies have also indicated

that adeno-associated virus (AAV) and integration-deficient lentivirus are ineffective to fulfill

gene editing in SSCs [47]. Currently, novel electroporation devices are increasingly used to


transfect SSCs [46-50]. In our study, we employed a Neon electroporator to deliver the large

CRISPR-Cas9 vectors into GS cells. The cells harboring CRISPR-Cas9 vectors were then

enriched and subjected to clonal isolation. To achieve single cell-derived GS cell clones, we

first cultured single GS cells in a full 96-well plate, but failed to get any cell clones. Because

the initial cell density has been reported to have a big influence on subsequent GS cell

culture [45], we then attempted to pick single cell-derived patches. We found that GS cells

attached slowly to MEFs and that many single cells readily aggregated before attachment.

We therefore used laminin instead of MEF feeder cells. GS cells rapidly attach to laminin,

thereby greatly facilitating monoclonal isolation when single GS cells are plated at a low

density. Collectively, the entire optimized protocol (elaborated in ‘Materials and methods’

section) is less laborious and more efficient compared to recent reports [48-50]. Using this

optimized protocol, we successfully generated an Nsmce2-/- GS cell line lacking the NSMCE2

subunit of the SMC5/6 complex.

The SMC5/6 complex has been shown to play pivotal roles in many important

biological processes, in particular, genomic integrity maintenance and DNA damage repair

[3-5]. In yeast, all subunits of the SMC5/6 complex are indispensable [11] and also in mice

KO of SMC6 leads to embryonic lethality [32]. Consistently, deprivation of SMC5 in mouse

ES cells leads to abnormal mitotic progression, accumulation in G2 of the cell cycle and

apoptosis [27]. In our preliminary experiments, we attempted to use CRISPR-Cas9 to knock-

out Smc5 and Smc6 in GS cells. However, we observed cell death of more than 90% of the

transfected cells, which may indicate that the deletion of one of these genes is lethal to GS

cells. In contrast, Nsmce2 KO did not influence the survival of GS cells, and we eventually

generated an Nsmce2-/- GS cell line. NSMCE2 depletion can destabilize the SMC5/6

complex, characterized by the reduced expression of SMC5 and/or SMC6 [61, 62]. Using

Western blot analysis, we showed that the protein levels of SMC5 and SMC6 were not

affected by NSMCE2 depletion in GS cells. Indeed, the stable presence of SMC5 and SMC6

does not mean that the whole SMC5/6 complex remains stable. However, combined with the

fact that we did not detect any other significant phenotype, it does suggest that NSMCE2 is

not essential for the SMC5/6 stability and function in GS cells.

We have recently found that the removal of NSMCE2 in U2OS cells generated

significant differences in phenotypes, including slower cell growth, accumulation of cells in

G0-1 of the cell cycle, as well as a decreased plating efficiency [28]. In line with these, an

earlier study in human MCF-7 breast cancer cells showed that knockdown of Nsmce2

resulted in slower cell growth and impaired G1-S transition. Ectopic expression of the full-

length NSMCE2, but not its SUMO ligase-inactive mutant, rescued the reduced cell growth,

68 Chapter 3

implying that the normal growth of these breast cancer cells requires the NSMCE2 SUMO

ligase function [63]. However, consistent with a study conducted in chicken DT40 cells [61],

we found that the viability and growth of GS cells was not influenced by the removal of

NSMCE2. Nsmce2-/- GS cells showed similar doubling time, EdU incorporation and cell cycle

progression as their Nsmce2+/+ counterparts. Furthermore, GS cells showed normal cellular

morphology without nuclear abnormalities, such as micronuclei or nucleoplasmic bridges that

are observed in mouse and human Nsmce2-deficient fibroblasts [31, 62]. These disparities

imply cell type-specific roles of NSMCE2.

We have recently found that SMC6 protein expression specifically marks differentiating

spermatogonia [16], the spermatogonial subpopulation irreversibly committed toward meiosis.

However, the exact role of NSMCE2 during spermatogonial differentiation has not been

elucidated. We therefore investigated whether absence of NSMCE2 would influence the

capacity of GS cells to differentiate. GS cells can be induced to differentiation by adding RA

to the culture medium [54, 55]. RA-induced differentiation will lead to downregulation of

SSC/progenitor cell markers (e.g. PLZF and OCT4), upregulation of differentiation markers

(e.g. STRA8), reduced self-renewal and increased apoptosis and eventually to a decline in

the total cell number [54, 55, 64]. Before RA-induced differentiation, Nsmce2-/- GS cells

exhibited routine expression profiles characteristic for undifferentiated spermatogonia. Here,

we found that Nsmce2-/- GS cells could be normally induced to differentiate, exhibiting similar

morphology and characteristics as their Nsmce2+/+ counterparts. Hence, NSMCE2 does not

seem to be involved in in vitro spermatogonial proliferation or differentiation.

NSMCE2 is extensively reported to be crucial for the response to DNA damage, mostly

in yeast [9-11, 61, 65, 66]. However, recent studies performed in multiple mouse and human

cell types have indicated that NSMCE2 is redundant for the repair of ionizing radiation (IR)-

induced DNA damage in mammalian cells [28, 31, 67]. Indeed, we have recently shown that

also in U2OS cells, NSMCE2 is redundant for the repair of double-strand breaks (DSBs)

induced by IR [28]. Nonetheless, in the same paper, we showed that CRISPR-Cas9-

mediated removal of NSMCE2 led to increased sensitivity to etoposide. Etoposide is a

cytotoxic agent that, by forming a complex with DNA and topoisomerase II, causes

replication-induced topological stress and DSBs at replication forks [68]. Surprisingly, here

we found that removal of NSMCE2 in GS cells did not cause increased sensitivity to

etoposide. Moreover, etoposide-induced DSBs marked by ɣ-H2AX were repaired efficiently

in both Nsmce2+/+ and Nsmce2-/- GS cells. Also in contrast to U2OS cells [28], etoposide-

induced ɣ-H2AX in GS cells did not co-localize with SMC6. It is thus plausible that GS cells

hold different mechanisms to resolve topological stress than somatic cells, which might not

rely on NSMCE2 or even the SMC5/6 complex.


To gain a broader perspective on the overall molecular effects of NSMCE2 removal, we

finally conducted a whole transcriptome RNA-seq analysis for Nsmce2+/+ and Nsmce2-/- GS

cells. We previously hypothesized that, by altering chromatin structure, the SMC5/6 complex

would influence spermatogonial gene transcription [15, 16]. GSEA did indicate the

downregulation of gene sets involved in chromatin regulation, such as H3K27Me3 or

H3K4Me2 related to histone methylation and cancer development, suggesting that chromatin

architecture or function is somehow affected by the absence of NSMCE2. However, removal

of NSMCE2 only led to 9 DEGs, of which none were present in these gene sets. Of the 9

DEGs, Sgsm1 showed a very significant downregulation without NSMCE2. SGSM1, as well

as its two paralogs SGSM2 and SGSM3, all consist of RUN and TBC motifs and have been

reported to orchestrate small G protein-mediated signaling transduction. Unlike SGSM2 and

SGSM3, which show ubiquitous expression in a wide range of tissues, SGSM1 is primarily

expressed in human brain, heart and testes [69]. Moreover, in human testes, SGSM1 is

highly expressed in spermatogonia (Human Protein Atlas, v15 [58]). In our present study, we

also demonstrated the high protein level of SGSM1 in GS cells. However, while removal of

NSMCE2 significantly downregulated Sgsm1 transcripts, its protein level was not affected.

Perhaps the SGSM1 protein level was somehow stabilized in the absence of NSMCE2.

Alternatively, assuming that SGSM1 protein stability is not affected by NSMCE2, the

detected lower Sgsm1 mRNA level was still sufficient to ensure the wildtype protein level.

Nevertheless, the detected Sgsm1 downregulation in our study and localization in human

spermatogonia indicate an unknown role of this protein in GS cells. To acquire more

knowledge about this, loss-of-function studies for SGSM1 could be performed in the future.

The absence of evident phenotypes and the minimal transcriptome variation caused by

NSMCE2 removal raise the question what the exact role of NSMCE2 in GS cells is. Recently,

a case report described 2 female patients with heterozygous frameshift mutations in Nsmce2

that result in decreased expression of NSMCE2. These patients exhibited serious

phenotypes like primordial dwarfism, extreme insulin resistance and, notably, primary ovarian

failure [62]. Given that NSMCE2 is required for yeast meiosis [70], the possibility remains

that further downstream steps beyond spermatogonial proliferation and differentiation require

the function of NSMCE2. On the other hand, the specific expression of SMC6 in

differentiating spermatogonia [16], the stable presence of SMC5 and SMC6 without NSMCE2

and the observed lethality of GS cells after transfection with CRISPR-Cas9 plasmids

targeting Smc5/6 all suggest that the SMC5/6 complex plays important roles in

spermatogonia, but that this spermatogonial function of SMC5/6 is not affected by NSMCE2

removal.

70 Chapter 3

Male germline stem cells are responsible for the lifelong daily production of millions of

sperm and the transmission of genetic information to the offspring. Decades of studies have

well demonstrated that this unique adult stem cell population is largely distinct from somatic

cells in terms of cellular activities, cell fate commitment, developmental plasticity, chromatin

architecture and remodeling as well as (epi)genetic features [71-73]. They also hold unique

mechanisms to maintain genomic stability [74], which can partially explain the divergent roles

of NSMCE2 and SMC5/6 in GS cells. It has been known that the SMC5/6 complex, including

NSMCE2, is essential for genome integrity maintenance in somatic cells, demonstrated by,

e.g. the embryonic lethality of SMC6 or NSMCE2 KO [31, 32] and the role of NSMCE2 in the

prevention of cancer and aging in adult mice [31]. Nonetheless, the SMC5/6 complex in male

germline stem cells seems to function normally without NSMCE2. Hence, how germline stem

cells safeguard genomic integrity, and the role of SMC5/6 herein, remain to be further

investigated, especially given the current technical development such as CRISPR-Cas9 and

the potential clinical application of these cells, for instance, in fertility preservation, curing

spermatogenic failure or preventing transmission of genetic diseases [59].

Supplementary data This is linked to the online version of the paper at http://dx.doi.org/10.1530/REP-17-

0173.

Declaration of interest The authors declare that there is no conflict of interest that could be perceived as

prejudicing the impartiality of the research reported.

Funding This study has been supported by an AMC Fellowship, the People Programme (Marie

Curie Actions) of the European Union’s Seventh Framework Programme (CIG 293765) to

Geert Hamer and the China Scholarship Counsel (CSC) number 201306300081 to Yi Zheng.

Acknowledgements The authors thank Ieva Masliukaite and Dr Kaijun Liu, for assistance with FACS

analysis and sorting, respectively, and Saskia K M van Daalen for assistance with picking

single cell-derived patches. In addition, they thank Oscar Fernandez-Capetillo for kindly

providing the NSMCE2 antibody and MEF extracts.


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Chapter 4

On the increasing sensitivity of differentiating spermatogonia to DNA damage

Yi Zheng

Aldo Jongejan

Callista L. Mulder

Saskia K.M. van Daalen

Sebastiaan Mastenbroek

Grace Hwang

Philip W. Jordan

Sjoerd Repping

Geert Hamer

Submitted

78 Chapter 4

Abstract Lifelong mammalian male fertility is maintained through an intricate balance between

spermatogonial proliferation and differentiation. DNA damage in spermatogonia, for instance

caused by chemo- or radiotherapy, can induce cell cycle arrest or germ cell apoptosis,

possibly resulting in male infertility. Because genetic aberrations in spermatogonia can be

transmitted to future generations, these cells are generally more radiosensitive, and hence

more prone to undergo apoptosis, than somatic cells. Among spermatogonial subtypes the

response to DNA damage is differentially modulated; undifferentiated spermatogonia are

relatively radio-resistant, whereas differentiating spermatogonia are very radiosensitive. To

investigate the molecular mechanisms underlying this difference, we used an in vitro system

consisting of primary cultured undifferentiated mouse spermatogonia that can be induced to

differentiate. Using RNA-sequencing analysis, we analyzed the response of undifferentiated

and differentiating spermatogonia to ionizing radiation (IR). At the RNA level, both

undifferentiated and differentiating spermatogonia showed a very similar response to IR.

However, at the protein level, undifferentiated spermatogonia showed a much stronger

upregulation of p53 in response to IR than differentiating spermatogonia. Our results suggest

that the difference in radiosensitivity between undifferentiated and differentiating

spermatogonia is largely caused by properties, like chromatin architecture, proliferation

activity, protein content or post-translational modifications that are already induced upon

differentiation, rather than an alternative gene expression pattern in response to irradiation in

both cell types.

Keywords Spermatogonia; Differentiation; Spermatogonial markers; DNA damage response;

Transcriptome

Transcriptome of irradiated spermatogonia 79

Introduction Spermatogenesis is an intricate process that takes place in the seminiferous tubules

within the testis. In mammals, the entire process of spermatogenesis is comprised of three

consecutive phases: a mitotic phase (spermatogonial proliferation and differentiation), a

meiotic phase (spermatocyte meiotic divisions to generate haploid spermatids) and

spermiogenesis (elongation and maturation of spermatids) [1]. For continuous

spermatogenesis spermatogonial stem cells (SSCs) are essential. SSCs can be defined as a

subpopulation of undifferentiated spermatogonia able to generate and maintain donor-

derived spermatogenesis when transplanted into infertile recipient testes [2, 3]. Continuous

spermatogenesis requires a constant balance between SSC self-renewal, proliferation and

differentiation [1]. Within the seminiferous tubules, spermatogenesis occurs in an

orchestrated spatio-temporal fashion in which specific germ cell types are grouped in specific

stages of the seminiferous epithelium. The undifferentiated spermatogonia may divide freely

during all of these epithelial stages. In contrast, differentiating spermatogonia are irreversibly

committed towards meiosis and their subsequent divisions are strictly dictated by the

epithelial stage in which they are present [4].

To guarantee the continuous generation of healthy and functional sperm, DNA integrity

is constantly being monitored during spermatogenesis [1]. DNA damage, for instance caused

by gonadotoxic chemicals or ionizing radiation (IR), can result in gene mutations or

chromosomal aberrations and often leads to spermatogenic arrest and subsequent male

infertility. Indeed, in the human, treatment with chemo- or radiotherapy in adult males often

results in impaired fertility [5]. Because genetic aberrations can potentially be transmitted to

future generations when DNA damage checkpoints mechanisms fail during spermatogenesis,

spermatogenic cells are thought to be generally more radiosensitive than somatic cells [6].

Because genetic aberrations in SSCs have the potential to result in lifelong generation

of mutated sperm, one might expect that the SSCs, when compared to all other

spermatogonia, are most prone to undergo apoptosis in response to IR. However, this

appears not to be the case. It turns out that differentiating spermatogonia are actually much

more radiosensitive and show a stronger apoptotic response [7]. Among the undifferentiated

spermatogonia, the self-renewing SSCs are most resistant to DNA damage induced by either

the alkylating agent busulfan or IR [8-10]. Evidently, while damaged differentiating

spermatogonia are more easily sacrificed, preservation of SSCs, and thus long-term fertility,

seems to outweigh a certain risk of mutated offspring.

What determines the differences in radiosensitivity among spermatogonial subtypes is

currently unknown. One process that characterizes spermatogonial differentiation is the

80 Chapter 4

condensation of repetitive sequences surrounding the centromeres in pericentromeric

heterochromatin domains. From spermatogonial differentiation and onwards through meiosis,

these heterochromatic regions are marked by the presence of SMC6, a major subunit of the

SMC5/6 complex known to be involved in the repair of DNA double-strand breaks (DSBs) [11,

12]. Within these regions the SMC5/6 complex has been postulated to inhibit aberrant

homology-driven recombinational repair of DSBs that would otherwise easily occur between

repetitive sequences [12]. Hence, this changed chromatin architecture in differentiating

spermatogonia may profoundly influence their response to DNA damage.

Apart from chromatin architecture, also several DNA damage response proteins are

differentially regulated during spermatogonial differentiation. For instance, phosphorylated

histone H2AX (ɣ-H2AX), usually marking DSBs, has been described to increase with

spermatogonial differentiation [13, 14] and is highly expressed in intermediate and B

spermatogonia [15]. The DNA damage response protein p53 has been found to be induced

in all spermatogonia by irradiation, but knockout of p53 seems to predominantly affect the

apoptotic response of the undifferentiated spermatogonia [16-18]. Nevertheless,

transplantation assays of mutated SSCs revealed that deficiency in a specific p53 pathway

(Trp53-Trp53inp1-Tnfrsf10b) actually increased survival of SSCs after irradiation [19]. The

same study also reported that the apoptosis-inducing protein BBC3 was specifically active in

differentiating spermatogonia after irradiation [19].

To investigate the relation between the IR-induced DNA damage response and

spermatogonial differentiation, we used an established culture system for undifferentiated

mouse spermatogonia [20, 21]. In this culture system, primary isolated mouse SSCs, then

referred to as male germline stem (GS) cells, can propagate in vitro for years without losing

SSC properties [21]. GS cells can also be induced to differentiate by adding retinoic acid (RA)

to the culture medium [22, 23]. Moreover, by way of RNA-sequencing (RNA-seq), the

transcriptome of RA-induced differentiating GS cells was reported recently [23]. To gain

insights into the differential DNA damage responses of undifferentiated and differentiating

spermatogonia, we investigated the transcriptomes of irradiated and non-irradiated GS cells

with or without RA treatment.

Materials and methods Animals

Neonatal (4-5 d.p.p) DBA/2J male mice were used for GS cell isolation, and adult (~8

weeks) C57BL/6J male mice were used for irradiation and immuno-histochemical analysis.

For histological analysis on neonatal testis sections, 8 d.p.p old C57BL/6J male mice were


used. All animal procedures were in accordance with and approved by the animal ethical

committee of the Academic Medical Center, University of Amsterdam or in accordance with

the National Institutes of Health and US Department of Agriculture criteria approved by the

Institutional Animal Care and Use Committees of Johns Hopkins University.

GS cell culture A mouse GS cell line was established as previously reported [20, 24]. Briefly, testes

were harvested from neonatal DBA/2J male mice, and after removing the tunica albuginea,

testicular tissues were mechanically dissociated and subjected to a collagenase-trypsin

dissociation to obtain a single-cell suspension. Germ cells were enriched by an overnight

differential plating and cultured in a medium mainly composed of StemPro-34 SFM medium

(Thermo Fisher Scientific), StemPro-34 Supplement (Thermo Fisher Scientific), 1% fetal

bovine serum (FBS), recombinant human GDNF (15 ng/ml, Peprotech), recombinant human

bFGF (10 ng/ml, Peprotech), as well as other components as previously reported [20]. The

cells were cultured on mitotically inactivated mouse embryonic fibroblasts (MEFs) since the

third passage, and were refreshed every 2-3 days and passaged every 5-7 days at a ratio of

1:4-6. The cells were maintained at 37°C in an atmosphere of 5% CO2 in air.

RA treatment Before RA treatment, GS cells cultured on MEFs were transferred to laminin (20 µg/ml,

Sigma-Aldrich)-coated wells. On the next day, GS cells were treated with 2µM all-trans-RA

(Sigma-Aldrich) in culture medium for 48-72 hours. In control groups, vehicle (0.1% ethanol

in medium) was applied to the cells.

Ionizing irradiation (IR) Before IR treatment, GS cells cultured on MEFs were transferred to laminin (20 µg/ml,

Sigma-Aldrich)-coated wells. On the next day, GS cells were subjected to 1 Gy of IR emitted

by a 137Cs source (95% β-emission). To prepare irradiated mice, adult C57BL/6J male mice

were exposed to a whole-body IR (1 Gy).

Quantitative-real time PCR (Q-PCR) Total RNA was extracted from GS cells which had been subjected to RA treatment for

2 days or IR 3 hours before, using ISOLATE II RNA Mini Kit (Bioline) and following the

protocol provided by the manufacturer. After treatment with DNase (Qiagen) and tests for

82 Chapter 4

genomic DNA free, RNA samples were reversely transcribed, using SensiFAST cDNA

Synthesis Kit (Bioline). The synthesized cDNA was then used for Q-PCR reactions, using the

Roche LightCycler 480 platform (the 384-well plate format). The Q-PCR reaction was

performed in a 10µl volume system including 2× LightCycler 480 SYBR Green I Master

(Roche). Ppt2 and Mtg1 were used as reference genes, and the data were analyzed using

the -ΔΔCt method. Data were presented as the mean ± standard error of mean (SEM) of 3

independent experiments (n=3). Differences between groups were assessed using the

Student’s t-test. P˂0.05 was considered statistically significant and P˂0.01 was considered

extremely significant. The primers for Q-PCR analysis are listed in Table 1.

Table 1: Primer sequences for Q-PCR analysis.

Gene Forward primer Reverse primer Product size (bp)

Ppt2 CCTGCTGGACTATATCAATGAGAC TCTCGGAACCCTTGTACCTG 114

Mtg1 CACGATGTAGCACGCTGGTT GGGTTTCGACCTGAAAATGGG 128

Plzf TCTCGGAACCCTTGTACCTG ACCGAAAGAGGTGGAGACTGA 132

Oct4 CACGAGTGGAAAGCAACTCA CTTCTGCAGGGCTTTCATGT 125

Stra8 GGAAGGCAGTTTACTCCCAGTC GATTCCCATCTTGCAGGTTGA 144

Clu CAGTTCCCAGACGTGGATT GGGCAGGATTGTTGGTTG 157

Ntrk3 TTTGGGGTGTCCATAGCAG AGCCACAGGACCCTTCATT 120

Wnt16 CTTCCCATCAGAAACACCACA GCGGCAGTCCACAGACATTA 114

Agtr2 GAAGAACAGAATTACCCGTGAC AGGGAAGCCAGCAAATGA 80

Bmp2 ATCTGTACCGCAGGCACTC ACGGCTTCTTCGTGATGG 112

Sarm1 CAAGGAGATTGTGACTGCTTTA GGTACTCATGGGACCATTTGA 143

Insm1 GGTGTTCCCCTGCAAGTACT CTATTCTCAGACGGGTGGC 90

Slit2 ATGGAGAACAGAATCAGCACC TCGCAGTCCCGAGAAACA 124

Adgrg1 AGCCAAGTCCTGGGTGAGA TTGATGCCGGGTCTTCAA 154

Myog GTCCCAACCCAGGAGATCA AACAGACATATCCTCCACCGT 107

Pax6 AACAGACATATCCTCCACCGT TATCATAACTCCGCCCATTC 135

Vsx2 CACTACCCAGATGTCTACGCC CACTTCTCCCTCTTCCTCCAC 116

Adora1 AACCCAGCATCCTCATCTACA GTGGTCGTTCCAAATCTTCA 125

Sfrp2 GTGGTCGTTCCAAATCTTCA GCTCTTTGTCTCCAGGATGAT 130

Irf1 AATGCGGATGAGACCCTG ATGTCCCAGCCGTGCTTA 129

Pdx1 AATCCACCAAAGCTCACGC CGGGTCCTCTTGTTTTCCTC 82

Bbc3 GAGCGGCGGAGACAAGAAGA ATCCCTGGGTAAGGGGAGGA 96

Plk2 ATGTGGAACCCCAAATTATCTC GGTCTTCCTAGCAGCATCGTAT 114

Trp53inp1 GACACCAGTGATTCCTGCTTC GGACTTGTTTCCACCTTGATAG 122


Ddias TGTCCTTGAAAGTGGCAGAA GTGTAAACCAGTGGCCGTAA 98

Ccng1 ATAATGGCCTCAGAATGACTGC CCAAGATGCTTCGCCTGTAC 160

Klhl42 CAATCCCATCACCAACGAG TAGAAACAGCCTGCCCACC 116

Psrc1 TGCCCACCGTGAGTTCTT GTGGGTGATTCCTTCTTTATGC 139

Eda2r ACTTGTGCTGTCATCAATCGG CGTGTCTTTCGGTAGAACCTG 98

Pdrg1 GGAAGGAGCCAAGGTGAAGT CCTCGGCCAACTCCTCTAC 117

Sesn2 AACTACATCCACTGCGTCTTTG CATCCTACGGGTCGTCTTCT 135

Slc2a10 TACTTGTTCCTGAAACCAAAGG TCCAGGCGATGGTACTGAA 121

Far2 TTATTGGAACACCGTCAGCC CAGCATTCTGGGTTTCCTTC 85

Western blot Proteins were extracted from the cells and quantified with Qubit Protein Assay Kit

(Thermo Fisher Scientific). Then Western blot analysis was performed as reported previously

[12, 24, 25], using the LI-COR Odyssey imaging system (LI-COR Biosciences). The primary

antibodies used were mouse anti-PLZF (1:100; D-9, Santa Cruz Biotechnology), mouse anti-

OCT4 (1:200; C-10, Santa Cruz Biotechnology), rabbit anti-STRA8 (1:500; ab49602, Abcam),

rabbit anti-p53 (1:100; FL-393, Santa Cruz Biotechnology), rabbit anti-GAPDH (1:400; FL-

335, Santa Cruz Biotechnology) and mouse anti-β-actin (1:5,000; A1978, Sigma-Aldrich). For

quantification of the relative p53 expression, p53 band intensity is divided by that of GAPDH.

Data were presented as the mean ± SEM of 4 independent experiments (n=4). Differences

among groups were assessed using the one-way ANOVA followed by LSD test. P˂0.05 was

considered statistically significant and P˂0.01 was considered extremely significant.

Immunocytochemistry (ICC) and immunohistochemistry (IHC) For ICC, GS cells were grown on laminin (20 µg/ml, Sigma-Aldrich)-coated glass

coverslips in 24-well plates. In case of IR treatment, GS cells were fixed in 4%

paraformaldehyde (PFA) at 3 hours post IR. Cells were permeabilized and blocked as

previously described [24, 25], followed by 4°C overnight incubation with the following primary

antibodies: mouse anti-PLZF (1:50; D-9, Santa Cruz Biotechnology), mouse anti-OCT4 (1:50;

C-10, Santa Cruz Biotechnology), rabbit anti-LIN28A (1:1,000; ab46020, Abcam), rabbit anti-

ID4 (1:100; M106, CalBioreagents), mouse anti-ɣ-H2AX (1:20,000; 05-636, Merck Millipore)

and guinea pig anti-SMC6 (1:400; custom made, peptide: KRPRQEELEDFDKDGDEDE [24,

25]). Replacing primary antibodies with phosphate buffered saline (PBS) containing 0.5%

bovine serum albumin (BSA) was used as negative controls. On the next day, the cells were

washed and incubated with the corresponding host-specific secondary antibodies (Alexa

84 Chapter 4

Fluor 488 or 555, 1:1,000; Thermo Fisher Scientific), and counterstained with DAPI. The cells

were mounted on glass slides with the Prolong Gold anti-fade mountant (Thermo Fisher

Scientific) and later visualized under the microscope. Microscopy was performed as

previously described [24].

For IHC, testes were collected from normal adult and neonatal mice, or the

experimental and control mice at 3 hours post IR, fixed in diluted Bouin’s solution or 4% PFA,

and embedded in paraffin. Testis sections were sliced at a thickness of 5µm. After

deparaffinization and rehydration, testis sections were alternatively subjected to microwave-

mediated antigen retrieval in sodium citrate buffer (0.01M, pH 6.0), followed by blocking in

Super Block (ScyTek Laboratories) or PBS with 1% BSA and 0.1% Tween-20, for 1 hour at

room temperature (RT). Then the sections were incubated with primary antibodies diluted in

Normal antibody Diluent (ImmunoLogic) or PBS containing 0.5% BSA and 0.1% Tween-20,

at 4°C overnight. The primary antibodies used were rabbit anti-WNT16 (1:100; H-96, Santa

Cruz Biotechnology), rabbit anti-SLIT2 (1:80; ab7665, Abcam), rabbit anti-PDX1 (1:1,000;

ab47267, Abcam), rabbit anti-ADORA1 (1:600; ab82477, Abcam), rabbit anti-p53 (1:100; FL-

393, Santa Cruz Biotechnology), rabbit anti-BBC3 (1:250; ab9643, Abcam), rabbit anti-PLK2

(1:100; H-90, Santa Cruz Biotechnology), rabbit anti-PDRG1 (1:100; 16968-1-AP,

Proteintech) and rabbit anti-SESN2 (1:100; 10795-1-AP, Proteintech). The isotype rabbit IgG

was used in negative controls. On the next day, the sections were washed and incubated

with Powervision Poly-HRP-Anti-mouse/rabbit/rat secondary antibody (ImmunoLogic) for 1

hour at RT. After washing, the sections were stained with diaminobenzidine (DAB) and

counterstained with hematoxylin. Then the slides were dehydrated, embedded in Entellan

(Merck Millipore), and visualized under the microscope. Microscopy was performed as

previously described [12].

RNA-sequencing (RNA-seq) analysis Total RNA was extracted from GS cells which had been subjected to RA treatment for

2 days and/or IR 3 hours before, using ISOLATE II RNA Mini Kit (Bioline) and following the

protocol provided by the manufacturer. After treatment with DNase (Qiagen) and tests for

genomic DNA-free, RNA samples from 3 independent experiments (biological triplicates)

were sent to BGI Tech Solutions (HongKong) where libraries were constructed (TruSeq,

160bp) and paired-end sequencing was performed (101bp, Illumina HiSeq 4000). RNA-seq

analysis was conducted as described previously [24]. In brief, clean reads were subjected to

quality control, and then aligned to UCSC mm10 GRCm38.p4 GTF using HISAT2 (v2.0.4)

[26]. Counts were obtained using HTSeq (v0.6.1) [27]. Count tables were made and all

genes without counts in any of the samples were removed, whilst genes with more than 1


count-per-million reads (CPM) in 2 of the samples were kept. Genes were re-annotated using

biomaRt and mm10 of Ensembl. Count data were transformed to log2-counts per million

(logCPM) using voom, estimating the mean-variance relationship. Differential expression was

assessed using a moderated t-test and the linear model framework from the limma package.

Benjamini-Hochberg false discovery rate was used to correct for multiple testing of the

resulting p-values. The entire analysis was performed using R v3.2.2 and Bioconductor v3.0

[28, 29].

Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG)

analyses were conducted using DAVID bioinformatics resources 6.8 [30, 31], and Gene Set

Enrichment Analysis (GSEA) [32, 33] was used to analyze differentially expressed gene sets.

Accession numbers

All sequencing data have been submitted to NCBI (SRA) and will be available under

the accession number: SUB2838515 (BioProject ID PRJNA392875).

Results Culture, differentiation and irradiation of GS cells

First, we established a GS cell line according to a previously published and well-

demonstrated protocol [20]. GS cells can be maintained on either a feeder layer of mouse

embryonic fibroblasts (MEFs) or in laminin-coated wells [34]. Like previously described [34],

GS cells cultured on MEFs formed typical grape/morula-like colonies, whereas they started

to form chain-like structures when seeded in laminin-coated wells (Figure 1A). The cultured

GS cells were positive for putative SSC/progenitor markers PLZF, LIN28A, OCT4 and ID4

(Figure 1B), indicative of their undifferentiated spermatogonial properties.

To induce GS cell differentiation, we transferred them to laminin-coated wells and

added RA to the culture medium. Multiple studies have shown that RA can drive

spermatogonial differentiation both in vivo and in vitro [22, 23, 35-37]. Consistent with these

reports, exposure to RA for 3 days made the GS cells gradually lose the structure

characteristic for undifferentiated spermatogonia and display an enlarged cell size and round

morphology with clear cellular boundaries (Figure 1C), indicative of spermatogonial

differentiation. Q-PCR and Western blot analyses showed substantial downregulation of the

SSC self-renewal genes Plzf and Oct4, while RNA and protein levels of the differentiation

marker Stra8 markedly increased after 3 days of RA exposure (Figure 1D, E). These results

demonstrate that in vitro spermatogonial differentiation was successfully induced in our

culture system.

86 Chapter 4

To investigate their response to induced DSBs, GS cells on laminin were subjected to

1Gy of IR, a dose that causes substantial DNA damage but does not necessarily kill these

cells [7]. To visualize the DSBs, we stained the cells with ɣ-H2AX, a widely used DSB marker.

Before IR, the cells displayed a weak ɣ-H2AX staining. Three hours post IR, clear nuclear ɣ-

H2AX foci were observed (Figure 1F), demonstrating the induction of DSBs. In line with our

recent findings showing that, in contrast to somatic cells [25], SMC6 does not co-localize with

ɣ-H2AX at etoposide-induced DSBs in spermatogonia [24], spermatogonial SMC6 did not co-

localize with IR-induced ɣ-H2AX foci (Figure 1F, right panel).


Figure 1: In vitro culture, differentiation and irradiation of GS cells. (A) Representative images of

GS cells cultured on MEFs or laminin. Bar = 100µm. (B) PLZF, LIN28A, OCT4 and ID4 staining of GS

cells on laminin. NC: omitting primary antibodies. Bar = 10µm. (C) Representative images of GS cells

(on laminin) with/without exposure to RA. Asterisks (*) refer to the long extensions typical for

undifferentiated spermatogonia. Bar = 25µm. (D) Q-PCR analysis of Plzf, Oct4 and Stra8 expression

in GS cells with/without exposure to RA. Data are presented as the mean ± standard error of mean

(SEM), n=3. *: P˂0.05; **: P˂0.01. (E) Western blot analysis of PLZF, OCT4 and STRA8 expression in

GS cells with/without exposure to RA. β-actin or GAPDH is used as the loading control. (F) ɣ-H2AX

and SMC6 staining in GS cells with/without IR. NC: omitting primary antibodies. Left panel: bar =

10µm; right panel: bar = 5µm.

Spermatogonial differentiation induces a high transcriptomic change Differentiating spermatogonia are known to be much more radiosensitive than

undifferentiated spermatogonia [7]. To study the underlying molecular mechanism causing

this increased radiosensitivity, we performed a whole transcriptomic RNA-seq analysis for

GS cells with/without RA and/or IR treatment (Figure 2A). A multidimensional scaling (MDS)

plot and heat map showed that RA treatment triggered a much larger transcriptomic change

than IR (Figure 2B). 1,748 upregulated and 966 downregulated differentially expressed

genes (DEGs, adj.P<0.05 and fold change>2) resulted from RA treatment (Table S1).

Upregulated DEGs included markers for differentiating spermatogonia and (pre-)meiotic

germ cells such as c-Kit, Stra8, Rec8, Prdm9. Likewise, well-known marker genes for

SSCs/progenitors, e.g. Plzf, Oct4, Gfra1, c-Ret, Nanos2, Nanos3, Lin28A, Id4 and Pax7,

showed a significant downregulation (Table S1). Gene Ontology (GO) analysis demonstrated

that RA-induced upregulated genes were involved in cell adhesion, differentiation and

response to retinoic acid. Downregulated genes were mainly related to transcription,

proliferation, regulation of cell death and apoptosis (Figure 2C, Table S5). Kyoto

Encyclopedia of Genes and Genomes (KEGG) pathway analysis revealed that

differentiation-upregulated genes were enriched with pathways mediating extracellular matrix

(ECM)-receptor interaction, PI3K-Akt and Rap1 signaling, whereas downregulated genes

were primarily related to stem cell pluripotency regulation and cancer pathways (Figure 2D,

Table S6). We further conducted a Q-PCR analysis for 16 RA-induced DEGs representative

of the aforementioned cellular processes (Figure 2E). For most genes the Q-PCR results

were in line with the RNA-seq data, confirming the validity of the RNA-seq analysis.

88 Chapter 4

Figure 2: Transcriptomic profiles of GS cells in response to RA and IR treatment. (A) A

schematic overview of the experimental design. (B) The MDS plot (left panel) and heat map (right

panel) showing the expression profiles of 4 experimental groups of cells (-RA-IR, +RA-IR, -RA+IR,

+RA+IR). (C) GO term enrichment analysis of RA-induced up- and downregulated genes. (D) The

representative enriched pathways shown by KEGG analysis. (E) Q-PCR analysis of a panel of RA-

induced DEGs. Data are presented as the mean ± SEM, n=3. *: P˂0.05; **: P˂0.01. (F) Q-PCR

analysis of a panel of IR-induced DEGs. Data are presented as the mean ± SEM, n=3. *: P˂0.05.


Undifferentiated and differentiating spermatogonia display similar transcriptomic changes in response to IR

Also IR yielded transcriptomic variations in both undifferentiated and differentiating

spermatogonia (Figure 2B). Between the -RA-IR and -RA+IR cell populations, 31 and 8

DEGs (adj.P<0.05 and fold change>2) were up- and downregulated, respectively (Table S2).

KEGG analysis disclosed that IR-upregulated genes, including Bbc3, Gtse1, Ccng1, Cdkn1a

and Sesn2, were predominantly related to the p53 signaling pathway. To validate the RNA-

seq data, we used Q-PCR to quantify the expression of 12 IR-induced DEGs that are

implicated in the p53 signaling pathway, cell cycle arrest or apoptosis. The Q-PCR results

were in line with the RNA-seq data for most genes (Figure 2F).

Next, we investigated IR-induced DEGs in differentiating spermatogonia. Between the

+RA-IR and +RA+IR cell populations, there were 20 DEGs (adj.P<0.05 and fold change>2)

that all showed upregulation (Table S3). When comparing these 20 DEGs from the +RA-IR

vs +RA+IR comparison to the 39 DEGs from the -RA-IR vs -RA+IR comparison, we found

that 15 DEGs, including well-known IR response genes such as Bbc3, Gtse1 and Ccng1,

were upregulated by IR in both undifferentiated and differentiating spermatogonia. There

were 24 DEGs only belonging to the -RA-IR vs -RA+IR comparison, and 5 DEGs only

belonging to the +RA-IR vs +RA+IR comparison. One of the reasons that these 24 and 5

specific DEGs were found when simply comparing the (-RA-IR vs -RA+IR) to the (+RA-IR vs

+RA+IR) group is the threshold used in our study (adj.P<0.05 and fold change>2). Genes

can be found differentially expressed only in one group while being just below the threshold

in the other group. Moreover, expression of these DEGs can be influenced by RA-induced

differentiation independent of IR. To overcome these problems, we also directly compared

the -RA+IR and +RA+IR cell populations after correction for their respective baseline levels

of gene expression without exposure to IR (i.e. -RA-IR and +RA-IR, respectively). In this way,

the gene expression affected by differentiation independent of IR is separated from that in

response to IR. Apart from the genes affected by RA treatment independent of IR, no

significant DEGs (adj.P<0.05 and fold change>2) were detected between the -RA+IR and

+RA+IR cell populations (Table S4). Hence, undifferentiated and differentiating

spermatogonia do not show significantly differential gene expression in response to IR.

In order to find significant up- or downregulation of specific pathways or gene sets, we

performed a Gene Set Enrichment Analysis (GSEA). In contrast to GO or KEGG analysis,

GSEA is not based on observed DEGs but instead takes into account all minor variations at

90 Chapter 4

the whole transcriptome level. GSEA showed that mesenchymal transition pathways went up

in +RA+IR cell populations compared to their -RA+IR counterparts (Table S7).

Undifferentiated spermatogonia display a more robust p53 induction in response to IR Because the IR-upregulated genes were predominantly related to the p53 signaling

pathway and p53 is known to be upregulated in spermatogonia in response to IR [17], we

performed a Western blot analysis to investigate IR-induced p53 in both undifferentiated and

differentiating spermatogonia. Interestingly, we found that undifferentiated spermatogonia

displayed a much stronger induction of p53 than differentiating spermatogonia at 3 hours

post IR (Figure 3).

Figure 3: Western blot analysis of p53 protein levels in -IR-RA, -IR+RA, +IR-RA, +IR+RA cell

groups. GAPDH is used as a loading control.

p53 band intensity is divided by that of GAPDH.

Data are presented as the mean ± SEM, n=4. *:

P˂0.05; **: P˂0.01.

Protein localization of differentiation-induced genes in the testis To study the protein localization of differentiation-induced DEGs, we performed

immunohistochemistry (IHC) on testis sections. We examined the protein localization

patterns of 4 RA-induced DEGs (Wnt16, Slit2, Pdx1, Adora1) for which working antibodies

were available in adult mouse testes (Figure 4A-D).

In accordance with the RNA-seq and Q-PCR data showing differentiation-induced

upregulation of Wnt16 gene expression, WNT16 staining was observed in the cytoplasm of

pachytene spermatocytes, round and elongating spermatids (Figure 4A). Again, in line with

the RNA-seq and Q-PCR data showing differentiation-induced upregulation of Slit2 gene


expression, we observed SLIT2 staining in the nuclei of round spermatids. In addition, some

staining was observed in chromatid bodies in pachytene spermatocytes (Figure 4B).

92 Chapter 4

Figure 4: The protein localization of several RA-induced DEGs in the adult testis. Shown are

localization of (A) WNT16, (B) SLIT2, (C) PDX1 and (D) ADORA1 in adult testis sections. Examples of

several cell types are marked: A, type A spermatogonia; Int, intermediate spermatogonia; B, type B

spermatogonia; pL, pre-leptotene spermatocytes; L, leptotene spermatocytes; Z, zygotene

spermatocytes; P, pachytene spermatocytes; D, diplotene spermatocytes; R, round spermatids; E,

elongating spermatids; Ser, Sertoli cells; Ley, Leydig cells. Stages of the seminiferous epithelium are

indicated with Roman numerals. Bar = 20µm. Negative controls, using the isotype rabbit IgG, do not

show any staining apart from interstitial cells (Figure S1).

RNA-seq and Q-PCR analyses showed that Pdx1 was significantly downregulated by

RA-induced differentiation. In the testis, PDX1 staining was exclusively found in the nuclei of

a subpopulation of type A spermatogonia (Figure 4C). Spermatogonia with clear

heterochromatin patches at the rim of their nuclei, indicative of spermatogonial differentiation,

and intermediate and type B spermatogonia did not stain for PDX1. To pinpoint whether it is

the undifferentiated spermatogonia that are positive for PDX1 staining, we performed IHC on

testis sections from neonatal (8 d.p.p) mice. Indeed, PDX1 staining was observed in

undifferentiated spermatogonia but not in Sertoli cells (Figure 5A), consistent with its staining

pattern in adult testis sections.

While RA-induced differentiation significantly lowered the number of Adora1 transcripts,

ADORA1 protein localization in the testis appeared more dynamic. Like for PDX1, ADORA1

staining was observed in the nuclei of some but not all type A spermatogonia. Also

pachytene spermatocytes and round spermatids up to stage VII of the seminiferous

epithelium were stained (Figure 4D). To investigate whether ADORA1 positive

spermatogonia represent the undifferentiated spermatogonial population, we again

performed IHC on neonatal testis sections. However, unlike for PDX1, all neonatal testicular

cells, consisting of undifferentiated spermatogonia and Sertoli cells, were negative for

ADORA1 staining (Figure 5B).


Figure 5: The protein localization of PDX1 (A) and ADORA1 (B) in the neonatal testis. Asterisks

(*) indicate the positive undifferentiated spermatogonia. Bar = 20µm. Negative controls, using the

isotype rabbit IgG, do not show any staining apart from interstitial cells (Figure S1).

Protein localization of irradiation-induced genes in the testis To study the protein localization of irradiation-induced genes in vivo, adult mice were

subjected to a whole-body IR (1Gy), after which the protein localization patterns of 4 IR-

induced DEGs (Bbc3, Plk2, Pdrg1, Sesn2) for which working antibodies were available were

studied. IHC was performed on testis sections from experimental (3 hours post IR) and

control (no IR treatment) mice. p53 was used as a positive control for the testicular radiation

response [17]. As shown in Figure 6A, nuclear p53 staining was clearly induced in

spermatogonia after IR.

In line with our RNA-seq and Q-PCR data and a previous report [19], IR induced a

strong increase of BBC3 staining in all testicular cells (Figure 6B).

PLK2, on the other hand, although significantly induced at the RNA level in GS cells,

showed no significant difference in response to IR with regard to its localization in vivo. PLK2

staining was observed in pachytene spermatocytes and all subsequent germ cells but not in

spermatogonia, (pre-)leptotene or zygotene spermatocytes (Figure 6C).

Like PLK2, also PDRG1 staining in the testis was not influenced by IR treatment.

Although all germ cells displayed a vague cytoplasmic staining, clear nuclear PDRG1

staining was observed in spermatocytes and round spermatids (Figure 6D).

SESN2 staining was present in Sertoli cells before IR. After IR, SESN2 staining

appeared more intense and was clearly induced in spermatogonia and, albeit less intense

and consistent, in spermatocytes (Figure 6E).

Discussion By perfectly balancing self-renewal and differentiation, SSCs are capable of

maintaining lifelong male fertility. For rodents, long-term culture systems that retain SSC

capabilities have been well established [20, 46, 47]. In these culture systems,

undifferentiated spermatogonia, then termed GS cells, can proliferate exponentially in vitro

and, even after years of culture, still hold the capacity to initiate and maintain

spermatogenesis upon transplantation into recipient testes [21]. Moreover, not only self-

renewal but also spermatogonial differentiation can be induced in this culture system,

providing an advantageous in vitro system to interrogate this process. Several factors, such

as bone morphogenetic protein 4 (BMP4), activin A and RA, have been reported to be

involved in the regulation of spermatogonial differentiation [48-50]. Of these,

94 Chapter 4

RA, the active metabolite of vitamin A, plays pivotal roles in the transition of undifferentiated

spermatogonia into differentiating spermatogonia, as well as in the initiation and progression

of meiosis [50]. Several studies have demonstrated that RA can be used to induce

spermatogonial differentiation in culture [22, 23]. Specifically, RA exposure significantly

downregulates transcription factors essential for SSC self-renewal (e.g. PLZF, OCT4), while

it upregulates spermatogonial differentiation markers such as c-KIT and STRA8 [22]. Further

demonstrating spermatogonial differentiation, RA-treated GS cells exhibit a significantly

reduced colonization ability after transplantation [22]. Moreover, a recent article reported that

RA alone was able to induce GS cell differentiation into zygotene spermatocytes [23].

Intriguingly, differentiation increases the spermatogonial sensitivity to DNA damage in

vivo, for instance DNA damage induced by irradiation [7]. To study the increasing

radiosensitivity of differentiating spermatogonia, we first used RA to induce GS cell

differentiation as described previously [22, 23] and then compared the transcriptomes of

undifferentiated and differentiating spermatogonia. Our differentiation protocol induced the

transcriptomic alteration characteristic for spermatogonial differentiation. Markers for

differentiating spermatogonia and (pre-)meiotic germ cells were significantly upregulated,

whereas well-known markers for SSCs/progenitors or genes essential for SSC self-renewal

showed a substantial downregulation. GO and KEGG analyses further uncovered that, apart

from the expected response to RA, differentiation orchestrated the expression of genes

involved in diverse biological processes such as cell adhesion, cell proliferation and

differentiation, regulation of cell death and apoptosis, and gene transcription. Collectively, in

terms of transcriptomic variation, RA-induced differentiation of GS cells for a large part

mimics in vivo spermatogonial differentiation, supporting the use of this in vitro model for our

study.

We further examined the protein localization patterns of several RA-induced DEGs in

vivo. Wnt16 and Slit2 were upregulated by RA treatment, as shown by our RNA-seq and Q-

PCR data. Previous papers have reported that WNT signaling regulates the growth and

differentiation of stem cells including SSCs [38, 39]. As a member of the WNT family, WNT16

has been found to be associated with estrogen withdrawal and bone loss during aging [51],

and it regulates periosteal bone formation via activation of the canonical WNT signaling

pathway [52]. We observed WNT16 staining mainly in the cytoplasm of pachytene

spermatocytes and spermatids. SLIT2, a member of the SLIT family, has been reported to

inhibit endothelial cell proliferation and migration [40]. Similar to WNT16, SLIT2 staining was

observed in advanced germ cells, in this case the nuclei of round spermatids. Both protein


localization patterns illustrate the idea that genes not required until later stages of

spermatogenesis can already be expressed upon spermatogonial differentiation.

96 Chapter 4

Figure 6: The protein localization of several IR-induced DEGs in the adult testis. Shown are

localization of (A) p53, (B) BBC3, (C) PLK2, (D) PDRG1 and (E) SESN2 in adult testis sections

with/without IR treatment. Examples of several cell types are marked: A, type A spermatogonia; Int,

intermediate spermatogonia; B, type B spermatogonia; pL, pre-leptotene spermatocytes; L, leptotene

spermatocytes; Z, zygotene spermatocytes; P, pachytene spermatocytes; D, diplotene spermatocytes;

M, metaphase I spermatocytes; R, round spermatids; E, elongating spermatids; Ser, Sertoli cells; Ley,

Leydig cells. Stages of the seminiferous epithelium are indicated with Roman numerals. Bar = 20µm.

Negative controls, using the isotype rabbit IgG, do not show any staining apart from interstitial cells

(Figure S1).

Pdx1 and Adora1 showed a significant downregulation upon RA treatment. PDX1 is a

transcription factor that acts as a regulator of pancreatic development and β cell function [41].

Intriguingly, we found staining of PDX1 exclusively in a subpopulation of type A

spermatogonia. Further IHC analysis on neonatal testis sections demonstrated that the

spermatogonia positive for PDX1 staining were indeed undifferentiated spermatogonia.

PDX1 may thus be a suitable novel marker for these cells. Because Pdx1 is downregulated

upon spermatogonial differentiation, it may be involved in SSC self-renewal or proliferation.

ADORA1 belongs to the G protein-coupled receptor 1 family and plays a role in modulating

respiration and metabolism [53, 54]. Like PDX1, ADORA1 staining was observed in some but

not all type A spermatogonia in the adult testis. However, ADORA1 staining was absent from

spermatogonia in the neonatal testis. Also pachytene spermatocytes and a fraction of round

spermatids were stained, suggestive of a more dynamic role for ADORA1 during

spermatogenesis. Interestingly, ADORA1 staining disappeared from round spermatids after

stage VII of the seminiferous epithelium. This is exactly the stage at which spermatogonial

differentiation and meiotic initiation occur in response to RA signaling [55-57]. After stage VII,

the spermatid nuclei start to elongate while the cytoplasm and a more pronounced flagellum

move to the luminal side of the seminiferous tubule [55, 56]. It may thus be the case that, at

stage VII of the seminiferous epithelium, also spermiogenesis is regulated by RA signaling.

To unravel why differentiating spermatogonia are more radiosensitive than their

undifferentiated counterparts, we compared the molecular response of both undifferentiated

and differentiating spermatogonia to IR by way of RNA-seq. We found that, at the

transcriptome level, undifferentiated and differentiating spermatogonia displayed a similar

response to IR. When comparing undifferentiated and differentiating spermatogonia, there

were no IR-induced DEGs that could be attributed to spermatogonial differentiation. Thus, it

is likely that the increase in radiosensitivity of differentiating spermatogonia is mainly caused

by properties, like chromatin architecture, proliferation activity, protein content or post-


translational modifications that are already induced upon differentiation, rather than the

additional expression of genes that is induced by irradiation in both cell types. Our RNA-seq

data uncovered RA-induced upregulation of apoptosis-associated genes (e.g. apoptotic

peptidase activating factor 1 (Apaf1)) and downregulation of genes suppressing apoptosis

(e.g. Bcl2l1, Table S1). The increased radiosensitivity of differentiating spermatogonia may

thus be primed by the presence of genes that favor apoptosis over cell cycle arrest and DNA

damage repair.

Notably, while RNA-seq did not reveal significant DEGs between -RA+IR and +RA+IR

spermatogonia after correction for their respective baselines, GSEA, which does not depend

on observed DEGs but instead takes all the small contributions of the different genes and

looks at them in a pathway setting, did reveal upregulation of the epithelial-mesenchymal

transition (EMT) pathway in irradiated differentiating spermatogonia. EMT is a dynamic

developmental process by which epithelial cells that normally interact with basal membranes

convert to migratory cells with fibroblast-like morphology and mesenchymal secretory

characteristics [58]. SSCs are typically regarded as an intermediate cell category between

epithelial and mesenchymal cells, since they express markers for both epithelial cells (e.g.

CDH1 [59]) and mesenchymal cells (e.g. THY1 [60]). Hence, supported by our RNA-seq data

showing the downregulation of CDH1 and upregulation of THY1 by RA treatment (Table S1),

spermatogonial differentiation, characterized by movement from the basal membrane

towards the lumen, can be associated with enhanced EMT. GS cells can, albeit rarely,

spontaneously reprogram to pluripotency, becoming multipotent GS (mGS) cells.

Interestingly, EMT has recently been found to block this reprogramming of GS cells to

pluripotency [58]. Moreover, mGS cells have been shown to be more resistant to IR than GS

cells [19]. The other way around, it could thus make sense that differentiating spermatogonia,

which are associated with increased EMT and decreased level of pluripotency, are less

resistant to IR.

In addition to upregulation of the EMT pathway revealed by GSEA, differentiating

spermatogonia displayed a less robust increase of p53 protein levels in response to IR. p53

is a well-known tumor repressing factor. Activated by post-translational modifications, it

regulates the transcription of genes involved in cell cycle arrest and apoptosis [61, 62]. p53

can initiate apoptosis to eliminate damaged cells or, alternatively, arrest the cell cycle. The

latter will give the cells an opportunity to repair the damaged DNA before resuming the cell

cycle [61, 62]. Yet, how p53 signaling triggers one of these cascades remains unknown for

most cell types. Our Western blot data showed a more robust p53 induction in

undifferentiated spermatogonia in response to IR. As undifferentiated spermatogonia are

relatively radio-resistant, it seems plausible that spermatogonial p53 preferentially induces

98 Chapter 4

cell cycle arrest. Differentiating spermatogonia, which induce less p53 in response to IR,

would then be less likely to undergo cell cycle arrest and thus have fewer opportunities to

repair the inflicted DNA damage. This would then render differentiating spermatogonia more

radiosensitive.

We also examined the protein localization patterns of several IR-induced DEGs in vivo.

Our RNA-seq and Q-PCR data demonstrated a clear upregulation of Bbc3, Plk2, Pdrg1 and

Sesn2 in irradiated spermatogonia. As a positive control for the IR-induced DNA damage

response, we stained testis sections from irradiated and non-irradiated mice with an anti-p53

antibody. Consistent with a previous literature [17], p53 staining was clearly discerned in

spermatogonia after IR treatment. BBC3 is known to be a key determinant of the intrinsic IR-

induced p53-dependent apoptotic pathway [42]. A previous study showed that

spermatogonial BBC3 was specifically upregulated in differentiating spermatogonia by IR,

whereas its knockdown attenuated apoptosis and increased spermatogonial survival [19].

We found an increase of BBC3 staining in all testicular cells after IR treatment, suggesting a

general role in the testicular response to DNA damage. PLK2 is a serum-inducible kinase

that functions in cell proliferation [43]. Its expression initiates at the G1 phase of the cell cycle,

and it was recently found to modulate mitotic spindle orientation as well as mammary gland

development [63]. Our protein staining showed that IR treatment did not change the

expression pattern of PLK2 in the testis. Interestingly, PLK2 staining was not observed in

spermatogonia, (pre-)leptotene or zygotene spermatocytes but in more advanced germ cells,

in line with a previous article [64] and suggesting a role in meiosis and spermiogenesis.

PDRG1, modulated by p53, is known to be induced by ultraviolet (UV) irradiation [44] and

was recently identified as a novel tumor marker potentially functioning in cancer development

[65]. In addition, Pdrg1 transcripts have been shown to be highly expressed in human testes

[44]. Like that for PLK2, we found PDRG1 staining, irrespective of IR treatment, in pachytene

spermatocytes and round spermatids. Also SESN2 is a tumor suppressor implicated in the

p53 signaling pathway, modulating the cellular response to IR [66]. Before IR, we observed

SESN2 staining in the Sertoli cells. In the IR group, this staining appeared more intense.

Furthermore, after IR, also the spermatogonia were clearly stained, suggesting a role for

SESN2 in the spermatogonial response to IR.

Male cancer patients subjected to chemo- or radiotherapy are often confronted with

sub-fertility caused by spermatogonial apoptosis. Nevertheless, the specific DNA damage-

induced gene expression profiles of undifferentiated and differentiating spermatogonia have

not been studied so far. We took advantage of a well-established in vitro model of

spermatogonial differentiation to study the increasing radiosensitivity of differentiating

spermatogonia, and found no DEGs that were induced specifically in undifferentiated or


differentiating spermatogonia in response to IR. Nevertheless, at the protein level,

undifferentiated spermatogonia showed a stronger upregulation of p53 in response to IR than

differentiating spermatogonia. We therefore propose that the increased sensitivity of

differentiating spermatogonia to IR-induced DNA damage is largely determined by properties,

like chromatin architecture, proliferation activity, protein content or post-translational

modifications that are already induced upon differentiation. The difference in radiosensitivity

may reflect the divergent roles of undifferentiated and differentiating spermatogonia in

maintaining genome integrity and male fertility. When damaged, differentiating

spermatogonia can easily be sacrificed, thereby preventing the transmission of mutations to

the offspring. When the differentiating spermatogonia are lost, the SSCs are still there to

reinitiate spermatogenesis and preserve long-term fertility. In contrast, elimination of

undifferentiated spermatogonia could potentially remove the SSC pool and lead to

permanent infertility. No longer being able to reproduce is an evolutionary dead end. The

undifferentiated spermatogonia may have therefore developed a relatively high resistance to

DNA damage. From an evolutionary point of view, mutated offspring is better than no

offspring.

Author contribution Yi Zheng and Geert Hamer conceived and designed the experiments. Yi Zheng,

Callista L. Mulder, Saskia K.M. van Daalen and Grace Hwang performed the experiments. Yi

Zheng, Aldo Jongejan and Geert Hamer analyzed the data. Yi Zheng and Geert Hamer wrote

the manuscript. Sebastiaan Mastenbroek, Philip W. Jordan, Sjoerd Repping and Geert

Hamer read and revised the manuscript.

Conflict of interest There is no conflict of interest that could be perceived as prejudicing the impartiality of

the research reported.

Funding This study has been supported by an AMC Fellowship, the People Programme (Marie

Curie Actions) of the European Union’s Seventh Framework Programme (CIG 293765) to

G.H., the China Scholarship Counsel (CSC) number 201306300081 to Y.Z., National

Institutes of Health (NIH) K99/R00 HD069458 and NIH R01 GM117155 to P.W.J., and NIH

training grant CA009110 fellowship to G.Hw.

100 Chapter 4

Acknowledgment We thank Klaas Franken and the Laboratory for Experimental Oncology and

Radiobiology, AMC Amsterdam, for assistance with and use of their 137Cs source for IR.


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63. Villegas E, Kabotyanski EB, Shore AN, Creighton CJ, Westbrook TF, Rosen JM. Plk2

regulates mitotic spindle orientation and mammary gland development. Development 2014; 141:1562-

1571.

64. Jordan PW, Karppinen J, Handel MA. Polo-like kinase is required for synaptonemal complex

disassembly and phosphorylation in mouse spermatocytes. Journal Of Cell Science 2012; 125:5061-

5072.

65. Jiang L, Luo X, Shi J, Sun H, Sun Q, Sheikh MS, Huang Y. PDRG1, a novel tumor marker for

multiple malignancies that is selectively regulated by genotoxic stress. Cancer Biol Ther 2011; 11:567-

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66. Sanli T, Linher-Melville K, Tsakiridis T, Singh G. Sestrin2 modulates AMPK subunit expression

and its response to ionizing radiation in breast cancer cells. PLoS One 2012; 7:e32035.

106 Chapter 4

Supplementary data

Figure S1: Negative controls for IHC.

Tables S1-7 will be available on line upon acceptance of the paper.

Chapter 5

Spermatogonial stem cell autotransplantation and germline genomic editing: a future cure for spermatogenic failure and

prevention of transmission of genomic diseases

Callista L. Mulder#

Yi Zheng#

Sabrina Z. Jan

Robert B. Struijk

Sjoerd Repping

Geert Hamer

Ans M.M. van Pelt

#equal contribution

Human Reproduction Update

2016 Sep;22(5):561-73

108 Chapter 5

Table of Contents Introduction Methods Clinical prospects of SSCT to restore spermatogenesis

SSCT to restore fertility in adult cancer survivors

SSCT to enhance fertility in oligozoospermic or azoospermic men

Clinical prospects of SSCT and germline genomic editing CRISPR-Cas9 to genomically modify SSCs

Curing spermatogenic failure by transplantation of genetically modified SSCs

Preventing diseases in offspring by transplantation of genetically modified SSCs

Epigenetic editing of SSCs

Clinical and technical hurdles and drawbacks

Ethical issues

Concluding remarks

Prospects of spermatogonial stem cell transplantation 109

Abstract Background: Subfertility affects approximately 15% of all couples, and a severe male factor

is identified in 17% of these couples. Whilst the etiology of a severe male factor remains

largely unknown, prior gonadotoxic treatment and genomic aberrations have been

associated with this type of subfertility. Couples with a severe male factor can resort to ICSI,

with either ejaculated spermatozoa (in case of oligozoospermia) or surgically retrieved

testicular spermatozoa (in case of azoospermia) to generate their own biological children.

Currently there is no direct treatment for azoospermia or oligozoospermia. Spermatogonial

stem cell (SSC) autotransplantation (SSCT) is a promising novel clinical application currently

under development to restore fertility in sterile childhood cancer survivors. Meanwhile, recent

advances in genomic editing, especially the clustered regulatory interspaced short

palindromic repeats-associated protein 9 (CRISPR-Cas9) system, are likely to enable

genomic rectification of human SSCs in the near future.

Objective and rationale: The objective of this review is to provide insights into the prospects

of the potential clinical application of SSCT with or without genomic editing to cure

spermatogenic failure and to prevent transmission of genetic diseases.

Search methods: We performed a narrative review using the literature available on PubMed

not restricted to any publishing year on topics of subfertility, fertility treatments, (molecular

regulation of) spermatogenesis and SSCT, inherited (genetic) disorders, prenatal screening

methods, genomic editing and germline editing. For germline editing, we focussed on the

novel CRISPR-Cas9 system. We included papers written in English only.

Outcomes: Current techniques allow propagation of human SSCs in vitro, which is

indispensable to successful transplantation. This technique is currently being developed in a

preclinical setting for childhood cancer survivors who have stored a testis biopsy prior to

cancer treatment. Similarly, SSCT could be used to restore fertility in sterile adult cancer

survivors. In vitro propagation of SSCs might also be employed to enhance spermatogenesis

in oligozoospermic men and in azoospermic men who still have functional SSCs albeit in

insufficient numbers. The combination of SSCT with genomic editing techniques could

potentially rectify defects in spermatogenesis caused by genomic mutations or, more broadly,

prevent transmission of genomic diseases to the offspring. In spite of the promising

110 Chapter 5

prospects, SSCT and germline genomic editing are not yet clinically applicable and both

techniques require optimization at various levels.

Wider implications: SSCT with or without genomic editing could potentially be used to

restore fertility in cancer survivors to treat couples with a severe male factor and to prevent

the paternal transmission of diseases. This will potentially allow these couples to have their

own biological children. Technical development is progressing rapidly, and ethical reflection

and societal debate on the use of SSCT with or without genomic editing is pressing.

Keywords: spermatogonial stem cell autotransplantation / male infertility / male reproductive

disorders / germline editing / CRISPR-Cas9


Introduction Subfertility, defined as failure to achieve a clinical pregnancy after at least 12 months of

regular unprotected coitus [1], affects ~15% of all couples. In ~17% of these couples, a

severe male factor, defined as a total motile sperm count below 3x106, is present [2]. A

severe male factor may present as azoospermia (complete absence of spermatozoa in the

ejaculate) or oligozoospermia (low number of spermatozoa in the ejaculate). In case of a

severe male factor, a patient’s own biological children can be generated by ICSI with either

ejaculated spermatozoa (in case of oligozoospermia) or surgically retrieved testicular

spermatozoa by means of testicular sperm extraction (TESE) (in case of azoospermia).

Currently, no direct clinical treatment exists for a severe male factor.

A severe male factor is typically caused by a disturbance during spermatogenesis.

Spermatogenesis occurs in the seminiferous tubules inside the testis. Essential to this

process are spermatogonial stem cells (SSCs) that maintain a perfect balance between self-

renewal and differentiation into mature sperm, thereby sustaining fertility throughout a man’s

life. Both oligozoospermia and azoospermia can be due to a reduction in SSC numbers

throughout the testis as seen in testicular biopsies taken from these men [3-5]. In some of

these men, the testicular biopsies display focal spermatogenesis, where sperm is only

produced in a subset of seminiferous tubules. Although the production of spermatozoa is

limited due to the low number of SSCs, the existing SSCs are still functional and capable of

continuous self-renewal and differentiation [6].

Whilst the etiology of a severe male factor remains largely unknown, a few genetic

defects [7-9], including structural and numerical chromosomal abnormalities [10, 11] and

Y‐chromosome deletions [12, 13], have been identified to associate with a severe male

factor-induced subfertility. In addition, male subfertility can be attributed to previous

gonadotoxic treatment [14]. Both chemotherapy and radiotherapy are known to destroy the

SSC pool, resulting in oligozoospermia or azoospermia in a large proportion of cancer

survivors [15].

A possible future application that could directly treat a severe male factor might be

autotransplantation of SSCs. Transplantation of SSCs that are stored prior to cancer

treatment is proposed as a means to restore fertility in childhood cancer survivors. This

application involves transplantation of SSCs into the seminiferous tubules via the efferent

duct or rete testis [16, 17]. Upon SSC transplantation (SSCT), SSCs migrate to the basement

membrane of recipient seminiferous tubules, colonize the epithelium and undergo self-

renewal and differentiation so that permanent spermatogenesis is established. Therefore,

112 Chapter 5

SSCT should allow natural conception without further fertility treatment, making SSCT a

direct treatment for a severe male factor.

In case the severe male factor is due to a genomic mutation, SSCT can only be

successful if it is combined with correction of the mutation. Recent advances in genomic

editing, especially those with the clustered regulatory interspaced short palindromic repeats-

associated protein 9 (CRIPSR-Cas9) system, can allow for rapid, easy and highly efficient

genetic alterations of a wide array of cell types and organisms including human cells [18].

Thus, if genomic editing is combined with SSCT, it would in principle allow those suffering

from spermatogenic failure to have their own biological children. Furthermore, SSCT with

genomic editing may be used to prevent paternal transmission of genomic diseases.

In this review, the clinical prospects of SSCT with or without genomic editing for male

adult cancer survivors, for men with oligozoospermia and azoospermia and for carriers of

genomic diseases are discussed (Figure 1).

Methods We performed a narrative review using literature available on PubMed not restricted to

any publishing year on topics of subfertility, fertility treatments, (molecular regulation of)

spermatogenesis and SSCT, inherited (genetic) disorders, prenatal screening methods,

genomic editing and germline editing. For germline editing, we focussed on the novel

clustered regulatory interspaced short palindromic repeats-associated protein 9 (CRISPR-

Cas9) system. We included papers written in English only.

Clinical prospects of SSCT to restore spermatogenesis The first successful SSCT was reported in 1994 and resulted in fertility restoration in

sterile recipient mice upon transplantation of donor-derived SSCs [19]. Subsequent studies

substantiated the findings by showing restoration of spermatogenesis after SSCT in mice

[20-24] and other animal models [25-29], including non-human primates [30, 31]. In mice,

SSCs can be transplanted into the recipient seminiferous tubules by injecting the efferent

ducts, the rete testis or directly injecting seminiferous tubules [32]. Injection via the efferent

ducts is mostly employed in rodents [33]. Yet, due to the anatomic and size difference in

testes, ultrasound-guided transplantation via the intra-rete testis has been proved to be the

least invasive and most efficient and successful protocol for non-rodent species such as pigs

[26], bulls [27], primates [30, 31, 34] and ex vivo human testes [31, 35]. Crucial to this

procedure is the injection of cells via the rete testis, and optimization is needed in this


respect to augment success rates [36]. Inaddition, the quality and quantity of SSC niches in

the recipient testis makes a difference to the success of autologous transplantation [37]. To

date, multiple papers describe the generation of SSCT-induced healthy offspring in rodents

[21-24, 38, 39] and non-rodent large animal models such as goats and sheep [40-42]. A

breakthrough was recently achieved by Hermann et al. [30], who successfully performed

autologous and allogeneic transplantation of SSCs from rhesus monkey, leading to the

generation of donor-derived sperm in both cases. Subsequently, ICSI was conducted to

fertilize oocytes, and embryos with donor paternal origin were finally produced. This

demonstration in primates provides prospects for future clinical translation of SSCT.

For humans, SSCT has been proposed as a future clinical application for those men

with the risk of complete germ cell depletion that have no option to cryopreserve sperm, in

particular, prepubertal cancer patients [17, 43-45] and, theoretically, even in case of focal

spermatogenesis [46]. Prepubertal cancer patients especially rely on SSCT because

spermatogenesis is not initiated yet, which means that semen cryopreservation prior to

treatment is not an option. By opting for storage of a testicular biopsy before cancer therapy,

SSCT can be applied later in life when the patient is cured and expresses the wish to have

children [45, 47]. In this case, cryopreservation of biopsied testicular tissues constitutes an

important part of fertility preservation [48]. Currently the most popular avenue to preserve

pieces of testicular tissues is through controlled slow freezing [49-54], with the addition of

cryoprotective agents such as dimethyl sulphoxide or ethylene glycol with or without sucrose

[47, 49, 52, 53, 55-57]. Vitrification instead of controlled slow freezing has also been tested

with positive outcomes [58-60]. Nevertheless, protocols for tissue preservation require further

optimisation to maximize the viability of post-thawed human testicular tissues [47].

Following successful cryopreservation, conventional SSCT can be performed at a later

stage, which involves the transplantation of the patient’s SSCs into one’s own testis (i.e.

autotransplantation). After autotransplantation, SSCs migrate to the basement membrane of

the recipient seminiferous epithelium, from where they reinitiate spermatogenesis (Figure 1).

In this way, the patient can (re)gain his fertility and generate their own biological children by

natural conception, as shown in animal experiments [21, 40].

The efficiency of SSCT is highly dependent on the number of transplanted SSCs [61,

62]. Previous reports suggest that SSCT is unlikely to become clinically applicable without an

approach to successfully expanding human SSCs in vitro [37, 63]. In vitro expansion of SSCs

has been successfully demonstrated in studies using rodent SSCs [20, 64-67]. Even after 2

years of culture, the SSCs retained the capacity to colonize the basal membrane of

114 Chapter 5

seminiferous tubules and could further develop into healthy and functional sperm [64]. In

human trials, SSCs can first be isolated from biopsies and then subjected to primary culture

to increase their number for the sake of SSCT. Several culture systems have been

established for human adult SSCs [68-72], as well as for human prepubertal SSCs [73].

Presently (xeno)transplantation is the well-acknowledged and only available assay for

functionality of human SSCs. It is shown that in vitro propagated human SSCs are capable of

migrating to the niche at the basal membrane of the seminiferous tubules upon

xenotransplantation into the testis of immunodeficient mice [70, 73], indicating that these

cultured testicular cells still have SSC capabilities. Nevertheless, human spermatogenesis is

not initiated in the mouse model. This is not unexpected given the large phylogenetic

distance between mice and humans, and because of this, the murine niche cannot support

full development of human SSCs into sperm [74]. Despite that, by comparing the numbers of

migrated SSCs in the recipient testis after xenotransplantation of early and late passages

from primary human testicular cultures, it has been established that adult as well as

prepubertal SSCs can indeed proliferate in culture [70, 73]. Thus, the successful expansion

of human SSCs in culture paves the way for clinical applications of SSCT.

Once SSCT is clinically implemented for childhood cancer survivors, other patient

groups that have the risk of becoming subfertile or that suffer from subfertility might also

benefit from this treatment. In a recent study in azoospermic men, it was shown that these

men hold a positive attitude toward SSCT, which was persistent even after acknowledging

that a new experimental technique might have some risks for themselves or their offspring

[75].

SSCT to restore fertility in adult cancer survivors Due to the high sensitivity of spermatogonia to DNA damage [14, 76], spermatogonial

apoptosis and subsequent subfertility is a major side-effect of most cancer treatments. The

chance of becoming azoospermic temporarily or permanently after cancer therapy is highly

dependent on the type and dosage of the treatment [15]. A recent study has shown that 25%

of patients who underwent chemotherapy were azoospermic after 7-218 months (median: 40

months), with the highest chance of azoospermia in Hodgkin disease survivors (63%) [77].

Cryopreservation of semen prior to treatment is historically an effective and

inexpensive way to preserve fertility in adult male cancer patients. The cryopreserved semen

can be used to achieve a pregnancy by cervical or intrauterine insemination (IUI) or, in case

the quality of the semen is too low, by IVF or ICSI treatment. However, cancer patients often

show decreased fertility at the time of cancer diagnosis. In fact, Ragni et al. [78] report that

12% of patients who wish to store their semen are azoospermic at the time of cancer


diagnosis. Moreover, semen cryopreservation severely reduces sperm motility [79, 80],

sperm count [79] and DNA integrity [81]. Hampered sperm motility significantly decreases the

chance of live birth after IUI [82], and therefore the majority of patients who make use of

cryopreserved spermatozoa have to resort to IVF/ICSI. However, IVF/ICSI is costly and

burdensome, and requires ovarian hyperstimulation of the healthy female to retrieve oocytes,

which is not risk-free and leads to onset of the ovarian hyperstimulation syndrome in 1-8% of

stimulated women [83, 84]. In addition, it is well known that IVF/ICSI is associated with

adverse short-term outcomes including preterm birth, lower birthweight and a higher

prevalence of birth defects [85, 86].

Although SSCT does require a testis biopsy and treatment of the affected male, it

would avoid IVF/ICSI treatment of the healthy female partner since natural conception might

be feasible after SSCT. In the future, SSCT may therefore become an appealing alternative

for fertility preservation in adult cancer patients in a similar way as described for prepubertal

cancer patients by cryopreserving a testis biopsy before onset of cancer treatment.

SSCT to enhance fertility in oligozoospermic and azoospermic men Oligozoospermic and azoospermic men are capable of fertilization if they produce

morphologically normal sperm in the testis. In couples where the male is oligozoospermic or

azoospermic, ICSI is used to achieve fertilization with ejaculated or surgically retrieved

spermatozoa, respectively. However, since oligozoospermia and azoospermia may be

caused by a reduction in functional SSCs [3-5], a simple increase in SSCs may restore

spermatogenesis and fertility in these men. This is especially relevant to the patients who

display focal spermatogenesis at the histological level, in which some tubules show normal

spermatogenesis, while others display Sertoli cell-only syndrome. The tubules with normal

spermatogenesis harbour functional SSCs, and the hypothesis is that if these spermatogonia

are propagated in vitro and transplanted back into the testis, they will repopulate the empty

tubules and initiate spermatogenesis in these tubules. A recent article describes the

characteristics of cultured SSCs deriving from patients who suffer from focal

spermatogenesis due to a deletion of the azoospermia factor c (AZFc) region on the Y

chromosome [46]. In vitro propagated SSCs from these men with focal spermatogenesis

behaved similarly during culture and showed comparable gene expression of key

spermatogonial markers when compared to SSCs originating from healthy counterparts with

normal spermatogenesis. These results suggest that patients with oligozoospermia or

azoospermia as a result of focal spermatogenesis might also benefit from propagation and

transplantation of their own SSCs. Yet, this hypothesis needs to be demonstrated in clinical

trials. However, one drawback that should be accounted for is that if a

116 Chapter 5

mutation is present on the Y chromosome, as in the case of men with AZFc deletions, male

offspring of these men will harbour the same mutation and are likely to be oligozoospermic or

azoospermic too [87]. It makes sense that this also holds true for mutations on other

chromosomes. However, this problem also arises with other contemporary fertility treatment,

such as IVF/ICSI with or without TESE. In order to prevent transmission of these genetic

aberrations, additional measures have to be taken.


Figure 1: A schematic depiction of the proposed SSCT therapy. (A) A testicular biopsy is taken

from the patient and cryopreserved. From the biopsy, SSCs are propagated in vitro, during which

endogenous genomic defects may be repaired. Propagated SSCs are subsequently autotransplanted

to the testis and then colonize the testis and restore spermatogenesis, enabling the patient to father a

child without additional therapy. (B) The testicular histology of men with a severe male factor in

different patient groups. The histology may show various phenotypes throughout the testis. For male

(childhood) cancer survivors, a biopsy is cryopreserved prior to cancer therapy. Hence, thawing of the

cryopreserved biopsy is indispensable to the treatment. In vitro propagation is needed for all patient

groups, while genomic modification is only needed for those with a maturation arrest or carriers of

diseases. In male carriers of diseases with full spermatogenesis, all germ cells including spermatids

express the mutated genes, and local irradiation of the testis is required prior to transplantation to

remove the mutated endogenous spermatids. After (genomically modified) SSCT, testis histology

should, in theory, restore to full spermatogenesis.

Clinical prospects of SSCT and germline genomic editing While the use of SSCT for adult cancer patients or oligozoospermic and azoospermic

patients that display focal spermatogenesis seems rather straightforward, azoospermic

patients suffering from a maturation arrest in spermatogenesis cannot directly benefit from

SSCT because transplantation of the patient’s SSCs would result in the same arrested

phenotype and not cure their spermatogenic failure (Figure 1). However, in some cases, the

maturation arrest may be attributed to genetic mutations or arises from epigenetic

disturbances. Repair of these disorders in SSCs before SSCT would theoretically restore

spermatogenesis and subsequent fertility in these patients and in addition prevent the

transmission of the mutation to the offspring. The fact that human SSCs can propagate in

culture for extended periods of time enables (epi)genetic editing prior to transplantation.

With recent advances in the field of genomic editing, in particular the use of novel

techniques such as the CRISPR-Cas9 system, SSCT with genetically modified SSCs has

become feasible [88-91]. In addition, recent work has shown that epigenetic editing is also

possible with CRIPSR-Cas9 [92].

CRISPR-Cas9 to genomically modify SSCs Traditional genome editing mainly relies on homologous recombination in embryonic

stem cells (ESCs). Over the last decade, novel genome editing platforms such as zinc-finger

nucleases (ZFNs), transcription activator-like effector nucleases (TALENs) and most recently

the CRISPR-Cas9 system have been developed. These techniques are based on

118 Chapter 5

engineered nucleases that can cause a double-strand break of DNA, and are much less

laborious and time-consuming compared to traditional strategies. With the emergence of

these novel techniques (Table 1), the majority of cell types, including SSCs, can now be

targeted.

Table 1: An overview of different genome editing techniques.

Genome editing

systems

Homologous

recombination without

engineered

nucleases

Conventional

engineered nucleases (ZFNs

/ TALENs)

Novel

engineered nucleases

(CRISPR-Cas9)

Novel

engineered nucleases

(GeCKOa)

Target cells Mostly ESCs Most cell types Most cell types Most cell types

Approaches to

delivering

targeting vectors

Non-viral

transfection / viral

transduction/

microinjection

Non-viral


transduction /

microinjection

Non-viral


transduction /

microinjection

Lentiviral

transduction

Technical

difficulty

High High Low Intermediate

Targeting

efficiency

Off-target effects

Low

Low

Variable

Variable

Generally high

Generally low

High

Variable

Possible to target

a large scale of

genes in parallel?

No No No Yes

Suitable for the

clinic?

No, due to low

efficiency and the

typical

requirement of

ESCs

Not optimal Yes Currently not due

to lentiviral

transduction

aGeCKO, genome-scale CRISPR knockout; ESC, embryonic stem cell; ZFN, zinc-finger nuclease;

TALEN, transcription activator-like effector nuclease; CRISPR-Cas9, clustered regulatory interspaced

short palindromic repeats-associated protein 9.

Prior to clinical application of genomically modified SSCs, the safety of the patient

needs to be guaranteed. In a clinical setting, off-target effects are not acceptable, and

technical simplicity would be desirable. ZFNs and TALENs were utilized to successfully

manipulate mouse SSCs [90]. However, the required design and engineering of nucleases


necessary for both ZFNs and TALENs is strenuous and of high technical difficulty. A better

alternative to genetically modify SSCs seems to be the unprecedentedly simple CRISPR-

Cas9 system (Figure 2), and articles describing successful manipulation of rodent SSCs by

way of CRISPR are now available [88, 89, 91]. The CRISPR-Cas9 system not only bypasses

the engineering of nucleases but also generates far less off-target effects compared with

ZFNs [93]. According to a recent report, with genome-wide screens, no obvious off-target

genetic or epigenetic changes could be detected in a large SSCT experiment involving

CRISPR-Cas9-mediated gene targeting and transplantation of modified mouse SSCs [88].

Figure 2: The CRISPR-Cas9 system. The

Type II Streptococcus pyogenes clustered

regulatory interspaced short palindromic

repeats-associated protein 9 (CRISPR-Cas9)

(SpCas9) system, which is the simplest and

most extensively used CRISPR-Cas9

technology, is based on a guide-RNA (gRNA)

containing a specific 20 bp sequence to guide

the DNA endonuclease Cas9 to a

complementary target DNA sequence in the

genome where it induces a DNA double-strand

break (DSB). The 20-bp target genomic DNA

must be upstream of a specific sequence (5’-

NGG, where N represents a random nucleotide).

The Cas9-induced DSB occurs ~3-bp upstream

of the 5’-NGG, and can in theory be induced in any 20-bp genomic DNA sequence flanking 5’-NGG.

The Cas9-induced DSB will then be repaired by either homology-directed repair (HDR), which can

occur with the presence of DNA repair templates, or by non-homologous end joining (NHEJ). The

error-prone NHEJ creates insertions/deletions (indels) around the DSB point. Indels, especially when

occurring in early coding exons, can cause loss of gene function (gene knockout) by causing a frame

shift that can lead to formation of a pre-mature stop codon. In contrast, HDR uses a template

sequence for very precise repair of the DSB. Exogenous DNA repair templates (with the required

sequences placed between homology arms) can be provided to the cells together with other

components of the CRISPR-Cas9 system to create specific indels or modifications at target genomic

loci. Thus, the CRISPR-Cas9 system can be used to insert sequences or correct disease-causing

mutations in a very accurate way.

120 Chapter 5

Curing spermatogenic failure by transplantation of genetically modified SSCs A proportion of infertile men have non-obstructive azoospermia as a result of

spermatogenic arrest. However, in 41% of these patients a few germ cells escape the arrest

and form elongated spermatids, which can be extracted from testicular tissue by means of

TESE and used for fertilisation in an ICSI procedure [94]. When no sperm is found during this

procedure, no treatment options are currently available. Fortunately, Lim et al. [68] have

shown that spermatogonia from patients suffering from non-obstructive azoospermia due to a

maturation arrest are able to proliferate in their long-term culture system in a similar manner

as men with obstructive azoospermia. Therefore, SSCs might be used in future SSCT as a

valid option to treat these men. Very recently, Yuan et al. [95] employed TALENs to successfully rectify a point

mutation in the mouse c-kit gene that blocks spermatogonial differentiation, and after

correction spermatogenesis could be rescued, for the first time demonstrating that

spermatogenic failure-related genetic defects can be corrected by genome editing platforms.

Thus, it makes sense that after CRISPR-Cas9-mediated correction of the genetic defects

responsible for spermatogenic failure, SSCs from azoospermic men can subsequently be

used for SSCT, offering men with spermatogenic failure a patient-tailored treatment option.

Besides, in small cohorts of azoospermic men, various single nucleotide polymorphisms

(SNPs) associated with arrests during spermatogenesis have been identified [96-100],

forming additional candidate targets for genetic modification of SSCs. A recent paper reports

the use of CRISPR-Cas9 to interrogate male infertility-related SNPs in mice [101]. In addition,

the recently developed genome-scale CRISPR knockout (GeCKO) enables the targeting of a

variety of genes in parallel [102, 103]. Because infertility is often not believed to be a

monogenic disorder but is rather thought to be caused by a spectrum of genes [104], the

GeCKO system could serve as a prospective platform for gene therapy of such patients.

Hence, when the infertility-causing genetic mutations are known, SSCs could first be isolated

from testicular biopsies and propagated in vitro to increase their number. Subsequently, the

propagated SSCs could be co-transfected with CRISPR-related vectors (to cut DNA) and

exogenous DNA repair templates specific for the mutations (to induce homology-directed

repair, Figure 2) in a patient-specific manner. Finally, after selection and whole genome

(epi)genetic off-target analysis, the modified SSCs can be auto-transplanted into the testis to

initiate spermatogenesis and produce corrected sperm.


Preventing diseases in offspring by transplantation of genetically modified SSCs Carriers of inherited genetic diseases, albeit often fertile, have to make important

decisions when it comes to reproduction. Couples can opt to remain childless to prevent

diseases in their children, opt for adoption or resort to a germ cell donor. Alternatively, these

couples can attempt to have their own biological children and detect whether their

prospective children are carriers of the disease via prenatal testing during pregnancy. In case

the fetus is a carrier of the disease, the parents have to make the emotionally laden decision

whether to terminate their pregnancy. Couples can also opt for PGD as a preventive

measure for the birth of a child with a genetic defect. PGD is well established for monogenic

diseases such as cystic fibrosis [105], beta thalassemia [106] and Huntington’s disease [107].

In PGD, couples undergo IVF treatment in which a single cell is aspirated from each embryo

at the 6-8 cell stage to perform subsequent genetic testing for high-risk disease alleles. Only

unaffected embryos are transferred to prevent the birth of children with these severe genetic

diseases.

Even though PGD provides a solution for couples at risk for transmitting a genetic

disease, IVF treatment of the women, including ovarian hyperstimulation, is indispensable in

the process of PGD. Moreover, many embryos are created, while only a few will be used to

induce pregnancy.

Currently there is no way to prevent genetic diseases in the offspring without creating

affected embryos. Nevertheless, if the prospective father is the carrier of a disease allele, the

disease-causing mutation can theoretically be corrected in isolated SSCs during in vitro

culture. Subsequently, transplantation of the modified SSCs would result in genetically

normal sperm and therefore prevent transfer of the disease allele to the next generations.

Additionally, it would enable male carriers at risk for transmitting genetic diseases to naturally

conceive a healthy child without IVF or prenatal genetic testing.

Inspiringly, CRISPR-Cas9 has been shown to successfully repair mutations in disease-

causing genes in different species and cell types. In mice, mutations in the Crygc gene

(which causes cataracts) [108], dystrophin gene (which causes Duchenne muscular

dystrophy, DMD) [109] and a Fah mutation in hepatocytes [110] have been repaired by the

CRISPR-Cas9 system. In human trials, the CRISPR-Cas9 system has been used to

precisely correct the hemoglobin beta and dystrophin gene, in β-thalassemia [111] and DMD

patient-induced pluripotent stem cells [112], respectively. Another report describes

successful repair of the cystic fibrosis transmembrane conductor receptor locus in cultured

intestinal stem cells from patients with cystic fibrosis [113]. These reports raise the possibility

that CRISPR-Cas9 could be used to repair inheritable mutations through the germline.

122 Chapter 5

Interestingly, a recent article reports successful genome editing of mouse SSCs with

the CRISPR-Cas9 system [88]. Transplantation of the genetically modified SSCs led to fertile

offspring, in which a Crygc mutation causing cataracts was corrected. To our knowledge, this

is the first report that describes CRISPR-Cas9-mediated genome editing in SSCs in

combination with SSCT, thereby preventing diseases in the offspring. Furthermore, as the

transplanted SSCs were the cell lines derived from isolated and corrected single cells, this

method can generate healthy descendants at 100% efficiency, thereby averting the problem

of mosaicism. In addition to this pioneering work, two other recent articles also provide the

proof of concept by showing CRISPR-Cas9 and SSCT-induced germline transmission in

rodents [89, 91]. Hence, SSCT and the CRISPR-Cas9 system can be well combined in the

future to prevent the transmission of inheritable diseases to the offspring.

Epigenetic editing of SSCs Mechanisms that underlie cellular functioning are orchestrated by different layers of

transcriptional regulation. Apart from genetic factors, epigenetic regulation is key for the

proper functioning of a cell. Epigenetic traits are defined [114] as ‘heritable phenotypes

resulting from changes in a chromosome without alterations in the DNA sequence’, mediated

by several factors such as DNA methylation, histone modifications and higher order

chromatin structuring. Disruption of epigenetic regulation has been shown in an array of

complex diseases, including cancer, diabetes and cardiovascular diseases (reviewed in [115-

117]). Thus, in theory, epigenetic editing may be a valid alternative for genetic repair to

modify aberrant gene expression in SSCs from those at risk of transmitting diseases to their

own biological children.

The core of currently described epigenetic editing approaches is the fusion of an

epigenetic modulatory enzyme to a protein with a DNA binding domain in order to affect gene

expression and modulate local parameters, such as cytosine methylation and demethylation

or histone modification once the enzymatic construct is in place. This powerful approach has

been used to silence and/or activate specific DNA sequences by altering epigenetic

parameters of target genes (reviewed in [118], [119]). For example, a recent study describes

the guidance of the Ten-Eleven Translocation 2 DNA demethylation enzyme to the promoter

region of the intercellular adhesion molecule 1 gene, causing local demethylation and

reactivation of the gene where it was normally silenced [120]. Another group has reported the

development of a programmable molecular construct consisting of a nuclease-deactivated

Cas9 (dCas9) protein fused to the catalytic core of the acetyltransferase p300, which can be

used to modulate histone acetylation of any Cas9-targetable genomic location [92]. One


might imagine that epigenetic editing techniques might allow correction of the transcriptional

regulation of pivotal genes in various biological processes, including germ cell development.

Epigenetics have been shown to play a key role in normal germ cell development [121-123],

and allele-specific DNA methylation was altered in semen from men that suffer from

spermatogenic failure [124-128]. Local correction of abnormal DNA methylation or histone

modification of target genes in infertile men might improve the spermatogenic potential.

In theory, SSCT of epigenetically modified SSCs may also be applicable for inherited

epigenetic diseases. Epidemiological data point to human transgenerational epigenetic

inheritance, including the Dutch Famine Birth Cohort Study [129-132] and the Swedish

Overkalix population [133]. Additionally, a few (case) studies describe inheritance of a

disease-associated epimutation of a specific locus, such as the SNURF-SNRPN locus in

Prader-Willi and Angelman syndrome [134] and the cancer predisposing gene MLH1 [135].

However, one must realize that the epigenome is reset in an extensive way during early

embryonic development (reviewed in [136] and [137]). Even though some inherited

epigenetic marks seem to escape epigenetic reprogramming, it remains unclear whether

epigenetic germline editing combined with SSCT may benefit patients in the future. Therefore,

more research is needed before SSCT can be applied to cure heritable epigenetic diseases.

Clinical and technical hurdles and drawbacks The field of SSCT is uprising, and a variety of patient groups may benefit from this

therapy in the future. However, some hurdles still need to be overcome prior to its clinical

implementation. For one thing, safety of the patient and his offspring is of major concern [45].

Human SSCs may change (epi)genetically when exposed to an in vitro environment. Yet,

there is evidence of genetic stability of cultured human SSCs [138]. DNA methylation of

maternal and paternal imprinted genes in uncultured murine SSCs did not alter after

transplantation [21, 23], whereas cultured human SSCs showed changes in DNA methylation

in some selected regions of maternal and paternal imprinted genes [138]. Conclusively,

although the published data suggest that SSCT may be safe for the clinic, more (pre)clinical

studies are needed in this field to ensure safety for patients, as well as for their offspring.

Another challenge in cancer patients is that primary testicular cultures may be

contaminated with lingering cancer cells from leukemic or metastasized patients. While some

researchers were unable to successfully sort out cancer cells [139], others succeeded in

removing leukemic cells from testicular cultures [140] or from cell suspensions [141].

124 Chapter 5

In comparison, the clinical implementation of genomically modified SSCT is even more

challenging. Aside from the potential risks accompanying conventional SSCT, we currently

do not know whether genomic manipulation of SSCs harbours any extra risks for the patient

or his offspring. In addition, a major clinical drawback of germline therapy in fertile carriers of

diseases remains that the patients have to undergo local irradiation to deplete the testis of

endogenous mutated spermatogonia before SSCT. Otherwise, the testis of the recipient

father would produce two populations of sperm cells: those that arise from endogenous

SSCs carrying the disease-causing mutation as well as those from the corrected SSCs

introduced by transplantation. As a consequence, the semen of the father would contain a

mixed population of spermatozoa and children conceived by natural conception could be

derived from either a corrected or endogenous sperm cell. Although local irradiation has

been demonstrated to be an effective measure to deplete the testis of germ cells in animal

studies [41, 142], it can have a deleterious influence on outgrowth of seminiferous tubules,

especially in prepubertal testes [36]. Besides, it may cause damages to surrounding organs

and cells. Moreover, some endogenous spermatogonia might survive the irradiation and are

still capable of developing into spermatids, thereby risking the transfer of the genomic

aberrations. In this sense, the development of alternatives to exclude endogenous SSCs is

needed to better strike the balance between the benefits of SSCT and the potential risks of

the required total depletion of endogenous spermatogenesis.

In terms of genomic manipulation, some technical problems remain to be addressed.

First, the CRISPR-Cas9 components need to be delivered into cells. The way in which the

Cas9 nuclease is introduced (viral or non-viral) has significant clinical implications (Table 1).

In case of viral transduction, CRISPR-sequences, and possibly even residual viral

sequences, integrate into the genome of the patient. This raises serious safety concerns and

is not suitable for clinical application. However, non-viral delivery methods often fall short, as

they can be inefficient in gene delivery (e.g. liposome-mediated transfection), are laborious

(e.g. microinjection) or lead to significant cell mortality (e.g. electroporation). While SSCs

have been demonstrated to be refractory to most non-viral transfection approaches [143],

novel electroporation devices that are currently being used in some laboratories may be the

option to transfect SSCs with adequate efficiency [88-91, 144]. Alternatively, transfection of

the mRNA instead of the corresponding DNA vectors has been shown to be more efficient for

genome editing [90]. Also the recently developed novel method regarding direct intracellular

delivery of proteins might serve as another option for gene targeting [145].


In spite of the developments, one needs to realize that SSCT in combination with

genomic editing to cure or prevent diseases is only feasible when the genetic mutation is

known. Unfortunately, the genetic etiology remains elusive in many cases. More research on

the identification of genomic mutations that cause subfertility is necessary in this respect [7-

9]. A recent article describes that CRISPR-Cas9 was exploited to successfully eradicate

porcine endogenous retroviruses with the copy number as high as 62 in a porcine cell line

[146], showing the robustness and versatility of CRISPR. However, various genetic diseases,

including subfertility, are believed to be caused by an array of genes [104], which renders

genetic correction substantially more difficult as it would require simultaneous targeting of

various loci. A previous report shows that a maximum of five genes could be simultaneously

disrupted by microinjection of CRISPR components into mouse ESCs [147]. At present, the

novel GeCKO system seems to be the only possible approach to targeting a wide array of

genes in parallel. Nevertheless, GeCKO, which requires lentiviral transduction and

subsequent integration of CRISPR components into the genome, is considered unsafe for

clinical application at the moment. We still need to await further development in this regard

before we can target a variety of genes in parallel safely for clinical purposes.

Another important issue is the potential off-target alterations induced by the CRISPR-

Cas9 system. Recent studies have revealed that the 20-bp gRNA-DNA hybrid (Figure 2) has

the potential to tolerate 1-3 or even more sequence mismatches [18, 148, 149]. As a

consequence, normal genes containing high homology to the target sequence might also be

targeted. Reassuringly, whole genome sequencing of the CRISPR-Cas9-modified SSCs

showed no apparent off-target mutations [88]. Moreover, novel versions of CRISPR with

enhanced specificity but without the sacrifice of on-target activity have been developed

recently [150, 151], which will further facilitate its broad applications in the clinic.

Also, the targeting spectrum of the CRISPR-Cas9 system needs to be expanded. The

requirement of a specific sequence (e.g. 5’-NGG for type II SpCas9, Figure 2) following the

target is a principal constraint. While 5’-NGG occurs quite frequently in the human genome,

further extensions of the targeting range by development of other types of CRISPR that

recognize distinct sequences downstream of the targeting site [152, 153] would give broader

options for genome editing.

Ethical issues As genomic modification of SSCs leads to germline transmission of the modified trait to

the next generation, elaborate ethical reflection and an intensive societal debate on the

126 Chapter 5

acceptability of a clinical application of germline gene editing should precede the actual

clinical application of modified SSCs. Recently two groups employed the CRISPR-Cas9

system to achieve genome editing in human tripronuclear zygotes [154, 155]. Although the

zygotes used were unable to develop into viable embryos, these reports still initiated major

debates. Some people propose a complete ban on germline genomic editing [156], while

others request a moratorium on the clinical application of germline editing but suggest

permission for research in this field [157]. To date, multiple articles have been published to

broaden the discussion [158-162]. We strongly support the establishment of a societal

platform including molecular and (stem) cell biologists, medical professionals, ethicists,

politicians, citizens and most importantly patients, to discuss under which circumstances and

to what extent germline modification should be allowed.

Concluding remarks In this review, we explore different patient groups that may benefit from SSCT with or

without genomic editing. We conclude that the clinical implementation of SSCT can

potentially reach far beyond fertility preservation in childhood cancer patients. Conventional

SSCT could help adult cancer patients preserve their fertility, while fertility may also be

enhanced in oligozoospermic or azoospermic patients using this technology. Successful

genomic modification of SSCs by the novel CRISPR-Cas9 system in culture could repair

detrimental mutations, thereby treating patients with non-obstructive azoospermia and

carriers of diseases in the future.

The safety of the patient and the following generations is of paramount importance.

From our perspective, by no means should these techniques be employed in the clinic until

safety and efficiency has been demonstrated empirically. Therefore, more fundamental

research remains to be conducted. In addition, a societal and ethical debate should precede

the use of modified SSCs in a future clinical application of SSCT.

Nonetheless, SSCT, with or without genome editing, is a potential powerful platform

that potentially can be employed to cure infertility or even inheritable mutations in various

patient groups. Research in this field is thriving, and a revolution might be visible at the

horizon.

Authors’ roles C.L.M., Y.Z., G.H. and A.M.M.P. designed the outline of the review. C.L.M. and Y.Z.

drafted the original manuscript. S.Z.J. and R.B.S. gave substantial contribution to the

manuscript. C.L.M., Y.Z. and S.Z.J. designed and created the figures and table in this


manuscript. S.R., A.M.M.P. and G.H. critically reviewed and revised the manuscript and

approved the final version.

Funding ZonMW (TAS116003002), the China Scholarship Counsel (CSC) (201306300081), an

AMC Fellowship and the People Programme (Marie Curie Actions) of the European Union’s

Seventh Framework Programme (CIG 293765).

Conflict of interest The author(s) report no financial or other conflict of interest relevant to the subject of

this article.

128 Chapter 5

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Chapter 6

General discussion and implications for future research

140 Chapter 6

General discussion

In the current thesis we have shed light on the dynamic response to DNA damage

during spermatogonial development, with a specific focus on the role of the structural

maintenance of chromosomes (SMC) 5/6 complex. In addition, we have addressed the broad

and potentially large clinical prospects and implications of using CRISPR-Cas9 to treat

spermatogenic failure or to prevent the transmission of genetic abnormalities to human

offspring by transplantation of genetically modified human spermatogonial stem cells (SSCs).

In the current chapter, we put our results into perspective and provide guidance for future

research.

The role of the SMC5/6 complex during spermatogonial proliferation and differentiation

One striking characteristic of spermatogenesis is the continuous change in chromatin

architecture and function. Regulation of chromatin dynamics is not only required to guide

processes like meiosis and chromatin compaction during spermiogenesis but also essential

to safeguard genomic stability during spermatogonial development. Previous studies in

model organisms such as yeast and C. elegans have clearly demonstrated the involvement

of the SMC5/6 complex in chromatin modeling and genome stability maintenance during

meiosis [1]. Also in mouse and human testes, the localization of SMC5/6 subunits in meiotic

cells has been reported [2-4]. Interestingly, initial protein expression of SMC6 coincides

exactly with spermatogonial differentiation [4]. Yet, the function of SMC5/6 during

spermatogenesis remains largely unknown, highlighting the need for loss-of-function studies.

To this end we applied the novel CRISPR-Cas9 technique to the cultured primary mouse

SSCs, namely male germline stem (GS) cells, to generate spermatogonial cell lines lacking a

functional SMC5/6 subunit. Using our optimized protocol we successfully generated a stable

GS cell line devoid of non-SMC element 2 (NSMCE2) [5]. As a subunit of the SMC5/6

complex, NSMCE2 is a SUMO ligase essential for DNA damage repair in yeast [6]. In

mammalian somatic cell types NSMCE2 has significant phenotypes. In mice, NSMCE2

knockout (KO) leads to embryonic lethality, while it suppresses cancer and aging in adult

mice [7]. Patients with decreased expression of NSMCE2 display primordial dwarfism,

extreme insulin resistance and gonadal failure [8]. We employed CRISPR-Cas9 to generate

a Nsmce2-null human osteosarcoma cell line (U2OS), and found that NSMCE2 helps to

resolve topological stress, for instance replication-induced chromosome entanglements [6].

Somewhat surprisingly, when we switched to GS cells, Nsmce2-/- GS cells did not show any

significant phenotype compared with their Nsmce2+/+ counterparts, and the transcriptome

was only marginally affected by NSMCE2 deprivation [5]. This is in sharp contrast

General discussion 141

with what we and others have previously reported in somatic cell types [6-8]. It has long been

known that germ cells differ from somatic cells in relation to cell biology and (epi)genetic

attributes [9, 10]. Germ cells also have their unique ways to preserve DNA integrity [11]. Our

results again illustrate the unique genomic integrity maintenance pathways active in the

germline.

Notably, a trivial role of NSMCE2 at the spermatogonial stage does not mean that this

protein is absolutely dispensable for the entire process of spermatogenesis. NSMCE2 could

be involved in later developmental stages such as meiosis. This is not unlikely, considering

that yeast meiotic progression requires NSMCE2 function [12], and that patients with

decreased expression of NSMCE2 display primary ovarian failure [8]. Transplantation of

Nsmce2-/- GS cells into sterile recipient testes, which enables further development of these

cells, could provide an answer to this issue. We have also endeavored to generate Smc5-

and Smc6-null spermatogonial cell lines. However, we observed cell death for ˃90% of GS

cells after transfection with Smc5/6-targeting plasmids. Coincidentally, we also failed to

generate Smc6-null U2OS cell lines. We therefore propose that, despite the trivial role for

NSMCE2, the SMC5/6 complex as a whole is indispensable for spermatogonial and perhaps

also somatic cell viability. Given that reliable conditional KO systems are not available for the

earliest stages of spermatogenesis due to the lack of suitable promoters, applying an

inducible Cas9 system to GS cells for controlled loss-of-function studies, as reported recently

by Chen et al. [13], or performing rescue experiments after gene KO, could help to elucidate

the role of SMC5/6 in spermatogonia and spermatogenesis once combined with SSC

transplantation.

SSC differentiation A cornerstone of lifelong spermatogenesis is the perfect balance between SSC self-

renewal and differentiation. Too much self-renewal will lead to tumor-like spermatogonial

clusters while excessive differentiation will deplete the stem cell pool eventually leading to a

total loss of germ cells and infertility [14]. In the mouse, the SSCs are part of the greater pool

of undifferentiated spermatogonia and can divide to form two single spermatogonia or stay

connected via intercellular bridges to eventually form chains of usually 8 or 16 cells [15]. At a

fixed stage of spermatogenesis, and regulated via retinoic acid (RA) signaling, the

undifferentiated spermatogonia develop into differentiating spermatogonia. These

differentiating spermatogonia are no longer dividing “freely” but instead are committed

towards meiosis and subsequent spermiogenesis via a fixed program of divisions and

development. Because differentiation of spermatogonia is strongly negatively correlated with

142 Chapter 6

stem cell capacity [15, 16], unravelling the mechanisms for spermatogonial differentiation is

of great interest and importance.

In vitro cultured SSCs, termed GS cells, provide an advantageous system to

interrogate this matter. We and others have demonstrated that both self-renewal and

differentiation can occur in GS cell culture, and that exogenous factors such as RA can drive

GS cell differentiation in a controlled manner [5, 17, 18]. Previous in vivo studies have shown

that RA plays crucial roles in spermatogonial differentiation and the initiation of meiosis [19].

Consistently, we also found that exposure of GS cells to RA led to the significant

downregulation of SSC markers and genes involved in self-renewal, whereas differentiation

genes were upregulated [5]. The molecular alterations induced in vitro resemble those

occurred during normal spermatogenesis in vivo. Nonetheless, few target genes of RA have

been identified in spermatogenic cells. By high-throughput RNA-sequencing (RNA-seq), we

unraveled the transcriptomic profile of murine differentiating GS cells. Our results are in line

with a recent study that also used RA to induce differentiation of mouse spermatogonia in

vitro [18]. Our RNA-seq data showed a significant transcriptomic change induced by RA-

mediated spermatogonial differentiation. Specifically, differentiation affected a large number

of genes involved in diverse biological processes, e.g. cell adhesion, cell migration, cell cycle,

cell proliferation/differentiation, apoptosis/cell death and transcription regulation. Using our

RNA-seq data, novel markers that can separate undifferentiated from differentiating

spermatogonia can be identified. For instance, using our RNA-seq data, we have identified

and validated PDX1 as a novel marker for mouse undifferentiated spermatogonia.

Furthermore, by way of prevailing genetic modification tools such as RNA interference (RNAi)

or CRISPR-Cas9, and SSC transplantation, the exact function of genes found to be involved

in spermatogonial development can be investigated. In the future, a similar strategy, i.e. the

establishment of a reliable culture system for human SSCs followed by in vitro differentiation

and RNA-seq, could be employed to probe the mechanisms for human spermatogonial

development. Because human spermatogonia are thought to be very diverse, consisting of

various subtypes with differential stem cell capacity, modern single-cell sequencing

techniques need to be optimized and used [20]. Human spermatogonial biology is currently

not sufficiently understood, and more knowledge is required for the future use of human

spermatogonia in the clinic. We have recently successfully determined the transcriptomes of

six successive stages of human spermatogenesis through a combination of laser-capture

microdissection (LCM) and RNA-seq (Jan et al., Development, in press). Further optimization

of our established protocol to allow single-cell analysis would greatly benefit our pursuit of

understanding human spermatogonial biology.


As an alternative for SSC auto-transplantation, in vitro differentiation of SSCs to

functional haploid gametes can potentially be used to treat male infertility. Nonetheless, this

has remained challenging for decades, mainly due to the inability to reproduce complete

meiosis and spermiogenesis in vitro. A breakthrough in this respect was achieved recently,

when complete in vitro meiosis and formation of functional spermatid-like cells from

embryonic stem (ES) cell-derived primordial germ cell-like cells (PGCLCs) were

demonstrated [21]. However, no expression of SSC markers was detected during the

differentiation process, suggesting the failure of those PGCLCs to differentiate into SSCs,

probably due to the absence of growth factors essential for SSC self-renewal (e.g. GDNF

and bFGF). In the future, it would be intriguing to investigate whether SSCs could initiate and

maintain spermatogenesis in this culture system. Alternatively, instead of SSCs, induced-

pluripotent stem cells (iPSCs) could in theory be used to generate male gametes. Either way,

the development and optimization of in vitro gametogenesis methods hold great promise for

use in both fundamental research and the clinic.

The impact of DNA damage on spermatogonia and male fertility Male cancer patients subjected to chemo- or radiotherapy are typically faced with a

declined number of spermatogenic cells and therefore subfertility, largely due to the therapy-

induced DNA lesions and the resulting germ cell apoptosis. Especially, spermatogonia are

very sensitive to DNA damage [11] and severe spermatogonial apoptosis can result in SSC

depletion leading to permanent infertility. On the other hand, failure to repair DNA damage or

induce apoptosis can lead to transmission of genetic mutations or chromosomal aberrations

to the next generation. Hence, in spermatogenic cells, a prompt DNA damage response is of

overriding importance to the maintenance of transgenerational genomic stability. It is

therefore thought that, in comparison to somatic cells, germ cells show differential sensitivity

to DNA damage and hold unique mechanisms for DNA repair [11]. Specifically, they tend to

be more prone to undergo apoptosis, displaying a slower rate of DNA repair as well as a

higher incidence of unrepaired DNA damage in the surviving cells [22, 23].

Interestingly, the undifferentiated spermatogonia, including the SSCs, are more

resistant to DNA damage than differentiating spermatogonia [24-27]. However, the

mechanism that causes differentiating spermatogonia to undergo apoptosis more readily is

currently not known. Our data revealed that, at the transcriptome level, undifferentiated and

differentiating spermatogonia exhibited a highly similar response to IR. Therefore, it is likely

that the increase in radiosensitivity of differentiating spermatogonia is largely caused by

intrinsic properties, such as chromatin architecture, proliferation activity, gene expression

144 Chapter 6

and protein content that are already induced upon differentiation, rather than the additional

genes that are induced by irradiation in both cell types. Differentiating spermatogonia may

readily undergo apoptosis in response to irradiation to prevent the accumulation of genomic

alterations and thus reduce the hereditary risk. In contrast, apoptosis of the undifferentiated

spermatogonia will lead to permanent infertility if the SSCs are eliminated. Undifferentiated

spermatogonia are likely to be “programmed” to ensure continuation of gamete production,

even with a risk of genomic aberrations being transmitted to the offspring. Despite this, the

underlying molecular mechanisms modulating the balance between spermatogonial

apoptosis and survival, particularly in SSCs, remain largely elusive. Various genetically

modified animal models, of which the generation can now be greatly facilitated by novel

techniques such as CRISPR-Cas9, could be employed to further study the unique DNA

damage response and DNA repair pathways in different spermatogenic cells. In addition, our

optimized protocol for genome modification of mouse GS cells has enabled a more efficient

generation of GS cell lines with modified/removed genes involved in the spermatogonial DNA

damage response, facilitating further studies in this field.

SSC transplantation and its clinical translation In 1994, Brinster and colleagues reported successful SSC transplantation [28, 29].

They demonstrated that, upon transplantation into recipient testes, donor SSCs can go

through the blood-testis barrier and relocate to the basement membrane of seminiferous

tubules from where they can initiate and maintain complete spermatogenesis. Later,

transplantation became the standard assay to demonstrate the stem cell capacity of SSCs.

SSC transplantation has now successfully been recapitulated in a wide array of species,

including rodents, domestic animals and even non-human primates, with the generation of

healthy descendants or embryos from donor-derived sperm [30, 31]. The success of SSC

transplantation in various animal models, especially in non-human primates, has become an

important step to consider future use of SSC transplantation in a clinical setting. For instance,

SSC transplantation is currently being investigated as a promising treatment option for

childhood or adult cancer survivors who are at high risk of losing their fertility due to chemo-

or radiotherapy [32]. In the future, SSC transplantation might also be considered to improve

fertility of subfertile men, e.g. males suffering from oligozoospermia or azoospermia.

Moreover, in combination with currently available genome editing tools, SSC culture and

transplantation could be used to prevent transmission of genomic diseases from the affected

father to the offspring [33].

At the moment, apart from ethical and clinical considerations, clinical translation of SSC

transplantation is mainly hindered by technical hurdles. Above all, as shown by previous


animal studies, a sufficient number of transplanted SSCs is a prerequisite for successful

transplantation [34, 35]. Unfortunately, in contrast to most animal studies, typically only a

small testicular biopsy can be obtained from patients. As a consequence, SSCs from the

obtained biopsy must be propagated in vitro prior to transplantation. In rodents, long-term

culture of SSCs is well-established and rodent SSCs can proliferate in vitro for years while

maintaining stem cell capacity [36]. Several groups, including ours, have reported the culture

of human SSCs [37-42]. However, from these studies it is still impossible to determine if truly

functional, i.e. spermatozoa-producing, spermatogonia have been cultured in vitro. First,

there is no reliable molecular marker to distinguish SSCs from other testicular cells in culture.

Second, after xeno-transplantation into immunodeficient mice, only migration of cultured

human cells to the basal membrane of the recipient seminiferous tubules was reported [37,

38]. To unequivocally prove the presence of bona fide human SSCs in culture, de novo

testicular morphogenesis using cultured cells [43, 44] or auto-transplantation to the human

testis will most likely be required.

Once long-term culture of human SSCs has been established and optimized, the

(epi)genetic stability of cultured SSCs before and after transplantation needs to be

guaranteed. Moreover, for cancer patients, lingering cancer cells must be eliminated from the

primary SSC culture before auto-transplantation. Hence, extensive preclinical research is

needed to ensure the safety and efficacy of SSC transplantation.

SSCs and CRISPR-Cas9-mediated germline genomic editing Since SSCs can both initiate and maintain lifelong production of sperm that can be

used to generate offspring, they are an ideal target of gene manipulation to generate

genetically modified animal models. Genome editing used to depend on homologous

recombination which is rather inefficient and time-consuming. Yet, over the last decade,

more efficient techniques such as ZFNs, TALENs and, most recently, the CRISPR-Cas9

system have emerged. These tools use engineered nucleases that are able to trigger DNA

double-strand breaks (DSBs) at specific genomic loci, thereby greatly enhancing the DNA

mutation rate. Of these, CRISPR-Cas9 is currently most extensively used, enabling rapid and

efficient genome editing in a wide range of species and cell types including SSCs. To date,

several groups have reported precise genome modification of rodent SSCs by way of

CRISPR-Cas9 [45-47]. Moreover, after transplantation of genetically modified SSCs, a

genetic disease has been corrected in the offspring [46]. These accomplishments in rodents

bring closer the future clinical use of genetically modified human SSCs to cure

spermatogenic failure, or prevent transmission of genetic disorders to the offspring

146 Chapter 6

[33]. Our optimized protocol to generate genetically modified mouse GS cell lines more

efficiently may contribute to the realization of this goal in the future.

Although one can expect that human SSCs should be amenable to genome

modification, SSCs from large animals such as boars [48] and goats [49] are more refractory

to transfection/transduction than their rodent counterparts. Thus, it is likely that also human

SSCs are very refractory to gene transfection/transduction. In addition, while novel

electroporation devices are increasingly harnessed to transfect SSCs with adequate

efficiency, electroporation typically generates considerable cell death. Moreover, transfection

of SSCs most likely reduces cell proliferation and induces senescence [50]. Together, these

factors weaken the use of transfection/electroporation on the already sparse human SSCs in

culture. Genome modification of human SSCs could also be achieved via viral transduction

[51]. However, residual viral sequences can integrate into the genome of SSCs, raising

serious safety concerns and excluding its clinical application.

For males who wish to prevent the transmission of genomic disorders to the offspring

via transplantation of genetically modified SSCs, more issues need to be taken into

consideration. First, the disease-causing mutation needs to be corrected in SSCs. This can

only be achieved when the CRISPR-Cas9-mediated DSB is repaired via homology-directed

repair (HDR). Unfortunately, the incidence of HDR is rather low [52]. Second, the repaired

SSCs, if any, need to be isolated from the mixed cell population and then expanded from

single cells to establish stable and healthy SSC lines. Even in rodents, this procedure is

technically difficult and time-consuming. We and others have demonstrated that in rodents,

SSC expansion from single cells to a 6-well plate takes 2-3 months [5, 45]. It is expected to

be more time-consuming for human SSCs that have a longer doubling time [53]. Moreover,

although the established healthy SSC lines can be screened for (epi)genetic stability and the

absence of off-target effects, long-term cultured SSCs have been reported to have a

substantially impaired ability to regenerate full spermatogenesis after transplantation [54].

For males who wish to prevent transmission of genetic disorders to their offspring by

transplantation of corrected SSCs, another major barrier remains which is the depletion of

endogenous mutated germ cells before transplantation. Besides being necessary to prevent

mutated sperm from still being produced, endogenous spermatogonia may need to be

eliminated to provide access to the niche for transplanted and genetically corrected

spermatogonia. In rodents, the depletion of endogenous spermatogenesis is typically

achieved by injecting alkylating antineoplastic agents such as busulfan. However, for non-

rodent species the administration of busulfan at dosages to sufficiently eliminate germ cells is


usually lethal [31], making testicular irradiation the only option. However, irradiation of a

man’s testis is accompanied by serious side-effects [33]. Thus, auto-transplantation of

genetically modified human SSCs to produce healthy offspring is not a straightforward

procedure.

Because transplantation of genetically modified SSCs results in germline transmission

of modified genes to the offspring, also ethical issues remain an important topic of discussion.

Many of these discussions are likely to be similar to those that have been vigorously held

over the genome modification of human embryos. Initially, two groups from China reported

genome editing in human zygotes by CRISPR-Cas9, albeit at low efficiency and with a high

incidence of off-target effects [55, 56]. Recently, a study from the US demonstrated highly

effective modification of human embryos virtually without off-target effects [57], although the

validity of this study has been debated [58]. Collectively, these publications received

worldwide media attention and raised intensive social and ethical debates. Interestingly,

recent years witnessed a dramatic switch in terms of the public attitude towards human

germline editing, from the initial complete prohibition [59] to the recent conditional permission

at laboratory levels [60]. Debates on this matter are still ongoing and a broader audience

including patients, clinicians, researchers, ethicists and politicians need to be involved in

future discussions.

Implications for future research A major focus of this study is how genomic integrity maintenance is regulated during

spermatogonial development. To study the role of the SMC5/6 complex in this process we

optimized the use of CRISPR-Cas9 to edit the mouse spermatogonial genome [5]. Because

CRISPR-Cas9-mediated genome editing in combination with SSC transplantation could be

used to cure spermatogenic failure or prevent genetic diseases in the offspring, we also

addressed the clinical prospects of SSC transplantation, with or without germline genomic

editing [33].

Indeed, in vitro propagation of SSCs followed by transplantation could be a promising

treatment option for subfertile men. Nevertheless, unlike the establishment of long-term

culture systems for rodent SSCs, human SSC culture remains inefficient and requires

optimization. Recent publications about long-term culture of undifferentiated spermatogonia

from large animals such as pigs [61] and cattle [62] may provide clues on how to optimize the

culture of human SSCs. Besides, given that human SSCs transplanted to a recipient mouse

testis fail to undergo further differentiation due to their phylogenetic distance, the

spermatogenic ability of cultured human SSCs still needs to be investigated. This could for

instance be done by in vitro differentiation or in vivo developmental assays such as de novo

148 Chapter 6

testicular morphogenesis or auto-transplantation. Future studies should also focus on the

(epi)genetic stability of cultured human SSCs, the fitness of the recipient males subjected to

SSC transplantation, as well as the follow-up of SSC transplantation-generated offspring, if

any. For SSC transplantation with germline genomic editing, much more fundamental and

preclinical work needs to be conducted to guarantee the efficiency and safety of this

technique. Moreover, ethical and societal implications of germline editing need to be probed.

In addition, alternatives for SSC transplantation, such as three-dimensional (3D) or organ

culture, which may potentially allow in vitro differentiation and maturation of human SSCs to

functional sperm, could be developed and optimized.

An issue that cannot be ignored is that the etiology of spermatogenic failure is unknown

in most cases, largely due to the intricate regulatory mechanisms of spermatogenesis. Over

the last decades, research that uses (transgenic) mouse models has greatly extended the

knowledge about the regulatory mechanisms for SSC self-renewal and differentiation and

further spermatogenesis. Nevertheless, due to the phylogenetic distance, these data gained

via mouse studies may not fully translate to the human. An illustration of this point are the

difficulties in culturing human SSCs when using protocols initially developed for the mouse.

An important reason for this is the lack of knowledge about the signals or growth factors that

promote human SSC self-renewal. Establishment of an immortalized human spermatogonial

cell line could provide adequate cells for fundamental mechanistic studies [51]. Also, a recent

paper from our department, which unraveled the transcriptomes of human male germ cells at

distinct developmental stages, could provide clues for the optimization of culture systems for

human SSC proliferation/differentiation (Jan et al., Development, in press). In addition,

previous studies employing mouse models have well demonstrated that epigenetic factors

[10], non-coding RNAs such as miRNA [63] and long non-coding (lnc) RNA [64] play

important roles in regulating SSC fate and spermatogenesis. Single-cell sequencing could be

further exploited to uncover the expression profiles of non-coding RNAs in sub-clusters of

human germ cells. By using the novel CRISPR-Cas9 system or other genetic modification

strategies, their functions may be further annotated. Collectively, it is only when fundamental

studies and preclinical trials are well combined that the development of treatment options for

male subfertility can be accelerated.


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Chapter 7

Summary Samenvatting

156 Chapter 7

Summary

The aim of this thesis was to unravel the mechanisms that determine and regulate the

dynamic response to DNA damage during spermatogonial development, with a specific focus

on the role of the structural maintenance of chromosomes (SMC) 5/6 complex. To this end

we used CRISPR-Cas9-mediated genome editing to remove genes of interest from

spermatogonial stem cells (SSCs) in culture. Because cultured and genetically modified

SSCs can be (auto-)transplanted to recipient donor testes, our optimized method of germline

genome editing evoked clinical and ethical questions that were reviewed at the end of the

thesis. Following is the summary of our findings.

In Chapter 1 we provided an overview of spermatogenic failure, spermatogenesis,

SSCs, DNA damage repair, chromatin dynamics, the SMC5/6 complex and genome editing.

In Chapter 2 we aimed to gain more insights into the molecular mechanisms by which

the SMC5/6 complex works in mammalian cells. We therefore harnessed CRISPR-Cas9 to

generate a human osteosarcoma cell line (U2OS) that lacks NSMCE2 (non-SMC element 2),

a subunit of the SMC5/6 complex described to be essential for DNA repair. Using this cell

line, we were able to find that treatment with the topoisomerase II inhibitor etoposide

triggered an increased sensitivity in cells lacking NSMCE2. In contrast, NSMCE2 appeared

not essential for a proper DNA damage response or cell survival after DNA double-strand

break (DSB) induction by ionizing irradiation (IR). By way of immunoprecipitation and mass

spectrometry, we found that the SMC5/6 complex physically interacts with the DNA

topoisomerase TOP2A. We therefore propose that the SMC5/6 complex functions in

resolving TOP2A-mediated DSB-repair intermediates that are generated during replication.

In Chapter 3 we reported an optimized protocol to generate genetically modified

mouse male germline stem (GS) cell lines using CRISPR-Cas9. By this we generated a GS

cell line devoid of NSMCE2. We found that NSMCE2 was dispensable for proliferation,

differentiation and topological stress relief in mouse GS cells. Moreover, RNA-sequencing

analysis demonstrated that the spermatogonial transcriptome was only minimally affected by

the absence of NSMCE2. Only differential expression of Sgsm1 appeared highly significant,

but with SGSM1 protein levels being unaffected without NSMCE2. Hence, despite the

essential roles of NSMCE2 in somatic cells, genome integrity maintenance in the germline

seems to be regulated differently, without the requirement of NSMCE2.

Summary 157

In Chapter 4 we used in vitro cultured GS cells to investigate the influence of

spermatogonial differentiation on the response to DNA damage. Interestingly, the sensitivity

of spermatogonia to DNA damage increases during differentiation but the mechanisms

responsible for this have not been revealed before. We analyzed the transcriptomes of in

vitro differentiating and undifferentiated spermatogonia with and without IR treatment. At the

RNA level, both spermatogonial subtypes showed a very similar response to IR. When

comparing undifferentiated and differentiating spermatogonia, there were no IR-induced

differentially expressed genes (DEGs) that could be attributed to spermatogonial

differentiation. Nevertheless, in response to IR, differentiating spermatogonia did not

upregulate the DNA damage response protein p53 as much as undifferentiated

spermatogonia. Hence, the difference in radiosensitivity between undifferentiated and

differentiating spermatogonia is largely caused by intrinsic properties, like chromatin

architecture, gene expression, proliferation rate and protein content that are already induced

upon differentiation, rather than additional genes induced by irradiation in both cell types.

In Chapter 5 we reviewed the state of the art with respect to SSC transplantation and

genomic editing using CRISPR-Cas9, followed by envisioning the clinical prospects of SSC

transplantation, with or without genomic editing, to restore male fertility or prevent

transmission of genomic disorders. The clinical and technical hindrances, as well as ethical

issues, were disclosed as well. Despite the requirement of optimization at various levels,

SSC transplantation, with or without genomic editing, is still a promising platform that may be

used in the clinic in the future.

In Chapter 6 we discussed the results obtained in this thesis and uncovered

implications for future (pre)clinical and fundamental research.

158 Chapter 7

Samenvatting

Het doel van dit proefschrift is het ontrafelen van de mechanismen die de veranderlijke

reactie bepalen van zich ontwikkelende spermatogonia op DNA-schade, met een focus op

de rol van het “structural maintenance of chromosomes” (SMC) 5/6 complex in dit proces.

Om dit te bereiken hebben we de CRISPR-Cas9-methode voor genoomediting gebruikt om

relevante genen te verwijderen uit spermatogoniale stamcellen (SSCs). Omdat gekweekte

en genetisch gemodificeerde SSCs getransplanteerd kunnen worden naar een donortestis

roept onze geoptimaliseerde methode voor kiembaanmodificatie ook klinische en ethische

vragen op die beschreven staan aan het eind van dit proefschrift. Nu volgt een samenvatting

van onze bevindingen.

In hoofdstuk 1 geven we een overzicht van spermatogenese, spermatogenetisch falen,

SSCs, reparatie van DNA-schade, dynamiek van chromatine, het SMC5/6 complex en

genoom-modificatie.

In hoofdstuk 2 onderzoeken we de moleculaire mechanismen die de functie van het

SMC5/6 complex bepalen in zoogdiercellen. Hiertoe hebben we CRISPR-Cas9 gebruikt om

non-SMC element 2 (NSMCE2, een onderdeel van SMC5/6 dat essentieel is voor reparatie

van DNA-schade) te verwijderen uit humane osteosarcoma cellen. Gebruik makende van

deze cellijn hebben we gevonden dat gebrek aan NSMCE2 tot gevolg heeft dat cellen

gevoeliger worden voor topologische stress, in dit geval veroorzaakt door de topoisomerase

II-remmer etoposide. NSMCE2 bleek echter niet essentieel voor het repareren van DNA-

schade veroorzaakt door ioniserende straling. We hebben bovendien gevonden dat het

SMC5/6 complex fysiek bindt aan de DNA topoisomerase TOP2A. We stellen daarom voor

dat het SMC5/6 complex bijdraagt aan het repareren van TOP2A-gemedieerde DNA-schade

die veroorzaakt wordt door topologische stress die optreedt gedurende het repliceren van

DNA.

In hoofdstuk 3 beschrijven we een geoptimaliseerd protocol voor het gebruik van

CRISPR-Cas9 voor het maken van genetisch gemodificeerde muizenzaadcel-stamcellijnen

(GS-cellen). Met dit protocol hebben we GS-cellen gemaakt die geen NSMCE2 meer hebben.

Het blijkt dat in deze cellen NSMCE2 niet nodig is voor proliferatie, differentiatie of het

oplossen van topologische stress. Uit transcriptoom-analyse blijkt bovendien dat het

spermatogoniale transcriptoom bijna niet beïnvloed wordt door het afwezig zijn van NSMCE2.

Alleen expressie van het gen Sgsm1 leek significant te verschillen, ware het niet dat het eiwit

SGSM1 onveranderlijk aanwezig bleef. Ondanks de essentiële rol van NSMCE2 in

Samenvatting 159

somatische cellen lijkt het er dus op dat het bewaken van genomische integriteit anders

gereguleerd is in voortplantingscellen, niet noodzakelijkerwijs via NSMCE2.

In hoofdstuk 4 hebben we GS-cellen gebruikt om het gevolg van spermatogoniale

differentiatie op de reactie van deze cellen op DNA-schade te onderzoeken. Het is bekend

dat de stralingsgevoeligheid van spermatogoniën toeneemt als ze differentiëren. Wat hieraan

ten grondslag ligt is echter onbekend. We hebben het transcriptoom geanalyseerd van in

vitro differentiërende en ongedifferentieerde spermatogonia, met en zonder stralingsschade.

Beide spermatogoniale subtypen lieten een sterk vergelijkbaar genexpressiepatroon zien in

reactie op ioniserende straling. In deze vergelijking kwamen er geen differentieel tot

expressie gebrachte genen (DEGs) aan het licht veroorzaakt door spermatogoniale

differentiatie. Desondanks lieten differentiërende spermatogonia een minder sterke verrijking

van het DNA-schade eiwit p53 zien. In het algemeen bleek het verschil in

stralingsgevoeligheid tussen differentiërende en ongedifferentieerde spermatogonia vooral

bepaald te worden door intrinsieke eigenschappen zoals chromatine-architectuur,

genexpressie, proliferatie en eiwitten die al geïnduceerd waren door differentiatie, en niet

door genen die geïnduceerd werden door stralingsschade in beide celtypen.

In hoofdstuk 5 geven we een overzicht van de stand van zaken met betrekking tot

SSC-transplantatie en genoommodificatie middels CRISPR-Cas9, gevolgd door een

beschrijving van de klinische vooruitzichten van SSC-transplantatie (met of zonder

genoommodificatie) voor het herstellen van mannelijke vruchtbaarheid of het voorkomen van

het doorgeven van erfelijke ziekten aan het nageslacht. Klinische en technische problemen

en ethische kwesties zijn hierbij meegenomen. Ondanks dat verdere optimalisatie nog op

alle vlakken nodig is, blijkt SSC-transplantatie, met of zonder genoommodificatie, een

veelbelovende methode die wellicht ooit toepast zal worden in de kliniek.

In hoofdstuk 6 worden de resultaten beschreven in dit proefschrift bediscussieerd,

inclusief implicaties voor mogelijk toekomstig (pre)klinisch en fundamenteel onderzoek.

160 Acknowledgements

Acknowledgements

Over the last 4 years I was honored to join in this group as a PhD student and meet,

communicate and work with these fabulous people. It's somewhat sad that the farewell time

finally comes. Here I would like to convey my sincere gratitude to all the great people that I

have met during my stay in the Netherlands.

First, my promotor, prof. dr. Sjoerd Repping. Sjoerd, thank you very much for accepting

me as a PhD student in your group and thus giving me an opportunity to learn, be trained

and grow professionally. Although not occurring too often over the last 4 years, I do

appreciate each time of our conversations, as they were always stimulating and refreshing.

You always remind me of the big picture of my thesis. Besides, I am impressed by your

dedication to “good science” in the current blundering scientific community. This kind of

scientific attitude will always go along with me in my future journey.

Another person to whom I would like to convey my heartfelt gratitude is my co-promotor,

dr. Geert Hamer. Geert, I am happy that finally I can have an opportunity to let you know how

much I appreciate you and how happy I work with you. Absolutely this thesis would never

come true without you. I will never forget our first meeting in the airport, where you picked me

up upon my first arrival in the Netherlands. I am happy that I could work with you and I was

never pushed but had full liberty to arrange my experiments, which makes me think

independently and grow professionally. Moreover, I got a lot of valuable scientific feedback

from you, and was always reassured under tricky circumstance. Overall, I would be hard-

pressed to express all my gratitude to your understanding and support during my PhD study.

As I will stay in the field of spermatogenesis and stem cells, I am looking forward to our

future collaboration.

Absolutely I will acknowledge my doctorate committee members, dr. ir. W.M. Baarends,

prof. dr. N. Zelcer, prof. dr. C.J.F. van Noorden, dr. N.A.P. Franken and prof. dr. D.G. de

Rooij. Thank you for making up my thesis committee and spending precious time in reading

and judging my thesis.

The persons that I will not forget to acknowledge must include Saskia and Cindy. I am

grateful to you for teaching me important experimental skills and your help in my project!

Apart from technicians, I would like to convey my sincere gratitude to our dear secretary,

Beatrix. Thank you so much for your help in all aspects during my stay and study in the

Netherlands! Also, I appreciate the help and efforts from Aldo, our in-house bioinformatician.

It is your support and help that makes my thesis finally come true!

Acknowledgements 161

My dear colleagues, Callista, Jitske, Sabrina, Joana, Ieva, Myrthe, Robbert, Robin, Majid,

Febilla, Vera, Arno, Kai Mee, Emma, Hajar, Louise, Jan Willem, etc., thank you for your

accompanying during my PhD study! I miss the life that we discuss and have lunch together,

and the memory of our dinners and parties always stays in my mind. I am proud of you and I

wish you all the best in your future career!

I would also not forget the help from other principal investigators in CVV and members

of MORG: Ans, Sebastiaan, Gijs, Carrie, Marie, Remco, Truus and Souad. Thanks a lot for

your useful suggestions and help in my project. To talk with you is always compelling. Indeed,

what I benefit from our talks are not only scientific feedback, but also attitudes towards life

and dilemma.

Coming to a foreign country alone is always accompanied by some extent of

nervousness and apprehension. It’s reassuring to meet and know some people who speak

the same language and share the same culture. Thanks to my housemate Na Zhao, Gang

Wang, Chao Ding and Zemin Ren, I genuinely felt at home when I live with you. I am also

grateful to my dear friends Zongliang, Xiaoxia, Lihui, Xiuping, Jing Zhao, Wanhai, Xiao Yu,

etc. I will never forget the time that we have dinners and travel together, and I sincerely wish

you cheerfulness and success in your future journey!

My scientific career started in 2010 when I became a master student; therefore, I would

like to acknowledge prof. dr. Wenxian Zeng, who is the supervisor for my master’s study. It is

under your guidance and supervision that I gained some basic knowledge about

spermatogonial stem cells and spermatogenesis, which laid the groundwork for my PhD

study. Furthermore, thank you so much for your efforts in my job application. Now it’s with

great joy to rejoin your flourishing group. With your established platform and my experience

gained in these years, I believe we have good chances for significant scientific

accomplishments.

Finally, I would like to devote my infinite gratitude to the most important persons in my

life: my parents, for all the love and support during these years. It goes without saying that

everything in my life would not have been possible without your support and understanding.

Now my only wish for you is that you were healthy and happy every day!

The 4-year PhD study in the Netherlands is a fantastic journey in my life. Not only am I

trained to become an independent researcher, but also know the Dutch, the Netherlands, the

Europe and the world outside of China. Yes, life is often associated with frustration and

adversity. However, this journey is also surrounded by love, hope and a host of wonderful

memories of times, places, people and emotions. With these staying in my heart I will

definitely go ahead, no fear, no hesitation.

162 PhD portfolio

PhD portfolio

PhD period: 1.10.2013-30.9.2017

PhD supervisors: Prof. Dr. Sjoerd Repping

Dr. Geert Hamer

PhD training

Year Workload (Hours/ECTS)

General courses Oral Presentation in English (AMC)

The AMC World of Science (AMC)

Scientific Writing in English for Publication (AMC)

2014

2014

2017

22/0.8

20/0.7

42/1.5

Specific courses Laboratory Animal Science (Utrecht University) DNA Technology (AMC)

Advanced qPCR (AMC)

Bioinformatics Sequence Analysis (AMC)

2014

2014

2015

2016

80/3.0

60/2.1

20/0.7

30/1.1

Seminars, workshops and master classes Journal club

Weekly department seminars (ReproBio)

Progress report seminars

2013-2017

2013-2017

2013-2017

112/4.0

112/4.0

112/4.0

Oral presentations (invited) Bringing CRISPR/Cas9 to aneuploidy and refractory cells:

practical issues and suggestions, AMC CRISPR Symposium

2015

14/0.5

(Inter)national conferences 7th Meeting of the International Network for Young

Researchers in Male Fertility (Elsinore, Denmark) Symposium: Genome on Demand? Exploring the

Implications of Human Genome Editing (Amsterdam)

9th Dutch Society for Stem Cell Research Meeting (Utrecht)

10th Dutch Society for Stem Cell Research Meeting (Utrecht)

2014

2015

2016

2017

16/0.5

16/0.5

8/0.25

8/0.25

Total 672/23.9

About the author 163

About the author

The author of this thesis, Yi Zheng, was born on September 8th, 1988, in Yaan,

Sichuan, China, where is also the hometown of panda. From 2006 to 2010, as an

undergraduate student, he studied Animal Science at Sichuan Agricultural University, in

Sichuan, China.

After his bachelor graduation in June 2010, he enrolled in Northwest A&F University, in

Shaanxi, China, where he became a master student and studied Animal Genetics, Breeding

and Reproduction Science. During the 3-year master study, he focused on characterization

and in vitro culture of spermatogonial stem cells (SSCs) from pigs under the supervision of

Prof. Wenxian Zeng.

In 2013, he obtained his master’s degree and a scholarship from China Scholarship

Council (CSC) to support his PhD study abroad. With the funding and a strong interest in

spermatogenesis and SSCs, in October 2013 he joined Reproductive Biology Laboratory,

Center for Reproductive Medicine, Academic Medical Center, University of Amsterdam, The

Netherlands, as a PhD student under the supervision of Prof. Sjoerd Repping and Dr. Geert

Hamer, where he focused on genome integrity maintenance during spermatogonial

development. The results of the work during his PhD are presented in this thesis.

164 List of publications

List of publications Mulder CL#, Zheng Y#, Jan SZ, Struijk RB, Repping S, Hamer G, van Pelt AM. Spermatogonial stem

cell autotransplantation and germline genomic editing: a future cure for spermatogenic failure and

prevention of transmission of genomic diseases. Human Reproduction Update. 2016 Sep;22(5):561-

73. (#equal contribution)

Zheng Y, Jongejan A, Mulder CL, Mastenbroek S, Repping S, Wang Y, Li J, Hamer G. Trivial role for

NSMCE2 during in vitro proliferation and differentiation of male germline stem cells. Reproduction.

2017 Sep;154(3):81-95.

Verver DE#, Zheng Y#, Speijer D, Hoebe R, Dekker HL, Repping S, Stap J, Hamer G. Non-SMC

Element 2 (NSMCE2) of the SMC5/6 Complex Helps to Resolve Topological Stress. International

Journal of Molecular Sciences. 2016 Oct 26;17(11). pii: E1782. (#equal contribution)

Mulder CL, Catsburg LAE, Zheng Y, de Winter-Korver CM, van Daalen SKM, van Wely M, Pals S,

Repping S, van Pelt AMM. Long-term health in recipients of transplanted in vitro propagated

spermatogonial stem cells. Human Reproduction. 2017 Nov 18:1-10. doi: 10.1093/humrep/dex348.

Zhang P, Chen X, Zheng Y, Zhu J, Qin Y, Lv Y, Zeng W. Long-term propagation of porcine

undifferentiated spermatogonia. Stem Cells and Development. 2017 Aug 1;26(15):1121-1131.

Zhang P, Qin Y, Zheng Y, Zeng W. Phospholipase D family member 6 (PLD6) is a surface marker for

enrichment of undifferentiated spermatogonia in the pre-pubertal boars. Stem Cells and

Development. 2017 Nov 7. doi: 10.1089/scd.2017.0140.

Zheng Y, Zhang Y, Qu R, He Y, Tian X, Zeng W. Spermatogonial stem cells from domestic animals:

progress and prospects. Reproduction. 2014 Feb 3;147(3):R65-74.

Zheng Y, He Y, An J, Qin J, Wang Y, Zhang Y, Tian X, Zeng W. THY1 is a surface marker of porcine

gonocytes. Reproduction, Fertility and Development. 2014;26(4):533-9.

Zheng Y, Tian X, Zhang Y, Qin J, An J, Zeng W. In vitro propagation of male germline stem cells from

piglets. Journal of Assisted Reproduction and Genetics. 2013 Jul;30(7):945-52.

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