Abstract

BACKGROUND

There is an increasing body of evidence that human pluripotent stem cells (hPSCs) are prone to (epi)genetic instability during in vitro culture. This review aims at giving a comprehensive overview of the current knowledge on culture-induced (epi)genetic alterations in hPSCs and their phenotypic consequences.

METHODS

Combinations of the following key words were applied as search criteria: human induced pluripotent stem cells and human embryonic stem cells in combination with malignancy, tumorigenicity, X inactivation, mitochondrial mutations, genomic integrity, chromosomal abnormalities, culture adaptation, aneuploidy and CD30. Only studies in English, on hPSCs and focused on (epi)genomic integrity were included. Further manuscripts were added from cross-references.

RESULTS

Numerous (epi)genetic aberrations have been detected in hPSCs. Recurrent genetic alterations give a selective advantage in culture to the altered cells leading to overgrowth of abnormal, culture-adapted cells. The functional effects of these alterations are not yet fully understood, but suggest a (pre)malignant transformation of abnormal cells with decreased differentiation and increased proliferative capacity.

CONCLUSIONS

Given the high degree of (epi)genetic alterations reported in the literature and altered phenotypic characteristics of the abnormal cells, controlling for the (epi)genetic integrity of hPSCs before any clinical application is an absolute necessity.

Introduction

Two major types of pluripotent stem cells have been described in the human: embryonic stem cells (hESCs) and induced pluripotent stem cells (hiPSCs), and neither of these cells occur naturally in vivo. ESCs are obtained through the in vitro culture of the outgrowth of an inner cell mass of a preimplantation embryo. hiPSCs, on the other hand, are reprogrammed somatic cells, in which the pluripotent capacity is induced artificially.

hESCs and hiPSCs are very attractive cells to the field of regenerative medicine because of their capacity to differentiate virtually into any cell type of the adult human body, including gametes. Another use of these cells that is increasingly receiving attention is as research models. hESCs and hiPSCs can provide further knowledge when combined with the existing animal models for human monogenic diseases, or can be used as a first step for drug development and testing or toxicology tests.

There is an increasing body of evidence that hESCs and more recently, hiPSCs, are prone to genetic and epigenetic instability. It is not yet understood how these abnormalities arise, how they affect the cells and to which measure they hamper their clinical applicability and reliability as research models.

It is the objective of this review to provide a comprehensive overview of the current knowledge on the instability of the genome and epigenome of human pluripotent stem cells (hPSCs) and the impact this has on their phenotype.

Methods

We searched the Pubmed database using combinations of the following key words: human induced pluripotent stem cells and human embryonic stem cells in combination with malignancy, tumorigenicity, X inactivation, mitochondrial mutations, genomic integrity, chromosomal abnormalities, culture adaptation, aneuploidy and CD30. On March 2012, this retrieved 338 papers. We added 22 manuscripts from the authors' individual collections and other sources.

We initially excluded 53 articles, including 38 papers that reported work that was not carried out on hESCs or hiPSCs and 2 papers that were not in English. There were 306 full-text articles assessed for their relevance to this review. Finally, 111 manuscripts were included.

Results

Integrity of the nuclear genome

The type and size of the chromosomal abnormalities that have been described in hPSCs greatly varies depending on the methods that were used in the study. Whilst G-banding can readily identify subpopulations of abnormal cells, it is limited in the detection of small genetic changes. In contrast, comparative genomic hybridization (CGH), array-based CGH and single nucleotide polymorphism (SNP) arrays offer higher levels of resolution and reproducibility, though they suffer from rapid loss of sensitivity when applied to mosaic cultures. Conversely, interphase fluorescent in situ hybridization (FISH) is sensitive in identifying cells carrying specific chromosome rearrangements, amplifications and deletions but provides only information for one or a few loci (Baker et al., 2007; Josephson, 2007).

The first abnormal hESCs karyotypes were detected while performing G-banding and CGH analysis during long-term culture (Draper et al., 2004; Inzunza et al., 2004; Rosler et al., 2004). Since then, a large body of evidence has shown that hiPSCs frequently acquire aneuploidies, large chromosomal gains and losses, and chromosomal rearrangements as well as small abnormalities below the resolution of G-banding, with ∼60% of these abnormalities being recurrent (Taapken et al., 2011). Table I shows an overview of all the reported abnormalities in hPSCs.

Table I

Overview of chromosomal abnormalities in human pluripotent stem cells.

Line Passage number Karyotype analysis Abnormality References 
HS237 p61 CGH, G-banding 46,X,idic(X)(q21) Inzunza et al. (2004) 
H7.s6 p60  46,XX,der(6)t(6;17)(q27;q1)  
H7.s9 p29 G-banding, FISH 47,XX,+17 Draper et al. (2004) 
H7.s14 p27  46,XX,+17 [76%]  
 p22+17  46,XX,+17 [95%]  
H14.s9 p41  47,XY,+17  
H1.1A P75/0  47,XY,+12  
H1.1B P75/30  44,X,-Y,der(6)t(6;17;17)(6pterq15::17q25q25.3::17q11.2qter),del(10) (p11.2),+12,der(18)t(17;17;18)(17qterq11.2::17q25.3q25.1::18q23pter)  
HUES-1 p39 G-banding 46,XX,add(2)(q?) Cowan et al. (2004) 
HUES-3 p33–47  47,XY,+12/46,XY  
HUES-4 p29–47  47,XY,+12/46,XY  
HUES-5 p14  46,XX,inv(9)  
HUES-9 p9  46,XX,inv(9)  
H1, H7, H9 p28–37, p>48 Karyotype, microarray Trisomy 20 Rosler et al. (2004) 
hESC5 p38 G-banding 46,XY[14/28]/46,XY,t(X,17)(p11,p13)[14/28] Buzzard et al. (2004) 
p42  46,XY,t(X,17)(p11,p13)[20/20]  
H7-s6 NI G-banding 46,XX,der(6)t(6;17)(q27;q1) Andrews et al. (2005) 
 47,XX,+1,der(6)t(6;17)(q27;q1)  
 48,XX,+1,der(6)t(6;17)(q27;q1),+8  
BG01 p41 FISH, SNP dup(17)(17q) Maitra et al. (2005) 
H1 >80  del(18)(18q21.32q22.1)  
H7 p63  dup(20)(q11.21)  
H9 p78  dup(1)(q31.1q31.3)  
SA002/2.5 p205  dup(8)(q24.21),-13  
H7 >100 G-banding 47,XX,+1,der(6)t(6;17)(q27;q1) Enver et al. (2005) 
BG01 p65 G-banding, FISH 48,XY,+12,+17[80%]/49,XXY,+12,+17[20%] Mitalipova et al. (2005) 
BG02 P70  47,XY,+17[71%]/47,XY,+inv(17)(q11.2q21)[29%]  
BG02 p68  50,XXY,+12,+14,+17[85%]/51,XXY,+12,+14,+17,+20[15%]  
SA002 p14, p114 MLPA, FISH 47,XX,+13 Darnfors et al. (2005) 
SA121 p73, p117  46,-6p  
HS181  Q-banding, FISH, SKY 46,XX[5]/47,XX,del(7)(q11.2),+i(12)(p10)[21] Imreh et al. (2006) 
   XX,del(7)(q11.2),+i(12)(p10)  
HES-2 p51/28 G-banding 46,XX,dup(1)(q21.1q32.1) Herszfeld et al. (2006) 
HES-3 p55/10  46,XX/47,XX,+12  
p37/32  46,XX,t(1;6)(p22;q15)  
SA002 p19, p23 G-banding, FISH, aCGH 47,XX,+3 Caisander et al. (2006) 
H1 p114 SNP 48,XY,+12,+17[24]/49,XXY,+12,+17[3]/46,XY[3] Baker et al. (2007) 
H1a p53  47,XY,+12[3]/47,XY,+17[3]/48,XY,+12,+17[14]  
H14 p37  47,XY,+17[14]/46,XY[6]  
H14a p121  48,XY,+12,+der(17)del(17)(p12p13.3)hsr(17)(p11.2)[14]/48,XY,+i(12)(p10),+der(17)del(17)(p12p13.3)hsr(17)(p11.2)[6]  
H7 p209  49,XXX,+add(1)(p36),der(6)t(6;17)(q27;q1),+20[14]/48,XX,+add(1)(p36),der(6)t(6;17)(q27;q1),+20[6]  
H7a p51  47,XX,+17[17]/46,XX[3]  
HUES5 p19  52,XXXX,+inv(9)(p11q13),+12,+14,+17[6]/50,XX,+inv(9),+12,+14,+17[6]/46,XX,inv(9)[5]  
HUES6 p21  46,XX,der(13)t(13;17)(q13;q12)[20]  
HUES7 p20  48,XY,+12,+17[18]/50,XXY,+12,+17,+mar[14]/49,XY,+7,+12,+17[2]/46,XY[2]  
HUES10 p18  50,XXY,+9,+12,+17[15]/46,XY[5]  
HUES13 p33  48,XY,+7,+12,der(22;17)(q10;q10)[2]/46,XY,der(17;22)(q10;q10)[28]  
HUES14 p32  46,XY,der(10)t(10;17)(q26;q21)[20]  
HUES17 p24  48,XY,+12,+17[15]/46,XY[15]  
Shef1 p87  47,XY,+3[14]/46,XY[6]  
 p106  47,XY,+i(12)(p10)[9]/46,XY[11]  
Shef4 p79  47,XY,+17[20]/46,XY[10]  
Shef5 p39  47,XX,t(1;11)(p36;q13),trp(17)(p11.2p11.2),+20[19]/46,XX[1]  
HS181  FISH Trisomy 12 Gertow et al. (2007) 
H9 p46 G-banding 46,XX [18/20]/45,XX,-9 [1/20]/45,XX,-5[1/20] Chan et al. (2008) 
H9-Trypsin-Adapted  p61/13  49,XX,+9,+12,+17[8/10]/48,XX,+12,+17[2/10]  
p64/16  46,XX[2/14]/50,XXX,+9,+12,+17[8/14]/49,XX,+9,+12,+17[2/14]/48,XX,dic(1;3)(q42;p21),+9,+12,+17[1/14]/50,XX,+5,+9,+12,+17[1/14]  
H-Re-collagenase p62/6/8  46,XX[18/21]/49,XX,+9,+12,+17[2/21]/45,XX,-13[1/21]  
p69/6/15  46,XX[19/20]/45,XX,-9[1/20]  
BGO1-trypsin-adapted p40/7  47,XY,+17[13/20]/48,XY,+12,+17[6/20]/47,XY,+i(11)(q10),+12,+17,-13[1/20]  
BGO1-re-collagenase p55/8/14  47,XY,+17[10/22]/48,XY,+12,+17[10/22]/47,XY,del(9)(p12),+12,+17[1/22]/47,XY,+12,+17,−19[1/22]  
     
HSF6-trypsin/EDTA p30  48,XX,+17,+20  
VUB01 p135,276 G-banding, FISH, aCGH 46,XY,dup(20)(q11.21) Spits et al. (2008) 
VUB01.2 p55  46,XY,-18  
VUB02 p252  46,XY,dup(20)(q11.21)  
VUB03-DM1 p98,152  46,XX,dup(20)(q11.21)  
VUB04-CF p66,117  46, XX, dup(5)(q14.2qter),del(18)(q21.2qter)  
VUB06.2 p21  47,XX,+17  
VUB07.2 p77  46,XX,dup(20)(q11.21)  
p122,131  46,XX,dup(3)(q26.33q27.2),dup(20)(q11.21)  
VUB08_MFS p49  47,XX,+22  
VUB09_FSHD.2 p40  46,XX,dup(8)(q24.21)  
p88  46,XX,dup(11)(q24.2q25)  
p100  46,XX,dup(11)(q24.2q25),dup(20)(q11.21)  
VUB13_FXS p43  46,XX,dup(5)(q21.3qter),del(18)(q12.1qter)  
p45  45,X,-X,dup(5)(q21.3qter),del(18)(q12.1qter)  
VUB20_CMT1A p7,18  46,XX,dup(5)(q13.2)  
VUB26 p10  46,XX,dup(7)(q33qter),del(18)(q23qter)  
SA01 p83 BAC, aCGH dup(20)q11.21 Lefort et al. (2008) 
H1 p41  dup(20)q11.21  
VUB05-HD p103  dup(20)q11.21  
FY-hES-5 p27 FISH 46,XX,+13,der(13)(q10;q10) Sun et al. (2008) 
FY-3PN p44  69,XXX  
  aCGH, oligo-microarray 46,XY,dup(20)(q11.21),del(6)(p21.32),del(7)(q34),del(22)(q11.23) Wu et al. (2008) 
HSF6   46,XX,dup(1)(q21.3),del(4)(q13.2),del(6)(p21.32),del(19)(p12),del(22)(q11.23)  
HS181 P71/17 G-banding, aCGH, SKY 47,XX,+12[26%] Catalina et al. (2008) 
P71/21  47,XX,+12[31%],+20  
P71/30  47,XX,+12[89%],+20  
SHEF3 p51/10  47,+14[15–36%]  
SNUhES4 p21+6 G-banding, FISH 47,XY,+12/46,XY Seol et al. (2008) 
NI  47,XXY/46,XY  
p7+39  47,XXY  
NA NA  47,XY,+8/46,Y,i(X)(q10)/47,XXY  
NA NI G-banding 47,XY,+16 Liao et al. (2009) 
HS306 p35 SNPs dup(4)(q22.1q22.2) Närvä et al. (2010) 
CCTL-12 p143  dup(5)(q14.2q14.3)  
I3.2 p55  dup(10)(q11.21q11.22)  
H7-s6 p128,132  del(10)(q21.2q21.3)  
H7-s6 Tera p125,127  del(10)(q21.2q21.3)  
HS401 p53  del(15)(q11.2)  
H1 p61  del(15)(q11.2)  
H9 p25,34  dup(15)(q11.2)  
HS237 p135  dup(18)(q21.32q21.33)  
CCTL-14 p38,49  dup(20)(q11.21)  
H1 p56 aCGH, SNPs 46,dup(1)(q32.2),dup(22)(q12.2) Hovatta et al. (2010) 
HiPSC18 p58  46,XY[11/20]/47,XY,+12[9/20] Mayshar et al. (2010) 
 p63  47,XY,+12  
chHES-3 p34 G-banding, FISH 46,XX,dup(1)(p32p36) Yang et al. (2010) 
p44–99  46,XX,dup(1)(p32p36),t(1;6;4)(q25;q23;p16), ins(4;1)(p16;q21q25),t(7;8)(q32;q13)  
p142–153  47,XXX,dup(1)(p32p36),t(1;6;4)(q25;q23;p16), ins(4;1)(p16;q21q25),t(7;8)(q22;q22)  
p188–197  46,XX,dup(1)(p32p36),t(1;6;4)(q25;q23;p16), ins(4;1)(p16;q21q25),t(7;8)(q32;q13),+mar(15?)  
AA-03 p16 SNPs 46,XX,t(2;19)(p21;p13.1)[3]/46,XX[27] Amps et al. (2011) 
AA-05 p16  47,XX,+11[9]/46,XX,del(10)(p12)[6]/44,XX,+add(1)(p1),-9,der(15)t(9;15)(p12;p11)[2]/46,XX[9]  
p30  46,XX,der(9)t(1;9)(q12;q11),der(15)t(9;15)(q12;p11.2)[30]  
AA-07 p10  47,XX,+17[4]/46,XX[24]  
p36  47,XX,+17[9]/46,XX[21]  
BB-04 p94  46,X,der(Y)t(Y;15)(q12;q21)[30]  
BB-09 p93  47,XY,+12[4]/46,XY[26]  
C-02 p17, p61  47,XX,+13[30]  
CC-05 p25, p36  46,XX,r(18)(p11.3q23)[30]  
D-01 p22  47,XX,+i(12)(p10),der(18)t(17;18)(q1;q23)[7]/46,XX[23]  
p51  47,XX,+i(12)(p10),der(18)t(17;18)(q1;q23)[11]/46,XX[19]  
F-01 p84  46,XX,der(15)t(15;17)(p11;q21)[10]  
F-02 p39  46,XY,add(20)(p11.2)[2]/46,XY[27]  
p56  46,XY,add(20)(p11.2)[1]/46,XY[29]  
p72  46,XY,add(20)(p11.2)[2]  
F-03 p202  46,XX,add(10)(p11)[20]  
p152  46,XX,add(10)(p11)[12]/46,XX[10]  
FF-02 p68  46,XX,dup(13)(q12q32)[5]/46,XX[25]  
HH-01 p14  59∼64,XX,add(X)(p11.2),add(X)(q2),+der(1)t(1;15)(q44;q15), +der(1)t(1;15)(q44;q15),add(p22),i(2)(q10),+der(2)t(2;3)(q23;p13), der(3)t(3;21)(p13;q22),t(4;17)(q28;q25),+5,+add(6)(p25),+7,+9,+10, +add(11)(q23),+12,der(13)t(6;13)(q21;q34),+14,+17,+19, +der(20)t(8;20)(q12;q13.3),+der(22)t(3;22)(p13;p11),+mar1,+mar2[cp10]  
I-01 p54  47,XY,+1[2]/46,XY[27]  
p211  47,XY,+17[6]/46,XY[24]  
p128  46,XY,add(22)(q13)[3]/46,XY[27]  
I-03 p29  47,XY,+8[30]  
p103  47,XY,+8[1]/46,XY[29]  
I-04 p115  47,XX,+12[2]/46,XX[28]  
J-01 p71  46,XY,der(10)t(1;10)(q13;p13),der(22)t(20;22)(q11.2;p11)[17]/46,XY,der(7)t(7;17)(q36;q21),der(10)t(1;10)(q13;p13), der(22)t(20;22)(q11.2;p11)[4]/46,XY,der(17)t(17;17)(p13;q11)[4]/ 47,XY,+i(12)(p10),der(17)t(17;17)(p13;q11)[4]/46,XY[4]  
J-02 p96  46,XY,inv(11)(q21q23)[2]/46,XY[28]  
K-01 p160  47,XY,+17[10]/46,XY[20]  
K-05 p92  47,XX,t(1;11)(p?36;q13),trp(17)(p11.2),+20[30]  
L-02 p35  47,XY,+20[30]  
   47,XY,+20[27]/46,XY[3]  
 p42  47,XY,+20[30]  
LL-02 p22, p64  48,XY,+add(17)(p11.2),+add(17)(p11.2)[30]  
NN-01 p9  46,XX,del(10)(p11.2)[26]/46,XX,der(10)t(10;17)(p11.2;q25), del(17)(q25)[5]/46,XX,der(10)t(10;10)(p13;q11.2)[3]/46,XX[1]  
p35  46,XX,der(7)(t(1;7)(q21;q11.2)[6]/46,XX,idem,add(22)(q13.2)[7]/46,XX,idem,der(16)add(16)(q2)add(16)(p13)[4]/46,XX,idem,add(22)(q13.2),add(17)(q25)[2]/46,XX,idem,der(7)add(7)(p22)trp(7)(q32q34),add(22)(q13.3)[8]  
NN-02 p19, p26  46,XX,i(7)(p10)[27]/46,XX,add(7)(q1)[3]  
NN-03 p9  47,XX,i(7)(p10),+16[5]/46,XX,i(7)(p10)[25]  
NN-07 p15, p25  46,XX,i(7)(p10),inv(9)(p13q13)[30]  
PP-107 p25, p81  45,XX,der(21;22)(q10;q10[30]  
Q-02 p81  47,XX,+der(1)(t(1;1)(p?21.2;q11)[29]/46,XX[1]  
Q-04 p66  47,XY,+20[10]  
RR-01 (iPS) p16, p110  47,XX,+12  
RR-03 (iPS) p9  46,XX,inv(5)(p13q22)[15]/46,XX[15]  
RR-05 (iPS) p9  47,XX,+12[1]/46,XX[29]  
S-04-004-DL p115  50,XXYY,+12,+17,der(18)t(13;18)(q12;q23)[30]  
SS-02 p20  46,X,add(X)(p1)[4]/46,XX[26]  
p35  46,X,add(X)(p1)[22]/46,XX[8]  
U-04 p78  47,XY,+12[3]/46,XY[27]  
UU-01 p41  46,XX,dup(20)(q11.21)[3]/47,XX,+12[2]/46,XX[6]  
 p104  46,XX,dup(20)(q11.21)[6]/46,XX[17]  
UU-02 p14  45,X[25]/46,XX[5]  
 p41  45,X[30]  
UU-03 p25  48,XX,+12,+17[10]/46,XX,del(6)(q15q23),del(18)(q21.3)[26]  
X-04 p136  46,XX,der(18)t(5;18)(q15;q21)[30]  
YY-01 p43  47,XX,+i(17)(p10)[30]  
MR90-YZ1 p24 G-banding, FISH, aCGH Trisomy 12 Martins-Taylor et al. (2011) 
HDFa-TK2 p31  46,XX,t(6;16)(q15;q13),t(13;14)(q14.1;q24.3)  
HiPSC line NI  dup(1)(q31.3)/dup(2)(p11.2)/del(17)(q21.1)  
ICF-TK4   46,XX,der(21)t(17;21)(p11.2;q22.3)  
iPSC#1  G-banding 47,XX,+1[85%] Elliott et al. (2011) 
iPSC#3   47,XX,+20[35%]  
iPSC#1  aCGH dup(1)(p36.33-q44),dup(3)(q26.33),dup(5)(q23.1–q33.2),del(7)(q31.33), dup(8)(q24.21), dup(9)(q31.2), del(11)(p15.4),dup (14)(q23.2), dup (20)(q11.21),dup (22)(q13.31)  
iPSC#3   dup(2)(q37.2),del(3)(p24.3),dup( 3)(q26.33),dup(5)(q35.3),dup(8)(q24.21), dup(9)(q31.2),dup(20)(p13–q13.33),dup (22)(q13.33)  
Line Passage number Karyotype analysis Abnormality References 
HS237 p61 CGH, G-banding 46,X,idic(X)(q21) Inzunza et al. (2004) 
H7.s6 p60  46,XX,der(6)t(6;17)(q27;q1)  
H7.s9 p29 G-banding, FISH 47,XX,+17 Draper et al. (2004) 
H7.s14 p27  46,XX,+17 [76%]  
 p22+17  46,XX,+17 [95%]  
H14.s9 p41  47,XY,+17  
H1.1A P75/0  47,XY,+12  
H1.1B P75/30  44,X,-Y,der(6)t(6;17;17)(6pterq15::17q25q25.3::17q11.2qter),del(10) (p11.2),+12,der(18)t(17;17;18)(17qterq11.2::17q25.3q25.1::18q23pter)  
HUES-1 p39 G-banding 46,XX,add(2)(q?) Cowan et al. (2004) 
HUES-3 p33–47  47,XY,+12/46,XY  
HUES-4 p29–47  47,XY,+12/46,XY  
HUES-5 p14  46,XX,inv(9)  
HUES-9 p9  46,XX,inv(9)  
H1, H7, H9 p28–37, p>48 Karyotype, microarray Trisomy 20 Rosler et al. (2004) 
hESC5 p38 G-banding 46,XY[14/28]/46,XY,t(X,17)(p11,p13)[14/28] Buzzard et al. (2004) 
p42  46,XY,t(X,17)(p11,p13)[20/20]  
H7-s6 NI G-banding 46,XX,der(6)t(6;17)(q27;q1) Andrews et al. (2005) 
 47,XX,+1,der(6)t(6;17)(q27;q1)  
 48,XX,+1,der(6)t(6;17)(q27;q1),+8  
BG01 p41 FISH, SNP dup(17)(17q) Maitra et al. (2005) 
H1 >80  del(18)(18q21.32q22.1)  
H7 p63  dup(20)(q11.21)  
H9 p78  dup(1)(q31.1q31.3)  
SA002/2.5 p205  dup(8)(q24.21),-13  
H7 >100 G-banding 47,XX,+1,der(6)t(6;17)(q27;q1) Enver et al. (2005) 
BG01 p65 G-banding, FISH 48,XY,+12,+17[80%]/49,XXY,+12,+17[20%] Mitalipova et al. (2005) 
BG02 P70  47,XY,+17[71%]/47,XY,+inv(17)(q11.2q21)[29%]  
BG02 p68  50,XXY,+12,+14,+17[85%]/51,XXY,+12,+14,+17,+20[15%]  
SA002 p14, p114 MLPA, FISH 47,XX,+13 Darnfors et al. (2005) 
SA121 p73, p117  46,-6p  
HS181  Q-banding, FISH, SKY 46,XX[5]/47,XX,del(7)(q11.2),+i(12)(p10)[21] Imreh et al. (2006) 
   XX,del(7)(q11.2),+i(12)(p10)  
HES-2 p51/28 G-banding 46,XX,dup(1)(q21.1q32.1) Herszfeld et al. (2006) 
HES-3 p55/10  46,XX/47,XX,+12  
p37/32  46,XX,t(1;6)(p22;q15)  
SA002 p19, p23 G-banding, FISH, aCGH 47,XX,+3 Caisander et al. (2006) 
H1 p114 SNP 48,XY,+12,+17[24]/49,XXY,+12,+17[3]/46,XY[3] Baker et al. (2007) 
H1a p53  47,XY,+12[3]/47,XY,+17[3]/48,XY,+12,+17[14]  
H14 p37  47,XY,+17[14]/46,XY[6]  
H14a p121  48,XY,+12,+der(17)del(17)(p12p13.3)hsr(17)(p11.2)[14]/48,XY,+i(12)(p10),+der(17)del(17)(p12p13.3)hsr(17)(p11.2)[6]  
H7 p209  49,XXX,+add(1)(p36),der(6)t(6;17)(q27;q1),+20[14]/48,XX,+add(1)(p36),der(6)t(6;17)(q27;q1),+20[6]  
H7a p51  47,XX,+17[17]/46,XX[3]  
HUES5 p19  52,XXXX,+inv(9)(p11q13),+12,+14,+17[6]/50,XX,+inv(9),+12,+14,+17[6]/46,XX,inv(9)[5]  
HUES6 p21  46,XX,der(13)t(13;17)(q13;q12)[20]  
HUES7 p20  48,XY,+12,+17[18]/50,XXY,+12,+17,+mar[14]/49,XY,+7,+12,+17[2]/46,XY[2]  
HUES10 p18  50,XXY,+9,+12,+17[15]/46,XY[5]  
HUES13 p33  48,XY,+7,+12,der(22;17)(q10;q10)[2]/46,XY,der(17;22)(q10;q10)[28]  
HUES14 p32  46,XY,der(10)t(10;17)(q26;q21)[20]  
HUES17 p24  48,XY,+12,+17[15]/46,XY[15]  
Shef1 p87  47,XY,+3[14]/46,XY[6]  
 p106  47,XY,+i(12)(p10)[9]/46,XY[11]  
Shef4 p79  47,XY,+17[20]/46,XY[10]  
Shef5 p39  47,XX,t(1;11)(p36;q13),trp(17)(p11.2p11.2),+20[19]/46,XX[1]  
HS181  FISH Trisomy 12 Gertow et al. (2007) 
H9 p46 G-banding 46,XX [18/20]/45,XX,-9 [1/20]/45,XX,-5[1/20] Chan et al. (2008) 
H9-Trypsin-Adapted  p61/13  49,XX,+9,+12,+17[8/10]/48,XX,+12,+17[2/10]  
p64/16  46,XX[2/14]/50,XXX,+9,+12,+17[8/14]/49,XX,+9,+12,+17[2/14]/48,XX,dic(1;3)(q42;p21),+9,+12,+17[1/14]/50,XX,+5,+9,+12,+17[1/14]  
H-Re-collagenase p62/6/8  46,XX[18/21]/49,XX,+9,+12,+17[2/21]/45,XX,-13[1/21]  
p69/6/15  46,XX[19/20]/45,XX,-9[1/20]  
BGO1-trypsin-adapted p40/7  47,XY,+17[13/20]/48,XY,+12,+17[6/20]/47,XY,+i(11)(q10),+12,+17,-13[1/20]  
BGO1-re-collagenase p55/8/14  47,XY,+17[10/22]/48,XY,+12,+17[10/22]/47,XY,del(9)(p12),+12,+17[1/22]/47,XY,+12,+17,−19[1/22]  
     
HSF6-trypsin/EDTA p30  48,XX,+17,+20  
VUB01 p135,276 G-banding, FISH, aCGH 46,XY,dup(20)(q11.21) Spits et al. (2008) 
VUB01.2 p55  46,XY,-18  
VUB02 p252  46,XY,dup(20)(q11.21)  
VUB03-DM1 p98,152  46,XX,dup(20)(q11.21)  
VUB04-CF p66,117  46, XX, dup(5)(q14.2qter),del(18)(q21.2qter)  
VUB06.2 p21  47,XX,+17  
VUB07.2 p77  46,XX,dup(20)(q11.21)  
p122,131  46,XX,dup(3)(q26.33q27.2),dup(20)(q11.21)  
VUB08_MFS p49  47,XX,+22  
VUB09_FSHD.2 p40  46,XX,dup(8)(q24.21)  
p88  46,XX,dup(11)(q24.2q25)  
p100  46,XX,dup(11)(q24.2q25),dup(20)(q11.21)  
VUB13_FXS p43  46,XX,dup(5)(q21.3qter),del(18)(q12.1qter)  
p45  45,X,-X,dup(5)(q21.3qter),del(18)(q12.1qter)  
VUB20_CMT1A p7,18  46,XX,dup(5)(q13.2)  
VUB26 p10  46,XX,dup(7)(q33qter),del(18)(q23qter)  
SA01 p83 BAC, aCGH dup(20)q11.21 Lefort et al. (2008) 
H1 p41  dup(20)q11.21  
VUB05-HD p103  dup(20)q11.21  
FY-hES-5 p27 FISH 46,XX,+13,der(13)(q10;q10) Sun et al. (2008) 
FY-3PN p44  69,XXX  
  aCGH, oligo-microarray 46,XY,dup(20)(q11.21),del(6)(p21.32),del(7)(q34),del(22)(q11.23) Wu et al. (2008) 
HSF6   46,XX,dup(1)(q21.3),del(4)(q13.2),del(6)(p21.32),del(19)(p12),del(22)(q11.23)  
HS181 P71/17 G-banding, aCGH, SKY 47,XX,+12[26%] Catalina et al. (2008) 
P71/21  47,XX,+12[31%],+20  
P71/30  47,XX,+12[89%],+20  
SHEF3 p51/10  47,+14[15–36%]  
SNUhES4 p21+6 G-banding, FISH 47,XY,+12/46,XY Seol et al. (2008) 
NI  47,XXY/46,XY  
p7+39  47,XXY  
NA NA  47,XY,+8/46,Y,i(X)(q10)/47,XXY  
NA NI G-banding 47,XY,+16 Liao et al. (2009) 
HS306 p35 SNPs dup(4)(q22.1q22.2) Närvä et al. (2010) 
CCTL-12 p143  dup(5)(q14.2q14.3)  
I3.2 p55  dup(10)(q11.21q11.22)  
H7-s6 p128,132  del(10)(q21.2q21.3)  
H7-s6 Tera p125,127  del(10)(q21.2q21.3)  
HS401 p53  del(15)(q11.2)  
H1 p61  del(15)(q11.2)  
H9 p25,34  dup(15)(q11.2)  
HS237 p135  dup(18)(q21.32q21.33)  
CCTL-14 p38,49  dup(20)(q11.21)  
H1 p56 aCGH, SNPs 46,dup(1)(q32.2),dup(22)(q12.2) Hovatta et al. (2010) 
HiPSC18 p58  46,XY[11/20]/47,XY,+12[9/20] Mayshar et al. (2010) 
 p63  47,XY,+12  
chHES-3 p34 G-banding, FISH 46,XX,dup(1)(p32p36) Yang et al. (2010) 
p44–99  46,XX,dup(1)(p32p36),t(1;6;4)(q25;q23;p16), ins(4;1)(p16;q21q25),t(7;8)(q32;q13)  
p142–153  47,XXX,dup(1)(p32p36),t(1;6;4)(q25;q23;p16), ins(4;1)(p16;q21q25),t(7;8)(q22;q22)  
p188–197  46,XX,dup(1)(p32p36),t(1;6;4)(q25;q23;p16), ins(4;1)(p16;q21q25),t(7;8)(q32;q13),+mar(15?)  
AA-03 p16 SNPs 46,XX,t(2;19)(p21;p13.1)[3]/46,XX[27] Amps et al. (2011) 
AA-05 p16  47,XX,+11[9]/46,XX,del(10)(p12)[6]/44,XX,+add(1)(p1),-9,der(15)t(9;15)(p12;p11)[2]/46,XX[9]  
p30  46,XX,der(9)t(1;9)(q12;q11),der(15)t(9;15)(q12;p11.2)[30]  
AA-07 p10  47,XX,+17[4]/46,XX[24]  
p36  47,XX,+17[9]/46,XX[21]  
BB-04 p94  46,X,der(Y)t(Y;15)(q12;q21)[30]  
BB-09 p93  47,XY,+12[4]/46,XY[26]  
C-02 p17, p61  47,XX,+13[30]  
CC-05 p25, p36  46,XX,r(18)(p11.3q23)[30]  
D-01 p22  47,XX,+i(12)(p10),der(18)t(17;18)(q1;q23)[7]/46,XX[23]  
p51  47,XX,+i(12)(p10),der(18)t(17;18)(q1;q23)[11]/46,XX[19]  
F-01 p84  46,XX,der(15)t(15;17)(p11;q21)[10]  
F-02 p39  46,XY,add(20)(p11.2)[2]/46,XY[27]  
p56  46,XY,add(20)(p11.2)[1]/46,XY[29]  
p72  46,XY,add(20)(p11.2)[2]  
F-03 p202  46,XX,add(10)(p11)[20]  
p152  46,XX,add(10)(p11)[12]/46,XX[10]  
FF-02 p68  46,XX,dup(13)(q12q32)[5]/46,XX[25]  
HH-01 p14  59∼64,XX,add(X)(p11.2),add(X)(q2),+der(1)t(1;15)(q44;q15), +der(1)t(1;15)(q44;q15),add(p22),i(2)(q10),+der(2)t(2;3)(q23;p13), der(3)t(3;21)(p13;q22),t(4;17)(q28;q25),+5,+add(6)(p25),+7,+9,+10, +add(11)(q23),+12,der(13)t(6;13)(q21;q34),+14,+17,+19, +der(20)t(8;20)(q12;q13.3),+der(22)t(3;22)(p13;p11),+mar1,+mar2[cp10]  
I-01 p54  47,XY,+1[2]/46,XY[27]  
p211  47,XY,+17[6]/46,XY[24]  
p128  46,XY,add(22)(q13)[3]/46,XY[27]  
I-03 p29  47,XY,+8[30]  
p103  47,XY,+8[1]/46,XY[29]  
I-04 p115  47,XX,+12[2]/46,XX[28]  
J-01 p71  46,XY,der(10)t(1;10)(q13;p13),der(22)t(20;22)(q11.2;p11)[17]/46,XY,der(7)t(7;17)(q36;q21),der(10)t(1;10)(q13;p13), der(22)t(20;22)(q11.2;p11)[4]/46,XY,der(17)t(17;17)(p13;q11)[4]/ 47,XY,+i(12)(p10),der(17)t(17;17)(p13;q11)[4]/46,XY[4]  
J-02 p96  46,XY,inv(11)(q21q23)[2]/46,XY[28]  
K-01 p160  47,XY,+17[10]/46,XY[20]  
K-05 p92  47,XX,t(1;11)(p?36;q13),trp(17)(p11.2),+20[30]  
L-02 p35  47,XY,+20[30]  
   47,XY,+20[27]/46,XY[3]  
 p42  47,XY,+20[30]  
LL-02 p22, p64  48,XY,+add(17)(p11.2),+add(17)(p11.2)[30]  
NN-01 p9  46,XX,del(10)(p11.2)[26]/46,XX,der(10)t(10;17)(p11.2;q25), del(17)(q25)[5]/46,XX,der(10)t(10;10)(p13;q11.2)[3]/46,XX[1]  
p35  46,XX,der(7)(t(1;7)(q21;q11.2)[6]/46,XX,idem,add(22)(q13.2)[7]/46,XX,idem,der(16)add(16)(q2)add(16)(p13)[4]/46,XX,idem,add(22)(q13.2),add(17)(q25)[2]/46,XX,idem,der(7)add(7)(p22)trp(7)(q32q34),add(22)(q13.3)[8]  
NN-02 p19, p26  46,XX,i(7)(p10)[27]/46,XX,add(7)(q1)[3]  
NN-03 p9  47,XX,i(7)(p10),+16[5]/46,XX,i(7)(p10)[25]  
NN-07 p15, p25  46,XX,i(7)(p10),inv(9)(p13q13)[30]  
PP-107 p25, p81  45,XX,der(21;22)(q10;q10[30]  
Q-02 p81  47,XX,+der(1)(t(1;1)(p?21.2;q11)[29]/46,XX[1]  
Q-04 p66  47,XY,+20[10]  
RR-01 (iPS) p16, p110  47,XX,+12  
RR-03 (iPS) p9  46,XX,inv(5)(p13q22)[15]/46,XX[15]  
RR-05 (iPS) p9  47,XX,+12[1]/46,XX[29]  
S-04-004-DL p115  50,XXYY,+12,+17,der(18)t(13;18)(q12;q23)[30]  
SS-02 p20  46,X,add(X)(p1)[4]/46,XX[26]  
p35  46,X,add(X)(p1)[22]/46,XX[8]  
U-04 p78  47,XY,+12[3]/46,XY[27]  
UU-01 p41  46,XX,dup(20)(q11.21)[3]/47,XX,+12[2]/46,XX[6]  
 p104  46,XX,dup(20)(q11.21)[6]/46,XX[17]  
UU-02 p14  45,X[25]/46,XX[5]  
 p41  45,X[30]  
UU-03 p25  48,XX,+12,+17[10]/46,XX,del(6)(q15q23),del(18)(q21.3)[26]  
X-04 p136  46,XX,der(18)t(5;18)(q15;q21)[30]  
YY-01 p43  47,XX,+i(17)(p10)[30]  
MR90-YZ1 p24 G-banding, FISH, aCGH Trisomy 12 Martins-Taylor et al. (2011) 
HDFa-TK2 p31  46,XX,t(6;16)(q15;q13),t(13;14)(q14.1;q24.3)  
HiPSC line NI  dup(1)(q31.3)/dup(2)(p11.2)/del(17)(q21.1)  
ICF-TK4   46,XX,der(21)t(17;21)(p11.2;q22.3)  
iPSC#1  G-banding 47,XX,+1[85%] Elliott et al. (2011) 
iPSC#3   47,XX,+20[35%]  
iPSC#1  aCGH dup(1)(p36.33-q44),dup(3)(q26.33),dup(5)(q23.1–q33.2),del(7)(q31.33), dup(8)(q24.21), dup(9)(q31.2), del(11)(p15.4),dup (14)(q23.2), dup (20)(q11.21),dup (22)(q13.31)  
iPSC#3   dup(2)(q37.2),del(3)(p24.3),dup( 3)(q26.33),dup(5)(q35.3),dup(8)(q24.21), dup(9)(q31.2),dup(20)(p13–q13.33),dup (22)(q13.33)  

In the column passage: for cells cultured sequentially in different culture conditions, the number of passages in each condition are separated by a /.

In the column: abnormality the different karyotypes are separated by / and the number of metaphases with a given karyotype is indicated between [ ]. NI, not indicated in the manuscript.

Aneuploidy

In hESCs, acquired aneuploidy of all chromosomes but four has been reported at least once. Gains of chromosome 12 (Cowan et al., 2004; Draper et al., 2004; Mitalipova et al., 2005; Baker et al., 2007; Gertow et al., 2007; Catalina et al., 2008) and 17 (Draper et al., 2004; Mitalipova et al., 2005; Baker et al., 2007) have been repeatedly reported as the most frequent karyotypic changes in hESCs. Recently, the International Stem Cell Initiative screened 125 hESC lines worldwide at early and late passages and highlighted that althought most hESC lines remained karyotypically normal, there was a progressive tendency to acquire extra chromosomes after long-term culture, commonly affecting chromosomes 1, 12, 17 and 20 (Amps et al., 2011). In addition, the acquisition of additional copies of chromosome X has also been observed by different research groups (Mitalipova et al., 2005; Baker et al., 2007; Seol et al., 2008; Sun et al., 2008), in one case in up to one-third of their lines (Baker et al., 2007).

hiPSCs also frequently acquire aneuploidies during long-term culture (Chin et al., 2009; Ben-David et al., 2010; Mayshar et al., 2010; Martins-Taylor et al., 2011; Taapken et al., 2011). Conversely, although gain of chromosome 12 is also predominant in hiPSCs (Elliott et al., 2010; Mayshar et al., 2010; Amps et al., 2011; Martins-Taylor et al., 2011; Taapken et al., 2011), trisomy 17 has never been identified in hiPSCs and trisomy 8 is found more frequently in hiPSCs than in hESCs (Taapken et al., 2011).

Structural variants

Chromosomal structural variants have also been repeatedly reported in hiPSCs, with their sizes varying according to the resolution of the method used in the study. Although numerous translocations, inversions, losses and gains have been reported (Table I), only losses on chromosomes 10p13pter, 18q21qter and 22q13qter and gains on chromosomes 1, 12, 17 and 20 recurrently appear in hESCs. Recurrent chromosome 1 amplification mainly involve two regions: 1p36 (Baker et al., 2007; Yang et al., 2010) and 1q31.3 (Maitra et al., 2005; Herszfeld et al., 2006), while gains of chromosomes 12 and 17 affect the entire p and the q arms, respectively (Maitra et al., 2005; Baker et al., 2007; Amps et al., 2011) and the gains on chromosome 20 are mostly restricted to 20q11.21 (Amps et al., 2011). In hiPSCs, gains of 1q31.3 and 17q21.1 and loss of 8q24.3 are also recurrently detected (Martins-Taylor et al., 2011).

It appears that the occurrence of specific genomic aberrations is evolutionary conserved. One of the hotspots of chromosomal aberrations in mouse PSC is a gain of 11qE2, which is fully syntenic to the common aberration 17q25 in hiPSCs; while rhesus macaque PSCs recurrently acquire a gain in chromosome 16q, syntenic to the hotspot in human 17q (Ben-David and Benvenisty, 2012).

Mechanisms of origin, low-grade mosaicism and selective advantage of chromosomal abnormalities

Whole chromosome gains and losses in cultured cells can be explained by anaphase lagging or chromosome non-disjunction during mitotic division, while structural variants can be caused by DNA damage. Albeit that hPSCs are very proficient in the fast repair of DNA damage, and cells that do not repair on time will differentiate or die (Momcilovic et al., 2010; for a review, see Tichy, 2011), the accuracy of such repair is not always guaranteed (Bogomazova et al., 2011). The repair of DNA double strand breaks (DSBs) by homologous recombination could explain the loss of heterozygosity and structural variants may arise from DSB repair by non-allelic homologous recombination, non-homologous end joining or microhomology-mediated end joining (Hastings et al., 2009; Kidd et al., 2010).

It is not known how frequently hESCs spontaneously mutate through either abnormal chromosome segregation or acquiring small amplifications or deletions. Nevertheless, it is possible to gain some insight by considering the existing data on low-grade mosaicism in hESCs, despite the fact that the results are not always concordant.

In the work of Inzunza et al. (2004), single-cell CGH analysis was used to investigate the karyotype of three hESC lines. This also allowed the authors to obtain information on low-grade mosaicism in the culture. The 24 cells analysed from two of the lines showed a completely balanced chromosomal content, while in the third line, five out of 24 cells carried a monosomy X. Rosler et al. (2004) carried out G-banding on three hESC lines in a feeder-free culture at numerous time points. They reported that 20% of the cultures contained at least some aneuploid cells, and the frequency of aneuploidy did not appear to increase with continuous subculturing of the cells. Similarly, Catalina et al. (2008) detected trisomy 14 in SHEF-3 at a relatively constant mosaicism rate of 15–36% during prolonged culture. Forsyth et al. (2006) found that hESCs undergo DNA breaks, gaps and exchanges in 0.14–0.44% of studied metaphases, but that these did not seem to lead to gross structural or numerical chromosomal aberrations, as evidenced by the normal karyotypes they found in Giemsa-stained and G-banded metaphases. Conversely, Lim et al. (2011) found that 4–20% of Giemsa-stained hESCs metaphases are structurally abnormal and 10–25% are aneuploid. Finally, Peterson et al. found that 18–35% of the metaphases of hiPSCs were mosaic aneuploid. Strikingly, most of the abnormal metaphases showed losses, frequently of several chromosomes, which contrasts with the fact that the majority of abnormal karyotypes described in the literature consist of chromosome gains (Peterson et al., 2011). It is likely that the majority of these changes do not confer any selective advantage and remain in culture in a state of random drift or are selected out due to a deleterious effect. In any case, if the highest estimations of low-grade mosaicism are correct, hESCs appear to be quite prone to chromosome segregation errors.

Abnormalities acquired during long-term culture most likely start as a single random event in one cell in the dish, but some have the potential to take over the culture due to selective advantage.

For instance, hESCs carrying a trisomy 12 have a proliferative advantage and noticeable shortening in population doubling time (Cowan et al., 2004), leading to the mutant cells taking over the culture (Imreh et al., 2006; Catalina et al., 2008). Imreh et al. (2006) showed that in a mosaic culture, the population of cells carrying a trisomy 12 increases from 30% at passages 33–68% at passage 72. Catalina et al. (2008) found that the karyotypically normal hESC line HS181 acquired chromosomal changes shortly after being transferred to a feeder-free culture system. A gain of chromosome 12 was first detected at passage 17 in 26% of the cells and the percentage of the variant cells gradually increased to 31% at passage 21 and 89% at passage 30. It has been suggested that the selective advantage of the gain of chromosome 12 is due to the presence of the pluripotency regulators NANOG and GDF3 on this chromosome (Draper et al., 2004; Baker et al., 2007). The overexpression of NANOG and GDF3 that was found in the hiPSC18 line with trisomy 12 (Mayshar et al., 2010) suggests that the selective advantage mechanisms for this trisomy are identical in hESCs and hiPSCs.

Similarly, hESCs commonly acquire additional copies or partial gain on 17q during prolonged culture (Draper et al., 2004; Mitalipova et al., 2005; Baker et al., 2007). In the work of Draper et al. (2004), trisomy 17 was detected in two different sublines of H7 and one subline of H14 after few months in culture. Particularly, FISH analysis on one H7 subculture showed trisomy 17 in 76% of the cells at passage 22 and in 95% of the cells after an additional 17 passages. Interestingly, trisomy of mouse chromosome 11, which is syntenic to human chromosome 17, has been found in almost 20% of murine ESC lines (Sugawara et al., 2006). The non-random gain of chromosome 17q material indicates that this region encompasses genes that are important for cell survival and proliferation. It has been suggested that BIRC5 (also called survivin), a gene involved in apoptosis, plays a key role in explaining the chromosome 17 trisomies (Draper et al., 2004; Baker et al., 2007).

Concerning small structural variants, so far, gain of 20q11.21 is the commonest and probably the most significant recurrent subkaryotypic abnormality observed in hPSCs. It has been detected in over 20% of the hESC lines worldwide (Amps et al., 2011) and is repeatedly reported by different research groups (Maitra et al., 2005; Lefort et al., 2008; Spits et al., 2008; Wu et al., 2008; Elliott et al., 2010; Närvä et al., 2010). Furthermore, 20q11.21 gain is also common in human cancers (Scotto et al., 2008; Beroukhim et al., 2010; Tabach et al., 2011), which strongly suggests a selective advantage for this mutation both in hPSCs and cancer cells. The minimal amplicon of this mutation is 0.55 MB and includes the genes HM13, ID1 and BCL2L1 (Amps et al., 2011). Notably, ID1 has been found to promote self-renewal in stem cells (Romero-Lanman et al., 2012) and BCL2L1 encodes the anti-apoptotic protein Bcl-xL. In some cases, the region of amplification also encompasses DNMT3B, an important pluripotency-associated gene (Spits et al., 2008; Närvä et al., 2010; Martins-Taylor et al., 2011). Finally, duplications of other regions containing pluripotency genes such as MYC at 8q24.21 and SOX2 at 3q26.3–q27 have also been detected in hiPSCs (Elliott et al., 2010) and hESCs (Spits et al., 2008).

Integrity of the mitochondrial genome

Mitochondrial DNA (mtDNA) is a small (16.6-kb) circular and self-replicating molecule that encodes 13 essential proteins of the mitochondrial oxidative phosphorylation complexes, 2 rRNA and 22 tRNA genes. mtDNA mutations are random and occur more frequently compared with genomic DNA mutations due to the presence of free-radical generating enzymes, potentiated by poor DNA repair mechanisms and the lack of protective histones. Undifferentiated hESCs contain only a few immature mitochondria but with differentiation the number of mitochondria increases and their morphology undergoes dramatic functional changes and becomes that of mature cells. Upon reprogramming, mitochondria undergo a complex remodelling and acquire the mitochondrial embryonic-like state, thus, hiPSCs show a mitochondrial complement similar to that of natural hESCs (Suhr et al., 2010).

Very few data are available on the stability of the mitochondrial genome in hPSCs. Using a mitochondrial resequencing oligonucleotide array, Maitra et al. (2005) identified six heteroplasmic mutations occurring in two of the nine hESC lines (Maitra et al., 2005). hiPSCs also seem to harbour a large number of acquired mtDNA point mutations, both hetero- and homoplasmic, that do not appear to affect their pluripotent capacity (Prigione et al., 2011).

Biomarkers of genetic instability

Because the commonly used cytogenetic methods for the detection of karyotypically abnormal cells are capable only of low-throughput analysis on small numbers of cells, there is a general interest in the identification of biomarkers of genetic instability in hESCs and hiPSCs. This would greatly facilitate the development of culture methods that preserve genomic integrity, and would eliminate abnormal cells from cultures prior to their use in a clinical or research setting.

In 2006, Herszfeld et al. (2006) reported that CD30, a member of the tumour necrosis factor receptor superfamily, was expressed on chromosomally abnormal but not on normal hESCs, and that CD30 expression protected hESCs against apoptosis. CD30 seemed a very promising marker as it is a cell surface protein that could be used in FACS sorting of normal and abnormal cells. Nevertheless, several reports soon followed, reporting that CD30 could be found in hESCs with both a normal and abnormal karyotype (Lagarkova et al., 2008; Thomson et al., 2008; Harrison et al., 2009; Mateizel et al., 2009), and that CD30 disappeared upon differentiation, making it rather a marker of undifferentiated hESCs than of transformation (Lagarkova et al., 2008; Mateizel et al., 2009). No correlation was found with the method of passaging (Thomson et al., 2008) or the levels of apoptosis (Harrison et al., 2009) but it was observed that hESCs grown in mTESR and knock-out serum replacement medium expressed CD30 (Lagarkova et al., 2008; Mateizel et al., 2009), whereas hESCs in medium containing fetal calf serum did not show CD30 expression (Mateizel et al., 2009). Finally, the same group that initially reported CD30 as a biomarker published a study that identified ascorbate as the compound that activates CD30 by promotor demethylation (Chung et al., 2010).

More recently, Gerwe et al. (2011) compared the membrane proteomic patterns of karyotypically normal and abnormal hESCs using a high-throughput approach. Their data showed that each hESC line had their own membrane proteomic signature but unfortunately failed to identify a biomarker specific to abnormal cells.

X chromosome inactivation

X chromosome inactivation (XCI), the transcriptional silencing in females of the majority of the genes of one X chromosome, is one of the earliest epigenetic marks that appear during mammalian development. XCI is initiated in the X-inactivation centre of the X chromosome, which contains a gene encoding X-inactive specific transcript (XIST). The XIST gene, which is expressed solely from the inactivated X chromosome, is crucial for XCI because its non-coding RNA directly interacts with and coats the inactive X chromosome. After the completion of XCI, the two X chromosomes in female cells are distinguished by differential transcription of the XIST gene, DNA methylation and epigenetic chromatin modifications such as histone H3 lysine 27 trimethylation (H3K27me3), histone H4 lysine 20 methylation (H4K20me1) and core histone variant macroH2A1 (Brockdorff, 2002).

In hESCs, the inactivation status of the X chromosome seems to be not only hyper-variable between different lines but also between subcultures and even different passages of the same line (Hall et al., 2008; Shen et al., 2008; Silva et al., 2008; Liu et al., 2011). An interesting example is the NIH-approved hESC line H9, in which XCI has been studied in several different laboratories with very variable findings (see Table II).

Table II

Determination of X chromosome inactivation profiles in different sublines of H9, as reported by different groups.

 XCI status XIST expression Cot-1 exclusion H3K27me3; H4K20me1 macroH2A Barr body Reference 
Undifferentiated cells No XCI Negative NA NA NA NA Dhara and Benvenisty (2004
EB differentiation Random XCI Positive NA NA NA NA  
Trophoblast differentiation Skewed XCI Positive NA NA NA NA  
Undifferentiated cells XCI Positive Yes NA Negative Yes Hoffman et al. (2005) 
Spontaneous differentiation (colony periphery) XCI Positive Yes NA Positive Yes  
EB differentiation XCI Positive NA NA Highly positive NA  
Neural progenitors XCI Positive NA NA Highly positive NA  
Undifferentiated cells (H9-Ware P31-35) No XCI Negative (∼16% positive) NA NA NA NA Hall et al. (2008) 
Spontaneous differentiation (H9-Ware P31-35) XCI Positive NA NA NA NA  
Undifferentiated cells (H9-Stein P48) XCI Negative NA H3K27me3 positive Positive Yes Hall et al. (2008) 
Spontaneous differentiation XCI Negative NA H3K27me3 positive Positive Yes  
Undifferentiated cells XCI Negative Yes H3K27me3 negative NA NA Silva et al. (2008) 
Spontaneous differentiation XCI Negative NA NA NA NA  
Undifferentiated cells (different sublines) Skewed XCI 1-100% positive NA 1–100% positive, coupled with XIST Idem H3K27me3/H4K20me1 + in early passaged XIST cells NA Shen et al. (2008) 
 XCI status XIST expression Cot-1 exclusion H3K27me3; H4K20me1 macroH2A Barr body Reference 
Undifferentiated cells No XCI Negative NA NA NA NA Dhara and Benvenisty (2004
EB differentiation Random XCI Positive NA NA NA NA  
Trophoblast differentiation Skewed XCI Positive NA NA NA NA  
Undifferentiated cells XCI Positive Yes NA Negative Yes Hoffman et al. (2005) 
Spontaneous differentiation (colony periphery) XCI Positive Yes NA Positive Yes  
EB differentiation XCI Positive NA NA Highly positive NA  
Neural progenitors XCI Positive NA NA Highly positive NA  
Undifferentiated cells (H9-Ware P31-35) No XCI Negative (∼16% positive) NA NA NA NA Hall et al. (2008) 
Spontaneous differentiation (H9-Ware P31-35) XCI Positive NA NA NA NA  
Undifferentiated cells (H9-Stein P48) XCI Negative NA H3K27me3 positive Positive Yes Hall et al. (2008) 
Spontaneous differentiation XCI Negative NA H3K27me3 positive Positive Yes  
Undifferentiated cells XCI Negative Yes H3K27me3 negative NA NA Silva et al. (2008) 
Spontaneous differentiation XCI Negative NA NA NA NA  
Undifferentiated cells (different sublines) Skewed XCI 1-100% positive NA 1–100% positive, coupled with XIST Idem H3K27me3/H4K20me1 + in early passaged XIST cells NA Shen et al. (2008) 

XCI, X chromosome inactivation; NA, not available.

There is now a general consensus to divide hESC sublines into three classes according to their XCI status (Silva et al., 2008). Class I cells possess two active X chromosomes and inactivate one of the X chromosomes upon differentiation. In the undifferentiated state they do not express XIST or other XCI markers such as H3K27me3, H4K20me1 and macroH2A1. Class II cells show XIST expression and other XCI marks in both the undifferentiated and the differentiated state. Cells of class III have lost XIST coating and do not show other XCI marks. Moreover, they do not reactivate XIST expression upon differentiation (Hall et al., 2008; Shen et al., 2008; Silva et al., 2008). However, they do retain an inactivated X chromosome as demonstrated by the presence of Cot-1 RNA exclusion.

Bruck and Benvenisty (2011) performed a meta-analysis comprising the expression of the entire set of X-genes and could further divide the class II and class III sublines into lines with full and partial inactivation. Moreover, they found that the partial inactivation of the X chromosome always involved the region of the chromosome surrounding the XIST transcription site.

It has been suggested that these three classes could represent an epigenetic progression of hESC cultures with class I being the closest approximation of ground state pluripotency, a delicate epigenetic state prone for progression to class II due to suboptimal culture conditions, while class III would characterize culture-adapted pluripotent cells (see Fig. 1) (Hall et al., 2008; Shen et al., 2008; Silva et al., 2008; Lengner et al., 2010). In support of this hypothesis are the results of Shen et al. who reported that subcultures exhibiting excessive cell death during passaging and displaying a morphologically abnormal nucleus tended to lose XCI marks and thus progressed from class II to class III XCI (Shen et al., 2008). Lengner et al. (2010) found that hESC derivation at 5% (physiologic) O2 not only helped to maintain pluripotency and suppress differentiation but also resulted in preventing precocious XCI. Chronic exposure to 20% (atmospheric) O2, on the other hand, led to irreversible XCI. Initiation of XCI was also observed in response to cellular stress induced by harsh freeze–thaw cycles or through induction by stress-inducing compounds, but could be prevented by the addition of antioxidants or the small molecules sodium butyrate and 3-deazaneplanocin A to the culture media (Lengner et al., 2010; Diaz Perez et al., 2012). These molecules could also be applied to reprogramme class II cells to the class I XCI state; however, this strategy was not successful for cells in the third XCI state (Diaz Perez et al., 2012).

Figure 1

Overview of the epigenetic progression X chromosome inactivation (XCI) in undifferentiated hESC. Class I cells possess two active X chromosomes (Xa), inactivating one of the X chromosomes only at differentiation. Class II cells have inactivated one of the X chromosomes, characterized by a dense XIST coating and presence of repressing histone modifications, while class III cells still have an inactive X chromosome (Xi) but have lost most or all of the epigenetic marks associated with XCI. This loss of epigenetic marks often results in partial reactivation of the inactivate X chromosome.

Figure 1

Overview of the epigenetic progression X chromosome inactivation (XCI) in undifferentiated hESC. Class I cells possess two active X chromosomes (Xa), inactivating one of the X chromosomes only at differentiation. Class II cells have inactivated one of the X chromosomes, characterized by a dense XIST coating and presence of repressing histone modifications, while class III cells still have an inactive X chromosome (Xi) but have lost most or all of the epigenetic marks associated with XCI. This loss of epigenetic marks often results in partial reactivation of the inactivate X chromosome.

One could argue that, whereas in mice embryos it has been shown that X reactivation occurs in the inner cell mass (ICM), in the human embryo XIST expression, and thus XCI persists in the blastocyst stage (van den Berg et al., 2009). This would thus suggest that cells within the human ICM have undergone XCI and thus, that the ground state in hESCs would be represented by class II cells, rather than class I cells. A recent study by O'Leary et al. (2012) investigated the XCI status during the transition of the inner cell mass towards a first hESC colony. In the earliest outgrowths, termed post-ICM intermediate, they consistently observed clear nuclear H3K27me3 foci, indicative for XCI. However, in two out of four established female lines, these foci could no longer be found, whereas differentiation of these lines resulted in reappearance of the H3K27me3 foci. This would suggest that X reactivation takes place during early hESC derivation (O'Leary et al., 2012). Strikingly, many studies on XCI in hESC report cultures exhibiting non-random XCI patterns (Shen et al., 2008; Liu and Sun 2009; Dvash et al., 2010; Lengner et al., 2010), whereas adult human females display random XCI, both in embryonic as well as in extra-embryonic tissues (Moreira de Mello et al., 2010). There are two plausible explanations for this phenomenon. First, recent studies suggest that an imprinted mechanism of X inactivation may take place during human preimplantation development, as is known in the mouse (van den Berg et al., 2011), while XCI is only completed at around the time of the blastocyst stage embryo (van den Berg et al., 2009). It is not known how many cells of the preimplantation embryo are involved in the derivation of hESCs but it is possible that only a limited number of cells attribute to an hESC outgrowth. Therefore, the random or non-random XCI pattern of an hESC line could be directly related to the cells it was originally derived from and could reflect the epigenetic status of the embryo at the time of derivation and/or the limited number of ancestor cells needed to derive an hESC line. This hypothesis could be supported by the fact that a skewed XCI pattern can already be observed before passage 15 (Dvash et al., 2010).

Another explanation may be a possible growth advantage of cells that inactivated a specific X chromosome. Lengner et al. (2010) found that hESC lines derived and cultured at physiological oxygen levels had no XCI in the undifferentiated state and acquired random XCI upon differentiation. However, after prolonged culture of the same line at atmospheric oxygen conditions, the undifferentiated cells displayed skewed XCI patterns. These results support the idea that hESCs undergo XCI in response to stress induced by suboptimal culture conditions and that the best-adapted cells would be clonally selected. Ben-Yosef et al. (2012) found a significant overrepresentation of female lines when performing a meta-analysis on a total of 545 hESC lines. This overrepresentation was found to originate from the actual derivation process rather than from unequal representation of male and female embryos. The authors suggested a possible XCI-related growth advantage as one of the possible explanations for these striking results.

Although hiPSCs are derived from somatic cells in which XCI is already established, they also display variable XCI patterns. Some studies found that hiPSCs display a non-random XCI pattern and therefore concluded that hiPSCs retain the inactive X chromosome from their somatic cell source (Tchieu et al., 2010; Amenduni et al., 2011; Ananiev et al., 2011; Pomp et al., 2011; Cheung et al., 2012; Mekhoubad et al., 2012), while others reported the derivation of hiPSCs that had reactivated the inactive X and displayed two active X chromosomes (Kim et al., 2009; Marchetto et al., 2010). Bruck and Benvenisty (2011) studied X-linked gene expression in 10 female hiPSC lines and could divide them, as they did for hESCs, into three categories: hiPSC lines with no XCI, with partial XCI or with full XCI. A recent study of Mekhoubad et al. (2012) demonstrated that during culture ‘erosion’ of XCI may occur, characterized by loss of XIST expression and foci of H3K27me3, as well as transcriptional derepression of several genes of the inactivated X chromosome (Mekhoubad et al., 2012). As, to our knowledge, there are no reports of hiPSC lines displaying a random XCI pattern, it is unclear whether the hiPSC lines that show full or partial XCI have not or only partially been reactivated, or whether they did reactivate the somatic inactivated X chromosome upon reprogramming and underwent XCI similar to their embryo-derived counterparts.

Whether the XCI state influences the (epi)genetic or functional properties of hPSCs is still a question. In the undifferentiated state, hESC lines in all three classes of XCI express a broad range of pluripotency markers and no or very low lineage-specific markers (Silva et al., 2008). Also the global gene expression or epiproteomic signature of hESC lines does not seem to alter before or after spontaneous completion of XCI during culture (Hoffman et al., 2005; Tanasijevic et al., 2009). Moreover, the preservation of normal imprinting patterns was reported for hESC lines without XCI and with both random and non-random XCI (Liu and Sun, 2009; Lengner et al., 2010).

However, Silva et al. (2008) reported that, upon differentiation in embryoid bodies, class I and II cells showed fair to robust embryoid body growth with the clear presence of lineage-specific markers, whereas for class III cells, embryoid bodies showed varying phenotypes with often scant or poor growth and in one case poor expression of endodermal markers. In another study, class III lines were reported to display an altered response to BMP4 upon trophoblast differentiation (Tanasijevic et al., 2009). The XCI state might thus have little effect in undifferentiated hESCs but a more direct influence upon differentiation of hESCs.

Abnormal DNA methylation

In the first report on epigenetic changes during culture of hESCs, the authors found promoter methylation changes in 3 out of the 14 analysed genes (Maitra et al., 2005). Since then, several groups have reported that undifferentiated hESCs and hiPSCs carry abnormal CpG island methylation patterns and that these changes are stably passed on to the differentiated cells (Bibikova et al., 2006; Shen et al., 2006; Allegrucci et al., 2007; Calvanese et al., 2008; Doi et al., 2009; Amps et al., 2011; Lister et al., 2011; Nishino et al., 2011; Nazor et al., 2012). Whilst it appears that all hESCs undergo overall CpG methylation changes with time in culture, most of the changes have been reported to happen during the earliest stages post-derivation (Allegrucci et al., 2007), and there does not seem to be a group of genes that changes consistently during long-term culture (Bibikova et al., 2006; Amps et al., 2011). On the other hand, hiPSCs undergo extensive epigenetic changes during reprogramming, and gradually converge to an hESC-like epigenome with time in culture (Nishino et al., 2011). Nevertheless, several global methylation studies have shown that a number of regions remain differently methylated between hESCs and hiPSCs, even after long-term culture (Doi et al., 2009; Lister et al., 2011; Nishino et al., 2011).

The study of Doi et al. (2009) shows that some loci in hiPSC cells remain incompletely reprogrammed, whereas others are aberrantly reprogrammed. They also identified 71 differentially methylated regions (DMRs) between hESCs and hiPSCs, 51 showing hypermethylation and 20 showing hypomehtylation. Lister et al. (2011) showed that hiPSCs display significant reprogramming variability, including somatic memory and aberrant reprogramming of DNA methylation. Differentiation of these cells shows that errors in CpG methylation are transmitted at a high frequency, providing an hiPSC reprogramming signature that is maintained after differentiation. Furthermore, they identified 1175 DMRs between hESCs and hiPSCs that were different at least in one hiPSC or hESC line. An interesting finding in this study is that, whilst somatic cells contain negligible levels of cytosine methylation in non-CpG sites, in hESCs and hiPSCs 20–30% of their methylated cytosines are non-CpG, making this a specific trait of hPSCs (Ramsahoye et al., 2000; Lister et al., 2009; Lister et al., 2011). Although the non-CpG methylation levels and distribution were very similar between hESCs and hiPSCs on a whole-genome scale, they found very large non-CpG DMRs in hiPSCs. In general, the DMRs were due to hypomethylation in the hiPSCs, and proximal to centromeres and telomeres. The authors propose that the localized failure to restore non-CpG methylation in these large regions could be mechanistically linked to the presence of particular histone modifications that impart a regional chromatin conformation that is refractive to remethylation at non-CpG sites during reprogramming. Supporting this hypothesis, they identified significant regional enrichment of H3K9me3 that was spatially correlated with non-CpG DMRs.

Nishino et al. (2011) found 20 DMRs that were strongly associated with aberrant methylation during the reprogramming process. These DMRs were consistent in 15 of the 20 studied samples and some of them overlapped with those described in the work of Lister et al. (2011). In this work they also studied the methylation profiles at different time points between passages 4 and 40, and showed that, interestingly, the number of DMRs between hESCs and hiPSCs decreases over time, and the hiPSCs become increasingly similar to hESCs. The most drastic decrease in DMRs was on the X chromosome of female hiPSCs; the DMRs nearly completely disappeared towards passage 40. They also found that new DMRs rapidly appear and gradually disappear during culture, but the number of newly appearing DMRs decreases with passaging. After passage 40, hiPSCs still maintain ∼100 DMRs on their autosomes, implying that they do not become identical to hESCs, albeit very similar. The authors propose a model in which during reprogramming, the cells undergo a critical step of hypermethylation, followed by waves of aberrant methylation that decrease with time in culture, while the hiPSCs converge to the hESC state.

Imprinted genes also appear vulnerable to disrupted DNA methylation. In hESCs, several imprinted genes have been found to show variable allelic expression (Rugg-Gunn et al., 2005; Kim et al., 2007, Rugg-Gunn et al., 2007). In the work of Rugg-Gunn et al. (2005), the authors found that H19 changed from mono to biallelic expression after long-term culture in one out of the three hESC lines of the study. The number of cells expressing the biallelic form increased with time in culture, but the mechanism did not seem to be methylation related. SLC22A18 and GNAS had a predominantly monoallelic expression, although some low and stable expression of the second allele was found in the culture, with no changes occurring during >100 passages. In a later and larger study, Rugg-Gunn et al. (2007) found that nearly all hESC lines possess a relatively stable expression pattern of imprinted genes, and that some genes seemed to be more stable than others. SNRPN, IPW and KCNQ1OT1 were highly stable, whilst H19, IGF2 and MEG3 appeared more variable. Another interesting finding of this study is the differences that were found amongst two sublines of the hESC line TE03 that were cultured in different laboratories. Whilst one subline expressed IGF2 biallelically, the other had relatively normal IGF2 expression. Array analysis showed also contrasting XIST expression between these sublines.

On the other hand, hiPSCs appear to have an overall normal monoallelic expression of imprinted genes (Pick et al., 2009). On a methylation level, Nishino et al. reported that, out of the 87 imprinted loci they studied in hiPSCs, only MEG3 and H19 showed aberrant methylation. MEG3 was abnormally methylated in 6/15 iPSC lines, and H19 was aberrantly hypermethylated in all hiPSC and hESC lines. The significance of these abnormalities for the future use of these cells is still unclear. Although the hypermethylation of MEG3 lead to a lack of expression, MEG3-negative iPSCs were not significantly different from the other lines (Nishino et al., 2011). On the other hand, work on mouse cells have shown that although iPSCs without Meg3 expression (due to aberrant hypermethylation of Dlk1-Dio3) are still able to differentiate into cell types of the three germ layers, they fail to support entirely iPSC-derived animals (Stadtfeld et al., 2010).

Conversely, the overall methylation changes found during time in culture are small compared with the differences observed between different hESC lines (Bibikova et al., 2006; Allegrucci et al., 2007; Amps et al., 2011), and in vitro culture may be responsible for only a small percentage of the changes (Calvanese et al., 2008). Furthermore, the causes of the variation in methylation patterns between hESC lines remain elusive. There is no relationship with other genomic alterations or a laboratory effect (Allegrucci et al., 2007; Amps et al., 2011), although the change in hESCs to a serum-free culture system seems to increase the epigenetic instability (Allegrucci et al., 2007). On the other hand, in hiPSCs, the fingerprint-like methylation patterns seem to originate from the significant reprogramming variability these cells display (Lister et al., 2011; Nishino et al., 2011).

Culture conditions, reprogramming and (epi)genetic instability

The role of the culture conditions on the (epi)genetic stability of hESCs and hiPSCs has received much attention. It has been hypothesized that the mechanical passaging of hESCs would reduce the appearance of aneuploid clones, which at first seemed more common upon enzymatic or chemical passaging methods (Buzzard et al., 2004; Draper et al., 2004; Mitalipova et al., 2005; Imreh et al., 2006). In the work of Mitalipova et al. (2005), hESC lines, BG01 and BG02, maintained a normal karyotype for over 40 passages when mechanically passaged, and developed aneuploidies after switching to either non-enzymatic cell dissociation buffer or enzymatic (collagenase/trypsin) passaging. The authors also show that there were important gene expression differences between chemically or enzymatically passaged hESCs, when compared with mechanically passaged lines. Conversely, Thomson et al. (2008) studied hESC lines passaged by enzymatic and chemical methods, and found that both methods equally induced genomic instability. In addition, Catalina et al. (2008) reported that regardless of the culture conditions and passaging methods, some hESC lines seemed more susceptible to karyotypic instability than others (Catalina et al., 2008). This finding is supported by the fact that numerous other research groups have reported genetic changes in the hESC lines cultured in their laboratory with no apparent correlation with how the lines were maintained or passaged (Table I). Recently, Taapken et al. (2011) found that in hiPSCs, the karyotypic changes and their frequency did not correlate with the reprogramming methods or the culture substrates, but to the time in culture (Taapken et al., 2011).

Another focus point has been the oxygen tension during culture. The environment of the mammalian reproductive tract is hypoxic. Nevertheless, hESCs, which are derived from preimplantation embryos, are typically cultured under 21% (room) oxygen tension. Several studies have investigated the impact of hypoxic culture conditions on the levels of spontaneous differentiation and maintenance of the pluripotent state in hESCs. In most cases, the authors conclude that hESCs kept in lower oxygen tensions are less prone to differentiation (Ezashi et al., 2005; Forsyth et al., 2006; Westfall et al., 2008; Chen et al., 2009; Lim et al., 2011). In two of these reports, the incidence of chromosomal aberrations was also evaluated. Forsyth et al. (2006) found that culturing hESCs at room oxygen tension resulted in a significant increase in the frequency of spontaneous chromosome breaks, gaps and exchanges during metaphase. On the other hand, their results also showed that none of the aberrations observed during metaphase would lead to gross structural abnormalities. Conversely, Lim et al. (2011) found a higher aneuploidy rate in hESCs cultured under 21% oxygen tension when compared with mild hypoxia (12% oxygen) (Lim et al., 2011). These authors also found that the cells have a lower proliferation rate in hypoxic conditions, which is in line with previously published data (Chen et al., 2009), and hypothesize that the slow mitotic division that occurs under mild hypoxia may allow hESCs to accurately segregate chromosomes, thereby decreasing the incidence of aneuploidy.

Finally, it has been suggested that hiPSCs acquire genetic changes during the reprogramming process. Although in the first works on the genomic integrity of hiPSCs, high-resolution microarray profiling failed to find any specific genomic alteration that could be induced by the reprogramming or that could be correlated with the ability of cells to be reprogrammed (Lowry et al., 2008; Chin et al., 2009), more recent work indicates the contrary (Martins-Taylor et al., 2010; Mayshar et al., 2010; Gore et al., 2011; Hussein et al., 2011). hiPSCs have now been shown to carry chromosomal abnormalities at very early passages, suggesting that these abnormalities were acquired during reprogramming or early culture due to selective pressure during the reprogramming process (Martins-Taylor et al., 2010; Mayshar et al., 2010). Furthermore, hiPSCs carry two times more copy number variations (CNVs) than the cells prior to reprogramming and when compared with hESC lines. The majority of those CNVs are unique to hiPSCs, also suggesting that they emerged during reprogramming (Martins-Taylor et al., 2010; Hussein et al., 2011). Interestingly, the number of CNVs decreased during passaging, probably due to selection favouring less damaged cells (Hussein et al., 2011). The apparently discordant results of these studies (Lowry et al., 2008; Chin et al., 2009; Martins-Taylor et al., 2010; Mayshar et al., 2010; Hussein et al., 2011) may be harmonized by the finding of Gore et al. (2011). In their work, they report protein-coding point mutations in hiPSCs. An in-depth analysis of the progenitor fibroblasts revealed that approximately half of these mutations existed in the source cells at very low frequencies and the other half would have originated during or shortly after reprogramming process. These findings show that the progenitor cells may carry mutations and possibly CNVs as a very low-grade mosaicism, and that the bottleneck of the reprogramming process drastically increases their frequency in the obtained hiPSCs.

Functional consequences of genetic changes

Differentiation capacity

Genomic changes in pluripotent stem cells may not only result in a selective advantage of the cells during in vitro culture, but may also influence their functional characteristics upon differentiation. Generally, a decreased differentiation potential is observed that can be attributed to an increased capacity to self-renew.

Culture-adapted hESCs display an increased cloning efficiency (Herszfeld et al., 2006) and a higher mitotic index, which translates into decreased doubling times in culture (Herszfeld et al., 2006; Werbowetski-Ogilvie et al., 2009). Some of the culture-adapted hESC lines have been described as clearly morphologically different from non-adapted cell lines, for instance a lack of well-defined colony edges and loss of the fibroblast-like cells surrounding the colony (Werbowetski-Ogilvie et al., 2009). Moreover, in culture-adapted cultures, the pluripotency-associated markers, SSEA3 and POU5F1, were overexpressed when compared with karyotypically normal hESCs and the expression of these markers was not exclusively detected within the hESC colonies, as in normal hESCs, but also in small clusters and even individual cells surrounding the colonies (Werbowetski-Ogilvie et al., 2009). Gradual withdrawal of basic fibroblast growth factor (bFGF, FGF-2) from the culture medium induces spontaneous differentiation with loss of SSEA3 expression in karyotypically normal hESCs but not in hESCs bearing chromosomal alterations, pointing to a reduced dependence on bFGF to remain undifferentiated (Herszfeld et al., 2006; Werbowetski-Ogilvie et al., 2009).

The comparison of the expression of embryonic lineage markers during embryoid body differentiation of karyotypically normal and abnormal hESCs reveals that, although there is evidence of extensive differentiation in all embryoid bodies, culture-adapted hESCs show altered gene-expression patterns. The loss of alpha-fetoprotein induction in the culture-adapted cells is especially marked, suggesting a reduced capacity to produce extra-embryonic endoderm (Fazeli et al., 2011). Similar results were obtained by Werbowetski-Ogilvie et al. (2009) who reported a reduced haematopoietic potential and more immature neuronal differentiation patterns in hESCs with a gain of 20q11.1q11.2.

In an in vivo differentiation setting, karyotypically normal hiPSCs usually produce mature teratomas, while karyotypically abnormal hiPSCs tend to lead to relatively immature teratomas. These teratomas contain a much higher proportion of primitive, poorly differentiated or undifferentiated cells, and they are formed by predominantly immature mesenchymal and primitive neural tissues (Herszfeld et al., 2006; Yang et al., 2008; Werbowetski-Ogilvie et al., 2009). Furthermore, teratoma-initiating cell frequencies are >400 times higher in culture-adapted hESCs than in wild-type variants. Contrary to culture adapted hESCs, cells with a normal karyotype lose the ability to form a teratoma when injecting <10–25 000 cells (Werbowetski-Ogilvie et al., 2009). Finally, work on mouse ESCs has shown that aneuploid cell lines have a lowered contribution to all tissues (somatic and germline) in adult chimaeras (Liu et al., 1997; Longo et al., 1997). Longo et al. (1997) showed that lines with >50% of chromosomally abnormal cells failed to transmit to the germline, and this failure was related to the aneuploid status rather than a loss of totipotent capacity. Similar results were obtained by Liu et al. (1997) who demonstrated that mESCs carrying a trisomy of chromosome 8 had a selective advantage during in vitro culture while they rarely contributed to the germ line upon chimaera formation.

Malignancy

Many of the genomic abnormalities reported after long-term culture of hiPSCs have also been associated with cancer cells, particularly malignant embryonic carcinoma cells (ECCs). Although ECCs share a common protein expression pattern with hESCs, mainly associated with pluripotency and development, a significant difference in the expression is observed in an important subset of proteins, with down-regulation of early developmental markers (for instance DNMT3B) and overexpression of markers that have been attributed to several malignancies, such as mitogen-activated protein kinase kinase-1 (MAP3K1) (Dormeyer et al., 2008; Chaerkady et al., 2011). The commonality of genetic alterations suggests a common mechanism of selective advantage. However, whether these genomic alterations drive karyotypically abnormal hPSCs towards a malignant phenotype is still a point of discussion.

In the undifferentiated state, karyotypically abnormal hESCs show an up-regulation of a number of oncogenes, concurrent with a downregulation of genes related to differentiation, which can be interpreted as a first step towards malignant transformation (Yang et al., 2008). Similarly, Werbowetski-Ogilvie et al. (2009) found that the expression microarray signature for 440 cancer-associated genes of an hESC line with an amplification of 20q11.1q11.2 is more similar to that of an embryonic carcinoma cell line than of an hESC line with normal genetic content. Furthermore, they showed that, similarly to malignant cancer cells, undifferentiated cells from genetically abnormal lines migrate faster in three-dimensional collagen gels than the wild-type cell lines. Further evidence for the similarities between chromosomally abnormal hESCs and cancer cells was provided by the fact that statins (proliferation inhibitors of various human cancer cells) are able to inhibit proliferation and induce apoptosis in an hESC subline carrying duplications of chromosomes 12 and 17, comparable to the effects on a human breast adenocarcinoma line, while the same cell line with a normal karyotype is not affected by this drug, even after long-term exposure (Gauthaman et al., 2009).

Of greater concern is that certain properties of the culture-adapted hESCs can spread within a culture dish from abnormal to normal cells through cell-to-cell contact. Co-culture of transformed-hESCs with normal hESCs leads to an enhanced self-renewal, higher expression of pluripotent markers and loss in normal terminal differentiation programmes in the normal hESCs (Werbowetski-Ogilvie et al., 2011).

The inheritability of certain ‘undesirable’ traits has also been shown in in vitro differentiation experiments of chromosomally abnormal hESCs. Neural precursor-like cells derived from altered hESCs show a higher percentage of cells in the S-phase when compared with normal hESC-derived progenitors. This fraction of S-phase cells is comparable to that of the undifferentiated parental line. Upon injection of these abnormal hESC-derived neural precursors into immunodeficient mice, well-defined tumours are formed that contain a high fraction of Ki67-positive cells, demonstrating that the cells continue to proliferate in vivo (Werbowetski-Ogilvie et al., 2009).

Gopalakrishna-pillai and Iverson (2010) differentiated diploid and trisomic hESCs into astrocyte progenitor cells and found that the gene expression profile of differentiated trisomic astrocyte progenitors displays significant similarities to those of a malignant astrocytoma cell line and glioblastoma samples. Many of the up-regulated transcripts in trisomic hESC-derived astrocyte progenitors and astrocytoma cells encode proteins associated specifically with astrocytomas or implicated in cancer in general. The down-regulated transcripts included several markers of normal differentiated astrocytes (Gopalakrishna-Pillai and Iverson, 2010).

Finally, as previously mentioned, hESC lines with genomic alterations tend to form fairly immature teratomas (Herszfeld et al., 2006; Yang et al., 2008). Often, nested regions of POU5F1-positive cells are observed, classifying them as grade III teratomas (Yang et al., 2008; Werbowetski-Ogilvie et al., 2009). However, these altered hESC-derived teratomas do not form malignant teratocarcinomas as known for ECCs (Herszfeld et al., 2006), nor do they cause retention of metastatic cells in any tissue of the recipient animals (Werbowetski-Ogilvie et al., 2009).

Conclusions

Human pluripotent stem cells frequently acquire (epi)genetic aberrations during in vitro culture. Strikingly, many of the observed alterations are recurrently observed. This marked (epi)genetic instability is possibly caused by suboptimal culture conditions, although this needs to be proved. The culture adaptations often lead to increased survival and proliferation of undifferentiated stem cells resulting in a selective advantage in culture that leads to overgrowth of abnormal, culture-adapted cells as well as an altered differentiation capacity of the affected sublines. Moreover, they are highly similar to adaptations observed in malignancies, and premalignant transformation has been reported in several culture-adapted pluripotent stem cell lines. The functional effects of these alterations are not yet fully understood, but suggest a (pre)malignant transformation of affected cells with decreased differentiation and increased proliferative capacity.

In conclusion, the high degree of genetic and epigenetic alterations reported in the literature and the altered phenotypic characteristics of the affected cells urge for a stringent quality control system, and make the screening of the (epi)genetic integrity of human pluripotent stem cells before any application an absolute must.

Authors' roles

H.N., M.G. and C.S. searched the public databases, read the papers and co-wrote the manuscript. All authors equally contributed to this work.

Funding

This work was supported by the Methusalem grant of the Research Council of the VUB. C.S. is a postdoctoral fellow at the Fonds voor Wetenschappelijk Onderzoek Vlaanderen.

Conflict of interest

The authors declare no conflict of interest.

Acknowledgements

The authors wish to acknowledge Prof. Karen Sermon for the critical reading of the manuscript.

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