Download UVA radiation causes DNA strand breaks, chromosomal

Oncogene (2008) 27, 4269–4280
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ORIGINAL ARTICLE
UVA radiation causes DNA strand breaks, chromosomal aberrations
and tumorigenic transformation in HaCaT skin keratinocytes
K Wischermann1, S Popp1, S Moshir1, K Scharfetter-Kochanek2, M Wlaschek2, F de Gruijl3,
W Hartschuh4, R Greinert5, B Volkmer5, A Faust5, A Rapp6, P Schmezer7 and P Boukamp1
1
Division of Genetics of Skin Carcinogenesis, German Cancer Research Center (DKFZ), Heidelberg, Germany; 2Department of
Dermatology and Allergic Diseases, University of Ulm, Ulm, Germany; 3Department of Dermatology, Leiden University Medical
Center, Leiden, The Netherlands; 4Department of Dermatology, University of Heidelberg, Heidelberg, Germany; 5Division of
Molecular Cell Biology, Dermatology Center, Buxtehude, Germany; 6Sir William Dunn School of Pathology, Oxford, UK
and 7Division of Toxicology and Cancer Risk Factors, German Cancer Research Center (DKFZ), Heidelberg, Germany
The role of UVA-radiation—the major fraction in
sunlight—in human skin carcinogenesis is still elusive.
We here report that different UVA exposure regime
(4 5 J/cm2 per week or 1 20 J/cm2 per week) caused
tumorigenic conversion (tumors in nude mice) of the
HaCaT skin keratinocytes. While tumorigenicity was not
associated with general telomere shortening, we found new
chromosomal changes characteristic for each recultivated
tumor. Since this suggested a nontelomere-dependent
relationship between UVA irradiation and chromosomal
aberrations, we investigated for alternate mechanisms of
UVA-dependent genomic instability. Using the alkaline
and neutral comet assay as well as c-H2AX foci formation
on irradiated HaCaT cells (20–60 J/cm2), we show a dosedependent and long lasting induction of DNA single and
double (ds) strand breaks. Extending this to normal
human skin keratinocytes, we demonstrate a comparable
damage response and, additionally, a significant induction
and maintenance of micronuclei (MN) with more acentric
fragments (indicative of ds breaks) than entire chromosomes particularly 5 days post irradiation. Thus, physiologically relevant UVA doses cause long-lasting DNA
strand breaks, a prerequisite for chromosomal aberration
that most likely contribute to tumorigenic conversion of
the HaCaT cells. Since normal keratinocytes responded
similarly, UVA may likewise contribute to the complex
karyotype characteristic for human skin carcinomas.
Oncogene (2008) 27, 4269–4280; doi:10.1038/onc.2008.70;
published online 31 March 2008
Keywords: DNA double strand breaks; tumor formation;
chromosomal aberrations; telomere length; radiation
regimes
Correspondence: Dr P Boukamp, Division of Genetics of Skin
Carcinogenesis, German Cancer Research Center (DKFZ), Im
Neuenheimer Feld 280, Postfach 101949, Heidelberg D-69120,
Germany.
E-mail: [email protected]
Received 14 March 2007; revised 19 December 2007; accepted 15
February 2008; published online 31 March 2008
Introduction
The incidence of skin cancer has rapidly increased
during the last decades, making skin cancer—comprised
of basal cell carcinoma (BCC), squamous cell carcinoma
(SCC) and malignant melanoma (MM)—the most
frequent cancer worldwide (reviewed in Boukamp,
2005b). It is now well accepted that UV radiation is
the major risk factor, and that cumulative UV damage is
responsible for the development and progression of
SCCs, while BCCs and especially MM depend more on
intermittent UV overexposure (Armstrong and Kricker,
2001). From the three UV components—UVA (wavelengths 320–400 nm), UVB (280–320 nm) and UVC
(200–280 nm)—only UVB and UVA radiation reach
the earth while UVC radiation is fully absorbed by the
ozone layer. Similarly, most of the UVB radiation is
absorbed. Consequently, the UV radiant energy reaching the earth consists of only B5% UVB and B95%
of UVA.
UVB radiation is thought to be the major risk factor
for human skin carcinogenesis because its energy is
sufficient to enter the nuclei of epidermal cells and to
directly damage DNA by causing cyclobutane pyrimidine dimers (CPDs) and pyrimidine(6-4)pyrimidine
photoproducts at dipyrimidine sites, thereby giving
rise to very specific mutation patterns. Such C to T
transitions are particularly frequent in the p53 tumor
suppressor gene that is mutated in 50–90% of the
human SCCs (summarized in Boukamp, 2005b).
Recently, UVA radiation also gained more interest.
Although the photon energy may be insufficient to
damage DNA directly, UVA radiation is able to
penetrate into the epidermis down to the proliferative
basal cells and even further to the dermal compartment
where it contributes to photoaging. In addition,
evidence is increasing that UVA may also be relevant
for skin cancer development. Albino hairless mice
exposed to UVA irradiation (330 nm) developed
tumors after >400 days and even long-wave UVA
irradiation (>340 nm) has been found to induce skin
carcinomas (for review see de Gruijl, 2002). Furthermore, the immortal human HaCaT keratinocytes
UVA causes tumorigenic conversion of HaCaT keratinocytes
K Wischermann et al
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became tumorigenic when irradiated with 24 J/cm2 UVA
for 18 weeks (He et al., 2006).
The action of UVA is thought to be indirect by
activating cellular chromophores and inducing photooxidative cellular stress (Wondrak et al., 2006). The
thereby generated reactive oxygen species (ROS) can
modify guanine to 8-oxo-7,8-dihydro-70 -deoxyguanosine (8-oxo-guanine) resulting in transversion mutations
by changing guanine to thymine during replication
(Cadet et al., 2005)—now considered as UVA signature
mutations. ROS are also able to damage DNA most
preferentially by inducing DNA-protein cross-linking,
alkali labile sites as well as CPDs. This said, Ono
and co-workers showed that longer wavelength UVA
(310–340 nm) contributes significantly to induction of
mutations and appearance of hotspots at cytosine
phosphate guanine-associated dipyrimidine sites (Ikehata and Ono, 2007). Furthermore, mutations in UVAexposed epidermis (including wavelength bordering
UVB) were dominated by C to T transitions thus
making broadband UVA a considerable contributor to
UV-dependent mutations (Ikehata et al., 2003, 2004). In
addition, ROS are involved in generating DNA single
strand (ss) breaks (Kielbassa et al., 1997). If ss breaks
are not repaired during the G1 phase of the cell cycle
they can generate double strand (ds) breaks during
S-phase which then can result in chromosomal aberrations such as amplifications, deletions or translocations
(for review see Limoli et al., 2002; Thompson and
Limoli, 2003); all hallmarks of skin carcinomas. Interestingly, CPDs can also cause ds breaks in S-phase
(Garinis et al., 2005), suggesting that they too may
contribute to chromosomal instability.
To determine the functional consequence of UVA
radiation on human skin keratinocytes, we utilized the
immortal human HaCaT cells (Boukamp et al., 1988).
These cells represent an early stage of skin carcinogenesis because they harbor UVB type-specific p53
mutations (Lehman et al., 1993) as well as some
chromosomal abnormalities typical for human skin
SCCs (Lehman et al., 1993; Boukamp et al., 1997; Popp
et al., 2002). While the HaCaT cells remained nontumorigenic through long-term passaging, they became
tumorigenic by transduction with the Harvey-ras
oncogene, increased temperature or stroma modulating
growth factors known to be relevant for tumorigenicity
(Boukamp et al., 1990b, 1995, 1997, 1999; Skobe and
Fusenig, 1998; Obermueller et al., 2004). Thus, the
intrinsic stable nontumorigenic phenotype, together
with the sensitivity for tumorigenic conversion upon
distinct changes, make HaCaT cells a highly suitable
model to study the role of UVA in skin carcinogenesis.
Exposing HaCaT cells to UVA radiation, we now
show that UVA is able to induce tumorigenicity upon
different treatment regimes. Since tumorigenicity was
associated with new chromosomal aberrations, we
additionally explored the immediate damage response
to UVA radiation and determined its role in inducing
DNA ss and ds breaks and the formation of micronuclei
as indicators of chromosomal aberrations. We also show
that normal human skin keratinocytes exhibit a similar
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damage response pattern as HaCaT cells, thus providing
evidence for a role of UVA-dependent chromosomal
damage in human skin carcinogenesis.
Results
UVA-induced tumorigenic conversion of the HaCaT cells
In a first exploratory experiment we irradiated HaCaT
cells with a steadily increasing dose (from 7 J/cm2 during
the first week up to 42 J/cm2 during the last week) of
UVA radiation over a period of 42 days. This treatment
caused cell death of almost all cells except one colony
that was further expanded. Different from the control
HaCaT cells, nodules developed in four of the eight
injection sites upon injection into nude mice. Of these,
two regressed while the other two continuously expanded
and were classified as invasively growing welldifferentiated SCCs (Supplementary Figures 1A and
B). Recultivation of one of these SCCs and its genetic
comparison (multiplex fluorescence in situ hybridization;
M-FISH) with the control HaCaT cells clearly demonstrated its HaCaT origin (containing the original
HaCaT marker chromosomes) and four new chromosomal aberrations (Supplementary Figure 1C and D).
Being present in all metaphases, the new aberrations
clearly document the clonal origin of the tumor.
Tumorigenic conversion is induced upon defined UVA
treatment regimes
Since tumorigenicity had occurred upon UVA treatment, we next asked whether distinct and physiological
relevant doses would be sufficient for tumorigenic
conversion. We applied a dose of 20 J/cm2 which equals
a 1 h exposure around noon in mid Europe (Federal
Office for Radiation Protection, 2006) to two parallel
cultures of HaCaT cells (series A and B for each
treatment) either as a single dose once per week—
allowing a recovery time of 6 days—or as a split dose of
5 J/cm2 four times per week thereby mimicking continuous low-dose UVA exposure. Different from the
exploratory experiment, toxicity was negligible; the
cells continued to proliferate and required passaging
once a week.
All cells were tested for tumorigenicity after 5 and 15
weeks of continuous UVA treatment. With the exception of one rapidly expanding nodule (5-week time
point) which by histology was identified as a water- and
blood-filled epidermal cyst (data not shown), the control
cells remained nontumorigenic (Figure 1a). Series A cells
irradiated with 4 5 J/cm2 per week also remained nontumorigenic (data not shown). Small nodules occurring
shortly after injection regressed within 2 months.
However, series B cells irradiated with 4 5 J/cm2
(4 5 J/cm2 B) developed tumors. After 5 weeks, tumor
growth occurred in 2/6 and after 15 weeks of continuous
UVA treatment in 5/8 injection sites (data not shown
and Figure 1b). Treatment of 20 J/cm2 once a week
caused tumor growth in both series. Series A cells
formed tumors in 3/6 injection sites already after 5
UVA causes tumorigenic conversion of HaCaT keratinocytes
K Wischermann et al
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Figure 1 Tumor growth and histology after distinct doses of UVA irradiation. (a–c) Tumor growth curves of nonirradiated HaCaT
control cells (a) and UVA-irradiated HaCaT cells from series B treated with 4 5 J/cm2 for 15 weeks (b) and series B cells treated with
1 20 J/cm2 for 15 weeks (c). (d) Tumor histology representative for the well-differentiated SCCs formed by the UVA-treated HaCaT
cells. Bar ¼ 100 nm.
weeks treatment (data not shown). Unfortunately, these
cells were then lost due to infection. Series B cells
(1 20 J/cm2 B) developed one tumor (1/6 injection
sites) after 5 weeks while five tumors formed in the eight
injection sites after 15 weeks (data not shown and
Figure 1c). As exemplified in Figure 1d, all tumors were
histologically classified as well-differentiated invasively
growing SCCs.
Tumorigenicity is associated with new chromosomal
changes
To determine whether these defined exposure regimes
may also be associated with changes in the chromosomal make up, we performed M-FISH analysis on
the cells irradiated for 5 weeks either with 4 5 or
1 20 J/cm2 per week prior to injection into the mice as
well as with cells re-established from six tumors formed
by the UVA irradiated cells. As shown in Figure 2a,
UVA-treated HaCaT cells prior to injection showed the
HaCaT-specific aberrations and aberrations that had
occurred during passaging. In addition, we found
aberrations in individual metaphases (Supplementary
Table 1). Since the number was slightly increased in the
UV-irradiated versus nonirradiated control cultures,
this suggested that some chromosomal changes may
have occurred upon UVA treatment.
Of the six tumor-derived lines four originated from
series B HaCaT cells irradiated with 4 5 J/cm2 per
week. Tumor 1 (T1) developed after the 5-week
treatment and tumors T2, T3 and T4 after the 15-week
treatment. Two cell lines derived from series B HaCaT
cells irradiated with 1 20 J/cm2 once per week for 15
weeks (tumors T5 and T6). As exemplified in Figures
2b–d, M-FISH analysis clearly identified the original
HaCaT markers and a common set of aberrations—
representing a subpopulation of the untreated HaCaT
cells. In addition and in agreement with the tumor cells
established from the exploratory experiment, all tumorderived lines showed tumor-specific karyotypic changes.
First, the number of chromosome 15 was reduced in
all metaphases (Table 1). Second, three cell lines
(T4, T5 and T6) exhibited 3–5 additional recurrent
aberrations, that is, the same aberration present in all
metaphases, that were unique for the respective
tumor-derived lines (Figures 2b–d and Table 1) demonstrating that in these tumors one subclone with a
specific aberration profile dominated during tumor
growth. The other three cell lines (T1, T2 and T3)
were characterized by different subpopulations, each
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Figure 2 Genetic characterization of the tumors induced by the two UVA treatment regimes. M-FISH analysis from HaCaT cells
irradiated with 4 5 J/cm2 for 5 weeks prior to s.c. injection (a) and HaCaT cells recultivated from tumors from series B 4 5 J/cm2 for
15 weeks, T4 (b), series B 1 20 J/cm2 for 15 weeks, T5 (c) and series B 1 20 J/cm2 for 15 weeks, T6 (d). All karyotypes show the
typical early ‘HaCaT markers’ (t(3q;4q), del(7p), i(9q), t(4;18)) as well as culture-derived markers (t(1;2), t(4;11), t(5;9), t(6;7), t(7;9),
t(5;8), del(10p), t(8;11), t(6;14), t(8;15), t(16;20), t(7;17), del(19), del(20), i(22q), del(Xp). In addition the recultivated tumors show extra
recurrent chromosomal aberrations (in all metaphases analysed). 4 5 J/cm2 B 15 wk T4: t(14;16), t(4;18;13), del(13) and del(17) (b),
1 20 J/cm2 B 15 wk T3: t(X;2), t(13;2;13), del(t(19;22)), del(1) and del(7) (c) and 1 20 J/cm2 B 15 wk T4: i(10q); gain t(5;8) and
del(16) (d).
Table 1
4 5 J/cm B for
5 weeks (T1)
2
Chromosomal aberrations characterizing the different cell lines established from the UVA-dependent tumors
4 5 J/cm2 B for
15 weeks (T2)
Aberrations present in all metaphases
del(15)
del(15)
4 5 J/cm2 B for
15 weeks (T3)
4 5 J/cm2 B for
15 weeks (T4)
1 20 J/cm2 B for
15 weeks (T5)
1 20 J/cm2 B for
15 weeks (T6)
del(15)
del(15) t(14;16)
t(4;18;13) del(13)
del(17)
del(15) t(X;2)
t(13;2;13)
del(t(19;22))
del(1) del(7)
del(15) i(10q)
gain t(5;8) del(16)
Aberrations present in only a subset of metaphases
del(3) or del(7)
gain 12 or X
t(X;16)
+ del(2q)
dup t(8;15)
del(13q)
characterized by distinct individual aberrations (at least
be detectable in three metaphases each, see Table 1).
Thus, besides selection for a pre-existing HaCaT
subpopulation all tumor-derived lines had gained new
chromosomal aberrations, which obviously had provided a selective growth advantage in vivo and, accordingly, were crucial for the tumorigenic phenotype of the
respective HaCaT populations.
UVA irradiation induces DNA damage in HaCaT cells
Chromosomal aberrations (translocations, deletions,
amplifications) require the formation of DNA strand
breaks which further progress to chromosomal breakage. To determine whether UVA can induce DNA
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strand breaks we irradiated HaCaT cells with 60 J/cm2
and performed the alkaline comet assay—detecting ss
and ds breaks as well as alkali-labile sites—either
directly or after a recovery period of 2 and 4 h. While
the nonirradiated mock-treated control cells exhibited
almost no DNA damage (average DNA in tail: 1.96%
±0.53), the average DNA in tail of irradiated cells
increased significantly (40.35% ±4.68; Po0.0001).
After 2 h B84% of the DNA damage was repaired
leaving 6.45% (±2.88) DNA in the tail (Figure 3a). This
decreased further to 3.94% (±2.95) after 4 h demonstrating a repair of 91%. Plotting the DNA in tail for
each individual cell additionally clarified that UVA
irradiation of 60 J/cm2 had caused significant damage in
UVA causes tumorigenic conversion of HaCaT keratinocytes
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Figure 3 DNA damage in HaCaT cells after UVA irradiation. (a) The alkaline comet assay was used to measure DNA damage
directly after irradiation with 60 J/cm2 UVA (0 h) as well as after 2 and 4 h recovery time. Damage is expressed as DNA in tail (%).
Examples of cells without irradiation (control), directly (UVA 0 h), 2 h (UVA 2 h) and 4 h (UVA 4 h) after irradiation are shown in the
lower panel. A representative experiment is shown. (*Po0.05; **Po0.001 (Mann–Whitney rank sum test)) (b) The neutral comet
assay was used to measure DNA ds breaks directly after irradiation with 20, 40 and 60 J/cm2 UVA. Damage is expressed in DNA in tail
(%). Examples of control and cells irradiated with 60 J/cm2 are shown in the lower panel. The mean of three experiments is shown
(**Po0.001 (t-test)). (c and d) Ds breaks are detected by g-H2AX immunofluorescence staining in G1-enriched cells. While control
cells harbor no or only a few foci (c), the amount of foci increases after irradiation with 60 J/cm2 UVA (d).
about every cell (Supplementary Figure 2A) and that
although most of the cells had completely repaired their
damage within 4 h, a fraction of 15–20% still showed
X20% of DNA in tail. Thus, UVA induced DNA
strand breaks and a small fraction of cells maintained
damage for longer time periods.
UVA induces double strand breaks in HaCaT
keratinocytes
Since the alkaline comet assay does not differentiate
between ss and ds breaks we performed the neutral
comet assay as well as immunofluorescence staining for
g-H2AX, both indicative for DNA ds breaks (Rogakou
et al., 1998; Olive, 1999; Sedelnikova et al., 2002;
Rothkamm and Lobrich, 2003). HaCaT keratinocytes
were irradiated with 20, 40 and 60 J/cm2 UVA and the
neutral comet assay was performed directly thereafter.
While nonirradiated control cells on average showed
B23% (±4.3) DNA in tail, cells irradiated with
20 J/cm2 UVA showed 42.93% (±3.7) and this further
increased to 48.82% (±4.8) with 40 J/cm2 and 55.67%
(±4.2) with 60 J/cm2 (Figure 3b, Supplementary Figure
2B). In all cases the increase was statistically significant
(Po0.001). Thus, UVA induced DNA ds breaks in a
dose-dependent manner.
We next performed immunofluorescence staining for
g-H2AX. Since it is suggested that also during S-phase
ds breaks occur that are highlighted by g-H2AX staining
(Thompson and Limoli, 2003), HaCaT cells were
enriched in the G1 phase of the cell cycle (Supplementary Figures 3A and B) before being irradiated with 60 J/
cm2 UVA and incubated for 30 min to allow maximal
foci induction (Rapp and Greulich, 2004). Furthermore,
it was recently shown that UVC radiation induced
g-H2AX phosphorylation mainly as a diffuse, even pannuclear staining which was discussed as being dependent
on nucleotide excision repair (Marti et al., 2006).
Different from this, we here found induction of distinct
g-H2AX foci. While nonirradiated control cells showed
on average 2.3 foci per nucleus, the number strongly
increased in UVA-irradiated cells (Figures 3c and d).
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Most importantly, the majority (B65%) showed on
average 6 foci per nucleus while such nuclei were only
detected in about 10% of the control population.
Telomere length is not altered upon UVA treatment
Another potential site for damage that can cause
genomic instability, are the telomeres. We, therefore,
performed telomere length restriction fragment (TRFL)
analysis with UVA-treated HaCaT cells prior to
injection as well as tumor-derived cell lines. As
exemplified in Figure 4a, telomere length of the
irradiated pre-injected cells—irrespective of treatment
regime and time of treatment—was very similar to that
of the nonirradiated control cells (mean TRFL of
B2.5 kb). Thus, even the 15-week treatment did not
cause obvious changes in telomere length.
Different from this and with one exception (T4), all
recultivated tumors exhibited an altered telomere length
(Figure 4b). Three lines seemed to contain two populations with different telomere length. In two, the original
mean TRFL of B2.5 kb predominated (TA and T1)
over a second peak with a mean TRFL of B5 kb while
in one cell line the population with longer telomeres
(mean TRFL of B6 kb, T2) predominated. Two lines
showed a mean TRFL of B3.5 kb (T3 and T6) and
two of B6 kb (TB and T5). Thus, instead of potential
telomere shortening we rather observed subpopulations
with about double the original telomere length. It is
noteworthy that the three genetically heterogeneous
tumor-derived lines (T1, T2 and T3) were among those
showing two different telomere lengths.
Figure 4 Telomere length determination of the different HaCaT
variants. (a) Telomere restriction fragment length (TRFL) measurement of the HaCaT cells treated with UVA radiation either
with 4 5 J/cm2 per week for 5 and 15 weeks or with 1 20 J/cm2
per week for 15 weeks. (b) TRFL from recultivated UVAdependent tumor cells either show a similar telomere length as
the cells prior to injection (T4), increased mean TRFL (T3, TB, T5,
T6) or exhibit two populations with different mean TRFLs (TA,
T1, T2). The respective mean TRFLs are marked in the different
samples. Size of the marker lambda HindIII digested is indicated in
the left margin.
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UVA irradiation induces DNA damage also in normal
human keratinocytes
Since the immortal HaCaT keratinocytes harbor p53
mutations (Lehman et al., 1993) which could influence
the UVA-dependent damage response, we next investigated normal human skin keratinocytes. G1-enriched
keratinocytes (Supplementary Figures 3C and D) were
irradiated with 60 J/cm2 UVA and analysed for g-H2AX
foci formation directly after irradiation (0 h) as well as
after 10, 20, 30, 60 and 360 min (Figure 5a). Nonirradiated control cells exhibited on average 1–2 foci per
nucleus. While directly after UVA irradiation there was
no change, damage increased steadily during the first
30 min post irradiation with an average of 5 foci
per nucleus and even after 6 h another slight increase
was seen.
Since in HaCaT cells damage and repair was not
evenly distributed, we also evaluated each individual
nucleus (Figure 5b). The number of nuclei with 1–5 foci
decreased from 68% after 1 h to 48% after 6 h and,
accordingly, the number of foci-negative nuclei increased from 20 to 28%. Interestingly, the number of
nuclei with 6–10 foci also increased from 10 to B20%.
Thus, while repair took place in some cells, damage
persisted in others and here the amount of ds breaks was
even increased after 6 h. Although we cannot firmly rule
out that an excess of g-H2AX foci was due to replication
processes or repair, it has to be stressed that 83% of the
cells were in the G1-phase at the time of irradiation
(Supplementary Figures 3C and D).
UVA induces ds breaks already at ‘low’ doses in normal
keratinocytes
To determine whether lower doses of UVA radiation
would also induce a distinct damage profile, we
irradiated normal human skin keratinocytes with 10,
20, 30, 40 and 60 J/cm2 UVA and analysed for g-H2AX
foci after 1 h—identified as the time for maximal foci
induction in these cells (Figure 5c). Keratinocytes
irradiated with 10 J/cm2 UVA showed 2.26 foci per
nucleus, values very similar to that of nonirradiated
controls (1–2 foci per nucleus). In contrast, 20 J/cm2
UVA irradiation induced a major increase in foci
formation with an average of 5–6 foci/nucleus. Interestingly, there was no further increase with higher doses
including 60 J/cm2. Although this suggested that 20 J/
cm2 was as effective as 60 J/cm2, we observed some
difference when classifying the individual damage
response. Eighteen percent of the human keratinocytes
were negative (0 foci), 67% showed 1–5 and 15% 6–10
foci after 20 J/cm2 UVA treatment (Figure 5d). After
irradiation with 60 J/cm2, 20% of cells were also without
foci, while the number of highly damaged keratinocytes
was slightly increased with B60% of the cells with 1–5
and >20% with 6–10 foci.
Finally, we performed the alkaline and neutral comet
assays. Irradiation of normal human keratinocytes with
30 and 60 J/cm2 UVA in the alkaline comet assay
resulted in a dose-dependent increase in DNA damage
from 9.62% (±2.6%) DNA in tail in nonirradiated
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control cells to 23.83% (±3.7%; Po0.001) after 30 J/
cm2 and 42.54% (±5.5%; Po0.001) after 60 J/cm2
UVA (Figure 5e; Supplementary Figure 2C). Also the
neutral comet assay provided evidence for UVAdependent formation of ds breaks, though the induction
was only minor (Supplementary Figure 2D).
UVA irradiation causes micronuclei formation in normal
keratinocytes
As a consequence of ds breaks and chromosomal
breakage, micronuclei (MN) are formed. Therefore,
MN formation represents an established marker for
genomic instability (Fenech, 2002). Since it is suggested
that the cytokinesis-block MN assay allows a better
precision because data obtained are not confound by
altered cell division (Fenech, 2000), we treated the
keratinocytes with cytochalasin-B for 24 h after irradiation
with 20 J/cm2. To distinguish between MN originating
from whole chromosomes or acentric fragments,
we additionally performed centromere staining and
determined the fraction of MN-positive cells as well as
the ratio between MNs with and without centromeres in
B1000 cells.
Investigating keratinocytes from three different
donors, the number of MN-positive cells nearly doubled
from 1–1.8% in nonirradiated control cells to 2–3.2%
after UVA irradiation and this increase was significant
for two keratinocytes strains (w2-test; Figure 6a). Concerning the kind of MNs, we found some inter-personal
variation. While one strain showed only induction of
centromere-containing MNs, the other two also showed
MNs without centromeres (32 and 65%, respectively;
Figures 6a and b). This indicates, that UVA is able to
induce both, a nondisjunction and thereby loss of entire
chromosomes, as well as DNA ds breaks thereby
causing chromosome breakage that result in acentric
fragments. Whether there is a preference for one or the
other may well be donor specific.
To determine whether MN induction by UVA was
only a short-term effect or may be longer lasting, we
next irradiated keratinocytes with 10, 20 and 40 J/cm2
and cultured them for 5, and in part 10, consecutive days
without cytochalsin-B blockade. While only B1% of
nonirradiated control cells showed MN, we observed a
significant dose-dependent increase of MN-positive cells
5 days after irradiation with 1.8% after 10 J/cm2, 2.6%
after 20 J/cm2 and 4.4% after 40 J/cm2 UVA (Figure 6c,
Cochran–Armitage test, Po0.00001). While both, the
fraction of MN with and without centromeres increased,
MNs without centromeres dominated with increasing
UVA doses (20 and 40 J/cm2). Interestingly, also
keratinocytes cultured for 10 days post irradiation
(20 J/cm2) still showed twice the amount of MN-positive
cells (B6%) compared to their nonirradiated controls
Figure 5 DNA damage in normal keratinocytes after UVA irradiation. (a) G1-enriched normal human keratinocytes were irradiated with
60 J/cm2 UVA and the number of g-H2AX foci/nucleus determined 0,
10, 20, 30, 60 and 360 min post irradiation. (b) Classification of nuclei
carrying 0, 1–5, 6–10 and >10 g-H2AX foci 60 and 360 min post
irradiation with 60 J/cm2. (c) g-H2AX foci formation was determined
1 h post irradiation in G1-enriched cells irradiated with 10, 20, 30, 40
and 60 J/cm2 UVA and the average number of foci/nucleus was
calculated. (d) Keratinocytes were irradiated with 20 and 60 J/cm2
UVA and the number of g-H2AX foci/nucleus classified into three
groups: no (0), 1–5 and 6–10 foci. (e) DNA damage was measured
directly after irradiation by the alkaline comet assay following 30 and
60 J/cm2 UVA irradiation and being expressed as DNA in tail (%);
(**Po0.001 (Mann–Whitney rank sum test)). All data represent a
mean of at least two independent experiments.
Oncogene
UVA causes tumorigenic conversion of HaCaT keratinocytes
K Wischermann et al
4276
Figure 6 Micronuclei induction in normal keratinocytes after UVA irradiation. (a) Micronuclei (MN) formation in keratinocytes
from three different donors were analysed with the cytokinesis-blocked MN-assay 24 h after irradiation with 20 J/cm2 UVA. An
additional all-centromere staining shows MNs without centromeric signals (dark part of the bar), indicative for chromosomal
breakage, as well as MNs with centromeric signals (light part of the bar) indicative for loss of whole chromosomes. (b) Representative
cells with MN without centromeric signals (two left panels) and MN with centromeric signals (two right panels). (c) MN formation in
keratinocytes irradiated with 10, 20 and 40 J/cm2 5 d post irradiation. MNs are induced in a significant dose-dependent manner. The
majority of micronuclei lack centromere staining. In (a) and (c) the results of representative experiments are shown (*Po0.05;
**Po0.001 (w2-test)).
(2–3%; data not shown), demonstrating long-lasting
damage.
Discussion
In the present paper, we report that defined treatment
regimes (1 20 or 4 5 J/cm2 per week) of environmentally highly relevant UVA doses cause tumorigenic
conversion of HaCaT skin keratinocytes allowing them
to form well-differentiated SCCs in nude mice. Tumorigenicity was characterized by distinct chromosomal
Oncogene
aberrations present in the cells of the recultivated
tumors. In an approach to assess DNA damage
formation by UVA and its mutagenic outcome we show
that UVA radiation caused DNA ss and ds breaks in
HaCaT cells and that some of this damage was
maintained even several hours post irradiation. Furthermore, we demonstrate that this damage profile is not
restricted to the immortal HaCaT cells but is similarly
seen in normal human keratinocytes. We even provide
evidence for a long-lasting increase in micronuclei
formation with about half of the MNs containing either
entire chromosomes or acentric fragments, that is,
UVA causes tumorigenic conversion of HaCaT keratinocytes
K Wischermann et al
4277
chromosomal parts that require chromosomal breakage
in order to occur. All this strongly implies that UVA
radiation—most likely through oxidative damage
(Kielbassa et al., 1997; Cadet et al., 2005)—induces
DNA ss and ds breaks. Whether the ds breaks arise
directly as or following ss breaks processing, is currently
unclear and remains to be addressed. However, the
study shows that UVA is able to induce genomic
instability by causing DNA strand breaks which
presumably induce chromosomal breakage and, upon
repair, chromosomal changes as observed in the
tumorigenic HaCaT cells. Furthermore, since similar
long-lasting damage is seen in normal human keratinocytes and since skin SCCs are characterized by a high
frequency of chromosomal changes (reviewed in Boukamp, 2005b), it is tempting to suggest that UVA
radiation may contribute to skin cancer development
and progression in a similar way.
This assumption is also substantiated by a recent
study showing that irradiation of HaCaT cells with as
little as 10 J/cm2 UVA reduced long-term plating and
colony forming efficiency (Phillipson et al., 2002). Since
this was not related to an extended population doubling
time but to partial cell death, the authors reasoned that
UVA induced delayed cell death. Also we found an
increased death rate 14 h (40%) post irradiation while
directly after irradiation (up to 4 h) and later on (after 5
days) cell death was negligible. Although we cannot
exclude that during this phase all ‘long-lasting’ damaged
cells died, we found—again in line with the previous
report—that this delayed cell death correlated with
increased MN formation. The authors interpreted this
as an indication for persistent genomic instability and
hypothesized that UVA leads to an increasing frequency
of chromosomal rearrangements (Phillipson et al., 2002)
something that we now show for the recultivated
tumorigenic HaCaT cells. Accordingly, in fibroblasts
low doses of UVA induced MNs at a level comparable
to 1 Gy ionizing radiation, supporting the view that
UVA is a potential inductor of chromosomal aberrations (Emri et al., 2000). Thus, with our study we
establish a first association between UVA-dependent
damage, chromosomal aberrations and their functional
consequences for carcinogenesis.
Having performed Comet-FISH analysis on human
lymphocytes, we previously suggested that 50 J/cm2
UVA induced a number of chromatid strand breaks in
chromosomes in a nonrandom manner (Rapp et al.,
2000). While lacking correlation between strand breaks
and chromosome size, an inverse correlation was
established between the density of active genes and
sensitivity toward UVA radiation. In the HaCaT cells
studied here, we found many different chromosomes
being involved in the aberrations prior to injection.
Furthermore, since all changes were only seen in a single
metaphase obviously none of them provided a growth
advantage in vitro. In vivo, on the other hand, and most
likely due to a massive selection pressure, only few cells
survived and gave rise to tumors of genetically defined
subpopulations. These were either of monoclonal
origin (all carrying the same aberrations) or derived
from several clones (different aberrations in different
subpopulations). However, in these also the aberrationinvolved chromosomes differed. Thus, we have to
conclude that damage and formation of chromosomal
aberrations is a rather random process and that different
aberrations are able to contribute to tumorigenicity.
Different from, for example, chronic myeloid leukemia where the formation of specific translocation
chromosomes is accepted as being crucial for tumor
development (Tefferi et al., 2005), the role of most
chromosomal aberrations in solid tumors is still elusive.
This is also the case for the aberrations identified for the
different tumor-derived cell lines. One exception may be
loss of chromosome 15, the only common aberration.
We previously showed that loss of chromosome 15
correlated with loss of the angiogenesis inhibitor TSP-1
which in turn correlated with the angiogenic switch
required for malignant conversion (Folkman and
Hanahan, 1991; Bleuel et al., 1999). We recently also
showed that loss of chromosome 15 occurs in human
skin carcinomas and that this correlated with progression from benign to malignant tumor phenotype
(Burnworth et al., 2006). Accordingly, loss of chromosome 15 seen here, may also have provided a selective
advantage and thereby contributed to the high
frequency of tumorigenic conversion (three out of the
four different HaCaT populations irradiated with 4 5
or 1 20 J/cm2).
Convincing evidence for the role of UVA in tumorigenic conversion of HaCaT cells is the rapid onset.
Already a five-week-UVA-treatment is sufficient to
induce tumorigenicity in three of the four populations.
We previously showed that mere cultivation even
for >6 years and >300 passages did not lead to
tumorigenicity (Boukamp et al., 1997). Accordingly,
mock-treated control cells remained nontumorigenic.
However, increased temperature caused tumorigenic
conversion and, similar as shown here for UVA, was
correlated with induction of DNA ds breaks and new
chromosomal aberrations (Boukamp et al., 1999). Since
UVA as well as increased temperature cause oxidative
stress, they may well function through a similar pathway. As a major difference, however, tumorigenicity was
already evident after 5-week UVA treatment and further
increased after 15 weeks, while heat-induced tumorigenic conversion required >15 weeks (Boukamp et al.,
1999). Thus, tumorigenicity can be induced by highly
relevant UVA doses in a reproducible manner after only
few weeks of irradiation thereby making UVA a potent
tumor promoter for HaCaT cells.
One potential mechanism of induction of chromosomal aberrations could be critically short telomeres
leading to fusion bridge-breakage cycles (Murnane and
Sabatier, 2004). ROS, as induced by UVA radiation, are
known to cause breaks in G-rich sequences (von
Zglinicki et al., 2000). Actually, it was shown that
telomeric DNA was 5 to 10-fold more assailable than
nontelomeric DNA (Oikawa and Kawanishi, 1999).
Thus, telomeres should be optimal substrates for UVA
radiation. On the other hand, UVA-induced ROS
depend on the up-to-now unknown nuclear sensitizers.
Oncogene
UVA causes tumorigenic conversion of HaCaT keratinocytes
K Wischermann et al
4278
If they are not present at telomeres, these genomic
regions can be underrepresented in the DNA fragmentation due to the limited diffusion range of the UVA
generated ROS. Comparing telomere restriction
fragment lengths from HaCaT control cells with UVAtreated cells prior to injection as well as tumor-derived
cells, we did not find telomere shortening and we have
preliminary evidence that UVA radiation may also not
account for rapid telomere erosion when investigating
individual cells (D Krunic, K Wischermann and P
Boukamp, unpublished results). In fact, some of the
tumor-derived lines even had increased mean telomere
lengths. Thus, a telomere length-dependent mechanism
of genomic instability appears unlikely.
Instead we propose that UVA radiation induces ss
and ds breaks (the latter either directly or through
clustered and processed ss breaks) in chromosomal sites.
These are not repaired to completion immediately but a
small fraction of cells maintains the damage for
extended periods. Such breaks are prerequisite for
chromosomal breakage, which in turn is prerequisite
for chromosomal aberrations as identified in the
tumor-derived HaCaT cells. This finding in conjunction
with an increased formation of micronuclei 10 days
post irradiation, therefore, underlines the role of UVAdependent damage in inducing persistent genomic
instability, which finally results in chromosomal
aberrations. Furthermore, since we obtained a similar
damage profile with HaCaT cells and normal keratinocytes, this may well reflect the general action spectrum of
UVA radiation and may suggest a distinct role of UVA
in the multi-step process of skin carcinogenesis.
Materials and methods
Cell culture
HaCaT cells were routinely cultured as previously described
(Boukamp et al., 1988). Normal human keratinocyte cultures
were established from human trunk skin (approved by the
Ethics Commission of the Medical Faculty, University of
Heidelberg, Germany) and passaged as described (Boukamp
et al., 1990a). Second passage keratinocytes (grown in
keratinocyte growth medium, Epilife, Cascade Biologics,
Tebu-bio GmbH, Offenbach, Germany) were used for
irradiation experiments at a density of 70–90%.
G1 phase: enrichment of HaCaT and normal keratinocytes
HaCaT cells (80% confluent) were cultured in RPMI-1640
medium without cystein and methionine (No. R7513, Sigma,
Deisenhofen, Germany) for 48 h and for additional 14 h in
RPMI-1640 with 400 mM L-Mimosine (Sigma; adopted from
Krude, 1999). Normal keratinocytes (80% confluent) were
incubated in keratinocyte growth medium without pituitary
extract (Promocell, Heidelberg, Germany) in the presence of
10 ng/ml TGF-1 (R&D Systems, Wiesbaden, Germany) for
48 h. Cells were stained with 30 mg/ml propidium iodide and
the G1 enrichment was controlled using standard fluorescenceactivated cell sorting techniques (Supplementary Figure 3).
UVA irradiation of cells
For the exploratory experiment HaCaT cells were irradiated
with a daily increasing UVA dose from 7 to 42 J/cm2 (2–12 h
Oncogene
per day) over a 6-week period (irradiance of 1.0 mW/cm2 from
Philips PL-S 9W/10 blacklight lamps 330–400 nm). Cells were
irradiated in phenol-red-free culture medium and closed lid,
yielding a transmission of >70% UVA. For comparing single
versus split doses, the high intensity UVA source Mutzhas
UVASUN 3000, equipped with UVASUN safety filters and a
WG305 filter (emitting wavelengths in the 320–450 nm range;
Mutzhas et al., 1981), was used. Fluences were determined
with a UVA meter (Dr Hönle, Martinsried, Germany)
equipped with an UVA-sensor calibrated for emission from
315–400 nm. The emission spectrum was checked with an
Optronics OL-754 spectroradiometer (Optronics, Orlando,
FL, USA). Cells were irradiated in phosphate-buffered
saline (PBS) without lid at a distance of 40 cm for 100 s for
5 J/cm2 UVA and 400 s for 20 J/cm2, respectively (irradiance
50 mW/cm2). For DNA damage and repair analyses the
Phillips HB 404 tanning lamp (irradiance 21 mW/cm2)
equipped with an UVB blocking filter (Hoya UV34, Galvoptics, Basildon, UK), an infrared filter (M-UG2, Schott,
Grünenplan, Germany) and ventilators for cooling were used.
The fluence was measured with a Vilbert–Lourmat VLX-3W
radiometer. HaCaT cells and normal human keratinocytes
plated on glass slides were irradiated on ice in PBS in a
distance of 16 cm to the UVA source for 7.03 min (10 J/cm2) to
42.3 min (60 J/cm2). The cells were either analysed immediately
or after an incubation time as indicated. For each experiment
nonirradiated control cells were treated accordingly.
For all three UVA lamps, spectral characteristics were
comparable in emitting UVA with a maximum at 365 nm while
the UVB fractions was negligible with less then 0.005%
(Supplementary Figure 4).
Tumorigenicity test
About 5 106 cells in 100 ml were injected subcutaneously into
each side of the back of three (5 weeks) and four (15 weeks)
nude mice (Swiss/c nu/nu backcrosses). Tumor growth was
measured weekly and tumor size calculated in mm3 (Parangi
et al., 1996). Tumors were excised and formalin fixed for
standard hematoxilin/eosin staining. For recultivation, tumors
were shortly rinsed in ethanol (80%) and culture medium,
minced into 1 mm3 pieces and plated as explant cultures. After
tumor cell outgrowth, the cells were treated according to
Boukamp et al. (1988). Tumor cell lines were established from
4 5 J/cm2 5 weeks animal 3 right side (T1), 4 5 J/cm2 15
weeks animal 1 left side (T2), 4 5 J/cm2 15 weeks animal 4
right side (T3), 4 5 J/cm2 15 weeks animal 2 left side (T4),
1 20 J/cm2 15 weeks animal 3 left side (T5) and 1 20 J/cm2
15 weeks animal 4 left side (T6).
Alkaline and neutral comet assay
The alkaline comet assay was performed either directly after
irradiation or after a recovery of 2 or 4 h in complete culture
medium, according to Wischermann et al. (2007). The neutral
comet assay was performed directly after irradiation according
to the protocols described by Olive and Banath (Olive et al.,
1993). For both assays all steps were carried out on ice or in
the cold to minimize repair processes. For each data point two
to three areas on parallel slides were scored with 51–60 cells
each and DNA in tail (%) was calculated for each cell using
the image-analysis software ‘Komet 4’ (Kinetics Imaging Ltd,
Liverpool, UK). The median of DNA in tail (%) was
calculated for each area and the presented values are the
means of the medians of each data point. At least two
independent experiments were performed. The Mann–Whitney
rank sum test and t-test were used to determine statistically
significant differences between control and irradiated cells.
UVA causes tumorigenic conversion of HaCaT keratinocytes
K Wischermann et al
4279
Detection of g-H2AX
HaCaT cells and normal human keratinocytes (irradiated and
nonirradiated control cells) grown on glass slides were fixed
in methanol/acetone (1:1) for 10 min at 20 1C. Cells were
rehydrated in PBS for 15 min and permeabilized in PBS
containing 0.5% Triton X-100 for 20 min. After blocking for
30 min in 3% bovine serum albumin (BSA) in PBS, the cells
were incubated with mouse anti-g-H2AX (Upstate, Charlottesville, VA, USA; 1:100 in blocking solution) for 1 h and after
washing (3 10 min) with a secondary anti-mouse Alexa 488
antibody (1:800; Molecular Probes/Invitrogen, Karlsruhe,
Germany) containing 5 mg/ml bisbenzimid (Aventis Pharma,
Frankfurt, Germany) for 30 min. The average number of
g-H2AX foci/nucleus was determined using the ImageJ software (Wayne Rasband, http://rsb.info.nih.gov/ij/) or the cells
were classified into four groups (0, 1–5, 6–10 or >10 foci)
according to the number of foci per nucleus.
Micronuclei assay
For the cytokinesis-blocked MN assay, normal keratinocytes
were irradiated with 20 J/cm2 UVA and cultivated in the
presence of 4 mg/ml cytochalasine B (Sigma Aldrich, Munich,
Germany) for 24 h. Cells were fixed with methanol and acetone
for 5 min each at 20 1C, blocked with 3% BSA and incubated
with a human anti-all-centromere antibody (1:100; Antibodies
Inc., Davis, CA, USA) for 30 min at 37 1C. After washing in
PBS, cells were incubated with a rabbit anti-human-FITC
(1:200; Dako, Hamburg, Germany) for 30 min at 37 1C and
nuclei were counterstained with 0.1 mg/ml 40 ,6-diamidino2-phenylindole
dihydrochloride
(DAPI;
Vectashield,
Wertheim, Germany). From 1000 binucleated cells the number
of cells with MN (expressed as % MN-positive cells) and the
ratio of MNs with and without centromere signals was
determined. For long-term maintenance of MNs, the cells
were irradiated with 10, 20 and 40 J/cm2 UVA and cultivated
for 5 and 10 days without Cytochalasin-B block. MN staining
was performed as described.
Statistical significant differences between control and
irradiated cells were assessed with the w2-test (Cohen, 1960)
and significance of dose-dependency with the Cochran–
Armitage test (Cochran, 1954; Armitage, 1955).
Multiplex fluorescence in situ hybridization analysis
Metaphase spreads were prepared and combinatorial labeling
of whole chromosome painting probes and multicolorhybridization was performed as described (Popp et al., 2000).
Telomere length measurement by Southern blot analysis
Genomic DNA was isolated using DNAzol (Gibco BRL,
Eggenstein, Germany). DNA (5 mg) was digested with Rsa I
(Roche, Mannheim, Germany) and subjected to electrophoresis.
Telomere length measurements by Southern blot was
performed as described (Figueroa et al., 2000).
Acknowledgements
We thank Cees Guikers for the first experiments with daily
exposures, Heinrich Steinbauer for performing the tumor
experiments, Heidi Holtgreve for her excellent technical
assistance in M-FISH hybridization and Stefan Henning for
comparing the different UVA-sources. This work was supported in part from the Sander Stiftung, Deutsche Krebshilfe
e.V. and EU (LSHC-CT-2004-502943) (both to PB), QLK41999-01084 (to PB and KSK) as well as the German Research
Foundation (SCHA 411/10-1,-2) and the European Community Marie Curie Program MEIF-CT-2005-023821 (to AR).
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Supplementary Information accompanies the paper on the Oncogene website (http://www.nature.com/onc)
Oncogene