Advances in Anthropology
2012. Vol.2, No.2, 49-56
Published Online May 2012 in SciRes (
Copyright © 2012 SciRes.
Genotoxicity Assessment of Birch-Bark Tar—A Most Versatile
Prehistoric Adhesive
A. Baumgartner1,2, M. Sampol-Lopez1, E. Cemeli1, T. E. Schmid3, A. A. Evans4,
R. E. Donahue4, D. Anderson1
1Division of Biomedical Sciences, University of Bradford, Bradford, UK
2Department of Pediatric Cardiology, University of Leipzig, Leipzig, Germany
3Department of Radiotherapy and Radiooncology, Klinikum Rechts der Isar,
Technische Universität München, Munich, Germany
4Division of Archaeological, Geographical and Environmental Sciences, University of Bradford, Bradford, UK
Received February 22nd, 2012; revised March 30th, 2012; accepted April 11th, 2012
In the Mesolithic, birch-bark tar was commonly utilized across Europe and much of Asia as an adhesive
to bind, seal and coat surfaces, but also quite frequently it was found to be chewed. The tar is known to
contain biomarker triterpenoid compounds like betulin, crucial in preserving food and for medical appli-
cations. Aqueous, ethanolic and DMSO extracts were prepared from solid birch-bark tar and evaluated in
vitro for the induction of DNA damage using Comet, micronucleus and sister-chromatid-exchange assays.
Additionally, apoptosis induction was assessed. For the ethanolic extract, only the Comet assay showed a
significant increase of DNA damage. All three extracts were able to significantly induce apoptosis. Thus,
birch-bark tar seems capable of inducing genotoxic damage as well as apoptotic effects possibly originat-
ing from the triterpenoids’ antimicrobial properties. We examine why prehistoric tar is found with tooth
marks, the beneficial effects of birch-bark tar, and evidence for increased genotoxic risk upon exposure.
Keywords: Birch-Bark Tar; Genotoxicity; Micronuclei; Comet Assay; Apoptosis; Anthropology
Birch-bark tar has long been known as an adhesive. Dating
back to the Middle Palaeolithic period, pieces of birch-bark tar
were found in a lignite open-mining pit near Königsaue, Ger-
many, being made by Neanderthal hunter-gatherers around
48,000 years before present (BP) (radiocarbon dating) (Grün-
berg, 2002) and possibly as early as 80,000 BP (geo-strati-
graphic dating) (Koller et al., 2001). By the start of the Meso-
lithic period around 9500 BP, birch-bark tar was extensively
used across Europe from Norway to Greece (Stern et al., 2006)
and this continued during the Neolithic period (Regert, 2004).
So far, little is know about the actual manufacturing process in
prehistoric times despite the wide-spread common use of this
pyrolysed plant tar throughout Europe. The tar was not only
employed for adhering, sealing, caulking and waterproofing
(Charters et al., 1993) but also as an antifungal and antibacterial
agent (Urem-Kotsou et al., 2002). Interestingly, birch-bark tar
was also chewed (Aveling, 1997)—probably due to these pre-
cise antimicrobial properties (Aveling & Heron, 1999). By as-
sessing the light stable isotope δ13C content, prehistoric birch-
bark tar from Greece can easily be distinguished from that of
northern Europe. So, even if we know little about the manufac-
turing process in early prehistoric times, it does seem evident
that such adhesives were probably made and used locally rather
than acquired through long-distance trading (Stern et al., 2006).
Five triterpenoid constituents from the lupane family: betulin
(lup-20(29)-ene-3β,28-diol), lupeol (lup-20(29)-ene-3β-ol), be-
tulinic acid, lupenone and betulone are characteristic for the
white birch bark, directly linking the tar chemically to the bo-
tanical source (Figure 1). The pentacyclic triterpene alcohol,
betulin, can be found in concentrations up to 30% of dry weight
of birch bark (Betula pendula) (Alakurtti et al., 2006). On the
other hand, other tar components, i.e. transformants of betulin,
allobetulene (lupa-2,20(29)-diene-28-ol), and of lupeol, lupa-
diene (lupa-2-20(29)-diene), act as biomarkers of human action
originating from pyrolytic dehydration during the manufactur-
ing process (Dudd & Evershed, 1999; Regert, 2004). When
heating (pyrolysis) the bark of birch trees (e.g. European white
Figure 1.
Triterpenes of the outer birch bark of Betula
pendula, B. papyrifera and B. neoalaskana.
Modified according to (Krasutsky, 2006).
birch Betula pendula Roth) under reducing conditions with
limited air supply and a temperature of over 300˚C, the distilla-
tion product is an odorous dark-brown viscous pitch (Dudd &
Evershed, 1999).
Despite the mentioned beneficial qualities as a common tool
for craftsmen, birch-bark tar originates from destructive heat
distillation also generating an abundance of possible genotoxic
substances, volatile and nonvolatile. As yet no toxicological
studies have been undertaken to investigate the genotoxic po-
tential of birch-bark tar. This present in-vitro study uses solvent
extracts of water, ethanol and dimethyl sulfoxide (DMSO) from
birch-bar tar to evaluate its genotoxic potential. Whole blood
cultures as well as separated lymphocytes were treated in vitro
with birch-bark tar extracts and analysed concentrating on clas-
sical cytogenetic endpoints such as induction of micronuclei
(MN assay) and sister chromatid exchanges (SCE assay) as
well as changes in DNA integrity (Comet assay). Additionally,
flow cytometry (FCM) was applied to evaluate the tar’s poten-
tial to induce cellular apoptosis or necrosis.
Evidence for Us e as a M ed icinal Produ ct
There is ethnographic evidence for birch-bark tar and other
products from birch trees in healing practices among traditional
cultures located in the European and Asian boreal forests. The
Yakut of Siberia, renowned for their desire for and care of
horses, will use birch-bark tar to close wounds and use heated
birch bark for bandaging (Sieroszewski et al., 1993: p. 361).
Birch-bark tar was made by the Fisher Lapps in Inari in the
same way they recover tar from pine. They dug a hole on the
side of a hill; the bark was rolled up to form a pipe and filled
with more bark and resinous wood. The pipe was place in the
hole on a slant with a pot or wooden container placed at the
lower end. It was then covered with peat, sand and stone.
Burning extended from a few hours to a whole day (Itkonen,
The Lapps appear to use a variety of remedies, including
birch derived treatments, for toothaches. The person may chew
birch-bark tar or pine pitch or simply rub the tooth with tar or
pine pitch. When the ache is setting in, they will bite on a piece
of birch wood until the pain disappears. Another interesting
practice, suggestive that there is a medicinal property within the
tree, occurs in winter when a chrysalis from a branch of birch is
warmed up and the larvae taken out and pressed into the cavity
in a tooth (Itkonen, 1984). Among the Inari-Lapps the elderly
tend to have better teeth than younger Lapps and although their
teeth may be much worn, they seem to have suffered very little
from tooth disease. They suggest it is because they chew [birch]
bark, which keeps their teeth in good condition (Itkonen, 1984).
Rheumatism is also treated with birch treatments by the
Lapps. Tar water is made from pine pitch, birch-bark tar, or
both and drunk. Young, resinous birch leaves will be layered on
a bed or put in a sack and the patient, nude, will lie on them.
Gout and rheumatism will also be treated by placing tinder on a
piece of birch bark and blowing on it to start a small fire. The
birch bark is held against the infected area and the fire blown
on until it burns through the birch bark and burns into the skin,
water and the rheumatism will flow from the burnt hole in the
skin. The hole should be kept open as long as possible. The
same process may also be used with a birch gnarl (Itkonen,
1984). For coughing, tar water is poured on a hot flagstone and
the steam is breathed by the patient. Prior to bed, birch catkins
or birch bark will be chewed by the patient. The saliva will be
swallowed and the chew spat out. In some regions, bark from
an old birch will be boiled and taken two or three times each
day. Whooping cough is treated with a tea made from boiling
birch catkins. Birch bark also is used in a variety of ways to
treat diarrhea. Birch bark would be ground and mixed with
meal and water and then baked. The resulting cake is eaten
plain. Birch bark will also be boiled until the water turns red
and has a very strong taste. It is mixed with sugar before being
taken. Dry birch bark may also be chewed. The medicinal ac-
tion of birch bark is attributed to tanning agents within the bark
(Itkonen, 1984: p. 918). This is also the explanation for the
value of willow bark, which is used in a large number of reme-
dies, but, research has shown that it is the occurrence of sali-
cylic acid in the bark.
Material and Methods
Chemical Extracts and Treatment of Cells Template
A sample of experimentally produced birch-bark tar was
provided for analysis by the Hunter-Gatherer Laboratory of the
Division of Archaeological, Geographical and Environmental
Sciences, at the University of Bradford. Three different solvents,
water, ethanol and DMSO, were used to prepare extracts with a
concentration of 100 mg/ml each. The suspension was thor-
oughly mixed and kept for 7 days at 37˚C. After removal of the
non-dissolvable precipitate, the extracts were then diluted to
working stock concentrations of 10, 100 and 1000 µg/ml. Hu-
man lymphocytes were treated in vitro with final concentrations
of 0.1, 1.0 and 10 µg/ml in culture.
Blood Samples, Culture and Treatment of
Lymphocytes for the MN, SCE and FCM-Apoptosis
After informed consent, blood from healthy volunteers (do-
nor A, male, age 33, and donor B, female, age 28) was obtained
in heparinised vacutainers (Greiner) by venepuncture. Ethical
approval was provided by the University of Bradford’s Re-
search Ethics Subcommittee involving human subjects (refer-
ence number 0405/8). Whole blood (0.5 ml each) was added to
plastic culture flasks (25 cm2, Corning) containing RPMI 1640
medium with Glutamax (Gibco), 15% foetal bovine serum (FBS,
Sigma) and 1% penicillin-streptomycin solution (Gibco), This
basic culture medium was supplemented with 1% (v/v) phyto-
haemagglutinin-M (PHA-M, Gibco) to initiate proliferation of
T-lymphocytes in vitro. For the SCE assay, additional supple-
mentation with 10 µM of 5-bromo-2’-deoxy-uridine (BrdU;
Sigma) for cell cycle control was needed. Cultures were incu-
bated for 72 hours at 37˚C in 5% CO2 in air. For the MN, SCE
and FCM-apoptosis assays, the cultures were treated 24 hours
after the start. The volume of treatment stock solutions added
was kept constant to 1% of the total cell culture volume. For the
MN assay, 44 hours after the start, cytochalasin B (Sigma) was
added to produce a final concentration of 6 µg/ml in order to
arrest cytokinesis for the rest of the incubation time. Cells were
then harvested after the end of the 72 hours culture period. For
the SCE assay, cells were arrested in metaphase 2 hours before
the end of the culture by adding colcemid (Sigma) to a final
concentration of 0.1 µg/ml. The cultures for the FCM-based
apoptosis assay received no supplementations. Cells treated
with mitomycin C (Sigma) served as a positive control for the
Copyright © 2012 SciRes.
MN assay (0.2 µM) and for the SCE and FCM-apoptosis assays
(0.1 µM).
Lymphocyte Isolation for the Comet Assay
Seven healthy individuals (4 males and 3 females) with an
average age of 30.00 ± 2.25 years volunteered to donate blood.
Lymphocytes were isolated using a Ficoll gradient (Lympho-
prep, Axis-Shield) according to the manufacturer’s instructions.
Once a pellet with lymphocytes was obtained, it was resus-
pended in FBS and transferred into a cryovial containing DMSO
(9 parts FBS, 1 part DMSO). This cell suspension was frozen at
–20˚C overnight and then transferred to –80˚C for long-term
Cell Viability for Lymphocytes
Cell viability was measured directly after treatment by Try-
pan blue exclusion (Krause et al., 1984; Pool-Zobel et al., 1992)
and fluorescein diacetate/ethidium bromide (Hartmann & Speit,
1997) (all chemicals from Sigma). The treatment concentrations
selected for the present investigation resulted generally in vi-
ability rates over 90%, but always higher than the required 75%
(Henderson et al., 1998).
Metaphase Prep aration for SCE Assay and
Differential Giemsa Staining
After 72 hours, the cell suspensions were transferred to Fal-
con tubes and washed twice with phosphate-buffered saline
(PBS, Sigma) at 37˚C. Each washing step was followed by a
centrifugation (190 g for 10 min) and removal of the super-
natant. The subsequent hypotonic treatment was carried out
with 75 mM KCl (Sigma) for 20 min at 37˚C. After centrifuga-
tion for 8 min and removal of the supernatant, the cells were
fixed in Carnoy’s solution (3 parts methanol/1 part glacial ace-
tic acid, both from Sigma). The fixation step was repeated out
three times. Chromosome preparations were obtained by pipet-
ting 20 µl of cell suspension onto clean glass slides. After air-
drying, the preparations were aged for a minimum of one week
at room temperature. Four slides per treatment (two cultures per
treatment and each culture prepared in duplicate) were em-
ployed. For differential staining, fluorescence plus Giemsa
(FPG) staining was applied (Perry & Wolff, 1974). Briefly,
slides with aged metaphase spreads were incubated in 1 µg/ml
bisbenzimide (Sigma)/Weise buffer (8 mM Na2HPO4, 3.6 mM
KH2PO4, pH 7.2) solution for 20 min in the dark at room tem-
perature, then rinsed and incubated in PBS for 90 min under a
UV-A/B lamp. After a final 30 min incubation at 60˚C in 2x
SSC (Sigma), the slides were rinsed in pure water and chroma-
tin was stained with 5% Giemsa (BDH) in Weise buffer during
a 5 min incubation.
Cytokinesis-Block MN Assay
After 72 hours, the cultures were transferred into Falcon
tubes, centrifuged (190 g for 8 min) and the supernatant was
removed. This was followed by a hypotonic treatment in 75
mM KCl for 15 min at 4˚C followed by another centrifugation.
The cell pellets were then subjected to fixation with fresh Car-
noy’s solution. To each preparation three drops of formalde-
hyde (38%, BDH) were added. The fixation was repeated twice
without the addition of formaldehyde. Then, the cell suspension
was dropped on clean glass slides (20 µl per drop, 2 drops per
slide) and allowed to air-dry. Four slides per treatment (two
cultures per treatment and each culture prepared in duplicate)
were employed. Slides were stained with 5% Giemsa in phos-
phate buffer (pH 6.8) for 10 min, rinsed in pure water and air-
dried before being mounted with cover-slips. The scoring crite-
ria were described in detail by Fenech and colleagues (Fenech
et al., 2003; Fenech, 2007).
Comet Assa y
DNA strand breaks were measured with the alkaline Comet
assay using the method previously described (Anderson et al.,
1997; Anderson et al., 1998) and reviewed (Tice et al., 2000).
In brief, microscope slides were covered with a basic layer of
1% normal melting-point agarose in water (Invitrogen). The
slides were dried overnight and then stored at room temperature.
Cryovials with frozen lymphocytes were quickly thawed at
37˚C and added to 1 ml PBS containing 1% volume of the
chemical treatment. After a treatment interval of 30 min at 37˚C
in Eppendorf® tubes, followed by a centrifugation at 700 g
(table-top centrifuge) for 5 min, 900 µl of the supernatant were
discarded and the pellet resuspended. The remaining lympho-
cyte suspension (100 l) was then mixed with 100 l of 1% low
melting-point agarose in PBS (Invitrogen). Of this suspension,
100 l were pipetted on an agarose-coated slide. Slides were
covered with cover slips and left on an ice-cold surface until the
gel set. After removing the cover slip, a protective third layer of
0.5% low melting point agarose in PBS was added, spread us-
ing a cover slip and again allowed to set for approximately 5
min on an ice-cold surface. Then, the slides were immersed in
lysis solution (2.5 M NaCl, 100 mM EDTA, 10 mM Tris, 1%
Triton X-100 and 10% DMSO, pH set to 10 with NaOH, all
chemicals from Sigma). The slides were subjected to electro-
phoresis in a pH 13.5-electrophoresis buffer (1 mM EDTA and
300 mM NaOH). The DNA was allowed to unwind within 30
min at 4˚C in the electrophoresis buffer prior to electrophoresis
being performed at 4˚C for 30 min at a constant 25 volts (0.75
V/cm). The amperage was kept at 295 - 300 mA (adjusted by
removing or adding electrophoresis buffer). Tris buffer (0.4 M,
pH 7.5; Sigma) was used to neutralise the alkali buffer. Then,
50 µl ethidium bromide (20 µg/ml; Sigma) were added onto
each slide, covered with cover slips and analysed within 3
hours. The slides were examined at ×400 magnification on a
fluorescence microscope (Leica, UK) equipped with a BP546/10
excitation filter and a 590 nm barrier filter. Fifty nuclei were
scored and evaluated per concentration. Seven independent
experiments were performed with isolated lymphocytes from 7
different subjects. A computerised image analysis system (Ko-
met 4.0, Kinetic Imaging, Liverpool, UK) was used. In the
Comet assay, Olive tail moment (OTM) was the parameter of
choice since it was reported to provide good correlation with
the dose of genotoxic agents used (Kumaravel & Jha, 2006).
Flow Cytometry (FC M) Apoptosis Ass a y
After 72 hours of treatment, the tubes containing the treat-
ment were centrifuged; the supernatant discarded and replaced
by 40% RPMI 1640 with 40% FBS and 20% DMSO. In the
first instance, the pellets were frozen overnight at –20˚C and
then transferred to –80˚C. The tubes were shipped in dry ice to
the Department of Radiotherapy and Radiooncology in Munich.
Copyright © 2012 SciRes. 51
Copyright © 2012 SciRes.
and DMSO were evaluated in vitro for their genotoxic potential
to induce DNA damage in lymphocytes from healthy individu-
After storage at –80˚C, the pellets were thawed and centrifuged
for 10 min at 150 g at 8˚C. The supernatant was discarded and
replaced by 5 ml RPMI 1640 + 20% FBS and newly centri-
fuged in order to wash the pellet. The pellet was resuspended in
the described medium, transferred to cell culture flasks and
incubated overnight at 37˚C. Next day, the cultures were trans-
ferred into tubes, centrifuged at 150 g, the supernatant dis-
carded and resuspended in 5 ml Annexin buffer (BD) and cen-
trifuged again in the same conditions. The supernatant was
discarded and the pellets were resuspended in 100 µl Annexin-
V-FITC/Annexin buffer (1:100) and left in the dark for 15 min.
Using the alkaline Comet assay we investigated the DNA
damage generated in vitro in lymphocytes based on seven ex-
periments with lymphocytes from seven volunteers. The extent
of the DNA damage on a single-cell level as seen in the Comet
assay was described by the Olive tail moment (OTM) parame-
ter. While the aqueous extract did not induce any DNA damage
within the chosen dose range (0.1, 1 and 10 µg/ml), the ethano-
lic extract gave a clear dose response almost doubling the OTM
when compared to the negative control with p < 0.001 for all
three doses (Figure 2). The DMSO extract on the other hand
only induced slight DNA damage above the negative control,
which was statistically significant at 0.1 µg/ml (p < 0.05) and
10 µg/ml (p < 0.01) but not at a dose of 1 µg/ml.
The volume was increased up to 495 µl with Annexin buffer.
Immediately before the analysis, 5 µl propidium iodide (Invi-
trogen) were added to achieve a 1:100 final dilution. Cells were
analysed on a Becton Dickinson FACSCalibur flow cytometer
using Cell Quest Software. Only single cells were gated for
fluorescence analysis. The induction of genetic damage was also assessed by using
the cytokinesis-block micronucleus (MN) assay. In Table 1, the
results of the MN assay are listed. None of the extracts was able
to increase the MN frequencies in vitro at a dose of 1 µg/ml.
However, there was a trend visible which showed that the
DMSO extract almost doubled the MN frequency in lympho-
cytes from both healthy individuals (donors A and B), but this
increase was not significant when compared to the negative
control. The nuclear division index (NDI) however, yielded in a
highly significant decrease (1.77; p < 0.001) for donor A when
treating with the 1 µg/ml DMSO extract. Donor B on the other
hand did not show a decline in NDI for the DMSO extract.
Neither pure water nor DMSO showed any positive effect.
Statistical Analysis
For the Comet assay, after verifying normality (Shapiro-
Wilks test) data were analysed by a one-way ANOVA with a
Dunnett’s post-hoc test using mean ± SE of tail moments (n =
7). For the MN assay, a one-tailed Fisher’s exact test was ap-
plied to all parameters except the nuclear division index (NDI)
where a χ2-test was applied. For the SCE assay, a t-test was
applied for SCE and χ2-test for the proliferation rate index
(PRI). Data from the FCM-apoptosis assay were statistically
assessed with a χ2-test with Yate’s correction. The statistical
software used was Statistica 6.1 from StatSoft Inc. (Tulsa, OK,
USA). For the ethanolic extract, a significant decrease in NDI from
1.87 to 1.67 (p < 0.001) was seen only in lymphocytes from
donor A, however, it seems 1% ethanol on its own entirely
contributed to this decrease in the NDI. This was not observed
for donor B. For the aqueous extract, a significant decrease in
In this study, three birch-bark tar extracts in water, ethanol
Figure 2.
Dose responses for aqueous (H2O), ethanolic (ETOH) and DMSO extracts of prehistoric birch-bark tar using the alkaline Comet assay. Data were
assessed for normality using the Shapiro-Wilks test. Data resulted in a normal distribution. Data were analysed by one-way ANOVA with a Dunnett’s
post-hoc test. For statistical interpretation, each concentration for each single extract was compared to the negative control (*p < 0.05, **p < 0.01, ***p
< 0.001). Data shown correspond to mean ± SE of tail moments in seven individuals.
Table 1.
Evaluation of aqueous, ethanolic and DMSO extracts of prehistoric birch-bark tar with the micronucleus (MN) assay.
Treatment concentration NDI % BiMN per
1000 BiBi containing
MN NPB per
1000 BiBiBuds per
1000 Bi MN per
1000 Mono Mono containing
Negative control 1.87 59.137.86 7.86 0.00 0.00 6.90 6.90
H2O 1% 1.85 62.755.87 5.87 0.00 0.00 3.91 3.91
EtOH 1% 1.68*** 63.717.91 7.91 0.00 0.99 7.11 7.11
DMSO 1% 1.93* 70.013.91 3.91 0.00 0.00 1.30 1.30
Birch-bark tar H2O extract, 1 µg/ml 1.88 64.049.82 6.88 0.00 0.00 7.77 7.77
Birch-bark tar EtOH extract, 1 µg/ml 1.67*** 64.035.99 5.99 0.98 0.00 0.00 0.00
Birch-bark tar DMSO extract, 1 µg/ml 1.77*** 67.7814.00 14.00 0.00 1.00 3.98 3.98
Donor A
MMC 0.2 µM 1.79* 58.7829.67
*** 29.67 *** 0.00 0.00 3.77 3.77
Negative control 1.54 44.899.94 9.94 0.00 0.00 3.89 3.89
Birch-bark tar H2O extract, 1 µg/ml 1.38*** 30.2213.56 13.56 0.00 0.00 3.94 3.94
Birch-bark tar EtOH extract, 1 µg/ml 1.52 42.768.12 8.12 0.00 0.00 2.85 2.85
Birch-bark tar DMSO extract, 1 µg/ml 1.76*** 55.9717.86 16.87 0.00 0.00 2.91 2.91
Donor B
MMC 0.2 µM 1.66*** 51.7830.00
** 30.00 ** 0.00 1.00 9.28
* 7.73
Values for the parameters scored: NDI (nuclear division index), % Bi (% binucleated cells), MN per 1000 Bi (total number of MN per 1000 binucleated cells), Bi con-
taining MN (number of binucleated cells containing MN out of 1000 binucleated), NPB per 1000 Bi (total number of nucleoplasmic bridges per 1000 binucleated cells),
BiBuds per 1000 Bi (total number buds per 1000 binucleated cells), MN per 1000 Mono (total number of MN per 1000 cells) and Mono containing MN (number of
mononucleated cells containing MN out of 1000 cells). Statistical comparisons were carried out between the negative control and each of the treatments. One-tailed
Fisher’s exact test was applied to MN per 1000 Bi, Bi containing MN, NPB per 1000 Bi, BiBuds per 1000 Bi, MN per 1000 Mono and Mono containing MN. χ2-test
was applied to NDI (*p < 0.05, **p < 0.01 and ***p < 0.001).
NDI was shown only donor B together with a non-significant
increase in MN. When focussing on other parameters like the
frequency of MN in mononucleated cells, nuclear buds and
nuclear plasmatic bridges (NPB), no significant increases or
modulations were recorded.
The SCE assay is able to detect an increased exchange of
DNA stretches between two sister chromatids of a duplicating
chromosome after treatment. When using this assay, the aque-
ous and the ethanolic extract (1 µg/ml) exhibited statistically
significant decreases in the frequency for SCEs per 50 cells (see
Table 2), but only for donor A. For donor B, no effect was seen
regarding SCEs.
This present investigation also evaluated the induction of
apoptosis employing a flow cytometry based apoptosis assay
(Vermes et al., 1995). Lymphocytes were treated in culture
similarly as for the MN and SCE assays, however, without any
addition of cell cycle inhibitors. Table 3 shows that the sol-
vents 1% ethanol and 1% DMSO were capable of significantly
inducing apoptosis (both slightly above 2.5 fold, p < 0.001).
For donor A, all the extracts showed a highly significant dose
response (p < 0.001) with increasing apoptotic frequencies. The
aqueous, ethanolic and DMSO extracts showed increases of
6.2-fold, 7-fold and 11.9-fold, respectively, when compared to
the negative control. Even when comparing the ethanolic and
the DMSO extracts to the solvent controls they still showed
increases of 2.9-fold and 4.6-fold, respectively. For donor B,
apoptosis was statistically significantly induced by the birch-
bark tar extracts, however, at much lower levels when com-
pared to donor A.
In 1991, the Tyrolean Iceman “Oetzi” was found in a slightly
Table 2.
Evaluation of aqueous, ethanolic and DMSO extracts of prehistoric birch-
bark tar with the sister-chromatid exchange (SCE) assay.
Treatment concentration SCE/cell ± S.E.PRI
Negative control 7.12 ± 0.44 1.74
H2O 1% 6.00 ± 0.28 1.98**
EtOH 1% 8.30 ± 0.92 1.89*
DMSO 1% 7.54 ± 0.68 1.85
Birch-bark tar H2O extract, 1 µg/ml 4.02 ± 0.26*** 1.82
Birch-bark tar EtOH extract, 1 µg/ml 5.52 ± 0.38** 1.89*
Birch-bark tar DMSO extract, 1 µg/ml 6.90 ± 0.38 1.96**
Donor A
MMC 0.1 µM 12.16 ± 1.00*** 1.98**
Negative control 7.82 ± 0.54 1.84
Birch-bark tar H2O extract, 1 µg/ml 7.06 ± 0.56 1.87
Birch-bark tar EtOH extract, 1 µg/ml 7.36 ± 0.61 1.81
Birch-bark tar DMSO extract, 1 µg/ml 7.78 ± 0.69 1.83
Donor B
MMC 0.1 µM 15.36 ± 0.86*** 1.65**
Values for the parameters scored: SCE/cell ± S.E. (mean of sister-chromatid ex-
changes for 50 cells ± standard error) and PRI (proliferative rate index). Statistical
comparisons were carried out between the negative control and each of the treat-
ments. The t-test was applied for SCE and χ2-test for PRI (*p < 0.05, **p < 0.01
and ***p < 0.001).
retrieving glacier in the Alps having died approximately in the
Late Neolithic around 5200 BP. The flint stones of his arrows
as well as the copper blade of his hatchet were fixed onto
wooden shafts with an organic agglutinant (Sauter et al., 2000).
High levels of betulin and lupenol and lower levels of betulinic
Copyright © 2012 SciRes. 53
Table 3.
Evaluation of aqueous, ethanolic and DMSO extracts of prehistoric birch-
bark tar with the FCM-apoptosis assay.
Treatment/Concentration (µg/m l ) Apoptosis (%)
H2O 1% (Negative control) 5.66
EtOH 1% 14.29***
DMSO 1% 14.65***
Birch-bark tar H2O extract, 1 µg/ml 35.01***
Birch-bark tar EtOH extract, 1 µg/ml 42.06***
Birch-bark tar DMSO extract, 1 µg/ml 67.24***
Donor A
MMC 0.1 µM 33.12***
H2O 1% (Negative control) 3.95
Birch-bark tar H2O extract, 1 µg/ml 12.41***
Birch-bark tar EtOH extract, 1 µg/ml 17.68***
Birch-bark tar DMSO extract, 1 µg/ml 14.92***
Donor B
MMC 0.1 µM 31.55***
The values for the apoptosis are presented as a percentage of the apoptotic events
and the total gated. Two independent experiments were performed on 2 subjects.
The χ2-test with Yate’s correction was applied (*p < 0.05, **p < 0.01 and ***p <
0.001). As a positive control mitomycin C (MMC) was used.
acid showed that this adhesive was birch-bark tar originating
exclusively from the bark of Betula pendula birch trees (Hayek
et al., 1989; Sauter et al., 2000). It was produced via destructive
dry distillation of dried bark under anoxic conditions with tem-
peratures around 340˚C leaving betulin and lupenol as bio-
markers almost unchanged. Higher temperatures in the produc-
tion process (>400˚C) would have otherwise resulted in heat
degradation markers like lupenone (lup-20(29)-ene-3-one) and
lupadiene (lupa-2,20(29)-diene) (Regert & Rolando, 1996; Koller
et al., 2001), which were only found in small concentrations. In
addition to the terpenoids high amounts of the lipid material
suberin can be found in the tar (Modugno et al., 2006) prevent-
ing water from penetrating the material.
Betulin, lupenol and especially betulinic acid have recently
been proven to exhibit anti-malarial, anti-HIV, anti-fungal and
antibacterial properties, and to be selectively cytotoxic against a
number of tumour types (Yogeeswari & Sriram, 2005; Alakurtti
et al., 2006; Dominguez-Carmona et al., 2010). This might have
been the reason why our ancestors used birch-bark tar to pre-
serve fermented beverages (Urem-Kotsou et al., 2002). It has
been argued that the tar of the birch bark was also quite gener-
ally used as “chewing gum” in prehistoric northern Europe
stretching back at least 9,000 years (Aveling, 1997; Aveling &
Heron, 1999). Potential explanations for this behaviour include
the possible use as a narcotic, in rituals or for medical reasons
due to the tar’s antimicrobial properties. But the explanation for
chewing the tar could be much simpler such as for removing
milk teeth or just to pacify children as most chewers were 6 to
15 years old (Aveling, 1997; Aveling & Heron, 1999). Ethno-
graphic information presented earlier showed that not long ago
birch-bark tar was used for various dental and gum problems,
which would also explain the tooth or teeth impressions in tar
recovered from prehistoric contexts. Whatever its use, it seems
that in prehistoric times birch-bar tar was a commonly used
product which came in prolonged contact with skin and the oral
cavity. Inhalation of more volatile chemical components cannot
be excluded when the tar was made and used in its viscous form
after heating for various processes.
In this study we investigated the genotoxic potential of
birch-bark tar. Knowing the fact that e.g. betulin and betulinic
acid as pure chemicals are almost insoluble in aqueous media
(Drag et al., 2009), three different extraction solvents generally
used as solvents in toxicology were chosen due to their differ-
ent hydrophilic properties: pure water, ethanol and DMSO. At
the highest dose of 10 µg/ml the ethanolic extract of birch-bark
tar generated an OTM of 4.59 ± 0.91 in the Comet assay, which
shows a highly significant induction of DNA damage (Figure
2). The induced damage ranged almost at the same level as that
of hydrogen peroxide, a genotoxin widely used as a positive
control, which generated at 100 µM OTM values of 5.59 ± 0.36
(data not shown in Figure 2). The DMSO extract induced only
slight but significant DNA damage while the aqueous extract
did not show any induction of damage. It has been shown that
betulinic acid can generate reactive oxygen radicals (ROS) and
also inhibit topoisomerase I, contributing to the DNA damage
seen with the Comet assay (Eiznhamer & Xu, 2004). Despite
the fact that triterpenes are the most abundant group of com-
pounds in birch-bark tar as seen with gas chromatography
(Charters et al., 1993; Aveling & Heron, 1999), other genotoxi-
cally active compounds even if lower concentrated might con-
tribute to the overall genotoxic damage.
In our study, the micronucleus assay only showed a trend
towards the induction of micronuclei (Table 1). The 2-fold
increase compared to the control for the DMSO extract, how-
ever, was not found to be statistically significant. Only the nu-
clear division index was significantly decreased for the ethano-
lic and the DMSO extract. For the sister-chromatid exchange
assay, no significant induction of SCEs was seen (Table 2).
Lupenol and betulinic acid, compounds found in birch bark
have been shown to be non-toxic for normal cells but highly
potent chemopreventive and chemotheraputic agents alleviating
inflammation and cancer (Saleem et al., 2009). This fact most
likely supports the use of birch-bark tar as an antimicrobial in
prehistoric times as no adverse effects were obvious. Due to the
remarkable biological and medical aspects of birch-bark triter-
penes, especially the abundant triterpenoids betulin, betulinic
acid and lupenol these compounds are nowadays forming a new
class of anti-cancer bioactive compounds (Krasutsky, 2006).
Betulin by itself has proven to also bind to GABA receptors
thus showing beneficial central nervous system effects in vivo
in mice (Muceniece et al., 2008).
Early apoptosis is characterised by the translocation of phos-
phatidylserine (PS) from the inner to the outer side of the lipid
bilayer of the plasma membrane, disturbing the normal asym-
metrical distribution of PS. Once exposed to the extracellular
environment, PS serves as a signal for phagocytosis (Schlegel
& Williamson, 2001). The exposed PS can also act as an anchor
for a Ca2+-dependent phospholipid binding protein, Annexin V.
Conjugated to a fluorochrome, Annexin V-FITC can quantita-
tively and rapidly detect early apoptotic cells when used in a
FCM-apoptosis assay (Vermes et al., 1995). To exclude ne-
crotic cells the vital dye propidium iodide, which can only
penetrate leaky plasma membranes and stain nuclear chromatin,
is used. The results for such a flow cytometry apoptosis assay
in our study showed for the aqueous, ethanolic and the DMSO
extracts of birch-bark tar a significant induction of apoptotic
cells—more pronounced for donor A showing a more distinct
dose-response than for donor B (Table 3). It has been shown
Copyright © 2012 SciRes.
that betulinic acid can induce apoptosis in sensitive cells in a
p53 and CD95 independent fashion functioning possibly through
a mitochondrial-mediated pathway, however, it can also gener-
ate reactive oxygen species (ROS) and inhibit topoisomerase I
(Eiznhamer & Xu, 2004).
While the Comet assay detects genetic integrity changes
mostly in the form of alkaline labile sites, single- and double-
strand breaks, the MN assay and the SCE assay primarily target
cytogenetic changes in the cell. These changes have already
escaped inherent repair mechanisms and have manifested them-
selves for instance as MN, i.e. chromosomal fragments. When
taking the results of the all three employed assays into account,
it seems that the genotoxic impact of birch-bark tar caused
DNA damage, but did not develop into permanent cytogenetic
damage—maybe even due to induced apoptosis. However, this
should not diminish the potential health risk of a genotoxin as
deficiency in mismatch repair of DNA damage can result in
increased mutations and lead to cancer (Abdel-Rahman, 2008).
The birch-bark tar used for this study has gone through a
process of aging lasting at least a few years and may have lost
the more volatile portions through evaporation while resinifica-
tion has made the tar/pitch rather solid. Due to a loss of volatile
substances the tar might have lost some of its genotoxic impact,
but this study clearly shows that birch-bark tar has the potential
to damage DNA of exposed cells and to induce apoptosis pos-
sibly due to the characteristic compounds found in the bark of
birch trees. The antimicrobial and apoptosis-inducing aspects
made the tar material a useful tool for specific preservative
and/or medical procedures.
We shall never know for sure whether the individual use of
birch-bark tar on skin or in the oral cavity had done more harm
than good. Our results indicate that birch-bark tar used by our
ancestors increased their genotoxic risk, but was also beneficial
to them due to the innate birch-bark compounds. Our review of
the ethnographic literature also indicates that it was, and may
still be, used for medicinal purposes by some peoples and pro-
vides a much more logical explanation for the tooth marks ob-
served in prehistoric birch-bark tar. Additional studies will be
necessary to further characterise the potential genotoxicity and
medicinal potential of birch-bark tar in greater detail.
The authors want to thank the former MSc students Ms Lin
“Betty” Liu and Mr. Khaled Lamin for their help in the labora-
tory. A number of grants did support this collaborative study:
Dr Randolph Donahue was awarded a grant of the University of
Bradford “Research Matters” for Palaeopharmacology. Dr Adrian
Evans’ research has been funded by NERC, UK (grant NER/S/
A/2004/12213). Ms Magdalena Sampol-Lopez was supported
by The DaVinci Scholarship provided by CAEB of the Balearic
Islands, Spain.
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