(Received for publication, March 13, 1997, and in revised form, May 22, 1997)
From the Genetisches Institut, Justus-Liebig-Universität, Heinrich-Buff-Ring 58-62, D35392 Giessen, Germany
The mouse lysozyme downstream enhancer was previously colocalized with the DNase I-hypersensitive site in the chromatin of mature macrophages. This hypersensitive site was shown to be macrophage differentiation-dependent. Demethylation of CpG sequences within the enhancer is correlated with lysozyme expression in mature macrophages. Binding of the GABP heterotetrameric transcription factor to the enhancer core element (MLDE), only seen in vivo on the demethylated MLDE element in macrophages, is inhibited by DNA methylation. Here, we analyzed the DNA sequences required for demethylation. In electrophoretic mobility shift experiments we found that in addition to the complete methylated MLDE the hemimethylated form of the lower strand inhibits GABP binding as well. Therefore, GABP is unlikely to be the mediator of demethylation. In addition, we show by stable DNA transfections of methylated mouse lysozyme enhancer sequences that MLDE-flanking sequences are required for demethylation. We narrowed down these DNA elements to two short regions of 163 and 79 base pairs on either side of the MLDE, each of which is sufficient to mediate demethylation of the GABP site.
The mouse genome contains two lysozyme genes, a Paneth cell
(P-lysozyme)1 and a
macrophage-specific (M-lysozyme) gene generated by a gene duplication
event (1). The M- and P-lysozyme genes are arranged in tandem with the
coding regions separated by 5 kb (Fig. 1). Analysis of the M-lysozyme
gene domain by DNase I digestion identified multiple hypersensitive
(HS) sites in the 5 and 3
M-lysozyme gene-flanking regions in
macrophage and myeloid precursor cell lines (Fig.
1) (2). Only a single site in the
3
-flanking region (HS3) was dependent on the differentiation state of
the cell line and correlated with M-lysozyme gene expression (2).
Transfection analysis of the flanking regions identified a single
enhancer downstream of the M-gene which overlapped the HS3 site and is limited to the subregion HS3.2 (2, 3). Analysis of the HS3 region
methylation state in M-lysozyme-expressing and nonexpressing cells
demonstrated a correlation between undermethylation of this region
with both the presence of the HS3 site and expression of the M-lysozyme
gene. Further fine mapping identified a central core enhancer (MLDE),
which is bound by a heterotetrameric GABP complex (4). We found that
GABP binding to the MLDE is methylation-sensitive (4). Thus, very
likely, macrophage-specific demethylation of the single CpG
dinucleotide within the MLDE is a mechanism to confer tissue-specific
enhancer activity.
In other systems, methylation of CpG dinucleotides has been correlated with transcriptional inactivity as well (for review, see Refs. 5 and 6). In two cases, DNA transfections have identified quite complex DNA regions required for tissue-specific demethylation (7, 8). Recent achievements in demethylating DNA in vitro (9) showed an involvement of RNA and that tissue-specific proteins are required for the specificity of the reaction.
Here, we wanted to analyze the mechanisms mediating macrophage-specific demethylation of the single CpG site within the mouse lysozyme enhancer core MLDE. We identified two short DNA regions of 163 and 79 bp which are required and sufficient for demethylation. The two fragments are overlapping, but the sequence in common is not sufficient for demethylation.
The pHS3/6(1-1166)-tkCAT construct contains a
1.17-kb Asp718I/PstI fragment of the mouse
lysozyme 3-enhancer region including the MLDE sequence in a
ptkCAT
H/N vector (NdeI/HindIII fragment deleted from pBL-CAT2, (10)), plus additional restriction sites from
the psk(
) polylinker of a subcloning vector. For psk(
)HS3.2 construction, an EcoRI/PstI fragment from pmLg219
(4) was subcloned into the EcoRI/PstI site of
psk(
). For pHS3.2-tkCAT construction, an
XhoI/BamHI fragment of the pskHS3.2 plasmid was
inserted into the SalI/BamHI site of
ptkCAT
H/N.
pMLDEtkCAT(HS3.2(121-171) (3)) was reconstructed because of a
sequencing error in the published oligonucleotide sequence. The
corrected bases are shown in bold letters:
5-CTATAGGTAGGCAGGAAGTAGAAGGTGGGACTTCCGGGAGGAGAGTGGAAC-3
. For pHS3.2(153-188) construction, the synthetic double-stranded oligonucleotide (5
-TCCGGGAGGAGAGTGGAACTCTGGGAGATAGTCAAG-3
) was inserted into the Klenow-filled SalI site of ptkCAT
H/N.
pHS3.2(82-219) and pHS3.2(146-219) were generated from pHS3.2-tkCAT
by exonuclease III deletions of the
SphI/EcoRV-digested vector. HS3.2(1-181) and
HS3.2(1-163) were constructed as follows. First, a modified pskHS3.2
(the SalI site 5
to the insert was deleted by digesting with XhoI/ClaI, blunt ending with Klenow and
religating) was digested with SalI plus PstI and
treated with exonuclease III to generate deletions. After religation
and verification of the deletions by DNA sequencing, the deleted
fragments were subcloned into ptkCAT
H/N using
HindIII/XbaI.
Mouse cell lines RAW264 (ATCC TIB71), M1 (ATCC TIB192), RMB-3 and J774-1.6 (11) were grown in Dulbecco's modified Eagle's medium (Life Technologies, Inc.) supplemented with 10% fetal bovine calf serum, 100 µg/ml streptomycin, and 100 µg/ml penicillin. EL4 cells (ATCC TIB39) were grown in RPMI 1640 (Life Technologies, Inc.) supplemented with 10% fetal bovine calf serum, 100 µg/ml streptomycin, and 100 µg/ml penicillin.
In Vitro MethylationMethylation was carried out in vitro on 40-50 µg of plasmid DNA by incubating with HpaII methylase (Fermentas 3 units/µg) overnight at 37 °C according to the supplier's instructions. The reaction was terminated by incubating for 20 min at 65 °C, extracting with phenol:chloroform (1:1), and precipitating with EtOH. The degree of methylation was tested by HpaII digestion.
Stable Transfection, DNA Isolation and DigestionFor the RAW264 cell line, 107 cells were resuspended in 400 µl of Dulbecco's modified Eagle's medium including 10 µg of reporter plasmid, 1.5 µg of pPUR (12) and electroporated at 300 volts and 900 microfarads with an Easyject Gene Pulser (Eurogenetec). The cells were replated in a 15-cm tissue culture dish and selected for adherent cells. After 2-3 days, the cells were selected for puromycin resistance, using 6.5-7 µg puromycin (ITC)/ml medium (60% fresh medium, 40% RAW-conditioned medium). Growing colonies were expanded as cell pools for DNA isolation. For the EL4 cell line, 0.75 × 107 cells were resuspended in 400 µl of RPMI including 20 µg of reporter plasmid, 3 µg of pPUR, and electroporated at 250 volts, and 900 microfarads. The cells were transferred to a 15-cm dish. Selection for puromycin-resistant cells was started after 48 h using 7 µg of puromycin/ml of RPMI medium. After an additional 24 h a fetal calf serum centrifugation2 was performed as follows to remove the dead cells. The cells were centrifuged (800 rpm/5 min), and the cell pellet was resuspended in 10 ml of phosphate-buffered saline, separated into two vials, and centrifuged again (800 rpm/5 min). The cell pellet was resuspended 1 ml of phosphate-buffered saline, overlaid on 5 ml of fetal calf serum from the same batch that was used for the culture medium, and centrifuged again. The pellet with live cells was resuspended in 10 ml of RPMI including 7 µg/ml RPMI (60% fresh RPMI, 40% EL4 conditioned medium). The two vials were combined and transferred to a 15-cm cell culture dish. Growing colonies were expanded as cell pools for DNA extraction. The genomic DNA isolation was performed as described previously (13) with the following modifications. The DNA was precipitated with 1 volume of isopropyl alcohol, removed with a flame sealed Pasteur pipette, rinsed in 70% EtOH, air dried, and disssolved in TE buffer (TE: 10 mM Tris-HCl, 1 mM EDTA, pH 7.6). About 30-50 µg of genomic DNA was digested twice with 8-10 units/µg of an appropriate restriction enzyme for 5 h and split into three aliquots, for MspI digestion, HpaII digestion, and control, respectively. One µg from each of the three aliquots was used for LM-PCR.
Identification of Positive Clones and LM-PCR AssayOne µl
of genomic DNA (0.5-2 µg of DNA) was PCR amplified with
plasmid-specific primer pairs (C-TK with an annealing temperature of
64 °C or Forw25-P3 at 58 °C; 30 cycles), separated on an agarose gel, and visualized after staining. For tkCAT constructs
C
(5
-TGGCGGGTGTCGGGGCTGGC-3
) and TK (5
-GCCCCGACTGCATCTGCGTG-3
)
primers were used. Integration of pskHS3.2 was checked with Forw25
(5
-CGCCAGGGTTTTCCCAGTCACGACG-3
) in combination with P3
(5
-CGAGCTTCTTTCTCTGCATCCCTTCATCCGC-3
). First strand DNA synthesis and
ligation were performed according to Ref. 14. The primers
C16 (5
-
AGCAGACAAGCCCGTC-3
, 52 °C) for the tk constructs or For19
(5
-CTCGAGGTCGACGGTATCG-3
, 50 °C) for psk constructs were used.
Ligation was performed using the annealed linker (L23
(5
-GGTGACCCGGGAGATCTGAATTC-3
), L2 (5
-GAATTCAGATC-3
)). Amplification
and end labeling were performed as described by Ausubel et al.
(37) on a Perkin-Elmer thermal cycler. For amplification the
primer sets L23/
C18 (5
-TGTTGGCGGGTGTCGGGG-3
) for tk constructs or
L23/Forw.19 for pskHS3.2 were used at annealing temperatures of
64 °C and 50 °C, respectively. End labeling was performed at an
annealing temperature as indicated with 32P end-labeled
C38 (5
-TGGCTTAACTATGCGGCATCAGAGCAGATTGTACTGAG-3
, 70 °C) or
S5 (5
-GTCCACCGCCTTCTGATTGGTCTGATAAAGAGCTG- 3
, 68 °C) depending on
the plasmid constructs. The samples were electrophoresed on a 4-6%
polyacrylamide, 8 M urea gel and autoradiographed. The size
of the fragments was verified by control digestion and
32P-labeled DNA marker.
Nuclear protein extracts from RAW264 and M1 cells were
prepared as described by Nickel et al. (4). Genomic cytosine
sequencing and DNase I footprinting were performed as described (13).
For generation of hemimethylated MLDE probes, 100 pmol of
oligonucleotide, synthesized with a single 5-methylcytosine CpG site,
was annealed with the same amount of unmethylated complementary strand
oligonucleotide. To generate methylated MLDE, complementary
oligonucleotides were synthesized with 5-methylcytosine CpG sites and
annealed. 30 pmol of the double-stranded oligonucleotides were labeled
with [-32P]ATP using T4 polynucleotide kinase
according to the supplier's instructions (New England Biolabs).
Labeled oligonucleotides were separated on a 5% polyacrylamide gel.
After electrophoresis the gel was autoradiographed, and the band was
excised. The DNA was eluted and used for electrophoretic mobility shift
assay experiments, which were performed as described (4) with the
modification that each sample contained 37 µg of yeast-RNA
(Amersham).
Previously we have determined that methylation of the
single CpG within the enhancer core MLDE is sufficient to regulate
binding of heterotetrameric GABP (4). Therefore a possible mechanism for demethylation of this site might be that after DNA replication the
GABP complex would be able to bind to the hemimethylated DNA. Thereby,
the activity of the maintenance methylase could be inhibited, generating a fully demethylated DNA after a second round of
replication. This type of competitive inhibition of the maintenance
methylase has been suggested for the transcription factor Sp1 (15).
Such a mechanism would require that the transcription factor GABP can bind at least to the hemimethylated DNA. Here, we tested this possibility by competing GABP binding with the double-stranded MLDE
oligonucleotide with a single 5-methylcytosine CpG site in either the
upper (sense) strand or the lower (antisense) strand. Electrophoretic
mobility shift assays were performed with nuclear extract from RAW
macrophage cells as well as from M1 myeloblasts (Fig.
2). Both extracts generated several
retarded complexes, with the upper complex (complex a, Fig. 2)
consisting of the GABP heterotetramer bound to the palindromic DNA,
whereas complex b is generated by an unknown protein (4). As expected,
methylation of both strands of the single CpG site inhibits GABP
binding and therefore is inactive in competition, whereas unmethylated
DNA competes efficiently. Using the sense and antisense hemimethylated MLDE oligonucleotide as competitor, a strand-specific effect is seen.
Lower strand (antisense) methylation inhibits GABP binding and is
therefore ineffective in competition similar to the effect seen with
the competitor methylated on both strands. In contrast, upper strand
(sense) methylation shows almost no interference in GABP binding and is
therefore an efficient competitor (Fig. 2).
Thus, an indirect demethylation by binding of GABP to the hemimethylated DNA after replication is unlikely since such a binding would only be possible for the hemimethylated upper strand. Such a mechanism might be conferred by another protein (complex) which is able to bind the methylated DNA (complex b, Fig. 2). To test this possibility or to identify other mechanisms for demethylation, we designed additional experimental strategies.
All of the CpG Dinucleotides within the Mouse MLDE Region Are Specifically Demethylated in MacrophagesThe mouse M-lysozyme
downstream enhancer contains several CpG dinucleotides that are
potential targets for methylation. It has been shown previously that
one of these is located within the enhancer core element MLDE and is
specifically demethylated in macrophage cells (13). In contrast to
macrophages, immature macrophages and T lymphocytes contain a
5-methylcytosine at this site on both strands. Here we extended this
analysis on the entire HS3.2 region mediating full enhancer activity in
DNA transfections (2, 3). Detection of methylated CpG dinucleotides was
performed with cytosine sequencing in combination with LM-PCR (see
"Experimental Procedures"). The genomic DNA was isolated from the
following three mouse cell lines: mature macrophages (J774-1.6),
immature macrophages (RMB-3), and T lymphocytes (EL4). Five CpG
sequences are found within the HS3.2 region, sites 1, 2, 4, and 5 are
shown in Fig. 3, and site 3 has been
described previously (13). All of the sites follow the methylation and
demethylation pattern seen for site 3 (13); they are methylated in the
T cell line and in the immature macrophages (Fig. 3). In contrast, the
mature macrophage line J774-1.6 displays all of these sites in the
cytosine sequencing reaction, indicating that all five sites are
demethylated (Fig. 3). We have previously published that the central
part of the HS3.2 region containing CpG site 3 is bound in
vitro by GABP. This in vitro footprint was identical
for protein extracts from macrophage cells or from non-macrophage cells
(13). Therefore, we analyzed the flanking sequences in an in
vitro footprint reaction as well (Fig. 3C) and found
additional sequences protected. Again, no major difference was seen for
the EL4 and the J774 extract.
Cis-elements for Demethylation
To analyze the mechanisms
involved in the macrophage-specific demethylation of the lysozyme
downstream enhancer, we wanted to identify the DNA sequences required
for the removal of methyl groups from the CpG dinucleotide within the
enhancer core element. For these experiments we choose different DNA
fragments from the downstream region harboring the chromatin-DNase I
hypersensitive sites HS3-HS6 (see Fig. 1). This stretch of DNA
sequence contains several CpG dinucleotides (see above), only one of
which is part of the recognition sequence for the restriction enzyme
HpaII (CCGG). This sequence is located in the center of the
downstream enhancer within the binding site of the heterotetrameric
transcription factor GABP (4). Therefore, this sequence could be easily
methylated in vitro with the help of the HpaII
DNA methylase. Such a methylated DNA was transfected into recipient
cell lines, and stable transfectants were identified by their
resistance to puromycin. Position effects on the methylation pattern of
transgenes have been reported (16); therefore, we wanted to minimize
the effects of particular integration sites by pooling about 50 colonies to analyze the methylation of the transfected DNA fragments.
Such a strategy has been used successfully for the light chain
enhancer (8). In a first series of experiments we tested a long (1.2 kb) fragment from the downstream region HS3/6 (see Fig. 1). We wanted
to know whether such a DNA element after in vitro
methylation could be found demethylated when integrated into the genome
of a macrophage cell line. After transfection into RAW cells and
subsequent isolation of the DNA from pooled cell clones, the genomic
DNA was digested either with the restriction enzyme HpaII or
with the restriction enzyme MspI. MspI digestion
serves as a control, since the enzyme cuts both methylated DNA and
unmethylated DNA. In contrast, digestion with HpaII is only
possible with unmethylated DNA. To avoid difficulties in
restriction enzyme digestion caused by the viscosity of the large
genomic DNA, we routinely used an additional restriction enzyme to
digest the DNA into larger fragments (NdeI,
BamHI, PstI; see restriction site map in Fig.
4). In addition, the amount of the large
fragment generated by the first restriction enzyme represents the
undigested (i.e. methylated) material after HpaII
digestion. Thus, the efficiency of demethylation can be judged from the
ratio of intensities of the "smaller" fragment
(HpaII-digested) relative to the "larger" fragment
(HpaII-resistant). In contrast, the relative intensities of
bands generated in different digestions (different lanes) are of no
relevance. The fragments were visualized by LM-PCR with primers
specific for the transfected DNA. Cell pools generated with the
methylated HS3/6 fragment always showed an almost complete digestion
with the enzyme HpaII (Fig. 4), indicating a successful demethylation of the DNA. At this point we were worried about a
possible artifact, since Weiss et al. (9) showed that
nonspecific demethylation activity is present during the genomic DNA
isolation procedure even after proteinase K digestion. On the other
hand, these authors showed that EDTA abolished this activity
completely. We tested this possibility and found that our extraction
procedures in the presence of 5 mM EDTA did not allow
additional nonspecific in vitro demethylation (data not
shown). In addition, as seen below, depending on the DNA sequence
tested, we found the methylated CpG being preserved after transfection
and DNA isolation.
Next we focused just on the 200-bp enhancer fragment HS3.2 and found
that this short fragment was similarly demethylated (Fig. 4). This was
surprising since a similar analysis of the demethylation within the
intronic chain enhancer showed that more than 1 kb of DNA was
required for successful demethylation in lymphocytes (8).
Similarly, in transient transfections of the methylated
-actin
promoter about 800 bp of DNA was required to demonstrate demethylation
in myelocytes (7). We wondered whether the transcriptional activity of
the transfected reporter gene may cause or influence the
demethylation or whether the observed demethylation on the short
HS3.2 fragment would be seen in the context of a different vector as
well, which does not show transcriptional activity of its own in
eukaryotic cells. Therefore we introduced the HS3.2 fragment into the
prokaryotic Bluescript vector psk. The demethylation analysis of the
pooled transfection clones showed a demethylation similar to that
observed in conjunction with the tkCAT reporter (Fig. 4). Therefore, we
were assured that the demethylation observed is not the consequence of
transcriptional activity in its vicinity, but rather that
demethylation is caused by the HS3.2 fragment itself.
Previously we have shown (13) that demethylation of the endogenous
downstream enhancer is restricted to macrophage cells. T lymphocytes
that do not express the lysozyme gene show a methylation of the CpG
site within the downstream enhancer HpaII sequence. Therefore, we wanted to know whether the transfected methylated DNA
shows this preference for demethylation in macrophages as well. We
transfected methylated HS3.2 DNA as well as the methylated GABP-binding
fragment (MLDE) into a T cell line (EL4). Analysis of the transgene
showed that the majority of HS3.2 fragments was resistant to
HpaII digestion and therefore has maintained the methylation
at this site (Fig. 4C). Similarly, propagation of the methyl
group was seen for the transfected MLDE sequence as well (Fig.
4C). This tissue specificity of the HS3.2 fragment encouraged us to use even shorter enhancer fragments to delineate the
DNA elements sufficient for demethylation in macrophages. From the
HS3.2 region we generated four overlapping fragments that were
methylated by HpaII methylase and transfected into RAW cells
similar to the previous experiments. Surprisingly, all of these
subfragments showed a demethylation for the majority of the
molecules (Fig. 5). Even the shortest
fragment (HS3.2(146-224)), only 78 bp in length, was
HpaII-digestible. The other extreme, fragment HS3.2(1-163),
showing only an overlap with the previous fragment of 17 bp, was
digestible by HpaII as well. Such a result may be explained
by two different mechanisms. Either the short region just overlapping
between the two fragments HS3.2(146-224) and HS3.2(1-163) would
contain all of the information required to direct the demethylation
activity to this CpG dinucleotide, or redundant cis-elements for
demethylation occur in the enhancer, one in HS3.2(146-224) and one in
HS3.2(1-163). To distinguish between these two possibilities we
transfected the MLDE (4), which contains a little more than the
overlapping sequence (bp 120-171), into RAW cells (Fig.
6). The transgene fragment was not
digestible at all with HpaII, in contrast to the complete digestion with the control enzyme (MspI) and in contrast to
the unmethylated transfection analyzed by HpaII digestion
(Fig. 6). Similarly, we tested the demethylation of the fragment
HS3.2(153-188). In addition to the methylation site, this fragment
contains a footprint region seen in vivo in macrophage cells
(footprint 4).3 Methylation
of this fragment shows a result similar to that seen with the MLDE
fragment; the methyl group is maintained in the transgene fragment
(Fig. 6). Other attempts to identify the nucleotide sequences required
for demethylation more precisely resulted in variable degrees of
demethylation.
Thus we can conclude that both of the fragments HS3.2(146-224) and
HS3.2(1-163) contain all of the cis-elements required for demethylation (Fig. 7) and that these
elements very likely are redundant within the HS3/6 downstream enhancer
region of the mouse M-lysozyme gene.
Many examples have been identified linking sequence-specific DNA
demethylation with differentiation of a particular tissue or cell type
(for review, see Refs. 5 and 6). In these cases, it has been shown that
the differentiation-dependent expression of a
tissue-specific gene is correlated with the demethylation of flanking
sequences. Such a demethylation is usually restricted to a specific
region or at least to specific genes. For example, expression-linked
demethylation has been seen for the genes coding for chicken
vitellogenin, human dihydrofolate reductase, mouse collagen IV, rat
-actin, mouse
chain, human estrogen receptor, human galectin-1,
or mouse pyruvate dehydrogenase E1
subunit, to name only a few (7,
8, 17-22). Other genes, not expressed in this particular tissue,
remain methylated. Such a correlation has been found for the mouse
M-lysozyme gene as well. The M-lysozyme gene is inactive in
non-macrophage cells and shows a methylated CpG dinucleotide within the
single HpaII recognition sequence of the downstream enhancer
(3). It has been shown that during the differentiation of the
multipotent FTCP-A4 cell line toward macrophages the enhancer loses its
methyl groups (3). Similarly, the human lysozyme gene has been found to
be demethylated depending on the differentiation as seen in
ex vivo cultures of hematopoetic progenitor cells, whereas
the gene coding for myeloperoxidase showed unaltered demethylation
(23). Other myeloid-specific demethylation events have been shown as
well to correlate with transcriptional activity, such as the
c-fms gene and the regulatory region of the tumor necrosis
factor-
gene (24, 25).
Many of the published examples could not distinguish between a role for
the demethylation being required for transcriptional activity or a
possible demethylation-inducing function mediated by transcription. In
some cases, methylation-dependent repressor binding or
inhibition of transcription factor binding could be demonstrated (5,
26-29). For the mouse lysozyme gene, it was shown that within the core
part of the downstream enhancer the heterotetrameric transcription
factor GABP is required for full enhancer activity (4). In addition, it
was demonstrated that the methylation of a single CpG dinucleotide
within the enhancer core region inhibits in vitro DNA
binding of GABP (4). A similar sensitivity in methylated DNA binding
was demonstrated for GABP and other ETS proteins in the context of
binding sites that differ from the mouse lysozyme GABP binding site
(30-32). This suggests that demethylation of this enhancer is a
prerequisite for enhancer activity and therefore, for transcription.
Here we show that the cis-element-dependent demethylation
can be observed independent of the type of the neighboring DNA or
promoter context. The 224-bp fragment HS3.2 can be stably transfected
either fused to a eukaryotic reporter gene or to a prokaryotic vector
and will be demethylated in both cases. Therefore, the lysozyme
downstream enhancer confers at least two functions. One function is to
mediate the demethylation during differentiation, and another function
is to activate gene transcription by the bound enhancer factors. A
similar dual activity has been demonstrated for the -actin promoter
and the
enhancer (7, 8).
Our transfection data suggest that the MLDE fragment within the downstream enhancer is not sufficient to mediate demethylation. This fragment harbors the single HpaII site and binds the heterotetrameric GABP factor in the absence of methylation. Here we have shown that even a single methyl group on the lower strand is sufficient to inhibit GABP binding, whereas the hemimethylated upper strand does not interfere with binding. Since GABP binding is impaired even by hemimethylated DNA, this factor cannot be the cause for demethylation. This is in contrast to the mechanism observed in the context of Sp1 binding (15). Sp1 is able to bind methylated DNA and after replication prevents the maintenance methylase from modifying the newly synthesized DNA strand. Such a Sp1-like activity could have been envisioned to be utilized by an unknown factor that has been found to bind to the GABP response element even in the case of a fully methylated DNA (4). If this factor would indeed play such a role, this function would not be sufficient for demethylation, since we have shown that the methylated MLDE fragment is not demethylated (Fig. 6).
Within the group of fragments mediating demethylation, there seems to be a bias in demethylation efficiency: all of the fragments extending up to the position 224 (Fig. 7; i.e. fragments HS3/6, HS3.2, 82-224, 146-224) mediate 90-100% demethylation. In contrast, fragments with a downstream deletion (Fig. 7; i.e. fragments 1-181, 1-163) mediate demethylation for only 70-80% of the molecules. Nevertheless, two different sets of fragments, overlapping in the GABP site only, confer demethylation. Computer analysis of the sequences flanking the GABP site did not reveal any consensus in common, which otherwise might have been an indication for trans-acting proteins required for demethylation. In addition, in vitro footprinting showed only one protected region in addition to the GABP site (Fig. 3C).
One of the minimal regions required and sufficient for
demethylation is only 78 bp in length (HS3.2(146-224)) (Fig. 7).
None of the sequences analyzed for conferring demethylation in the other model systems could be delineated to such a small fragment. For
both the -actin promoter and the
enhancer, DNA fragments of 800 bp to more than 1000 bp in length were required for demethylation (7,
8). Despite the complexity of the
enhancer it could be shown that
the absence of a single enhancer factor (nuclear factor-
B) leads to
the failure of this fragment to undergo selective demethylation (33).
Previous results on in vitro demethylation of hemimethylated
DNA by chicken embryonic extracts suggested a glycosylase activity (34,
35). Ras-induced overall demethylation in mouse embryonal P19 cells
could be followed in vitro as well (36). In contrast to
embryonic cells and tissues, demethylation specific for differentiation
and for particular sites may utilize a different mechanism. Recent
achievements with in vitro demethylation showed that RNA is
involved in sequence-specific demethylation (9). Nevertheless,
these authors showed that the tissue specificity of the demethylation
reaction involves proteins as well.
Taken together, these and our results demonstrate that the demethylation activity is mediated via enhancer elements and may be organized in a manner similar to that of the enhancer elements; that is, specific modules within the enhancer regions are either sufficient by themselves to mediate demethylation or have to act in combination with other modules.
We thank M. Cross for technical hints in macrophage transfections, H. Wahn for excellent technical assistance, and A. Baniahmad for critically reading the manuscript.