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The nucleus of the cell contains our genetic material, which must be tightly and
neatly packaged into an area roughly one-tenth the size of the cell (1), and yet able to be
accessed for replication, transcription, and repair. The observation of this central
structure common to most cell types was made in the early 1830’s, described by Robert
Brown as “this areola, or nucleus of the cell as perhaps it might be termed…is more or
less distinctly granular…There is no regularity as to its place in the cell; it is not
unfrequently, however, central or nearly so.” (2). Microscopic details of cellular and
nuclear morphology quickly became an interest to many biologists, and concepts of our
modern day cell theory were formally established.
Although spontaneous generation and free cell formation were originally believed
to be the source of new cells (3, 4), emerging evidence suggested fission of one cell to
create two new cells was the correct concept (5). To decipher between the theories of de
novo cell formation and cellular division, German biologist Walther Flemming developed
a variety of fixatives and stains to visualize resting and dividing salamander cells under a
microscope, which lead to his description of “nuclear multiplication involving
metamorphosis of the nuclear mass” (6), now known as mitosis (Figure 1). This dividing
mass localized only to the nucleus of the cell, and was stainable by Flemming’s dyes,
thus termed “chromatin”. Flemming also observed that chromatin could condense and
form thread-like structures termed chromosomes, which were then passed onto daughter
cells (7), disproving the idea of free cell formation, and laying the basis for the field of
nuclear biology.
In addition to histological characterization of the cell and nucleus, German
biochemist Albrecht Kossel initiated studies to identify the chemical composition of the
nucleus using avian erythrocytes, which presented “very favourable conditions for the
chemical investigation” (8). Nuclear extraction using acid and salt yielded a precipitate
of a basic compound, which Kossel inexplicably named “histone”. Histone was also
noted to purify with nucleic acids, and this complex was collectively known as
nucleoprotein (9). Further studies of histones were mostly carried out in cells with a
large nuclear to cytoplasmic ratio, such as sperm cells, and histones identified in nuclei of
sea urchin gonads were initially thought to be a type of hormone involved in sexual
reproduction and cell division (10). Later, calf thymus provided another tissue with a
suitable nuclear mass to extract histones, but most other types of animal and plant tissue
originally yielded no measurable amount of a basic nuclear protein component (11).
Further investigation and improved fractionation and chromatographic techniques,
however, demonstrated that most types of somatic cells contained histones (11-13),
and these basic proteins were a main constituent of chromosomes, similar to the
Figure 1. Schematic of mitosis. After the genetic material is replicated in S-phase, the
cell divides during mitosis (M-phase). The M-phase of the cell cycle consists of 4 major
stages: prophase, condensing of replicated chromosomes; metaphase, alignment of
chromosomes at the center of the mitotic spindle; anaphase, separation of sister
chromosomes toward opposite spindle poles; and telophase, chromosomes decondense
nuclear envelope reassembles. Cytokinesis, the division of cytoplasm to create two new
daughter cells, occurs simultaneously during the last stages of M-phase. Images were
modified from (7).
nucleoprotein complex Kossel had described almost 60 years previously. With the
discovery that inheritable information was encoded in nucleic acids (DNA) contained
within the nucleus (14), the significance of understanding why histones were the main
constituents of chromosomes heightened (15).
Originally, the basic histone compound was thought to contain a single protein,
but fractionation using gradient column chromatography lead to the identification and
classification of 5 major histone groups (16-19). The original nomenclature for each
fraction was based on the amino acid composition and point of elution from the column,
and they included the very-lysine rich F1 fraction, the intermediate fraction F2, which
later was resolved into 3 separate fractions F2a1, F2a2, and F2b, and the arginine-rich
fraction F3 (Figure 2) (20). These original fractions were subjected to much more
rigorous extraction procedures to further separate out what was thought to be a highly
complex mixture of proteins within each fraction. The general school of thought
suggested that such a high abundance of histone protein in the nucleus must be the result
of a multitude of unique proteins (20). However, only limited separation of the already
established fractions was possible. Electrophoresis of whole histone extracts and
individual histone fractions in polyacrylamide gels (21) further resolved the homogeneity
of each fraction. Fraction F2a1 (Histone H4, see Figure 2) from calf thymus was the first
histone protein to have its complete amino acid sequence determined (22). Following
identification of individual amino acid sequences in the other 4 major fractions (23-27),
the evidence demonstrated that only 5 distinct histones (Histone H1, H2A, H2B, H3, and
H4) were present in the majority of eukaryotic tissues studied. Sequencing studies also
revealed a strong evolutionary conservation of amino acids from calf thymus to pea
Calf Thymus
NaCl & EtOH Wash
EtOH & HCl
Weight (kDa): 17
Figure 2. Original purification scheme for histone proteins. Five total fractions of basic
protein were purified from calf thymus and characterized based on amino acid content.
The original fraction name, current nomenclature (labeled as Histone), and molecular
weight are shown for each fraction.
pod (22, 28, 29). Later, the discovery of histones in yeast (30) and fungi (31, 32)
suggested that histones were a vital component of nuclear structure and function in many
forms of life.
Heterochromatin and Euchromatin
In earlier observations of the physical state of the nucleus, Walther Flemming
described condensation of chromatin during nuclear division, which returned to its
diffuse resting state after cellular division (Figure 1). The change from diffuse to dense
states of chromatin required a cell to proceed through phases of mitosis, but observations
in additional types of resting and dividing cells suggested that condensed chromatin did
not always return to a more diffuse state. In studies performed in the 1920’s, E. Heitz
observed that after the final phase of mitosis, a portion of the condensed chromatin of
moss cells remained coiled within the nucleus during interphase, which was termed
“heterochromatin”, while the rest of the diffuse chromatin within the resting nucleus was
termed “euchromatin” (33). Higher magnification of calf thymus nuclei using electron
microscopy (EM) revealed similar dense and diffuse areas of chromatin (34), suggesting
this was a common state of the nucleus in multiple types of cells. The function of
heterochromatin in a non-dividing cell was a topic of study to both geneticists and
chemists alike, and complementary findings in both fields helped to define the role of
histones in chromatin regulation.
Genetic Studies
Resting chromosomes were studied in simple biological systems, such as the
mealy bug, an insect related to the aphid. In the mealy bug, 5 chromosomes existing in a
diploid state had been characterized. Only in the females do all 10 chromosomes exist as
euchromatin, while in the males, one set of chromosomes exist as heterochomatin. A
euchromatin haploid gene set is passed on to offspring from both male and female, but
only in male offspring does the paternal set become heterochromatin and genetically inert
(Figure 3A), as demonstrated through informative irradiation experiments of either
parental male or female insects (35). When parental females were irradiated, the male
offspring had chromosomal aberrations only in the euchromatin, while irradiation of
parental males resulted in aberrations in heterochromatic regions of male offspring. To
analyze the function of heterochromatin, male parents were given increasing doses of
irradiation to create genetic mutations, and the ratio of male to female progeny was
scored. With higher doses of irradiation, female progeny numbers drastically declined,
while numbers of male progeny remained similar to non-irradiated control experiments.
These data lead to the conclusion that mutated genetic material passed on to the female
offspring was genetically active and lethal, but male progeny survived because the
paternal chromosomes became heterochromatic and genetically inert (Figure 3B).
Similar observations of heterochromatic inactivation had also been made in studies of the
second copy of the X chromosome in female mice (36, 37).
Chemical Studies
Because heterochromatin was genetically inert, and histones were a main
component of chromatin, it was hypothesized that chemical interactions between basic
histones and nucleic acids were necessary for chromatin regulation. Previous hypotheses
Figure 3. Genetics of heterochromatin and euchromatin in the mealy bug. A. Female
chromosomes (in red) are passed on and remain as euchromatin in both male and female
offspring, while male chromosomes (black) remain as euchromatin in females but
become heterochromatin in male offspring. B. Irradiation of male parents to induce
lethal mutation (gray stars) results in damaged DNA being passed on to both offspring.
The father’s DNA remains as euchromatin in female offspring, and expression of the
lethal mutations cause death, whereas male offspring silence the damaged DNA,
preventing death.
regarding the physiological role of histones suggested that they may function to inhibit
gene expression and regulate cellular differentiation (13, 38). The content of histone
protein was analyzed from fractions of calf thymocyte interphase nuclei, collected by
gradient centrifugation to separate dense or diffuse chromatin. A higher abundance of
histone in the heterochromatin was believed to be the mechanism of keeping portions of
the genetic material silenced, but interestingly, the histone content between
heterochromatin and euchromatin was no different in interphase calf thymocytes (39).
Similarly, histone extracts from both female and male mealy bugs were analyzed by gel
electorphoresis, and no differences in histone content or mobility were identified (40).
Although the levels of histones were unchanged between dense and diffuse
chromatin areas, histones did have a role in the structural aspect of keeping the
heterochromatin condensed. Proteolytic digestion of nuclei with trypsin converted the
mass of dense chromatin to a loose network of fibrous material, as trypsin specifically
cleaves after lysine and arginine residues, which populate histone amino terminal
sequences (41). Although trypsin could target other nuclear proteins, the disruption of
heterochromatin was believed to be the result of structural imbalances in the absence of
histone. Complete depletion of arginine-rich histones from calf thymus nuclei using
concentrated ethanol (EtOH) solution had little effect on heterochromatin structure, but
when lysine-rich histones were extracted with citric acid solution, the resulting
heterochromatin resembled that which had been digested with trypsin. Reconstitution of
the citric acid-extracted nuclei with one or two times the amount of lysine-rich histones,
but not arginine-rich histones, yielded a re-condensed form of chromatin (34). The
lysine-rich histone fraction, or histone H1, seemed to crosslink the DNA strands together,
demonstrating the importance of histones in chromatin structure, although how histones
actually distributed themselves among the genetic material remained in question.
A Repeating Unit
The double helical structure of DNA was discovered through X-ray diffraction,
and this same technique was used to gain an understanding of the structure of the DNAhistone complex, or nucleohistone. Chromatin fibers were purified and gently stretched
out and allowed to relax under multiple conditions before obtaining an X-ray diffraction
pattern. A single diffraction pattern was consistently seen from long fibers, over the
course of at least 5 separate experiments, suggesting chromatin fibers consisted of a
common repetitive structure. The fiber was roughly 100Å in diameter, and the DNA
diffraction suggested in addition to the normal double helical structure, the DNA
molecule was bending or folding in an additional coiled-coil motif, or superhelix
conformation (42). The data also demonstrated that the lysine-rich histone fraction is
unnecessary for forming the same diffraction pattern as seen with whole nucleohistone
fibers (43). Using improved techniques to visualize a more precise chromatin structure,
chromatin fibers were depleted of lysine-rich histones by salt extraction or trypsin
digestion and closely examined using EM (44). Purified chromatin fibers, regardless of
the species or tissue of origin, resembled long, flexible chains containing spherical
particles spaced rather evenly along the fiber, like “beads on a string” (45). The diameter
of each particle, or nucleosome, measured under the microscope correlated with the
diameter of nucleohistone measured by X-ray diffraction. Linkage of the nucleosomes
was established by a 10-15Å thick fibril, similar to the diameter of a naked strand of
DNA. Direct lysis of nuclei on EM grids allowed for visualization of native chromatin
structure, containing all 5 histone proteins, and the conformation was similar, although
much more compact (44). The identification of repeating nucleosome units on strands of
DNA did not, however, reconcile how histones actually interacted with DNA or with
each other.
In solution, total histone tended to form aggregates due to harsh denaturing
protocols, but milder extraction procedures prevented histone aggregation, and histones
could be analyzed by gel filtration chromatography. In this assay, two protein peaks
eluted from the column, with the first containing histone H1, but also histone H3 and
histone H4, whose molecular weights were each less than that of histone H1 (see Figure
2). Histones H2A and H2B eluted in a second fraction, even though their molecular
weights were more closely related to H3 and H4, which suggested H3 and H4 formed a
high affinity dimer. Cross-linkage of histone protein mixtures through their amino
groups established that histones H3 and H4 not only formed a heterodimer, but could
homodimerize, as well as form a tetrameric complex of (H3)2(H4)2. Histone H1 was not
able to form dimers with itself, or any of the other histones, confirming its 0.5:1 molar
ratio to the other histones (46), and lack of necessity as a structural component of
individual nucleosomes. Histones H2A and H2B more commonly formed heterodimers
(H2A, H2B), and cross-linking an equimolar ratio of H2A, H2B, H3, and H4 together
only yielded oligomers of (H2A, H2B), (H3, H4), and (H3)2(H4)2 (45, 47).
Reconstitution of a nucleosome in vitro using adenovirus-2 DNA and purified individual
histones from multiple species required 4 of the 5 histones (minus histone H1). The same
experiment, when repeated with 4 histones and λ phage DNA which does not associate in
vivo with histones, could form nucleosomes in vitro as well, indicating there was not a
sequence specific recognition requirement in DNA for core histone binding (44). The
definitive structure of the nucleosome came after multiple attempts at crystallization of
the DNA-histone complex, initially being resolved at 7Å (48), 3.1Å (49), and 2.8Å (50)
by X-ray crystallography. At 2.8Å, DNA remained in the B conformation, and 146 bp of
the double helix wrapped in a superhelical conformation around a central octamer of
histones, 2 each of H2A, H2B, H3, and H4. Increasing the length of DNA from 146 bp
to 147 bp drastically improved crystal formation and yielded an enhanced 1.9Å structure
Figure 4. Nucleosome structure. DNA (modeled in purple and gray strands) is wrapped
roughly 1 ¾ times around the octamer of histones H2A (green), H2B (orange), H3 (red),
and H4 (blue), represented by ribbon diagrams. The N-terminal tails of the histones are
highly unstructured and protrude from the nucleosome core between the DNA double
helix strands. (PDB 1KX5)
(51), with further resolution of the unstructured N-terminal histone tails (Figure 4). Each
histone within the protein octamer interacted specifically with the DNA phosphate
backbone to keep such a compact and coiled shape within the nucleus, and each
nucleosome formed a further condensed quarternary structure by being linked through
small stretches of DNA and the single linker histone H1.
Histone Modifications
Resolution of the nucleosome structure defined the placement of histones within
the proteinaceous core and established the contact points between DNA bases and histone
residues, which kept the DNA restrained in the superhelical conformation through salt
linkages and hydrogen bonds (50). The histones themselves formed a conserved
secondary structure termed the histone-fold motif, consisting of a general helix-loophelix-loop-helix domain (Figure 5) (49, 50). The strong tertiary structure in the histone
core created by anti-parallel histone-fold binding of dimers, and hydrogen bonding
association (through the phosphate backbone) of 121 of 146 bp of the DNA helix to the
protein core, resulted in a tight binding complex. This structure limited access to the
nucleosome for essential processes such as transcription and replication. A point of
regulation existed in the unstructured random-coil N-terminal histone regions, which
extended from the proteinaceous core as tails, and could be modified to essentially
remodel the higher-order chromatin structure.
N-terminal Histone Tails
The N-terminal regions of histone proteins are highly enriched for basic residues
Amino Acid Residue
Figure 5. The histone fold motif. The core histone proteins have a conserved secondary
structure consisting of a helix-loop-helix-loop-helix motif required for their tight
association to homo- and heterodimerization. The helix domain is represented by the
helical structure for each histone at the approximate amino acid residue. The dotted line
represents the region of each N-terminal tail which could not be resolved through X-ray
crystallography. Data modified from (49) and (50).
such as lysine and arginine, and are the least likely regions to form secondary structures,
based on NMR conformation studies (52). These regions are also much more susceptible
to low levels of trypsin digestion compared to the interacting regions of histone in the
protein core, and cleavage of 20-40 N-terminal amino acids resulted in greater access to
staphylococcal nuclease digestion of DNA in extracted chromatin preparations
(53). These random-coil N-terminal tails of histones associated with DNA in a
protective manner, and were required for forming the higher order chromatin structure.
When trypsinized (H3)2(H4)2 tetramers were mixed with complete (H2A)(H2B) dimers
(and to a lesser extent, trypsinized (H2A)(H2B) dimers mixed with complete (H3)2(H4)2
tetramers), only the lower order “beads on a string” conformation could form, similar to
chromatin depleted of histone H1 linker (54, 55). The N-terminal tails were the only
portions of the histone octamer to be functional on the outside face of the DNA helix,
passing through channels formed by the minor grooves of DNA (50, 51). This allows for
the histone tails to interact with chromatin remodeling enzymes, which in turn leads to
the recruitment of transcription, repair, and replication machinery. Cross-talk between
histone tails and cellular machinery is a vital process, and one of the first lines of
regulation occurs by covalent modification of lysine, serine, arginine, and threonine
residues located within the first 20-40 amino acids of histone tails.
The Histone Code
The idea that histones were involved in the regulation of DNA function was first
introduced in the 1950’s (13), and experimental proof demonstrated that while naked
DNA was genetically active and capable of supporting RNA synthesis, addition of
histones to DNA in a concentration-dependent manner prevented this DNA-dependent
RNA synthesis (56). But in heterochromatin, which is genetically inactive, there is no
difference in the ratio of DNA to histone compared to the euchromatin DNA-histone
ratios (39, 40). Acid extraction of histones from nuclei could enhance RNA polymerase
function as well (41), but weakening of electrostatic bonds between the histone proteins
and DNA through high acid concentrations was unlikely to occur physiologically. So
how did histones, then, know when and where to control genetic regulation? Under
physiological conditions, the basic residues of histone N-terminal tails served as
acceptors of a number of post-translational modifications, and creating small pockets of
neutral charge which could slightly loosen a histone’s hold on DNA.
Histone modifications include acetylation, methylation, phosphorylation,
ubiquitination, sumoylation, and ADP-ribosylation. Most of these modifications occur
on the N-terminal tails, but a small number can be inserted internally in the proteinaceous
core. The combinations of these modifications result in a “histone code” (57) that can be
read by many cellular proteins required for nuclear functions such as transcription,
replication, chromosome condensation, and DNA repair. The following summaries
represent a handful of the complex combinations of modifications that exist for regulation
of DNA through core histone modifications.
Transcriptional Regulation
The most general and well-studied modification on histones is that of acetylation,
which is commonly associated with transcriptional activation, and will be discussed in
more detail in the next section. Most lysine residues in the first 20-30 amino acids of the
N-terminal tails of histones H2A, H2B, H3, and H4 can accommodate this modification
(Figure 6).
Methylation on lysine and arginine residues is commonly associated with
heterochromatin condensation and transcriptional repression. Some exceptions exist in
regard to specific H3 lysines, which are preferentially targeted by methyl groups during
transcriptional activation, such as H3 lysine 4 (H3K4) (58) and H3K36 (59). In contrast
to acetylation, lysines can exist in either mono-, di-, or tri-methylated states, and the
amount of this modification can correlate with the degree of function. For example,
tri-methylated H3K4 exists only at transcriptionally active genes sites, while dimethylated H3K4 can exist at both active and silent gene regions (60), suggesting that
Figure 6. Post-translational modifications of human histone N-terminal tails. See text for
higher degrees of methylation at H3K4 denote more specificity for transcriptional
activation. To regulate states of transcriptional activation, lysine residues that are
commonly acetylated during gene transcription, such as H3K9 and H3K27 (61), must be
switched from the “on” to the “off” state, and this is done through replacing the acetyl
group with 1-3 methyl groups (Figure 6). For example, after removal of the acetyl group,
site-specific enzymes such as the SUV39H1 histone methyltransferase target H3K9 for
tri-methylation (62) once the lysine residue has been primed by mono-methylation (63).
A small degree of phosphorylation occurs on histone H3 serine 10 (H3S10) in
interphase cells, and in the context of acetylation of H3K9 and H3K14, is a marker of
transcriptional activation. H3S10 phosphorylation also prevents methylation of H3K9
(62) and this modification is thought to act as a regulatory switch between modes of
transient activation (acetylation) and stable repression (methylation) through dynamic
modulation of H3K9, which is essential in genes that may need to be turned on quickly in
G0-G1 transitions (64) or inflammatory responses (65, 66).
In addition to small covalent additions such as acetylation and methylation,
transcriptional states of the tightly bound nucleosomal DNA are controlled by large
protein additions, which can bridge or wedge the nucleosome structure to recruit and
accommodate transcriptional machinery. Ubiquitin and the small ubiquitin-like modifier
(SUMO) are both similar in size to a histone protein, but can be added to specific lysine
residues to function in opposing manners in transcriptional regulation. Relatively low
levels of ubiquitin or SUMO modifications exist in vivo on histones, with only H2AK119
(67) and H2BK120 (68) sites shown to contain ubiquitin (Figure 6). All 4 core histones
have been shown to contain a SUMO modification in yeast and mammalian cells, but
only specific lysine residues of H2A (K126), H2B (K6/7 and K16/17) in yeast, and nonspecific N-terminal H4 lysines in both yeast and mammalian cells have been identified
(69, 70). Similar to the differences noted in methylation patterns and regulation of
transcription, the type of control exhibited by ubiquitin is dependent on the lysine residue
it modifies. Mono-ubiquitination of H2BK120 (H2BK120ub1) serves to establish
methylation of H3K4 and H3K79 (reviewd in (71)), both markers of active transcription,
while H2AK119ub1 participates in gene silencing (72), although it is unknown if there
are other histone modifications regulated by H2A ubiquitination. On the other hand,
sumoylation, regardless of what lysine residue or histone it modifies, can prevent both
ubiquitination and acetylation, playing a consistent role in transcriptional repression (69,
DNA Replication and Chromosome Condensation
Acetylation and phosphorylation have a dynamic role in regulation of the S- and
M-phases of the cell cycle. During replication, acetylation of H4 N-terminal lysines by
the histone acetyltransferase (HAT) HBO1 is required for proper S-phase progression and
incorporation of BrdU (73), while H3 and H4 acetylation by HATs is required for
continued replication origin firing (74).
After replication, the chromosomes must condense for distribution between
dividing cells. Heavy phosphorylation of H3S10 is a hallmark of mitotic chromosomes,
in which it is suggested that this histone modification recruits chromosome condensation
machinery in the beginning of M-phase (75, 76). Another prime phosphorylation event
during mitosis occurs on histone H3 threonine 3 (T3) (Figure 6), with similar timing to
the presence of H3S10 phosphorylation, although phosphorylation of H3T3 is necessary
for proper chromosome alignment (77). In contrast, failure of proper phosphorylation of
H3S10 in most biological systems does not result in a defect of normal chromosome
segregation through the progression of mitosis (78). Of those systems that do not have a
mitotic defect when S10 phosphorylation is prevented, either by kinase-specific
inhibition or S→A mutational analysis (reviewed in (79)), a redundant set of
phosphorylation events on histone H4S1 and H2AS1 (Figure 6) may exist to signal in a
similar manner as H3 phosphorylation (80). In agreement with the histone code, the
combination of phospho-H3S10 with additional phosphorylation sites on histones during
the cell cycle may be the signal needed by the cellular machinery to distinguish between
chromatin condensation progression or the transcriptional activation processes that
correlate with phosho-H3S10 in the presence of H3 acetylated lysines.
DNA Repair
The close proximity of DNA to the histone core prevents easy access for genome
maintenance when DNA damage occurs. Very specific types of histone modifications
exist to aid in the repair process and recruitment of repair complexes to the site of
different types of damage. Di-methylation, but not mono- or tri-methylation, of H4K20
functions in recognition of DNA double strand breaks (DSB) by signaling a G2/M phase
arrest (81, 82). UV-light-induced single strand lesions which are typically repaired by
nucleotide excision repair (NER) are marked by ubiquitination of H2AK119 (83) and Nterminal ubiquitination of histones H3 and H4 (84). Covalent attachment of an ADPribose molecule to specific residues in histone H1 and to a glutamic acid residue of H2B
(H2BE2) (Figure 6) results from DNA damage (85-87). The function of mono-ADPribosylation on histone residues is still not fully characterized or understood, but it is
possible that this initial modification can yield chains of poly-ADP-ribose to loosen
chromatin architecture similar to nuclear depletion of histone H1, or help specify the type
of damage that needs to be repaired in combination with other histone modifications (88).
One of the most well-characterized markers of both DNA DSB and single-strand
lesions is phosphorylation of an H2A histone variant, H2AX (89, 90). H2AX is an
evolutionary divergent variant of H2A (91), making up from 2-25% of a cell’s total H2A
content. The C-terminal domain of H2AX contains a highly conserved serine residue
(S139), which is an acceptor of a phosphate group within minutes of induction of DNA
damage (90). Phosphorylated H2AX (γH2AX) can be detected as distinct foci in nuclei
of damaged cells, and directs the recruitment of DNA repair machinery to the correct
location (reviewed in (92)). γH2AX is unique only to DNA lesions, and has no role in
other chromatin modifying functions, so it seems to be a vital signaling mechanism for
genomic maintenance. But, H2AX-null mice are viable, although more susceptible to
DNA damage and increased genomic instability (93, 94). This demonstrates that
responding to DNA damage heavily relies on, but is not dependent on, H2AX
phosphorylation, and a cell can turn to other histone modifications as part of the histone
code for repairing DNA damage.
Histone Acetylation
Acetylation is the enzymatic reaction of a HAT transferring the acetyl group from
Coenzyme A (CoA) to a lysine residue contained within the histone amino acid sequence.
The lysine residues are contained primarily in the basic N-terminal tails of histones,
which elicit a postive charge, attracting negatively charged nucleosomal cores into a tight
complex. Acetylation of these lysine residues neutralizes the positively-charged tails and
loosens the chromatin structure to allow for retained binding of HATs and recruitment of
transcription factors or other chromatin remodeling complexes to initiate, promote, and
regulate gene transcription.
Acetylation of histones was first identified in the 1960’s, as a result of the
difficulty in identifying the complete N-terminal peptide sequence of histones through
traditional methods. Multiple techniques were used to identify what type of moiety
would be masking the N-terminal region from identification, and only when hydrolysis of
histone fractions yielded acetate did it become clear that acetylation was a natural part of
the histone protein (95). Isolated nuclei could incorporate C14-labeled sodium acetate
into histones as an acetyl group, similar to the kinetics of radiolabeled uridine uptake into
nuclear RNA (96, 97), so a potential connection was made between histone acetylation
and function of RNA polymerase. Varying degrees of acetylated histone, ranging from
unmodified to highly acetylated, were added to histone-depleted nuclear extract and RNA
synthesis was monitored after addition of calf thymus or E. coli RNA polymerase. The
acetylated proteins could indeed form a histone-DNA complex similar to non-acetylated
proteins, and in a dose-dependent manner, regardless of the source of polymerase,
acetylated histone increased the ability of the nuclear extract to produce a labeled RNA
molecule (Figure 7) (96).
This important post-translation modification has since been associated with
transcriptional activation, and it is no surprise that multiple families of HAT enzymes
Calf Thymus Polymerase
E. Coli Polymerase
Figure 7. Inhibition of RNA polymerase by deacetylated histones. RNA polymerase
from both a mammalian and bacterial source were incubated with either no histone or
increasingly acetylated histones and nuclear extract, and the amount of a radiolabeled
RNA molecule was quantitated as a percent of RNA polymerase activation (modified
from (96)). AH, acetylated histones.
exist for its regulation. The first isolation of a cellular fraction containing HAT activity
was roughly 30 years ago, and in the age of genome sequencing, yeast served as the first
source for identification of the genes possessing this HAT function, Gcn5 and Hat1 (98,
99). These two enzymes are part of the GNAT (Gcn5 N-acetyltransferases) family of
HATs, which have human homologues including GCN5 and PCAF (reviewed in (100)).
The MYST (MOZ-YBF2/SAS3-SAS2-TIP60)) enzymes make up the second major
family of HATs, which includes the human enzymes TIP60, MOZ, and HBO1,
homologous to the yeast enzymes Esa1, Sas3, and Sas2, respectively (reviewed in (101)).
The GNAT family has a substrate specificity primarily for lysine residues on histone H3,
H4, and H2B tails, and the MYST family has a similar specificity for H3, H4, and H2A
histone tails (reviewed in (102)). This redundant targeting of similar lysine residues by
multiple HATs is regulated by their involvement in different large multi-protein
complexes, ranging from 500 kDa to 7 MDa in size. For example, both the PCAF and
GCN5 complexes target similar H3 lysine residues for acetylation, and while the 10subunit complex containing PCAF shares similarity with 6 subunits of the GCN5
complex, the GCN5 complex has an additional 9 subunits which differ from the PCAF
complex (reviewed in (103)), suggesting these differences may dictate distinct and nonredundant regulatory functions. For example, in vitro deletion studies of either Pcaf or
Gcn5 in chicken DT40 cells demonstrated that a phenotype was only present in cells
lacking Gcn5, and deletion of Pcaf did not affect cell viability. Interestingly, loss of
Gcn5 greatly up-regulated expression of Pcaf in a compensatory way, suggesting that
both enzymes can function in overlapping ways, but are also required for specific cellular
functions (104). Similar to the DT40 cell experiments, Pcaf deletion in vivo resulted in
viable mice with no observable phenotype other than compensation by increased
expression of Gcn5 (105). On the other hand, deletion of Gcn5 did not yield viable mice,
and embryonic lethality of double knockout mice (Pcaf-/-/Gcn5-/-) is compounded by loss
of both genes (106).
The amount of acetylation by HATs is regulated in a very dynamic way, and in
the context of active gene transcription, acetylation can occur and then be turned over
within 1-5 minutes (reviewed in (107)). The necessity for controlling acetylation on
histones has been implied in the previous sections, by its role in the crosstalk between
histone modifications and transcriptional regulatory factors, and acetylation is turned
over by a complementary family of enzymes, histone deacetylases.
Histone Deacetylases
Identification and Classification
After the identification of HAT enzymatic activity in cellular extracts (96, 97) and
the fact that acetylation had a more rapid turnover rate than the histones themselves
(108), enzymatic removal, as an alternative to quick degradation, was hypothesized to be
responsible for deacetylation of histones. An enzymatic nuclear fraction specific for
removal of an acetyl group on the ε-amino group of lysine residues was identified in calf
thymus extracts, which could deacetylate the known forms of histones (109-112).
Separation of this activity into distinct fractions was first performed in plants, and was
suggestive of the idea that the multiple roles of histone modification supported the
existence of multiple histone deacetylase (HDAC) enzymes (113). Purification and
cloning of a mammalian histone deacetylase enzyme HD1 (now known as HDAC1) (114)
demonstrated strong homology to a previously isolated yeast gene Rpd3, identified in a
screen for transcriptional repressors, but with unknown function (115). Purification and
analysis of the HDA and HDB histone deacetylase complexes from yeast lead to the
characterization of Rpd3 as an HDAC, and identification of a second enzyme, Hda1
(116). Rapid identification of HDAC family members ensued, and based on the
conservation of their amino acid sequences and presence of a deacetylase domain, the 18
known human HDACs can be grouped into 5 specific classes, which will be described in
relation only to their function as bona fide histone deacetylases, although they have nonhistone substrates (reviewed in (117)).
Class I
Class I HDACs share homology with the yeast Rpd3 enzyme, and include human
HDAC1, -2 (118), -3 (119), and -8 (120). The class I HDACs are expressed in most
tissues, and are predominantly localized to the nucleus. Their catalytic deacetylase
domain requires a Zn2+ ion to mediate the release of acetate and form a free lysine residue
(121). This enzymatic activity is commonly present in large, multi-subunit complexes
that require HDACs for their function. HDAC1 and -2 are very similar in their homology
and function, and can interact with each other (122). Thus, they are both found in the
Sin3 complex and the Mi-2/NuRD complex, in which the subunits are conserved from
humans to lower species such as Drosophila and C. elegans (123). The core Sin3 protein
has no known function on its own and relies on the interactions of 9 other subunits to
direct its function (124-126). The Sin3 complex can interact with multiple types of
adapter proteins, such as chromatin remodeling enzymes in addition to HDACs, and
transcription factors, which help to target the complex to specific regions of DNA for
repression (127). The Mi-2/NuRD complex is composed of 13 subunits, with 4 subunits
identical to those of the Sin3 complex, 2 of which are HDAC1 and HDAC2 (128-130).
In addition to the ability of the Mi-2/NuRD complex to repress transcription through
histone deacetylation, the Mi-2α and β subunits are responsible for an ATPase activity,
which is thought to mobilize nucleosomes to a more condensed formation after gene
transcription (131). Among the other class I HDACs, HDAC3 forms a separate complex
through binding with the proteins N-CoR or SMRT, and will be discussed further in the
next section. To date, HDAC8 is the only class I HDAC not known to form a higher
order complex for its function (132).
To determine the precise role of an HDAC in its representative complex, one can
use the power of genetics and animals models to delete or overexpress a specific gene
and analyze the consequences. The class I HDACs have been well-studied, and currently
there are knockout mouse models of Hdac1, -2, and -3, but to date, there are no reports of
an Hdac8 knockout model. Deletion of Hdac1 in mice revealed a vital role for this
enzyme in embryonic development, as embryos lacking Hdac1 did not survive past
embryonic day 10.5 (e10.5). Early embryos lacking Hdac1 contained a defect in
proliferation, which correlated with increased expression of p21 and p27, both of which
are cell cycle inhibitors. Up-regulation of both Hdac2 and Hdac3 was seen, and
increased enzymatic activity of Hdac2 was observed, but these increases in other class I
enzymes were not enough to compensate for the loss of Hdac1 (133). Studies in
embryonic stem cells (ESC) lacking Hdac1 also revealed similar abnormalities in cell
cycle regulation, but slightly more compensation by Hdac2 in this system (134).
Deletion of Hdac2 revealed a very different role for this enzyme in mouse
development. Although there was a small amount of embryonic lethality, most Hdac2null mice were born, although approximately half of those pups died within the first 3
weeks of life. Of those that did not survive during the postnatal time frame, Hdac2-/hearts had increased proliferation and thickened ventrical walls. The remaining Hdac2-/mice that survived into adulthood had no noticeable defects until stress was put upon the
heart. When adult Hdac2-/- mice were stressed with aortic restriction, the normal response
of cardiac hypertrophy was absent, suggesting an important role for Hdac2 in the
transmission and/or response of exogenous cellular signals (135).
Whole animal deletion of Hdac3 resulted in embryonic lethality before e7.5, and
Hdac3-null mouse embryonic fibroblasts (MEF) displayed increased apoptosis and DNA
damage (136). To understand how Hdac3 functions in adult animals, a conditional
knockout approach was taken to delete Hdac3 in a tissue-specific manner, which will be
the focus of this dissertation work in the coming chapters.
Class IIa, IIb, and IV
Originally, the second class of HDACs, which shares homology with the yeast
Hda1 enzyme, was comprised of human HDAC4, -5, -6, -7, -9, -10, and -11 (137-141).
Only recently have the functions of these HDACs been analyzed more completely, which
has lead to separation of class II HDACs into further distinct classes.
Class IIa HDACs include 4, -5, -7, and -9, and are roughly double the size of class
I HDACs due to a conserved, elongated region N-terminal to the deacetylase domain,
which is required for binding essential co-regulators. Class IIa enzymes have a much
more tissue-specific expression pattern compared to the ubiquitously expressed class I
enzymes. Primarily, all 4 class IIa members are expressed in the heart and skeletal
muscle, and HDAC4, -5, and -9 show expression in the brain, while HDAC7 is highly
expressed in the lung and thymus (142). Consistent with their expression pattern are their
interactions with a number of transcription factors which require this histone deacetylase
activity to repress transcription. For example, the class IIa enzymes have a conserved
binding region in their N-terminus for the calcium-dependent transcription factor
myocyte enhancer factor 2 (MEF2) (Figure 8A) (143). MEF2 functions in differentiation
of muscle cells, immune cells such as thymocytes, and neuronal cells, and requires the
nuclear localization and binding of class II HDACs to repress its transcription (144).
Signaling through the calcium/calmodulin-dependent protein kinase (CaMK) regulates
this binding by phosphorylating both HDACs and MEF2 (145, 146), releasing MEF2 to
interact with transcriptional co-activators (Figure 8B). So until there is a need to respond
to an intracellular signal, HDACs function to sequester MEF2, as well as other
transcription factors, in a repressed state.
In addition to the tissue specificity that differentiates class IIa from class I
HDACs, the class IIa enzymes also have a nuclear export signal (NES) to drive them
from the nucleus (147). As with the CaMK signaling, the phosphorylation events not
only release MEF2, but initiate a conformational change within the HDAC protein to
uncover the NES. Conserved binding sites for the chaperone protein 14-3-3 are also
present in class IIa HDACs, and 14-3-3 proteins serve to shield the nuclear localization
signal (NLS) and sequester HDACs once they are in the cytoplasm (Figure 8B)
Figure 8. Regulation of cellular localization of class IIa HDACs. A. Schematic of the
secondary structure common to the class IIa HDACs. Green box, MEF2 binding site;
blue box, 14-3-3 binding site; NES, nuclear export signal; NLS, nuclear localization
signal. B. In resting cells, class IIa HDACs can be complexed with MEF2, repressing its
transcriptional function. Upon activation of CaMK signaling, both MEF2 and IIa
HDACs are phosphorylated, releasing MEF2 from IIa HDACs to activate transcription,
and causing a conformation change in IIa HDACs, revealing a NES. 14-3-3 chaperone
proteins bind and conceal the NLS of IIa HDACs, and help to export and retain IIa
HDACs from the nucleus.
(148, 149). The cytoplasmic function of class IIa HDACs, other than a preventative
measure of their transcriptional repression, is unknown, although they are enzymatically
inactive once exported from the nucleus.If all 4 class IIa HDACs are expressed in similar
tissues and are regulated in the same way, questions arise concerning their redundancy or
need for multiple HDACs in the same tissues. In vivo deletion of these 4 genes using
mouse models revealed non-redundant functions, and unexpected phenotypes. Deletion
of either Hdac5 or Hdac9 resulted in viable mice, but unregulated heart hypertrophy
occurred after exposure to cardiac stress (150). Similarly, deletion of Hdac4 yielded
viable mice, but revealed a role for Hdac4 in repressing chrondrocyte hypertrophy during
bone development (151). Interestingly, Hdac7 is the only class IIa member that resulted
in embryonic lethality when deleted, due to weakened vascular structure at e11.0 (152).
Although these are seemingly different phenotypes, the commonality shared between
them consists of similar co-factor interactions the HDACs have in each specific tissue.
The class IIb HDACs include HDAC6 and -10. These enzymes differ from the
IIa enzymes by their tissue, as well as subcellular localization and overall protein
composition. Each enzyme is described as having two tandem deacetylase domains. In
HDAC6, both are completely functional (137). In HDAC10, only the first domain
contains a catalytically active site, while the second domain is leucine-rich and
homologous to the first domain, but lacks an active pocket for deacetylase activity (153,
154). Both class IIb enzymes predominantly localize to the cytoplasm, although
HDAC10 can also be present in the nucleus (153). The tissue specificity of the class IIb
enzymes include liver and kidney, but also heart and pancreas for HDAC6, and spleen for
HDAC10 (137, 140, 155).
Both class IIb enzymes can deacetylate histones in vitro (140, 153), but evidence
for in vivo specificity is lacking. Instead, HDAC10 acts as a co-repressor through
interactions with complexes occupied by class I HDACs in the nucleus (153), and
currently, the HDAC10 cytoplasmic function is undefined. On the other hand, even
though potential nuclear localization has been reported for HDAC6 (156), the main
function of HDAC6 has been specifically linked to its cytoplasmic localization. HDAC6
can localize to microtubule networks, and deacetylate its substrate, α-tubulin (157-159).
Acetylation and deacetylation of tubulin regulates its polymerization and stabilization,
thus HDAC6 has an important regulatory role in cell structure and motility. In support of
these data, mice lacking Hdac6 are viable but have significantly increased levels of
acetylated tubulin in all cell types examined, with only minor phenotypes associated with
bone and immune function, suggesting hyperacetylated tubulin is not detrimental under
normal circumstances (160).
HDAC11 is currently the only known HDAC with characteristics of both class I
and class II HDACs, which is why it is designated on its own as a class IV enzyme. Its
protein size and nuclear localization is reminiscent of a class I enzyme, yet its expression
in human tissues is limited to brain, heart, skeletal muscle, and kidney (141). The precise
molecular function of HDAC11 is currently unknown, because it can deacetylate histones
in vitro, but does not associate with any major co-repressor complexes, yet associates
with HDAC6. HDAC11 is highly conserved in many organisms (161), demonstrating
that although its function is unknown, it is likely that there is a vital role for this enzyme
in many forms of life.
Class III (Sirtuins)
This last class of HDACs has the least conservation to class I, II, or IV family
members. Originally identified in yeast as Silent Information Regulators (SIR) (162), or
sirtuins, SIR2 was found to have 3 more homologous proteins in yeast (163), and 7
homologous in humans, called SIRT1-7 (164, 165). These enzymes associate with
heterochromatic regions and telomeres, and can regulate longevity through metabolic
regulation in yeast (166, 167). Class III enzymes require NAD+ instead of Zn2+, and in
addition to histone deacetylase activity, they have the ability to enzymatically add an
ADP-ribose moiety to amino acids (168, 169).
The interest in studying enzymes involved in such diverse and relevant roles has
lead to the deletion of each of the 7 mammalian SIRT family members in individual
knockout mouse models. Deletion of a majority of the sirtuin genes resulted in viable
mice, with differing phenotypes. Both the Sirt2-/- and Sirt5-/- mice were viable with no
discernable phenotype in any tissue studied (170, 171). Both Sirt3-/- and Sirt4-/- mice
were viable as well, but had more pronounced phenotypes associated with metabolic
disruption. In Sirt3-/- mice, an increase in mitochondrial acetylation was observed, and
the metabolic enzyme glutamate dehydrogenase (GDH) was hyperacetylated specifically
(171). In Sirt4-/- mice, the same enzyme GDH is affected, but lack of GDH ADPribosylation in both liver mitochondria and pancreas occurs in the absence of Sirt4, which
disrupts insulin signaling (172). Deletion of Sirt1 and Sirt7 had more detrimental
phenotypes, both relating to heart dysfunction and p53 regulation. Sirt1-/- mice had only
a 10% survival rate at birth, and of those, two-thirds died postnatally due to defective
heart development. Studies in Sirt1-/- MEFs revealed that p53 was a target of
deacetylation by Sirt1, but the cells did not show any increased sensitivity to DNA
damage, even though hyperacetylated p53 is a more stable and active form of the protein
(173). In Sirt7-/- mice, increases in hyperacetylated p53 were also found, which lead to
increased apoptosis in cardiac tissue, both at basal levels and when tissue was stressed.
This increased level of cell death lead to a decreased life span by roughly 55% in Sirt7-/mice (174). The Sirt6-/- mice exhibit phenotypes pertaining both to DNA damage and
metabolism. Although Sirt6-/- mice are born viable, within 2 weeks they exhibit sharp
deterioration and death by 21-24 days of age. Analysis showed a drastic reduction in
adipose tissue, low to undetectable blood glucose levels, and increased thymocyte
apoptosis. In addition, Sirt6-/- MEFs were more prone to DNA damaging agents and
chromosome aberrations, suggesting the combination of metabolic imbalances and
genomic instability leads to rapid degeneration (175).
Inhibitors and Disease Treatment
As can be pointed out from the information presented thus far on histone
deacetylases, their role in the processes that control access to DNA are highly regulated,
and disruption of their enzymatic function, or deletion altogether as in mouse models,
demonstrates drastic consequences when HDACs are inhibited. Conversely,
overexpression or increased activation of HDACs can be unfavorable to a cell too, as
demonstrated by the amount of references that profile levels of HDACs in disease.
Cancer is the primary disease that cites increased HDAC function, but numerous other
diseases, such as neurodegenerative disorders (176), lupus (177), heart disease (178),
multiple sclerosis (179), and HIV (180) have seen beneficiary results when HDACs are
inhibited in these respective diseases.
Histone deacetylase inhibition was uncovered almost directly after the enzymatic
activity of histone deacetylation was isolated and characterized. Treatment of malignant
cells such as Friend erythroleukemia or HeLa cells with the compound n-butyrate could
induce morphological changes and a “switching to a non-malignant differentiating cell”
(181). Analysis of nuclear extracts by gel electrophoresis from n-butyrate treated cells
showed slower migrating bands of histone proteins, suggesting an increase in modified
protein. Phosphorylation, as a modification, was ruled out based on data demonstrating
incubation of histones with a bacterial phosphatase did not alter modified histone gel
migration (181). Instead, column filtration analysis of modified histones verified that the
increased molecular weight of histone proteins was due to an ε-N-acetyl-lysine moiety
(181). Rates of acetylation were measured in n-butyrate treated cells, and increases in
modified histones were not the result of increased acetyl transferase activity (182). Short
chain fatty acids such as n-butyrate can be metabolized to acetyl CoA, increasing the
cellular pool of the reactant necessary for an acetylation reaction. Specific metabolic
studies including n-butyrate, acetate (which can be metabolized to acetyl CoA also), and
propionate and isobutyrate, which are both metabolized to succinyl CoA, concluded in
inhibition of histone acetylation with each individual fatty acid, so increased acetylation
was not dependent on increases in the required reactant (183). Alternatively, turnover of
existing acetyl groups on histones was delayed, demonstrating n-butyrate treatment could
inhibit the deacetylase enzymatic function (182), because of competitive inhibition by
increased concentrations of a molecule that mimics acetate structure (183). But, nbutyrate has non-specific side effects in cell culture unrelated to HDAC inhibition (184,
185), and is a relatively weak inhibitor in vivo due to being rapidly metabolized, yielding
a short half-life when infused into the blood stream (186).
Some years later, in a screen for compounds that induce differentiation in a model
erythroleukemia cell line, the anti-fungal compounds trichostatin A (TSA) and C (187)
were identified to have strong differentiation properties and could inhibit cell cycle
progression, with TSA needing a lower concentration for a stronger effect than TSC
(188). Through studies using a murine tumor cell line unaffected by TSA, the target
mechanism by which this compound acted was through specific inhibition of the histone
deacetylase enzymatic fraction isolated from TSA-sensitive cell lines (189).
Inappropriate recruitment and function of HATs and HDACs is a common
hallmark of cancer, most characterized in promyelocytic leukemia (PML), acute myeloid
leukemia (AML), and acute lymphoblastic leukemia (ALL) (190-197), but also
associated with origins of solid tumors, such as glioblastomas, gastric, colorectal, breast,
and cervical cancers (198-206). Development of naturally occurring and synthetically
derived compounds to inhibit HDACs is based on modulating tumor-related phenotypes
such as cell differentiation, cell cycle arrest, apoptosis (extrinsic, intrinsic, or mitotic
catastrophe), accumulation of reactive oxygen species (ROS), and angiogenesis, which
have been thoroughly studied in malignant cell culture (207). Currently, there are 4 main
classes of HDAC inhibitors (HDI), based on their chemical structure, and an emerging
class of hybrid molecules (Figure 9).
The smallest and simplest types of compounds are the short-chain fatty acids (or
aliphatic acids), which include n-butyrate and valproic acid (VPA). While n-butyrate is a
natural bacterial by-product occurring from fiber fermentation, VPA is a synthetic
Figure 9. Histone deacetylase inhibitors. See text for details.
compound originally used in the treatment of epilepsy and bipolar disorder (208). Both
of these compounds work in the millimolar range to inhibit class I and II HDAC activity,
and have an important use for understanding HDAC function both in vitro and in vivo.
Their weaker binding to the HDAC binding pockets makes aliphatic acids the least
effective HDI used in the clinical setting (209), but VPA is currently in Phase II clinical
trials for use in combination with alkylating agents and radiation for treatment of patients
with glioblastomas (
A second class of HDI called benzamides shares no structural homology to
common HDIs such as n-butyrate or TSA. Efficacy of inhibition of these synthetic
compounds is in the micromolar range (209), and the compound MS-275 (Figure 9) has
anti-proliferative effects in both cell culture and xenograft nude mouse models of 7
different human tumor lines (210). This compound is also much more isoform-specific in
its HDAC inhibition, with selectivity for class I enzymes in the order of HDAC1/HDAC2
>HDAC3 >>HDAC8 (211). Although MS-275 is a more selective compound, it lacks
the potency of other classes of HDI currently in clinical trials, thus it is currently in Phase
I and II clinical trails in combination with other chemotherapeutic drugs such as Erlotinib
(kinase inhibitor) and azacitidine (DNA methylase inhibitor).
The HDI class of cyclic peptides are naturally occurring compounds isolated from
bacterial and fungal species, and can work in the nanomolar range of concentration. The
compound FK-228, or depsipeptide (Figure 9), is a “pro-drug” compound activated in
vivo through metabolism of the compound red FK, which leads to formation of reduced
sulfur bonds that are thought to interact with the zinc ion within the active site of HDACs
(212). FK-228 primarily inhibits class I HDACs at nanomolar concentrations, while
Figure 10. SAHA binding at the active site in the Aquifex aeolicus histone deacetylaselike protein (HDLP). Left hand panels represents ribbon diagram (top) and space filling
model (bottom) of HDLP, in which the active is clearly seen centrally located (white
box). The right hand panels shows how SAHA can fit into the active site and block
substrates from entering the binding pocket. (PDB 1C3S)
unable to inhibit class II enzymes such as HDAC4 and HDAC6 at this lower range (212).
FK-228 can induce growth arrest and increase apoptosis in human lymphoma and
leukemia cell lines, and can prolong survival in nude mice inoculated with lymphoma
cells (213), although toxicity has been reported for FK-228 at high concentrations in vivo
(214). Additionally, inhibition of angiogenesis has been observed with use of FK-228,
although these effects may be regulated through non-HDAC mechanisms (215, 216).
Based on these data, FK-228 is in Phase I and II clinical trials currently for multiple
types of blood and solid tumors as both a chemotherapeutic and anti-angiogenic
The final main class of HDI are the hydroxamates, which are the most abundant
type of HDI compounds. Their structural composition allows for the compounds to fit
within the catalytic pocket of HDACs while preventing access to the active site (Figure
10). TSA is classified as a hydroxamate compound, and works in the nanomolar range to
inhibit class I and II HDACs, but has undesirable toxic side effects. This naturally
occurring compound serves as a base for modeling synthetic hydroxamate molecules,
such as suberoylanilide hydroxamic acid (SAHA). SAHA works in the micromolar
range, and inhibits both class I and class II HDACs. Its activity as an anti-cancer drug
works through the common mechanisms of inducing growth arrest, cell death, and
increasing ROS (217-220). It also promotes degradation of the AML fusion protein
RUNX1-MTG8 (221), and may have anti-angiogenic properties as well (222). Currently,
SAHA (marketed under the name vorinostat) is the only HDI in Phase II/III clinical trials
for malignant mesothelioma, and approved by the FDA for cutaneous T-cell lymphoma
The aliphatic acids, benzamides, cyclic peptides, and hydoxamates are all
reversible inhibitors of HDAC function, but a few irreversible HDAC-binding
compounds have been isolated and manipulated to further exploit HDAC inhibition. The
epoxide compound trapoxin, isolated as a naturally occurring metabolite from fungus,
could induce morphological changes in transformed fibroblasts, similar to TSA (223).
Trapoxin could induce histone acetylation and inhibit partially purified HDACs
irreversibly with nanomolar concentrations. Structurally, trapoxin is a cyclic
tetrapeptide, which has an extended epoxide group (Figure 9) required for its inhibition of
class I and IIa HDACs (224), but this moiety is unstable in vivo, thus preventing its use as
an HDI (209). Hybrids, as they are called, consisting of combinations of structures from
known HDIs, were synthesized to create novel HDI compounds with a better chance at in
vivo usefulness. By utilizing the functional group from TSA that interacts with the zinc
ion in the HDAC catalytic domain, this domain replaced the epoxyketone function groups
of trapoxin, thus creating a reversible and potent HDI called cyclic hydroxamic acidcontaining peptide (CHAP1) (225) (Figure 9). Similarly, the functional group of TSA
was combined with the majority of the benzamide MS-275 to create SK-7041 (226)
(Figure 9). These hybrid molecules inhibit HDACs with nanomolar concentrations,
similar to unmodified TSA, while also having antiproliferative effects on cancer cell lines
and in vivo tumor growths (225, 226). Interestingly, both CHAP1 and the SK-7041 are
HDAC isoform-specific in their inhibition. CHAP1 can definitively inhibit HDAC1 and
HDAC4 enzymatic activity (225), while SK-7041 preferentially inhibits activity of
HDAC1 and -2, but not -3, -4, -5, or -6 (226).
Generation of hybrid compounds from already-existing HDI is leading the way
for more thoughtful drug development for chemotherapeutic agents. Steps are gradually
being made toward identifying an HDI with low toxicity, and possibly targeted to inhibit
specific isoforms. Additionally, these newer HDI are utilized in combination with
proven chemotherapeutic agents currently used in the clinic. For example, treatment of
chronic myeloid leukemia (CML) patients with the Bcr/Abl kinase inhibitor Gleevac
results in a high rate of cancer remission, but some patients can develop drug resistance,
thus resulting in relapse. The compound SK-7041 has recently been investigated as a
follow-up or compound treatment for CML in addition to Gleevac, with very effective
and promising results relating to induction of apoptosis and expression of cell cycle
inhibitory genes in vitro (227).
Of note, studies in cancer cell lines and tumor-bearing nude mice utilizing
naturally occurring and synthetically derived HDI can induce very beneficial changes in
cancer cells, yet leave normal cells mostly unharmed. The significance of this is that it
allows for one to treat cancer patients on a whole with HDI, without too much worry
about off-target effects in more quiescent, normal tissues. Indeed, specific deletion of
HDAC3, as well as HDAC2 and HDAC1 to a certain extent, leads to DNA damage
followed by apoptosis in proliferating, but not serum-starved (non-proliferating)
fibroblasts, suggesting a mechanism by which HDIs preferentially target highly
proliferative cancer cells (136). These data also demonstrate the importance of
understanding the function of each individual HDAC globally and their regulation in a
tissue specific manner.
Histone Deacetylase 3
Of the class I HDACs, histone deacetylase 3 (HDAC3) was the third member to
be fully sequenced and characterized (119, 228, 229). HDAC3 was classified as a class I
HDAC based on its homology to the well-studied enzymes HDAC1 and HDAC2, and
similar to the other class I enzymes, HDAC3 is expressed in most tissue types. The high
degree of similarity between these enzymes suggested that they had redundant roles in
the cell, but as previously described, each class I HDAC has distinct functions, which
can not be compensated for by the other enzymes within the same class.
HDAC3 has been mapped to the 5q31 chromosomal region in humans, and
similarly to a homologous region within murine chromosome 18 (206, 230, 231). There
is roughly 50-60% sequence homology between human HDAC3 and HDAC1 or HDAC2
in their N-terminal deacetylase domain. However, the C-terminal region of HDAC3
varies from any other known HDAC protein sequence, and the last 30 amino acids of the
C-terminal domain are also required for the histone deacetylase enzymatic ability of
HDAC3 (232, 233). Subcellular localization of class I enzymes is primarily nuclear, but
HDAC3 differs from other class I enzymes because of the presence of putative NES
signals (232, 233), which can direct HDAC3 to the cytoplasm and cell membrane
dependent on cell type and context (Figure 11) (234-236).
An oligomerization domain exists within the N-terminal 120 amino acids of
HDAC3, in which the protein can self-associate to form both dimers and trimers (232),
yet purified HDAC3 alone is enzymatically inactive (237). Instead, its enzymatic activity
is regulated by protein-protein interactions (other than with itself) and post-translational
Catalytic Domain
T390 S405
Figure 11. Structural organization of the human HDAC3 protein. Oligo, oligomerization
domain; N/S, N-CoR/SMRT binding domain; NLS, nuclear localization signal; NES,
nuclear export signal; black circles, phosphorylated residues.
modifications. Mapping of the HDAC3 protein sequence using phosphobase detection
databases has identified putative phosphorylation sites at a threonine and two different
serine residues specifically within the C-terminal domain (Figure 11) (238, 239).
Definitively, serine 424 (S424), which is located in a consensus sequence for casein
kinase 2 (CK2), is a direct site of phosphorylation by CK2 (239), and DNA-dependent
protein kinase (DNA-PK) utilizes HDAC3 as a substrate, potentially at residues T390
and/or S405 (Figure 11) (238). Regulation of HDAC3 phosphorylation is significant, as
increased phosphorylation of HDAC3 increases its basal enzymatic activity, although the
presence or absence of phosphorylation does not alter its subcellular localization or
interactions within protein complexes (238, 239).
The majority of modified and unmodified forms of HDAC3 exist in a large
protein complex utilized by multiple nuclear hormone receptors (NR) and transcription
factors to aid in the regulation of gene repression. Components of this ~2 MDa complex
include nuclear co-repressor (N-CoR) or silencing mediator for retinoid and thyroid
hormone receptors (SMRT), transducin β-like 1 (TBL1), TBL1-related protein (TBLR1),
and G-protein pathway suppressor 2 (GPS2), in 1:1 stoichiometric ratios with HDAC3
(240-243). Other class I HDACs can be found in sub-stoichiometric ratios with NCoR/SMRT/HDAC3 (242), or independently in a separate population of NCoR/Sin3/NURD complexes (244).
The N-CoR and SMRT proteins are highly homologous to each other and are
often considered interchangeably in relation to the HDAC3 repressor complex, yet they
are encoded by distinctly separate regions of the genome and have non-redundant
functions (245-250). Both contain binding domains to recruit NRs and mediate
repression, but preferentially are recognized by different types of these hormone- and
ligand-regulated transcription factors (251, 252). The repression activity of NCoR/SMRT/HDAC3 complexes is completely dependent on the catalytic activation of
HDAC3 by the presence of SANT (Swi3/Ada2/N-CoR/TFIIIB) domains (253, 254) and a
deacetylase activation domain (DAD) in the N-CoR/SMRT proteins, thus resulting in
transcriptional repression through histone deacetylation (237, 255). The roles of the
additional co-factors TBL-1, TBLR-1, and GPS2 are more likely to act in stabilization of
and substrate recognition by the N-CoR/SMRT/HDAC3 complex (242, 256).
The enzymatic activity of HDAC3 seems to be specific towards certain histone
tail lysine residues. When acetylated histone peptides or purified oligonucleosomes
reconstituted from recombinant histones were treated with purified, active HDAC3
complexes, strong specificity of deacetylation toward both histone H3 and K5/K12 of
histone H4 were reported (257-259). Similarly, the acetylation status of an endogenous
promoter (RARγ2), activated by all-trans retinoic acid (ATRA) and repressed by HDAC3,
in mouse NIH 3T3-L1 and human embryonic kidney (HEK) 293 cells was monitored.
Although histone H3 lysine residues were quickly deacetylated upon removal of ATRA,
the kinetics of H4 lysine deacetylation showed a specific and non-random pattern, with
rapid reversal of H4K5ac occurring first, followed by H4K8ac and H4K12ac.
Knockdown of Hdac3 expression using siRNA demonstrated that Hdac3 was required to
deacetylate those lysine residues when a cell necessitated repression of RARγ2 (260).
Additionally, the requirement of Hdac3 in vivo seems to have similar specificity when
global deacetylation is examined by western blot of total histone H3 and H4 (see Chapter
IV and (261)). Overall, these data suggest that HDAC3 has the ability to deacetylate
multiple histone lysine residues if forced in vitro, but under specific conditions at
designated promoter regions, the substrate specificity may differ in an in vivo context.
Classical targets of HDAC3 enzymatic activity are histone residues, but much
more complex data is emerging that HDAC3, as well as other HDAC family members,
have important functions in deacetylating non-histone substrates. Acetylation can affect
the stability, activity, and localization of proteins. Through deacetylation, HDAC3
regulates the transcription factors SRY and GCMa during embryogenesis (262, 263).
Interpretation of cellular signaling by MEF2, p53, and RelA are all controlled by
HDAC3-targeted deacetylation (264-266). Deacetylation by HDAC3 also negatively
regulates transcriptional elongation through targeting of the CDK9 subunit of RNA
polymerase II-required machinery (267). Interestingly, HDAC3 can regulate chromatin
remodeling by deacetylation of the HATs PCAF and CBP (263, 264). A potential
requirement exists for HDAC3 to not only target these known histone and non-histone
substrates, but also those of class IIa HDACs, in which an interaction with HDAC3/NCoR/SMRT is sufficient for their own enzymatic activity (268).
Transcription Factor and Nuclear Receptor Interactions
Loss of HDAC3 is detrimental to cell viability (136, 233), possibly through
deregulation of transcriptional control, although treatment of cells with HDI elicits only a
2-10% change in transcriptional profiles (269-272). Additional primary or secondary
transcription-independent factors such as triggering cell cycle checkpoints in S- or Mphase, mitochondrial-mediated apoptosis, and destabilization of oncogene/Hsp90
chaperone interactions, cannot be completely ruled out (221, 273-276).
In the context of transcription-dependent cell requirements, HDAC3 interacts with
a diverse number of transcriptional co-factors to mediate their function. The transcription
factor Krüppel-like factor 6 (KLF6) is defined as a tumor suppressor (277), but has an
additional role of utilizing HDAC3 for modulation of adipocyte differentiation (278). An
HDAC3 repressor complex also prevents c-Jun mediated gene expression, and mutations
in c-Jun, which mimic the avian viral isoform v-Jun, destabilize HDAC3 binding,
activating their oncogenic transforming potential (279, 280). In AML, the co-repressor
proteins MTG8 (myeloid translocation gene 8) and MTG16 can become fused to the
hematopoietic transcription factor RUNX1 as a result of chromosome translocations.
While both MTGs and RUNX1 can bind HDAC3 (discussed further in Chapter 3), the
inappropriate recruitment of HDAC3 by the fusion protein results in a switch from a
regulated transcription factor that can both activate and repress transcription, to an
unregulated repressor of RUNX1 target genes, providing a mechanism for the
development of AML (195, 281). The tumor suppressor RB directly interacts with E2F
transcription factors to prevent progression through G1- to S-phase of the cell cycle by
repressing E2F target genes. This repression mechanism includes RB binding to HDAC3
(282, 283). The RB/HDAC3 complex can additionally be recruited to peroxisome
proliferator-activated receptor γ (PPARγ), an important NR involved in metabolic
regulation (284).
PPARγ is classified into the family of NRs that utilize lipid intermediates as
activating ligands, while the classical NRs, which respond to endocrine hormones,
include estrogen receptor (ER), glucocorticoid receptor (GR), thyroid hormone receptor
(TR), and vitamin D receptor (VDR), and the NR group of orphan receptors have no
identified endogenous ligand for activation (reviewed in (285-287)). N-CoR and SMRT
were originally identified based on their interactions with NRs. More specifically,
PPARγ and TR preferentially recruit N-CoR/SMRT (242, 288-291), which in turn
requires the function of HDAC3 for repression mediated by these two receptors.
PPARγ was the third isotype to be described in the PPAR family of transcription
factors. Unlike its counterparts PPARα and PPARβ/δ, PPARγ is highly expressed in
adipose tissue, with only low to non-detectable expression in all other tissues, and is
strongly considered a master regulator of adipocyte differentiation and cellular energy
homeostasis. PPARγ exists as 2 isoforms, γ1 and γ2, which differ at their N-termini
through differential promoter usage and splicing. The PPARγ2 isoform is preferentially
expressed in adipose tissue, while PPARγ1 can be found at slightly higher levels in
tissues such as heart, spleen, and gut in addition to adipose tissue (292-298). Activation
of PPARγ occurs through binding of endogenous fatty acids such as linoleic acid,
arachidonic acid, and prostaglandins (299-302), although this diverse spectrum of
endogenous fatty acids, which have ranging affinities for PPARγ, has yet to be fully
characterized as to their biological relevance.
Synthetic PPARγ ligands (thiazolidinediones or TZD) have been developed to
clinically treat diabetic patients, due to their high affinity for PPARγ (303, 304). TZDs
have an insulin-sensitizing effect on liver, muscle, and adipose tissue (305-308), although
adverse side effects include increased adiposity and reversible congestive heart failure
due to plasma volume expansion (309, 310). Yet PPARγ still remains an important
biological target of pharmacological inhibitors to treat metabolic disorders.
In agreement with current models of NR regulation, N-CoR/SMRT/HDAC3
complexes bind to PPARγ in the absence of ligand to repress transcription, but in the
presence of ligand, a conformation change occurs that displaces co-repressor complexes
and recruits co-activators (Figure 12). The same is true for that of TR. Inactive TR
interacts with co-repressors (242, 290), but binding of the activating ligand, thyroid
hormone (T3), displaces these complexes and allows for recruitment of the HATs and
other chromatin modifying enzymes (311). T3 acts on 5 known isoforms of TR, TRα1,
TRα2, TRα3, TRβ1, and TRβ2 (312). The TRα isoforms are ubiquitously expressed,
with higher expression of TRα1 in skeletal muscle and brown adipose tissue. Less focus
has been put on TRα2 and -3, which may act as dominant negative isoforms of TRα1.
TRβ1 is also ubiquitously expressed, with its highest expression in the brain, kidney, and
liver. The TRβ2 isoform has very specific expression patterns in the pituitary gland and
certain regions of the brain. Thyroid deficiency can lead to abnormal neurological
development and muscular and cardiac deficiency, while hyperthyroidism can affect
metabolism in both the liver and adipose tissue (reviewed in (313)). Additionally,
knockout mouse models deficient in TRs demonstrate that TRs are required for normal
development, and although each TR isoform responds to T3 by activating transcription,
there is only a small degree of redundant function between each one (314).
All NRs control transcription through DNA binding, and in the case of PPARγ
and TR, each NR is bound at their respective response element sequences regardless of
the presence or absence of ligand (Figure 12). Both recognize the common response
element AGGTCA (and slight variations) in DNA sequences. Monomers or homodimers
of TR can recognize this half-site, or a direct repeat (DR) with a 4-bp gap (DR4) between
each half-site (312), while PPARγ recognizes the peroxisome proliferator response
Figure 12. Transcriptional regulation of nuclear receptors. The nuclear receptors TR and
PPARγ bind to the consensus DNA sequence AGGTCA either in the presence or absence
of ligand. A. Before binding of ligand, nuclear co-repressor complexes containing
HDAC3 bind nuclear receptors to keep them in a transcriptionally repressed state. B.
Upon ligand binding, a conformation change occurs, displacing the HDAC3-co-repressor
complex, and recruitment of chromatin remodeling enzymes and transcriptional
machinery occurs. NR, nuclear receptor; HAT, histone acetyltransferase; SWI/SNF,
chromatin remodeling complex; RNA Pol, RNA polymerase.
element (PPRE) DR1 (315). Although TR can interact with itself to regulate
transcription, enhanced binding to its response element is obtained by heterodimerizing
with retinoid X receptor (RXR) (316), while PPARγ requires binding to RXR for any
type of transcriptional function (317). The TR/RXR dimer responds only to T3 activation
to tightly regulate target gene expression, while the PPARγ/RXR dimer is a more
permissive complex that can become activated by both the RXR ligand 9-cis retinoic acid
and the numerous PPARγ ligands (317).
Although TR and PPARγ may have an added level of control through their
binding partner RXR, this only adds a layer of complexity as to how HDAC3 can
functionally keep track of the “when” and “where” of its enzymatic repressor ability in
relation to NR regulation. Thus, the aim of this dissertation is to begin to characterize the
requirements of HDAC3 in vivo by using a conditional mouse model to delete Hdac3 in
hematopoietic and liver tissues to understand its action on NRs and other transcription