Leukemia (1999) 13, 1473–1480
 1999 Stockton Press All rights reserved 0887-6924/99 $15.00
http://www.stockton-press.co.uk/leu
REVIEW
Molecular control of cell cycle progression in primary human hematopoietic stem
cells: methods to increase levels of retroviral-mediated transduction
MA Dao and JA Nolta
Division of Research Immunology/Bone Marrow Transplantation, Childrens Hospital of Los Angeles, and Departments of Pediatrics and
Craniofacial Developmental Biology, University of Southern California School of Medicine, 4650 Sunset Blvd, Mailstop 62, Los Angeles, CA
90027, USA
Pluripotent hematopoietic stem cells (HSC) are the ideal targets
for gene transfer because they can repopulate a sublethally
irradiated recipient, giving rise to all lineages of blood cells.
Thus, introduction of a corrected gene into HSC (stem cell gene
therapy) should ensure persistent transmission of the gene. To
date, the most efficient mode of gene delivery is via Moloney
murine leukemia virus (MoMuLV)-based retroviral vectors
which stably integrate into the genome of the target cell. The
quiescent nature of HSC and the fact that MoMuLV-based retroviral vectors can only integrate into dividing cells are major
obstacles in gene therapy. While increasing efforts have been
directed toward identifying growth factors which facilitate
division of primary hematopoietic progenitor and stem cells,
little is known about the molecular mechanisms which these
cells use to enter cell cycle. In this review, we will discuss the
correlation between the hematopoietic inhibitory and growth
factors and their impact on the regulation of the cell cycle
components.
Keywords: hematopoietic stem cells; retroviral-mediated transduction; cell cycle; CDK inhibitors; p27 kip-1
Introduction
Cell cycle is commonly thought to consist of four orderly
steps: G1, S, G2, and M. In reality, the cell cycle process is
more complex, and is comprised of 12 sequentially overlapping steps: each of the above four steps is further broken down
into early, mid, and late stages. The 12 steps are distinguished
by the appearance/disappearance and activation/inactivation
of different intracellular positive and negative regulators. Positive contributors include the cyclins and the cyclin-dependent
kinases (CDK) (Figure 1). Negative regulators involve the
cyclin-dependent kinase inhibitors (CDKI) and the retinoblastoma product (pRb). Overexpression and knockout studies
have defined the importance of each cyclin, CDK, and CDKI
at specific stages of the cell cycle in development. In addition,
much research has been focused on the regulation of these
cell cycle proteins via signaling cascades initiated at the surface membrane of the cells in response to surrounding factors,
such as cell–cell and cell–environment interactions. There is
evidence supporting the notion that mechanisms by which
cell cycle entry, progression, and exit are regulated may vary
between different cell types. The majority of research done
on mammalian cell cycle has been performed on fibroblasts.
Recently, there has been a growing interest in understanding
the molecular aspects of cell cycle entry in the field of hematopoiesis, as gene therapists acknowledge one of the greater
barriers to their success: the inability to stably introduce a corrective gene into nondividing pluripotent stem cells via retroCorrespondence: JA Nolta; Fax: 323 660 1904
Received 12 April 1999; accepted 15 June 1999
Figure 1
The cyclin-dependent kinases that mediate cell cycle
progression through the G1 phase in human hematopoietic progenitors, and their inhibitors that can be reduced to induce the onset of
S phase. For cell cycle to proceed through the G0/G1 phase into the
S phase, the complexes cyclin D/cdk4(6), cyclin E/cdk2, and cyclin
A/cdk2 must be sequentially assembled and activated. TGF␤ induces
an increase in p15 levels, which blocks assembly of cyclin D/cdk4(6).
In hematopoietic cells, treatment with anti-TGF␤ neutralizing antibody causes a dramatic reduction in p15 levels. The CDKI p27 blocks
activity of cyclin E/cdk2 and cyclin A/cdk2. Quiescence and serum
starvation cause an increase in p27 levels in fibroblastic cells. The
natural regulators of p27 levels in hematopoietic cells are not yet
known. However, treatment with antisense oligonucleotides can
greatly reduce p27 levels in hematopoietic progenitor and stem cells,
allowing cell cycle progression, when used in conjunction with
anti-TGF␤ neutralizing antibodies.21
viral-mediated transduction. Theoretically, integration of the
gene of interest into pluripotent and multipotential hematopoietic stem cells would ensure that the gene is transmitted into
all lineages of blood cells.
Hematopoietic stem cells
Hematopoiesis is the study of blood cell development. Primitive hematopoietic progenitors can be obtained from numerous sources including bone marrow, cord blood, mobilized
peripheral blood, and fetal liver. The vast majority of hematopoietic progenitors are quiescent, and reside in the G0/G1
phase of the cell cycle. Although there is still controversy over
the definitive surface markers of true pluripotent hematopoietic stem cells (HSC), there is unanimous agreement that HSC
have the ability to reconstitute a sublethally irradiated recipient with all blood cell lineages. In vitro studies have greatly
expanded the list of external factors which positively and/or
negatively regulate the proliferation and differentiation of
hematopoietic progenitor cells. These external factors include
cytokines, chemokines, cell–cell interactions, and cell–
Molecular control of hematopoietic stem cell cycle progression
MA Dao and JA Nolta
1474
stromal interactions, and have been thoroughly reviewed
elsewhere.1–5
While the study of hematopoietic progenitor cells in vitro
is useful and informative, the results do not necessarily predict
the behavior of true, reconstituting hematopoietic stem cells.
In vivo studies are necessary to define the cell surface phenotypes and ex vivo culture requirements of reconstituting
human HSC. Animal xenograft models of gene therapy provide a system to study culture, homing, and transduction of
multipotent, engrafting human hematopoietic stem cells, and
to monitor the duration and effects of an introduced gene in
an in vivo setting. Some interesting and informative studies
have been done using pre-immune fetal sheep as recipients
of the human cells, and other pre-immune large animal models are under investigation.6–8 However, due to the high cost
and level of surgical expertise involved with these models, the
most commonly used host for engineered human cells is the
immune deficient mouse. In the most easily done
murine/human xenograft models (NOD/SCID and bnx/hu),
there are no surgical manipulations required, but only an
intraperitoneal or tail vein injection of the transplant inoculum. In addition to the ease of manipulation of human-tomouse transplantation, the murine bone marrow has proven
to be an excellent microenvironment for the development and
differentiation of human hematopoietic cells, both in vivo and
in vitro.9,10 Four types of immune deficient mouse xenograft
systems have been used with success to study retroviralmediated transduction of human hematopoietic stem cells: the
scid/hu thy/liv, scid/hu bone, NOD/SCID/hu, and bnx/hu
systems (reviewed in Refs 8, 11 and 12).
We established the bnx/hu xenograft co-transplantation system, which uses beige/nude/xid (bnx) mice as recipients of
human CD34+ hematopoietic progenitor cells that have been
marked ex vivo by retroviral vectors. Long-term hematopoiesis
(up to 18 months post-transplantation) from the human stem
and progenitor cells is sustained by co-transplantation of
human bone marrow stromal cells engineered to produce
human IL-3. Production of IL-3, which is species-specific, in
the mice has no discernable effect on murine hematopoiesis.13
Reciprocally, human hematopoietic cells do not undergo
long-term hematopoiesis in bnx mice in the absence of
human IL-3. From the transplanted human cells, all myeloid
lineages, erythroid progenitors, and T and B lymphocytes
develop.11,14–16 Clonogenic human hematopoietic progenitors
have been recovered for as long as 18 months.17 The bnx/hu
xenograft model allows investigation of the impact of different
ex vivo transduction protocols on both the efficiency of gene
transfer into stem cells and the maintenance of pluripotentiality.11,14–16,18–21 The major goal for gene therapy of both
acquired and inborn diseases of the hematopoietic system is
transduction of long-lived pluripotent stem cells, so that genetically corrected mature blood cells could be produced for
the lifetime of the recipient. Animal xenograft models currently provide the best surrogate models for testing homing
and hematopoiesis from reconstituting human HSC after culture and transduction. Following the general discussions on
gene therapy and cell cycle regulation in the next sections,
we will discuss the use of the bnx/hu xenograft system to test
the effects of inducing cell cycle entry by molecular intervention (to allow increased levels of retroviral-mediated gene
transfer) on long-term hematopoiesis from the manipulated
stem cells.
Hematopoietic stem cell gene therapy
One of the major subdivisions of research in gene therapy is
the development of gene delivery vehicles which can allow
permanent integration of the therapeutic gene into the host
genome. Three aspects must be taken into account when
designing viral vectors to act as gene delivery vehicles. First,
the vector capsid must be able to make contact with the cell
membrane via receptor mediated mechanisms or via envelope-membrane fusion, followed by endocytosis. The use of
gibbon ape leukemia virus (GALV) envelope partially satisfies
this requirement since hematopoietic primitive progenitors
have been shown to express better levels of receptors for
GALV22,23 than the previously used amphotropic receptors.24
To enhance entry of the viral vector into the cytoplasm, many
research groups are now exploring the use of VSV-G pseudotyped vectors which do not rely on receptor numbers to infect
cells, but fuse through the phospholipid bilayer.25,26 Second,
once in the cytoplasm, the viral particle carrying the gene of
interest must traverse through another lipid bilayer membrane,
the nuclear membrane, to arrive in the nucleus. This step is a
persistent problem with the use of Moloney murine leukemia
virus-based vectors because the viral nucleoprotein complex
can only reach the target DNA to integrate when the cell
enters mitosis and the nuclear membrane disintegrates (Figure
2).27 Lentiviral vectors are based on the HIV-1 virus, and can
enter the nucleus of nondividing cells.28–31 To date, MoMuLVbased vectors remain the most commonly used mode of gene
delivery because they meet the third criteria of a gene delivery
vehicle: they can stably integrate into the host genome, unlike
adenoviral vectors which remain episomal.32,33 It is not yet
proven whether lentiviral vectors can stably integrate with
high efficiency in hematopoietic progenitors, although active
Figure 2
The major hurdle to successful transduction of hematopoietic stem cells using Moloney murine leukemia-based retroviral
vectors is the quiescence of the cells. Breakdown of the nuclear membrane is required for the viral nucleoprotein complex to gain access
to the host cell chromosomes. Mitogenic factors or a reduction in
CDKI levels are required to induce cell cycle entry in primary human
hematopoietic stem cells. During mitosis, the nuclear membrane disintegrates, and the large retroviral nucleoprotein complex can gain
access to the host cell chromosomes. The reverse-transcribed proviral
DNA integrates into the chromosomes of one daughter cell, and is
passed to the progency of that cell. The challenge in human hematopoietic stem cell gene therapy, using Moloney murine leukemia-based
retroviral vectors, is to induce cell cycle entry to allow integration
without inducing terminal differentiation.
Molecular control of hematopoietic stem cell cycle progression
MA Dao and JA Nolta
research in that area is ongoing, and encouraging results have
been obtained with long-term expression.29,34
Because retroviral vectors require target cell proliferation for
successful gene insertion, efforts have been directed toward
identifying conditions which will induce active cycling of the
hematopoietic cells. Following the demonstration that the
combination of interleukin-3 (IL-3) and interleukin-6 (IL-6)
promoted the entry of quiescent murine hematopoietic progenitor cells into active cell cycle,35 we and others determined
that culturing human marrow cells in IL-3 and IL-6 greatly
increased the percentages of clonogenic progenitor cells
(CFU-C) which can be transduced with retroviral vectors.36,37
Subsequently, we and others demonstrated that additional
cytokines, primarily G-CSF, stem cell factor, and flt-3 ligand
can increase cell cycle, and enhance the extent of transduction into hematopoietic progenitors above and beyond what
could be obtained with IL-3 and IL-6 alone.37–40 The presence
of an underlying marrow stromal cell layer or a fragment of
fibronectin greatly enhances both survival and transduction of
committed and more primitive hematopoietic progenitors.41–44
However, in contrast to the relative ease of transduction of
committed progenitors, the more primitive, reconstituting HSC
have proven to be much more difficult to transduce, due to
their quiescence.
While stimulation with cytokines can readily recruit hematopoietic progenitors, such as CD34+ cells, into S phase, to
allow retroviral integration (Figure 2), the quiescent stem cells
are more refractory to the stimulatory effects of cytokines.
CD34+/CD38− cells are a primitive, quiescent population that
has been demonstrated to be greatly enriched for lymphomyeloid repopulating cells.7,38 This cell population lacks the more
mature, differentiated hematopoietic progenitors that are contained within the bulk CD34+ population.45–48 We previously
determined that retroviral-mediated transduction of a portion
of the long-term reconstituting stem cells contained within the
CD34+/CD38− population from human bone marrow required
at least 5 days in culture.38 By culturing HSC for extended
periods in vitro, to promote cell division, one might concomitantly induce their progression toward terminal differentiation,
depending on the cytokines and ex vivo culture conditions
used. Animal xenograft models have been very valuable in
determining ex vivo culture conditions which promote the
lowest levels of differentiation. Using the bnx/hu xenograft
system, our group determined that HSC lose homing capacity
and/or commit to differentiation during culture with serum
and cytokines, if kept in suspension with no opportunity to
engage their integrin receptors, for 72 h. By culturing the cells
on plates coated with the COOH-terminal fragment of fibronectin, the long-term reconstituting capacity could be at
least partially maintained.18 We also determined that the
capacity of HSC derived from human bone marrow to
undergo B lymphopoiesis following transplantation into bnx
mice is lost after 24 h in culture (even in cells maintained
on FN support). The current view is that retroviral-mediated
transduction should be done in hours, not days, to maintain
the highest degree of stem cell reconstitution potential.
In order to determine methods to induce cell cycle entry in
as short a period as possible, we and other research groups
have focused our effort in understanding how cellular components of the hematopoietic cell cycle are regulated. By understanding the internal factors that regulate cell cycle entry, one
may be able to manipulate the external factors to obtain a
balance of forces for rapid cell cycle induction without promoting terminal differentiation. In the final section of this
review, we describe the recent progress made by our laboratory toward this goal.
G0/G1 to S phase transition
The vast majority of hematopoietic progenitors are in the
G0/G1 phase of the cell cycle. In the presence of a combination of growth factors and the proper microenvironment, a
portion of the CD34+ cells can be stimulated to enter the S
phase of the cell cycle. However, as mentioned above, the
more primitive hematopoietic progenitors have been shown
to respond to growth factors significantly less efficiently, and
are much more difficult to recruit into S phase.38,45,48,49 The
fate of the cell, whether to remain quiescent, self renew, differentiate, or undergo apoptosis is determined in mid G1 where
the external factors can exert negative or positive influences.
For instance, TGF␤, a pleiotropic factor, inhibits cell cycle
entry in mid G1 via downregulation of cyclins and/or cyclindependent kinases or via upregulation of cyclin-dependent
kinase inhibitors.50 Stem cell factor is an example of a cytokine which has been shown to enhance cell cycle entry by
downregulation of specific cyclin kinase inhibitors, in hematopoietic progenitors and cell lines.51 Thus, to induce cell cycle
entry in hematopoietic stem cells, it is imperative to understand and identify the key cellular proteins involved in driving
the cell cycle forward, and the mechanisms by which they
can be controlled in vitro.
The critical point during the cell cycle is the mid G1 into
early S phase. Within this transitional step, there is a restriction point (R), which functions as the ‘all-or-none’ gate; once
a cell passes the restriction point, it is irreversibly committed
to cell cycle progression. Prior to R, the cell rests in a G0/G1
stage where its fate depends solely on the external survival
stimuli which include cytokines, chemokines, and mesenchymal cells. Progression through G1 and entry into S phase
requires the kinase activity of three cyclin/cdk complexes:
cyclin D/cdk4 (or cdk6) and cyclin E/cdk2 which are involved
in sequential inactivation of the retinoblastoma product (pRb);
and cyclin A/cdk2 which is required for DNA synthesis. A
decrease in either the formation of these complexes or in their
kinase activity can result in blockage or delay in cell cycle
entry.
The cyclin D/cdk4(6) complex is one of the critical driving
forces for cell cycle entry. The formation of the complex can
be regulated at two stages: synthesis of the individual components and their association. Expression of both D-type cyclins
and their catalytic partners is tissue-restricted. While epithelial
cells express cyclin D1, hematopoietic cells express cyclin D2
and D3. Hematopoietic cell lines, for instance 32Dcl3, have
been demonstrated to express cdk4, whereas CD34+ primary
hematopoietic progenitors express cdk6 (Ref. 52, and Dao and
Nolta, unpublished results).
Cyclin D is a major rate limiting factor in G1 phase progression (Figure 1). In response to mitogen stimulation, cyclin
D synthesis is induced in G1. The protein normally has a short
half-life, but there is an accumulation of cyclin D in the
nucleus as cells approach the G1/S transition. Then, following
S phase entry, the cyclin D protein level declines and disappears from the nucleus.53 Both growth inhibitory and growth
promoting factors can modulate cyclin D synthesis. For
instance, in rat intestinal epithelial cells, a decrease in cyclin
D1 synthesis occurs in response to TGF␤ 1.54 In response to
IL-3, there is an increase in cyclin D2 and D3 in the hematopoietic cell line 32Dcl3.55 In addition, there is evidence that
1475
Molecular control of hematopoietic stem cell cycle progression
MA Dao and JA Nolta
1476
pRb modulates the level of cyclin D. In Rb−/− cells, the level
of cyclin D is reduced as compared to cells containing pRb.
Co-transfection of Rb-deficient cell lines with a cyD1 promoter-luciferase construct and the Rb gene resulted in stimulation of the cyclin D1 promoter.56
The association of cyclin D with cdk4 and cdk6 is blocked
by mechanisms that have not yet been determined in many
cell types, and mitogenic stimulation can induce assembly
factor(s) that facilitate formation of the complex.57 The
MEK/ERK signalling pathway has been implicated in promoting the formation of active cyclin D/cdk4 complexes. Cheng et
al postulated that the MEK-mediated enhancement of complex
formation could occur through phosphorylation of molecular
chaperones such as cdc37 or assembly factors such as
p21.58,59
Mitogen stimulation can also modulate the level of cdk4.
Treatment of 32Dcl3 cells with TGF␤ resulted in a decrease
in cdk4 protein levels, whereas treatment with IL-3 induced
an increase in the cdk4 levels in these cells.60 Association of
cyclin D with cdk4(6) can be inhibited by the Ink4 family of
cyclin-dependent kinase inhibitors, resulting in G1 arrest. The
Ink4 family consists of p15, p16, p18, and p19, in mammalian
cells. Similar to cyclin D and cdk4(6), individual Ink4 members are modulated by different factors in different cells. For
instance, p19Ink4d has been shown to be induced in 32Dcl3
cells when stimulated to differentiate along the macrophage
lineage.61 Treatment of the human histiocytic lymphoma cell
line, U937, with onconase (a 12 kDa protein homologous to
pancreatic RNase A, isolated from amphibian oocytes)
resulted in induction of p16Ink4a accompanied by G1 arrest.62
Onconase shows cytostatic and cytotoxic activity in vitro,
inhibits growth of tumors in mice and is currently in phase III
clinical trials.63–65
Cell cycle arrest was also observed when p15Ink4b was
induced in mink lung epithelial cells following treatment with
TGF␤.66 Our laboratory demonstrated that TGF␤ addition to
primary hematopoietic progenitors in culture caused an
increase in p15Ink4b levels, and that the CDKI levels could be
dramatically decreased in the same cells by addition of neutralizing antibody to TGF␤.21 Members of the Ink4 family
primarily mediate G1 arrest by binding directly to cdk4 or
cdk6, and thus preventing their association with the regulatory
partner, cyclin D, in mid to late G1. Inhibition of cdk2 by p16
occurs through a different mechanism, since the two proteins
do not directly associate. CDK4 is sequestered by p16, which
leads to a reassortment of cyclin–CDK inhibitor complexes,
with more p27 binding to cdk2.67,68
Once assembled, the cyclin D/cdk4(6) complex is regulated
by a cyclin-activating kinase (CAK) which phosphorylates the
threonine-172 residue on cdk4; the complex is then activated
to phosphorylate pRb.69,70 In the absence of this activation
step, the cyclin D/cdk4(6) complex lacks kinase activity. For
instance, a cdk4 mutant which contains a nonphosphorylatable residue, alanine, in place of threonine-172, associates
with cyclin D, but the complex is inactive. A cdk4 mutant
which contains a phosphorylatable residue, serine, in place of
threonine-172, forms an active cyclin D/cdk4 complex which
phosphorylates pRb.69,70 Therefore, while assembly of the
complex can proceed in the absence of phosphorylation, activation of its kinase activity requires CAK phosphorylation of
threonine-172 residue on cdk4.
In late G1, cyclin E/cdk2 association results in an active kinase which then hyperphosphorylates and inactivates pRb at
G1/S transition. Regulation of the complex has been shown to
primarily exist at the transcriptional level. Cyclin E is synthe-
sized periodically. There is a burst of cyclin E transcription
immediately prior to S phase. As S phase progresses, there is
a diminution in cyclin E levels as a result of its dissociation
from its catalytic partner cdk2; degradation of dissociated
cyclin E occurs via the ubiquitin-proteasome mediated pathway.71 Two forms of cyclin E have been identified: a 48 kDa
cyclin E and a 43 kDa splice variant cyclin E which lacks the
cyclin box. Only the 48 kDa isoform of cyclin E associates
with cdk2 to phosphorylate histone H1 and pRb, suggesting
the cyclin box as the interaction site between cdk2 and cyclin
E.72 Overexpression of cyclin E precipitated cell cycle entry by
accelerating G1, and microinjection of anti-cyclin E antibodies
blocked entry into S phase.73
The presence of cyclin E alone does not ensure S phase
entry; cyclin E must associate with its catalytic partner, cdk2,
to drive cell cycle progression. Mink lung epithelial cells
treated with TGF␤ do not show modulation in the levels of
cyclin E or cdk2. However, there is a reduction in cyclin
E/cdk2 complex levels due to a decrease in the stable interaction between the two components, in TGF␤-treated cells.74
Cell cycle arrest can also occur in the presence of preformed
cyclin E/cdk2 complexes, because cdk2 remains inactive until
it is activated via phosphorylation on its tyrosine-15 residue.75
For instance, although senescent human fibroblasts have been
shown to express higher levels of cyclin E than early-passage
fibroblasts, these cells cannot be induced to enter S phase in
the presence of mitogen: the kinase activity of cyclin E/cdk2
complex formed was low due to the accumulation of unphosphorylated and thus, inactive cdk2.75
The kinase activity of cyclin E/cdk2 can also be inhibited
by induction of the corresponding cyclin dependent kinases,
p21cip-1 and p27kip-1, which bind to cyclin/cdk complexes. The
negative effect of p27 on the kinase activity of cyclin E/cdk2
has been examined via transfection of HELA cells with p27
alone and via co-transfection of p27 with cyclin E. Transfection with p27 induced G1 arrest. There was no decrease in
cdk2 levels; however there was a significant reduction in
cdk2-associated kinase activity. Co-transfection of p27 with
cyclin E restored the cdk2 kinase activity and removed the G1
block.76 Inhibition of cyclin E/cdk2 kinase activity has also
been linked to TGF␤-mediated cell cycle arrest in LPS-stimulated murine B lymphocytes. In the presence of TGF␤, these
cells expressed an increase in p27Kip1 levels, which inhibited
the kinase activity of cyclin E/cdk2 complexes.77 Of interest,
the cyclin E/cdk2 complex has been shown to phosphorylate
p27 on threonine-187 in late G1.78 Following phosphorylation, p27 is degraded via the ubiquitin–proteasome pathway.79 In summary, there are three regulatory steps involving
cyclin E/cdk2 complex function at the G1/S transition: at the
level of transcription of cyclin E, at the level of cdk2 phosphorylation, and at the level of modulation of kinase activity
by cdk inhibitors.
The initiation of DNA replication depends on the
expression of an additional regulatory protein, cyclin A. The
protein exists as two forms: cyclin A and cyclin A1.80–83
Unlike other G1 related regulatory proteins, cyclin A is not
commonly associated with tumorigenesis. Expression of cyclin
A1, on the other hand, correlates with numerous hematological malignancies.80,82,83 RT-PCR analysis of peripheral blood
from patients with CML or ALL showed expression of cyclin
A1 mRNA while peripheral blood from healthy donors did not
express cyclin A1.83
Cyclin A expression is regulated in a cell cycle-related
fashion. In HELA cells arrested in G1 phase, cyclin A
immunostaining was negative.84 As the cells began DNA syn-
Molecular control of hematopoietic stem cell cycle progression
MA Dao and JA Nolta
thesis, cyclin A first appeared and then accumulated as the
cells progressed through S phase. As the cells proceeded into
mitosis, cyclin A disappeared. In interphase, cyclin A staining
was exclusively nuclear whereas in prophase, there was diffuse staining for cyclin A throughout the cell. No detection
of cyclin A was found in cells at metaphase, anaphase, or
telophase.84 The expression of cyclin A is primarily regulated
at the transcriptional level. NIH 3T3 cells transfected with
cyclin A promoter/luciferase constructs had constant luciferase activity during exponential growth. When these cells were
synchronized and arrested in G1 following treatment with
nocodazole, there was a significant reduction in luciferase
activity. Stimulation with serum recruited cells to enter S
phase along with full activation of the cyclin A promoter.85 In
vivo footprinting analysis of 5⬘ flanking sequences of the
murine cyclin A gene was done to identify regions that were
shielded from digestion by bound transcription factors. There
was a direct correlation between the protection of a GC-rich
sequence and the repression of cyclin A transcription in
G0/early G1.86 This indicates binding of a cell cycle-responsive
transcription factor in G0/early G1 cells, and was not observed
at later stages of the cell cycle. Moreover, the footprint is
present in dimethyl sulfoxide-induced differentiating and not
in proliferating Friend erythroleukemia cells. A mutation
within the GC-rich region relieved the transcriptional
repression, resulting in a constitutively active cyclin A
promoter.86
Cyclin A is critical at the onset of DNA replication and at
the G2/M phase. Microinjection of anti-cyclin A antibody into
G0/G1-arrested HS68 cells (non-immortalized human
fibroblasts) inhibits DNA synthesis following serum stimulation. The inhibition was not observed in cells arrested at the
beginning of S phase. When HELA cells, which had already
proceeded through DNA synthesis, were microinjected with
anti-cyclin A antibody, there was complete but reversible cell
cycle arrest: reinjection of cyclin A restored the cells’ ability
to undergo mitosis and cytokinesis.84 Flow cytometry
immunofluorescence analysis that detected a nuclear-bound
form of cyclin A in G1 and G2 further demonstrated its
involvement in the G1/S and G2/M transition phases.87
Expression of cyclin A alone is not sufficient for cell cycle
progression. First, cyclin A complexes with cdk2 to form a
partially active kinase. Then, only following phosphorylation
by CAK on Thr160 of cdk2, the kinase complex becomes fully
activated to phosphorylate various substrates. For instance, the
E2F–pRB and E2F–p107 complexes can be disrupted via phosphorylation by cyclin A/cdk2, resulting in increased levels of
free E2F.88 In addition, cyclin A has also been shown to downmodulate the binding of E2F-1 to the E2 promoter by phosphorylating E2F-1.88 Induced expression of exogenous cyclin
A in early G1 has been shown to result in phosphorylation of
pRb, suggesting its role at the G1/S transition.89
The kinase activity of cyclin A/cdk2, and therefore progression through S phase, is tightly regulated by p27Kip1. In
studies using fibroblast cell lines transfected with a Tet-cyclin
A construct, it was shown that induction of cyclin A
expression in the presence of serum resulted in premature reentry into the cell cycle, compared to treatment with serum
alone. While the level of exogenous cyclin A level remained
constant throughout the cycle, its activity fluctuated from
being nearly undetectable in early G1 to peaking in late G1.89
When extracts from quiescent cells were mixed with isolated
preformed/active cyclin A/cdk2 complexes, the kinase activity
of cyclin A/cdk2 was abolished. When extracts from quiescent
cells were first depleted of p27Kip1 prior to mixing with cyclin
A/cdk2, the kinase activity of cyclin A/cdk2 was not
inhibited.90 In a further level of control of cyclin A levels by
p27Kip1, the CDKI was recently demonstrated to activate a latent protease that cleaves cyclin A at the N-terminus.91 In
addition, p27Kip1 has also been shown to indirectly modulate
the transcription of cyclin A by abrogating the cyclin E/cdk2mediated transactivation of the cyclin A promoter.92 Thus,
p27Kip1 is a prominent negative regulator of cyclin A/cdk2
kinase activity.
Manipulation of hematopoietic stem cell cycle to enhance
retroviral-mediated gene therapy
We rationalized that since p27Kip1 is a major factor in preventing cell cycle entry by antagonizing cyclin A induction,
levels, and activity, and thus in maintaining the quiescent state
in fibroblastic cells, that it might also be playing an important
role in hematopoietic stem cell quiescence (Figure 1). We had
previously demonstrated that a reduction in levels of another
CDKI important in maintaining quiescence, p15, could be
attained in primary human hematopoietic stem and progenitor
cells by neutralizing TGF␤ in the medium. However, a
reduction in TGF␤ levels was not sufficient to bring 4-HCresistant CD34+ cells (reconstituting stem cells which are
synchronized in G0) into cycle, to allow transduction.21 We
therefore studied the effects of a reduction in p27Kip1 levels
on hematopoietic stem cell cycle.
Since the extracellular factors that control p27Kip1 levels in
primary human hematopoietic stem cells are so far unknown,
we chose to reduce levels through the use of antisense oligonucleotides (ON). After lengthy studies, we determined that
lipofectants, commonly used to augment ON delivery to many
cell types, are toxic to the hematopoietic reconstituting and
long-term culture initiating cells (Dao and Nolta, unpublished
data). However, entry of the ON into an average of 68% of
the cells, 12–18 h after addition, could be achieved by soaking the cells in a high concentration of the ON: 50 ␮M. Unfortunately, this concentration of ON is very cost-ineffective, and
we are striving to find more natural methods of p27Kip1
reduction in hematopoietic stem cells, to apply to clinical
gene therapy trials.
The studies done to date therefore targeted the p27Kip1
mRNA using antisense oligonucleotides (ON). C-5 propyne
modified phosphorothioate anti-p27 16-mers93 were introduced into human CD34+ cells, and 4-HC-resistant CD34+
cells, at a concentration of 50 ␮M in serum-free medium with
IL-3, IL-6, and SCF.21 Mis-sense ON bearing the same modifications (mis-p27) and medium alone were used as controls.
A significant increase in the number of colony-forming units
was obtained from 10 000 CD34+ cells incubated for 12 h with
anti-p27 ON, as compared to the same number of mis-p27treated control cells, and those cultured in medium alone.21
The anti-p27 ON also caused an increase in the number of
colony-forming units from the 4-HC-resistant progenitors
(which are synchronized in the G0 phase of the cell cycle) in
comparison to those treated with mis-sense p27 ON, but did
not permit retroviral-mediated transduction, when used alone.
Both neutralization of TGF␤ and antisense ON to p27 kip-1
were required to obtain transduction of a portion of the 4-HC
resistant CD34+ cells.21 This phenomenon is likely due to the
cross-regulation of p15 and p27 levels through subcellular
localization, as described in elegant studies by Reynisdottir
et al.94
We next used the combination of anti-TGF␤ antibodies to
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MA Dao and JA Nolta
1478
reduce p15 levels, and antisense ON to p27 kip-1 to attempt
transduction of CD34+38− cells, a primitive and quiescent
stem cell population. We had previously demonstrated that
this population could only be transduced after an incubation
period of 5–7 days with cytokines, using the ‘old’ technology,
that was not based on the molecular status of the hematopoietic stem cells.38 Cells treated with anti-TGF␤ antibody and
antisense ON to p27kip-1 expressed a significant increase in
levels of cyclin A protein and stained positive for Ki67,
characteristic of cells entering S phase.21 Very little cyclin A
or Ki67 expression was detected in stem cells exposed to antiTGF␤ antibodies alone, without p27 reduction, verifying the
importance of p27 in maintaining stem cell quiescence.
As a follow-up to the in vitro studies, we transplanted bnx
mice with the treated, transduced CD34+38− cells to address
an important concern: do primitive progenitors recruited into
cycle by the combination of anti-TGF␤ Ab and anti-p27 ON
retain their multipotentiality and capacity for long-term
engraftment? The supposedly ‘improved’ transduction technology would be of little practical use, if cells induced into
cycle were also forced into terminal differentiation. Bone marrow CD34+/38− cells were pre-incubated in serum-free
medium for 12 h with anti-TGF␤ Ab and anti-p27ON vs the
scrambled mis-sense oligonucleotide. Retroviral supernatant
from the PG13/LN line was then added for an additional 12
h. Only cells that had received the dual combination of antiTGF␤ Ab and anti-p27 ON had significant CFU growth and
levels of transduction.21 Cells from each experimental condition were co-transplanted with human stroma producing
hIL-3 into bnx mice. Mice were harvested and analyzed by
FACS, human-specific CFU assay, neo PCR, and inverse PCR
11 months post-transplantation. Human CD45+ cell
engraftment levels were equivalent between mice transplanted with cells treated with anti-TGF␤ Ab and anti-p27 ON
and mice transplanted with cells treated with mis-p27 ON
alone.21 There was an equal distribution of the human hematopoietic lineages that had developed in the two groups. Bone
marrow samples were further analyzed by neo PCR and
human-specific CFU plating for G418 resistant colonies. Only
mice that had been transplanted with human progenitors
transduced with anti-TGF␤ Ab and anti-p27 ON had neo
gene-marked human cells in their marrow.21 Equivalent numbers of human-specific colonies developed from the bone
marrow of the two groups of mice, but G418-resistant CFU
only grew from mice transplanted with human progenitors
transduced with anti-TGF␤ Ab and anti-p27 ON.21 The recovery, from the bnx bone marrow, of human T lymphocyte and
myeloid clones with many different clonal integration sites
demonstrated that multiple primitive human hematopoietic
stem and progenitor cells had been transduced, using the
optimized conditions.21
Therefore, primitive hematopoietic progenitors induced into
cell cycle entry by the combination of anti-TGF␤ Ab and antip27 ON can be transduced by retroviral vectors in a 24-h
period, and retain the ability to reconstitute and durably
engraft immune deficient mice. The conditions that are currently used in our laboratory for transduction by Moloney
murine leukemia-based retroviral vectors are the following:
fibronectin support, serum-free medium, the cytokines IL-3,
IL-6, SCF, and Flt3 ligand, anti-TGF␤ antibody, and antisense
ON to p27kip-1.18,21 Retroviral supernatant is currently being
produced with 10% fetal calf serum, and added in equal volumes to the transduction medium to generate a final concentration of 5%. Surprisingly, medium containing 5% serum was
demonstrated to be better at recruiting hematopoietic stem
and progenitor cells into S phase than total serum-free
medium, perhaps due to an upregulation of p27 levels beyond
what could be neutralized by the ON in serum-free culture
(Dao and Nolta, manuscript in preparation). Serum-free conditions are known to upregulate p27 levels in fibroblastic
cells, and the same may be true in HSC.
The conditions described in the latter portion of this review
have been demonstrated to allow retroviral-mediated transduction, in a 24-h period, of 20–25% of primitive human
hematopoietic cells that are capable of long-term (1 year)
multilineage hematopoiesis in a xenograft model.21 We are
currently studying the factors that prevent the remainder of
the human hematopoietic stem cells from being rapidly
recruited into S phase by a reduction in the levels of the CDK
inhibitors p15INK4B (using anti-TGF␤ neutralizing antibodies
and medium with low serum) and p27kip-1 (using antisense
oligonucleotides). In addition, the extracellular factors that
control p27 levels and activity in primitive human hematopoietic cells are currently under investigation. Our eventual goal
is to develop methods to recruit at least 50% of the primitive
hematopoietic cells into a single cell cycle ex vivo, to allow
retroviral-mediated transduction. Our hope is that the genecorrected cells would then return to quiescence, relatively
unaffected by the manipulation, and home normally to the
bone marrow of the gene therapy recipients, to allow production of corrected, mature progeny for the lifetime of the
patient.
Acknowledgements
This work was supported by a grant from the NIH NHLBI
(SCOR No. 1-P50-HL54850), a grant from the NIH NIDDK
(1RO1DK53041), start-up funding from the John Connell
Gene Therapy Foundation, a Career Development Award
from the Childrens Hospital of Los Angeles Research Institute,
and a James A Shannon Directors’ Award from the NIH
NIDDK.
References
1 Whetton AD, Spooncer E. Role of cytokines and extracellular
matrix in the regulation of haemopoietic stem cells. Curr Opin
Cell Biol 1998; 10: 721–726.
2 Verfaillie CM. Adhesion receptors as regulators of the hematopoietic process. Blood 1998; 92: 2609–2612.
3 Kincade PW, Oritani K, Zheng Z, Borghesi L, Smithson G, Yamashita Y. Cell interaction molecules utilized in bone marrow. Cell
Adhes Commun 1998; 6: 211–215.
4 Simmons PJ, Haylock DN. Use of hematopoietic growth factors
for in vitro expansion of precursor cell populations. Curr Opin
Hematol 1995; 2: 189–195.
5 Simmons PJ, Levesque JP, Zannettino AC. Adhesion molecules in
haemopoiesis. Baillières Clin Haematol 1997; 10: 485–505.
6 Kawashima I, Zanjani ED, Almaida-Porada G, Flake AW, Zeng H,
Ogawa M. CD34+ human marrow cells that express low levels of
Kit protein are enriched for long-term marrow-engrafting cells.
Blood 1996; 87: 4136–4142.
7 Civin CI, Almeida-Porada G, Lee MJ, Olweus J, Terstappen LW,
Zanjani ED. Sustained, retransplantable, multilineage engraftment
of highly purified adult human bone marrow stem cells in vivo.
Blood 1996; 88: 4102–4109.
8 Zanjani ED, Almeida-Porada G, Flake AW. The human/sheep xenograft model: a large animal model of human hematopoiesis. Int
J Hematol 1996; 63: 179–192.
9 Berardi AC, Meffre E, Pflumio F et al. Individual CD34+CD38low
CD19−CD10− progenitor cells from human cord blood generate B
lymphocytes and granulocytes. Blood 1997; 89: 3554–3564.
Molecular control of hematopoietic stem cell cycle progression
MA Dao and JA Nolta
10 Rawlings DJ, Quan SG, Kato RM, Witte ON. Long-term culture
system for selective growth of human B-cell progenitors. Proc Natl
Acad Sci USA 1995; 92: 1570–1574.
11 Dao MA, Nolta JA. Use of the bnx/hu xenograft model of human
hematopoiesis to optimize methods for retroviral-mediated stem
cell transduction (Review). Int J Mol Med 1998; 1: 257–264.
12 Dick JE, Bhatia M, Gan O, Kapp U, Wang JC. Assay of human
stem cells by repopulation of NOD/SCID mice. Stem Cells 1997;
15: 199–203; Discussion 204–207.
13 Nolta JA, Hanley MB, Kohn DB. Sustained human hematopoiesis
in immunodeficient mice by cotransplantation of marrow stroma
expressing human interleukin-3: analysis of gene transduction of
long-lived progenitors. Blood 1994; 83: 3041–3051.
14 Nolta JA, Dao MA, Wells S, Smogorzewska EM, Kohn DB. Transduction of pluripotent human hematopoietic stem cells demonstrated by clonal analysis after engraftment in immune-deficient
mice. Proc Natl Acad Sci USA 1996; 93: 2414–2419.
15 Dao MA, Hannum CH, Kohn DB, Nolta JA. FLT3 ligand preserves
the ability of human CD34+ progenitors to sustain long-term hematopoiesis in immune-deficient mice after ex vivo retroviralmediated transduction. Blood 1997; 89: 446–456.
16 Dao MA, Pepper KA, Nolta JA. Long-term cytokine production
from engineered primary human stromal cells influences human
hematopoiesis in an in vivo xenograft model. Stem Cells 1997;
15: 443–454.
17 Nolta JA, Arakawa-Hoyt J, Dao MA, Barsky L, Uribe L, Puck J,
Weinberg KI. In vivo generation of IL-2 responsive human T lymphocytes in a chimeric mouse model of stem cell ␥c gene therapy
for X-SCID. Blood 1999 (submitted).
18 Dao MA, Hashino K, Kato I, Nolta JA. Adhesion to fibronectin
maintains regenerative capacity during ex vivo culture and transduction of human hematopoietic stem and progenitor cells. Blood
1998; 92: 4612–4621.
19 Dao MA, Yu XJ, Nolta JA. Clonal diversity of primitive human
hematopoietic progenitors following retroviral marking and longterm engraftment in immune-deficient mice. Exp Hematol 1997;
25: 1357–1366.
20 Nolta JA, Kohn DB. Retroviral-mediated transduction of human
hematopoietic stem cells. In: Boyle A (ed). Current Protocols in
Human Genetics. John Wiley: 1997, unit 13.7.
21 Dao MA, Taylor N, Nolta JA. Reduction in levels of the cyclindependent kinase inhibitor p27(kip-1) coupled with transforming
growth factor beta neutralization induces cell-cycle entry and
increases retroviral transduction of primitive human hematopoietic cells. Proc Natl Acad Sci USA 1998; 95: 13006–13011.
22 Miller AD, Garcia JV, von Suhr N, Lynch CM, Wilson C, Eiden
MV. Construction and properties of retrovirus packaging cells
based on gibbon ape leukemia virus. J Virol 1991; 65: 2220–2224.
23 Kiem HP, Andrews RG, Morris J et al. Improved gene transfer into
baboon marrow repopulating cells using recombinant human fibronectin fragment CH-296 in combination with interleukin-6,
stem cell factor, FLT-3 ligand, and megakaryocyte growth and
development factor. Blood 1998; 92: 1878–1886.
24 Orlic D, Girard LJ, Jordan CT, Anderson SM, Cline AP, Bodine
DM. The level of mRNA encoding the amphotropic retrovirus
receptor in mouse and human hematopoietic stem cells is low and
correlates with the efficiency of retrovirus transduction. Proc Natl
Acad Sci USA 1996; 93: 11097–11102.
25 Agrawal YP, Agrawal RS, Sinclair AM et al. Cell-cycle kinetics and
VSV-G pseudotyped retrovirus-mediated gene transfer in bloodderived CD34+ cells. Exp Hematol 1996; 24: 738–747.
26 Burns JC, Friedmann T, Driever W, Burrascano M, Yee JK. Vesicular stomatitis virus G glycoprotein pseudotyped retroviral vectors:
concentration to very high titer and efficient gene transfer into
mammalian and nonmammalian cells (see comments). Proc Natl
Acad Sci USA 1993; 90: 8033–8037.
27 Miller DG, Adam MA, Miller AD. Gene transfer by retrovirus vectors occurs only in cells that are actively replicating at the time
of infection. Mol Cell Biol 1990; 10: 4239–4242.
28 Naldini L, Blomer U, Gallay P et al. In vivo gene delivery and
stable transduction of nondividing cells by a lentiviral vector.
Science 1996; 272: 263–267.
29 Miyoshi H, Smith KA, Mosier DE, Verma IM, Torbett BE. Transduction of human CD34+ cells that mediate long-term engraftment of
NOD/SCID mice by HIV vectors. Science 1999; 283: 682–686.
30 Kafri T, van Praag H, Ouyang L, Gage FH, Verma IM. A packaging
cell line for lentivirus vectors. J Virol 1999; 73: 576–584.
31 Miyoshi H, Blomer U, Takahashi M, Gage FH, Verma IM. Development of a self-inactivating lentivirus vector. J Virol 1998; 72:
8150–8157.
32 Neering SJ, Hardy SF, Minamoto D, Spratt SK, Jordan CT. Transduction of primitive human hematopoietic cells with recombinant
adenovirus vectors. Blood 1996; 88: 1147–1155.
33 Frey BM, Hackett NR, Bergelson JM et al. High-efficiency gene
transfer into ex vivo expanded human hematopoietic progenitors
and precursor cells by adenovirus vectors. Blood 1998; 91:
2781–2792.
34 Case SS, Price MA, Jordan CT et al. Stable transduction of quiescent CD34(+)CD38(−) human hematopoietic cells by HIV-1-based
lentiviral vectors. Proc Natl Acad Sci USA 1999; 96: 2988–2993.
35 Suda T, Suda J, Ogawa M. Proliferative kinetics and differentiation
of murine blast cell colonies in culture: evidence for variable G0
periods and constant doubling rates of early pluripotent hemopoietic progenitors. J Cell Physiol 1983; 117: 308–318.
36 Bodine DM, Karlsson S, Nienhuis AW. Combination of interleukins 3 and 6 preserves stem cell function in culture and enhances
retrovirus-mediated gene transfer into hematopoietic stem cells.
Proc Natl Acad Sci USA 1989; 86: 8897–8901.
37 Nolta JA, Kohn DB. Comparison of the effects of growth factors
on retroviral vector-mediated gene transfer and the proliferative
status of human hematopoietic progenitor cells. Hum Gene Ther
1990; 1: 257–268.
38 Dao MA, Shah AJ, Crooks GM, Nolta JA. Engraftment and retroviral marking of CD34+ and CD34+CD38− human hematopoietic
progenitors assessed in immune-deficient mice. Blood 1998; 91:
1243–1255.
39 Luskey BD, Rosenblatt M, Zsebo K, Williams DA. Stem cell factor,
interleukin-3, and interleukin-6 promote retroviral-mediated gene
transfer into murine hematopoietic stem cells. Blood 1992; 80:
396–402.
40 Petzer AL, Hogge DE, Landsdorp PM, Reid DS, Eaves CJ. Selfrenewal of primitive human hematopoietic cells (long-term-culture-initiating cells) in vitro and their expansion in defined
medium. Proc Natl Acad Sci USA 1996; 93: 1470–1474.
41 Moore KA, Deisseroth AB, Reading CL, Williams DE, Belmont JW.
Stromal support enhances cell-free retroviral vector transduction
of human bone marrow long-term culture-initiating cells. Blood
1992; 79: 1393–1399.
42 Moritz T, Patel VP, Williams DA. Bone marrow extracellular
matrix molecules improve gene transfer into human hematopoietic
cells via retroviral vectors. J Clin Invest 1994; 93: 1451–1457.
43 Moritz T, Dutt P, Xiao X et al. Fibronectin improves transduction
of reconstituting hematopoietic stem cells by retroviral vectors:
evidence of direct viral binding to chymotryptic carboxy-terminal
fragments. Blood 1996; 88: 855–862.
44 Wells S, Malik P, Pensiero M, Kohn DB, Nolta JA. The presence
of an autologous marrow stromal cell layer increases glucocerebrosidase gene transduction of long-term culture initiating cells
(LTCICs) from the bone marrow of a patient with Gaucher disease.
Gene Therapy 1995; 2: 512–520.
45 Hao QL, Shah AJ, Thiemann FT, Smogorzewska EM, Crooks GM.
A functional comparison of CD34+CD38− cells in cord blood and
bone marrow. Blood 1995; 86: 3745–3753.
46 Hao QL, Smogorzewska EM, Barsky LW, Crooks GM. In vitro
identification of single CD34+CD38− cells with both lymphoid and
myeloid potential. Blood 1998; 91: 4145–4151.
47 Rawlings DJ, Quan S, Hao QL et al. Differentiation of human
CD34+CD38− cord blood stem cells into B cell progenitors in vitro.
Exp Hematol 1997; 25: 66–72.
48 Shah AJ, Smogorzewska EM, Hannum C, Crooks GM. Flt3 ligand
induces proliferation of quiescent human bone marrow
CD34+CD38− cells and maintains progenitor cells in vitro. Blood
1996; 87: 3563–3570.
49 Jordan CT, Yamasaki G, Minamoto D. High-resolution cell cycle
analysis of defined phenotypic subsets within primitive human
hematopoietic cell populations. Exp Hematol 1996; 24: 1347–
1355.
50 Florenes VA, Bhattacharya N, Bani MR, Ben-David Y, Kerbel RS,
Slingerland JM. TGF-beta mediated G1 arrest in a human melanoma cell line lacking p15INK4B: evidence for cooperation
1479
Molecular control of hematopoietic stem cell cycle progression
MA Dao and JA Nolta
1480
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
69
70
71
72
73
between p21Cip1/WAF1 and p27Kip1. Oncogene 1996; 13:
2447–2457.
Takahira H, Gotoh A, Ritchie A, Broxmeyer HE. Steel factor
enhances integrin-mediated tyrosine phosphorylation of focal
adhesion kinase (pp125FAK) and paxillin. Blood 1997; 89:
1574–1584.
Della Ragione F, Borriello A, Mastropietro S et al. Expression of
G1-phase cell cycle genes during hematopoietic lineage. Biochem
Biophys Res Commun 1997; 231: 73–76.
Baldin V, Lukas J, Marcote MJ, Pagano M, Draetta G. Cyclin D1
is a nuclear protein required for cell cycle progression in G1.
Genes Dev 1993; 7: 812–821.
Ko TC, Sheng HM, Reisman D, Thompson EA, Beauchamp RD.
Transforming growth factor-beta 1 inhibits cyclin D1 expression
in intestinal epithelial cells. Oncogene 1995; 10: 177–184.
Ando K, Ajchenbaum-Cymbalista F, Griffin JD. Regulation of G1/S
transition by cyclins D2 and D3 in hematopoietic cells. Proc Natl
Acad Sci USA 1993; 90: 9571–9575.
Muller H, Lukas J, Schneider A et al. Cyclin D1 expression is regulated by the retinoblastoma protein. Proc Natl Acad Sci USA 1994;
91: 2945–2949.
LaBaer J, Garrett MD, Stevenson LF, Slingerland JM, Sandhu C,
Chou HS, Fattaey A, Harlow E. New functional activities for the
p21 family of CDK inhibitors. Genes Dev 1997; 11: 847–862.
Cheng M, Sexl V, Sherr CJ, Roussel MF. Assembly of cyclin Ddependent kinase and titration of p27Kip1 regulated by mitogenactivated protein kinase kinase (MEK1). Proc Natl Acad Sci USA
1998; 95: 1091.
Stepanova L, Leng X, Parker SB, Harper JW. Mammalian
p50Cdc37 is a protein kinase-targeting subunit of Hsp90 that
binds and stabilizes Cdk4. Genes Dev 1996; 10: 1491–1502.
Ando K, Griffin JD. Cdk4 integrates growth stimulatory and inhibitory signals during G1 phase of hematopoietic cells. Oncogene
1995; 10: 751–755.
Adachi M, Roussel MF, Havenith K, Sherr CJ. Features of macrophage differentiation induced by p19INK4d, a specific inhibitor
of cyclin D-dependent kinases. Blood 1997; 90: 126–137.
Juan G, Ardelt B, Li X, Mikulski SM, Shogen K, Ardelt W, Mittelman A, Darzynkiewicz Z. G1 arrest of U937 cells by onconase is
associated with suppression of cyclin D3 expression, induction of
p16INK4A, p21WAF1/CIP1 and p27KIP and decreased pRb phosphorylation. Leukemia 1998; 12: 1241–1248.
Mikulski SM, Viera A, Deptala A, Darzynkiewicz Z. Enhanced in
vitro cytotoxicity and cytostasis of the combination of onconase
with a proteasome inhibitor. Int J Oncol 1998; 13: 633–644.
Deptala A, Halicka HD, Ardelt B, Ardelt W, Mikulski SM, Shogen
K, Darzynkiewicz Z. Potentiation of tumor necrosis factor induced
apoptosis by onconase. Int J Oncol 1998; 13: 11–16.
Schein CH. From housekeeper to microsurgeon: the diagnostic
and therapeutic potential of ribonucleases (published erratum
appears in Nat Biotechnol 1997; 15: 927). Nat Biotechnol 1997;
15: 529–536.
Ewen ME, Sluss HK, Whitehouse LL, Livingston DM. TGF beta
inhibition of Cdk4 synthesis is linked to cell cycle arrest. Cell
1993; 74: 1009–1020.
McConnell BB, Gregory FJ, Stott FJ, Hara E, Peters G. Induced
expression of p16(INK4a) inhibits both CDK4- and CDK2-associated kinase activity by reassortment of cyclin-CDK-inhibitor complexes. Mol Cell Biol 1999; 19: 1981–1989.
Mitra J, Dai CY, Somasundaram K, El-Deiry WS, Satyamoorthy K,
Herlyn M, Enders GH. Induction of p21(WAF1/CIP1) and inhibition of Cdk2 mediated by the tumor suppressor p16(INK4a). Mol
Cell Biol 1999; 19: 3916–3928.
Matsushime H, Quelle DE, Shurtleff SA, Shibuya M, Sherr CJ, Kato
JY. D-type cyclin-dependent kinase activity in mammalian cells.
Mol Cell Biol 1994; 14: 2066–2076.
Kato JY, Matsuoka M, Strom DK, Sherr CJ. Regulation of cyclin
D-dependent kinase 4 (cdk4) by cdk4-activating kinase. Mol Cell
Biol 1994; 14: 2713–2721.
Clurman BE, Sheaff RJ, Thress K, Groudine M, Roberts JM. Turnover of cyclin E by the ubiquitin–proteasome pathway is regulated
by cdk2 binding and cyclin phosphorylation. Genes Dev 1996;
10: 1979–1990.
Sewing A, Ronicke V, Burger C, Funk M, Muller R. Alternative
splicing of human cyclin E. J Cell Sci 1994; 107: 581–588.
Ohtsubo M, Theodoras AM, Schumacher J, Roberts JM, Pagano
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
M. Human cyclin E, a nuclear protein essential for the G1-to-S
phase transition. Mol Cell Biol 1995; 15: 2612–2624.
Koff A, Ohtsuki M, Polyak K, Roberts JM, Massague J. Negative
regulation of G1 in mammalian cells: inhibition of cyclin Edependent kinase by TGF-beta. Science 1993; 260: 536–539.
Dulic V, Drullinger LF, Lees E, Reed SI, Stein GH. Altered regulation of G1 cyclins in senescent human diploid fibroblasts:
accumulation of inactive cyclin E-Cdk2 and cyclin D1-Cdk2 complexes. Proc Natl Acad Sci USA 1993; 90: 11034–11038.
Kwon TK, Nordin AA. Overexpression of cyclin E and cyclindependent kinase inhibitor (p27Kip1): effect on cell cycle regulation in HeLa cells. Biochem Biophys Res Commun 1997; 238:
534–538.
Bouchard C, Fridman WH, Sautes C. Effect of TGF-beta1 on cell
cycle regulatory proteins in LPS-stimulated normal mouse B lymphocytes. J Immunol 1997; 159: 4155–4164.
Morisaki H, Fujimoto A, Ando A, Nagata Y, Ikeda K, Nakanishi
M. Cell cycle-dependent phosphorylation of p27 cyclin-dependent kinase (Cdk) inhibitor by cyclin E/Cdk2. Biochem Biophys
Res Commun 1997; 240: 386–390.
Vlach J, Hennecke S, Amati B. Phosphorylation-dependent degradation of the cyclin-dependent kinase inhibitor p27. EMBO J
1997; 16: 5334–5344.
Yang R, Nakamaki T, Lübbert M, Said J, Sakashita A, Freyaldenhoven BS, Spira S, Huynh V, Müller C, Koeffler HP. Cyclin A1
expression in leukemia and normal hematopoietic cells. Blood
1999; 93: 2067–2074.
Yang R, Muller C, Huynh V, Fung YK, Yee AS, Koeffler HP. Functions of cyclin A1 in the cell cycle and its interactions with transcription factor E2F-1 and the Rb family of proteins. Mol Cell Biol
1999; 19: 2400–2407.
Yang R, Morosetti R, Koeffler HP. Characterization of a second
human cyclin A that is highly expressed in testis and in several
leukemic cell lines. Cancer Res 1997; 57: 913–920.
Kramer A, Hochhaus A, Saussele S, Reichert A, Willer A,
Hehlmann R. Cyclin A1 is predominantly expressed in hematological malignancies with myeloid differentiation. Leukemia 1998;
12: 893–898.
Pagano M, Pepperkok R, Verde F, Ansorge W, Draetta G. Cyclin
A is required at two points in the human cell cycle. Embo J 1992;
11: 961–971.
Henglein B, Chenivesse X, Wang J, Eick D, Brechot C. Structure
and cell cycle-regulated transcription of the human cyclin A gene.
Proc Natl Acad Sci USA 1994; 91: 5490–5494.
Huet X, Rech J, Plet A, Vie A, Blanchard JM. Cyclin A expression
is under negative transcriptional control during the cell cycle. Mol
Cell Biol 1996; 16: 3789–3798.
Stivala LA, Scovassi AI, Bianchi L, Prosperi E. Nuclear binding of
cell cycle-related proteins: cyclin A versus proliferating cell
nuclear antigen (PCNA). Biochimie 1995; 77: 888–892.
Suzuki-Takahashi I, Kitagawa M, Saijo M, Higashi H, Ogino H,
Matsumoto H, Taya Y, Nishimura S, Okuyama A. The interactions
of E2F with pRB and with p107 are regulated via the phosphorylation of pRB and p107 by a cyclin-dependent kinase. Oncogene
1995; 10: 1691–1698.
Resnitzky D, Hengst L, Reed SI. Cyclin A-associated kinase activity
is rate limiting for entrance into S phase and is negatively regulated in G1 by p27Kip1. Mol Cell Biol 1995; 15: 4347–4352.
Eblen ST, Fautsch MP, Anders RA, Leof EB. Conditional binding
to and cell cycle-regulated inhibition of cyclin-dependent kinase
complexes by p27Kip1. Cell Growth Differ 1995; 6: 915–925.
Bastians H, Townsley FM, Ruderman JV. The cyclin-dependent
kinase inhibitor p27(Kip1) induces N-terminal proteolytic cleavage of cyclin A. Proc Natl Acad Sci USA 1998; 95: 15374–15381.
Zerfass-Thome K, Schulze A, Zwerschke W, Vogt B, Helin K, Bartek J, Henglein B, Jansen-Durr P. p27KIP1 blocks cyclin E-dependent transactivation of cyclin A gene expression. Mol Cell Biol
1997; 17: 407–415.
St Croix B, Florenes VA, Rak JW, Flanagan M, Bhattacharya N,
Slingerland JM, Kerbel RS. Impact of the cyclin-dependent kinase
inhibitor p27Kip1 on resistance of tumor cells to anticancer
agents. Nat Med 1996; 2: 1204–1210.
Reynisdottir I, Massague J. The subcellular locations of p15(Ink4b)
and p27(Kip1) coordinate their inhibitory interactions with cdk4
and cdk2. Genes Dev 1997; 11: 492–503.