REVIEW Molecular control of cell cycle progression in primary ... - Nature

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.


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


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-


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

1477


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. Baillie`res 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.


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


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, Lu¨bbert M, Said J, Sakashita A, Freyaldenhoven BS, Spira S, Huynh V, Mu¨ller 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.