CELL CYCLE MARKERS

Abstract

The present invention relates to nucleic acid reporter constructs and host cells transfected with said constructs. The invention also relates to methods which are useful for determining the cell cycle status of a mammalian cell and for determining the effect of a test agent on the cell cycle position of a mammalian cell.

Description

FIELD OF THE INVENTION

The present invention relates to G1/S cell cycle specific markers and methods for determining the transition between G1-phase and S-phase of the cell cycle in mammalian cells.

BACKGROUND OF THE INVENTION

Eukaryotic cell division proceeds through a highly regulated cell cycle comprising consecutive phases termed G1, S, G2 and M. Disruption of the cell cycle or cell cycle control can result in cellular abnormalities or disease states such as cancer which arise from multiple genetic changes that transform growth-limited cells into highly invasive cells that are unresponsive to normal control of growth. Transition of normal cells into cancer cells can arise though loss of correct function in DNA replication and DNA repair mechanisms. All dividing cells are subject to a number of control mechanisms, known as cell-cycle checkpoints, which maintain genomic integrity by arresting or inducing destruction of aberrant cells. Investigation of cell cycle progression and control is consequently of significant interest in designing anticancer drugs (Flatt, P. M. and Pietenpol, J. A. Drug Metab. Rev., (2000), 32(3-4), 283-305; Buolamwini, J. K. Current Pharmaceutical Design, (2000), 6, 379-392).

Accurate determination of cell cycle status is a key requirement for investigating cellular processes that affect the cell cycle or are dependent on cell cycle position. Such measurements are particularly vital in drug screening applications where:

• i) substances which directly or indirectly modify cell cycle progression are desired, for example, for investigation as potential anti-cancer treatments;
• ii) drug candidates are to be checked for unwanted effects on cell cycle progression; and/or
• iii) it is suspected that an agent is active or inactive towards cells in a particular phase of the cell cycle.

Traditionally, cell cycle status for cell populations has been determined by flow cytometry using fluorescent dyes which stain the DNA content of cell nuclei (Barlogie, B. et al, Cancer Res., (1983), 43(9), 3982-97). Flow cytometry yields quantitative information on the DNA content of cells and hence allows determination of the relative numbers of cells in the G1, S and G2+M phases of the cell cycle. However, this analysis is a destructive non-dynamic process and requires serial sampling of a population to determine cell cycle status with time.

Cell cycle progression is tightly regulated by defined temporal and spatial expression, localisation and destruction of a number of cell cycle regulators which exhibit highly dynamic behaviour during the cell cycle (Pines, J., Nature Cell Biology, (1999), 1, E73-E79). For example, at specific cell cycle stages some proteins translocate from the nucleus to the cytoplasm, or vice versa, and some are rapidly degraded. For details of known cell cycle control components and interactions, see Kohn, Molecular Biology of the Cell (1999), 10, 2703-2734.

U.S. Pat. No. 6,048,693 describes a method for screening for compounds affecting cell cycle regulatory proteins, wherein expression of a reporter gene is linked to control elements which are acted on by cyclins or other cell cycle control proteins. In this method, temporal expression of a reporter gene product is driven in a cell cycle specific fashion and compounds acting on one or more cell cycle control components may increase or decrease expression levels.

U.S. Pat. No. 6,159,691 relates to a method for assaying for putative regulators of cell cycle progression. In this method, nuclear localisation signals (NLS) derived from the cell cycle phase specific transcription factors DP-3 and E2F-1 are used to assay the activity of compounds which act as agonists or antagonists to increase or decrease nuclear localisation of an NLS fused to a detectable marker.

A number of researchers have studied the cell cycle using traditional reporter molecules that require the cells to be fixed or lysed. For example Hauser & Bauer (Plant and Soil, (2000), 226, 1-10) used β-glucuronidase (GUS) to study cell division in a plant meristem and Brandeis & Hunt (EMBO J., (1996), 15, 5280-5289) used chloramphenical acetyl transferase (CAT) fusion proteins to study variations in cyclin levels.

WO 03/031612 describes DNA reporter constructs and methods for determining the cell cycle position of living mammalian cells by means of cell cycle phase-specific expression control elements and destruction control elements.

None of the reports referred to above utilise cell cycle phase-dependent location control elements as a means of studying the cell cycle. However, Gu et al. (Mol Biol Cell., 2004, 15, 3320-3332) have recently investigated the function of human DNA helicase B (HDHB) and shown that it is primarily nuclear in G1 and cytoplasmic in S phase, that it resides in nuclear foci induced by DNA damage, that the focal pattern requires HDHB activity, and that HDHB localization is regulated by CDK phosphorylation.

None of the preceding methods specifically describe sensors which can be stably integrated into the genome and used to indicate G1 and S phases of the cell cycle. Consequently, methods are required that enable these phases of the cell cycle to be determined non-destructively in a single living mammalian cell, allowing the same cell to be repeatedly interrogated over time, and which enable the study of the effects of agents having potentially desired or undesired effects on the cell cycle.

SUMMARY OF THE INVENTION

The present invention describes a method which utilises key components of the cell cycle regulatory machinery in defined combinations to provide novel means of determining cell cycle status for individual living cells in a non-destructive process providing dynamic read out.

The present invention further provides DNA constructs, and stable cell lines containing such constructs, that exhibit translocation of a detectable reporter molecule in a cell cycle phase specific manner, by direct linkage of reporter signal switching to a G1/S cell dependent location control sequence.

This greatly improves the precision of determination of cell cycle phase status and allows continuous monitoring of cell cycle progression in individual cells. Furthermore, it has been found that key control elements can be isolated and abstracted from functional elements of the cell cycle control mechanism to permit design of cell cycle phase reporters which are dynamically regulated and operate in concert with, but independently of, endogenous cell cycle control components, thus providing the means for monitoring cell cycle position without influencing or interfering with the natural progression of the cell cycle.

According to a first aspect of the present invention, there is provided a nucleic acid reporter construct comprising a nucleic acid sequence encoding a detectable live-cell reporter molecule operably linked to and under the control of:

i) at least one cell cycle independent expression control element, and

ii) a G1/S cell cycle phase-dependent location control element; wherein the translocation of said reporter construct within a mammalian cell is indicative of the cell cycle position.

It will be understood that translocation is defined as the detectable movement of the reporter from one sub-cellular location to another, typically from the nucleus to the cytoplasm or vice versa. It will be further understood that the term ‘live cell’, as it relates to a reporter molecule, defines a reporter molecule which produces a detectable signal in living cells and is thus suitable for use in live-cell imaging systems, such as the IN Cell Analyzer (GE Healthcare).

The term, ‘operably linked’ indicates that the elements are arranged so that they function in concert for their intended purposes, e.g. transcription initiates in a promoter and proceeds through the DNA sequence coding for the reporter molecule of the invention.

Suitably, the expression control element controls transcription over an extended time period. Preferably, the expression control element is the ubiquitin C promoter which provides transcription over an extended period which is required for the production of stable cell lines.

Suitably, the cell cycle phase-specific dependent location control element is selected from the group consisting of Rag2, Chaf1B, Fen1, PPP1R2, helicase B, sgk, CDC6 or motifs therein such as the phosphorylation-dependent subcellular localization domain of the C-terminal special control region of helicase B (PSLD). Preferably, the phase-specific dependent location element is the phosphrylation-dependent subcellular localization domain of the C-terminal special control region of helicase B (PSLD).

A human helicase B homolog has been reported and characterised (Taneja et al J. Biol. Chem., (2002), 277, 40853-40861); the nucleic acid sequence for MGC clone NM 033647 and the corresponding protein are given in SEQ ID No. 1 and SEQ ID No. 2, respectively. The report demonstrates that helicase activity is needed during G1 to promote the G1/S transition. Gu et al (Mol. Biol. Cell., (2004), 15, 3320-3332) have shown that a small C-terminal region of the helicase B gene termed the phosphorylation-dependent subcellular localization domain (PSLD) is phosphorylated by Cdk2/cyclin E and contains NLS and NES sequences. Gu et al (Mol. Biol. Cell., (2004), 15, 3320-3332) carried out inhibitor-based studies (cells halted in G1 with mimosine etc, G2 with colchecine etc) on cells that had been transiently transfected with plasmid encoding an EGFP-BGal-PSLD fusion (beta-galactosidase was included in the construct as an inert group to make the whole fusion protein similar in size to the complete helicase B) expressed from a CMV promoter. Cells inhibited in G1 exhibited EGFP signal predominantly in the nucleus, whilst cells inhibited in other phases of the cell cycle exhibited predominantly cytoplasmic EGFP signal. These researchers concluded that the PSLD was directing translocation of the reporter from the nucleus to the cytoplasm at the G1/S phase of the cell cycle.

Suitably, the live-cell reporter molecule is selected from the group consisting of fluorescent protein, enzyme reporter and antigenic tag. Preferably, the fluorescent protein is selected from Green Fluorescent Protein (GFP) and a functional GFP analogue in which the amino acid sequence of wild type GFP has been altered by amino acid deletion, addition, or substitution. Preferably, the GFP is Enhanced Green Fluorescent Protein (EGFP), Emerald or J-Red.

Optionally, the enzyme reporter is halo-tag (Promega Corporation, USA).

Suitably, the cell cycle phase-dependent location control element is PSLD.

Preferably, the reporter molecule is a GFP and the cell cycle phase-dependent location control element is PSLD. More preferably, the reporter molecule is EGFP and the cell cycle phase-dependent location control element is PSLD.

Most preferably, the reporter construct comprises a CMV promoter, a PSLD and EGFP.

Preferably, the reporter construct comprises a human ubiquitin C promoter, a PSLD and a green fluorescent protein.

Suitably, the construct additionally comprises an inert group to increase the size of the expressed protein. The purpose of such a group is to allow the translated protein to be comparable in size to the ‘parent’ protein if, for example, only a portion of the protein has been used as the cell cycle phase-dependent location control element (e.g. only the PSLD domain of the complete helicase B protein).

For example, the inert group is βGal.

According to a second aspect of the present invention, there is provided a nucleic acid reporter construct comprising an expression vector comprising:

a) a vector backbone comprising:

i) a bacterial origin of replication; and

ii) a bacterial drug resistance gene;

b) a cell cycle independent expression control element; and

c) a G1/S cell cycle phase-dependent location control element; and

d) a nucleic acid sequence encoding a reporter molecule.

Optionally, the construct additionally contains a eukaryotic drug resistance gene, preferably a mammalian drug resistance gene.

Expression vectors may also contain other nucleic acid sequences, such as polyadenylation signals, splice donor/splice acceptor signals, intervening sequences, transcriptional enhancer sequences, translational enhancer sequences and the like. Optionally, the drug resistance gene and reporter gene may be operably linked by an internal ribosome entry site (IRES), which is cell cycle independent (Jang et al., J. Virology, (1988), 62, 2636-2643) rather than the two genes being driven by separate promoters. The pIRES-neo and pIRES vectors commercially available from Clontech may be used.

The construction and use of expression vectors and plasmids are well known to those of skill in the art. Virtually any mammalian cell expression vector may be used in connection with the cell cycle markers disclosed herein. Examples of suitable vector backbones which include bacterial and mammalian drug resistance genes and a bacterial origin of replication include, but are not limited to: pCI-neo (Promega), pcDNA (Invitrogen) and pTriEx1 (Novagen). Suitable bacterial drug resistance genes include genes encoding for proteins that confer resistance to antibiotics including, but not restricted to: ampicillin, kanamycin, tetracyclin and chloramphenicol. Eurkaryotic drug selection markers include agents such as: neomycin, hygromycin, puromycin, zeocin, mycophenolic acid, histidinol, gentamycin and methotrexate.

The DNA construct may be prepared by the standard recombinant molecular biology techniques of restriction digestion, ligation, transformation and plasmid purification by methods familiar to those skilled in the art and are as described in Sambrook, J. et al (1989), Molecular Cloning—A Laboratory Manual, Cold Spring Harbor Laboratory Press. Alternatively, the construct can be prepared synthetically by established methods, eg. the phosphoramidite method described by Beaucage and Caruthers, (Tetrahedron Letters, (1981), 22, 1859-1869) or the method described by Matthes et al (EMBO J., (1984), 3, 801-805). According to the phosphoramidite method, oligonucleotides are synthesised, eg. in an automatic DNA synthesizer, purified, annealed, ligated and cloned into suitable vectors. The DNA construct may also be prepared by polymerase chain reaction (PCR) using specific primers, for instance, as described in U.S. Pat. No. 4,683,202 or by Saiki et al (Science, (1988), 239, 487-491). A review of PCR methods may be found in PCR protocols, (1990), Academic Press, San Diego, Calif., U.S.A.

During the preparation of the DNA construct, the gene sequence encoding the reporter must be joined in frame with the G1/S cell cycle phase-dependent location control element. The resultant DNA construct should then be placed under the control of one or more suitable cell cycle phase independent expression control elements.

In a third aspect of the present invention, there is provided a polypeptide encoded by a nucleic acid construct as hereinbefore described.

In a fourth aspect of the present invention, there is provided the use of a polypeptide as hereinbefore described in the third aspect for determining the cell cycle position of a mammalian cell.

In a fifth aspect of the present invention, there is provided a host cell transfected with a nucleic acid construct as hereinbefore described. The host cell into which the construct or the expression vector containing such a construct is introduced may be any mammalian cell which is capable of expressing the construct.

The prepared DNA reporter construct may be transfected into a host cell using techniques well known to the skilled person. One approach is to temporarily permeabilise the cells using either chemical or physical procedures. These techniques may include: electroporation (Tur-Kaspa et al, Mol. Cell Biol. (1986), 6, 716-718; Potter et al, Proc. Nat. Acad. Sci. USA, (1984), 81, 7161-7165), a calcium phosphate based method (eg. Graham and Van der Eb, Virology, (1973), 52, 456-467 and Rippe et al, Mol. Cell Biol., (1990), 10, 689-695) or direct microinjection.

Alternatively, cationic lipid based methods (eg. the use of Superfect (Qiagen) or Fugene6 (Roche) may be used to introduce DNA into cells (Stewart et al, Human Gene Therapy, (1992), 3, 267; Torchilin et al, FASEB J, (1992), 6, 2716; Zhu et al, Science, (1993), 261, 209-211; Ledley et al, J. Pediatrics, (1987), 110, 1; Nicolau et al, Proc. Nat. Acad. Sci., USA, (1983), 80, 1068; Nicolau and Sene, Biochem. Biophys. Acta, (1982), 721, 185-190). Jiao et al, Biotechnology, (1993), 11, 497-502) describe the use of bombardment mediated gene transfer protocols for transferring and expressing genes in brain tissues which may also be used to transfer the DNA into host cells.

A further alternative method for transfecting the DNA construct into cells, utilises the natural ability of viruses to enter cells. Such methods include vectors and transfection protocols based on, for example, Herpes simplex virus (U.S. Pat. No. 5,288,641), cytomegalovirus (Miller, Curr. Top. Microbiol. Immunol., (1992), 158, 1), vaccinia virus (Baichwal and Sugden, 1986, in Gene Transfer, ed. R. Kucherlapati, New York, Plenum Press, p 117-148), and adenovirus and adeno-associated virus (Muzyczka, Curr. Top. Microbiol. Immunol., (1992), 158, 97-129).

Examples of suitable recombinant host cells include HeLa cells, Vero cells, Chinese Hamster ovary (CHO), U2OS, COS, BHK, HepG2, NIH 3T3 MDCK, RIN, HEK293 and other mammalian cell lines that are grown in vitro. Preferably the host cell is a human cell. Such cell lines are available from the American Tissue Culture Collection (ATCC), Bethesda, Md., U.S.A. Cells from primary cell lines that have been established after removing cells from a mammal followed by culturing the cells for a limited period of time are also intended to be included in the present invention.

In a preferred embodiment, the cell line is a stable cell line comprising a plurality of host cells according to the third aspect.

Cell lines which exhibit stable expression of a cell cycle position reporter may also be used in establishing xenografts of engineered cells in host animals using standard methods. (Krasagakis, K. J et al, Cell Physiol., (2001), 187(3), 386-91; Paris, S. et al, Clin. Exp. Metastasis, (1999), 17(10), 817-22). Xenografts of tumour cell lines engineered to express cell cycle position reporters will enable establishment of model systems to study tumour cell division, stasis and metastasis and to screen new anticancer drugs.

Use of engineered cell lines or transgenic tissues expressing a cell cycle position reporter as allografts in a host animal will permit study of mechanisms affecting tolerance or rejection of tissue transplants (Pye & Watt, J. Anat., (2001), 198 (Pt 2), 163-73; Brod, S. A. et al, Transplantation (2000), 69(10), 2162-6).

To perform the method for determining the cell cycle position of a cell, cells transfected with the DNA reporter construct may be cultured under conditions and for a period of time sufficient to allow expression of the reporter molecule at a specific stage of the cell cycle. Typically, expression of the reporter molecule will occur between 16 and 72 hours post transfection, but may vary depending on the culture conditions. If the reporter molecule is based on a green fluorescent protein sequence the reporter may take a defined time to fold into a conformation that is fluorescent. This time is dependent upon the primary sequence of the green fluorescent protein derivative being used. The fluorescent reporter protein may also change colour with time (see for example, Terskikh, Science, (2000), 290, 1585-8) in which case imaging is required at specified time intervals following transfection.

According to a sixth aspect of the present invention, there is provided a method for determining the cell cycle position of a mammalian cell by monitoring the expression of the reporter molecule and detecting signals emitted by the reporter using an appropriate detection device. If the reporter molecule produces a fluorescent signal, then, either a conventional fluorescence microscope, or a confocal based fluorescence microscope may be used. If the reporter molecule produces luminescent light, then a suitable device such as a luminometer may be used. Using these techniques, the proportion of cells expressing the reporter molecule may be determined. If the DNA construct contains translocation control elements and the cells are examined using a microscope, the location of the reporter may also be determined. In the method according to the present invention, the fluorescence of cells transformed or transfected with the DNA construct may suitably be measured by optical means in for example; a spectrophotometer, a fluorimeter, a fluorescence microscope, a cooled charge-coupled device (CCD) imager (such as a scanning imager or an area imager), a fluorescence activated cell sorter, a confocal microscope or a scanning confocal device, where the spectral properties of the cells in culture may be determined as scans of light excitation and emission.

In the embodiment of the invention wherein the nucleic acid reporter construct comprises a drug resistance gene, following transfection and expression of the drug resistance gene (usually 1-2 days), cells expressing the modified reporter gene may be selected by growing the cells in the presence of an antibiotic for which transfected cells are resistant due, to the presence of a selectable marker gene. The purpose of adding the antibiotic is to select for cells that express the reporter gene and that have, in some cases, integrated the reporter gene, with its associated promoter, into the genome of the cell line. Following selection, a clonal cell line expressing the construct can be isolated using standard techniques. The clonal cell line may then be grown under standard conditions and will express reporter molecule and produce a detectable signal at a specific point in the cell cycle.

Cells transfected with the nucleic acid reporter construct according to the present invention may be grown in the absence and/or the presence of a test agent to be studied and whose effect on the cell cycle of a cell is to be determined. By determining the proportion of cells expressing the reporter molecule and, optionally, the localisation of the signal within the cell, it is possible to determine the effect of the test agent on the cell cycle of the cells, for example, whether the test system arrests the cells in a particular stage of the cell cycle, or whether the effect is to speed up or slow down cell division.

Thus, according to a seventh aspect of the present invention, there is provided a method of determining the effect of a test agent on the cell cycle position of a mammalian cell, the method comprising:

• a) expressing in the cell in the absence and in the presence of the test agent a nucleic acid reporter construct as hereinbefore described; and
• b) determining the cell cycle position by monitoring signals emitted by the reporter molecule wherein a difference between the emitted signals measured in the absence and in the presence of the test agent is indicative of the effect of the test agent on the cell cycle position of the cell.

In a eighth aspect of the present invention, there is provided a method of determining the effect of a test agent on the cell cycle position of a mammalian cell, the method comprising:

• a) expressing in said cell in the presence of said test agent a nucleic acid reporter construct as hereinbefore described;
• b) determining the cell cycle position by monitoring signals emitted by the reporter molecule, and
• c) comparing the emitted signal in the presence of the test agent with a known value for the emitted signal in the absence of the test agent; wherein a difference between the emitted signal measured in the presence of the test agent and the known value in the absence of the test agent is indicative of the effect of the test agent on the cell cycle position of the cell.

In a ninth aspect of the present invention, there is provided a method of determining the effect of a test agent on the cell cycle position of a mammalian cell, the method comprising:

• a) providing cells containing a nucleic acid reporter construct according to the invention;
• b) culturing first and second populations of the cells respectively in the presence and absence of a test agent and under conditions permitting expression of the nucleic acid reporter construct; and
• c) measuring the signals emitted by the reporter molecule in the first and second cell populations; wherein a difference between the emitted signals measured in the first and second cell populations is indicative of the effect of the test agent on the cell cycle position of the cell.

According to a tenth aspect of the present invention, there is provided a method of determining the effect of the mammalian cell cycle on the expression, translocation or sub-cellular distribution of a first detectable reporter which is known to vary in response to a test agent, the method comprising:

• a) expressing in the cell in the presence of the test agent a second nucleic acid reporter construct according to the present invention;
• b) determining the cell cycle position by monitoring signals emitted by the second reporter molecule; and
• c) monitoring the signals emitted by the first detectable reporter, wherein the relationship between cell cycle position determined by step b) and the signal emitted by the first detectable reporter is indicative of whether or not the expression, translocation or sub-cellular distribution of the first detectable reporter is cell cycle dependent.

Suitably, the test agent is a form of electromagnetic radiation or is a chemical entity. Preferably, the test agent is a chemical entity selected from the group consisting of drug, nucleic acid, hormone, protein and peptide. Most preferably, the test agent is selected from a peptide or protein that is expressed in the cell under study.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 is the localisation of HDHB in the nucleus or cytoplasm.

(A) Cytoplasmic and nuclear extracts of U2OS cells were analyzed by denaturing gel electrophoresis and western blotting with antibody against recombinant HDHB, α-tubulin, and PCNA. Immunoreactive proteins were detected by chemiluminescence.

(B) GFP-HDHB transiently expressed in U2OS cells in contrast to endogenous HDHB. Control cells were transfected with pEGFP-C1 vector alone.

(C) GFP-tagged HDHB transiently expressed in microinjected U2OS cells were visualized by fluorescence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.

(D) FLAG-tagged HDHB transiently expressed in microinjected U2OS cells were visualized by fluorescence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.

(E) Walker A (MutA) of GFP-HDHB transiently expressed in microinjected U2OS cells were visualized by fluorescence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.

(F) Walker B mutants (MutB) of GFP-HDHB transiently expressed in microinjected U2OS cells were visualized by fluorescence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.(G) U2OS cells transiently expressing GFP-HDHB wt, MutA, and MutB were extracted with 0.5% Triton X-100 before fixation and fluoresence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.

FIG. 2 is the GFP-HDHB nuclear focus formation increases upon DNA damage.

(A) U2OS cells transiently expressing GFP-HDHB were treated with DMSO (control), 20 μM etoposide, 10 μM camptothecin, or 1 μM mitomycin C as indicated.

(B) The number of large distinct GFP-HDHB nuclear foci per cell in two independent assays is shown, with standard deviation in brackets.

FIG. 3 is the subcellular localization of GFP-HDHB is cell cycle-dependent.

(A) GFP-HDHB transiently expressed in U2OS cells in G1 or S phase was visualized by fluorescence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.

(B) The subcellular localization of GFP-tagged HDHB in asynchronous, G1, and S phase U2OS cells was quantified. The number of GFP-positive cells with a given distribution pattern was expressed as a percentage of the total number of GFP-positive cells in that experiment.

(C) Cytoplasmic and nuclear extracts of synchronized U2OS cells (G1 and S phase) were analyzed by denaturing gel electrophoresis and western blotting with antibody against recombinant HDHB, α-tubulin, and PCNA. Immunoreactive proteins were detected by chemiluminescence.

FIG. 4 is the identification of a domain required for nuclear localization of HDHB.

(A) Schematic representation of the HDHB protein showing seven potential phosphorylation sites for CDK (SP or TP), the putative subcellular localization domain (SLD) and phosphorylated SLD (PSLD), the Walker A and Walker B motifs. Amino acid residue numbers are indicated below the protein.

(B) GFP- and FLAG-tagged HDHB and their C-terminal truncation mutants are depicted. The C terminus of HDHB SLD and PSLD was fused to a GFP-âGal reporter to create GFP-βGal-SLD and GFP-βGal-PSLD respectively.

(C) GFP-HDHB-ΔSLD was transiently expressed in U2OS cells in G1 or S phase and visualized by fluorescence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.

(D) The subcellular localization of GFP-HDHB-ΔSLD in asynchronous, G1, and S phase U2OS cells was quantified and expressed as a percentage of the total number of GFP-positive cells in that experiment.

FIG. 5 is the GFP-βGal-PSLD subcellular localization pattern varies with the cell cycle. (A) GFP-βGal, GFP-βGal-SLD, and GFP-βGal-PSLD were transiently expressed in U2OS cells in G1 (left) and S phase (right) of the cell cycle. Nuclei were stained with Hoechst dye. Bar, 10 μm.

(B) The subcellular localization of GFP-βGal, GFP-βGal-SLD, and GFP-βGal-PSLD in asynchronous, G1, and S phase U2OS cells was quantified and expressed as a percentage of the total number of GFP-positive cells in that experiment.

FIG. 6 is the identification of a functional rev-type nuclear export signal (NES) in SLD of HDHB.

(A) Alignment of the putative NES in HDHB with those identified in other cell cycle-related proteins (Henderson and Eleftheriou, 2000; Fabbro and Henderson, 2003). Superscripts above the amino acid sequence indicate residue numbers. Thick arrows point to the conserved aliphatic residues in the NES. Two pairs of residues in the putative NES in HDHB were mutated to alanine as indicated by the thin arrows to create Mut1 and Mut2. (B) GFP- and FLAG-tagged HDHB were transiently expressed in asynchronously growing U2OS cells with (+) or without (−) LMB to inhibit CRM1-mediated nuclear export. The subcellular localization of GFP-HDHB and FLAG-HDHB in asynchronous, G1, and S phase cells was quantified and expressed as a percentage of the total number of GFP-positive cells in that sample. (C) GFP-HDHB and GFP-βGal-PSLD carrying the Mut1 or Mut2 mutations, or the corresponding proteins without the mutations, were transiently expressed in asynchronous U2OS cells and visualized by fluorescence microscopy. Cells showing the most frequently observed fluorescence pattern are shown. Nuclei were stained with Hoechst dye. Bar, 10 μm. (D) The subcellular localization of wild type and mutant GFP-HDHB and GFP-βGal-PSLD in asynchronous U2OS cells was quantified and expressed as a percentage of the total number of GFP-positive cells in that sample.

FIG. 7 is the cell cycle-dependent phosphorylation of FLAG-HDHB in vivo.

(A) U2OS cells transiently expressing FLAG-HDHB (lane 1) and its truncation mutants 1-1039 (lane 2) and 1-874 (lane 3) were labeled with [32P] ortho-phosphate. Cell extracts were immunoprecipitated with anti-FLAG resin. The precipitated proteins were separated by 7.5% SDS-PAGE, transferred to a PVDF membrane, and detected by autoradiography (top) or western blotting (bottom). The positions of marker proteins of known molecular mass are indicated at the left.

(B) FLAG-HDHB expressed in U2OS cells was immunoprecipitated with anti-FLAG resin, incubated with (+) or without (−) λ-phosphatase (λ-PPase) in the presence (+) or absence (−) of phosphatase inhibitors, as indicated, and analyzed by SDS-PAGE and immunoblotting with anti-HDHB antibody.

(C) U2OS cells expressing FLAG-HDHB were arrested at G1/S (top) or at G2/M (bottom), and then released from the block. FLAG-HDHB was harvested at the indicated time points, immunoprecipitated with anti-FLAG resin, treated with (+) or without (−) λ-PPase, and analyzed as in (B).

FIG. 8 is the identification of a major in vivo phosphorylation site in HDHB.

(A) Phosphoamino acid markers (left) and phosphoamino acids from in vivo 32P-labeled FLAG-HDHB (right) were separated in two dimensions and visualized by autoradiography. Some incompletely hydrolyzed phosphopeptides remained near the origin (+).

(B) Wild type and mutant FLAG-HDHB proteins were radiolabeled with orthophosphate in vivo, immunoprecipitated, separated by SDS-PAGE, and analyzed by autoradiography (top) and immunoblotting with anti-HDHB (bottom).

(C). Tryptic phosphopeptides of 32P-labeled wild type and S967A mutant FLAG-HDHB were separated in two dimensions and visualized by autoradiography.

FIG. 9 is the identification of cyclin E/CDK2 as the potential G1/S kinase of HDHB S967.

(A) Tryptic phosphopeptides from FLAG-HDHB phosphorylated in vivo as in FIG. 7C, or recombinant HDHB phosphorylated in vitro by purified cyclin E/CDK2 or cyclin A/CDK2, were separated in two dimensions, either individually or as a mixture, and visualized by autoradiography.

(B) Proteins that co-immunoprecipitated with FLAG vector (lanes 1, 4) or FLAG-HDHB (lanes 2, 5) expressed in U2OS cells were analyzed by immunoblotting with antibodies against HDHB (lanes 1-6), cyclin E (lanes 1-3), or cyclin A (lanes 4-6). One tenth of the cell lysate used for immunoprecipitation was analyzed in parallel as a positive control (lanes 3, 6).

FIG. 10 is the subcellular localization of HDHB is regulated by phosphorylation of S967.

(A) GFP fluorescence in U2OS cells transiently expressing GFP-HDHB with the S967A or S967D mutation was examined by fluorescence microscopy. Nuclei were stained with Hoechst dye. Bar, 10 μm.

(B) Subcellular localization of GFP-HDHB S967A and S967D expressed in asynchronous, G1, and S phase U2OS cells was quantified.

FIG. 11 is the vector map of pCORON1002-EGFP-C1-PSLD.

FIG. 12 is the vector map of pCORON1002-EGFP-C1-βGal-PSLD.

FIG. 13 is the EGFP-C1-PSLD transiently expressed in U2-OS cells. Image obtained on In Cell Analyzer 1000, 10× objective (Hoechst nuclear stain also shown to highlight a number of untransfected/low expressing cells).

FIG. 14 is the EGFP-C1-PSLD stably expressed in U2-OS cells. IN Cell Analyzer 1000, 10× objective.

FIG. 15 is the flow cytometry data comparing brightness and homogeneity of signal for representative stable cell lines developed with pCORON1002-EGFP-C1-PSLD, pCORON1002-EGFP-C1-Gal-PSLD and the parental U2OS cell line.

DETAILED DESCRIPTION OF THE INVENTION

Plasmids

pGFP-HDHB and mutant derivatives of it were created by inserting the full-length HDHB cDNA as a Bglll/NotI fragment (Taneja et al., J. Biol. Chem., (2002) 277, 40853-40861) into the NotI site of the pEGFP-C1 vector (Clontech, Palo Alto, Calif.). pFLAG-HDHB was constructed by inserting a HindIII/NotI fragment containing full-length HDHB cDNA into the NotI site of pFlag-CMV2 vector (Eastman Kodak Co., Rochester, N.Y.). Tagged HDHB-.SLD (1-1039) was constructed by cleaving the tagged HDHB plasmid with Nrul following the coding sequence for residue 1034 and with NotI in the polylinker and replacing the small fragment by a duplex adaptor oligonucleotide with a blunt end encoding residues 1035 to 1039, a stop codon, and an overhanging NotI-compatible 5′ end. To create pFLAG-HDHB (1-874), Stul-digested pFLAG-HDHB DNA was treated with Klenow polymerase to generate blunt ends and ligated into the pFLAG-CMV2 vector. To generate pEGFP-βGal, a DNA fragment encoding E. coli β-galactosidase (βGal) was amplified by PCR from the pβGal-control vector (Clontech) and inserted in frame at the 3′ end of the GFP coding sequence in pEGFP-C1, using the HindIII restriction site. The HDHB coding sequence for amino acids 1040-1087(SLD) and 957-1087(PSLD) were PCR amplified and inserted in frame at the 3′ end of the βGal cDNA in pEGFP-βGal to create pGFP-βGal-SLD and pGFP-βGal-PSLD respectively. The HDHB Walker A and Walker B mutants, MutA and MutB, were described previously (Taneja et al., J. Biol. Chem., (2002) 277, 40853-40861). The NES mutants and phosphorylation site mutants were created in the HDHB cDNA by site-directed mutagenesis (QuikChange, Stratagene, La Jolla, Calif.) according to the manufacturer's protocol using Pfu Turbo polymerase (Stratagene) and oligonucleotides containing the desired DNA sequence changes as primers in the PCR reactions.

The correct DNA sequence of the substitution mutations was confirmed by DNA sequencing.

pCORON1002-EGFP-C1-PSLD.

pCORON1002-EGFP-C1-PSLD was constructed by PCR amplification of the 390 bp PSLD region from the DNA construct pEGFP-Cl-Gal-PSLD (see above). Introduction of a NheI restriction enzyme site at the 5′ end and a SalI restriction enzyme site at the 3′ end of the PSLD fragment allowed for sub-cloning into the in house vector pCORON1002-EGFP-C1. The resulting 6704 bp DNA construct pCORON1002-EGFP-C1-PSLD, contains an ubiquitin C promoter, a bacterial ampicillin resistance gene and a mammalian neomycin resistance gene (FIG. 11). The nucleic acid sequence of the vector is shown in SEQ ID No. 3.

pCORON1002-EGFP-C1-Gal-PSLD.

pCORON1002-EGFP-C1-Gal-PSLD was constructed by NheI and XmaI restriction enzyme digest of the DNA construct pEGFP-Cl-Gal-PSLD (see above). The 4242 bp EGFP-C1-Gal-PSLD fragment was then ligated into the NheI and XmaI restriction enzyme digested pCORON1002 vector. The resulting 9937 bp DNA construct pCORON1002-EGFP-C1-Gal-PSLD, contains an ubiquitin C promoter, a bacterial ampicillin resistance gene and a mammalian neomycin resistance gene (FIG. 12). The nucleic acid sequence of the vector is shown in SEQ ID No. 4.

Antibodies

Anti-HDHB antibody was generated against purified recombinant HDHB (Bethyl Laboratories, Montgomery, Tex.) and affinity-purified on immobilized HDHB (Harlow & Lane, Antibodies: A laboratory manual. Cold Spring Harbor Laboratory). Initial characterization of these antibodies revealed that they were not ideal for indirect immunofluorescence or immunoprecipitation, but detected purified recombinant HDHB and endogenous HDHB in human cell extracts by western blotting.

Cell Culture, Synchronization, and Microinjection

U2OS cells were cultured as exponentially growing monolayers in Dulbecco-modified Eagle medium (DMEM) (Gibco BRL Lifetechnologies, Carlsbad, Calif.) supplemented with 10% fetal bovine serum (FBS) (Atlanta Biologicals, Norcross, Ga.) at 37° C. Exponentially growing U2OS cells were arrested at G1/S in incubation in DMEM containing 5 mM thymidine (Sigma-Aldrich, St. Louis, Mo.), for 24 h. To release the cells into S phase, the medium was aspirated and the cells washed three times with warm DMEM plus 10% FBS, and incubated in fresh DMEM plus 10% FBS. Exponentially growing U2OS cells were arrested in G2/M for 16 h in DMEM containing 30 ng/ml nocodazole (Sigma-Aldrich). To release cells into G1, mitotic cells were collected by gently shaking them off, washed three times with DMEM plus 10% FBS, and then plated on glass coverslips for microinjection, or in culture dishes for further manipulation.

Cell cycle synchronization was verified by flow cytometry as described previously (Taneja et al., J. Biol. Chem., (2002) 277, 40853-40861). In experiments to block nuclear protein export, cells were cultured for 3 h in DMEM containing 10 ng/ml of leptomycin B (LMB) (gift from Dr. M. Yoshida) and 10 μM cycloheximide (Calbiochem, San Diego, Calif.) to prevent new protein synthesis. Cells plated on glass coverslips were microinjected as described (Herbig et al., 1999) except that plasmid DNA rather than protein was injected.

Fluorescence Microscopy

For indirect immunofluorescence staining, cells were washed three times with phosphate buffered saline (PBS), fixed with 3.7% formaldehyde in PBS for 20 min, permeabilized for 5 min using 0.2% Triton X-100, and incubated with 10% FBS in PBS for 45 min. FLAG-HDHB was detected by staining with mouse monoclonal anti-FLAG antibody (Sigma-Aldrich) at a dilution of 1:100 in PBS plus 10% FBS for 2 h at room temperature. After washing, the cells were incubated with Texas Red-conjugated goat anti-mouse secondary antibody (Jackson ImmunoResearch Laboratories, West Grove, Pa.) at a dilution of 1:100 in PBS plus 10% FBS for 1 h at room temperature. After three washes, the cells were incubated for 10 min with Hoechst 33258 at a concentration of 2 μM in PBS. Coverslips were mounted in ProLong Antifade (Molecular Probes, Eugene, Oreg.). Images were obtained with a Hamamatsu digital camera using the Openlab 3.0 software (Improvision, Lexington, Mass.) on the Zeiss Axioplan 2 Imaging system (Carl Zeiss Inc.). The number of cells that exhibited each pattern of subcellular localization was counted and expressed as a percentage of the total number of cells scored (100 to 150 cells in each experiment). The subcellular distribution of each protein was quantitatively evaluated in at least two independent experiments.

For GFP fluorescence, cells were washed three times with phosphate-buffered saline (PBS), fixed with 3.7% formaldehyde in PBS containing 2 μM Hoechst 33258 for 20 min. Coverslips were mounted in ProLong Antifade (Molecular Probes) and fluorescent images were taken and evaluated as described above.

For Triton X-100 extraction, cells were washed twice with cold cytoskeleton buffer (CSK, 10 mM HEPES [pH 7.4], 300 mM sucrose, 100 mM NaCl, 3 mM MgCl2), and extracted for 5 min on ice with 0.5% Triton X-100 in CSK buffer (supplemented with 1× protease inhibitors) and then fixed as described above.

Flourescence microscopy was conducted using a confocal imaging system (In Cell Analyzer 1000, GE Healthcare, Amersham, UK) on cells transfected with the pCORON1002-EGFP-C1-PSLD or the pCORON1002-EGFP-C1-Gal-PSLD vectors.

DNA Damage Response Assay

U2OS cells (80-90% confluent) were transfected with pGFP-HDHB according to the manufacturer's protocol (Lipofectamine 2000, Invitrogen). At 4 h after transfection, cells were treated with DMSO (control), 20 μM etoposide, 10 μM camptothecin, or 1 μM mitomycin C. After 20 h, cells were extracted with Triton X-100 buffer and then fixed for immunofluorescence as described above. Distinctive GFP-HDHB nuclear foci were counted in more than 100 cells in each independent assay.

Electroporation

Asynchronously growing U2OS cells (5×10⁶) were trypsinized, collected by centrifugation, and resuspended in 800 μl of 20 mM HEPES (pH 7.4), 0.7 mM Na2HPO4/NaH2PO4, 137 mM NaCl, 5 mM KCl, 6 mM glucose at a final pH of 7.4. Ten μg of DNA was added and the mixture was transferred to a 0.4 cm electroporation cuvette (BioRad, Hercules, Calif.). Electroporation was performed using a Gene Pulser II apparatus and Gene Pulser II RF module (BioRad) at 300 V, 600 μF. Cells were then plated in tissue culture dishes, and 1 h later, washed with fresh medium and cultured for another 23 h.

Transfection with pCORON1002-EGFP-C1-PSLD Vector or the pCORON1002-EGFP-C1-Gal-PSLD Vector

U-2OS cells were transiently transfected with either the pCORON1002-EGFP-C1-PSLD vector (FIG. 11) or the pCORON1002-EGFP-C1-Gal-PSLD vector (FIG. 12) using FuGENE 6 (Roche Biochemicals). Various FuGENE 6: DNA ratios were investigated. Working with transiently transfected cells proved difficult due to low transfection efficiency and heterogeneity of expression.

U-2OS cells were transfected with either the pCORON1002-EGFP-C1-PSLD vector or the pCORON1002-EGFP-C1-Gal-PSLD vector using FuGENE 6. Stable clones expressing the recombinant fusion protein were selected using 1 mg/ml Geneticin G418 (Sigma). Isolated clones were chosen, FACS analysed, grown in culture and stocks frozen.

Metabolic Phosphate Labeling

U2OS cells (2.5×10⁶) were transiently transfected with wild type or mutant FLAGHDHB by electroporation. After 24 h, cells were incubated in phosphate-depleted DMEM (Gibco BRL Lifetechnologies) for 15 min and then radiolabeled with 32P-H3PO4 (0.35 mCi per ml of medium; ICN Pharmaceuticals Inc., Costa Mesa, Calif.) for 4 h. Phosphate-labeled FLAG-HDHB was immunoprecipitated from extracts, separated by 7.5% SDS/PAGE, and then transferred to a polyvinylidene difluoride (PVDF) membrane as described below.

Cell Extracts, Immunoprecipitation, and Western Blotting

At 24 h after transfection, FLAG-HDHB-transfected cultures to be analyzed by immunoprecipitation and immunoblotting were lysed in lysis buffer (50 mM Tris-HCl pH 7.5, 10% glycerol, 0.1% NP-40, 1 mM DTT, 25 mM NaF, 100 μg/ml PMSF, 1 μg/ml aprotinin, 1 μg/ml leupeptin) (0.5 ml per 35 mm or 1 ml per 60 mm dish or 75 cm flask). The extract was scraped off the dish, incubated for 5 min on ice, and centrifuged for 10 min at 14 000 g. Samples of the supernatant (0.5 to 1 mg of protein) were incubated with 10 μl anti-FLAG agarose (Sigma) on a rotator for 2 h at 4° C. The agarose beads were washed three times with lysis buffer. Immunoprecipitated proteins were transferred to a PVDF membrane and analyzed by western blotting with anti-HDHB-peptide serum (1:5000), anti-cyclin E antibody (1:1000), and anticyclin A antibody (1:1000) (Santa Cruz Biotechnology Inc., Santa Cruz, Calif.), and chemiluminescence (SuperSignal, Pierce Biotechnology Inc., Rockford, Ill.).

For selective nuclear and cytoplasmic protein extraction, 80-90% confluent U2OS cells were harvested by trypsinization and washed with PBS. They were resuspended and lysed in 10 mM Tris-HCl [pH 7.5], 10 mM KCl, 1.5 mM MgCl2, 0.25 M sucrose, 10% glycerol, 75 μg/ml digitonin, 1 mM DTT, 10 mM NaF, 1 mM Na3VO4, 100 μg/ml PMSF, 1 μg/ml aprotinin, and 1 μg/ml leupeptin for 10 min on ice, and centrifuged at 1000×g for 5 min. The supernatant fraction was collected as the cytosolic extract. The pellet washed, resuspended in high salt buffer (10 mM Tris-HCl [pH 7.5], 400 mM NaCl 1 mM EDTA, 1 mM EGTA, 1 mM DTT, 1% NP-40, 100 μg/ml PMSF, 1 μg/ml aprotinin, and 1 μg/ml leupeptin), and rocked for 10 min at 4° C. After sonication, the suspended material, containing both soluble and chromatin-bound protein, was analyzed as nuclear extract. Proteins in the nuclear and cytoplasmic extracts were analyzed by 8.5% SDS-PAGE, followed by western blotting with antibodies against α-tubulin, PCNA (both Santa Cruz Biotechnology), and recombinant HDHB.

Protein Phosphatase Reactions

FLAG-HDHB bound to anti-FLAG beads was incubated with 100 U of λ-phosphatase (New England Biolabs, Beverly, Mass.) in phosphatase buffer (50 mM Tris-HCl [pH 7.5], 0.1 mM EDTA, 0.01% NP-40) for 1 h at 30° C. The reaction was carried out in the presence or absence of phosphatase inhibitors (5 mM Na3VO4, 50 mM NaF). The proteins were separated by 7.5% SDSPAGE (acrylamide-bisacrylamide ratio, 30:0.36) and HDHB was detected by western blotting with anti-HDHB-peptide serum and chemiluminescence.

Tryptic Peptide Mapping and Phosphoamino Acid Analysis

At 24 h after transfection, radiolabeled FLAG-HDHB-transfected cultures to be used for immunoprecipitation and phosphoamino acid or phosphopeptide mapping were processed as above, except that lysis buffer was substituted by RIPA buffer (50 mM Tris-HCl [pH7.5], 150 mM NaCl, 1% NP-40, 0.5% deoxycholic acid, 1% SDS, 50 mM NaF, 1 mM EDTA, 5 mM Na3VO4, 100 μg/ml PMSF, 1 μg/ml aprotinin, and 1 μg/ml leupeptin). Immunoprecipitated proteins were separated by 7.5% SDS-PAGE and transferred to PVDF membranes. The membranes containing radiolabeled HDHB were rinsed well with deionized H2O twice before visualization of phosphoproteins by autoradiography. The phosphoproteins were then excised, and the membrane pieces were re-wet with methanol followed by water. The membranes were blocked with 50 mM NH4HCO3 containing 0.1% Tween 20 (Sigma-Aldrich) for 30 min at room temperature and washed three times with 50 mM NH4HCO3 before enzymatic cleavage of phosphoproteins from the PVDF with L-(tosylamido-2-phenyl)ethyl chloromethyl ketonetreated bovine pancreatic trypsin (Worthington, Lakewood, N.J.). The peptides were then subjected to two-dimensional phosphopeptide mapping or phosphoamino acid analysis as described in detail elsewhere (Boyle et al., Meth. Enzymology, (1991), 201, 110-149).

Cyclin-Dependent Kinase Reactions In Vitro

Kinase reactions using purified cyclin/CDK (200 pmol/h) (provided by R. Ott and C. Voitenleitner) and purified recombinant HDHB (Taneja et al., J. Biol. Chem., (2002) 277, 40853-40861) as the substrate were performed as described previously (Voitenleitner et al., Mol. Cell. Biol., (1999), 19, 646-56).

HDHB Resides in Nuclear Foci or in the Cytoplasm

To determine the subcellular localization of endogenous HDHB, nuclear and cytoplasmic proteins were selectively extracted from human U2OS cells, separated by denaturing gel electrophoresis, and analyzed by western blotting (FIG. 1A). The presence of PCNA and α-tubulin in each extract was first monitored to assess the extraction procedure. PCNA was enriched in the nuclear extract and not in the cytoplasmic fraction, while α-tubulin was found primarily in the cytoplasmic fraction, validating the fractionation. HDHB was detected in both the nuclear and cytoplasmic fractions (FIG. 1A). The cytoplasmic HDHB migrated more slowly than the nuclear fraction (FIG. 1A), suggesting the possibility of post-translational modification.

These results could indicate either that HDHB was distributed throughout the cell, or that a mixed population of cells contained HDHB in either the nucleus or the cytoplasm. To distinguish between these alternatives, HDHB was localized in situ in single cells. Since endogenous HDHB was not detectable by indirect immunofluorescence with antisera (data not shown), GFP- and FLAG-tagged HDHB were expressed in human U2OS cells by transient transfection. Transiently over-expressed tagged HDHB accumulated in greater amounts than the endogenous HDHB within 24 h (FIG. 1B). Since prolonged over-expression of tagged or untagged HDHB was cytotoxic, all experiments were conducted in the shortest time period possible (usually 24 h). Tagged HDHB localization was analyzed in individual cells by fluorescence microscopy. Both GFP-HDHB and FLAG-HDHB displayed two major patterns of localization, either in the nucleus in discrete foci or in the cytoplasm (FIG. 1C, D). The localization patterns of HDHB tagged with the GFP protein were not detectably different than those of HDHB tagged with the FLAG peptide. GFP-HDHB transiently expressed in primary human fibroblasts was also observed in either the nucleus or the cytoplasm (data not shown).

To test whether the subcellular localization of HDHB depended on its biochemical activity, the conserved lysine of the Walker A motif in GFP-HDHB was substituted by alanine (MutA) or the conserved glutamate of the Walker B motif was replaced by glutamine (MutB), crippling HDHB helicase activity (Taneja et al., J. Biol. Chem., (2002), 277, 40853-40861). Although DNA polymerase alphaprimase associated with both mutants, ATP stimulated single-stranded DNA binding of MutB, but not MutA (Taneja et al., J. Biol. Chem., (2002), 277, 40853-40861). These mutant forms of GFP-HDHB accumulated either in the nucleus, sparing the nucleoli, or in the cytoplasm of transfected cells, with few cells showing GFP-HDHB in both compartments (FIG. 1E, F). Interestingly, nuclear foci of GFP-HDHB were not obvious with the Walker A mutant (FIG. 1E). The focal pattern of the Walker B mutant was more variable, with some cells resembling the Walker A pattern (FIG. 1F, left panels) and others containing a small number of foci less prominent than in cells expressing wild type GFP-HDHB (FIG. 1F, middle panels). Interestingly, when the cells were extracted with detergent before fixation to remove soluble HDHB, GFP-HDHB wt maintained the distinct focal staining pattern, while MutB displayed a more obvious focal staining pattern, but with many smaller foci than those of the wild type (FIG. 1G). MutA staining was lost, indicating that MutA was largely soluble and not bound to nuclear structures. The results suggest that nucleotide binding of HDHB is required for nuclear focus formation and that the foci are bound to detergent-insoluble nuclear structures.

GFP-HDHB Nuclear Focus Formation Increases Upon DNA Damage

Since HDHB localization in nuclear foci depends on its biochemical activity, it is likely that HDHB executes its function in those nuclear foci. The specificity of HDHB for DNA as a substrate and its sequence homology with bacterial RecD and T4 dda proteins (Taneja et al., J. Biol. Chem., (2002) 277, 40853-40861) suggests that HDHB might be involved in DNA damage signaling, processing, or repair. Moreover, DNA damage induces nuclear foci that are thought to contain damage-sensing and -processing proteins (Nelms et al., Science, (1998) 280, 590-592; van den Bosch et al. EMBO Rep., (2003), 4, 844-849). To test whether HDHB may reside in DNA damage foci, U2OS cells transiently expressing GFP-HDHB were treated with the DNA damaging agents etoposide, camptothecin, and mitomycin C, or with DMSO as a control (FIG. 2).

The number of large GFP-HDHB nuclear foci per cell more than doubled upon exposure to etoposide and camptothecin compared to the control (FIG. 2), strongly suggesting that HDHB was recruited to foci generated by DNA damage. The observed increase in foci was dose-dependent (not shown). Mitomycin C treatment had little effect on GFP-HDHB nuclear foci, suggesting that only specific types of DNA damage may attract HDHB.

Identification of a Cell Cycle-Dependent Subcellular Localization Domain in HDHB

The ability of Walker A and Walker B mutants of HDHB to inhibit the onset of S phase (Taneja et al., 2002), together with the nuclear or cytoplasmic localization of tagged HDHB (FIG. 1), raised the question of whether the subcellular localization of HDHB might be cell cycle dependent. To test this possibility, U2OS cells were arrested in G2/M with nocodazole, released into G1 for three hours, and then microinjected pGFP-HDHB DNA into their nuclei. GFP-HDHB expression was easily detectable six hours later, when approximately 70% of G1 phase cells had accumulated the fusion protein primarily in the nuclei (FIG. 3A, B). In contrast, when cells were synchronized at G1/S with thymidine, released into S phase, and then microinjected with pGFP-HDHB DNA, more than 70% of S phase cells had accumulated the fusion protein predominantly in the cytoplasm (FIG. 3A, B). Selective extraction of U2OS cells in G1 and S phase revealed that endogenous HDHB was mostly nuclear in G1 and cytoplasmic in S phase (FIG. 3C). However, endogenous HDHB was clearly detectable in both subcellular fractions. The mobility of the S phase HDHB was slightly retarded compared to the G1 phase protein. These results indicate that the subcellular localization of HDHB is regulated in the cell cycle and that GFP-tagged HDHB reflects the localization of the endogenous untagged helicase.

There are two primary mechanisms to target a protein to the nucleus or cytoplasm in a cell cycle-dependent manner. One is that the protein carries its own nuclear location signal (NLS) and/or nuclear export signal (NES), motifs that are recognized by nuclear import or export machinery (Gorlich & Kutay, Rev. Cell Dev. Biol., (1999), 15, 607-660; Hood & Silver, Biochim. Biophys. Acta., (2000), 1471, M31-M41; Weis, Cell, (2003), 112, 441-451; Fabbro & Henderson, Exp. Cell. Res., (2003) 282, 59-69). Another is that the protein lacks a targeting signal, but can bind to another protein that has a NES and/or NLS. Prompted by the identification of C-terminal nuclear location signals in Bloom's syndrome helicase and other RecQ-family helicases (Hickson, Nature Rev. Cancer, (2003) 3, 169-178), a possible subcellular localization domain (SLD) was identified at the extreme C-terminus of HDHB (FIG. 4A). To determine whether this putative SLD was important for HDHB localization, a truncation mutant of HDHB (GFP-HDHB-.SLD) was generated that lacks the C-terminal 48 residues containing the SLD (FIG. 4B). The expression vector was microinjected into U2OS cells in G1 or S phase and the subcellular localization of the fusion protein was examined by fluorescence microscopy six hours later (FIG. 4C). Over 95% of the cells accumulated the fusion protein in the cytoplasm, regardless of the cell cycle timing of HDHB expression (FIG. 4C, D). This result suggests that HDHB may carry a NLS that is impaired or abolished by the C-terminal deletion in GFP-HDHB-ΔSLD.

To determine whether the C-terminal domain of HDHB was sufficient for nuclear localization, a bacterial β-galactosidase (βGal) was used as a reporter protein because it has a molecular mass (112 kDa) close to that of HDHB and does not contain subcellular localization signals (Kalderon et al., Cell, (1984), 39, 499-509). As a control, we created a GFP-βGal expression vector (FIG. 4B) and monitored the subcellular localization of the fusion protein after microinjection of the expression vector into U2OS cells. As expected, GFP-βGal protein accumulated primarily in the cytoplasm (FIG. 5). In contrast, GFP-βGal-SLD was found in both the nucleus and cytoplasm in asynchronous or synchronized U2OS cells (FIG. 5), suggesting that SLD contains a NLS, but was not sufficient for nuclear localization of the reporter protein. Reasoning that perhaps the neighboring potential CDK phosphorylation sites might affect subcellular localization in the cell cycle (FIG. 4A), a GFP-βGal-PSLD was constructed, in which the C-terminal 131 residues of HDHB, containing the putative SLD and the cluster of potential CDK phosphorylation sites, were appended to the C-terminus of GFP-βGal (FIG. 4B). When GFP-βGal-PSLD plasmid DNA was transiently expressed in asynchronous and synchronized U2OS cells, GFP-βGal-PSLD was found in the nucleus in over 90% of G1 phase cells, and in the cytoplasm in more than 70% of S phase cells (FIG. 5). In contrast with the focal pattern observed for nuclear GFP-HDHB in G1, GFP-βGal-PSLD protein was distributed evenly throughout the nucleus in G1, sparing only the nucleoli (FIG. 5). Taken together, these data suggested that the subcellular localization of HDHB is dependent on the cell cycle, and that the C-terminal PSLD domain of HDHB plays a major role in regulating the subcellular localization of the protein.

Identification of a Functional Rev-Type NES in HDHB

A number of proteins that shuttle between the nucleus and cytoplasm have been demonstrated to contain a NES similar to the prototype NES of HIV rev protein (FIG. 6A). Proteins containing a rev-type NES require the export factor CRM1 (also called exportin 1) to bind and transport proteins from the nucleus to the cytoplasm (reviewed by Weis, Cell, (2003), 112, 441-451). A fungal metabolite, leptomycin B (LMB), specifically inhibits CRM1 activity in nuclear protein export (Wolff et al., Chem. Biol., (1997), 4, 139-147; Kudo et al., Exp. Cell. Res., (1998), 242, 540-547. Inspection of the PSLD sequence in HDHB revealed a putative rev-type NES (LxxxLxxLxL, where L can also be I, V, M, or F, and x is any amino acid) (FIG. 6A). To determine whether the cytoplasmic localization of HDHB requires a functional NES, expression plasmids for GFP-HDHB or FLAG-HDHB DNA were microinjected into asynchronous, G1, and S phase cells in the presence and absence of LMB. The localization of the fusion proteins was examined by fluorescence microscopy and quantified. In the presence of LMB, both fusion proteins accumulated in the nucleus independently of the cell cycle (FIG. 6B and data not shown), consistent with the possibility that HDHB contains a rev-type NES that functions through CRM1. However it is also possible that HDHB may not be a direct cargo of CRM1 and that its export may be indirectly mediated through some other protein(s). To assess whether the putative NES in HDHB was functional, we mutated Val/Leu and Leu/Leu of the NES motif to alanine to create NES mutants 1 and 2 (FIG. 6A). GFP-HDHB and GFP-βGal-PSLD harboring these NES mutations were transiently expressed in either asynchronous or synchronized U2OS cells. Both NES mutant fusion proteins accumulated in the nucleus in more than 80% of cells, no matter when they were expressed in asynchronous or synchronized cells (FIG. 6C, D; data not shown). The focal pattern of nuclear GFP-HDHB was somewhat more diffuse with the NES mutants than with wild type GFP-HDHB (compare FIG. 6C with FIG. 1B). The results indicate that the NES mutations specifically impaired the export of both GFP-HDHB and GFP-βGal-PSLD, arguing that the PSLD region of HDHB contains a functional NES.

FLAG-HDHB is Phosphorylated in a Cell Cycle-Dependent Manner In Vivo.

The cluster of potential CDK phosphorylation sites in the PSLD domain of HDHB (FIG. 4A) suggested that phosphorylation of HDHB might regulate its subcellular localization in the cell cycle. If so, one would expect the PSLD region of HDHB to be phosphorylated in a cell cycle-dependent manner. To first test whether HDHB undergoes phosphorylation in PSLD, U2OS cells were transiently transfected with expression plasmids for wild type and C-terminally truncated forms of FLAG-HDHB, radiolabeled with phosphate, and then FLAG-HDHB was immunoprecipitated from cell extracts. Immunoprecipitated proteins were analyzed by denaturing gel electrophoresis, immunoblotting, and autoradiography (FIG. 7A). A radiolabeled band of FLAG-HDHB was detected at the same position as the immunoreactive HDHB band (FIG. 7A, lanes 1). Truncated FLAG-HDHB lacking SLD was also robustly phosphorylated in vivo (lanes 2), while truncated FLAG-HDHB (1-874) lacking PSLD was not significantly phosphorylated (lanes 3). These results demonstrate that SLD is not required for HDHB phosphorylation, while PSLD is required, and suggest that the phosphorylation sites probably reside in PSLD.

To examine the timing of HDHB phosphorylation in the cell cycle, it would be convenient to detect phosphorylation without the use of radiolabeling. Since phosphorylation often reduces the electrophoretic mobility of a protein in denaturing gels, transiently expressed FLAG-HDHB was immunoprecipitated and its mobility examined before and after treatment with λ-phosphatase (λ-PPase) (FIG. 7B). Without λ-PPase treatment, FLAG-HDHB was detected in western blots in two very closely migrating bands (lane 1), while dephosphorylated FLAG-HDHB migrated as a single band at the mobility of the faster band of the doublet (lane 2). When λ-PPase inhibitors were present in the reaction, FLAG-HDHB migrated as a doublet identical to the mock-treated protein (lane 3). These data suggest that the electrophoretic mobility of FLAG-HDHB was reduced by phosphorylation and that this assay may be suitable to track HDHB phosphorylation in the cell cycle.

To determine whether HDHB is phosphorylated in a cell cycle-dependent manner, U2OS cells transiently expressing FLAG-HDHB were arrested in G1/S by adding thymidine to the medium or in G2/M by adding nocodazole to the medium. The cells were released from the blocks for different time periods, and FLAG-HDHB was immunoprecipitated from cell extracts.

The immunoprecipitated material was incubated with or without λ-PPase and then analyzed by denaturing gel electrophoresis and western blotting (FIG. 7C). The mobility of FLAG-HDHB from cells arrested at G1/S was increased by λ-PPase treatment, suggesting that the protein was phosphorylated at G1/S (FIG. 7C, upper panel). A similar mobility shift was detected after phosphatase treatment of FLAG-HDHB for at least nine hours after release from the G1/S block (upper panel), as well as in cells arrested at G2/M (FIG. 7C, lower panel). However, after the cells were released into G1 for four and eight hours, FLAG-HDHB migrated as a single band that was much less affected by phosphatase treatment (FIG. 7C, lower panel). By twelve hours after release from the G2/M block, when most of the cells were entering S phase (data not shown), the mobility of FLAG-HDHB was again increased by phosphatase treatment, restoring the pattern observed in nocodazole-arrested cells (lower panel). These results strongly suggest that phosphorylation of FLAG-HDHB is cell cycle-dependent, with maximal phosphorylation from G1/S through G2/M and minimal phosphorylation during G1.

Serine 967 is the Major Phosphorylation Site of Ectopically Expressed HDHB.

To map the phosphorylation sites in FLAG-HDHB, we first wished to determine what amino acid residues were modified. Phosphoamino acid analysis of in vivo radiolabeled FLAG-HDHB revealed that phosphoserine(s) was the major phosphoamino acid of FLAG-HDHB in vivo (FIG. 8A). Assuming that the cell cycle-dependent phosphorylation sites of HDHB are located in PSLD between residues 874 and 1039 (FIG. 7A), that these sites are modified by CDKs, and that phosphoserine is the major amino acid modified (FIG. 8A), only four of the seven potential CDK sites would remain as candidate sites. To test each of these sites individually, FLAG-HDHB expression plasmids with the corresponding serine to alanine mutations were constructed. Cells transiently transfected with these plasmids were radiolabeled with orthophosphate in vivo and FLAG-HDHB was immunoprecipitated and analyzed by autoradiography and western blotting (FIG. 8B). The results showed that FLAG-HDHB and three of the mutant proteins were phosphorylated approximately equally, while the S967A mutant protein was only weakly phosphorylated (FIG. 8B). This result suggested that S967 might be the primary site of HDHB phosphorylation in vivo. Consistent with this interpretation, an electrophoretic mobility shift after phosphatase treatment of immunoprecipitated FLAG-HDHB was detected with three of the mutant proteins, but not with S967A protein (data not shown).

To confirm that S967 was the major phosphorylation site in HDHB in vivo, tryptic phosphopeptide mapping was carried out with wild type and S967A mutant FLAG-HDHB that had been metabolically radiolabeled with orthophosphate (FIG. 8C). One predominant radiolabeled peptide and a weakly labeled peptide were observed with the wild type protein (left panel). The predominant phosphopeptide was absent in the S967A protein, but the weakly labeled peptide remained detectable (FIG. 8C, right panel). The results provide additional evidence that serine 967 is a prominent phosphorylation site in HDHB in vivo.

Identification of Cyclin E/CDK2 as a Kinase that Potentially Modifies HDHB in G1/S

To test whether CDKs can actually modify HDHB, as suggested by the timing of HDHB phosphorylation in the cell cycle and the identification of S967 as a primary site of modification, purified cyclin E/CDK2 or cyclin A/CDK2 were incubated with purified recombinant HDHB and radiolabeled ATP in vitro. After the kinase reactions, the proteins were separated by denaturing gel electrophoresis, transferred to a PVDF membrane, and detected by autoradiography. The results revealed that recombinant HDHB could be phosphorylated strongly by both cyclin E/CDK2 and cyclin A/CDK2 (data not shown). The radiolabeled HDHB bands were then further processed for tryptic phosphopeptide mapping. Peptides from each digestion were separated in two dimensions, either individually or after mixing with tryptic peptides from in vivo phosphorylated FLAG-HDHB, and visualized by autoradiography (FIG. 9A). HDHB peptides phosphorylated by cyclin E/CDK2 and cyclin A/CDK2 yielded patterns essentially identical to those observed in the in vivo labeled peptide map, with one major spot and one minor spot (FIG. 9A). When the in vitro and in vivo labeled peptides were mixed and separated on one chromatogram, they co-migrated (FIG. 9A, right). These data argue that the major phosphopeptides modified in vitro by cyclin E/CDK2 and cyclin A/CDK2 in purified recombinant HDHB were the same ones modified in vivo in FLAG-HDHB.

Since cyclin E activity in human cells rises in late G1, while cyclin A activity rises later coincident with the onset of S phase (Pines, 1999; Erlandsson et al., 2000), it was important to try to distinguish whether one of these kinases might preferentially modify HDHB. Cyclin subunits frequently form a complex with the substrate proteins that they target for phosphorylation (Endicott et al., 1999; Takeda et al., 2001). To test whether cyclin E or cyclin A could associate with HDHB, FLAG-HDHB and associated proteins were immunoprecipitated from extracts of cells transfected with either FLAG-HDHB expression vector or empty FLAG vector as a control. The cell extracts and the immunoprecipitated material were analyzed by western blotting (FIG. 9B). Cyclin E clearly co-precipitated with FLAG-HDHB, but cyclin A did not (FIG. 9B, lanes 2 and 5), suggesting that FLAG-HDHB may interact preferentially with cyclin E in vivo. It is conceivable that this interaction may be required for phosphorylation of HDHB by cyclin E/CDK2 in vivo, and if so, mutations in HDHB that prevent its association with cyclin E would abrogate phosphorylation by cyclin E/CDK2. To test the possibility that the FLAG-HDHB mutant S967A was not phosphorylated in vivo (FIG. 8B, C) due to an inability to bind to cyclin E, FLAG-HDHB-S967A and associated proteins were immunoprecipitated from extracts of transfected cells and analyzed by western blotting. Co-precipitation of cyclin E with the mutant protein was as robust as with wild type FLAG-HDHB (data not shown).

Phosphorylation of Serine 967 is Critical for Regulation of HDHB Localization.

The data above indicate that subcellular localization and phosphorylation of ectopically expressed HDHB were regulated in a cell cycle-dependent manner with maximal phosphorylation of HDHB from G1/S to G2/M, coinciding with the period when HDHB accumulated in the cytoplasm. These results, together with the identification of S967 as the major in vivo phosphorylation site in HDHB, suggest that phosphorylation of S967 may regulate the subcellular localization of HDHB. To test this idea, expression plasmids for wild type GFP-HDHB and the mutants S967A, S984A, S1005A, and S1021A were microinjected into synchronized U2OS cells. Wild type GFP-HDHB accumulated in nuclear foci of cells in G1, but in the cytoplasm of cells in S phase as expected (not shown). However, regardless of cell cycle timing, GFP-HDHB-S967A localized in nuclear foci in about 70% of the fluorescent cells (FIG. 9A, B). The other three substitution mutants localized in either the nucleus or the cytoplasm like wild type GFP-HDHB (data not shown). In an attempt to mimic the phosphorylation of S967, serine 967 was mutated to aspartic acid, GFP-HDHB-S967D was expressed in asynchronous and synchronized U2OS cells, and the subcellular distribution of the mutant fusion protein was examined.

About 60% of the cells expressing GFP-HDHB-S967D displayed cytoplasmic fluorescence in asynchronous, G1 phase, and S phase cells (FIG. 10A, B), demonstrating that the S967D mutation mimicked phosphorylated S967. The data strongly suggest that phosphorylation of serine 967 is critical in regulating the subcellular localization of HDHB.

HDHB Resides in Nuclear Foci Inducible by DNA Damage

During G1 of the cell cycle, wild type GFP-HDHB resides in prominent nuclear foci that are associated with detergent-insoluble nuclear structures (FIG. 1, 2, 3). The topoisomerase inhibitors etoposide and camptothecin induced many more HDHB nuclear foci, suggesting that the helicase activity may be recruited to process sites of DNA damage (FIG. 2). Many proteins involved in DNA double strand break repair and recombination show a similar focal relocalization in response to DNA damage at different times in the cell cycle, including Rad52, RPA, Mre11, Ddc1 Ddc2, Rad9, Rad24, Rad51, Rad53, Rad54, and Rad55 (e.g. Lisby et al., 2003). In support of this notion, one of the helicase-defective point mutants (MutA) did not associate with detergent-insoluble foci (FIG. 1), indicating that focus formation depended on the biochemical activity of HDHB. Also consistent with this interpretation, the PSLD peptide of HDHB, which lacks the helicase domain (FIG. 4A), directed a βGal reporter to the nucleus during G1, but did not produce the focal localization pattern (FIG. 5). Intriguingly, MutB also localized in a focal pattern in a detergent-resistant manner, indicating that the ability of HDHB to bind to single-stranded DNA in the presence in ATP may be sufficient for the focal localization. These results suggest that the main function of HDHB may be to process endogenous DNA damage, possibly caused by incomplete topoisomerase reactions, during G1 phase of the cell cycle. This interpretation provides a simple explanation for the ability of helicase-defective HDHB mutants to block G1/S progression when the mutant protein is injected into cells in early G1 (Taneja et al., J. Biol. Chem., (2002), 277, 40853-40861). The MutB form of HDHB would associate with the damage, but could not process it properly, interfering with the activity of the endogenous helicase and leading to cell cycle arrest in late G1.

Another possible function considered for HDHB was DNA replication (Taneja et al., J. Biol. Chem., (2002), 277, 40853-40861). However, the predominantly cytoplasmic localization of HDHB during S and G2/M argues that HDHB is probably not directly involved in genomic DNA replication (FIG. 3). Consistent with this idea, the helicase-defective HDHB did not affect DNA synthesis when injected into the nucleus of cells in late G1 or S phase (Taneja et al., 2002), probably because the injected protein was phosphorylated by CDK2 associated with cyclins E or A and rapidly targeted to the cytoplasm. Indeed, export to the cytoplasm may serve as a means to sequester HDHB when its function is not needed or might be detrimental in the nucleus. HDHB activity is not affected by CDK phosphorylation in vitro (Taneja and Fanning, unpublished data), and prolonged over-expression of helicase-proficient HDHB mutants that cannot be exported prevents the G1/S transition and eventually results in apoptosis (Gu and Fanning, unpublished data).

A C-Terminal Domain of HDHB Confers Cell Cycle-Dependent Localization

A 131-residue domain, PSLD, is sufficient to target HDHB or a βGal reporter to either the nucleus or the cytoplasm in a cell cycle-dependent manner (FIG. 5). A rev-type NES resides in this domain (FIG. 6), but its activity or accessibility to the nuclear export machinery depends on phosphorylation of PSLD, primarily on serine 967, at the G1/S transition (FIG. 7-10). S967 is a perfect match to the consensus CDK substrate recognition motif (S/T)PX(K/R). Both cyclin E/CDK2 and cyclin A/CDK2 can modify HDHB in vitro, but the ability of cyclin E/CDK2 to complex with HDHB in cell extracts suggests that it may be the initial kinase that modifies HDHB at the G1/S transition (FIG. 9). Phosphorylation of HDHB in PSLD appears to persist through the latter part of the cell cycle, correlating well with the predominantly cytoplasmic localization of HDHB. However, it is not possible yet to distinguish whether HDHB undergoes dephosphorylation at the M/G1 transition (FIG. 7C) or is perhaps targeted for proteolysis and rapidly re-synthesized in early G1, when it would enter the nucleus.

Another open question is the mechanism by which phosphorylation of PSLD regulates nuclear export of HDHB. The data provide strong evidence that PSLD contains active targeting signals that are independent of protein context (FIG. 4-5). Since mutant HDHB with an inactivated NES is nuclear even when it is expressed during S phase and thus presumably phosphorylated (FIG. 6), it is probably that the NLS is probably not inactivated or masked by phosphorylation and that the primary target of CDK regulation is the NES. Extending this reasoning, the NES may be masked during G1 when the CDK motifs in PSLD are unmodified, and that the NES is liberated when S967 becomes phosphorylated, leading to NES recognition by nuclear export factors (FIG. 5, 6). How might phosphorylation of PSLD liberate the NES? Structural studies of a rev-type NES have shown that it forms an amphipathic α-helix, with the leucines aligned on one side of the helix and charged residues on the other side (Rittinger et al., Mol. Cell. Biol. (1999), 4, 153-166). Since the SLD of HDHB contains both the rev-type NES and an NLS, and the basic residues likely to serve as the NLS are interspersed through the NES, the NES and NLS may reside on opposite faces of an amphipathic helix. Additional sequences in PSLD would mask the NES intramolecularly, allowing only the NLS to be recognized. Phosphorylation of S967 would alter the conformation of the mask in PSLD to expose the NES, without affecting exposure of the NLS.

Transfection of Cells

U-2OS cells were transiently transfected with pCORON1002-EGFP-C1-PSLD (FIG. 11), pCORON1002-EGFP-C1-Gal-PSLD (FIG. 12) or J-Red derivatives of the above vectors. Cells transiently expressing EGFP-C1-PSLD were obtained (FIG. 13) but proved difficult to work with due to the heterogeneity of expression and variable signal. Stable clones expressing the recombinant fusion proteins were selected using 1 mg/ml G418 (Sigma) or hygromycin, where appropriate. Isolated primary clones (˜60 per construct) were analysed by flow cytometry to confirm the level and homogeneity of expression of the sensor and where appropriate secondary clones were developed using methods above.

Consequently, US-OS cells stably expressing EGFP-C1-PSLD (Clone 22; FIG. 14) and EGFP-C1-Gal-PSLD—were selected which were homogeneous in nature and provided a bright, uniform signal. These cells are much more useful for providing sensitive, stable and uniform assays for investigating the cell cycle and the effect of agents upon the cell cycle. The fluorescent signals generated by the stable cell lines developed with pCORON1002-EGFP-C1-Gal-PSLD and pCORON1002-EGFP-C1-PSLD are shown in FIG. 15.

Certain aspects of the invention disclosed hereinabove has been published in Molecular Biology of the Cell (15: 3320-3332, July 2004) and electronically published as MBC in press, 10.1091/mbc.E04-03-0227 on May 14, 2004, under the title of “Cell Cycle-dependent Regulation of a Human DNA Helicase That Localizes in DNA Damage Foci”, the disclosure of which is incorporated herein by reference in its entireties.

The foregoing is illustrative of the present invention and is not to be construed as limiting thereof. Although a few exemplary embodiments of this invention have been described, those skilled in the art will readily appreciate that many modifications are possible in the exemplary embodiments without materially departing from the novel teachings and advantages of this invention. Accordingly, all such modifications are intended to be included within the scope of this invention as defined in the claims. Therefore, it is to be understood that the foregoing is illustrative of the present invention and is not to be construed as limited to the specific embodiments disclosed, and that modifications to the disclosed embodiments, as well as other embodiments, are intended to be included within the scope of the appended claims. The invention is defined by the following claims:

Claims

1. A polypeptide construct comprising a detectable live-cell reporter molecule linked via a group having a molecular mass of less than 112, 000 Daltons to at least one cell cycle phase-dependent location control element, the location of which said element changes during G1 and S phase, wherein the translocation of said construct within a mammalian cell is indicative of the cell cycle position.

2. A polypeptide construct according to claim 1 wherein said group has a molecular mass of less than 100,000 Daltons.

3. A polypeptide construct according to claim 1 wherein said group has a molecular mass of less than 50,000 Daltons.

4. A polypeptide construct according to claim 1 wherein said group has a molecular mass of less than 25,000 Daltons.

5. A polypeptide construct according to claim 1 wherein said group has a molecular mass of less than 10,000 Daltons.

6. A polypeptide construct according to claim 1 wherein said group has a molecular mass of less than 1,000 Daltons.

7. A polypeptide construct according to claim 1 wherein said group has a molecular mass of less than 700 Daltons.

8. A polypeptide construct according to claim 1 wherein said group has a molecular mass of less than 500 Daltons.

9. A polypeptide construct according to claim 1 wherein the group is a polypeptide.

10. A polypeptide construct according to claim 9 wherein said polypeptide group is a heptapeptide.

11. The polypeptide construct of claim 10, wherein said heptapeptide is Gycine-Asparagine-Glycine-Glycine-Asparagine-Alanine-Serine (GNGGNAS).

12. A polypeptide construct according to any preceding claim, wherein the cell cycle phase-specific dependent location control element is selected from the group of peptides consisting of Rag2, Chaf1B, Fen1, PPP1R2, helicase B, sgk, CDC6 or motifs therein such as the phosphorylation-dependent subcellular localization domain of the C-terminal special control region of helicase B (PSLD).

13. A polypeptide construct according to any of claims 1 to 12 wherein the live-cell reporter molecule is selected from the group consisting of fluorescent protein, enzyme reporter and antigenic tag.

14. A polypeptide construct according to claim 13 wherein said fluorescent protein is selected from Green Fluorescent Protein (GFP), Enhanced Green Fluorescent Protein (EGFP), Emerald and J-Red.

15. A polypeptide construct according to claim 13 wherein said enzyme reporter is halo-tag (Promega).

16. A polypeptide construct according to any preceding claim, wherein the cell cycle phase-dependent location control element is PSLD.

17. A polypeptide construct according to any preceding claim, wherein the reporter molecule is EGFP and the cell cycle phase-dependent location control element is PSLD.

18. A polypeptide construct comprising the amino acid sequence of SEQ ID No. 5.

19. A nucleic acid construct encoding any of the polypeptide constructs according to any preceding claim.

20. The nucleic acid construct of claim 19, wherein said construct additionally comprises and is operably linked to and under the control of at least one cell cycle independent expression control element.

21. The nucleic acid construct of claim 20, wherein said expression control element is either an ubiquitin C promoter or a CMV promoter.

22. The nucleic acid construct according to any of claims 19 to 21 comprising a CMV promoter, and sequences encoding PSLD and EGFP or J-Red.

23. The nucleic acid construct according to any of claims 19 to 21 comprising a ubiquitin C promoter, and sequences encoding PSLD and EGFP or J-Red.

24. A vector comprising any of the nucleic acid constructs of any of claims 19 to 23.

25. A vector according to claim 24, wherein said vector is either a viral vector or a plasmid.

26. A vector according to claim 25, wherein said viral vector is an adenoviral vector or a lentiviral vector.

27. A host cell transfected with a nucleic acid construct according to any of claims 19 to 23.

28. The host cell according to claim 27, wherein said cell is a human cell.

29. A stable cell line comprising one or more of the host cells according to either of claims 27 or 28.

30. Use of a polypeptide construct according to any of claims 1 to 18 for determining the cell cycle position of a mammalian cell.

31. A method for determining the cell cycle position of a mammalian cell, said method comprising: a) expressing in a cell a nucleic acid construct according to any of claims 19 to 23; and b) determining the cell cycle position by monitoring signals emitted by the reporter molecule.

32. A method of determining the effect of a test agent on the cell cycle position of a mammalian cell, said method comprising: a) expressing in said cell in the absence and in the presence of said test agent a nucleic acid construct according to any of claims 19 to 23; and b) determining the cell cycle position by monitoring signals emitted by the reporter molecule wherein a difference between the emitted signals measured in the absence and in the presence of said test agent is indicative of the effect of the test agent on the cell cycle position of the cell.

33. A method of determining the effect of a test agent on the cell cycle position of a mammalian cell, said method comprising: a) expressing in said cell in the presence of said test agent a nucleic acid construct according to any of claims 19 to 23; and b) determining the cell cycle position by monitoring signals emitted by the reporter molecule, c) comparing the emitted signal in the presence of the test agent with a known value for the emitted signal in the absence of the test agent; wherein a difference between the emitted signal measured in the presence of the test agent and said known value in the absence of the test agent is indicative of the effect of the test agent on the cell cycle position of the cell.

34. A method of determining the effect of a test agent on the cell cycle position of a mammalian cell, said method comprising: a) providing cells containing a nucleic acid construct according to any of claims 19 to 23; b) culturing first and second populations of said cells respectively in the presence and absence of a test agent and under conditions permitting expression of the nucleic acid reporter construct; and c) measuring the signals emitted by the reporter molecule in said first and second cell populations; wherein a difference between the emitted signals measured in said first and second cell populations is indicative of the effect of said test agent on the cell cycle position of said cell.

35. A method of determining the effect of the mammalian cell cycle on a cellular process measurable by a first detectable reporter which is known to vary in response to a test agent, said method comprising: a) expressing in said cell in the presence of said test agent a second nucleic acid reporter construct according to any of claims 19 to 23; b) determining the cell cycle position by monitoring signals emitted by the second reporter molecule; and c) monitoring the signals emitted by said first detectable reporter. wherein the relationship between cell cycle position determined by step b) and the signal emitted by the first detectable reporter is indicative of whether or not said cellular process is cell cycle dependent.

36. Use of a polypeptide construct according to any of claims 1 to 18 for measuring CDK2 activity in a cell.

37. A method for measuring CDK2 activity in a cell, said method comprising the steps of a) expressing a nucleic acid construct according to any of claims 19 to 23 in a cell, and b) determining CDK2 activity by monitoring signals emitted by the reporter molecule.

38. A method for determining the effect of a test agent on CDK2 activity of a mammalian cell, said method comprising: a) expressing in said cell in the absence and in the presence of said test agent a nucleic acid construct according to any of claims 19 to 23; and b) determining CDK2 activity by monitoring signals emitted by the reporter molecule wherein a difference between the emitted signals measured in the absence and in the presence of said test agent is indicative of the effect of the test agent on the activity of CDK2.

39. A method of determining the effect of a test agent on CDK2 activity of a mammalian cell, said method comprising: a) expressing in said cell in the presence of said test agent a nucleic acid construct according to any of claims 19 to 23; and b) determining the cell cycle position by monitoring signals emitted by the reporter molecule, c) comparing the emitted signal in the presence of the test agent with a known value for the emitted signal in the absence of the test agent; wherein a difference between the emitted signal measured in the presence of the test agent and said known value in the absence of the test agent is indicative of the effect of the test agent on the CDK2 activity of the cell.

40. The method according to any of claims 37 to 39, wherein said test agent is a form of electromagnetic radiation or is a chemical entity.