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Originally published In Press as doi:10.1074/jbc.M608165200 on October 26, 2006

J. Biol. Chem., Vol. 281, Issue 52, 40330-40340, December 29, 2006
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Deficiency of the Zinc Finger Protein ZPR1 Causes Defects in Transcription and Cell Cycle Progression*

Laxman Gangwani1

From the Program in Molecular Medicine, University of Massachusetts Medical School, Worcester, Massachusetts 01605

Received for publication, August 24, 2006 , and in revised form, October 24, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The zinc finger protein ZPR1 is present in both the cytoplasm and nucleoplasm. Cell cycle analysis demonstrates that ZPR1 undergoes major changes in subcellular distribution during proliferation. ZPR1 is diffusely localized throughout the cell during the G1 and G2/M phases of the cell cycle. In contrast, ZPR1 redistributes to the nucleus during S phase and ZPR1 exhibits prominent co-localization with the survival motor neurons protein and the histone gene-specific transcription factor NPAT in subnuclear foci, including Cajal bodies that associate with histone gene clusters. ZPR1 deficiency causes disruption of survival motor neurons and NPAT localization within the nucleus, blocks S phase progression, and arrests cells in both the G1 and G2 phases of the cell cycle. These changes in subnuclear architecture and cell cycle progression may be caused by transcriptional defects in ZPR1-deficient cells, including decreased histone gene expression.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
ZPR1 was initially identified as a cytoplasmic zinc finger protein that interacts with tyrosine kinase receptors, including the epidermal growth factor receptor, in quiescent mammalian cells (1). In mitogen-stimulated cells, ZPR1 does not interact with epidermal growth factor receptors, but instead forms complexes with eukaryotic translation elongation factor-1{alpha} (2). Mutational analysis indicates that ZPR1 residues necessary for this interaction with eukaryotic translation elongation factor-1{alpha} are required for normal cellular proliferation (2). Indeed, genetic analysis indicates that Zpr1 is an essential gene (2-4).

Recent studies have demonstrated that ZPR1 also interacts with complexes formed by the survival motor neurons (SMN)2 protein (5). Mutations of the telomeric copy of the Survival Motor Neurons 1 (SMN1) gene that result in low level expression of the full-length SMN protein are a major cause of spinal muscular atrophy (SMA) (6-9). SMA is an autosomal recessive disease of early childhood that is characterized by the loss of {alpha}-motor neurons in the anterior horns of the spinal cord that result in muscle weakness, respiratory failure, and death (10). In humans, two copies of the SMN gene, SMN1 (telomeric) and SMN2 (centromeric), are located on chromosome 5q13 (6). These human SMN genes are very similar. However, transcripts from the SMN2 gene undergo alternative splicing and results in the production of low levels of full-length SMN mRNA and high levels of a truncated mRNA that lacks sequences derived from exon 7 (11, 12). In contrast to humans, mice have only one copy of the Smn gene. Homozygous deletion of Smn in mice results in embryonic lethality (13). The function of SMN in cell survival is unclear. SMN forms multiprotein complexes that are present in both the cytoplasm and nucleus. Cytoplasmic SMN is involved in the assembly of the Sm-core domain of small ribonuclear particles (snRNPs) (14-16), whereas nuclear SMN may be involved in pre-mRNA splicing (17). ZPR1 is part of the cytoplasmic preimport snRNP complex formed by SMN, Sm proteins, snurportin 1, and importin beta (18). In the nucleus, ZPR1 co-localizes with SMN in nuclear gems and Cajal bodies (4, 5).

The reduced amounts of full-length SMN protein causes disruption of multiprotein SMN complexes, including disruption of SMN-ZPR1 complexes in SMA patients (5). The co-localization of ZPR1 with SMN in nuclear bodies, including gems and Cajal bodies, observed in normal human cells is not observed in cells derived from SMA patients (5). ZPR1 is required for accumulation of SMN in subnuclear bodies, including gems and Cajal bodies (4). It is established that humans with SMA express low levels of ZPR1 (19). Furthermore, the reduced expression of ZPR1 is sufficient to cause neurodegeneration in mice (20). It is possible that ZPR1 may act as a modifier gene that may contribute to the severity of SMA disease (19-21).

Although it is established that Zpr1 is an essential gene in mice (4), the molecular mechanism that accounts for the requirement of ZPR1 for cell survival is unclear. The purpose of this study was to examine the effect of ZPR1 deficiency on cell survival. I show that ZPR1 deficiency causes defects in histone gene expression, prevents DNA synthesis, and causes disruption of the subnuclear localization of proteins that are associated with DNA synthesis, including the DNA polymerase subunit proliferating cell nuclear antigen (PCNA) and the histone gene transcription factor, NPAT.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture—HeLa cells (Strain PV, American Type Culture Collection (ATCC)) and wild type human fibroblasts (WI-38, ATCC) were cultured in minimal Eagle's medium (MEM) with 10% fetal bovine serum (FBS), 2 mM glutamine, 100 units/ml penicillin, 100 units/ml streptomycin, and 1 mM sodium pyruvate.

HeLa cells were synchronized by double thymidine block (22) by sequential incubation of the cells in serum-free medium supplemented with 2 mM thymidine (12-16 h), growth medium with 24 µM deoxycytidine (9 h), and serum-free medium with 2 mM thymidine (12-16 h). The cells were released from the second thymidine block by washing with serum-free medium and the subsequent addition of growth medium supplemented with 24 µM deoxycytidine. The cells were harvested at different time intervals after release from the second thymidine block.

WI-38 cells were arrested in the G0 phase of the cell cycle by serum starvation for 72 h in medium supplemented with 0.1% FBS. WI-38 cells were released from growth arrest by incubation in fresh growth medium with 10% FBS.

Antisense Oligonucleotides—Second generation optimized antisense oligonucleotides were purchased from Oligos Etc. Inc. The experimental conditions optimized for knockdown of ZPR1 expression using antisense human ZPR1 oligonucleotide 5'-CATGGCCACCACGCG-CAATT-3' and control oligonucleotide 5'-CACGGCTACCTCGCACAAGT-3' (scrambled sequence) were employed (5). The oligonucleotides labeled with or without FITC (0.4 nmol) in 100 µl of Opti-MEM (Invitrogen) were mixed with 100 µl of Opti-MEM containing 6 µl of Lipofectin (Invitrogen), diluted to 1 ml with Opti-MEM, and incubated (270 min) with subconfluent HeLa cells in 35-mm dishes at 37 °C. One ml of Dulbecco's modified Eagle's medium with 20% FBS was added and the cells were incubated at 37 °C (30 h). Transfection efficiency of FITC-conjugated oligonucleotides was monitored by flow cytometry. Greater than 95% of the cells were transfected with antisense oligonucleotides. Cell lysates were prepared and examined by immunoblot analysis.

BrdUrd and BrUrd Incorporation Assay—Cells were labeled by incubation in medium supplemented with 50 µM BrdUrd (1 h). BrdUrd-labeled cells were detected by flow cytometry and immunofluorescence microscopy using a BrdUrd detection kit (BD Biosciences). Cells (95-98%) were gated to exclude dead cells and aggregates. The distribution of cells in different phases of the cell cycle was calculated by the Dean-Jett-Fox method using Flow-Jo software. The average of three independent experiments (mean ± S.D.; n = 3) is presented. The cells were also labeled with 2 mM 5'-bromouridine (BrUrd) for 2 h, and released for 2 h by substituting fresh medium without BrUrd for detection of in vivo transcription activity. BrUrd-labeled cells were detected using FITC-coupled anti-BrdUrd antibody (BD Biosciences).

Immunofluorescence Analysis—Cells were cultured on glass coverslips, rinsed with phosphate-buffered saline (PBS; 4.3 mM Na2HPO4, 1.4 mM KH2PO4, 137 mM NaCl, and 2.7 mM KCl, pH 7.4), and then briefly with chilled methanol. The cells were fixed at -20 °C with methanol (5 min) and acetone (2 min). The coverslips were blocked with 3% bovine serum albumin in PBS with 0.5% Tween 20 (PBS-T) for 30 min at 25 °C in Coplin jars and then incubated with primary antibodies for 1 h at 25°C (5). Double labeling (ZPR1/SMN) was carried out by sequential incubations (1 h) with anti-SMN (clone 2B1) (23), Texas Redconjugated anti-mouse IgG secondary antibody (Jackson ImmunoResearch), and then with FITC or Alexa 488-conjugated anti-ZPR1 (clone LG1, mouse monoclonal antibody raised against full-length human protein) (5) at 25 °C. Double labeling (ZPR1/NPAT) was carried out by sequential incubations (1 h) with anti-NPAT (Transduction Lab), Texas Redconjugated anti-mouse IgG secondary antibody (Jackson ImmunoResearch), and then with FITC or Alexa 488-conjugated anti-ZPR1 (clone LG1) at 25 °C. Double labeling (ZPR1/p80 Coilin) was carried out by sequential incubations (1 h) with rabbit anti-p80 Coilin (number R288) (24) and Texas Red-conjugated anti-rabbit IgG secondary antibody and then with Alexa 488-conjugated anti-ZPR1 (clone LG1) at 25 °C. Double labeling (ZPR1/PCNA) was carried out by sequential incubations (1 h) with rabbit anti-PCNA (Santa Cruz Biotechnology) and Texas Redconjugated anti-rabbit IgG secondary antibody and then with FITC-conjugated anti-ZPR1 (clone LG1) at 25 °C.

Processed coverslips were mounted on slides with mounting medium (Vectashield) containing 4',6'-diamino-2-phenylindole (DAPI). Fluorescence microscopy was performed using a Zeiss inverted microscope (Axiovert M200) or a confocal laser scanning microscope (Leica TCS SP2) equipped with a 405-nm diode laser. All the images were processed similarly and acquired under identical conditions by keeping all microscope parameters constant, including PMT voltage, aperture (pin hole), scan speed, and excitation and emission wavelengths.

Northern Blot Analysis—HeLa cells were harvested after 30 h of transfection with antisense oligonucleotides. Total cellular RNA was extracted using the RNeasy kit (Promega Corp.) according to the instructions of the manufacturer. Total RNA (10 µg) was separated by electrophoresis using denaturing formaldehyde-agarose gel and transferred overnight onto a Zetaprobe membrane (Bio-Rad) by capillary action using 10x SSC buffer (3 M NaCl, 0.3 M sodium citrate). The RNA was cross-linked to the membrane by UV irradiation using a UV Stratalinker 2400 (Stratagene). The blots were prehybridized in 5x SSC, 5x Denhardt's solution, 0.5% SDS, and 10 µg/ml sonicated salmon sperm DNA for 2 h at 65°C. Probes (32P-labeled) specific for the H4 histone (H4/n coding region) (25) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) genes were generated by random priming (Stratagene). RNA blots were washed in buffers starting at 2x SSC (0.1% SDS), followed by 1x SSC and a final rinse in 0.1x SSC (0.1% SDS) at 65 °C. Blots were subjected to autoradiographic exposure overnight at -70 °C or analyzed by using a Storm 860 PhosphorImager (Amersham Biosciences).

In Situ Run-on Transcription—In situ run-on transcription was performed as described previously (26). HeLa cells were processed after 30 h of transfection with antisense oligonucleotides. Cells were rinsed with PBS and then with 10 mM phosphate buffer, pH 7.2 (PB), supplemented with 100 mM potassium acetate, 30 mM KCl, 1 mM MgCl2, 1 mM ATP, 1 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride, 100 µg/ml bovine serum albumin, and 20 units/ml RNasin. The cells were permeabilized using the same buffer supplemented with 0.05% Triton X-100 (5 min at 25 °C). The cells were washed with PB and then incubated in PB supplemented with 100 µM ATP, CTP, GTP, and BrUTP (30 min at 37 °C), washed with PB, fixed with 4% paraformaldehyde (15 min at 25 °C), washed with PBS, and stored in 70% ethanol at 4 °C. Double labeling (ZPR1/RNA) was carried out by sequential incubations (1 h) in PBS plus 0.2% Triton X-100 with anti-ZPR1 (clone LG1), Alexa 546-conjugated anti-mouse IgG secondary antibody, and Alexa 488-conjugated anti-BrdUrd (clone PRB1, Molecular Probes) at 25 °C. Processed coverslips were mounted on slides with mounting medium containing DAPI. Fluorescence microscopy was performed using a confocal laser scanning microscope (Leica TCS SP2) equipped with a 405-nm diode laser.


Figure 1
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  		FIGURE 1.
Cell cycle-dependent localization of ZPR1. A, HeLa cells were synchronized using a double thymidine block and released by incubation in medium supplemented with 10% FBS. The cells were examined by immunofluorescence microscopy to detect ZPR1 (FITC, green) and DNA (DAPI, blue). The scale bar is 10 µm. B, HeLa cells were synchronized using a double thymidine block and released by incubation in medium supplemented with 10% FBS to obtain cells in S phase and G1 phase. Synchronized cells incubated with FBS (4 h) were also subsequently incubated (8 h) with 3 µM aphidicolin, 2 mM hydroxyurea (H-Urea), or 0.25 µM nocodazole. The cells were examined by immunofluorescence analysis to detect ZPR1 (green) and DNA (DAPI, blue). The scale bar is 10 µm. C, HeLa cells were synchronized using a double thymidine block and released by incubation in medium supplemented with 10% FBS to obtain cells in different phases of the cell cycle. Synchronized cells released from thymidine block were also subsequently incubated with medium containing 10% FBS for 16 h (G1 phase), 3 µM aphidicolin for 8 h (S phase), and 0.25 µM nocodazole for 10 h (G2/M Phase). HeLa cells were starved for 12 h in serum-free medium (G0 phase). The expression of ZPR1, SMN, and beta-actin was examined by immunoblot analysis of cell lysates. Quantitation of immunoblots was performed using MetaMorph software. The relative amounts (mean ± S.D.; n = 4) of proteins normalized to beta-actin during different phases of the cell cycle are presented as the bar graph.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Cell Cycle-dependent Distribution of ZPR1—ZPR1 is present in both the cytoplasm and the nucleus of mammalian cells and redistributes from the cytoplasm to the nucleus in response to serum or mitogens (1, 5). The mechanism of nuclear redistribution of ZPR1 in response to growth proliferation signals is unclear. To define the function of ZPR1 in cell growth and development, I examined the subcellular distribution of ZPR1 during progression of the cell cycle using immunofluorescence microscopy in human cells. Staining of synchronized HeLa cells that are at the boundary of G1/S (G1 phase) with anti-ZPR1 antibodies shows that ZPR1 is present in both the cytoplasm and nucleus (Fig. 1A). ZPR1 is diffusely localized within the nucleus of G1 phase cells. ZPR1 begins to accumulate in subnuclear bodies during progression into the S phase (Fig. 1A). The number of ZPR1 containing nuclear bodies increases to ~8 during S phase in HeLa cells. The amount of ZPR1 (detected by immunoblot analysis) does not change as the cells progress through S phase (Fig. 1C), but the overall fluorescence intensity dramatically decreases during S phase (Fig. 1A), most likely because of epitope masking that may occur when ZPR1 accumulates in subnuclear bodies. In contrast to the accumulation in subnuclear bodies (gems and Cajal bodies) during S phase, ZPR1 was diffusely localized throughout the cell during mitosis (metaphase, anaphase, and telophase) (Fig. 1A). These changes in the subcellular localization of ZPR1 contrast with the predominantly cytoplasmic localization of ZPR1 in serum-starved cells growth arrested in G0 that express decreased level of ZPR1 (Fig. 1C) (1, 3, 5). To test the role of the cell cycle phase in the determination of the subcellular localization of ZPR1, I examined the effect of drugs that are known to cause cell cycle arrest. HeLa cells synchronized using a double thymidine block were incubated with serum (4 h) and subsequently with aphidicolin (S phase), hydroxyurea (S phase), or nocodazole (M phase) (Fig. 1B). Immunofluorescence microscopy demonstrated that ZPR1 was localized to subnuclear bodies, including gems and Cajal bodies, following treatment with aphidicolin or hydroxyurea. In contrast, ZPR1 was diffusely localized throughout the cells after treatment with nocodazole. It is possible that the loss of punctate nuclear staining of ZPR1 may be caused by the loss of nuclear membrane during G2/M phase or nocodazole treatment. Together, these data confirm that ZPR1 undergoes cell cycle dependent changes in subcellular localization, including an accumulation in subnuclear bodies during S phase (Fig. 1B). The nuclear accumulation observed during S phase suggests that ZPR1 may play a role during this phase of the cell cycle.


Figure 2
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  		FIGURE 2.
Distribution of SMN and ZPR1 during S phase of the cell cycle. A, serum-starved WI-38 fibroblasts were re-stimulated by incubation in fresh medium containing 10% FBS. The cells were labeled (1 h) with BrdUrd at different times after serum stimulation and then examined by flow cytometry using propidium iodide (PI) and an antibody to BrdUrd. The distribution of cells in different phases of the cell cycle was calculated by the Dean-Jett-Fox method using Flow-Jo software. The average of three independent experiments (mean ± S.D.; n = 3) is presented. B, serum-starved WI-38 fibroblasts were re-stimulated with serum. The cells were labeled (1 h) with BrdUrd at different times after serum stimulation and then examined by immunofluorescence microscopy to detect the subcellular localization of ZPR1 (red) and BrdUrd (green). Nuclei were stained with DAPI (blue). The scale bar is 10 µm.

 
ZPR1 Accumulates in Subnuclear Bodies during DNA Synthesis—To examine the role of ZPR1 containing subnuclear bodies (foci) during S phase of the cell cycle, I examined the number of ZPR1 foci present in cells during cell cycle progression in normal human WI-38 diploid fibroblasts. Cells growth arrested in the G0/G1 phase of the cell cycle by serum starvation were incubated with fresh medium supplemented with 10% FBS. The progression of cells through S phase was examined by pulse labeling cells with BrdUrd and analysis by flow cytometry (Fig. 2A). These data indicated that WI-38 cells enter S phase at ~19 h following serum stimulation and that a large S phase population of cells is detected after 24 h (Fig. 2A).

Immunofluorescence analysis demonstrated that ZPR1 diffusely localized throughout the nucleoplasm and cytoplasm of serum-starved WI-38 fibroblasts (Fig. 2B). A similar diffuse localization of ZPR1 was observed at 6 h following serum stimulation (Fig. 2B) when the cells are predominantly in the G1 phase of the cell cycle (Fig. 2A). The BrdUrd incorporation was not detected in starved and cell released for 6 h (Fig. 2B). However, at 19 h, when cells enter S phase (Fig. 2A), ZPR1 accumulation in subnuclear bodies and BrdUrd incorporation was observed in cells (Fig. 2B). The increase in the number of ZPR1 subnuclear bodies was observed during S phase progression from 19 to 24 h as indicated by increased incorporation of BrdUrd (Fig. 2B). The cells containing different numbers of ZPR1 foci were examined by immunofluorescence analysis (Table 1). The subpopulation of cells containing 2 or more ZPR1 subnuclear foci and labeled with BrdUrd was examined by immunofluorescence and FACS analysis (Table 2). A variable number of subnuclear bodies were detected in individual focal planes, but analysis of multiple focal planes throughout the nucleus indicated that the prominent ZPR1-positive subnuclear bodies increased from 2 to 4 during S phase of the cell cycle in WI-38 fibroblasts.


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  		TABLE 1
Number of ZPR1 foci in cultured WI-38 cells

Cultured WI-38 (randomly cycling) cells stained with monoclonal antibody to ZPR1 (clone LG1) were examined by immunofluorescence microscopy.

 


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  		TABLE 2
Number of ZPR1 foci in cells present in S phase of the cell cycle

Cultured WI-38 (randomly cycling) cells labeled with BrdUrd and stained with monoclonal antibody to ZPR1 (clone LG1) were examined by immunofluorescence (IF) microscopy and fluorescence-activated cell sorter (FACS) analysis.

 
The Co-localization of SMN and ZPR1 Is Cell Cycle-dependent—ZPR1 interacts with the SMN complex and both proteins co-localize in subnuclear bodies (5). To test whether this co-localization is cell cycle-dependent, I examined the subcellular distribution and localization of SMN and ZPR1 during cell cycle progression in normal human WI-38 diploid fibroblasts.

Immunofluorescence analysis demonstrated that ZPR1 and SMN are both diffusely localized throughout the nucleoplasm and the cytoplasm of serum-starved WI-38 fibroblasts (Fig. 3A). A similar diffused subcellular distribution of ZPR1 and SMN was observed at 6 h following serum stimulation (Fig. 3A) when cells are predominantly in G1 phase of the cell cycle (Fig. 2A). However at 19 h, when cells enter S phase of the cell cycle (Fig. 2), both ZPR1 and SMN were observed to co-localize in subnuclear bodies (Fig. 3A). Prominent nuclear bodies containing SMN and ZPR1 were observed in early S phase (19 h) and an increased number of subnuclear bodies were detected as the cells progressed through S phase (24 h). The increase in number of prominent ZPR1-positive subnuclear bodies from 2 to 4 during S phase of the cell cycle in diploid fibroblasts contrasts with observations made using HeLa (aneuploid human tumor) cells that indicated a larger (and more variable) number of ZPR1-positive subnuclear bodies that increased to approximately eight ZPR1-positive subnuclear bodies during S phase progression (Fig. 1). Together, these data indicate that both SMN and ZPR1 redistribute similarly during the cell cycle and accumulate in subnuclear bodies, including gems and Cajal bodies, in a cell cycle-dependent manner. These findings are consistent with our previous findings that the interaction of ZPR1 with SMN is reduced in serum-starved (G0/G1-phase) cells and increased in starved cells treated with serum (S phase) (5).

Localization of ZPR1 with the Transcription Factor NPAT—The localization of ZPR1 to subnuclear bodies during S phase of the cell cycle (Fig. 2B) in human diploid fibroblasts resembles the localization of the transcription factor NPAT (27, 28). NPAT is essential for histone gene expression (29, 30) and is known to localize with histone gene clusters on human chromosomes 1q21 and 6p21 during S phase when active histone gene expression occurs (27, 28). The increase in NPAT-positive subnuclear bodies from 2 to 4 in diploid cells during S phase progression is the result of constitutive interaction with chromosome 6p21 and an S phase-specific induction of an interaction with chromosome 1q21. In contrast, aneuploid human tumor cells with additional copies of chromosomes 1 and 6 exhibit a larger and more variable number of NPAT-positive subnuclear bodies that also increase in number during S phase progression.

The cell cycle-dependent changes in NPAT localization (27, 28) are similar to the changes in ZPR1 localization that I have observed (Figs. 1 and 2). Both NPAT (28) and ZPR1 (5) show similar localizations and are present in a subset of Cajal bodies. To test whether the S phase localization of ZPR1 occurs in NPAT-positive subnuclear bodies, I performed immunofluorescence analysis of WI-38 human diploid fibroblasts. NPAT and ZPR1 were found to be diffusely localized throughout the nucleoplasm and the cytoplasm of serum-starved WI-38 fibroblasts (Fig. 3B). In early S phase (19 h after serum stimulation), ZPR1 and NPAT were found to co-localize in two prominent subnuclear bodies. During S phase progression (24 h), ZPR1 and NPAT were found to co-localize in four prominent sub-nuclear bodies (Fig. 3B). Control studies demonstrated that SMN also co-localized with NPAT in these subnuclear bodies during S phase (data not shown). Because co-localizations of ZPR1 with SMN and ZPR1 with NPAT are similar and ZPR1 interacts with SMN, it is possible that ZPR1 also interacts with NPAT. However, co-immunoprecipitation of NPAT with ZPR1 was not detected. Together, these data indicate that ZPR1 is present in NPAT containing subnuclear bodies that co-localize with histone gene clusters on human chromosomes 1 and 6 during S phase of the cell cycle (27, 28).

ZPR1 Deficiency Causes Defects in Subnuclear Protein Localization—The distinctive localization of ZPR1 within the nucleus (Figs. 2 and 3) may have functional significance. As an initial approach to the examination of the role of ZPR1, I investigated the effect of ZPR1 deficiency on other proteins that also co-localize with Cajal bodies, including NPAT. We previously reported that disruption of the Zpr1 gene causes a pre-implantation lethal phenotype in mice (4). I therefore employed an alternative approach to examine the effect of ZPR1 deficiency using cultured cells. Transfection studies demonstrated that an antisense oligonucleotide can suppress ZPR1 expression (78 ± 2.7%, mean ± S.D.; n = 3) in assays using immunoblot analysis (Fig. 4) and immunofluorescence analysis (Fig. 5A). In contrast, a scrambled sequence oligonucleotide caused no change in ZPR1 expression. Control studies were performed using small interfering RNA-mediated gene suppression with transfected double-stranded RNA (Dharmacon) (data not shown). The extent of knockdown of ZPR1 protein expression was comparable in small interfering RNA-transfected and antisense oligonucleotide-transfected cells and similar results were obtained in experiments using both procedures.

Two proteins that co-localize with ZPR1 during S phase in prominent subnuclear bodies are SMN (Fig. 3A) and NPAT (Fig. 3B). Suppression of ZPR1 expression in HeLa cells prevented the accumulation of both SMN and NPAT in these sub-nuclear bodies (Fig. 5, A and B). These data suggest that ZPR1 is required for the normal co-localization of SMN and NPAT in subnuclear bodies that associate with histone gene clusters during cell proliferation (27, 28). This effect of ZPR1 deficiency on the localization of SMN and NPAT might reflect a specific role of ZPR1 or may result from a more general defect in nuclear architecture that occurs in response to reduced ZPR1 function.


Figure 3
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  		FIGURE 3.
Co-localization of ZPR1 with NPAT and SMN during S phase of the cell cycle. A, serum-starved WI-38 fibroblasts were re-stimulated with serum and examined by immunofluorescence microscopy to detect the subcellular localization of ZPR1 (green) and SMN (red). Nuclei were stained with DAPI (blue). The scale bar is 10 µm. B, serum-starved WI-38 fibroblasts were re-stimulated with serum and examined by immunofluorescence microscopy to detect the subcellular localization of ZPR1 (green) and NPAT (red). Nuclei were stained with DAPI (blue). The scale bar is 10 µm.

 
To test whether ZPR1 deficiency might cause defects in nuclear architecture, I examined the localization of Cajal bodies, which are known to associate with several specific chromosomal loci, including histone gene clusters (31). I therefore examined Cajal bodies by investigating the localization of p80 Coilin, a cytological marker for these subnuclear bodies (24). Immunofluorescence microscopy of HeLa cells demonstrated that there was partial overlap of ZPR1 with p80 Coilin in punctate foci (Fig. 5C). ZPR1 localized with p80 Coilin in some Cajal bodies, but was also present in structures that do not contain p80 Coilin (gems). In contrast, ZPR1 localized with most of the SMN containing subnuclear bodies (Fig. 5A). This co-localization of ZPR1 with both Cajal bodies and gems is similar to the reported localization of SMN (23, 32, 33). Suppression of ZPR1 expression resulted in a diffuse localization of p80 Coilin within cells (Fig. 5C). These data suggest that ZPR1 deficiency causes a defect in the organization of subnuclear bodies, including Cajal bodies. This conclusion is consistent with results of the analysis of cells derived from Zpr1-/- embryos (4).


Figure 4
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  		FIGURE 4.
ZPR1 deficiency does not cause defects in the expression of NPAT, SMN, or PCNA. HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense). Control experiments were performed using cells transfected with an oligonucleotide with a scrambled sequence (Scramble). The cells were harvested at 30 h post-transfection. The expression of ZPR1, SMN, NPAT, and PCNA was examined by immunoblot analysis of cell lysates. Quantitation of immunoblots was performed using MetaMorph software. The relative amounts (mean ± S.D.; n = 3) of proteins normalized to beta-actin are presented as bar graph.

 
ZPR1 Deficiency Causes Defects in DNA Replication—The co-localization of ZPR1 with NPAT containing subnuclear bodies that are known to co-localize with histone gene clusters during S phase (Fig. 3B) suggests that ZPR1 may play a role during DNA replication because it is known that defects in histone expression impair S phase progression (34, 35). To test this hypothesis, I examined the effect of ZPR1 deficiency on DNA replication. Exponentially growing HeLa cells were transfected with antisense or scrambled sequence oligonucleotides and incubated in medium (30 h). The cells were pulse-labeled by incubation with BrdUrd and subsequently examined using flow cytometry. ZPR1 deficiency severely impaired the ability of the cells to enter S phase and the cells accumulated at both the G1 and G2/M phases of the cell cycle (Fig. 6A). These data indicate that ZPR1 is required for DNA replication during S phase of the cell cycle. The pattern of growth arrest caused by ZPR1 deficiency is similar to that caused by deficiency of transcription factor NPAT, which is essential for histone gene expression (29, 30).

To obtain independent evidence that ZPR1 deficiency caused an S phase defect, I examined the effect of ZPR1 deficiency on the localization of PCNA, a subunit of DNA polymerase {delta} that is frequently used to detect proliferating cells in the S phase of the cell cycle. Knockdown of ZPR1 expression in HeLa cells did not alter PCNA expression (Fig. 4). Immunofluorescence analysis identified control cells with PCNA that was diffusely localized in the nucleoplasm (Fig. 6B). ZPR1 deficiency markedly altered the subnuclear localization of PCNA and caused an accumulation of PCNA within distinct subregions of the nucleus (Fig. 6B). These data indicate that ZPR1 is required for the normal localization of PCNA and are consistent with the observation that ZPR1 deficiency causes a major defect in S phase of the cell cycle.


Figure 5
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  		FIGURE 5.
ZPR1 deficiency causes mislocalization of SMN, NPAT and p80 Coilin. A, HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense), or a scrambled sequence antisense oligonucleotide (Scramble) and incubated for 30 h. The cells were fixed and stained with antibodies to ZPR1 (green) and SMN (red). Nuclei were stained with DAPI (blue). The subcellular distribution of ZPR1 and SMN was examined by immunofluorescence analysis. The scale bar is 8 µm. B, the subcellular distribution of ZPR1 (green) and NPAT (red) was examined by immunofluorescence analysis. Nuclei were stained with DAPI (blue). The scale bar is 10 µm. C, the subcellular distribution of ZPR1 (green) and p80 Coilin (red) was examined by immunofluorescence analysis. Nuclei were stained with DAPI (blue). The scale bar is 8 µm.

 
ZPR1 Deficiency Causes Defects in Transcription—The mechanism that accounts for block in S phase progression caused by ZPR1 deficiency (Fig. 6A) may be related to defects in histone gene expression. For example, both ZPR1 and NPAT co-localize in subnuclear bodies that associate with histone gene clusters (Fig. 3B) and ZPR1 deficiency causes mislocalization of NPAT (Fig. 5B). This mislocalization is likely to have functional significance because it is established that the transcription factor NPAT is essential for histone gene expression and that NPAT deficiency blocks S phase progression and causes growth arrest in both the G1 and G2 phases of the cell cycle (29, 30, 36). The growth defect caused by NPAT deficiency resembles that caused by ZPR1 deficiency.

To test whether ZPR1 deficiency caused defects in histone gene expression, I examined the expression of histone H4 mRNA by Northern blot analysis (Fig. 7A). Equal amounts of total RNA isolated from control cells and from cells transfected with antisense or scrambled sequence oligonucleotides were examined. Similar amounts of GAPDH mRNA were detected in each sample. However, a selective loss of histone H4 expression was observed in ZPR1-deficient cells. Quantitative analysis of histone gene mRNA expression normalized to GAPDH (histone H4)/(GAPDH) indicated that 60% reduction in histone H4 expression was detected in cells lacking ZPR1 (Fig. 7A). These data indicate that ZPR1 deficiency may cause decreased transcription of HISTONE H4 genes and are consistent with the observation that ZPR1 deficiency causes mislocalization of the transcription factor NPAT.

The observation that ZPR1 deficiency causes decreased expression of HISTONE H4 genes raises a question concerning the specificity of this defect. Does ZPR1 deficiency cause a defect in the expression of a subset of genes (including histone genes), or does it cause a generalized defect in the expression of many genes? To address this question, I examined total RNA expression in control and ZPR1-deficient cells using an in situ transcription run-on assay (Fig. 7B). Control cells and cells transfected with a scrambled sequence oligonucleotide were found to express high levels of RNA. In contrast, ZPR1-deficient cells were found to express very low levels of RNA in this run-on assay. A similar decrease in total RNA expression caused by ZPR1 deficiency was observed using cells pulse-labeled with BrUrd (Fig. 7C). The quantitation of fluorescence intensity of BrUrd incorporated RNA shows that ZPR1 deficiency causes 65.06 ± 5.05% (mean ± S.D.; n = 30) reduction in RNA transcription. ZPR1 deficiency also decreased the expression of rRNA in the nucleolus (Fig. 7C). This result is consistent with our previous finding that ZPR1 deficiency causes a decrease in 35 S-precursor rRNA in yeast cells (3). These data indicate that ZPR1 deficiency causes a decrease in total RNA expression that may be a result of reduced transcription of a specific subset of genes, including histones genes and rRNA.

It is established that spliceosomal snRNPs play an important role in pre-mRNA splicing that is essential for RNA expression. ZPR1 is part of the snRNP complex formed by cytoplasmic SMN (18). To test whether reduced expression of ZPR1 causes defects in the subcellular localization of snRNPs in HeLa cells, the effect of ZPR1 antisense oligonucleotides on subcellular distribution of snRNPs using antibodies against Sm proteins was examined. Immunofluorescence analysis shows that ZPR1 deficiency causes decrease in the cytoplasmic pool of snRNPs and increased accumulation of snRNPs in the nucleus (Fig. 7D). This is consistent with our previous finding that ZPR1 deficiency causes mislocalization of spliceosomal snRNPs in cells derived from Zpr1-/- embryos (4). Together, these data suggest that a decrease in total RNA transcription may be caused by defects in pre-mRNA splicing. Because cell cycle progression requires new gene expression, this defect in RNA expression is sufficient to explain the defect in S phase progression and arrest in both the G1 and G2 phases of the cell cycle observed in ZPR1-deficient cells.

Inhibition of Transcription Causes Mislocalization of ZPR1—The observation that ZPR1-deficient cells exhibit severe defects in RNA expression (Fig. 7) raises questions concerning the role of decreased transcription in the control of protein localization within the nucleus. To test whether decreased transcription may influence subnuclear protein localization, I examined the effect of treatment of cells with actinomycin D, a drug that inhibits transcription and is known to cause Cajal body disruption (37). This drug caused disruption of the subnuclear localization of ZPR1, SMN, and p80 Coilin (Fig. 8, A and B). Actinomycin D also caused the redistribution of PCNA from a diffuse localization in the nucleoplasm to an accumulation within distinct subregions of the nucleus (Fig. 8C). These changes in the localization of SMN, p80 Coilin, and PCNA caused by actinomycin D are similar to the changes in subnuclear distribution caused by ZPR1 deficiency (Figs. 5 and 6). These data suggest that decreased gene expression may contribute to the effects of ZPR1 deficiency on subnuclear protein localization and cell proliferation.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Targeted gene ablation studies indicate that Zpr1 is an essential gene (2-4). However, the mechanism that accounts for the requirement of ZPR1 for cell viability is unclear. The results of the present analysis of cultured cells using a knock-down approach indicate that ZPR1 is necessary for normal cell cycle progression. ZPR1 deficiency blocks S phase progression and causes growth arrest in both the G1 and G2 phases of the cell cycle (Fig. 6). The cause of cell cycle arrest in ZPR1-deficient cells is probably related to the finding that these cells exhibit severe defects in RNA expression, including histone mRNA expression (Fig. 7).


Figure 6
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  		FIGURE 6.
ZPR1 is required for DNA synthesis. A, HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense), or a scrambled sequence oligonucleotide (Scramble) and incubated for 30 h. The cells were labeled (1 h) with BrdUrd and examined by flow cytometry using propidium iodide (PI) and an antibody to BrdUrd. The distribution of cells in different phases of the cell cycle was calculated by the Dean-Jett-Fox method using Flow-Jo software. The average of three independent experiments (mean ± S.D.; n = 3) is presented. B, HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense), or a scrambled sequence antisense oligonucleotide (Scramble) and incubated for 30 h. The cells were fixed and stained with antibodies to ZPR1 (green) and PCNA (red). Nuclei were stained with DAPI (blue). The subcellular distribution of ZPR1 and PCNA was examined by immunofluorescence analysis. The scale bar is 10 µm.

 
Nucleocytoplasmic Transport of ZPR1—The redistribution of ZPR1 and SMN between the cytoplasm and the nucleus in serum-stimulated cells is similar (5). This redistribution of ZPR1 and SMN is related to cell cycle-dependent changes in subcellular localization. The mechanism of nuclear recycling of these proteins is unclear, but nuclear export of ZPR1 may require cyclophilin A peptidyl-prolyl isomerase activity (38) and nuclear import of ZPR1 may be facilitated by snurportin1 and importin beta (18). Nuclear import of SMN requires snRNP and may also depend on snurportin1 and importin beta (18, 39). However, it appears that ZPR1 and SMN can be independently transported into the nucleus (39). This independent transport raises questions concerning the requirement of SMN and ZPR1 for nuclear localization of these proteins. However, ZPR1 deficiency does cause mislocalization of SMN in the nucleus and SMN deficiency does cause mislocalization of ZPR1 (4, 5). Together, the experimental evidence available indicates that the role of ZPR1 may be to regulate the accumulation of SMN in subnuclear bodies rather than to regulate the nuclear import of SMN.


Figure 7
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  		FIGURE 7.
ZPR1 deficiency causes defects in RNA transcription. A, HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense), or a scrambled sequence oligonucleotide (Scramble) and incubated for 30 h. Total RNA was isolated and examined by Northern blot analysis using histone H4/n and GAPDH probes. The amount of histone H4 and GAPDH mRNA was quantitated by PhosphorImager analysis. The relative amount (mean ± S.D.; n = 4) of histone H4 mRNA normalized to GAPDH is presented as bar graph. B, HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense), or a scrambled sequence oligonucleotide (Scramble) and incubated for 30 h. The cells were permeabilized and in situ run-on transcription was investigated by incubation with nucleotides, including Br-UTP. After fixation, the cells were stained with antibodies to ZPR1 (red) and BrdUrd (green) and examined by immunofluorescence microscopy. Nuclei were stained with DAPI (blue). The scale bar is 8 µm. C, HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense), or a scrambled sequence oligonucleotide (Scramble) and incubated for 30 h. The cells were labeled in vivo with 2 mM 5'-bromouridine. After fixation, the cells were stained with antibodies to ZPR1 (red) and BrUrd (green) and examined by immunofluorescence microscopy. Nuclei were stained with DAPI (blue). The scale bar is 8 µm. D, HeLa cells were transfected without (Control) or with a ZPR1 antisense oligonucleotide (Antisense), or a scrambled sequence oligonucleotide (Scramble) and incubated for 30 h. HeLa cells were stained with antibodies to ZPR1 (red) and Sm proteins (green). Nuclei were stained with DAPI (blue). The scale bar is 10 µm.

 
ZPR1 Co-localizes with NPAT in Subnuclear Bodies That Are Required for Histone Gene Expression—Immunofluorescence analysis demonstrates that the subcellular localization of ZPR1 is regulated during the cell cycle. ZPR1 is diffusely localized throughout the nucleus and cytoplasm of diploid human fibroblasts that are growth arrested by serum starvation (Fig. 2). Nuclear accumulation of ZPR1 is observed at an early stage following serum stimulation, although large amounts of ZPR1 remain present in the cytoplasm. However, during S phase of the cell cycle, ZPR1 prominently localizes to four subnuclear bodies. Co-immunofluorescence analysis indicates that these S phase ZPR1-positive foci also stain for NPAT and SMN (Fig. 3). It is established that this localization of NPAT represents an accumulation of this transcription factor with histone gene clusters on human chromosomes 1q21 and 6p21 (27, 28). The association of NPAT with the histone gene cluster on chromosome 6p21 is constitutive, whereas the association with chromosome 1q21 is induced during S phase. The number of NPAT-positive foci therefore increases from 2 to 4 during S phase. This localization is consistent with the essential role of NPAT in histone gene expression during S phase of the cell cycle (29, 30).


Figure 8
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  		FIGURE 8.
Inhibition of transcription causes mislocalization of ZPR1. A, HeLa cells were untreated (Control) or treated with actinomycin D (Act-D) (1 µg/ml) for 2 h and examined by immunofluorescence microscopy. The cells were stained with antibodies to ZPR1 (green) and SMN (red). Nuclei were stained with DAPI (blue). The scale bar is 8 µm. B, HeLa cells were stained with antibodies to ZPR1 (green) and p80 Coilin (red). Nuclei were stained with DAPI (blue). The scale bar is 8 µm. C, HeLa cells were stained with antibodies to ZPR1 (green) and PCNA (red). Nuclei were stained with DAPI (blue). The scale bar is 8 µm.

 
Immunofluorescence analysis demonstrates that ZPR1 colocalize with NPAT in subnuclear bodies. Both ZPR1 and NPAT exhibit a similar localization in subnuclear bodies that co-localize with histone gene clusters during S phase of the cell cycle. Notably, NPAT is mislocalized in ZPR1-deficient cells (Fig. 5). ZPR1 can therefore influence the association of NPAT with histone gene clusters (Fig. 5) and can decrease NPAT-dependent histone gene expression (Fig. 7). The mechanism that accounts for this effect of ZPR1 may be indirect because ZPR1 deficiency blocks S phase progression and consequently may cause NPAT mislocalization as a consequence of cell cycle arrest. However, it is possible that the effect of ZPR1 to disrupt NPAT localization is a direct effect and that the cell cycle arrest is a secondary consequence of NPAT mislocalization because it is known that NPAT deficiency blocks S phase progression and arrests cells in both the G1 and G2 phases of the cell cycle (29, 30). Similarly, it is known that NPAT deficiency causes disruption of Cajal bodies; for example, the localization of p80 Coilin (29). The effect of ZPR1 deficiency to cause protein mislocalization within the nucleus could therefore be directly caused by NPAT mislocalization (Fig. 5B), but could also result from secondary effects of ZPR1 deficiency, including decreased RNA expression (Fig. 8).

Role of ZPR1 in RNA Expression—ZPR1 deficiency causes a profound inhibition of transcription (Fig. 7, B and C). This defect in gene expression is sufficient to explain the defect in cell cycle progression observed in ZPR1-deficient cells (Fig. 6A). The mechanism that causes transcription arrest is unclear. The mechanism may be a direct effect of ZPR1 deficiency, but it is likely that the transcription defect is a secondary consequence of ZPR1 deficiency, including altered nuclear architecture (Figs. 5 and 6). Because ZPR1 deficiency causes defects in the subcellular localization of snRNPs in HeLa cells lacking ZPR1 (Fig. 7D) and cells derived from Zpr1-/- embryos (4), it is possible that the effect of ZPR1 deficiency on RNA expression is a secondary consequence of defects in pre-mRNA splicing.


   FOOTNOTES
 
* This work was supported by grants from the Families of Spinal Muscular Atrophy and the Muscular Dystrophy Association (to L. G.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 To whom correspondence should be addressed: 373 Plantation St., Worcester, MA 01605. Tel.: 508-856-7844; Fax: 508-856-3210; E-mail: laxman.gangwani{at}umassmed.edu.

2 The abbreviations used are: SMN, survival motor neurons; SMA, spinal muscular atrophy; snRNP, small ribonuclear particles; PCNA, proliferating cell nuclear antigen; MEM, minimal essential medium; FBS, fetal bovine serum; FITC, fluorescein isothiocyanate; BrdUrd, bromodeoxyuridine; BrUrd, bromouridine; PBS, phosphate-buffered saline; DAPI, 4',6'-diamino-2-phenylindole; GAPDH, glyceraldehyde-3-phosphate dehydrogenase. Back


   ACKNOWLEDGMENTS
 
I thank Dr. Gary Stein for providing the histone H4/n probe, Dr. Roger J. Davis for discussions and suggestions, and Kathy Gemme for expert administrative assistance.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
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A. K. Mishra, L. Gangwani, R. J. Davis, and D. G. Lambright
Structural insights into the interaction of the evolutionarily conserved ZPR1 domain tandem with eukaryotic EF1A, receptors, and SMN complexes
PNAS, August 28, 2007; 104(35): 13930 - 13935.
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