Abstract
purpose. This study focuses on the identification of regulatory elements that contribute to lens-specific expression of the human CP49 gene within the 5′-flanking DNA sequences.
methods. The DNA sequence upstream of the human CP49 coding region was subcloned as a set of 5′ and 3′ deletion series. The constructs were transfected into lens (N/N1003A) and nonlens (NIH3T3) cell lines and chicken primary lens cultures, to test for promoter activity and specificity. To further test the specificity, a portion of the 5′ flanking DNA sequence was used to drive transgene expression in mice. The flanking DNA sequence was analyzed for potential transcription factor–binding sites.
results. The 5′-flanking DNA preferentially activated reporter gene expression in a lens-preferred manner when transfected into cultured cells. Transgene expression driven by the CP49 promoter region was lens specific. Analysis of the proximal promoter sequence revealed the presence of potential binding sites for the AP-1, AP-2, and OCT-1 transcription factors and the absence of TATA and CAAT boxes.
conclusions. The sequence upstream of the CP49 gene possesses promoter activity and is able to drive lens-preferred expression in both transfection and transgenic experiments. Promoter activity is dependent on the presence of the proximal 300 bp directly upstream of the coding region.
Ocular lens development commences with a thickening of the anterior surface ectoderm that establishes the boundary of the presumptive lens as the lens placode. Invagination of the thickened ectoderm creates the lens pit, which is then pinched off from the ectoderm as a hollow sphere of cells known as the lens vesicle. The epithelial cells at the posterior end of the lens vesicle undergo terminal differentiation to form primary lens fiber cells. During this process, the cells elongate to fill the lumen of the lens vesicle. The anterior portion of the lens is lined by a single layer of epithelial cells that remain mitotically active and undergo differentiation at the equator, or bow region, of the lens to continually add secondary fiber cells to the lens mass. In addition to cell elongation the differentiation process of lens epithelial cells to lens fiber cells involves the loss of the nucleus and other membrane-bound organelles and alterations in gene expression.
1
Lens fiber cell formation results in the upregulation of a limited collection of genes that includes members of the soluble crystallin gene family, major intrinsic protein (MIP), and the core components of the lens fiber–specific beaded filament CP49 and CP115.
2 The CP49 and CP115 genes have been classified as divergent members of the intermediate filament family on the basis of conserved primary DNA sequence and gene structure.
3 4 5 6 The expression of the beaded filament genes is unique to lens fiber cells, which allows the use of these genes as markers of differentiation. The CP49 and CP115 gene products are observed in lens fiber cells that are nearing full elongation.
7 Mutations in the CP49 have been linked to cataract formation, suggesting a role for the beaded filament, which is critical to lens clarity.
8 9
The characterization of crystallin promoters has identified a wide variety of transcriptional regulators involved in establishing the specific spatial and temporal expression patterns of the individual crystallin genes. Of these transcription factors, only a few are used in a widespread manner. Consensus-binding sites for Pax, Sox, and Maf factors have been located within the promoter-enhancer regions of several crystallin genes and are integral in achieving the high levels of expression observed in vivo.
10 11 12 13 The mutation or absence of any one of these factors in mice, accomplished through targeted gene deletion, results in abnormal eye and/or lens development, ranging from defective lens fiber cell differentiation to the complete absence of the lens.
13 14 15 16 17 Although the regulation of crystallin gene expression has been well characterized, knowledge of the mechanism(s) governing the expression of noncrystallin lens fiber cell genes is limited. It is not known whether regulators present in crystallin promoters influence lens fiber cell gene expression as a whole and also take part in the regulation of expression of CP49, CP115, and MIP, or whether a distinct set of factors is involved outside of the crystallin gene family.
In this study, the 5′-flanking and intronic DNA sequences of the human CP49 gene have been analyzed for potential regulatory elements. The transfection of cell lines and primary lens cultures and the production of transgenic animals were performed to test the proximal promoter for contributions to the cell type and differentiation-stage–specific expression pattern observed for the human CP49 gene. The results of these experiments suggest that the 5′-flanking DNA proximal to the human coding sequence confers lens-preferred expression of reporter genes. Sequence analysis of the 5′-flanking region does not identify consensus binding sites for transcription factors used in the regulation of crystallin genes, suggesting the possibility of separate control mechanism(s) for crystallin and noncrystallin gene regulation within the lens fiber cell.
5′-Deletion Series. DNA sequences flanking the 5′ end of the human CP49 gene were obtained from the screening of a human chromosome 3 HindIII λ-phage library. The isolated clone contained approximately 3 kb of upstream sequence, along with a portion of the coding sequence, and was shuttled into the pCR II vector by using PCR primers within the human CP49 rod 1B region and a primer flanking the insertion site of the phage. The entire portion of the 5′-flanking DNA was cloned into the pGL2-Basic vector (Promega, Madison, WI) as two restriction fragments, which resulted in the −3183/−19 clone, relative to the translation start site. The 5′ end of the sequence was excised as a 2.8-kb KpnI-BclI fragment from the pCRII clone. The 3′ end of the flanking sequence was subcloned into the pSP72 vector (Promega) as a blunted BanI fragment that spans the −911/−19 region of the upstream sequence. The proximal upstream sequence was then excised as a BclI-HindIII fragment and ligated, along with the 5′ end fragment, into the pGL2-Basic vector that had been cut with KpnI and HindIII. Subsequent deletion constructs were produced through restriction enzyme digestion of this parent clone or through PCR amplification of the 5′-flanking region. The integrity of the DNA sequence for PCR-amplified regions was confirmed by sequencing at the University of California Davis Division of Biological Sciences automated sequencing facility.
3′-Deletion Series. Oligonucleotide primers were obtained to manufacture a series of 3′ deletion constructs through PCR. Each 3′ deletion PCR primer was designed with a HindIII restriction site at the 5′ end. The 5′ oligonucleotide primer used in the production of the 3′ deletion series spanned the −382/−362 region of the 5′-flanking sequence and possessed a XhoI restriction site at its 5′ end. All the PCR reactions were cut with XhoI and HindIII and cloned into pGL2-Basic, cut with the same enzymes. Once again, DNA sequencing was used to confirm integrity of the inserted sequences generated by PCR.
Mouse CP49 Transfection Constructs. A P1 clone containing the coding region of the mouse CP49 gene and 5′-flanking sequence was obtained from Incyte Genomics (St. Louis, MO), through the screening of a mouse genomic P1 library, with CP49 exon I used as a probe. The 5′-flanking sequence of the CP49 gene was sequenced, and oligonucleotides were produced to generate the two constructs. The 5′ primer used spans the region of the flanking sequence from −924 to −903, and the 3′ primers correspond to the −38/−15 and −300/−277 portions of the upstream sequence. The primers possessed an additional 5′ sequence containing restriction enzyme sites to aid in subcloning (XhoI for the 5′ primer and HindIII for each of the 3′ primers). PCR amplification yielded two fragments that encompass the −924/−15 and −924/−277 flanking DNA sequences that were then cloned into the pGL2-Basic vector. Both clones were sequenced to verify that the amplified fragments were not altered.
N/N1003A Cells. The nontransformed rabbit lens epithelial cell line was maintained by culturing in Eagle’s minimal essential medium (EMEM; Sigma, St. Louis, MO) and 8% rabbit serum (Sigma) in a 37.0°C, 6.0% CO2, and humidified environment. Cells were seeded in 12-well plates (Falcon, Franklin Lakes, NJ) and grown to 60% to 80% confluence, typically within 24 to 30 hours, before the transfection was begun. Both serum and antibiotics were absent from the medium at the time of transfection. One hundred nanograms of pRL-TK (Promega), the internal standard, and 250 ng of each transfection construct was added to 50 μL of EMEM with nonessential amino acids. DNA condensing reagent (2 μL; Invitrogen, Carlsbad, CA) was then added to the diluted DNA, mixed, and incubated at room temperature for 15 minutes. Transfection reagent (2.5 μL; Lipofectamine; Invitrogen) was added to 50 μL of EMEM with nonessential amino acids. The transfection solution was then mixed with the DNA/Plus reagent solution and incubated at room temperature for 15 minutes. The cells were washed with 1 mL of EMEM with nonessential amino acids, and the medium was aspirated and replaced with 400 μL of fresh medium. The cells were covered with the DNA-transfection reagent mixture (100 μL) and incubated for 3 hours, at which time, 500 μL of fresh growth medium (EMEM supplemented with nonessential amino acids and 16% rabbit serum) was added to each well. At 24 hours after transfection, the medium was aspirated and replaced with 1 mL of EMEM complete with rabbit serum, nonessential amino acids, and gentamicin. The cells were harvested at 48 hours after transfection through addition of 250 μL of 1× passive lysis buffer from the a dual luciferase kit (Promega) and incubation for 15 minutes at room temperature. Lysates were cleared of particulate debris and stored at −80°C until analyzed.
NIH3T3 Cells. The cell line was maintained by culturing in Dulbecco’s minimum essential medium (DMEM; Invitrogen) supplemented with 25 mM HEPES (Invitrogen), 10% bovine calf serum (Hyclone, Logan, UT), and penicillin-streptomycin (Invitrogen). The cell line was cultured in the same environment as described earlier. The protocol for transfection of the NIH3T3 cell line was the same as that used for the N/N1003A cell line except that DMEM with HEPES was used and 3 μL of Plus reagent and 3 μL of transfection reagent replaced the volumes used in the N/N1003A protocol. At 3 hours, 500 μL of growth medium (DMEM with 20% bovine calf serum and 50 mM HEPES) was added, and, at 24 hours after transfection, the medium was replaced with 1 mL complete DMEM as described earlier.
Embryonic Chicken Lens Epithelial Cell Cultures. Lenses were harvested from 14-day-old chicken embryos and prepared for culturing as described by Menko et al.
18 Once isolated, the cells were cultured in medium 199 (Sigma) supplemented with 10% fetal calf serum (Sigma), cultured to confluence in a T-75 flask (Falcon, Franklin Lakes, NJ) coated with rat-tail collagen (Sigma), trypsinized, and seeded onto collagen-coated 12-well plates. The cells were grown until the proper confluence, 60% to 80%, was reached (36–48 hours), at which time the transfection was started. The protocol followed the same outline as that for NIH3T3 cells, with medium 199 replacing DMEM. Antibiotics and antifungal reagents were absent only for the initial 3-hour incubation at the start of the transfection. These reagents were then added with the growth medium. Two microliters of Plus reagent and transfection reagent were used for the transfection of the chicken lens epithelial cells.
Analysis of Transfected Cells. The lysates were assayed for each transfection experiment using the dual luciferase kit (Promega) and a luminometer (Dynex ML3000; Thermo Labsystems, Chantilly, VA). Twenty microliters of cell lysate was pipetted into individual wells of a flat-bottomed, 96-well microtiter plate (Microlite I; Thermo Labsystems). Luciferase assay reagent (100 μL, LARII; Promega) was dispensed into a well, and the luciferase activity allowed to integrate over a 10-second period. After the wells had been assayed, 100 μL of buffer (Stop n Glo; Promega) was dispensed into each well, and the luciferase activity from the internal standard pRL-TK was measured over a 10-second interval. Each construct within an experiment was transfected in triplicate. The summarized data are the result of at least three separate transfection experiments. Data were normalized within each experiment for each cell line and were used only when the internal standard levels were similar between the two cell lines, suggesting that the transfection efficiency across the experiments was similar and that differences in activity were not due to a significant disparity in transfection efficiency. The data from the luciferase experiments were then compared with the activity of the promoterless and enhancerless pGL2-Basic vector and expressed as multiples of increase over the activity of the pGL2-Basic vector.
Generation of the Transgenic Construct pβgal-Basic hCP49 -1859/-19. The −1859/−19 region of the 5′-flanking DNA sequence was shuttled from the pGL2-Basic vector into the pβgal-Basic vector (Clontech, Palo Alto, CA). The pβgal-Basic h49 −1859/−19 was then cut with XhoI and BamHI and the fragment gel purified. The inserted fragment contained all the h49 5′-flanking sequence and was ligated upstream of the coding region for the β-galactosidase gene along with an SV40 intron and polyA signal 3′ to the coding sequence. The fragment was microinjected into the fertilized pronuclei of a B6CBA mouse. Sixty-seven of the 250 injected pronuclei survived to term. Genomic DNA from the 67 mice was isolated by using a kit (DNeasy; Qiagen, Valencia, CA). A primer from the coding region of the β-galactosidase gene, within the pβGal-Basic vector corresponding to bases 228 to 252 of the vector sequence and a primer from the 5′-flanking DNA sequence of the human CP49 gene spanning the −227/−207 region were used in the PCR screen. PCR analysis confirmed the presence of the transgene within the genomic DNA of seven animals (three males and four females). The seven founder mice were then bred to a BL6 background. Offspring from the seven founder B6CBA transgenic mice and the BL6 mice were genotyped by PCR and subsequently analyzed for expression of the β-galactosidase reporter gene. All procedures involving animals were performed in compliance with the ARVO Statement for the use of Animals in Ophthalmic and Vision Research.
Tissue Lysate Preparation from Transgenic Animals. Samples were taken from several tissues and prepared in the following manner. The tissue samples were first rinsed with ice cold 1× PBS three times. A 1-mL aliquot of cell lysis buffer (100 mM potassium phosphate [pH 7.8], 0.2% Triton X-100, and 1 mM dithiothreitol [DTT]) was added to the tissue sample, and the samples were homogenized. The homogenate was shaken for 1 minute in lysis buffer after homogenization and subsequently centrifuged for 5 minutes at 13,000 rpm, to clear cellular debris. The supernatant was aspirated and total protein concentration was established by bicinchoninic acid (BCA) protein assay (Pierce, Rockford, IL).
β-Galactosidase activity from the tissue lysates was measured with a luminescent β-gal kit (Clontech). Thirty-five micrograms of total protein for each sample was pipetted into the wells of a flat-bottomed, 96-well microtiter plate. Two hundred microliters of reaction buffer and reaction substrate (Clontech) mixture was added to each sample and mixed. The reaction was then incubated for 1 hour at room temperature and assayed on a luminometer (Dynex ML3000; Thermo Labsystems) for a 10-second interval. The β-galactosidase levels were the mean results of at least three independent assays for each transgenic line.
Immunohistochemical Analysis of Transgenic Animals. The eyes of transgenic and wild-type animals were harvested and fixed in 4% paraformaldehyde in 0.1 M phosphate buffer at 4°C overnight. The tissue was then embedded in paraffin and sectioned for immunostaining. After deparaffinization and rehydration, the sections were treated for 15 minutes, at room temperature, with pepsin (Sigma), at a concentration of 1 mg/mL in 2.8% glacial acetic acid, followed by subsequent treatment with 3% H2O2. The tissue was then incubated in 10% normal goat serum (Invitrogen) for 30 minutes. The primary antibody, a rabbit polyclonal antibody to the bacterial β-galactosidase gene (Novus Biochemicals, Littleton, CO), was diluted 1:500 in PBS and the sections were exposed to the antibody solution for 90 minutes. After a wash in PBS, the sections were incubated with the secondary antibody solution, biotinylated goat anti-rabbit IgG (Zymed, Orange, CA), and then exposed to streptavidin-peroxidase (Zymed) for 15 minutes. Reactivity was visualized by a 3-minute incubation with 3,3 diaminobenzidine (DAB; Sigma). The sections were then counterstained in hematoxylin (Sigma), dehydrated, and coverslipped. For the preadsorbed primary antibody solution, an excess of purified bacterial β-galactosidase (Calbiochem, La Jolla, CA) was added to the diluted β-galactosidase antibody. The solution was then cleared by centrifugation.