In order to examine the function of HSF4 in lens formation in detail, we generated targeted disruption of the mouse Hsf4 gene by homologous recombination of 129S3 embryonic stem (ES) cells. In the vector, exons 3–5, which encode the DNA-binding domain, were replaced with the neomycin-resistance gene followed by the PGK cassette (Figure 1A). The ES cells (129S3 strain, derived from agouti) were electroporated with the linearized targeting vector under positive-negative selection 10, and eight correctly targeting clones were obtained (data not shown). ES cells with correctly targeting clones were injected into a C57Bl/6 blastocyst (black); the genotypes of the offspring were analyzed by PCR to identify wild-type (+/+), heterozygous (+/-), and homozygous (-/-) varieties (Figure 1B). As expected, the ratio of phenotypes was in accordance with Mendelian frequency. Hsf4^-/- mice are largely normal across all developmental stages except for cataract formation. Under slit-lamp detection, we observed a cataract phenotype in the Hsf4 knockout mouse (Figure 1C). Opacity of the lens of the Hsf4^-/- mouse appeared at an early postnatal stage and increased with age.

The structure of the Hsf4^-/- lens fibers became loose, and a vacuole-like cavity that appeared in lens fiber cells at day E15.5 (data not shown) became severe compared to wild-type; undegraded nuclei were clearly seen under a light microscope. However, the bow region of the lens, where the lens epithelial cells differentiate into fibers, was generally normal (Figure 1C). SEM images revealed loose fibers in the lenses of Hsf4 knockout mice that showed much less interaction than those of their wild-type counterparts (Figure 1D).

The Hsf4^-/- mouse lens is fragile and much lighter, but of a similar size, compared with its wild-type counterpart (Figure 2A). More than 90% of lens proteins are in the soluble form and include a variety of crystallins; in mammals, the crystallins are αA and αB; βB1, βB2, βB3, βA3/A1, and βA4; γA, γB, γC, γD, γE, γF, and γS. In mice, the γ-crystallins are the major contributor to the weight of the mature lens proteins. In humans, β-crystallins and γ-crystallins are in almost equal proportions, with a slightly lower amount of α-crystallins. Fujimoto et al. detected markedly reduced expression of γ(A-F)-crystallin genes in adult HSF4-null mice, even at 2 days old, but the expression levels of γ(A-F)-crystallin genes were normal in the lens of E15.5 HSF4-null embryos. They used chromatin IP (ChIP) analysis to show that HSF4 binds upstream of the γF-crystallin gene, suggesting that HSF4 regulates expression of the γF-crystallin genes 8. In contrast, Min et al. did not observe a significant reduction of γ-crystallin genes in 1-day-old to 28-day-old Hsf4-null mice, except the γF-crystallin gene, which showed reduced expression in 10-day-old Hsf4-null mice compared to wildtype mice 9. These conflicting results may be due to differences in Hsf4-null mice construction, differences in the genetic background of the mice being tested, and other unknown factors. However, neither Min et al. nor Fujimoto et al. studied another important crystallin protein, γS-crystallin. During maturation of the mouse lens, the level of γS-crystallin increases fivefold, replacing other types of crystallins and accounting for up to 15% of the total weight 11. This finding prompted us to examine the expression of γ-crystallins (especially γS-crystallin) in the lens of knockout and wild-type mice. We quantified the mRNA expression of γ-crystallins and assayed the transcriptional activity of Hsf4. We found that Hsf4^-/- mice have reduced expression levels of all subtypes of γ-crystallins directly after birth compared with wild-type mice, and there was almost a complete lack of γ-crystallin expression at 8 weeks old. In wild-type mice, the level of γS-crystallin mRNA indicates that γS-crystallin is only a small proportion of total γ-crystallin at birth, and the level increases to make it the major γ-crystallin at 8 weeks old (Figure 2B). The HSF4 and γS-crystallin genes are first expressed at the late embryonic stages and are up-regulated in late embryonic development 1213. The γ(A-F)crystallin genes are located in a gene cluster and are regulated by Sox1 and Maf, but the regulation of γS-crystallin gene (Crygs) has not been well characterized 14151617. To further evaluate the role of HSF4 in the regulation of γS-crystallin expression, we analyzed the expression of γS-crystallin mRNA in the human lens epithelial cell line SRA01/04 under conditions of hHSF4b overexpression 18. The γS-crystallin mRNA expression level was tenfold higher in hHSF4b-overexpressing cells than in the control (Figure 2C). As the γS-crystallin promoter region contains a heat-shock element (Crygs-HSE) that is less conserved than in the promoters of γ(A-F)-crystallin (Figure 2D), we asked whether HSF4b can interact directly with the endogenous promoter of CRYGS to regulate its expression. We used ChIP assays to test this hypothesis. The immunoprecipitation of solubilized chromatin prepared from HSF4b-expressing SRA01/04 cells with an anti-HSF4b antibody was followed by PCR using primers that target the potential HSF4b-binding site in the CRYGS promoter region. PCR yielded the expected band in the cells expressing HSF4b, and no band for the normal goat IgG control (Figure 2E). Additionally, we assumed that HSF4b would bind CRYGS-HSE to activate γS-crystallin transcription. Under this assumption, the CRYGS-luc vector, which contains an upstream sequence of the γS-crystallin gene, was constructed, and luciferase activity levels with and without overexpression of HSF4b were determined in the SRA01/04 cell line. The relative luciferase activity with overexpression of HSF4b was about sixfold greater than that of the system without overexpression. The dominant-negative HSF4b, which had lost the DNA-binding domain of HSF4b, had no significant effect on luciferase activity (Figure 2F). These results indicate that HSF4b could regulate the expression of γS-crystallin by binding to Crygs-HSE.

The γS-crystallin mutant rncat is a recessive cataract mouse model with a G to A transition point mutation at position 489 in exon 3 of the γS-crystallin gene 19. In heterozygous rncat mice, the lens is nearly normal. Our results suggested that HSF4 regulates the expression of the γS-crystallin gene (Figure 2). If this is true, lack of HSF4 would decrease the production of the wild-type γS-crystallin and might facilitate the cataract development. As reference, lens opacity of the rncat mouse appeared at 11 days old in the nuclear region 19. Interestingly, when we intercrossed the Hsf4^-/- mouse with the rncat mouse, the offspring (Hsf4^-/-/rncat/+) developed cataract in the posterior part of the lens, whereas the lenses of the Hsf4^+/+/rncat/+ mice remained basically clear (Figure 3A). The cataract in double null Hsf4^-/-/rncat/rncat mice developed earlier, appearing at 7 days old, and seemed more severe than that in Hsf4^+/+/rncat/rncat mice (Figure 3A). Lack of Hsf4 worsens the lens fiber defect in γS-crystallin mutation mouse rncat [see Additional file 1]. The mRNA levels of γS-crystallin in the Hsf4^-/-/rncat/+ and Hsf4^-/-/rncat/rncat mice were reduced by more than 90% compared with the Hsf4^+/+ and rncat/rncat mice (Figure 3B). To certain degree, our results further suggest that Hsf4 disruption reduces Crygs expression along with the cataract formation in the offsprings in such intercross.

An important aspect of Hsf4^-/- cataract formation is the abnormal development of lens fibers that cannot be explained completely by crystallin regulation. The loose fiber structure of the lens of the Hsf4 knockout mouse is similar to that of the Bfsp1/2 knockout mouse. Knockout of Bfsp1 or Bfsp2 resulted in a loose fiber structure with fewer connection proteins among the fibers 2021. Therefore, we quantified the levels of Bfsp1 and Bfsp2 mRNA in the lens of Hsf4 knockout mice. At birth, the level of Bfsp2 mRNA in Hsf4 knockout mice was half that of wild-type mice, whereas the level of Bfsp1 mRNA was similar to that of wild-type mice. At 8 weeks old, the levels of both Bfsp1 and Bfsp2 mRNA in the lens of Hsf4 knockout mice were reduced by more than sevenfold compared with wild-type mice, except for the level of Bfsp1 mRNA at birth (Figure 4A). It has been suggested that the 129S3 strain mouse has a deletion that causes the loss of exon 2 from Bfsp2 mRNA and dramatically reduces mRNA levels of Bfsp2. The Bfsp2 protein in this strain was undetectable by antisera to the wild-type protein 2223. Since we obtained the expected PCR product size from Hsf4^-/- mice using primers from the deletion region of the Bfsp2 gene (Figure 4B), there is at least one copy of the C57BL/6J Bfsp2 allele in the Hsf4^-/- mouse. Although Hsf4^-/- mice carry at least one copy of the highly expressed C57BL/6J Bfsp2 allele, Hsf4^-/- mice had a much lower level of expression of Bfsp2 compared to the 129S3 wild-type mice. Therefore, lack of Hsf4 results in reduced expression of Bfsp2.

Overexpression of hHSF4b in SRA01/04 cells up-regulated transcription of Bfsp1 and Bfsp2, and this up-regulation could be inhibited specifically by short hairpin RNA (shRNA) targeting HSF4b (Figure 4C). As both Bfsp1 and Bfsp2 promoters have the less-conserved HSE sequence (Figure 4D), we asked whether HSF4 can directly bind the promoter region of Bfsp1 and Bfsp2. We used ChIP assays and detected the expected bands of the promoter sequences of the Bfsp1 and Bfsp2 genes in cells expressing HSF4b (Figure 4E). This result suggests that HSF4 can bind to the promoter regions of the Bfsp1 and Bfsp2 genes. We then used the dual-luciferase system to assess whether HSF4 activates the expression of Bfsp1 and Bfsp2. Human Bfsp1-luc or Bfsp2-luc vectors were co-transfected with or without HSF4b into SRA01/04 cells. The relative luciferase activities of HSF4b co-transfected with Bfsp1-luc or Bfsp2-luc were approximately eightfold that of cells transfected with Bfsp1-luc or Bfsp2-luc alone (Figure 4F). These results indicate that HSF4 may bind specifically to the Bfsp1-HSE/Bfsp2-HSE sequences and regulate expression of the intermediate filament proteins Bfsp1 and Bfsp2.

To analyze changes in the lens components of HSF4^-/- mice, we used 2D electrophoretic (2D-E) analysis to generate maps of lens lysates from 8-week-old Hsf4^-/- and wild-type (129S3) mice, and identified selected spots using liquid chromatography (LC) with tandem mass spectrometry (MS/MS). By comparison with published 2D-E maps of the mouse lens 1124, it was possible to identify most of the crystallin proteins in the 2D-E maps of both Hsf4^-/- and wild-type mice. The lens of Hsf4^-/- and wild-type mice generally had similar 2D-E protein expression patterns with silver staining. However, alterations of some spots were observed in the 2D-E map of the Hsf4^-/- lens, which generally had less crystallin protein than the wild-type lens (Figure 5A). We selected and identified some altered spots in the lens map of Hsf4^-/- mice, where signals were either absent or weaker than in wild-type mice (Figure 5B). Interestingly, 5 of 21 spots were identified by MS as αA-crystallin. These αA-crystallin molecules had a molecular mass of 15–25 kDa and migrated to the acidic region (PI 4.5–6.0) of the 2D-E gel. Other differentially expressed spots were identified as αB-crystallin, βA1-crystallin, βB2-crystallin, γC-crystallin, γB-crystallin, heat shock protein 27, and 7 unknown proteins. Due to the limited amount of sample, the terminal sequences of selected spots were not characterized. A similar pattern has been reported during aging or cataractogenesis in mice 24. Truncation of αA-crystallin is probably caused by the activation of a class of calcium-activated proteases known as calpains, such as Lp82 or calpain2 2526, which have 'clipping' activity in vitro. We used real-time PCR to determine the levels of Lp82 and calpain2 mRNAs in the lens of Hsf4 knockout mice and of wild-type mice. The mRNA levels of Lp82 and calpain2 in Hsf4 knockout neonatal mice were reduced fourfold compared with wild-type mice; this reduction was even greater in 8-week-old mice (Figure 5C). Therefore, loss of αA-crystallin and reduced expression of Lp82 and calpain2 may be correlated with Hsf4 disruption.

A clone containing the entire mouse Hsf4 gene was isolated from a 129S3 mouse genomic library (Genome Systems). From this, a 2.8 kb 5'-flanking region EcoRI/KpnI fragment containing part of the promoter region was subcloned into X-pPNT. In X-pPNT, exons 3–5 and part of the promoter region were replaced by a 1.6 kb neomycin-resistance gene fragment. A downstream 5.0 kb SalI/NotI fragment was inserted into X-pPNT. These linearized segments were inserted into X-pPNT to generate pPNT-HSF4, which was linearized with EcoRI/NotI and electroporated into R1 ES cells. Two ES clones with a targeted disruption were identified from 131 clones isolated by the positive-negative G418/gancyclovir selection method. Targeted clones were identified by PCR and Southern blot analysis after HindIII digestion of genomic DNA, using a flanking probe 3' to the replacement. The ES clones were microinjected into C57Bl/6 blastocysts, and five chimeric mice were produced. These chimeras were bred with C57Bl/6 mice to generate progeny heterozygous for the mutation. Heterozygous mice were crossed to produce Hsf4^+/+, Hsf4^+/-, and Hsf4^-/- mice. Germline transmission of the mutations was determined by PCR analysis of tail DNA, as described above. All animal experiments were conducted under protocols approved by institutional Animal Care and Use Committees and according to the National Institutes of Health "Using Animals in Intramural Research" guidelines.

Mouse eyes were dissected, fixed in 4% paraformaldehyde for at least 2 weeks, embedded in paraffin, and cut into 6 μm thick sections. These paraffin-embedded sections were stained with hematoxylin and eosin.

The mice were sacrificed, the eyes were enucleated, and the corneas were removed. The remaining orbit was immediately placed into a fixative of 2% glutaraldehyde in 0.1 M sodium cacodylate buffer at pH 7.4. Tissues were fixed at 4°C for 2–3 days, with daily changes of the fixative. After washing overnight in 0.2 M sodium cacodylate buffer, the lens was removed from the posterior portion of the eye containing the retina. To expose the fibers and suture patterns, the lens capsule and superficial fibers were peeled away from the lens around its diameter. Both the fiber peelings and the remaining lens core were collected and processed for scanning electron microscopy (SEM). Specimens were post-fixed in 1% aqueous OsO₄ at 4°C for 12 hours and washed in 0.1 M sodium cacodylate buffer at pH 7.4. They were dehydrated in a series of graded ethanol and processed by critical point drying. A layer of gold 20 nm thick was deposited onto the surface of the dried samples, which were then examined using a Hitachi S-520 scanning electron microscope at 20 kV.

RNA was isolated using a Qiagen RNA-easy Mini Kit (Qiagen). A 1 μg sample of total RNA was reverse transcribed at 42°C for 1 hour in a total volume of 20 μL (Promega), with oligo(dT)₁₅ primers. At the end of the reaction, the mixture was heated at 70°C for 15 minutes to inactivate the reverse transcriptase. For Q-PCR (Table 1), 1 μL of cDNA was amplified for 45 cycles with a master mix (SYBR Green Supermix; ABI) using a thermocycler (MG). Melting curve analysis was done at the end of the reaction (after 45 cycles) to assess the quality of the final PCR products. The threshold cycle C(t) values were calculated by fixing the basal fluorescence at 0.05 unit. Five replicates were used for each sample, and the average C(t) value was calculated. The ΔC(t) values were calculated as C(t) sample – C(t) β-actin. The N-fold increase or decrease in expression was calculated by the ΔΔCt method using the fetal C(t) value as the reference point. The N-fold difference was determined as:

2^{-(ΔC(t)sample-ΔC(C(t)fetal)}

Electrophoresis of extracts was done under denaturing reducing conditions using 4%–12% Nupage Bis-Tris gels with Mops-SDS running buffer (Invitrogen). Approximately 30–50 μg of protein was used per lane for analysis as described in the protocol. Electrophoresis was for approximately 35–60 minutes at 200 V. After electrophoresis, the proteins were transferred to nitrocellulose membranes (Amersham) using the Surelock X-cell transfer system (Invitrogen). The blotted membranes were labelled with antibodies against HSF4 or HSF4b (Santa Cruz). The blots were washed in TBS/T, incubated using the ECL kit (Cell Signaling Technology. Inc.), and exposed to X-ray film (Kodak).

Human lens epithelial cells (SRA01/04) were grown and cultured in Dulbecco's modified Eagle's medium (DMEM) with low glucose (1 mg/ml) and supplemented with 15% fetal bovine serum, gentamicin (50 units/mL; Fluka), and a penicillin-streptomycin antibiotic mix (50 U/ml; PAA) at 36.5°C in an atmosphere of 5% CO₂. The methods for establishing primary cultures of human lens epithelial cells were as described 18. Cells from explant cultures were dissociated using trypsin-EDTA solution (PAA) and collected by centrifugation. The cells were subcultured in the initial medium at 36.5°C in a humidified atmosphere with 5% CO₂. The procedure was repeated for additional subcultures.

Cells were cultured as described above. Transient transfections were done using FuGENE 6 reagent (Roche) essentially according to the manufacturer's protocol but with minor modifications. SRA01/04 cells were plated in six-well dishes of regular medium at a density of 2 × 10⁶ cells/ml. A 2 μg sample of plasmid DNA was prepared in 100 μl of serum-free, antibiotic-free medium and incubated with 6 μl of FuGENE 6 reagent prepared separately in 100 μl of serum-free, 15% FBS medium. After 30 minutes, the DNA mix was added to the cells, which were then incubated for 24 hours at 37°C. The cells were washed in phosphate-buffered saline, supplemented with DMEM with 15% FBS, and incubated overnight at 37°C. In co-transfection experiments, the appropriate empty vector DNA was used to ensure similar concentrations of DNA under all conditions. In all transfections, 0.1 μg of Renilla luciferase plasmid (Promega, Madison, WI) was included to control for transfection efficiency. Protein concentration was determined by the Bradford assay. Firefly and Renilla luciferase activities were quantified using the Dual Luciferase kit (

Promega

Preparation of chromatin DNA from SRA01/04 cells expressing HSF4b or control plasmid pCDNA3.1(+) and subsequent ChIP assays were done with a chromatin immunoprecipitation assay kit (Upstate Biotechnology, Waltham, MA) following the manufacturer's instructions. HSF4b/DNA complexes were precipitated by anti-HSF4b antibody. Precipitated DNA was amplified by PCR using primers flanking the potential HSF4-binding site (positions -207 to -64) in the CRYGS promoter, positions -308 to -61 in the Bfsp1 promoter, or positions -351 to -81 in the Bfsp2 promoter. Goat IgG was used as the negative control. The primers are given in Table 1. The PCR products were resolved by electrophoresis in a 1.5% agarose gel.

Ten lenses from identical 8-week-old mice were homogenized by sonication in 1 ml of lysis solution containing a protease inhibitor cocktail (9.8 M urea, 2% Chaps, 1% DTT). The protein concentration was measured by Bradford assay, using bovine serum albumin as the standard, and samples were stored at -70°C.

Isoelectric focusing was done using immobilized pH gradient (IPG) gel strips (13 cm, pH 3–10), followed by molecular mass-based resolution by SDS-12% PAGE (Amersham). The silver-stained gel images were captured and image analysis was used (Imagemaster, Amersham) to determine the contribution of each spot to the total protein. The isoelectric point (pI) of each unmodified crystallin was extrapolated using the calculated pI and position as references. The spots captured from a Coomassie brilliant blue-stained gel were washed, dried, and digested with trypsin; then LC-MS/MS was used to determine the amino acid sequence of the peptides.