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Apoptosis (2018) 23:65–78 https://doi.org/10.1007/s10495-017-1437-4

ORIGINAL PAPER

Caspase cleavage of transcription factor Sp1 enhances apoptosis

Behzad Torabi1 · Samuel Flashner1 · Kate Beishline1 · Aislinn Sowash1 · Kelly Donovan1 · Garrett Bassett1 · Jane Azizkhan‑Clifford1

Published online: 13 December 2017 © Springer Science+Business Media, LLC, part of Springer Nature 2017

AbstractSp1 is a ubiquitous transcription factor that regulates many genes involved in apoptosis and senescence. Sp1 also has a role in the DNA damage response; at low levels of DNA damage, Sp1 is phosphorylated by ATM and localizes to double-strand break sites where it facilitates DNA double-strand-break repair. Depletion of Sp1 increases the sensitivity of cells to DNA damage, whereas overexpression of Sp1 can drive cells into apoptosis. In response to a variety of stimuli, Sp1 can be regulated through proteolytic cleavage by caspases and/or degradation. Here, we show that activation of apoptosis through DNA dam-age or TRAIL-mediated activation of the extrinsic apoptotic pathway induces caspase-mediated cleavage of Sp1. Cleavage of Sp1 was coincident with the appearance of cleaved caspase 3, and produced a 70 kDa Sp1 product. In vitro analysis revealed a novel caspase cleavage site at aspartic acid 183. Mutation of aspartic acid 183 to alanine conferred resistance to cleavage, and ectopic expression of the Sp1 D183A rendered cells resistant to apoptotic stimuli, indicating that Sp1 cleavage is involved in the induction of apoptosis. The 70 kDa product resulting from caspase cleavage of Sp1 comprises amino acids 184–785. This truncated form, designated Sp1-70C, which retains transcriptional activity, induced apoptosis when overexpressed in normal epithelial cells, whereas Sp1D183A induced significantly less apoptosis. Together, these data reveal a new caspase cleavage site in Sp1 and demonstrate for the first time that caspase cleavage of Sp1 promotes apoptosis.

Keywords Sp1 · Apoptosis · Protein stability · Transcription factor · DNA damage · Caspase cleavage · Extrinsic pathway · Intrinsic pathway

Introduction

Sp1 recognizes DNA elements known as GC boxes through its C-terminal, zinc-finger DNA-binding domain [1]. In the absence of a TATA box, Sp1 participates in the recruitment of specific transcription factors, general transcriptional machinery, and RNA polymerase II, as well as chromatin remodeling factors in order to facilitate initiation of tran-scription [2, 3]. Sp1 is ubiquitously expressed and mouse knockout is embryonic lethal at day 11 [4]. Sp1 regulates the transcription of a large number of genes that are involved

in cell proliferation and different signaling networks; many of the Sp1-regulated genes have opposing functions (e.g. pro- and anti-apoptotic, oncogenes and tumor suppressors). Sp1 activity is modulated by post-translational modifications that dictate the specific gene program. Since its discovery, several different post-translational modifications have been reported, including phosphorylation [5], acetylation [6], O-linked glycosylation [7–10], and SUMOylation [11]; these post-translational modifications enhance and/or reduce Sp1 transcriptional activity [5].

Sp1 is involved in the cellular response to DNA damage. DNA damage is sensed and monitored by the cell in order to maintain genomic stability. If DNA damage is too exten-sive to repair, the cell will undergo apoptosis. In response to DNA damaging agents that induce double strand breaks (DSBs), Sp1 localizes to break sites, is phosphorylated by ATM, and is required for DSB repair [12]. Cells depleted of Sp1 are more sensitive to DNA damage, which has been related to its ATM-dependent phosphorylation [13]. Sp1 has also been implicated in initiation of apoptosis.

Electronic supplementary material The online version of this article (https://doi.org/10.1007/s10495-017-1437-4) contains supplementary material, which is available to authorized users.

* Jane Azizkhan-Clifford Jane.Clifford@drexelmed.edu

1 Department of Biochemistry and Molecular Biology, Drexel University College of Medicine, Philadelphia, PA 19102, USA

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Overexpression of wild-type Sp1 has been shown to induce apoptosis in some cells [14–16]. In smooth muscle cells, Sp1 expression and phosphorylation were required for apoptosis through upregulation of Fas ligand [17, 18]. Furthermore, overexpression of a series of Sp1 deletion mutants revealed that the Sp1 DNA-binding domain was necessary for the activation of cell death [14], suggesting that these effects may be linked to transcription. Pro-apoptotic genes, such as Bax, TRAIL, fas, fas-ligand, caspase 8, and caspase 3 [14, 19], as well as anti-apoptotic genes such as Survivin and Bcl-2 [20–23] are regulated by Sp1. In addition, Sp1 activates transcription of caveolin-1, which is involved in the progression of senescence through an unknown mechanism [24, 25]. Apoptosis and senescence are safeguards against genomic instability [26].

Sp1 is also regulated in part through stability; however, the regulation of its stability is very complicated. Increased Sp1 stability is attributed to increases in to its phosphoryla-tion and glycosylation [11, 27, 28]. Decreases in Sp1 sta-bility can occur in response to glucose starvation, where-upon Sp1 was shown to undergo endo-proteolytic cleavage between amino acids 58–70 to facilitate proteasome-depend-ent degradation [27]. Degradation of Sp1 did not appear to require ubiquitination, although these experiments were done in vitro. Sp1 also contains a potential PEST sequence which could facilitate targeting Sp1 to the proteasome [29]. Additionally, caspases have been shown to cleave Sp1, including in T cells wherein apoptosis was induced by reti-noids [30]. Cleavage of Sp1 by caspase 3 and 7 during B-cell apoptosis has been reported, and one of the cleavage sites was mapped to aspartic acid 590 [31]. Other mechanisms of Sp1 destabilization include lysine 16 SUMOylation, which is linked to RNF-4-mediated degradation of Sp1 [32], and serine 59 phosphorylation by Cdk, which reportedly modu-lates SUMOylation and degradation [32].

We have discovered that induction of apoptosis by TRAIL ligand or high levels of DNA damage induces caspase-medi-ated cleavage of Sp1 at a previously uncharacterized site, aspartic acid 183. Cleavage at D183 results inaccumulation of the 70 kDa product comprising amino acids 184–785, which has previously been shown to be transcription-ally active [33]. Mutation of D183 reduced apoptotic cell death. Together, our data show that Sp1 cleavage at D183 is actively involved in apoptosis and is not simply a byproduct of cell death.

Materials and methods

Cell culture

U2OS, MDCK, MCF-7, NHDF (Normal Human Dermal Fibroblasts, Clonetics), and 293T cells were cultured in

DMEM supplemented with 10% FBS (Gemini Bio-prod-ucts cat# 900-108), l-glutamine (2 mM), sodium pyruvate (110 mg/L), 0.2 mg/mL penicillin, 60 ulg/mL streptomycin at 37 °C in 10% CO2. Human retinal pigment epithelial cells (ARPE-19; ATCC) utilized the same additives as the cell types above, with additional supplements including 4 mM L-glutamine, 50 nM insulin, and 15 ng/mL endothelial cell growth factor; ARPEs were cultured in DMEM containing 1 g glucose/mL and 5% CO2. The media from MCF-7 cells was also supplemented with 10 µg/mL insulin. MCF-10A cells were maintained in DMEM/F12 50:50, supplemented with 2  mM l-glutamine, 5% Horse Serum, 0.5  µg/mL Hydrocortisone, 10 µg/mL insulin, 0.1 µg cholera toxin/mL (Sigma), and 20 ng/mL EGF at 37 °C in 5% CO2. Retroviral packaging cells (GPG-293 cells) were maintained in DMEM supplemented with 10% FBS (heat inactivated), l-glutamine (2 mM), tetracycline (1 µg/mL), puromycin (2 µg/mL), and G418 (0.3 mg/mL). GPG-293 cells were maintained by split-ting 1:6 every 3–4 days and cultured at 37 °C in 5% CO2.

Transfection

U2OS, GPG-293 and 293T cells were transfected using Lipofectamine 2000 (Invitrogen) or Gendrill (Bamagen) following manufacturers’ protocols. Cell were trypsinized and resuspended at a concentration of 200,000 cells/mL. GPG-293 and 293T cells (2 × 106) were plated in 10 mL media in a 10 cm plate for the purposes of viral production. U2OS cells were plated at 2 mL/well in a standard 6 well plate (4 × 105 cells). Per 200,000 cells plated, 1 µg of DNA was mixed with 50 µL Opti-MEM (Invitrogen) and 3 µL of transfection reagent, followed by mixing with 50 µL of Opti-MEM and incubating for an additional 20 min before drop-wise addition to cells. A ratio of 1 µg of DNA to 3 µL of transfection reagent was maintained for all transfections. Media was changed the following day when using Gendrill or after 5 h when using Lipofectamine 2000. All transfec-tions took place in media without antibiotics.

Retroviral production

GPG-293 cells were used for the production of retroviruses expressing Sp1 and Bcl-2. The Sp1 constructs were in an LXSN retroviral vector, and Bcl-2 was in pBABE vector. GPG-293 selection media was replaced with media without antibiotic selection just prior to transfection. Transfection was carried out as described above. 5, 6, and 7 days post-transfection, viral media was collected and filtered through a 0.45 micron filter (Pall Life Sciences) and fresh media without antibiotics was added. Hexadimethrine bromide (8 µg/m), also known as polybrene (American Bioanalyti-cal), was added to the virus, which was aliquoted and stored at − 80 °C and thawed at room temperature prior to use.

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Lentivirus production

For shRNA-mediated gene silencing, we used two differ-ent Sp1-specific shRNAs, targeting either nt1805 or nt7276 of Sp1 cDNA (TRCN0000020448 and TRCN0000020444, respectively, Sigma-Aldrich). The nt7276 construct targets the 3′ UTR, which allows for expression of ectopic Sp1 with endogenous Sp1 knocked down. A control non-targeting vector from Sigma-Aldrich (Cat# SHC002) was used. Sp1, Sp1-D183A, and Sp1-70C cDNA were subcloned into an empty pLENTI backbone, a gift from Pantelis Tsoulfas (Addgene plasmid # 36097) [34]. The production of lenti-virus was carried out in 293T cells. To produce virus, 293T cells were plated 48 h prior to transfection in antibiotic-free media in a 10 cm plate. Cells (at ~ 80% confluence) were then transfected with 6 µg of vector, 2 µg of pRSV-REV, 2 µg of pMDLg/pRRE, and 2 µg of VSV-G (Addgene). Transfection was carried out as described above and virus was collected and filtered (0.45 micron) 48 h post-transfec-tion. Hexadimethrine bromide was added to obtain a con-centration of 8 µg/mL.

Viral transduction

MCF-10A cells were trypsinized and re-suspended in MCF-10A media at a concentration of 8 × 105 cells/mL. 500 µL of cell suspension was re-suspended in 1 mL of viral super-natant and incubated for 16 h at 37 °C in 1 well of a 6 well plate, followed by replacement of virus-containing media with 2 mL MCF-10A growth media. 48 h post-transduction, cells were selected with 0.3 mg/mL G418 for the LXSN Sp1 constructs and/or 1 µg/mL puromycin for shRNA knock-down. Cells were allowed to recover for a period of 1 week after Sp1 transduction to obtain a population with uniform Sp1 expression.

Caspase 3 knockdown by siRNA

MCF-10A cells were trypsinized and re-suspended at a concentration 100,000 cells/mL and 2 mL of cell suspen-sion was added to each well of a 6 well plate. Twenty-four hours after plating, 8 µL of Oligofectamine (Invitrogen) was combined with 22 µL of Opti-MEM (Invitrogen) for each well and incubated for 5 min. In a separate tube, 200 pmol (10 µL of a 20 µM stock) of caspase 3 siRNA (Dharmacon Cat#L-004307-00-0010) was combined with 160 µL of Opti-MEM and allowed to incubate for 5 min. The two tubes were then mixed, and the mixture was incubated for 20 min at room temperature in the dark. Media was removed from cells and replaced with 750 µL of fresh media. The 200 µL transfection mixture was then added dropwise to cells. Five

hours post-transfection, the media was replaced with 2 mL of growth media. Knockdown was measured 72 h post-trans-fection via western blot.

Apoptosis induction

MCF-10A cells were treated with TRAIL ligand (Peprotech Cat# 310-04) at 100 or 200 ng/mL final concentration. For UV treatments, media was removed before treatment with Spectrolinker XL-1000 UV cross-linker (Spectronics Cor-poration). Following UV exposure, media was immediately replaced on cells. All drugs were prepared in DMSO and added to cells as indicated in figure legends and/or results.

Senescence‑associated β‑galactosidase staining

NHDF cells were washed with PBS after 24 h treatment with UV. Cells were fixed in 2% formaldehyde/0.2% gluta-radlehyde in PBS for 5 min at room temperature. Cells were then washed with PBS twice prior to staining. Cells were stained with 40 mM Na2HPO4 (pH6.0 with HCl), 150 mM NaCl, 2 mM MgCl2, 5 mM K3Fe(CN)6, 5 mM K4Fe(CN)6, and 1 mg/mL X-gal. After 5 h of staining, cells were washed with PBS and imaged with an inverted phase microscope with a camera attached.

Site‑directed mutagenesis

Site directed mutagenesis to induce mutations D183A, D590A and D183A/D590A was performed with the Strata-gene Quick Change site directed mutagenesis kit following manufacturer’s protocol. D183A mutations were made using forward primer: 5′-CCC ACA GTT CCA GAC CGT TGC TGG GCA ACA GCT AGCA GTT TGC TGC-3′, reverse primer 5′-GCA GCA AAC TGC AGC TGT TGC CCA GCA ACG GTC TGG AAC TGT GGG-3′. D590A mutations were made using forward primer: 5′-GGA GAA AAC AGC CCA GCC GCC CAA CCC CAA GCC GG-3′ and reverse primer: 5′-CCG GCT TGG GGT TGG GCG GCT GGG CTG TTT TCT CC-3′.

Caspase cleavage assay

DNA fragments containing pLXSN-Sp1-HA with D183A, D590A, or D183A/D590A mutations were prepared with insertion of the T7 promoter and terminator with appropri-ate Kozak sequence via PCR. The forward primer 5′-GGC TAA TAC GAC TCA CTA TAG GGA CCA TGG ACT ACA AAG ACG ATG ACG ACA AGC TTG-3′ and reverse primer 5′-GGC TAT GCT AGT TAT TGC TCA GCT AGC TAG CGT AAT CTG GAA CAT CGT ATG GGT AGA AGC C-3′ were incubated with pLXSN-Sp1 vectors car-rying WT-SP1, D590-Sp1, or D183/D590 Sp1 sequence.

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PCR products were directly added to the TnT® T7 Quick for PCR DNA (Promega, product # L5540) supplemented with radioactive 35S labeled methionine following Manufacturers’ protocols. 10 µL of the TNT product was incubated with 10 µL (500 units) of purified active cleaved caspase-3 (Enzo Life Sciences) and incubated for 3 h at room temperature with 30 µL of cleavage buffer (10 mM Hepes pH 7.4, 0.1% CHAPS, 5 mM EDTA, 2 mM DTT). The control samples were incubated at room temperature with 40 µL of cleavage buffer. After incubation, 10 µL of 6× SDS sample buffer (0.35 M Tris–HCl, pH 6.8, 30% glycerol, 0.1 g/mL DTT, 0.1 g/mL SDS, 0.12 mg/mL bromphenol blue) was added to the reactions and the samples were boiled for 5 min at 95 C, followed by SDS-PAGE. Samples were then visualized by autoradiography.

In vitro cleavage assay of bacterially‑expressed GST‑Sp1

Protein was produced in BL21 E. coli. Cells were lysed in lysis buffer (50 mM Hepes pH7.4, 250 mM NaCl, 10 mM EDTA, 0.1% NP-40, 2  mM DTT, 10  mM NaF, 50  µM NaVO4, 5 mM Beta-GP, 1 mM PMSF, 10 µg/mL Leupep-tin, 15 µg/mL aprotinin, 8 M Urea). Solubilized lysate was diluted 10-fold with buffer and GST-Sp1 was purified with Glutathione Sepharose (Amersham Biosciences) for 1 h at 4 °C while rocking. 30 µL aliquots of beads were re-sus-pended in 20 µL of caspase cleavage buffer (10 mM Hepes, pH 7.4, 0.1% CHAPS, 5 mM EDTA, 2 mM DTT). Aliquots of GST-Sp1 bound beads were incubated with and without 1000 units of purified active cleaved caspase-3 (Enzo Life Sciences) for 5 h at 30 °C. Reactions were quenched by addi-tion of 20 µL 6× SDS sample buffer and boiling for 5 min. Proteins were analyzed on 8% SDS-PAGE gel and colloidal stained to visualize protein bands.

Annexin V/PI apoptosis assay

MCF-10A or NHDF cells were treated with various doses of UV. At the indicated time point, the media of both treated and untreated cells was collected and combined with trypsi-nized cells. Cells were then re-suspended in 1× annexin buffer (BD cat# 556454) at a concentration of 1000 cells/µL. 100 µL of the cell suspension was then added to a 96 well plate. A propidium iodide (BD cat# 556463) and Annexin V-FITC (BD cat# 556419) cocktail was prepared in the 1× annexin buffer. The cocktail contained 5 µL of propidium iodide and 5 µL of Annexin V and 90 µL of annexin V bind-ing buffer. 100 µL of the staining solution was then added to cells, followed by incubation for 15 min at RT. An additional 100 µL of annexin V binding buffer without annexin V or PI was added to each well to dilute samples for the flow cytometer. The samples were run on Guava Flow Cytometer

equipped with 96 well automated counter with green and red channel detectors to detect fluorescence from FITC-Annexin V and propidium iodide, respectively.

Western Blot

Primary Sp1 antibody was used as previously described [13]. Westerns for nucleolin (Santa Cruz Biotechnology cat# sc-55486), tubulin (Santa Cruz Biotechnology, cat# sc-8035) and GAPDH (Santa Cruz Biotechnology, cat# sc-365062) were performed as loading controls. GST antibody (Santa Cruz Biotechnology, cat# sc-459) was used to determine location of the cleavage fragment in GST-Sp1 purified from bacteria. Bcl-2 antibody (Calbiochem Cat#OP60) was used to determine levels of exogenous Bcl-2 in MCF-10A cells after transduction. The following antibodies were purchased from Cell Signaling Technology: HA (cat# 3724), full-length caspase 3 (cat# 9662), cleaved caspase 3 (cat# 9664), and cleaved caspase 6 (cat# 9761). γH2AX (cat# 05-636) and total H2AX (cat# 07-627) were purchased from Millipore. Secondary antibodies were purchased from Licor IRDye 680 Goat anti-Rabbit (cat# 926-32221) or IRDye 800CW Goat anti-Mouse (cat# 926-32210). Secondary antibodies were re-suspended in 5% BSA at 1:10,000.

Samples were resolved on denaturing polyacrylamide gels and transferred to PVDF membrane (Millipore cat# IPFL00010), which was blocked for 1 h in 5% BSA for all westerns except nucleolin and Sp1, which were blocked in 1% milk. The primary antibody was added and incubated overnight at 4 °C with rocking. The next day, the PVDF was washed 3 times for 10 min each in PBS containing 0.1% Tween, followed by incubation with secondary antibody for 2 h, three 10 min washes in PBS containing 0.1% Tween, and a 4th wash in PBS. The blots were exposed using Odys-sey Imager from Licor.

Results

Sp1 is cleaved in response to high levels of DNA damage

Treatment of MCF-10A cells with UV (10  mJ/cm2) resulted in a decrease in the amount of full-length Sp1 8 h after treatment, with no effect on the level of the loading control, nucleolin (Fig. 1a). By comparison, treatment with 1 mJ/cm2 UV had no effect on the level of full-length Sp1. The decrease in full-length Sp1 in response to 10 mJ/cm2 UV coincided with the appearance of a 70 kDa product. The antibody used to detect Sp1 recognizes an epitope at amino acids 524–543; the 70 kDa product was designated Sp1-70. To rule out the possibility that Sp1-70 was an anti-body artefact, we confirmed its appearance with multiple

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Sp1 antibodies (Fig. A1). To determine the time course of Sp1-70 appearance, cells were treated with 10 mJ/cm2 UV and analyzed every hour for up to 8 h after treatment. The 70 kDa cleavage fragment was detected as early as 5 h post treatment (Fig. 1b). To analyze the relationship between damage and Sp1-70, cells were treated with increasing doses of UV and analyzed at 8 h post treatment. Sp-70 was detectable with 2.5 mJ/cm2 UV treatment, and dose-dependently increased up to 20 mJ/cm2 UV, after which its appearance plateaued (Fig. 1c). Quantification of the full-length Sp1 and Sp1-70 bands revealed that the reduc-tion in full-length Sp1 inversely correlated with Sp1-70 (Fig. 1d), consistent with Sp1-70 arising from cleavage of full-length Sp1.

Cleavage of Sp1 is correlated with caspase activation

To determine whether cleavage of Sp1 was specific to the type of damaging stimulus, MCF-10A cells were treated with a panel of DNA damaging agents, including doxoru-bicin, bleomycin, or camptothecin. Cells treated with UV or doxorubicin were lysed 8 h after treatment; cells treated with camptothecin were lysed 24 h after treatment. Sp1-70 was detected after all treatments (Fig. 2a). Since the appear-ance of Sp1-70 is associated with moribund cells, lysates from cells treated with different DNA damaging agents were probed for the presence of cleaved caspase 3, a marker of apoptosis. We observed a direct correlation between

Fig. 1 Sp1 is cleaved in response to high levels of DNA damage. a MCF-10A cells were treated with 1 or 10 mJ/cm2 UV. At the indicated time points post-treatment, cells were lysed and immunoblotting for Sp1 was performed; blots were probed with nucleolin as a loading control. b MCF-10A cells were treated with 10 mJ/cm2 UV and lysed every hour for 8 h after treatment. c MCF-10A cells were treated with the indicated doses of UV and cells were lysed at 8 h post treatment. d Quantitation of the Western blot by Licor; the intensity of the Sp1 and Sp1-70 bands rela-tive to untreated was measured and normalized to nucleolin

0 1 4 8 0 1 4 8 hours 1 mJ/cm2 10 mJ/cm2

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Fig. 2 Sp1 cleavage correlates with caspase activation. a MCF-10A cells were treated with the indicated DNA damage agents and lysed at 8 h (doxorubicin and UV) or 24 h (bleomycin and camptothecin) after treatment or drug addition. b MCF-10A cells were treated with 100 or 200 ng/mL TRAIL for 8 h. a, b Lysates were prepared at the end

of treatment and immunoblots with antibodies against Sp1, cleaved caspase 3 or γ-tubulin were performed as indicated. c MDCK cells were treated with 10 or 100 mJ/cm2 UV. Eight hours after treatment, cells were lysed and immunoblotting was performed for Sp1 and γ-tubulin. Arrows to the right in each panel indicate Sp1-70

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the intensity of Sp1-70 and the level of cleaved caspase 3 (Fig. 2a). There are two established pathways for the induc-tion of apoptosis: the intrinsic pathway, generally activated by DNA damage, and the extrinsic pathway, mediated by receptor activation. Induction of either pathway converges on cleavage-mediated activation of executioner caspases, including caspase 3. To determine if Sp1 is also cleaved upon activation of the extrinsic pathway, MCF-10A cells were treated with 100 or 200 ng/mL TRAIL ligand, and ana-lyzed 8 h post-treatment. As shown in Fig. 2b, TRAIL ligand induced cleaved caspase 3 and Sp1 cleavage as indicated by the appearance of Sp1-70, demonstrating that activation of either the extrinsic or the intrinsic pathway can produce Sp1-70. Importantly, cleavage of Sp1 is not restricted to MCF-10A cells; Sp1-70 was observed in MDCK cells after UV treatment (Fig. 2c), as well as in U2OS cells after H2O2 treatment (data not shown). Together, these data suggest that the cleavage of Sp1 is linked to caspase activation and thus apoptosis. In contrast to multiple epithelial cell lines, Sp1-70 was not detectable in NHDF cells treated with up to 100 mJ/cm2 UV (Fig. A2a). Since the induction of Sp1-70 is correlated with apoptosis in other cell types, we sought to determine whether NHDF cells underwent apoptosis in response to UV treatment. Morphological changes observed in NHDF cells revealed enlarged and flattened cells, with no evidence of shrinking or nuclear blebbing. Quantifica-tion of apoptosis by Annexin V/propidium iodide labeling followed by flow cytometry, showed very few Annexin V/PI labeled NHDF cells compared to MCF-10A cells (Fig. A3b), suggesting that NHDF cells do not undergo classical apoptosis in response to UV treatment. This may account for the lack of Sp1-70 in these cells. Notably, UV-treated NHDF cells failed to proliferate and remained attached, albeit slightly enlarged, suggesting that they may be under-going senescence. To test this, we measured senescence by SA-β-galactosidase assay in UV-treated MCF-10A and NHDF cells. NHDF cells stained positive for β-galactosidase after treatment with 10 mJ/cm2 UV (Fig. A2c); in contrast, MCF10A were negative for β-galactosidase staining, sup-porting the premise that Sp1 cleavage and formation of Sp1-70 are linked specifically to apoptosis.

Blocking caspase activation prevents Sp1 cleavage

Bcl-2 is a potent inhibitor of apoptosis that inhibits caspase activity by preventing the release of cytochrome c from the mitochondria [35]. To further establish that the appearance of Sp1-70 is specific to apoptosis, Bcl-2 or RFP (red fluo-rescent protein, control vector) were overexpressed in MCF-10A cells. The level of Sp1-70 was significantly lower in UV-treated cells overexpressing Bcl-2 as compared to mock-treated or RFP control cells (Fig. 3a). Therefore, genera-tion of Sp1-70 is sensitive to modulation of the apoptotic

threshold. To more directly determine whether Sp1-70 is produced by caspase cleavage, we depleted cells of caspase 3 using siRNA, and treated cells with UV (10 mJ/cm2) or hydrogen peroxide (400 µM). Depletion of caspase 3 blocked the formation of Sp1-70 in response to hydrogen peroxide treatment, but not in response to UV treatment (Fig. 3b).

Since inhibition of caspase 3 can block the induction of Sp1-70 by H2O2 but not UV, Sp1 must also be cleaved by additional caspases activated by UV. Indeed, Sp1 is cleaved in MCF-7 cells, which are known to be caspase 3 deficient [36–39] (Fig. A3), although cleavage occurs at a much lower efficiency compared to MCF-10A cells. To evaluate the possibility that Sp1 cleavage can be mediated by multiple caspases, MCF-10A cells were treated with 50 µM of pan-caspase inhibitor Z-VAD FMK for 30 min prior to treatment with 10 mJ/cm2 UV or 100 ng/mL TRAIL ligand. Z-VAD FMK blocked the formation of Sp1-70 in response to UV or TRAIL treatment (Fig. 3c). Both Bcl-2 overexpression and Z-VAD FMK blocked the induction of Sp1-70, indicat-ing that global inhibition of caspase activation can prevent Sp1 cleavage in cells undergoing UV- or TRAIL-induced apoptosis. To analyze the difference between H2O2-induced cleavage and UV-induced cleavage, we monitored γ-H2AX levels to determine doses of UV or H2O2 that produce com-parable amounts of DNA damage. In MCF-10A cells treated with 10 mJ/cm2 UV or 400 µM hydrogen peroxide (which induce similar γ-H2AX signal), the level of activated cas-pases 3 and 6 was much higher in UV-treated cells (Fig. 3d). These data, as well as Fig. 3b implicate multiple caspases, including caspase 3, in the formation of Sp1-70. The varia-tion in the amount of Sp1-70 formed in H2O2-treated cells compared to UV-treated cells likely results from differences in the caspases activated by each treatment.

Sp1 is cleaved at aspartic acid 183 by caspase 3

After establishing that Sp1 is cleaved by caspases, we next sought to determine the precise location of the cleavage. A previous report indicated that Sp1 is cleaved by caspase 3 at aspartic acid 584 to produce a cleavage product of approximately 70 kDa [34]. At the time those studies were performed, misidentification of the initiator methionine had resulted in misnumbering of Sp1’s amino acids There-fore, aspartic acid 584 is actually aspartic acid 590. We used an Sp1 expression vector tagged with GST at the N-terminus and HA at the C-terminus to study the Sp1 cleavage products (illustrated in Fig. 4a). GST-Sp1-HA was purified from bac-teria and incubated with active caspase 3, which produced a 70 and 48 kDa product (Fig. 4b). Cleavage of Sp1 at D590 is predicted to produce a 100 kDa product (GST + 1-590 Sp1); no 100 kDa product was observed by immunoblotting with α-GST antibody. Rather, the α-GST antibody detected a 48 kDa fragment, which is the approximate length of a

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fusion protein containing GST and the first ~ 200 amino acids of Sp1. These data suggest that Sp1 is cleaved in the vicinity of amino acid residue 200 in vitro by caspase 3, sup-porting the presence of an additional cleavage site.

Analysis of the Sp1 amino acid sequence revealed two aspartic acid residues in the first 200 amino acids of Sp1, D183, which is highly conserved among species (Fig. A4), and the non-conserved D199. We mutated these aspartic acid residues to alanine and made 35S methionine labeled prod-ucts by coupled in vitro transcription/translation (described in methods) and incubated these products with active caspase 3. Mutation of D183 blocked cleavage of Sp1 by caspase 3, whereas mutation of D199 (not shown) or the previously established cleavage site D590, still gave rise to Sp1-70 (Fig. 4c). Therefore, aspartic acid 183 was required for caspase 3-mediated Sp1 cleavage. This was confirmed by expressing HA-tagged wild-type Sp1, D590A, D183/590A, or D183A in MCF10A cells and treating with 10 mJ/cm2 UV. After 24 h, α-HA antibody detected Sp1-70 in cells expressing wild-type Sp1 and Sp1-D590A, but failed to detect Sp1-70 in cells expressing D183A/D590A or D183A Sp1 (Fig. 4d). These results show that Sp1 is cleaved by caspases at aspartic acid 183 and that the 70 kDa product

includes the C-terminus. Given these data, Sp1-70 is here-after designated Sp1-70C.

Mutation of aspartic acid 183 renders cells resistant to UV‑induced apoptosis

We next sought to determine the functional significance of Sp1 cleavage. To address this, we depleted MCF-10A cells of endogenous Sp1 using shRNA 7276 (Sigma), which tar-gets the 3′ UTR, and then overexpressed Sp1 (wild-type and D183A) via retroviral transduction. Exogenous Sp1 lacks the 3′ UTR and is not targeted by this shRNA. Cells were selected with puromycin and G418 to derive cells depleted of endogenous Sp1 with uniform overexpresssion of exog-enous Sp1. During selection, cells expressing high levels of exogenous Sp1 underwent apoptosis. At the end of the puromycin selection period, exogenous Sp1 expression was equivalent among cells transduced with the different retro-viruses (Fig. A5). Cells were treated with 10 mJ/cm2 UV and analyzed for apoptosis by annexin V/propidium iodide labeling. As shown in Fig. 5, cells expressing Sp1 D183A or Sp1 D183/590A undergo significantly less apoptosis in response to UV, whereas cells expressing wild-type Sp1 or

50 mJ/cm2 25 mJ/m2

Mock Bcl-2 RFP

Sp1

Bcl-2

Nuc

Untreated

Mock Bcl-2 RFP Mock Bcl-2 RFP

a

d Sp1

C-casp 3

C-casp 6

γH2AX

H2AX

Ctrl H2O2 UV H2O2 UV

Sp1(long exp.)

4Hours 8 Hours

c

Z-VADTrailUV

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+

+ ++ +

+ +

b

tCasp 3

Ctrl

C3

10 mJ/cm2 UV 400 µM H2O2Untreated

Ctrl

C3

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C3

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Nuc

siRNA

Fig. 3 Blocking caspase activation prevents Sp1 cleavage. a MCF-10A cells were transduced with pBABE retrovirus expressing RFP (3) or Bcl-2 (2) or mock-treated (M). 24  h after transduction, cells were treated with indicated doses of UV and lysed after 8  h, fol-lowed by immunoblotting with the indicated antibody. b MCF-10A cells were transfected with non-targeting siRNA (Ctrl) or caspase 3 siRNA (C3). 72  h after transfection, cells were treated with 10  mJ/cm2 UV or 400 µM hydrogen peroxide. Eight hours after treatment,

immunoblotting was performed for Sp1 and full-length caspase 3. c 50 µM Z-VAD FMK was added to cells 30 min prior to treatment with 10 mJ/cm2 UV or 100 ng/mL TRAIL. Nucleolin was used as a loading control. d MCF-10A cells were treated with 400 µM hydro-gen peroxide or 10 mJ/cm2 UV and lysed after 4 and 8 h. Antibodies shown to the left of each panel were used for immunoblotting. Sp1-70 is indicated by arrow

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Sp1-D590A showed no significant difference from untreated controls. The difference in overall apoptosis suggests that caspase-mediated cleavage of Sp1 at D183 may be involved in the initiation of apoptosis.

Overexpression of full‑length Sp1 or Sp1‑70C induces apoptosis in RPE cells

Sp1 overexpression alone can induce apoptosis in many non-transformed cells [14]. To assess the function of Sp1 cleavage and Sp1-70C in apoptosis without the confound-ing effects of DNA damage, we overexpressed wild-type Sp1, the non-cleavable Sp1-D183A and Sp1-70C in reti-nal pigment epithelial cells (RPE). To get a high level of overexpression, the three different Sp1 cDNAs were subcloned into a pLENTI vector in which expression was driven by the CMV immediate early promoter. RPE cells in which Sp1 or Sp1-70C was overexpressed showed sig-nificant cell and nuclear shrinking and floating cells 96 and 120 hours post-transduction (Fig. A5A). In contrast, overexpression of the non-cleavable Sp1 D183A showed significantly less morphological evidence of apoptosis (Fig. A6a). Densitometry analysis of Western blot with cleaved caspase 3 revealed less cleaved caspase 3 at 96 and 120 hours post-transduction in cells with Sp1 D183A, compared to those transduced with wild-type Sp1 or Sp1-70C (Fig. A6b). RPE cells overexpressing HA tagged

a

d

100 kDa

70 kDa48 kDa

120 kDa

70 kDa

Caspase 3 - + - +70 kDa

100 kDa

cCaspase3- + - + - +

b

A B C DBDGST HA

D183 D590

Silver -GST -αα HA

Fig. 4 Sp1 is cleaved by caspase 3 at aspartic acid 183 in vitro and in cells. a Schematic of Sp1 showing aspartic acid cleavage sites (top). b GST-Sp1 (GST tag at N-terminus) was purified from BL-21 cells and incubated with active caspase 3. After SDS-PAGE, the gel was subjected to colloidal stain to detect protein (left panel); immunoblot with GST antibody was also performed on samples to detect N-termi-nal cleaved product (right panel). c U2OS cells were transfected with Sp1-HA (HA at C-Terminus) and treated with 10 mJ/cm2 UV. Immu-noblotting was performed with Sp1 (right panel) or HA (left panel).

c 35S labeled Sp1 made by in vitro coupled transcription-translation was untreated (−) or incubated with active caspase 3 (+) for 3 h; sam-ples were separated by SDS-PAGE and visualized by autoradiogra-phy. (d) MCF-10A cells were transduced with the indicated retrovirus expressing an N-terminal FLAG tag in lieu of GST (Sp1 wild-type and with mutation of the indicated residues). Cells were then exposed to 10 mJ/cm2 UV and immunoblotting was performed with an α-HA antibody to detect cleavage of exogenous Sp1

0

5

10

15

20

25

30

35

% C

ell D

eath

PI

Annexin VLate apoptosisEarly apoptosis

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D59

0A

D18

3/59

0A

D18

3A WT

D59

0A

D18

3/59

0A

D18

3A

Untreated UV (10 J/cm2)

Fig. 5 Non-cleavable Sp1 protects cells from apoptosis. MCF-10A cells were depleted of endogenous Sp1 and transduced with retro-viruses expressing Sp1 wild-type (WT) or with indicated aspartic acid mutations. Cells were untreated or exposed to 10  mJ/cm2 UV. Cell death was measured by staining with Annexin V and propidium iodide followed by flow cytometry. Early apoptotic cells stain with Annexin V and late apoptotic cells stain with both annexin V and PI

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(C-terminal) Sp1, Sp1-D183A or Sp1-70C for 120 h were subjected to Western blot to demonstrate the appearance of the Sp1-70C in RPE cells overexpressing WT Sp1 and not in the D183A mutant (Fig. 6a). Annexin V/PI stain-ing of these cells revealed the induction of apoptosis in cells overexpressing Sp1 or Sp1-70C, but not in those overexpressing the non-cleavable Sp1-D183A (Fig. 6b). Together, these data further support the conclusion that caspase-mediated cleavage of Sp1 at D183 may be involved in the initiation of apoptosis.

Caspase cleavage promotes accumulation of the Sp1 cleavage product

Sp1 is also regulated in part through its stability. To deter-mine whether there is differential stability in the mutants vs. full-length Sp1, we derived MCF-10A cells stably expressing exogenous wild-type Sp1 (pLXSN-Sp1-HA), Sp1-D183A (pLXSN-Sp1-D183A-HA), and the truncated form corresponding to the cleavage product Sp1-70C (pLXSN-Sp1-70C-HA). These cell lines were treated with the DNA damaging agent Adriamycin to induce apopto-sis, and analyzed by immunoblotting with α-HA antibody. The intensity of each exogenously-expressed protein was monitored over time by Licor quantification of the HA signal. At 12 and 24 h post-treatment, protein levels of the non-cleavable Sp1 D183A decreased compared to wild-type Sp1 and Sp1-70C. Full-length Sp1 and Sp1-70C showed comparable levels at both time points (Fig. 7).

Discussion

These studies demonstrate that induction of apoptosis by DNA damage, TRAIL, or overexpression of Sp1 induces caspase-mediated cleavage of Sp1 at aspartic acid 183, a site not previously reported to be cleaved by caspases. The 70 kDa cleavage product, Sp1184–785 (or Sp1-70C), previ-ously referred to as Sp1ΔA [40], retains the DNA-binding domain and has been shown to be transcriptionally active [33, 41]. DNA-damage-induced apoptosis was reduced in

HA

0

10

20

30

40

50

60

Vector control Sp1 Sp1-D183A Sp1-70C

%Ce

ll De

ath

Late Apoptosis

Early Apoptosis

a b

Fig. 6 Overexpression of Sp1 and Sp1-70C induces apoptosis in reti-nal pigment epithelial cells. ARPE-19 cells were transduced with len-tivirus containing CMV promoter-driven wild-type Sp1, Sp1 D183A, or Sp1-70C or vector control. a Cell lysates were obtained 120 h after transfection followed by immunoblotting with HA and α-tubulin anti-

bodies. b Cells were collected 120 h post-infection and stained with Annexin V/PI followed by flow cytometry. Early apoptotic cells stain with Annexin V and late apoptotic cells stain with both annexin V and PI

HA

Pro

tein

Lev

els,

Nor

mal

ized

to

-T

ubul

in C

ontr

ol

Full-length Sp1

Sp1-D183A

Sp1-70C

UT 12 hours 24 hours 48 hours

Fig. 7 Cleavage of Sp1 at Aspartic Acid 183 promotes accumula-tion of Sp1-70C. MCF-10A cells were transduced with wild-type Sp1 (pLXSN-Sp1-HA), Sp1 in which aspartic acid is mutated to ala-nine (pLXSN-Sp1-D183A-HA), or the C-terminal cleavage product (pLXSN-Sp1-70C). Cells were treated with 10  μM Adriamycin to induce DNA damage, and cell lysates were collected at 12, 24, and 48 h post-treatment. The signal intensity of the HA tag from each cell line was quantified using Image J and normalized to the α-tubulin loading control. Standard error bars indicate the results of triplicate measurements in 3 biological replicates

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cells expressing Sp1-D183A relative to wild-type Sp1 or Sp1-70C, indicating that caspase cleavage of Sp1 is involved in apoptosis. In the absence of damaging agents, overexpres-sion of Sp1 D183A in non-transformed cells produced less apoptosis than overexpression of wild-type Sp1 or Sp1-70C suggesting that the non-cleavable Sp1 is less pro-apoptotic than wild-type Sp1 or Sp1-70C. Taken together, our data imply that Sp1 plays a pivotal role in the maintenance of genomic stability through regulation of the apoptotic pro-gram (see Model Fig. 8). One mechanism that contributes to these observations is that caspase cleavage of Sp1 promotes the accumulation of a transcriptionally active form of Sp1 during apoptosis.

Caspases cleave at aspartic acid residues in a consensus recognition motif [42–44]. Global analysis of sites proteo-lytically cleaved during apoptosis revealed the frequency of specific amino acids surrounding cleaved aspartic acid resi-dues in 70 proteins [45]. Based on experiments performed in vitro and in vivo, Sp1 cleavage occurs at D183 in a QTVD sequence, which is not a previously recognized consensus motif [44].

In previous studies, we showed that depletion of Sp1 increased sensitivity to DNA damage [13]. Therefore, we had anticipated that depletion of Sp1 would increase sen-sitivity to apoptosis in response to DNA damage; however, we observed the opposite effect. In our earlier studies, we induced DNA damage at a low level and evaluated the effect of Sp1 depletion using colony survival assays. Here, we induced much higher levels of damage. These opposing roles of Sp1 suggest that Sp1 may function to signal DNA damage and/or recruit repair proteins during the initial DNA damage response, whereas at higher levels of DNA damage,

Sp1 may be involved in induction of apoptosis through its caspase-mediated cleavage.

D183 is a newly discovered caspase cleavage site that we have shown is actively involved in modulating levels of Sp1 and promoting apoptosis. The precise function of the sepa-rate domains that result from caspase cleavage at D183 is not known. The cleavage results in a transcriptionally active form of Sp1, Sp1184–785 or Sp1-70C. Overexpression of Sp1 leads to the induction of apoptosis in many non-transformed cell types [14–16], including retinal pigment epithelial cells. Studies involving overexpression of deletion mutants of Sp1 revealed that the DNA-binding domain of Sp1 is required and sufficient for the activation of cell death [14], consist-ent with a transcriptional mechanism. This may potentially involve transcriptional repression, as was suggested in an earlier gene expression array in Sp1 overexpressing cells [15]. Previous studies show that N-terminal deletion mutants of Sp1 comparable to Sp1-70C retain approximately 90% of the transcriptional activity of full-length Sp1 [46–48]. However, results varied by cell type and with altered loca-tions and number of Sp1 binding sites in target promoters. It is possible that removal of the N-terminal 183 amino acids alters the downstream target specificity in favor of a pro-apoptotic program. The gene expression program that results in apoptosis is clearly highly complex and variable, involving direct and indirect modulation of both pro- and anti-apoptotic targets.

Cleavage at D183 produces a product that is transcrip-tionally active, in contrast to cleavage at D590, a site previ-ously shown to be cleaved by caspase 3 and 7 during B-cell apoptosis [31]. Cleavage at D590 would separate the DNA-binding domain from the transcription activation domains, thereby producing a dominant negative transcription factor. Identification of D590 as a caspase cleavage site was based on sequencing the N-termini of the fragments and was not validated by mutagenesis [30]. We observed no cleavage at D590 in response to DNA damage, TRAIL, or overexpres-sion of Sp1. Conservation analysis revealed that the Sp1 sequence 180-QTVD-183 is conserved among frog, dog, mouse, rat, pig and human Sp1, whereas D590 is signifi-cantly more divergent and absent in several species (Fig. A4).

A variety of tumors overexpress Sp1, which is an indi-cator of poor prognosis [49–52]. This is likely due to the large number of Sp1-regulated genes that promote pro-liferation. However, tumors that overexpress Sp1 must also have mechanisms to avoid the induction of apopto-sis, which is a well-known characteristic of tumors that is responsible for a significant portion of chemotherapy resistance [53, 54]. Consistent with this premise, our data show that an increased apoptotic threshold (overexpressed Bcl2) was able to confer resistance to Sp1 overexpression and Sp1-70C. Tumors that overexpress Sp1 may also have

BCL-2 Z-VAD FMKCaspase Ac va on

Sp1 Cleavage

Apoptosis

Genomic Integrity

Sp1-D183A

Sp1 O/ETRAIL Excess DNA damage

Fig. 8 Model representing the major findings with Sp1 cleavage. A variety of pro-apoptotic stimuli induce cleavage of Sp1 at D183A. This cleavage is caspase- mediated and produces a transcription-ally active truncated form of Sp1. Expression of non-cleavable Sp1 (D183A) results in less  damage-induced apoptosis, implicating Sp1 cleavage in the induction of apoptosis. By promoting apoptosis through its cleavage, Sp1 contributes to the preservation of genomic integrity

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inactive pro-apoptotic gene targets; methylation of pro-apoptotic promoters is widely reported [55]. Sp1 overex-pression induces p53-dependent apoptosis in some cancer cells [16]; however, p53 is mutated in 50% of cancers and many of these mutations are in the p53 promoter [56]. Overexpression of Sp1 in tumors is also correlated with increased expression of the pro-survival factor survivin [57, 58]. Furthermore, apoptosis is dysregulated in many transformed cells due to mutations/deletions in key pro-apoptotic factors, or expression of proteins that disable their activity, such as papilloma virus E6 and E7 in HeLa cells. In non-transformed smooth muscle cells, expression and phosphorylation of Sp1 were shown to be required for apoptosis through Sp1-dependent up-regulation of Fas ligand [17, 18]. Whether these results are related to cleav-age of Sp1 at D183 is not yet clear, but they underscore the significant role of Sp1 in modulating apoptosis.

Several proteins involved in preserving genomic integrity appear to have roles in the DNA damage response as well as in apoptosis. Sp1 cleavage at D183 also produces an N-ter-minal product (Sp1-30N, Sp11–183). We have found that Sp1-30N is required and sufficient for localization of Sp1 to DNA double-strand- break sites; Sp1-30N is also sufficient for the recruitment of chromatin remodeling factors and to facilitate repair [12]. Sp1-30N contains several serine residues that are rapidly phosphorylated by ATM after DNA damage [5, 13, 59]. As such, caspase cleavage of Sp1 at double-strand-break sites would not affect the ability of Sp1 to promote repair, and would release the transcriptionally active Sp1-70C. Factors involved in regulating DNA repair and apoptosis must be tightly coupled to the DNA damage response. Many DNA damage response proteins, including BRCA-1, CHK1, MDC1, MLH1, FANCD2 and Rad51 are caspase substrates [60–72], suggesting that cleavage of damage response pro-teins may be critical for execution of apoptosis. Several of these factors, including BRCA-1, Rad51 and MLH1 appear to have a dual function in DNA repair and apoptosis [60, 68, 71], although the exact mechanisms are not fully understood. Mutation of the caspase cleavage site in BRCA-1 renders cells resistant to UV-mediated apoptosis [68]. Initially, it was thought that the degradation of BRCA-1 was required for apoptosis, but similar to our results with Sp1, cells that are deficient in BRCA-1 are resistant to apoptosis [64]. The impact of cleavage-resistant BRCA-1 or BRCA-1 depletion on apoptosis directly parallel the results observed with Sp1 [63]. ATM has a clear role in the DNA damage response upon treatment with various agents [73, 74]. Fibroblasts deficient in ATM have been shown to have enhanced sensi-tivity to DNA double-strand-breaks, and this phenotype was rescued when ATM was reintroduced into cells [75]. ATM is also critical for apoptosis, as ATM-deficient cells are more resistant to apoptosis [76–78]. These opposing functions suggest that ATM activity is critical for DNA repair and

cell survival, yet with higher levels of DNA damage, ATM deficiency leads to decreased apoptotic signaling.

Multiple other caspase substrates have a direct role in apoptosis. Mutation in the caspase cleavage site of the tran-scription factor Twist causes resistance to apoptosis [79]. Like Sp1, Twist has been shown to be overexpressed in a variety of tumors. Caspase cleavage of pRb also likely plays an active role in apoptosis [80]; cells expressing caspase cleavage-resistant pRb have reduced apoptosis in response to endotoxin. While mice expressing caspase cleavage-resistant pRb showed no increase in spontaneous tumor development, this mutation enhanced tumor development in mice with a p53 negative background. The observations presented here, together with data highlighting other proteins involved in both the DNA damage response and transcription, demon-strate the importance of caspase cleavage in the maintenance of a tumor-free organism.

Regulation of Sp1 protein levels is complex. Caspase cleavage of Sp1 results in a cleavage product, Sp1-70C, which accumulates to a higher level as compared to the caspase-resistant Sp1-D183A. Studies have shown that SUMOylation of Sp1 on lysine 16 facilitates Sp1 degradation through the recruitment of RNF4 [32], a SUMO-targeted ubiquitin ligase (STUbL). Cleavage of Sp1 at aspartic acid 183 may function to prevent RNF4-mediated degradation by physically separating the SUMO-modified N-terminus of the protein from the C-terminus. This would permit the C-ter-minal portion, which retains most of Sp1’s transcriptional activity [33], to function as an independent transcriptional regulator. There are other examples of proteins in which caspase cleavage results in a product with increased stabil-ity. REDD1 (regulated in development and DNA damage response), a TSC2 activator and mTORC1-dependent signal-ing pathway inhibitor, is cleaved by caspases in response to apoptotic stimuli. One of the products of caspase cleavage of REDD1 is stabilized, while the other two products undergo proteolytic degradation. The stabilized product is pro-apop-totic, whereas the full-length protein is anti-apoptotic [81]. Grim, an IAP antagonist in Drosophila, is ubiquitylated by Drosophila inhibitor of apoptosis 1, resulting in its rapid turnover; however, active caspases cleave Grim simultane-ously, separating the ubiquitylated lysine from the remainder of the protein to stabilize it, thereby enhancing apoptosis [82]. Like REDD1 and Grim, caspase cleavage of Sp1 may propagate death signaling through a pro-apoptotic amplifica-tion loop resulting in greater cell death.

Sp1 appears to play a dual role in the cellular response to DNA damage. Sp1 facilitates repair of DNA double-strand-breaks, which involves its ATM-dependent phosphorylation [12, 13]. As demonstrated here, at levels of DNA damage that induce apoptosis, Sp1 is cleaved by caspases, and the cleavage contributes to apoptosis. This effect might be medi-ated through retention of the N-terminal cleavage product

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at sites of DNA damage and release of the transcriptionally active Sp1-70C. It is tempting to speculate that Sp1-70C might modulate transcription of targets different from those of wild-type Sp1. One of the roles of caspase activation dur-ing apoptosis is to inactivate processes such as DNA repair or production of anti-apoptotic factors, in order to achieve the overall goal of the cell—the maintenance of genomic integrity. The cleavage of Sp1 and that of other factors dis-cussed above might go one step further and actually facili-tate apoptosis. Whether the cleavage of Sp1 at D183 modu-lates its transcriptional targets, or affects apoptosis without altering transcription, the caspase-mediated cleavage of Sp1 reported here is at a newly discovered site and is actively involved in the induction of apoptosis. Identification of Sp1 D183 mutations/deletions in tumors would certainly support the significance of our findings.

Funding This study was funded by College of Medicine, Drexel University.

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