XPB helicase regulates DNA incision by the Thermoplasma acidophilum endonuclease Bax1

DNA Repair 11 (2012) 286–293

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XPB helicase regulates DNA incision by the Thermoplasma acidophilum endonuclease Bax1 Heide M. Roth a , Johannes Römer a , Verena Grundler a , Bennett Van Houten b , Caroline Kisker a,∗∗ , Ingrid Tessmer a,∗ a b

Rudolf Virchow Center for Experimental Biomedicine, Würzburg, Germany Department of Pharmacology and Chemical Biology, University of Pittsburgh Cancer Institute, University of Pittsburgh, PA, USA

a r t i c l e

i n f o

Article history: Received 27 June 2011 Received in revised form 14 November 2011 Accepted 6 December 2011 Available online 9 January 2012 Keywords: Bax1 XPB Nucleotide excision repair Transcription Archaea

a b s t r a c t Bax1 has recently been identified as a novel binding partner for the archaeal helicase XPB. We previously characterized Bax1 from Thermoplasma acidophilum as a Mg2+ -dependent structure-specific endonuclease. Here we directly compare the endonuclease activity of Bax1 alone or in combination with XPB. Using several biochemical and biophysical approaches, we demonstrate regulation of Bax1 endonuclease activity by XPB. Interestingly, incision assays with Bax1 and XPB/Bax1 clearly demonstrate that Bax1 produces different incision patterns depending on the presence or absence of XPB. Using atomic force microscopy (AFM), we directly visualize and compare binding of Bax1 and XPB/Bax1 to different DNA substrates. Our AFM data support enhanced DNA binding affinity of Bax1 in the presence of XPB. Taken together, the DNA incision and binding results suggest that XPB is able to load and position Bax1 on the scissile DNA substrate, thus increasing the DNA substrate range of Bax1. © 2011 Elsevier B.V. All rights reserved.

1. Introduction Structure-specific nucleases play a crucial role in DNA processing. Repair nucleases must be tightly regulated to ensure that spurious incisions, which could affect genome stability, do not occur. This regulation is often mediated by protein–protein interactions [1–3]. The endonuclease Bax1 has been recently identified as an interaction partner of the nucleotide excision repair (NER)1 helicase XPB [4]. XPB is one of two highly conserved ATPases/helicases that are part of the TFIIH complex, which is an essential component of NER, as well as, transcription initiation. NER is responsible for the removal of a huge variety of bulky and structurally unrelated lesions such as UV-induced photoproducts and adducts caused by

∗ Corresponding author at: Rudolf-Virchow Center for Experimental Biomedicine, Structural Biology, Josef-Schneider-Strasse 2, 97080 Würzburg, Germany. Tel.: +49 0931 3180425. ∗∗ Corresponding author at: Rudolf-Virchow Center for Experimental Biomedicine, Structural Biology, Josef-Schneider-Strasse 2, 97080 Würzburg, Germany. Tel.: +49 0931 3180381. E-mail addresses: [email protected] (C. Kisker), [email protected] (I. Tessmer). 1 The abbreviations used are: NER, nucleotide excision repair; dsDNA, doublestranded DNA; ssDNA, single-stranded DNA; nt, nucleotides; bp, base pairs; AUC, analytical ultracentrifugation; SV, sedimentation velocity; AFM, atomic force microscopy. 1568-7864/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.dnarep.2011.12.002

polycyclic aromatic hydrocarbons [5]. Two subpathways, global genome repair (GGR) and transcription coupled repair (TCR), have been characterized. Non-functional NER proteins trigger several severe diseases such as xeroderma pigmentosum, Cockaynes Syndrome and trichothiodystrophy. Some of the phenotypes of these diseases cannot be explained by a deficient NER pathway alone, but implicate defective transcription of specific nuclear receptor dependent genes [6]. Recent evidence has suggested that not only TFIIH, but also other NER factors such as XPC, XPA, and the NER endonucleases XPF–ERCC1 and XPG are recruited to the transcription initiation complex [6,7]. The role of the NER nucleases in eukaryotic transcription initiation is, however, still controversial; functions in DNA demethylation, chromatin relaxation or the stabilization of protein interactions have been proposed [6]. The archaeal NER pathway or an involvement of the corresponding proteins in transcription initiation are still not well understood, although several archaic NER homologs have been investigated using in vitro activity assays [8–13] and in vivo studies [14,15]. The archaeal endonuclease Bax1 has been identified by its close association to XPB [4]. We recently characterized Bax1 from the euryarchaeon Thermoplasma acidophilum as a Mg2+ dependent, structure-specific endonuclease [16]. Subsequently, the XPB/Bax1 complex from the crenarchaeon Sulfolobus solfataricus was investigated [17]. Different incision properties and DNA substrate specificities, as well as requirements for divalent cations for this XPB/Bax1 complex were identified than those observed for

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Bax1 from T. acidophilum [16,17]. It has also been reported that binding of Bax1 to XPB stimulates XPB’s ATPase activity in Sulfolobus tokodaii [18]. However, the interaction between XPB and Bax1 and its consequences are not understood. Resolving the functions of Bax1 and their regulation by XPB in archaeal NER and/or transcription will help to provide deeper insight into the intricate controls in these systems. Here, we investigated effects of the T. acidophilum XPB–Bax1 interaction on Bax1 endonuclease function and activity. Employing a combination of biochemical activity assays and biophysical techniques such as atomic force microscopy (AFM), we were able to shed light onto the activity of Bax1 by itself and in complex with XPB. Our studies provide insights into a regulatory role of XPB for the endonuclease activity of Bax1 by loading and positioning the protein onto the scissile DNA substrate. 2. Materials and methods 2.1. Expression and purification of the XPB/Bax1 complex The XPB/Bax1 complex was co-purified by mixing E. coli BL21-CodonPlus® (DE3)-RIL cells (Stratagene) which separately expressed T. acidophilum Bax1 (46 kDa) and XPB (52 kDa) both via the expression vector pETM-11 providing an N-terminal His6 -tag. Cells were lysed in a buffer containing 50 mM Tris–HCl, 500 mM NaCl, 50 mM imidazole, pH 7.5 and the cleared lysate was applied to a 1 ml Ni-MAC® cartridge (Merck) as described [16]. Pooled fractions were diluted 1:10 with IEX-A buffer (50 mM Tris–HCl, 50 mM NaCl, pH 7.5) and subsequently subjected to cation exchange chromatography using a 1 ml HiTrap SP HP column (GE Healthcare). The protein complex was eluted by a two-step gradient with a final concentration of 1 M NaCl. Pooled fractions were further purified by size-exclusion chromatography using a HiLoadTM 16/60 SuperdexTM 200 column (GE Healthcare). Isocratic elution of the protein complexes was carried out in 20 mM Tris–HCl, 500 mM NaCl, pH 7.5. Fractions containing pure protein complex were identified by SDS–PAGE [19], pooled, concentrated and stored at −80 ◦ C. The single proteins Bax1 as well as XPB were purified as described [16]. The two XPB/Bax1 complex types (Supplemental Fig. S1) were analyzed by CD spectroscopy to confirm correct folding (Supplemental Fig. S2B). To demonstrate that the altered incision properties of the incision competent XPB/Bax1 complex do indeed stem from Bax1 activity, we also co-purified XPB with two different Bax1 mutants Y152A and D132A impaired in nuclease activity [16] following the same protocols as for the wildtype protein.

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with the nickase Nt.AlwI (New England Biolabs). The short, incised oligonucleotides were then separated from the DNA substrate by repeated heating to 65 ◦ C and using a 50 kDa cutoff centrifuge concentrator. Circular DNA substrates for AFM were derived from a modified pUC19 DNA plasmid (pUC19N; 2729 bp) containing two Nt.BstNBI nickase sites [20]. The plasmid was generously provided by S. H. Wilson’s laboratory (NIEHS). Relaxation of the DNA was achieved by incubation with Nt.BstNBI at 55 ◦ C for 2 h. For AFM images on XPB/Bax1 or Bax1 with supercoiled DNA, non-modified pUC19 plasmid DNA was employed. 2.3. Bio-Layer interferometry Bio-Layer interferometry was performed using an Octet Red System (Forté Bio) using streptavidin coated biosensors. The top DNA strand NDT (Supplemental Table 1) was 5 biotinylated and annealed to form either 3 overhang or double-stranded DNA (dsDNA) substrates. A 3 biotinylated version of the DNA substrate NDT was used to form 5 overhang substrates. DNA substrates and proteins were diluted in a buffer containing 20 mM MES, pH 6.5, 150 mM NaCl, 1 mM DTT, 1 mg/ml BSA and 10 mM CaCl2 . Streptavidin biosensors were coated with biotinylated DNA substrate by incubation for 10 min in a 100 nM DNA solution followed by a 2 min washing step in buffer. The biosensors were then immersed into protein solution for 10 min to allow association and subsequently transferred into buffer to detect protein dissociation. All steps were performed at 37 ◦ C and 1000 rpm in a 96-well plate containing 200 ␮l solution in each well. From changes in light interference over time upon binding and dissociation, the kinetic rate constants kon , koff , and the equilibrium dissociation constant KD were derived using a 1:1 binding model (ForteBio OctetRED Evaluation software 6.1). 2.4. Incision assays Incision assays were performed as described [16]. If not stated differently, proteins were applied at a concentration of 5 ␮M, the concentration of DNA was 20 nM. Incision reactions were performed at 45 ◦ C for 45 min in reaction buffer containing 20 mM MES pH 6.5, 150 mM NaCl, 1 mM DTT, 0.1 mg/ml BSA and 10 mM MgCl2 , unless otherwise stated. Gels were visualized using the PharosFXTM Imager System (BioRad) detecting the fluorescein emission at a wavelength of 521 nm. The software Quantity One (BioRad) was employed for quantification.

2.2. DNA substrates

2.5. Atomic force microscopy (AFM) imaging

For incision assays, oligonucleotides were purchased from Biomers, dissolved in 0.1× TE buffer pH 7.5, diluted to a concentration of 10 ␮M and annealed in the presence of 100 mM KCl. The top DNA strand carried a fluorescein for detection either at its 5 or 3 end, or as a fluorescein adducted thymine at position 26, as indicated (by an F) in the cartoons below the corresponding gels. Identical results were achieved with internally fluorescein labeled strands. Sequences of the top and bottom strands are shown in Supplemental Table 1. For AFM experiments, a 522 bp DNA fragment (uvrb gene from B. caldotenax) was amplified by PCR using forward (1) and reverse (2) primers: (1) 5 -TCCGGACGATTTTTTGATC-3 and (2) 5 GACCAATACATCGTATTTGCC-3 . To generate a 32 nucleotide (nt) 3 overhang, the PCR product was incised using the nickase Nt.BstNBI (New England Biolabs) according to the manufacturers’ protocol. A 30 nt 5 overhang was produced by cutting the PCR product

DNA substrates (see above) were pre-incubated at 65 ◦ C for 10 min and slowly cooled down to room temperature to remove any salt crystals formed during storage. Proteins (0.5–2 ␮M as indicated) and DNA (100 nM for linear DNA fragments, 20 nM for circular DNA) were incubated in AFM incubation buffer (20 mM Tris–HCl pH 7.5, 150 mM NaCl, 10 mM DTT, 10 mM CaCl2 ) for 30 min at 45 ◦ C. Linear DNA samples were diluted 75-fold, circular DNA samples 20-fold in AFM deposition buffer (25 mM HEPES pH 7.5, 25 mM Na-acetate, 10 mM Mg-acetate) after incubation, immediately deposited onto freshly cleaved mica, rinsed with deionized water and dried in a gentle stream of nitrogen. All images were collected on an MFP-3D-BIO atomic force microscope (Asylum Research) in oscillating mode using Olympus OMCL-AC240 silicon probes with spring constants of ∼2 N/m and resonance frequencies of ∼70 kHz. Images were captured at a scan size of 2 ␮m × 2 ␮m, a scan rate of 0.5 Hz and a resolution of 1024 × 1024 pixels.

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2.6. Statistical analysis of AFM images For analysis, AFM images were flattened to 1st order. Peak heights and AFM volumes were measured and calculated using Gwyddion (P. Klapetek and D. Neˇcas) and ImageSXM (S. Barret) software. AFM volumes can be translated into protein molecular weights by calibrating the instrument with proteins of known molecular weight to derive a standard linear relationship (Supplemental Fig. S3). For our set-up, we thus found V = 1.2 × (MW) − 5.9, where V is the AFM volume and MW is the molecular weight. Molecular weights were derived from the center positions of Gaussian fits to the distributions of measured volumes (±1 standard deviation) using the software Origin. To determine the approximate molecular weights of DNA-bound protein complexes, we subtracted the DNA volume that is covered by a protein complex from the total complex volume. The DNA volume was calculated using the same length as that covered by the bound protein complex, the measured DNA height, and a width of 2 nm. Positions of protein peaks on the DNA were measured using the NIH ImageJ software. The position distribution histograms are presented as occurrence probability Pi = ni /(Nbp,bins × ˙ni ) versus position, where i is the position on the DNA [21], ni is the number of complexes observed at position i, ˙ni = n is the total number of binding occurrences observed within the position range, and Nbp,bins is the number of DNA base pairs in each position bin. From maxima in the distribution of relative occupancies, specificities for particular DNA strand internal positions can be determined. Specificity S for different DNA fragment end types was calculated from the ratio of relative occupancies of DNA ends and DNA strand internal positions (X): S = N × X + 1 [21], where N is the number of binding sites on the DNA (here N = 522 bp for the blunt end DNA substrate and N = 490 bp for the 3 and 5 overhang DNA substrates). For analysis of protein coverage of circular plasmid DNA, protein–DNA complexes were visually inspected and counted using the AFM software (Asylum Research on Igor Pro). 3. Results We previously demonstrated that Bax1 acts as a structure specific endonuclease and does not require any additional protein to incise DNA [16]. Bax1 can, however, interact with the ATPase/helicase XPB. It has been reported that S. solfataricus XPB and Bax1 associate to functionally combine their helicase and nuclease activities [17]. Moreover, Bax1 has been shown to stimulate S. tokodaii XPB’s ATPase activity [18]. In the present study, we examined the effects of T. acidophilum XPB binding with regard to the DNA binding and incision properties of T. acidophilum Bax1. 3.1. XPB/Bax1 exists in two different complexes During large scale co-purification of XPB/Bax1 complexes we observed both an incision-competent, as well as an incision-incompetent XPB/Bax1 complex from T. acidophilum (Supplemental Fig. S1). Further biochemical characterization of these two complexes was performed to assess their biological relevance. The formation of the incision-competent XPB/Bax1 complex or the incision-incompetent XPB/Bax1 complex can be controlled by solution conditions (see Supplemental Information 1 and Supplemental Fig. S1). The two XPB/Bax1 complex forms are both folded correctly and are indistinguishable in their stoichiometry (Supplemental Fig. S2) and their DNA binding properties. Both species bound to DNA as a heterodimer with preference for ssDNA overhang structures and similar dissociation constants in the nM-range (Supplemental Fig. S4), excluding a potential decrease in DNA affinity as possible cause for incision incompetence. Protease

digestion patterns indicated small differences in local conformation for the two XPB/Bax1 complexes (Supplemental Fig. S2F), consistent with their slightly different elution and sedimentation properties. Further experiments were performed to characterize the incision competent complex. 3.2. XPB loads Bax1 onto DNA nonspecifically and as a heterodimeric complex The isolated XPB and Bax1 proteins both bind DNA independently. To investigate effects of the XPB/Bax1 interaction on Bax1 DNA binding properties, we directly visualized and compared complexes of Bax1 alone or in complex with XPB with different linear DNA substrates using AFM imaging (Fig. 1). Bax1 bound to these DNA substrates as a monomer (Fig. 1B and D), consistent with previous results which showed that the monomeric form of the protein is functionally most active [16]. We calculated a molecular weight of approximately 60 ± 20 kDa from the AFM volumes of DNA-bound Bax1 complexes (after subtraction of the covered DNA volume, see Section 2, and Supplemental Fig. S3 for AFM volume to molecular weight calibration curve), which is in good agreement with the molecular weight of 46 kDa based on the protein sequence. The XPB/Bax1 complexes bound to DNA as a heterodimer: from the AFM volumes of DNA-bound complexes we calculated an approximate molecular weight of 120 ± 40 kDa (after subtraction of the estimated covered DNA volume) (Fig. 1C and D). Under the applied incubation conditions (1 ␮M protein, 100 nM DNA), we observed non-saturating low coverage of the DNA fragments with XPB/Bax1 complexes (0.26 ± 0.15/DNA fragment). These experiments were carried out in the absence of ATP to prevent potential helicase activity of XPB and to support observation of the complexes in the original DNA-binding state. The position distributions of protein complexes on non-damaged DNA substrates revealed no preferred DNA strand-internal target sites for XPB/Bax1 or Bax1 (Fig. 1E), indicating no specificity for DNA sequence or base pair composition. The fragment ends of these linear DNA substrates contained single stranded regions, which provide a model target for both Bax1 and XPB [10,16,17,22] (see Section 2). Interestingly, however, more than 50% of XPB/Bax1 complexes were found to bind to DNA strand-internal sites (more than 40% for Bax1 at 2 ␮M protein concentration). Considering the vast excess of non-specific strandinternal binding sites over binding sites at DNA fragment ends, these data were consistent with an approximately 200-fold preference for DNA fragment ends (approximately 300-fold for Bax1; Fig. 1E and Section 2). We investigated binding of Bax1 and of the XPB/Bax1 complex to different DNA end structures in more detail using Bio-Layer interferometry (Fig. 2). From these data, we calculated equilibrium dissociation constants, KD . Bax1 bound to these DNA substrates with KD ’s in the low ␮M-range, independent of the nature of the DNA substrate. For the XPB/Bax1 heterodimer, however, we observed a binding preference of about 10-fold for substrates containing ssDNA-overhangs, as compared to dsDNA (7 ± 2 nM for ssDNA overhang versus 61 ± 13 nM for blunt end DNA; Fig. 2). Importantly, our results reveal that XPB/Bax1 binds to DNA with about two orders of magnitude higher affinity than Bax1 alone and with KD ’s similar to those of XPB alone (Fig. 2). We wanted to further understand the binding of Bax1 and XPB/Bax1 complexes to physiologically relevant DNA. Since the genome of T. acidophilum consists of one circular chromosome, which is negatively supercoiled, we compared binding of Bax1 and XPB/Bax1 complexes to circular relaxed (B-form DNA containing no ends) as well as supercoiled DNA using AFM imaging. Fig. 3A–C shows enhanced protein complex formation on relaxed circular DNA for the XPB/Bax1 complex compared to Bax1 in the

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Fig. 1. XPB/Bax1 binds to DNA non-specifically and as a heterodimer. AFM images of (A) a DNA duplex (522 bp) comprising a 3 ssDNA-overhang (32 nucleotides) in the absence of protein and in the presence of (B) Bax1 or (C) XPB/Bax1. In spite of the higher protein concentration used in the incubations with Bax1 (2 ␮M versus 1 ␮M for XPB/Bax1), the protein surface coverage in images with Bax1 alone is significantly lower than with XPB/Bax1, due to different AFM deposition behavior of the protein complexes. The scale bar (in (C)) is 200 nm, the surface area of (A), (B), and (C) is 1 ␮m × 1 ␮m. (D) Gaussian fits to the data (R2 ≥ 0.96, n = 61 for Bax1 and n = 115 for XPB/Bax1) provide AFM volumes for the bound proteins of 87 ± 32 nm3 and 167 ± 55 nm3 for Bax1 (shown in black) and XPB/Bax1 (shown in gray), respectively. After subtraction of the covered DNA volume (approximately 20 ± 5 nm3 and 30 ± 10 nm3 for Bax1 and XPB/Bax1, respectively), these volumes translate into molecular weights of 61 ± 23 kDa (consistent with the theoretical MW of a Bax1 monomer of 46 kDa) and of 119 ± 39 kDa (consistent with a heterodimeric complex of Bax1 and XPB), respectively. (E) The position distributions of Bax1 (black) and XPB/Bax1 (gray) complexes on DNA (n = 36 and 96, respectively) show preferential occupation of DNA fragment ends and no significant specificity for any DNA strand internal sequence or base pair composition.

absence of XPB (Fig. 3E, 5.1 ± 0.5 versus 1.9 ± 1.1 per plasmid DNA, respectively, for a 25–50 fold protein excess, N = 510–978). Furthermore, Fig. 3D confirms comparable XPB/Bax1 binding to the negatively supercoiled plasmid DNA. Taken together, these

Fig. 2. DNA binding affinities of Bax1, XPB and the XPB/Bax1 complexes towards different DNA substrates were determined using Bio-Layer interferometry. Representative data for 3 different concentrations for each protein from a single experiment are summarized in the figure. Only binding curves obtained in this experiment that could be accurately described by the fitting algorithm (R2 ≥ 0.99) were included in the diagram. All results could be repeatedly reproduced qualitatively. Bax1 binds dsDNA and ssDNA overhang substrates with affinities in the low ␮M-range. In contrast, XPB as well as XPB/Bax1 complexes prefer ssDNA overhang substrates over dsDNA with approximately one order of magnitude and, importantly, bind all DNA substrates with approximately two orders of magnitude better than Bax1.

AFM and Bio-Layer interferometry data consistently indicate that XPB facilitates binding of Bax1 to DNA. 3.3. XPB/Bax1 and Bax1 exert different incision activities We previously reported a complex from T. acidophilum between XPB and Bax1, which led to complete suppression of DNA incisions by Bax1 [16]. This finding was in contrast to results with the XPB/Bax1 complex from S. solfataricus proteins that showed DNA incision with a variety of different substrates [17]. Interestingly, in subsequent purifications, as presented in the supplement (see Supplemental Information 1 and Supplemental Fig. S1), we have identified a specific fraction eluting from the sizing column that contains an incision-competent XPB/Bax1 complex from T. acidophilum. To study the effect of XPB binding on Bax1 activity, we directly compared DNA incision by Bax1 and by this XPB/Bax1 complex, both from T. acidophilum. Incision assays showed that Bax1 cuts 3 overhang substrates leading to one distinct incision product, while XPB/Bax1 produced a pattern of multiple, different incision products (Fig. 4A). The different incision behavior of Bax1 and XPB/Bax1 was even more pronounced for 5 overhang DNA substrates: Bax1 by itself did not incise any of the 5 overhang substrates, whereas the XPB/Bax1 complex produced clearly visible incision products with these substrates (Fig. 4A and B). The occurrence of multiple incision products with the single stranded DNA overhangs for XPB/Bax1, but not for Bax1 alone (gray arrows in Fig. 4A), correlates with non-specific incision activity, which we observed for XPB/Bax1 (but not for Bax1 by itself) on purely single

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To further confirm that the new incision activities are intrinsic to the XPB/Bax1 complex, two different incision-compromised Bax1 variants, D132A and Y152A [16], were co-purified together with XPB to form a stable complex. Incision assays revealed reduced incision activities for the XPB/Bax1 complexes formed with these mutants. Furthermore, incubation of DNA substrate with identical concentrations of XPB in the absence of Bax1 showed no DNA incision (data not shown). Correlation of activities in Bax1 mutants and XPB/Bax1 mutant complexes supports Bax1 as the source of endonuclease activity in the complexes (Supplemental Fig. S6).

4. Discussion

Fig. 3. AFM imaging of Bax1 and XPB/Bax1 binding to circular DNA. AFM images of nicked, relaxed pUC19N DNA (A) in the absence of protein, (B) in the presence of 0.5 ␮M Bax1, and (C) in the presence of 0.5 ␮M XPB/Bax1 and of (D) pUC19 supercoiled DNA in the presence of 1 ␮M XPB/Bax1. DNA binding may be supported by protein–protein interactions on the DNA (gray arrow in (D)). Similar complex formation with relaxed and supercoiled DNA demonstrates the suitability of negatively supercoiled DNA as a target for the XPB/Bax1 system. AFM images of supercoiled pUC19 in the absence of protein and in the presence of Bax1 are presented in Supplemental Fig. S8. Images are 1 ␮m × 1 ␮m, scale bars 200 nm. (E) Quantitative analysis of the number of protein complexes bound to relaxed pUC19N DNA (n = 218 and 389 for Bax1 and XPB/Bax1, respectively) shows increased binding of XPB/Bax1 to DNA compared to Bax1 alone, for protein concentrations of 0.5–1 ␮M.

stranded DNA substrates (Fig. 4C). These data point towards a role of XPB in positioning Bax1 on the DNA to increase its substrate range. Intriguingly, while Bax1 required Mg2+ -ions for catalysis [16], we observed that the XPB/Bax1 complex efficiently incised ssDNAoverhang substrates in the presence of either 10 mM MgCl2 , CaCl2 , MnCl2 or EDTA (Fig. 4D). Results were comparable for DNA substrates with 3 and 5 ssDNA overhangs (data not shown). We speculate that a pre-bound Mg2+ ion in the incision-competent XPB/Bax1 complex may enable Bax1 incisions under these ionic conditions. Since XPB could affect the incision reaction through its helicase activity, incision assays were carried out both in the presence and absence of ATP (as well as ADP and the non-hydrolysable ATP analog AMPPNP). Results were identical under these different conditions (Supplemental Fig. S5), arguing against a requirement for the XPB helicase activity in aligning Bax1 incisions.

In the present study we have determined the effects of the XPB–Bax1 interaction on Bax1’s endonuclease function and activity using a combination of biochemical approaches and atomic force microscopy imaging. Other laboratories have previously focused on the helicase function or ATPase activity of XPB in the complex [17,18]. In contrast, our focus was on the nuclease activity of Bax1. Nucleases play essential roles in DNA repair and have to be tightly regulated to prevent spurious and potentially lethal DNA incisions. Bax1 is currently the only archaeal nuclease known that interacts with protein homologs of the eukaryotic nucleotide excision repair enzymes. We were able to express and purify both XPB and Bax1 from the same, euryarchaeal organism, T. acidophilum. This allowed us to directly compare DNA binding and incision properties of Bax1 alone and in complex with XPB to decipher the underlying nuclease control mechanism. We observed enhanced DNA binding as well as an increased substrate range for the XPB/Bax1 complex compared to Bax1. Interestingly, incision activity could partially be restored for two incision-compromised Bax1 mutants if in complex with XPB. We observed an approximately 5-fold increase in incision activity for these mutants in the presence of XPB: ca 30% and 50% incision for XPB/Bax1 D132A and XPB/Bax1 Y152A, respectively, compared to ca. 5% and 10% for Bax1 D132A and Y152A in the absence of XPB (Supplemental Fig. S6C and D; and [16]). This “rescue-effect” may be related to the mechanism that leads to restoration of Bax1 incisions on 5 overhang DNA substrates (which are not cut by Bax1 alone) in the XPB/Bax1 complex (Fig. 4B). The incision properties on ssDNA substrates support a mechanism of non-specific ssDNA incisions for the XPB/Bax1 complex. Lack of specificity of Bax1 within the XPB/Bax1 complex raises the question, if additional proteins may be required to regulate correct DNA incisions by controlled positioning and orientation of Bax1 on DNA. Notably, discrimination between damaged and non-damaged DNA by the archaeal NER helicase XPD has been reported [9]. Studies on eukaryotic cells suggest a concerted activity of both ATPases/helicases XPB and XPD to unwind DNA and to verify the presence of an NER substrate [23]. Further experiments will be required to shed light on potential required protein–protein interactions in archaeal NER. The effects of XPB/Bax1 complex formation on the activity of XPB from two different crenarchaeal Sulfolobus species have been investigated previously [17,18]. The euryarchaeon T. acidophilum (our data) and the crenarchaea S. solfataricus and S. tokodaii [17,18] have evolved in two separate groups within the archaeal kingdom [26]. The point of separation in evolution is still under discussion [26]. Some archaea also contain the prokaryotic UvrABC proteins in addition to eukaryotic NER homologs [27,28]. It remains to be elucidated whether in archaeal species with both prokaryotic and eukaryotic NER proteins these systems work in concert or have evolved to serve different roles. Interestingly, the presence and number of different NER homologs seems to be almost randomly distributed among different archaeal species [27,28]. Furthermore, XPB

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Fig. 4. XPB/Bax1 and Bax1 exert different incision activities. (A) Incision assays conducted with XPB/Bax1 and Bax1 (abbreviated by X/B and B, respectively) showed a different pattern of incision products for 3 overhang substrates NDT/NDB26 and NDT/NDB30 (sequences are listed in Supplemental Table 1). In contrast, 5 overhang substrates were only cut by XPB/Bax1. Furthermore, in support of modified Bax1 nuclease activity in the complexes, XPB could only partially rescue Bax1 incision activity of nuclease deficient mutants [16] (Supplemental Fig. S6). Sites of incision on the corresponding DNA substrate are schematically represented by gray and white arrows for XPB/Bax1 and Bax1, respectively; the incision efficiency is indicated by the length of the arrows. Both Bax1 and XPB/Bax1 were used at a final concentration of 5 ␮M under identical reaction conditions. DNA ladders were generated to serve as a reference. (B) The graph shows the quantification of incision products (sum of gray arrows for XPB/Bax1 in gray, product at white arrow for Bax1 in white) deduced from 7 and 3 independently performed experiments for 3 and 5 overhang substrates, respectively (5 ␮M Bax1 or XPB/Bax1, 20 nM DNA substrate NDT/NDB30 and NDT/NDBr30). (C) Incision assays with Bax1 and XPB/Bax1 employing a 50 nucleotide single-stranded DNA substrate (ssDNA) as well as a 3 ssDNA overhang substrate for comparison show incisions only for the 3 overhang substrate for Bax1. In contrast, the XPB/Bax1 complex is additionally able to incise the ssDNA substrate non-specifically. (D) Incision assays with various divalent cations or the chelating agent EDTA demonstrate that Bax1 incises DNA only in the presence of MgCl2 , whereas XPB/Bax1 incises DNA in the presence of all tested cations or EDTA. Bax1 as well as XPB/Bax1 were employed at a final concentration of 5 ␮M, the concentration of the 3 overhang DNA substrate NDT/NDB22 was 20 nM. Arrows indicate the different incision products for Bax1 and XPB/Bax1.

proteins from sulfolobus and thermoplasma species share in total only 39% sequence identity. In comparison, the helicase core domains of T. acidophilum and S. sulfolobus share 26% and 27% sequence identity to human XPB, respectively [22,29]. T. acidophilum and S. solfataricus Bax1 are only 22% identical in sequence. The diversity within the archaeal kingdom must be considered when comparing the activity of archaeal proteins from different species. Our results demonstrate a role for XPB in loading and correctly positioning Bax1 onto scissile DNA substrates to achieve specific DNA incisions. Surprisingly, the helicase activity of XPB was not involved in this positioning process although S. solfataricus XPB and Bax1 were reported to work together as a helicase-nuclease machinery (Supplemental Fig. S5) [17]. The crenarchaeal and the euryarchaeal XPB/Bax1 appear to exhibit different requirements for divalent cations for DNA incision [17]. In addition, site-directed mutagenesis suggested that different active sites are employed by complexes from the different species to incise DNA [16,17]. Interestingly, we also observe subtle differences in DNA substrate specificities between proteins from S. solfataricus and T. acidophilum, which can be partially explained by the different DNA superstructure in these organisms. In particular, in contrast to our data for T. acidophilum, the crenarchaeal protein complex can open and incise DNA bubble substrates (see Supplemental Fig. S5 and [17]). In all three domains of life, eukaryotes, prokaryotes and archaea, mechanisms have evolved to pack and compact DNA [30]. Hyperthermophilic archaea, such as S. solfataricus, contain a reverse gyrase, which

positively supercoils DNA to protect DNA from denaturation at high temperatures [31–34]. The genome of T. acidophilum, however, comprises one circular negatively supercoiled chromosome [32,35]. Our AFM data clearly demonstrate binding of XPB/Bax1 to negatively supercoiled DNA (Fig. 3D), mimicking physiological conditions in T. acidophilum. Negative supercoiling facilitates melting or bubble formation in the DNA, which is likely unnecessary at the high temperatures of hyperthermophiles [31]. Facilitated bubble formation may also explain the lack of ATP requirement for XPB/Bax1 incision activity for T. acidophilum. We hypothesize that DNA binding proteins may have adapted to the superstructure of the DNA present in the organism. T. acidophilum XPB might not require helicase activity to bind and to position Bax1 onto negatively supercoiled DNA. In contrast, S. solfataricus XPB/Bax1 may need to exploit its helicase activity to locally melt and unwind positively supercoiled DNA, creating an access point for Bax1 to subsequently perform the incision reaction. Further structural and functional analyses will be required to elucidate different superstructural specializations in the different archaeal species. Finally, our experiments also revealed two different types of XPB/Bax1 complexes, only one of which is competent in DNA incisions (Supplemental Fig. S1). Importantly, we did not observe interconversion of the two complex forms into each other (Supplemental Fig. S7), indicating that the incision-incompetent form may also represent a dead-end complex. Alternatively, an as yet unknown trigger protein and/or protein–DNA complex may initiate the interconversion of a nuclease inactive to an active complex.

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The presence of an incision-incompetent XPB/Bax1 complex in addition to the incision-competent form may point towards a regulation mechanism for Bax1 endonuclease activity. This notion is also consistent with the presence of two different XPBs (XPBI and XPBII) in S. solfatarius and S. tokodaii (Sto), only one of which is able to interact with Bax1 [4,18] and promote DNA incisions by Bax1 [17]. It has been shown, that interaction of StoBax1 with StoXPBII enhances XPB’s ATPase activity [18]. In eukaryotes, in the context of NER, an increase in ATPase activity of XPB is induced by its interaction with the protein p52 and is believed to lead to a conformational switch and ultimately a widening of the DNA bubble that is initially formed at an NER target site by the protein XPC [24]. Further proteins that are required for eukaryotic NER can then load onto the emerging DNA bubble. Because StoXPBI, which does not bind to StoBax1, shows high affinity to ssDNA, Ma and colleagues speculated that both XPBI and XPBII/Bax1 could function in dsDNA destabilization and thus bubble enlargement while XPBII/Bax1 is responsible for 3 incision of the DNA at a later step of the NER pathway [18]. The similar DNA binding properties of the two XPB/Bax1 complexes from T. acidophilum are consistent with this notion. Alternatively, the incision-incompetent XPB/Bax1 complex from T. acidophilum may serve to competitively suppress DNA incision under particular intracellular conditions or modulate interactions with further proteins. In support of this idea, roles in stabilization and modification of protein interactions by the eukaryotic NER endonuclease XPG have previously been reported [25]. 5. Conclusion In summary, we found that the interaction of XPB and the structure specific archaeal endonuclease Bax1 enhances binding to DNA and increases Bax1’s substrate range. XPB hence serves to load and orient Bax1 on the DNA. We discuss different approaches between species and the co-existence of two XPB/Bax1 complexes in T. acidophilum. While one complex supported DNA incisions by Bax1 on a broader substrate range than for Bax1 in the absence of XPB, the other complex completely suppressed incisions by Bax1. Nuclease inhibition may thus be a further aspect of the regulatory effect of XPB on Bax1activity. Conflicts of interest There are no conflicts of interest. Acknowledgments We would like to thank Hong Wang and Hermann Schindelin for critical reading of the manuscript, Michael Fried for helpful discussions, and Samual H. Wilson for providing pUC19N plasmid DNA. This work was supported by grants from the Deutsche Forschungsgemeinschaft (DFG; KI–562/2 Forschungszentrum FZ82) to CK and IT and NIH grant ES019566 (BVH). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.dnarep.2011.12.002. References [1] M. Skorvaga, K. Theis, B.S. Mandavilli, C. Kisker, B. Van Houten, The beta-hairpin motif of UvrB is essential for DNA binding, damage processing, and UvrCmediated incisions, The Journal of Biological Chemistry 277 (2002) 1553–1559. [2] Y. Zou, R. Walker, H. Bassett, N.E. Geacintov, B. Van Houten, Formation of DNA repair intermediates and incision by the ATP-dependent UvrB–UvrC endonuclease, The Journal of Biological Chemistry 272 (1997) 4820–4827.

[3] V. Mocquet, J.P. Laine, T. Riedl, Z. Yajin, M.Y. Lee, J.M. Egly, Sequential recruitment of the repair factors during NER: the role of XPG in initiating the resynthesis step, The EMBO Journal 27 (2008) 155–167. [4] J.D. Richards, L. Cubeddu, J. Roberts, H. Liu, M.F. White, The archaeal XPB protein is a ssDNA-dependent ATPase with a novel partner, Journal of Molecular Biology 376 (2008) 634–644. [5] A. Sancar, DNA excision repair, Annual Review of Biochemistry 65 (1996) 43–81. [6] N. Le May, J.M. Egly, F. Coin, True lies: the double life of the nucleotide excision repair factors in transcription and DNA repair, Journal of Nucleic Acids 2010 (2010). [7] N. Le May, D. Mota-Fernandez, R. Vélez-Cruz, I. Iltis, D. Biard, J.M. Egly, NER factors are recruited to active promoters and facilitate chromatin modification for transcription in the absence of exogenous genotoxic attack, Molecular Cell 38 (2010) 54–66. [8] S.C. Wolski, J. Kuper, P. Hanzelmann, J.J. Truglio, D.L. Croteau, B. Van Houten, C. Kisker, Crystal structure of the FeS cluster-containing nucleotide excision repair helicase XPD, PLoS Biology 6 (2008) e149. [9] N. Mathieu, N. Kaczmarek, H. Naegeli, Strand- and site-specific DNA lesion demarcation by the xeroderma pigmentosum group D helicase, Proceedings of the National Academy of Sciences of the United States of America 107 (2010) 17545–17550. [10] L. Fan, A.S. Arvai, P.K. Cooper, S. Iwai, F. Hanaoka, J.A. Tainer, Conserved XPB core structure and motifs for DNA unwinding: implications for pathway selection of transcription or excision repair, Molecular Cell 22 (2006) 27–37. [11] J. Rudolf, C. Rouillon, U. Schwarz-Linek, M.F. White, The helicase XPD unwinds bubble structures and is not stalled by DNA lesions removed by the nucleotide excision repair pathway, Nucleic Acids Research 38 (2010) 931–941. [12] L. Fan, J.O. Fuss, Q.J. Cheng, A.S. Arvai, M. Hammel, V.A. Roberts, P.K. Cooper, J.A. Tainer, XPD helicase structures and activities: insights into the cancer and aging phenotypes from XPD mutations, Cell 133 (2008) 789–800. [13] H. Liu, J. Rudolf, K.A. Johnson, S.A. McMahon, M. Oke, L. Carter, A.M. McRobbie, S.E. Brown, J.H. Naismith, M.F. White, Structure of the DNA repair helicase XPD, Cell 133 (2008) 801–812. [14] D.J. Crowley, I. Boubriak, B.R. Berquist, M. Clark, E. Richard, L. Sullivan, S. DasSarma, S. McCready, The uvrA, uvrB and uvrC genes are required for repair of ultraviolet light induced DNA photoproducts in Halobacterium sp. NRC-1, Saline Systems 2 (2006) 11. [15] S. Frols, M.F. White, C. Schleper, Reactions to UV damage in the model archaeon Sulfolobus solfataricus, Biochemical Society Transactions 37 (2009) 36–41. [16] H.M. Roth, I. Tessmer, B. Van Houten, C. Kisker, Bax1 is a novel endonuclease: implications for archaeal nucleotide excision repair, The Journal of Biological Chemistry 284 (2009) 32272–32278. [17] C. Rouillon, M.F. White, The XBP–Bax1 helicase-nuclease complex unwinds and cleaves DNA: implications for eukaryal and archaeal nucleotide excision repair, The Journal of Biological Chemistry 285 (2010) 11013–11022. [18] X. Ma, Y. Hong, W. Han, D. Sheng, J. Ni, G. Hou, Y. Shen, Single-stranded DNA binding activity of XPBI, but not XPBII, from Sulfolobus tokodaii causes doublestranded DNA melting, Extremophiles 15 (2011) 67–76. [19] U.K. Laemmli, Cleavage of structural proteins during the assembly of the head of bacteriophage T4, Nature 227 (1970) 680–685. [20] E.W. Hou, R. Prasad, K. Asagoshi, A. Masaoka, S.H. Wilson, Comparative assessment of plasmid and oligonucleotide DNA substrates in measurement of in vitro base excision repair activity, Nucleic Acids Research 35 (2007) e112. [21] Y. Yang, L.E. Sass, C. Du, P. Hsieh, D.A. Erie, Determination of protein–DNA binding constants and specificities from statistical analyses of single molecules: MutS-DNA interactions, Nucleic acids Research 33 (2005) 4322–4334. [22] T. Biswas, J.M. Pero, C.G. Joseph, O.V. Tsodikov, DNA-dependent ATPase activity of bacterial XPB helicases, Biochemistry 48 (2009) 2839–2848. [23] F. Coin, V. Oksenych, J.M. Egly, Distinct roles for the XPB/p52 and XPD/p44 subcomplexes of TFIIH in damaged DNA opening during nucleotide excision repair, Molecular Cell 26 (2007) 245–256. [24] L.C. Gillet, O.D. Scharer, Molecular mechanisms of mammalian global genome nucleotide excision repair, Chemical Reviews 106 (2006) 253–276. [25] S. Ito, I. Kuraoka, P. Chymkowitch, E. Compe, A. Takedachi, C. Ishigami, F. Coin, J.M. Egly, K. Tanaka, XPG stabilizes TFIIH, allowing transactivation of nuclear receptors: implications for Cockayne syndrome in XP-G/CS patients, Molecular Cell 26 (2007) 231–243. [26] C.J. Cox, P.G. Foster, R.P. Hirt, S.R. Harris, T.M. Embley, The archaebacterial origin of eukaryotes, Proceedings of the National Academy of Sciences of the United States of America 105 (2008) 20356–20361. [27] Z. Kelman, M.F. White, Archaeal DNA replication and repair, Current Opinion in Microbiology 8 (2005) 669–676. [28] D.W. Grogan, Stability and repair of DNA in hyperthermophilic Archaea, Current Issues and Molecular Biology 6 (2004) 137–144. [29] X. Huang, M. Miller, A time-efficient, linear-space local similarity algorithm, Advances in Applied Mathematics 12 (1991) 337–357. [30] M.F. White, S.D. Bell, Holding it together: chromatin in the Archaea, Trends in Genetics 18 (2002) 621–626. [31] P. Lopez-Garcia, P. Forterre, DNA topology in hyperthermophilic archaea: reference states and their variation with growth phase, growth temperature, and temperature stresses, Molecular Microbiology 23 (1997) 1267–1279. [32] R.G. Collin, H.W. Morgan, D.R. Musgrave, R.M. Daniel, Distribution of reverse gyrase in representative species of eubacteria and archaebacteria, FEMS Microbiology Letters 55 (1988) 235–240.

H.M. Roth et al. / DNA Repair 11 (2012) 286–293 [33] D.R. Musgrave, K.M. Sandman, J.N. Reeve, DNA binding by the archaeal histone HMf results in positive supercoiling, Proceedings of the National Academy of Sciences of the United States of America 88 (1991) 10397–10401. [34] G. Perugino, A. Valenti, A. D’Amaro, M. Rossi, M. Ciaramella, Reverse gyrase and genome stability in hyperthermophilic organisms, Biochemical Society Transactions 37 (2009) 69–73.

293

[35] T. Kawashima, N. Amano, H. Koike, S. Makino, S. Higuchi, Y. Kawashima-Ohya, K. Watanabe, M. Yamazaki, K. Kanehori, T. Kawamoto, T. Nunoshiba, Y. Yamamoto, H. Aramaki, K. Makino, M. Suzuki, Archaeal adaptation to higher temperatures revealed by genomic sequence of Thermoplasma volcanium, Proceedings of the National Academy of Sciences of the United States of America 97 (2000) 14257–14262.