High yield expression and purification of HIV-1 Tat1−72 for structural studies

Journal of Virological Methods 164 (2010) 35–42

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High yield expression and purification of HIV-1 Tat1−72 for structural studies Shaheen Shojania, Gillian D. Henry, Vincent C. Chen, Thach N. Vo, Hélène Perreault, Joe D. O’Neil ∗ Department of Chemistry, University of Manitoba, Winnipeg, MB, R3T 2N2, Canada

a b s t r a c t Article history: Received 23 June 2009 Received in revised form 17 November 2009 Accepted 17 November 2009 Available online 24 November 2009 Keywords: HIV Tat Transcriptional regulator Intrinsically disordered protein Cysteine-rich NMR spectroscopy

The HIV-1 transactivator of transcription (Tat) is a protein essential for virus replication. Tat is an intrinsically disordered RNA-binding protein that, in cooperation with host cell factors cyclin T1 and cyclin-dependent kinase 9, regulates transcription at the level of elongation. Tat also interacts with numerous other intracellular and extracellular proteins, and is implicated in a number of pathogenic processes. The physico-chemical properties of Tat make it a particularly challenging target for structural studies: Tat contains seven Cys residues, six of which are essential for transactivation, and is highly susceptible to oxidative cross-linking and aggregation. In addition, a basic segment (residues 48–57) gives the protein a high net positive charge of +12 at pH 7, endowing it with a high affinity for anionic polymers and surfaces. In order to study the structure of Tat, both alone and in complex with partner molecules, we have developed a system for the bacterial expression and purification of 6×Histidine-tagged and isotopically enriched (in 15 N and 13 C) recombinant HIV-1 Tat1−72 (BH10 isolate) that yields large amounts of protein. These preparations have facilitated the assignment of 95% of the backbone NMR resonances. Analysis by mass spectrometry and NMR demonstrate that the cysteine-rich Tat protein is unambiguously reduced, monomeric, and unfolded in aqueous solution at pH 4. © 2009 Elsevier B.V. All rights reserved.

1. Introduction The HIV-1 transactivator of transcription (Tat) is a small, intrinsically disordered protein (Shojania and O’Neil, 2006) that is essential to the viral life cycle (Liang and Wainberg, 2002; Swanson and Malim, 2008). During transcription of the HIV viral DNA, Tat protein is transported from the cytoplasm into the nucleus where it binds to a viral RNA stem-loop structure referred to as the transactivation response (TAR) element. Tat then recruits the pos-

Abbreviations: ␤ME, ␤-mercaptoethanol; CDK9, cyclin-dependent kinase 9; DSS, 2,2-dimethyl-2-silapentane-5-sulfonate; DTT, dithiothreitol; HSQC, heteronuclear single quantum coherence; Hexim1, hexamethylene bisacetamide-inducible protein 1; HAD, HIV-associated dementia; HIVE, HIV-associated encephalitis; IPTG, isopropyl-␤-d-thiogalactopyranoside; MALDI-TOF-MS, matrix assisted laser desorption/ionization time-of-flight mass spectrometry; N-TEF, negative transcription elongation factor; P-TEFb, positive transcription elongation factor b; Tat, transactivator of transcription; PAGE, polyacrylamide gel electrophoresis; SDS, sodium dodecylsulfate; TAR, transactivation response; TB, terrific broth; SPE, solid phase extraction; Gdn-HCl, guanidine hydrochloride; TCEP, tris(2-carboxyethyl) phosphine; TCP, tris(2-cyanoethyl)phosphine; THP, tris(hydroxypropyl)phosphine; SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis. ∗ Corresponding author at: Department of Chemistry, University of Manitoba, 144 Dysart Rd., 390 Parker Building, Winnipeg, MB, R3T 2N2, Canada. Tel.: +1 204 474 6697; fax: +1 204 474 7608. E-mail addresses: [email protected] (S. Shojania), [email protected] (G.D. Henry), vince [email protected] (V.C. Chen), [email protected] (T.N. Vo), [email protected] (H. Perreault), [email protected] (J.D. O’Neil). 0166-0934/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.jviromet.2009.11.021

itive transcription elongation factor b (P-TEFb), a hetero-dimeric complex of a regulatory cyclin T and cyclin-dependent kinase 9 (CDK9), which it activates by displacing Hexim1 (hexamethylene bisacetamide-inducible protein 1) from its cyclin T1 binding site (Schulte et al., 2005). Activated CDK9 then phosphorylates RNA polymerase II, components of the negative transcription elongation factor (N-TEF), and the transcription elongation factor Spt5, stimulating elongation of full-length viral transcripts (Yamaguchi et al., 1999; Bourgeois et al., 2002; Kim et al., 2002). Absence of Tat and low levels of CDK9 and cyclin T1 in resting CD4+ T-cells are implicated in HIV-1 latency that permits the virus to evade host immune responses and antiretroviral drugs (Lassen et al., 2004). The affinity of the Tat-cyclin T1-CDK9 complex for TAR is regulated through Tat acetylation by histone acetyl transferase (Bannwarth and Gatignol, 2005; Mujtaba et al., 2002) and it was recently shown that the suppression of the MicroRNA-silencing pathway is dependent on the histone acetyl transferase Tat cofactor PCAF (Triboulet et al., 2007). Tat may also be involved in derepression of heterochromatin, transcription initiation (Pumfery et al., 2003), and reverse transcription (Guo et al., 2003). In addition, Tat has been implicated in a number of extracellular activities including supporting endothelial cell proliferation (contributing to the development of Kaposi’s sarcoma; Ensoli et al., 1990; Albini et al., 1996a,b), inducing apoptosis of T cells (Goldstein, 1996), stimulating cell death of neurons (Nath et al., 1996; Pocernich et al., 2005), and contributing to oxidative stress (Pocernich et al., 2005; Westendorp et al., 1995). In general, the cytotoxic activities of Tat contribute to both immune

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and non-immune dysfunction resulting in an overall increase in the burden of the viral infection. The 101-residue Tat protein is encoded by two exons and expressed during the early stages of viral infection. The first tat exon defines amino acids 1–72 that encompass an acidic and proline-rich N-terminus (1–21), a cysteine-rich region (22–37), a core (38–47), a basic region (48–57), and a Gln-rich segment (58–72) (Derse et al., 1991); it activates transcription with the same proficiency as the full-length protein (Jeang et al., 1999; Kuppuswamy et al., 1989; Garcia et al., 1988; Smith et al., 2003). The second tat exon defines residues 73–101 and includes an RGD motif that may mediate Tat binding to cell surface integrins (Avraham et al., 2004). The biological function of the second exon-encoded polypeptide has been difficult to determine (Guo et al., 2003; Smith et al., 2003). There are also several laboratory viral strains that produce a Tat protein that is 86 residues long that likely originate from the HXB2 virus (subtype B) commonly found in Europe and North America (Opi et al., 2004). The 86 residue variants do not seem to be found in host viral isolates (Jeang, 1996) and it has been suggested that they arose as a consequence of tissue culture passaging that accidentally introduced a stop codon through a single nucleotide change (Neuveut and Jeang, 1996). A molecular structural description of Tat’s many activities requires high-resolution structure determination of Tat alone and in complex with its multiple interaction partners (Dyer et al., 2008). However, the physico-chemical properties of the protein make it a particularly challenging target for structural studies: Tat contains seven Cys residues and is highly susceptible to oxidative crosslinking that produces covalent multimers and irreversible protein precipitation. In a detailed study, Siddappa et al. (2006) showed that only the monomeric fully reduced Tat protein activates transcription. One approach to this problem uses the Equine Infectious Anemia Virus (EIAV) Tat protein that contains only two Cys residues and is more easily expressed and purified. This approach was used to determine the structure of a fragment of EIAV Tat in a complex with EIAV TAR RNA and EIAV Cyclin T1 (Anand et al., 2008). However, this strategy will not provide insight into the role of the six essential Cys residues in the human HIV-1 Tat protein. In addition, the basic segment of Tat (residues 48–57) gives the Tat1−72 protein a high net positive charge of +12 at pH 7 endowing it with a high affinity for anionic polymers and surfaces. For example, Tat binds to TAR RNA and heparin with dissociation constants of 12 nM (Dingwall et al., 1990) and 0.37 ␮M, respectively (Hakansson and Caffrey, 2003). Tat is also prone to adhering to glass and some chromatographic resins and this can be problematic for purification. Tat has a low amino acid sequence complexity, low overall hydrophobicity, and a high net positive charge suggesting that it is an intrinsically disordered protein and unlikely to crystallize. Indeed the structure of the EIAV Tat is mostly irregular (Anand et al., 2008). Consequently, NMR spectroscopy is the preferred method for Tat conformational analysis (Shojania and O’Neil, 2006). However, isotope labeling with 15 N and 13 C is essential for NMR analysis of disordered proteins because the backbone 1 H resonances span a very narrow chemical shift range (Zhang et al., 1997; Yao et al., 1997). Isotopic labeling is also essential for studying the structures of proteins in complexes as isotopic enrichment of one species in the complex allows filtering of the NMR signals to reduce the complexity of the spectra. In order to study the structure of Tat, both alone and in complex with partner molecules, we have developed a system for the bacterial expression and purification of 6×Histidine-tagged and isotopically enriched (in 15 N and 15 N/13 C) recombinant HIV1 Tat (residues 1–72; BH10 isolate) that addresses the difficulties described above and yields large amounts of monomeric, reduced protein. The methods have been used to prepare NMR-quality samples for backbone spectral assignment using three-dimensional

NMR spectroscopy and for 15 N NMR relaxation studies of the protein (Shojania and O’Neil, 2006). 2. Materials and methods 2.1. Plasmid construction The Tat expression vector was constructed from the E. coli codon-optimized exon 1 tat gene (residues 1–72 of the HIV-1 BH10 isolate) contained in pSV2tat72 obtained through the AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, NIH from Dr. Alan Frankel (Frankel and Pabo, 1988). The tat gene was PCR amplified using pSV2tat72 as template and the following forward (Nde I) and reverse (Bgl II) primers: Nde I: 5 -ATGATCGTCATATGGAACCGGTCGAC CCGCGT-3 ; Bgl II: 5 -CCGGGAGATCTTCACTGTTTAGACAGA GAAACCTGGTGGGTC-3 . The amplified DNA was then ligated into pUC18 (Pierce, Milwaukee, WI) that had been opened with Sma I. The insert was DNA sequenced and the resulting plasmid is referred to as pUC18tat. The tat exon 1 gene from pUC18tat was removed using Nde I and Bgl II and the purified fragment ligated into pET28b(+) (Novagen, Madison, WI) that had been opened with Nde I and BamH I. The expression vector was verified using the PCR primers for sequencing. The pET28tat plasmid was transformed into NovaBlue cells (Novagen, Madison, WI) for plasmid storage and into E. coli BL21(DE3)plysS cells for protein expression with an N-terminal 6× His segment and thrombin cleavage site that adds 20 residues to the 72 residue protein. Glycerol stocks of the cells were obtained from a single colony grown at 37 ◦ C to an OD600 of 0.68 in 2 mL of Luria–Bertani (LB) medium and subsequently divided into 0.07 mL aliquots. The 0.07 mL aliquots of the stock cells were then combined with 0.03 mL of a 70% glycerol solution, resuspended, flash frozen in liquid nitrogen, and then stored at −72 ◦ C for later use. 2.2. Expression of unlabeled His-tagged Tat1−72 To test the expression system, initial experiments were done using non-labeling conditions for the over-expression of Tat. Transformed cells from a 100-␮L glycerol stock were grown up in 50 mL of terrific broth (TB) (Sigma, St. Louis, MO) inoculated with 34 ␮g/mL chloramphenicol and 30 ␮g/mL kanamycin for 16 h at 37 ◦ C in a rotary shaker. A 10 mL aliquot of the overgrown culture was then added to 1 L of pre-incubated (37 ◦ C) TB (with 34 ␮g/mL chloramphenicol and 30 ␮g/mL kanamycin) in a 2-L baffled flask. Cell growth was monitored by optical density measurements at 600 nm until the measured reading was 0.8. Expression was then initiated by induction with 60 mg of isopropyl␤-d-thiogalactopyranoside (IPTG) (Sigma, St. Louis, MO). Cells were allowed to express for 5 h before the cell culture was put on ice for 15 min to halt the expression. The cells were then collected by centrifugation at 2600 × g for 15 min, sealed in bottles under an argon atmosphere prior to freezing in liquid nitrogen, and stored at −72 ◦ C. 2.3. Expression of 13 C/15 N-His-tagged Tat1−72 The following expression protocol was modified from published methods (Marley et al., 2001) to reduce the consumption of isotopically-labeled ingredients. As with the unlabeled expression, cell growth was initiated from a 100-␮L glycerol stock of the pET28tat-transformed cells into 50 mL of TB (with 34 ␮g/mL

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chloramphenicol and 30 ␮g/mL kanamycin) and grown for approximately 15 h. Four 10 mL aliquots of the 50 mL overgrown cell culture were then used to inoculate 4×2 L baffled flasks each containing 1 L of pre-incubated (37 ◦ C) TB (with 34 ␮g/mL chloramphenicol and 30 ␮g/mL kanamycin). Cells were grown at 37 ◦ C in a rotary shaker; growth was halted when the optical density of each flask reached 0.6–0.9. Flasks were submerged in crushed ice for 15 min to halt cell growth and then cells were collected by centrifugation at 2600 × g at 4 ◦ C for 15 min. Cell pellets were resuspended in 40 mL of M9 salts solution to wash away residual rich medium, pooled, and then centrifuged again at 2600 × g for 15 min. The single pooled pellet was then re-suspended in 10 mL of the M9 wash solution and then added to 1 L of pre-incubated (37 ◦ C) M9 minimal medium (with 34 ␮g/mL chloramphenicol and 30 ␮g/mL kanamycin). The medium, adapted from (Neidhardt et al., 1974), contained 0.7 g 15 NH4 Cl and 2 g of 13 C-glucose (Cambridge Isotope Laboratories Inc., Andover, MA) and was supplemented with vitamins and micronutrients (see Table S1 Supplemental). The cells were allowed to adjust to the new medium for 15 min and then over-expression was induced upon addition of 240 mg of IPTG. Expression was stopped after 5 h and cells were harvested by centrifugation at 2600 × g at 4 ◦ C. Cell pellets were re-suspended with 4×10 mL of M9 wash solution, pooled, and centrifuged again at 2600 × g for 15 min. The supernatant was removed and the bottle sealed in an argon atmosphere prior to freezing in liquid nitrogen for storage at −72 ◦ C. 2.4. Purification of His-tagged Tat1−72 Cell lysis was achieved by two freeze–thaw cycles, each with a 30-min incubation period at room temperature following complete thawing of the pellet. DNase I and RNase I (Sigma, St. Louis, MO) were added to the lysate (200 ␮g of each) and incubated at 37 ◦ C for 30 min. A 100 mL aliquot of extraction buffer (see Table 1) was added to the lysate and the mixture was microprobe sonicated (twice at 35% power with 30 s bursts and 30 s between bursts) using a Fisher Sonic Dismembrator Model 300 (Fisher Scientific, Norcross, GA). The lysate was then centrifuged at 17,000 × g for 30 min, and the supernatant poured over a 4-mL bed of TalonTM (cobaltSuperflowTM ) metal affinity resin (Clonetech, Palo Alto, CA) in a 10-mL polypropylene gravity flow column (QIAGEN Inc., Mississauga, ON). Because of the expectation of higher yields of unlabeled protein, the extract was usually divided into two identical portions to avoid saturating the cobalt metal affinity resin. The resin had been previously washed and equilibrated with the extraction buffer. The resin was washed with 20 mL of additional extraction buffer followed by 30 mL of wash buffer (see Table 1). Tat protein was released from the cobalt column with elution buffer (see Table 1) and 10×1 mL fractions collected. The fractions were pooled and serially dialyzed at 25 ◦ C in regenerated cellulose tubing with MWCO of 1000 Da (Fisher HealthCare, Houston, TX) against 1 L of degassed acetate buffer at pH 3 at concentrations of 0.1, 0.05, and 0.01 M (approximately 6 h each). A final dialysis was done against degassed water for 4 h. Degassing of the buffers was achieved by

Table 1 Protein purification buffers. Buffer

pH

Composition

Extraction

7.2

6 M guanidine hydrogen chloride (Gdn-HCl); 100 mM sodium phosphate; 10 mM tris(hydroxymethyl) amino-methane hydrochloride (Tris–HCl); 10 mM tris(2-carboxyethyl) phosphine (TCEP)

Wash

6.4

6 M Gdn-HCl; 50 mM sodium phosphate; 10 mM TCEP

Elution

4

6 M Gdn-HCl; 50 mM sodium acetate; 10 mM TCEP

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three cycles of purging the oxygen from the solution for 15 min with argon gas followed by removal of dissolved gases under a vacuum for 15 min. Each of the dialysis buffers were sealed under an argon atmosphere. A 1 mL aliquot was removed from the dialysate for near ultra-violet (UV) absorbance analysis and mass spectrometric analysis; the remainder of the dialysate was frozen and freeze-dried. 2.5. Electrophoresis Tat1−72 was prepared for electrophoresis by boiling for 15 min in 70 mM Tris–HCl buffer pH 6.8 containing 2% sodium dodecylsulfate (SDS), 0.1% bromophenol blue, 6% glycerol, 2% DTT and either with or without 10 mM TCEP. Separation was by SDS-polyacrylamide gel electrophoresis (PAGE) in Laemmli discontinuous gels (Laemmli, 1970) composed of a 4% acrylamide stacking gel and a 12.5% resolving gel. After electrophoresis, proteins were visualized by staining with Coomassie Brilliant Blue R250. The pre-stained protein molecular weight markers (Fermentas, Burlington, ON) were ␤-galactosidase (118 kDa), bovine serum albumin (BSA) (85 kDa), ovalbumin (47 kDa), carbonic anhydrase (36 kDa), ˇ-lactoglobulin (26 kDa), and lysozyme (20 kDa). 2.6. MALDI-TOF-MS A 10 ␮L aliquot of the dialysate (from the unlabeled Tat purification) in aqueous solution was subjected to solid phase extraction (SPE) to remove unwanted salts and buffers using a Millipore C18 ZipTipTM (Billerica, MA) following the manufacturer’s recommended protocol. SPE-treated samples were concentrated by aspirating the SPE tip with 2 ␮L of 50:50 acetonitrile/water with 0.1% TFA. Samples were then mixed with 2 ␮L of sinapinic acid (3,5dimethoxy-4-hydroxycinnamic acid) matrix solution (Sigma, St. Louis, MO) saturated in water and transferred to a Bruker ScoutTM (Billerica, MA) 384 stainless steel target. Mass spectrometric analysis was performed on a Bruker BiflexTM IV MALDI-TOF instrument operated in positive, linear mode with acceleration potentials of 21 and 17 kV for lenses 1 and 2, respectively. The instrument was externally calibrated with the [M+H]+ and [M+2H]2+ ions of BSA (m/z 66,431 and 33,215) and myoglobin (m/z 16,952.62 and 8476.81). 2.7. NMR sample preparation Freeze-dried protein was dissolved in 600 ␮L of degassed buffer containing 50 mM acetate-d4 /ammonium hydroxide, 20 mM MES (2-(N-morpholino)ethanesulfonic acid) (only in 13 C/15 N-labeled sample), 80 ␮M sodium sulfite, 0.02% sodium azide and 5% D2 O. The resulting protein solutions were at pH 4 (unlabeled) and pH 4.1 (13 C/15 N-labeled). The samples were put into 5 mm NMR tubes (Wilmad-Labglass, Buena, NJ) that had been purged with argon gas for 15 min and the dissolved protein was added to the sample tube under an argon atmosphere. The NMR tube caps were then sealed with Teflon® tape (DuPont, Wilmington, Delaware). The final protein concentration in the NMR tube was 1.5 mM (unlabeled) and 1 mM (13 C/15 N-labeled) monomeric Tat. 2.8. NMR data acquisition 1 H/15 N-heteronuclear single quantum coherence (HSQC) experiments for the 92-residue unlabeled (using the natural abundance of the 15 N isotope for the indirect dimension) and 13 C/15 N-labeled His-tagged Tat 1−72 were acquired on a 600 MHz Varian INOVA spectrometer equipped with a triple resonance probe head at 20.2 ◦ C, using standard Varian BioPack pulse sequences (Kay et al., 1992). The NMR probe temperature was calibrated

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with methanol (Cavanagh et al., 1996) and spectra were processed with NMRPipe (Delaglio et al., 1995). HSQC experiments were collected with 2048 complex points in the direct dimension for both samples. For the indirect 15 N dimension, 256 and 128 complex points were collected for the 13 C/15 N-labeled and the unlabeled samples, respectively. Sweep widths in both experiments were 12 ppm in the direct and 36 ppm in the indirect dimensions. A total of 192 transients were collected on the unlabeled Tat protein whereas 32 transients were collected for the 13 C/15 N-labeled protein. Spectra were apodized using a squared cosine bell function, zero filled to twice (13 C/15 N-labeled) or four times (unlabeled) the data set size, and linear predicted (forward-backward with eight prediction coefficients) prior to Fourier transformation in the indirect dimension. The dimensions of the resulting processed data sets were 4096 × 1024 points for both 1 H/15 N-HSQC experiments. Non-linear line shape fitting was performed on the peaks in the spectrum of the unlabeled Tat sample and the noise was subtracted from the result. The HSQC pulse sequence is sensitivity-enhanced and uses gradients for coherence selection and water suppression (Farrow et al., 1994). Radiation damping was suppressed with a water flip-back pulse (1.42 ms). 15 N decoupling during acquisition was done using the WALTZ-16 sequence (Shaka et al., 1983) with a 7.2-kHz field strength. 1 H chemical shifts were referenced to the water signal that resonates 4.82 ppm from 2,2-dimethyl2-silapentane-5-sulfonate (DSS) at 293 K (Cavanagh et al., 1996). 15 N and 13 C referencing were done indirectly relative to DSS as recommended (Wishart et al., 1995). 3. Results and discussion 3.1. Expression plasmid design A number of approaches to increasing Tat1−72 expression and simplifying the purification have been attempted in the past including construction of Tat fusion proteins (Magnuson et al., 1995) and the use of the Tat transduction domain’s affinity for negatively charged polymers for purification (Hakansson et al., 2001). We chose the pSV2tat72 vector (Frankel and Pabo, 1988) as the source of the tat gene because it is partially codon-optimized for expression in E. coli. As Tat has well described cytotoxic effects, the pET28 plasmid was chosen for expression because of the stringent control over the lacUV5 promoter owing to the presence of the lac repressor (lacI) (Studier et al., 1990); the plasmid is also less easily lost from the cell because of the kanamycin resistance gene. The choice of a pLysS-containing host was also made with the toxicity of Tat in mind but has the added advantage of more facile cell lysis by the endogenous T7 lysozyme. The Tat1−72 gene was cloned into pET28 to enable expression with a 20-residue N-terminal, thrombin cleavable, 6× His purification tag. Metal affinity chromatography has been used to rapidly purify many proteins to greater than 95% purity in a single step and rapid purification was considered important for the highly oxidation-prone Tat protein. Another important advantage of metal affinity purification for Tat is the ability to elute the protein from the resin at low pH, preventing disulfide bond formation. 3.2. Protein expression and purification Growth of E. coli BL21(DE3)plysS cells containing pET28tat in terrific broth (TB) typically yielded about 10 g of cells (wet weight) per liter of TB medium. Yields of E. coli are reduced by about half when the cells are grown in 1 L of 13 C/15 N labeling medium (M9) using cells from 4 L of TB, as described in MATERIALS AND METHODS. Typically, in both unlabeled and labeled protein purifications, the protein dialysate is free of visible precipitate. UV absorbance

measurements of the protein dialysate at 280 nm (calculated ε280 = 9090 cm−1 × M−1 ; Gill and von Hippel, 1989) were used to determine protein yields of His-tagged Tat. Typically, up to 20 mg of unlabeled protein and 15 mg of 13 C/15 N-Tat1−72 are recovered from cells grown in 1 L of TB and minimal medium, respectively. Attempts to remove the 6× His affinity tag followed by repurification on the metal affinity column resulted in significant loss of protein. Possible reasons for the problems associated with thrombin cleavage are: the protein contains a potential internal thrombin cleavage site between Lys-61 and Ala-62 (Harris et al., 2000) and the protein contains a possible thrombin inhibitory segment Arg-Pro-Pro (residues 76–78) (Hasan et al., 1996). NMR analysis (Shojania and O’Neil, 2006) (below) suggests that there is no interaction between the affinity tag and any other segment of the protein and this has been generally found to be the case for a large number of proteins containing polyhistidine purification tags (Edwards et al., 2000). Numerous reducing agents were tested for the production of reduced, monomeric Tat1−72 , and TCEP, in the presence of 6 M guanidine, was found to be the most effective. TCEP is both a stronger reducing agent and effective over a wider pH range (1.5–8.8) than thiol reducing agents such as ␤-mercaptoethanol (␤ME) and dithiothreitol (DTT) (Han and Han, 1994). Its use permitted the entire purification, from cell lysis at neutral pH to elution from the cobalt resin at pH 4, to be done in a strong reducing environment. This is not possible with thiol reducing agents as neither DTT (pKa = 9.2, 10.1) nor ␤ ME (pKa = 9.6) are effective reducing agents at low pH (Gough et al., 2002). According to the manufacturers (Clonetech and QIAGEN), DTT is not compatible with metal affinity resins but we encountered no incompatibility between TCEP and immobilized Co2+ and Ni2+ . ␤ ME can be used up to concentrations of 20 mM in metal affinity chromatography (according to the manual from QIAGEN) but its effectiveness as a reducing agent is much less than that of DTT or TCEP especially at lower pH values. One problem with TCEP is that it has three negative charges at neutral pH and at high concentrations tends to precipitate the highly basic Tat protein. Including 6 M guanidine in the extraction buffer and then removing the guanidine and TCEP together at low pH by dialysis overcame this problem. Tris(2cyanoethyl)phosphine (TCP) is also a strong reducing agent and has the added advantage that it is neutral, does not precipitate Tat, and can access less solvent exposed sulfhydryls. Unfortunately, it is less soluble than TCEP and is significantly less stable than TCEP, oxidizing more readily in air. Tris(hydroxypropyl)phosphine (THP) was also investigated as a reducing agent as it is miscible with water and neutral. However, THP is a viscous liquid and very reactive. To deal with the short lifetime, attempts were made to degas and seal protein samples containing THP in the NMR tubes. However, it was found that the high viscosity of THP-aqueous Tat mixtures significantly increased the protein NMR line-widths. Another reducing agent that was tested was sodium sulfite. A small amount was added to the NMR sample at low pH and has the advantage of being invisible to 1 H NMR spectroscopy but is only a mild reducing agent in comparison to TCEP. The use of 6 M guanidine throughout the purification has several advantages over alternatives such as 8 M urea with or without 1–2 M NaCl. In comparison to urea, guanidine solutions consistently yielded the highest amounts of soluble Tat in the initial cell lysates. Presumably, high concentrations of guanidine encourage dissociation of Tat from DNA, RNA, and other anionic molecules. Furthermore, removal of the guanidine during the washing steps of the protein while bound to the metal affinity resin resulted in very slow elution of the protein from the resin suggesting that the Arg-rich basic domain of the protein can interact with the nitrilotriacetic acid groups of the Sepharose resin that have lost their coordinated metal ion. In the presence of 6 M guanidine at pH 4,

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Fig. 1. SDS-PAGE. Coomassie-stained polyacrylamide gel of HIV-1 His-taggedTat1−72 purified using immobilized Co2+ -nitrilotriacetic acid affinity chromatography. Lanes 1 and 2 are different column fractions from one preparation without TCEP. Lane 3 is a column fraction from a different preparation without TCEP. Lane 4 molecular weight markers indicated in Materials and methods. Lanes 5 and 6 show the same material as in Lanes 1 and 2, respectively, with 10 mM TCEP.

70% of the Tat protein eluted from the immobilized metal resin in two 1 mL fractions (based on UV absorbance measurements with spectroscopic grade guanidine hydrogen chloride) whereas in the absence of guanidine the same amount of protein eluted in approximately 40 mL. Purification with denaturant used throughout each step followed by subsequent removal during dialysis was found to be the most efficient way to obtain large quantities of reduced, monomeric protein.

39

Fig. 2. MALDI-TOF-MS. MALDI-TOF-MS identification of monomeric His-tagged Tat1−72 .

Removal of the denaturant and reducing agent by dialysis necessitated the use of low pH buffers to maintain the cysteine thiol groups in their unreactive protonated state. Degassing the dialysis and NMR buffers as well as maintaining an argon atmosphere further reduces the possibility of oxidation of the protein. TCEP was not used as a reducing agent in the NMR samples because it tends to precipitate Tat at neutral pH. Instead, rigorous degassing of the sample buffer and addition of a mild reducing agent (sodium sul-

Fig. 3. Two-dimensional NMR of Tat. (a) The amino acid sequence of Tat1−72 . The 6× His purification tag is indicated in bold. Amide backbone regions of 1 H/15 N-HSQC spectra for: (b) naturally abundant 15 N in unlabeled His-tagged Tat1−72 ; (c) 13 C/15 N-labeled His-tagged Tat1−72 .

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fite) allowed preparation of NMR samples that were stable for more than 6 months. 3.3. MALDI-TOF-MS and SDS-PAGE Even in the presence of high concentrations of DTT and ␤ME, Tat1−72 forms a ladder of oligomerized protein on SDS-PAGE gels (Fig. 1: Lanes 1–3). Occasionally, large soluble oligomers form that can pass through the stacking gel but are too large to enter the running gel (Fig. 1: Lane 1). The dark background in Lanes 1–3 in Fig. 1 suggests that some oxidation is occurring during the electrophoresis of the proteins. The possibility that some of the bands observed in Lanes 1–3 are from protein impurities is ruled out by the observation that all of the bands disappear when 10 mM TCEP is included in the sample preparation buffer except the monomeric Tat protein and a very small amount of dimer (Fig. 1: Lanes 5–6). The high molecular weight oligomer in Lane 6 illustrates that whereas TCEP is excellent at maintaining the protein in a reduced monomeric state it is less effective at reversing the oxidation once high molecular weight oligomers have formed. No amount of TCEP will solubilize oxidized Tat1−72 once it has precipitated from solution. Note that, because of its high positive charge, the monomeric Tat protein and its oligomers electrophorese more slowly than expected on the basis of masses that are multiples of 10,509. As indicated in Fig. 2, the use of mass spectrometry, and in particular MALDI-MS, is an effective approach to ascertaining both the purity and the oligomeric state of the protein. A significant advantage of MALDI-MS over SDS-PAGE is that the former method can maintain the protein at a low pH where the cysteine residues

are protonated and unreactive. The MALDI-TOF mass spectrum shown in Fig. 2 indicates that there is one predominant peak at 10,519.8 Da corresponding to the [M+H]+ species for the unlabeled His-tagged Tat monomer (calculated MW 10,509.076 Da). A second, less intense peak at 5256.5 Da, is likely the [M+2H]2+ peak. These two peaks corresponding to the monomeric Tat protein make up 89% of the total intensity of the non-matrix related peaks. The additional weak peaks at 7376.4, 21,028.7, and 31,345.6 Da are most likely the [2M+3H]3+ , [2M+H]+ , and [3M+H]+ species, respectively. Similar peaks are often observed in MS and are usually ascribed to non-covalent protein oligomer formation mediated by interactions between basic residues (Arg, Lys, and His) and acidic residues (Asp and Glu) in proteins (Vertes et al., 2000). The low intensity of these peaks in the present spectrum may be explained by the high net positive charge on Tat at low pH suggesting that there is minimal Coulombic attraction between the proteins. 3.4. NMR spectroscopy Dissolution of freeze-dried protein dialysate at pH 4 usually yielded solutions free of visible precipitate and free of suspended material judging from the near UV absorption spectra. The natural abundance 1 H/15 N-HSQC spectrum of unlabeled Tat1−72 shown in Fig. 3(b) shows 64 of the 83 observable amide backbone resonances (non-proline and non-N-terminal) as well as 9 peaks corresponding to the Arg, Gln, and Asn side chain resonances. The 1 H/15 N-HSQC spectrum of the 13 C/15 N-labeled Tat1−72 (Fig. 3(c)) shows several additional resonances that correspond to missing backbone resonances from the unlabeled sample as well as some additional

Fig. 4. The cysteine region of Tat. Strip plots from an HNCACB spectrum of the Cys-rich region of 13 C/15 N-labeled His-tagged Tat1−72 . Indicated are the chemical shifts of the C␣ and C␤ resonances for the seven Cys residues, C54, C45, C47, C50, C51, C42, and C57 from left to right, and the residues preceding the cysteines.

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weaker resonances that correspond to backbone residues that are undergoing slow conformational exchange (ms–s range). In general, both samples show cross-peaks regionally clustered in a manner typical for denatured or disordered proteins: a Gly region, a Ser/Thr region and a region containing the rest of the backbone amides (Schwarzinger et al., 2000). The spectral dispersion of the resonances is also typical for proteins lacking regular secondary structure in that all backbone resonances lie within a 1.1-ppm range in the 1 H-dimension and within 20 ppm in the 15 N-dimension (Peti et al., 2001; Dyson and Wright, 1998). The peak widths of the resonances are broad relative to those of folded proteins of comparable size, which is indicative of conformational exchange on the intermediate NMR time scale (␮s–ms range). Line broadening can also result from hydrogen exchange with the water solvent. As these Tat samples are at low pH (∼ 4 in both cases) where the rate of hydrogen exchange is near its minimum (Wüthrich, 1986; Eriksson et al., 1995), it is also not likely that exchange with the solvent is the cause of line broadening. Hence, the broad line widths observed for Tat are most likely the result of conformational exchange in the ␮s–ms range as one would expect for a protein that lacks regular secondary structural elements. The observed chemical shifts of the cross-peaks do not differ significantly between the two samples indicating that the proteins are in the same conformational state. Peaks that are missing in the spectrum of the unlabeled protein correspond to those that are of relatively weak intensity in the 13 C/15 N-labeled protein and are therefore absent due to the sensitivity limitations of the natural abundance experiment. Many of the missing peaks in the natural abundance HSQC spectrum correspond to amide backbone resonances in the Cys-rich and core regions of the protein (Shojania and O’Neil, 2006). These resonances are the weakest in the spectrum and in some cases are those with multiple signals found in the HSQC spectrum of the 13 C/15 N-labeled Tat. The conformational exchange on the ␮s–ms time-scale in these regions implies possible transient structural formation, which may only become stabilized in the presence of zinc ions, binding to TAR, cyclin T1, or other binding partners. The lack of additional unassigned peaks in the NMR spectrum of the unlabeled Tat indicates a high level of purity since other unlabeled proteins in the sample would also produce signals through their 15 N natural abundance and confirms the MALDI-TOFMS analysis. The backbone assignment described in (Shojania and O’Neil, 2006) resulted in unambiguous assignment of 80 of the 83 observable (non-proline and non-N-terminal) backbone amide resonances. The assignments of the Cys residues are particularly informative as they confirm that all of the Cys residues are reduced; all of the 13 C˛ and 13 Cˇ chemical shifts (shown in Fig. 4) observed in the three-dimensional HNCACB (Wittekind and Mueller, 1993) spectrum, are in the range of the random coil chemical shifts of reduced cysteine (58.6 and 28.3 ppm) (Schwarzinger et al., 2000) differing significantly from those of oxidized cysteine (55.6 and 41.2 ppm) involved in disulfide bond formation. The chemical shift resonances for the Cys residues thus confirm the findings from the MALDI-TOF-MS analysis that the protein is in the reduced monomeric state and that the weak peaks in Fig. 2 most likely indicate the presence of non-covalent oligomers formed during the MS analysis. 4. Conclusions We have identified both an efficient method for the bacterial over-expression of uniformly labeled Tat1−72 with 15 N and 15 N/13 C, and a rapid purification protocol based on a 6× His affinity tag purification by metal affinity chromatography. This expression and purification system yields on the order of 15 mg of the uniformly labeled protein per litre of labeling medium and the yield could

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be increased with longer expression times. Both MALDI-TOF-MS and NMR spectroscopy have shown that the resulting protein is unambiguously reduced and monomeric in solution at pH 4. This expression system provided sufficient protein for detailed structural and dynamic analysis of Tat (Shojania and O’Neil, 2006) and will be used to study its interaction with potential binding partners using heteronuclear NMR methods. Thus far, this is the only system for the expression of uniformly 15 N/13 C-labeled Tat and paves the way for a number of potential studies of Tat interactions with TAR and host cell proteins involved in the regulation of HIV gene expression as well as those involved in Tat’s extracellular neurotoxic activities. Acknowledgements This work was supported by grants from the Natural Sciences and Engineering Research Council of Canada and the University of Manitoba; it was initiated with funding from the Medical Research Council of Canada and the Manitoba Health Research Council. We thank Kirk Marat for assistance and training at the NMR facility at the University of Manitoba and the Canadian Foundation for Innovation for funding the 600 MHz spectrometer. We also thank Richard Sparling of the Department of Microbiology at the University of Manitoba, and members of his lab for use of their glove bag. Appendix A. Supplementary Data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.jviromet.2009.11.021. References Albini, A., Benelli, R., Presta, M., Rusnati, M., Ziche, M., Rubartelli, A., Paglialunga, G., Bussolino, F., Noonan, D., 1996a. HIV-Tat protein is a heparin-binding angiogenic growth factor. Oncogene 12 (January(2)), 289–297. Albini, A., Soldi, R., Giunciuglio, D., Giraudo, E., Benelli, R., Primo, L., Noonan, D., Salio, M., Camussi, G., Rockl, W., Bussolino, F., 1996b. The angiogenesis induced by HIV1 Tat protein is mediated by the flk-1/kdr receptor on vascular endothelial cells. Nature Medicine 2 (December(12)), 1371–1375. Anand, K., Schulte, A., Vogel-Bachmayr, K., Scheffzek, K., Geyer, M., 2008. Structural insights into the cyclin T1-Tat-TAR RNA transcription activation complex from EIAV. Nat. Struct. Mol. Biol. 15 (12), 1287–1292. Avraham, H.K., Jiang, S., Lee, T.H., Prakash, O., Avraham, S., 2004. HIV-1 Tat-mediated effects on focal adhesion assembly and permeability in brain microvascular endothelial cells. Journal of Immunology 173 (November(10)), 6228–6233. Bannwarth, S., Gatignol, A., 2005. HIV-1 Tat RNA: the target of molecular interactions between the virus and its host. Current HIV Research 3 (January(1)), 61–71. Bourgeois, C.F., Kim, Y.K., Churcher, M.J., West, M.J., Karn, J., 2002. Spt5 cooperates with human immunodeficiency virus type 1 Tat by preventing premature RNA release at terminator sequences. Molecular and Cellular Biology 22 (February(4)), 1079–1093. Cavanagh, J., Fairbrother, W.J., Palmer, A.G., Skelton, N.J., 1996. Protein NMR Spectroscopy: Principles and Practice. Academic Press, San Diego. Delaglio, F., Grzesiek, S., Vuister, G.W., Zhu, G., Pfeifer, J., Bax, A., 1995. Nmrpipe: a multidimensional spectral processing system based on unix pipes. Journal of Biomolecular NMR 6 (November(3)), 277–293. Derse, D., Carvalho, M., Carroll, R., Peterlin, B.M., 1991. A minimal lentivirus Tat. Journal of Virology 65 (December(12)), 7012–7015. Dingwall, C., Ernberg, I., Gait, M.J., Green, S.M., Heaphy, S., Karn, J., Lowe, A.D., Singh, M., Skinner, M.A., 1990. HIV-1 Tat protein stimulates transcription by binding to a u-rich bulge in the stem of the TAR RNA structure. EMBO Journal 9 (December(12)), 4145–4153. Dyson, H.J., Wright, P.E., 1998. Equilibrium NMR studies of unfolded and partially folded proteins. Nature Structural Biology 5 (July), 499–503. Dyer, M.D., Murali, T.M., Sobral, B.W., 2008. The landscape of human proteins interacting with viruses and other pathogens. PLoS Pathog 4 (2), e32. Edwards, A.M., Arrowsmith, C.H., Christendat, D., Dharamsi, A., Friesen, J.D., Greenblatt, J.F., Vedadi, M., 2000. Protein production: feeding the crystallographers and NMR spectroscopists. Nature Structural Biology 7, 970–972. Ensoli, B., Barillari, G., Salahuddin, S.Z., Gallo, R.C., Wong-Staal, F., 1990. Tat protein of HIV-1 stimulates growth of cells derived from Kaposi’s sarcoma lesions of aids patients. Nature 345 (May(6270)), 84–86. Eriksson, M.A., Härd, T., Nilsson, L., 1995. On the pH dependence of amide proton exchange rates in proteins. Biophysical Journal 69 (August(2)), 329–339.

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