Utilizing RNA Aptamers To Probe a Physiologically Important Heme

Verderber , E., Lucast , L. J., VanDehy , J. A., Cozart , P., Etter , J. B. and Best , E. A. (1997) Role of the hemA gene product and delta-aminolevul...
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ARTICLE

Utilizing RNA Aptamers To Probe a Physiologically Important Heme-Regulated Cellular Network Jacquin C. Niles† and Michael A. Marletta†,‡,§,*

Departments of †Chemistry and ‡Molecular and Cell Biology, University of California, Berkeley, Berkeley, California 94720, § Division of Physical Biosciences, Lawrence Berkeley National Laboratory, Berkeley, California 94720

R

ecent advances in genomics and proteomics are increasing our understanding of transcriptional and post-transcriptional gene regulation and how various gene products integrate into networks (1–4). Understanding the functional importance of specific proteins in these contexts has been aided by several methods, including targeted gene knockouts/ mutant collections, RNA interference in permissive organisms, and yeast two-hybrid studies, used in combination with microarray transcriptional profiling and mass spectrometry (3, 5–10). While these strategies are facilitating the elucidation of protein function, less attention has been focused on the role biologically important small molecules play in modulating protein networks. It is known that small molecules can play important regulatory roles. For example, the cofactor NAD, in addition to a role in energy homeostasis, binds directly to Sir2 and orthologous histone deacetylases and regulates chromatin silencing, aging in response to caloric restriction, and repression of p53mediated apoptosis in response to DNA damage (11, 12). The reduced form of this cofactor, NADH, binds the transcription repressor Rex, facilitating high-affinity DNA binding of this protein, and regulates expression of several respiratory genes (13). Heme is also known to interact specifically with heme-responsive motifs on the transcription factor Hap-1, leading to transcriptional activation of genes involved in oxidative respiration (e.g., cytochrome c) and the response to oxidative stress (e.g., catalase and flavohemoglobin genes) (14). Similarly, heme binding to the transcription repressor Bach1 derepresses gene expression (15). Given the integration of small molecules into these critical circuits, broadly www.acschemicalbiology.org

A B S T R A C T Broadly applicable strategies facilitating direct and selective modulation of the intracellular levels of physiologically important small molecules are essential for dissecting their integral and multiple roles in cellular processes. Therefore, we have been exploring the suitability of RNA aptamers for this purpose. Using the Escherichia coli heme biosynthetic pathway as a simple model of a negative feedback regulated process, we show that heme-binding RNA aptamers, developed in vitro and expressed intracellularly, induce a heme-dependent growth defect in an E. coli heme auxotroph defective in converting ␦-aminolevulinic (␦-ALA) acid into downstream products. Relative to a control oligonucleotide, the aptamers also induce ␦-ALA accumulation in cells grown under heme-limiting conditions. Increasing the concentration of heme in the media completely reverses both the growth defect and ␦-ALA accumulation, except for two aptamers for which reversal is partial. Thus, these aptamers specifically target their cognate ligand in vivo and functionally modulate its intracellular concentration, demonstrating that RNA aptamers are useful tools for elucidating the role of heme and possibly other small molecules in regulating cellular networks.

*Corresponding author, [email protected].

Received for review June 14, 2006 and accepted August 8, 2006. Published online September 8, 2006 10.1021/cb6002527 CCC: $33.50 © 2006 by American Chemical Society

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Figure 1. Schematic of the E. coli heme biosynthetic pathway. tRNAGlu ⴝ glutamyl tRNA; GSA ⴝ glutamate-1semialdehyde; ␦-ALA ⴝ ␦aminolevulinic acid; PBG ⴝ porphobilinogen; HemA ⴝ glutamyl-tRNA reductase; HemL ⴝ glutamate-1-semialdehyde aminotransferase; HemB ⴝ ␦-aminolevulinic acid dehydratase.

applicable strategies that facilitate the systematic elucidation of their roles in these contexts are required to improve our understanding of cellular physiology. Nucleic acid aptamers have several characteristics that make them attractive tools in achieving this goal. First, aptamers naturally encoded in the 5=-untranslated region of several genes play integral roles in sensing and regulating gene expression in response to the intracellular concentrations of small molecules such as amino acids (16, 17), enzyme cofactors (18), purines (19, 20), and Mg2⫹ (21). Second, aptamers can be developed in vitro using the readily accessible systematic evolution of ligands by exponential enrichment (SELEX) procedure (22, 23) and can conceivably be evolved to bind any target, as exemplified by the wide range of small molecule, peptide, and protein aptamers previously described (24, 25). While proteins binding these small molecules may exist, aptamers offer greater flexibility since they can be “tailor-made” to have specifically desired properties (e.g., dissociation constants). Third, RNA aptamers can be expressed intracellularly, and their impact on cellular physiology can be examined using simple phenotypic screens (e.g., growth rate) or global approaches such as microarray, proteomic, and metabolomic analyses. Last, aptamers selectively targeting specific proteins have been used to investigate cellsignaling pathways, as demonstrated by inhibition of the mitogen-activated protein kinase (MAPK) pathway by aptamers targeting extracellular-regulated kinase (ERK1/2) in vitro (26) and cytohesin-2 in vivo (27), indicating that they are useful tools for probing important protein-based networks. To evaluate the hypothesis that nucleic acid aptamers can be used to explore the role of small molecules in regulating cellular pathways, we have used Escherichia 516

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coli heme biosynthesis as a model for a product feedback inhibited system (Figure 1). Reduction of transfer RNA (tRNA)Glu by glutamyl-tRNA reductase (HemA) is both the rate-determining (28) and regulated step in this pathway. There are at least two mechanisms governing bacterial HemA regulation, namely, (i) direct interaction of heme with HemA, causing a decrease in its enzymatic activity (29, 30), and (ii) heme-dependent proteolytic HemA degradation when intracellular heme is nonlimiting (31). In both cases, the net effect is that HemA is highly active and flux through the biosynthetic pathway is elevated when intracellular heme levels are low and conversely when the cell is replete with heme. Here we show that intracellular expression of in vitro evolved heme-binding RNA aptamers in E. coli predictably modulates this important biofeedback loop. RESULTS AND DISCUSSION In Vitro Aptamer Selection. Our first objective was to establish high-affinity, expressible heme-binding RNA aptamers for in vivo studies. Single-stranded DNA and 2=-NH2-modified RNA aptamers targeting other porphyrins, such as N-methylmesoporphyrin IX (NMM) and hematoporphyrin IX (HPIX) have previously been reported (32–35). While these aptamers bind heme, they are unsuitable for our purposes, since they cannot be expressed or loaded into E. coli at functionally important concentrations. Furthermore, attempts to convert both the DNA and 2=-NH2 RNA aptamers into expressible 2=-OH aptamers have resulted in significantly decreased binding affinity (35, 36), making this strategy highly inefficient. Thus, a 2=-OH RNA heme-binding aptamer library had to be evolved de novo for our experiments. Aptamers were generated using the in vitro SELEX method from a starting library containing 6 ⫻ 1013 unique sequences and a 50-nucleotide variable region. The RNA library, body-labeled with [␣]-33P-ATP to facilitate monitoring the selection process, was applied to a column containing immobilized mesoprotoporphyrin IX (MPIX) to retain aptamers having the ability to bind the protoporphyrin IX (PPIX) scaffold. After the column was washed to remove nonspecifically bound sequences, heme-binding aptamers were specifically eluted with 2.5 mM heme in selection buffer. Library selection was judged complete after eight rounds, when the recovered RNA postselection was the same as in the previous round (Figure 2, panel a). The ability of the round 7 selected library to bind PPIX was qualitatively assessed www.acschemicalbiology.org

ARTICLE relative to the round 2 library using fluorescence spectroscopy. Free PPIX, when excited at 400 nm gives an emission spectrum with a maximum at 620 nm, and this remains unchanged upon incubation with round 2 library. However, incubation of the round 7 selected library with PPIX results in a bathochromic shift in the emission maximum to 635 nm along with an increase in the emission intensity, indicative of a binding interaction (Figure 2, panel b). The rounds 6 and 8 DNA libraries were blunt-endcloned into a plasmid vector and transformed into E. coli, from which individual aptamers for sequencing, binding affinity characterization, and archiving were obtained. Sequence data obtained for two and 28 aptamers from the round 6 and 8 pools, respectively, are summarized in Table 1. Analysis using the MEME algorithm (37) revealed the occurrence of a highly conserved G-rich motif in the majority of sequenced clones (Table 1). Interestingly, 6-5 and several round 8 aptamers do not contain this consensus sequence. These may represent distinct heme-binding classes or RNA molecules with no significant heme affinity that bind well to the solid support in spite of negative selection. For 6-5, the latter appears to be the case (vide infra). The majority of sequenced aptamers, however, contain this G-rich motif, suggesting that it represents a conserved aptamer element that is important in heme binding. The motif selected in our experiments, where heme (FePPIX) is used to specifically elute 2=-OH-RNA aptamers, resembles those obtained during the selection of DNA and 2=-NH2-RNA aptamers to NMM and HPIX, respectively (32–34). This suggests that the PPIX scaffold is the predominant binding surface interacting with selected aptamers and that the electronic or steric changes upon metal coordination (heme), pyrrole N-methylation (NMM), and hydration of the vinyl side chains (HPIX) do not significantly alter this interaction. The observation that these aptamers bind interchangeably with the different PPIX derivatives supports this conclusion. Next, the heme-binding affinity of selected aptamers was determined using UV–vis spectroscopy monitored heme titrations. Upon binding heme, aptamers induce a shift in the Soret maximum from 396 nm for free heme to ⬃405–408 nm, with slight hyperchromicity. By titrating heme into aptamer containing solutions and monitoring by difference UV–vis spectroscopy, we determined apparent dissociation constants (Kd) for the binding interaction, and these are summarized in www.acschemicalbiology.org

Figure 2. Monitoring of the aptamer selection progress. a) The in vitro selection protocol and progress monitored by 33P-labeled RNA recovery at the end of each round; * indicates the amount of RNA (pmols) applied to the subtraction or selection column; ** indicates that Ve is the number of column volumes of selection buffer used during the washing step. b) Fluorescence emission spectra for PPIX only, PPIX ⴙ round 2 pool (negative control), PPIX ⴙ round 7 pool, and PPIX ⴙ PS2.M (positive control ⴝ DNA heme-binding aptamer reported in ref 25). The shift in emission maximum from 600 nm for free PPIX to 623 nm is indicative of a binding interaction between round 7 pool and PPIX.

Table 1. For the tested aptamers, the apparent Kd’s ranged between 190 and 450 nM, indicating a reasonably high-affinity interaction. A dissociation constant for 6-5 could not be determined, since the UV–vis spectrum for 6-5 with heme was indistinguishable from that of free heme (Figure 3). Co-incubating equimolar concentrations of 6-5 and 8-13, which binds heme and induces a shift in the heme Soret maximum, with one heme equivalent produces a spectrum practically identical to that of heme and 8-13 alone (data not shown). Indeed, if the heme-binding affinity of 6-5 and 8-13 were comparable, a Soret shift toward the 6-5/heme maximum would be expected. Thus, 6-5 does not contain the consensus G-rich motif and does not appear to have significant heme-binding affinity. The selected G-rich motif, which may form G-quartets (34), is directly involved in heme binding, as demonstrated by the ability of the chemically synthesized G-rich elements from 8-8 and 8-34 to also bind heme (data not shown), and this finding is consistent with earlier reports (33, 34). Interestingly, the apparent dissociation constants for the motifs from 8-8 and 8-34 are 1.8 and 4.5 ␮M, respectively, indicating that these constructs bind heme less tightly than the full-length aptamers by ⬃4–10-fold. This suggests that unidentified cis- or VOL.1 NO.8 • 515–524 • 2006

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TABLE 1.

The randomized region of sequenced aptamers

Aptamera

Screenedb

Sequencec

Kd (nM)d

6-3 6-5 8-1 8-2 8-3 8-5 8-6 8-7 8-8 8-12 8-13 8-19 8-20 8-26 8-27 8-28 8-30 8-31 8-32 8-33 8-34 8-35 8-37 8-39 8-40

Yes Yes Yes

UGGUUUCAGCGACAGGAGGGGUGUAGGUGGAUUGCUGUCCUUUGCGUGU CUUAUGCAGUUUUACAGGGGUGAUAUGACUGCACCAGUAGGGGGAUGUUC UGCUAGUGUUGUAUGCACGUGGAGGAGGAGGCGUACACUUGCUUUGUGGU (2) UGCUAGUAUUGUAUGCACGUGGAGGAGGAGGCAGAAAAGCGCUUUAGGGUGUUAC UGCAGAAGUUGCGUGUGGAGGAGGUGGCAUGACUGCUGUUAGGGUAGUUG GGGAUCUGGUUGAAACUGGAGGCCUAUAGAAGUUGGUUGUGUGUUUUGAG GUCCUGUGAUGGUAUUUUUCGUUCCGCUUACUUCGACAUGAGGCCCGGAUCCAUCUGAAU GAUUGACCGUAUGGAGGAUGCAAAGGGAGGGAGGUCACUUGAGUUAGUUA GCAGGAUGUGGAGGAGGCAUCUGCUGCAAUCGGGACUUGUGUCGAGUAUC (4) GCAUUGUCUGCGUGUGGAGGCAGGAGGCAAGAUAAGAGGUGAUGCGGUUG CAUGUUGGCGAUACGUCUAAACGGUGGGUUGUGGAGGAUUGAUUUAUACG AGUAGUGUCAGCGUGUGGUGGAGGUUGGCGACAUAUGUAGGGUGCGAUUG CGAAGGCACUUCAUGGGGUGGAGGAGGCAUGCGAGGUGUCCGGCGAGUGG CACACGUGACUGUGGAGGCAGCGGAGGCGAGUUAUGUGAUGUUAAGAGGU UAGGGUGAUUGUUGCUAGAGAUGGCAUGAAA UAUGUUAAGAGGCCACUGAUGCGCGUAGGUCUCUGGGGAUUGAGGAAGGU AGGUUGCGCUAGGUGAGGAAGGAGGUGUAGGUACGGCCUAUUGAGUGGGA CGUAGUCCAUGAGUGUCUUUAGCUAACGGUUGGUAGUGAACCAUAUCCUG GCCAAUGAGAGCUGUAGGAGGGCGGGACGUGCUUAGUGCGUGACACCGGA UUGUCCUGACUUGCUUGAACGUUAGCGUGAUGCGUUAUGCCCUGGAUGGG CACCAAUGACGGGGGUUAAGACGGAGGGAGAUGCAUCGGUGUGAAGCUGA UGCGCAAUACACGGUGAGGAGGUGGAGAGAUGUAGGUGCUUAGCAGUUGA CGUGAACGCAUGUGGUGGAGGAGGCGAUUGCACGUGGGACCGAGCAUUUG GAUGUANCGGUGUCUUAGCCUUGUGGGANUAGGGUGCGUAUGGGGAUGNC UGGACCGCAGCACGGCGCUCGUGGUAAGGCCGUAUGCCCAUCGAAUGAAG

188 ND 309

Yes Yes Yes Yes Yes

Yes

256 220 371 445

425

Aptamers with demonstrated PPIX scaffold binding are italicized, and ␦-ALA levels were determined for cells expressing aptamers shown in boldface. Aptamers screened in the E. coli growth assay are indicated. cThe conserved motif identified using MEME is displayed as a sequence logo (generated online at weblogo.cbr.nrc.ca/logo.cgi). G-rich regions within aptamer sequences are shown in italic boldface; two poly-G regions in the non-hemebinding control oligonucleotide 6-5 that do not conform to the G-rich consensus are underlined. The number of clones harboring a given sequence is shown in parentheses. dThe heme dissociation constants in select cases. ND ⫽ not detected.

a

b

trans-acting structural elements in the full-length aptamer may be stabilizing heme binding to the G-rich motif. Impact of Heme-Binding Aptamers on E. coli Growth. We hypothesized that intracellular aptamer expression will lead to heme sequestration and impair aerobic bacterial growth. The heme-permeable E. coli heme auxotroph RP523 was selected for these studies for two 518

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main reasons. First, prototrophic E. coli strains can significantly upregulate heme biosynthesis (38, 39), decreasing the likelihood that subtle growth phenotypes due to intracellular heme sequestration will be detectable. Second, growth of the auxotroph is strictly dependent upon heme added to the media, and this can be directly controlled experimentally. For testing in E. coli, www.acschemicalbiology.org

ARTICLE Figure 3. UV–vis spectral properties and titration curve for selected aptamers. a) UV–vis spectra for heme alone and with 6-5, 8-7, and 8-8. b) Representative difference spectra obtained during heme titrations with aptamers. c) Representative plot of ⌬A405nm versus log[heme] fit to eq 1 used to determine dissociation constants for heme binding to aptamers.

several characterized aptamers were cloned into the RNA expression vector pGFIB, which is a high-copy plasmid in which transcription is initiated by the strong and constitutively active llp promoter and terminated by the efficient rrnC terminator (40). From this plasmid, tRNAAla, which is similar in size to our aptamers, is synthesized in high abundance, with levels ⬃70 ⫻ that of chromosomally encoded tRNAAla (41). RP523 cells were transformed with several pGFIB– aptamer constructs, and production of full-length aptamer was ascertained by chain reaction (RT-PCR) for several aptamers (data not shown). To screen for an aptamer-induced growth defect, cells were grown aerobically overnight in 1 mL cultures supplemented with 0.5, 1, 2, and 4 ␮M heme added to the media. The extent of bacterial growth was determined by OD600 readings and normalized to RP523 harboring the nonheme-binding construct pGFIB–6-5. Overall, final bacterial density attained increases directly with heme concentration, as expected. For RP523 not harboring plasmid, the EC50 for maximal growth is ⬃1.5 ␮M, and heme concentrations ⱖ4 ␮M are no longer growth limiting (data not shown). These characteristics are also true for RP523 harboring pGFIB–6-5, although there is a slight decrease in the final bacterial densities achieved at a given heme concentration. The results of screening eight selected aptamers are summarized (Figure 4, panel a) and illustrate that RP523 harboring 8-12, 8-13, 8-19, and 8-35 show dramatically less growth relative to 6-5 at 0.5–1 ␮M heme, while 6-3, 8-1, and 8-7 show a trend toward decreased growth at 0.5 ␮M heme, but this is not statistically significant. Growth of RP523 expresswww.acschemicalbiology.org

ing 8-8 is not impaired relative to 6-5 at all of the tested heme concentrations. Thus, the aptamers are differentially effective at impairing bacterial growth. Importantly, the growth defect induced is fully reversed by increasing media heme concentration for cells expressing 6-3, 8-1, 8-7, 8-13, and 8-35 and partially so for 8-12 and 8-19 expressing cells. These data are consistent with the hypothesis that the aptamers are inhibiting growth by sequestering intracellular heme, and this effect can be completely or partially overcome by increasing the bioavailable heme concentration. Next, growth curves for RP523 expressing the selected aptamers 8-1, 8-7, 8-12, 8-13, and 8-35 and control oligonucleotide 6-5 were conducted at limiting (1 ␮M) and saturating (10 ␮M) heme. At 1 ␮M heme, bacteria expressing aptamers grew more slowly and reached final optical densities ⬃50% less than those of the control culture (Figure 4, panel b). At 10 ␮M heme, the growth curves for RP523 expressing 8-1, 8-7, 8-13, and 8-35 and the control oligonucleotide 6-5 were identical. Similar to the screening studies, this defect is not completely reversed for RP523 expressing 8-12, even though the growth rate and final OD600 increased with heme concentration. Overall, these data are consistent with the screening results and indicate that intracellular heme sequestration by aptamers slows bacterial growth and limits the total number of cell doublings. At nonlimiting heme concentrations, the heme-sequestration capacity of the aptamers is overcome, and sufficient heme is available to support growth comparable to that of control cultures. VOL.1 NO.8 • 515–524 • 2006

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heme concentrations. Between 4 and 10 ␮M heme, ␦-ALA levels among 6-5, 8-13, and 8-35 expressing cells are comparable, indicating that excess heme overcomes the sequestration effect of these aptamers. Interestingly, ␦-ALA levels in 8-12 relative to 6-5, 8-13, and 8-35 expressing cells remain elevated, even at nonlimiting heme concentrations. Altogether, there is a strong concordance between the heme dependence of growth and ␦-ALA levels in aptamer-expressing cells. Relative to control cells, a growth defect is observed at low heme concentrations, ␦-ALA levels are correspondingly high, and both parameters are normalized at high media heme concentrations. AptamFigure 4. Impact of intracellularly expressed aptamers on RP523 growth and ␦-ALA production as ers are not all equally bioactive. Intracellular a function of heme concentration. a) Final cell density measurements obtained for small-scale aptamer efficacy is a function of at least cultures containing initial heme concentrations of 0.5, 1, 2, and 4 ␮M. b) Representative growth three variables, namely, (i) target affinity curves for selected aptamers at limiting (1 ␮M; top panel) and nonlimiting (10 ␮M; bottom (Kd), (ii) intracellular concentration, and (iii) panel) heme. c) ␦-ALA concentration normalized to OD600nm for cells expressing control oligonucleotide 6-5 and aptamers 8-12, 8-13, and 8-35. attaining the properly folded structure required for productive target binding. As shown in Table 1, the aptamers all bind heme with Determination of ␦-ALA Levels in RP523 Expressing Aptamers. Heme auxotrophy in E. coli RP523 is due similar affinity, so this variable cannot explain the differto a mutation(s) in the hemB locus leading to absent ent growth phenotypes. Using quantitative RT-PCR, we ␦-aminolevulinic acid dehydratase (ALAD) activity (42). determined the intracellular levels of aptamers 8-1, 8-7, This enzyme is responsible for converting ␦-ALA into por- 8-12, 8-13, and 8-35 relative to control oligonucleotide phobilinogen in the heme biosynthetic pathway 6-5 in cells grown at 2 ␮M heme. The aptamers and (Figure 1). Since the hemA and hemL loci are intact in control oligonucleotide are present at similar levels RP523, this strain synthesizes ␦-ALA, and under heme- (Figure 5), indicating that dramatic differences in the limiting conditions, HemA activity increases, leading to intracellular concentration of the various aptamers do ␦-ALA accumulation. Conversely, when heme is nonlim- not account for their differential in vivo efficacy. In vivo iting, HemA activity is repressed, leading to decreased folding efficiency, therefore, is most likely the factor ␦-ALA levels. Therefore, if aptamers are limiting intracellular heme availability, ␦-ALA levels should be higher relative to those in RP523 expressing control oligonucleotide 6-5. To test this, OD600-corrected ␦-ALA levels in RP523 grown at 2–10 ␮M heme and expressing control oligonucleotide 6-5 and the aptamers 8-12, 8-13, and 8-35 were measured using LC/electrospray ionization (ESI)-MS in positive ions selected ion monitoring (SIM) mode, and these data are summarized (Figure 4, panel c). At 2 ␮M heme, ␦-ALA levels in aptamer expressing cells were ⬃1.5–4-fold higher than those in cells expressing 6-5, suggesting that at this heme con- Figure 5. Relative aptamer expression levels as centration, expressed aptamers are limiting intracellular determined by quantitative RT-PCR. 520

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ARTICLE responsible for the differential efficacy of aptamers at sequestering intracellular heme, with the aptamers demonstrating strong heme-dependent phenotypes reflecting those efficiently attaining folded structures competent for heme binding. Misfolding of in vitro selected aptamers when expressed intracellularly is a recognized challenge; however, sufficiently diverse libraries can be screened to identify those aptamers possessing the desired intracellular activity and hence the appropriately folded structure in vivo. Additionally, aptamers 8-12 and 8-19 are interesting in that the growth defect persists even at higher media heme concentrations. Similarly, in 8-12-expressing cells, ␦-ALA levels remain elevated. This may reflect the possibility that these aptamers fold intracellularly with the highest efficiency, and transport of heme into the cell becomes limiting prior to the aptamer heme-sequestration capacity being surpassed. While we cannot completely exclude additional “off-target” effects of these aptamers, there is a clearly demonstrated “on-target” effect as cells expressing 8-12 accumulate excess ␦-ALA. This suggests that intracellular heme limitation remains a reasonable explanation for the persistent growth defect observed in these aptamer-expressing cells, so that even at high heme concentrations in the culture media, the sensed intracellular heme levels may remain low due to sequestration. Overall, the ␦-ALA data are significant for two reasons: First, they validate the hypothesis that the tested aptamers are specifically targeting heme in vivo and, therefore, are acting “on-target”. Second, the data indicate that the aptamers can sequester intracellular heme and perturb the heme-mediated inhibition of the

heme biosynthetic pathway predictably and in an objectively measurable way, thus confirming their utility as tools for probing this heme-dependent cellular circuit. In this study, we have used in vitro SELEX to evolve and test the first reported expressible heme-binding RNA library in a cellular system. Selected aptamers, when expressed in an E. coli heme auxotroph, induce a growth defect and ␦-ALA accumulation at low heme concentrations, and both phenotypes are reversed upon increasing heme concentration in the growth media. Together, these data indicate that aptamers are selectively binding their cognate ligand in vivo. Furthermore, by restricting the bioavailable intracellular heme pool, with respect to both inhibiting cell growth and modulating heme-dependent repression of the heme biosynthetic pathway, these aptamers impact an important cellular regulatory circuit predictably, thereby validating this approach in efforts to elucidate other hemeregulated networks and possibly other small molecule regulated cellular processes. This strategy is inherently appealing because it can be used to explore a wide range of biologically important compounds, since the versatility of SELEX will enable discovery of suitable aptamers. While we have used an E. coli heme auxotroph to facilitate screening based on a growth phenotype, this is not an absolute requirement. Using aptamers to perturb intracellular pools of physiologically important small molecules in conjunction with global screening methods such as transcriptional profiling microarrays, proteomics, and metabolomics may augment and extend findings based on gene knockout and other available approaches.

METHODS

In Vitro RNA Transcription Reactions. Oligonucleotides were obtained from Integrated DNA Technologies, Inc. The randomized single-stranded DNA library with sequence GCC GGA TCC GGG CCT CAT GTC GAA [N]50T TGA GCG TTT ATT CTG AGC TCC C was synthesized on a 1 ␮mol scale and polyacrylamide gel electrophoresispurified prior to use. A 5=-HindIII primer (CCG AAG CTT AAT ACG ACT CAC TAT AGG GAG CTC AGA ATA AAC GCT CAA) and 3=-BamH1 primer (GCC GGA TCC GGG CCT CAT GTC GAA) were used for PCR amplification of the starting library and RT-PCR during library evolution. Prior to the first selection round, a single-stranded DNA library (10 ⫻ 100 ␮L reactions, 10 pmol DNA per reaction) in 10 mM TrisHCl, 50 mM KCl, 7.5 mM MgCl2, pH 8.3, 1 mM each dNTP, 2 ␮M each 5=- and 3=-primers, and Taq DNA polymerase (2.5 U) was subjected to six cycles of PCR (pre-PCR, 94 °C ⫻ 4 min, 57 °C ⫻ 5 min; PCR, 94 °C ⫻ 30 s, 57 °C ⫻ 60 s, 72 °C ⫻ 60 s, and a 7 min final extension). The double-stranded product was purified and extracted

Preparation of Selection and Subtraction Columns. MPIX (Frontier Sciences) was immobilized on oxirane-activated acrylic beads (Sigma-Aldrich) as previously described (34). Briefly, 3.1 mg (4.4 ␮mol) of MPIX was dissolved in 1 mL of DMSO, then diluted to 10 mL with 100 mM KH2PO4, pH 9.5, and added to 1.07 g of activated acrylic beads. This slurry was gently rotated for 2 d in the dark; then the beads were washed extensively with (i) 10 mM KH2PO4, pH 5, (ii) H2O, (iii) 0.1 M NaOH, (iv) H2O, (v) 100 mM KH2PO4, pH 8.0, and finally, (vi) 100 mM KH2PO4, pH 8, with 5% ␤-mercaptoethanol. The beads were gently rotated in this buffer in the dark for 2 d, before being washed extensively with H2O. Subtraction beads were prepared by treating 1 g of activated beads with 100 mM KH2PO4, pH 8, and 5% ␤-mercaptoethanol as above. Both selection and subtraction beads were stored in 10 mM KH2PO4, pH 7.2, 150 mM NaCl, 0.02% sodium azide buffer at 4 °C in the dark.

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from a 4% agarose gel, and 150 pmol was used as template for in vitro transcription using the Ampliscribe T7 Flash Kit (Epicentre Biotechnologies) spiked with 1 ␮L of ␣-33P-ATP (3000 Ci mmol–1, 10 ␮Ci ␮L–1, Perkin Elmer). Reaction times ranged from 4 h to overnight, at the end of which DNase I (1 U) was added at 37 °C for 30–60 min to digest the template DNA. RNA was purified by phenol–chloroform extraction followed by ethanol precipitation at –20 °C. Aptamer Selection. RNA (200–1000 pmol) in diethyl pyrocarbonate (DEPC)-treated water was denatured by heating to 70 °C for 5 min, allowed to cool to RT, and refolded in selection buffer (SB), with composition 100 mM Tris–acetate, 200 mM sodium acetate, 25 mM potassium acetate, 10 mM magnesium acetate, 0.05% Triton X-100, and 5% DMSO. For subtraction (rounds 1–3 and 8), RNA in ⬃200 ␮L of SB was added to ⬃100 ␮L of subtraction resin pre-equilibrated in SB and incubated at ambient temperature with gentle mixing for 30 min. The supernatant and 3 ⫻ 50 ␮L of SB washes of the subtraction resin were recovered and incubated with selection resin (200 ␮L) for 1 h at ambient temperature with gentle agitation. The selection resin was washed with 11 column volumes of SB and eluted with 2.5 mM hemin (Sigma-Aldrich) in SB (6 ⫻ 200 ␮L aliquots with 10 min between additions). The eluted RNA in each fraction was ethanol precipitated overnight at –20 °C with 20 ␮g of glycogen as a carrier, and amplified using 8 ⫻ 50 ␮L Ready-To-Go RT-PCR tubes (Amersham) and ⬃300 pmol each 5=- and 3=-primers. RT was carried out at 42 °C for 40 min, and the reverse transcriptase was inactivated at 95 °C ⫻ 5 min, followed by 18 PCR cycles (94 °C ⫻ 30 s, 57 °C ⫻ 60 s, 72 °C ⫻ 60 s) and a 7 min final extension. The RT-PCR products were pooled, concentrated, and purified using 4% agarose gel, and the desired length product was extracted and ethanol precipitated. DNA was resuspended in DEPC-treated H2O for the next round of in vitro transcription. At the sixth and eighth selection rounds, the evolved library was blunt end cloned into pSTBlue-1 vector (Invitrogen) and used to transform competent NovaBlue E. coli cells (Invitrogen). Single colonies were used for mini-prep cultures from which plasmid encoding a single aptamer was isolated for sequencing and archiving. Plasmids were sequenced at either Elim Biopharmaceuticals or the UC Berkeley Sequencing Facility using the SP6 primer. Determining Aptamer Heme-Binding Properties. Binding of rounds two and seven libraries to the protoporphyrin IX scaffold was qualitatively determined by fluorescence spectroscopy using a FluoroMax-2. Refolded library RNA (⬃2 ␮M) and PPIX in the more physiologic SHMCK buffer (20 mM N-2-hydroxyethylpiperazine-N=2-ethanesulfonic acid (Hepes), 120 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1 mM CaCl2, pH 7) were incubated at ambient temperature, and the fluorescence emission spectrum was measured after excitation at 400 nm. Binding of selected aptamers from the round 8 library to heme was determined by UV–vis spectroscopic difference titrations using a Cary 300 Bio spectrophotometer equipped with a dual cell Peltier accessory (Varian). Briefly, reference and sample cuvettes containing SHMCK only and refolded RNA (⬃1–2 ␮M) in SHMCK were prepared, and heme was added in 0.1–0.2 ␮M aliquots to each sample and reference cuvette pair at 25 °C while stirring continuously. Difference spectra were recorded every 5 min, and ⌬A405nm was plotted against log[heme] and fitted to eq 1 to determine aptamer apparent heme-binding dissociation constants. ⌬A405nm ⫽ m1 ⫹ (m2 – m1)/{1 ⫹ 10(–m3(m0–log(m4)))} (1) The variables are as follows: m0 ⫽ log[heme]; m1 ⫽ minimum ⌬A405nm; m2 ⫽ maximum ⌬A405nm; m3 ⫽ Hill coefficient; m4 ⫽ apparent Kd. Cloning of Aptamers into the RNA Expression Vector pGFIB. Plasmid pGFIB was obtained as a gift from Prof. William McClain (University of Wisconsin, Madison). With primers CCG GAA TTC AAT ACG ACT CAC TAT AGG GAG CTC AGA ATA AAC GCT CAA (5=-EcoRI) and GCC CTG CAG GGG CCT CAT GTC GAA (3=-PstI), full-length aptamers were PCR amplified from the archival plasmid, purified by

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4% agarose gel, extracted, and resuspended in ddH2O after ethanol precipitation. Aptamers (0.36–0.50 ␮g of DNA) were doubledigested with EcoR1 (20 U) and PstI (20 U) for 4 h at 37 °C in 1⫻ EcoR1 Unique Buffer (New England Biolabs), then ligated into pGFIB (⬃4.5 ␮g digested with 20 U of EcoR1 and 20 U of PstI for 5 h at 37 °C, then treated with 0.6 U of calf intestinal phosphatase at 37 °C for an additional 30 min) using the Rapid Ligation Kit (Roche). DH-5␣ E. coli cells were transformed using aliquots of the ligation reactions and grown overnight on Luria-Bertani plates supplemented with 50 ␮g mL–1 carbenicillin. Single colonies were selected for miniculture and plasmid isolation, and aptamer insertion was verified by sequencing using the M13 Forward primer. Screening Heme-Binding RNA Aptamers for in Vivo Function. pGFIB/aptamer constructs were used to transform the E. coli heme auxotroph RP523 obtained from the E. coli Genetic Stock Center (http://cgsc.biology.yale.edu/top.html). Cells were grown overnight on LB plates supplemented with 50 ␮g mL⫺1 carbenecillin and 15 ␮M hemin at 37 °C and stored in the dark at 4 °C. All growth experiments were done in LB containing 50 ␮g mL⫺1 carbenecillin and supplemented with the appropriate heme concentration as indicated. For high-throughput screening, overnight cultures (1 mL in 15 mL tubes) containing 0.5, 1, 2, and 4 ␮M hemin were inoculated with RP523 harboring specific pGFIB/aptamer constructs from starter cultures in the early to mid-log phase of growth and incubated at 37 °C and 250 rpm. Optical density measurements (OD600) at ⬃14–20 h were used to assess the extent of bacterial growth. For growth kinetics studies, 50 mL cultures in 250 mL Erlenmeyer flasks inoculated with mid-log phase starter cultures (⬃0.5 mL) and supplemented with 1 and 10 ␮M heme were grown at 37 °C and 250 rpm. OD600 readings were taken every 30–60 min to assess growth. Measurement of ␦-ALA Levels in RP523 E. coli. Cells were grown in 5 mL of LB containing between 2 and 10 ␮M heme and harvested by centrifugation after measurement of the OD600. Generally, cells expressing control oligonucleotide 6-5 and aptamers were harvested at similar OD600 values, dictated by the maximum OD600 attained by aptamer-expressing cells. This ensured that the degree of media heme depletion was similar between control and experimental cultures. Cell pellets were resuspended in 200 ␮L of 4% heptafluorobutyric acid (HFBA) and lysed by three freeze–thaw cycles followed by sonication for 5 min. Supernatants containing ␦-ALA were recovered after centrifugation at 14,000 rpm for 10 min and quantitated using an Agilent 1100 series liquid chromatograph/mass selective detector (LC/MSD) operated in positive ESI mode. A standard curve was constructed using authentic ␦-ALA (Sigma-Aldrich) detected by monitoring the m/z ⫽ 114 [M – H2O ⫹ H⫹]⫹ and 132 [M ⫹ H⫹]⫹ ions in SIMS mode. For the LC, a 150 mm ⫻ 3.9 mm, 5 ␮m Nova-Pak C18 column (Waters) with 25 mM HFBA, 5 mM ammonium acetate (solvent A) and 90:10 methanol/5 mM ammonium acetate (solvent B) as mobile phases were used. The column was eluted at a flow rate of 0.4 mL min⫺1, according to the following gradient: 5% B for 3 min; 5–25% B over 12 min; 25–100% B over 2 min; an isocratic phase at 100% B for 7 min; 100–5% B over 1 min. For the MSD, the drying gas flow rate and temperature were 12 L min⫺1 and 350 °C, respectively, and the nebulizer gas pressure was 35 psig. The capillary exit and fragmentor voltages were –3000 and 70 V, respectively. Quantitative RT-PCR. RP523 bacteria expressing 6-5, 8-1, 8-7, 8-12, 8-13, and 8-35 were grown in LB containing 2 ␮M heme. Cells were harvested at OD600 ⫽ 0.3 by adding ice-cold 95:5 ethanol/ water-saturated phenol to 11% v/v and centrifuging for 2 min at 4 °C. Supernatants were aspirated, and cell pellets were frozen in liquid nitrogen, then stored at –80 °C until needed. Total RNA was isolated using TRIzol (Invitrogen) according to the supplier’s protocol. Approximately 2 ␮g of RNA was digested with 5 U of RNasefree DNase I (Fermentas) in 10 mM Tris-HCl, pH 7.5, 2.5 mM MgCl2, 0.1 mM CaCl2 at 37 °C for 1.5 h. DNase I was inactivated at 75 °C for

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ARTICLE 15 min after adding EDTA to 1 mM. cDNA syntheses with ⬃200 ng of DNase I-treated RNA were performed using the Thermoscript RT-PCR System (Invitrogen). Target-specific primers for aptamers (GCC GGA TCC GGG CCT CAT GTC GAA) and 16S rRNA (GGT TAC CTT GTT ACG ACT T) as an internal reference were used for cDNA synthesis. Q-PCR reactions were carried out in triplicate per RNA dilution in 50 ␮L reactions containing 10 mM Tris-HCl, pH 8.5, 50 mM KCl, 1.5 mM MgCl2, 200 ␮M dNTPs, 0.1⫻ SYBR Green I (Molecular Probes), 1.5 U of Taq (Fermentas), and 200 nM primers. The primers for amplifying the aptamer library were also used for aptamer quantitation by Q-PCR. For 16S rRNA, primers 16S_462F (GTT AAT ACC TTT GCT CAT TGA) and 16S_801R (ACC AGG GTA TCT AAT CCT GTT) were used. The temperature program used for both cDNA targets was 95 °C ⫻ 10 min and 40 cycles of 95 °C for 30 s, 57 °C ⫻ 1 min, 72 °C ⫻ 1 min, followed by a melting curve. Buffer and no RT controls for each RNA sample were included to verify the absence of contaminating DNA in the DNase I-treated RNA samples. Expression levels reported are relative to an aptamer/16S rRNA ratio of 1 assigned to sample 6-5. Acknowledgments: We thank Prof. William H. McClain (University of Wisconsin, Madison) for providing plasmid pGFIB and Dr. Jasper Rine’s lab (University of California, Berkeley) for use of their Q-PCR instrument. This work is supported by NIH Grant 5 F32 AI058646 (J.C.N) and the Aldo DeBenedictis Fund (M.A.M). Note added after print publication: Because of a production error, the following references were misformatted: 1– 42. These errors do not affect the scientific integrity of the article. The electronic version was corrected and reposted to the web on October 20, 2006. This paper was originally posted September 8, 2006, and the electronic version was corrected and reposted to the web on October 20, 2006. An Addition and Correction may be found in ACS Chem. Biol. 1(9).

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