- Email: [email protected]
Assays for Investigating Transcription by RNA Polymerase IIin Vitro
METHODS: A Companion to Methods in Enzymology 12, 192–202 (1997) Article No. ME970471
Assays for Investigating Transcription by RNA Polymerase II in Vitro Daniel Reines,* Arik Dvir,† Joan Weliky Conaway,†,‡ and Ronald C. Conaway†,1 *Department of Biochemistry, Emory University School of Medicine, Atlanta, Georgia 30322; †Program in Molecular and Cell Biology, Oklahoma Medical Research Foundation, 825 N.E. 13th Street, Oklahoma City, Oklahoma 73104; and ‡Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, Oklahoma 73190
With the availability of the general initiation factors (TFIIB, TFIID, TFIIE, TFIIF, and TFIIH), it is now possible to investigate aspects of the mechanism of eukaryotic messenger RNA synthesis in purified, reconstituted RNA polymerase II transcription systems. Rapid progress in these investigations has been spurred by use of a growing number of assays that are proving valuable not only for dissecting the molecular mechanisms of transcription initiation and elongation by RNA polymerase II, but also for identifying and purifying novel transcription factors that regulate polymerase activity. Here we describe a variety of these assays and discuss their utility in the analysis of transcription by RNA polymerase II. q 1997 Academic Press
Messenger RNA synthesis is a major site for the regulation of gene expression. Eukaryotic messenger RNA synthesis is a complex biochemical process catalyzed by multisubunit RNA polymerase II and governed by the concerted action of a diverse collection of transcription factors (TF) that control polymerase activity at both the initiation and the elongation stages of transcription (1, 2). Over the past 10 years, substantial progress defining the molecular mecha1 To whom correspondence should be addressed. Fax: (405) 2711580.
nisms underlying the initiation and elongation stages of eukaryotic messenger RNA synthesis has been achieved. This progress is due in large part to two major advances: (i) the development of purified, reconstituted RNA polymerase II transcription systems amenable to detailed mechanistic analysis and (ii) the creation of novel assays for dissecting the mechanism of transcription in these systems. With the resolution and purification of the general initiation factors (TFIIB, TFIID, TFIIE, TFIIF, and TFIIH), basal transcription by RNA polymerase II can now be reconstituted with purified proteins from a large number of structurally diverse promoters. This ‘‘minimal’’ transcription system is proving valuable not only as a tool for investigating the mechanism of transcription by RNA polymerase II, but also as an assay system for identifying and purifying novel transcription factors that regulate polymerase activity. The development of well-defined transcription systems has been accompanied by a proliferation of powerful assays for investigating transcription. A number of these assays exploit specialized DNA templates. In this article, we describe the most commonly used variations of these assays and discuss their applications. Because basic procedures for purifying RNA polymerase II and the general initiation factors and for reconstituting optimized basal transcription have recently been thoroughly reviewed in Methods in Enzymology (Vols. 273 and 274), we will not discuss them here.
192
AID
1046-2023/97 $25.00 Copyright q 1997 by Academic Press All rights of reproduction in any form reserved.
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
DESCRIPTION OF THE METHOD General Assays for Measuring Promoter-Dependent Transcription by RNA Polymerase II RNA polymerase II will initiate transcription accurately in vitro from a large number of structurally diverse promoters. A variety of assays for detecting and quantitating initiation by RNA polymerase II from promoters are currently in use. These include the runoff transcription (3), G-less cassette (4), S1 nuclease, RNase protection, and primer extension assays (5, 6) (Fig. 1). Of these assays, the S1 nuclease, RNase protection, and primer extension assays are superior for confirming that RNA polymerase II is initiating transcription in vitro at the correct transcriptional start sites, but they are laborious and time-consuming to perform. The runoff transcription and G-less cassette assays are simpler to perform and are therefore preferred for most measurements of promoter-dependent transcription by RNA polymerase II. Although the runoff transcription assay is the method of choice for most purposes, it has several disadvantages that preclude its use in select circumstances. First, the runoff transcription assay cannot be used to investigate the role of DNA topology in transcription by RNA polymerase II, because it re-
FIG. 1. Commonly used assays of promoter-dependent transcription by RNA polymerase II. The asterisk in the bottom panel designates a radioactive label at the 5*-end of an oligonucleotide used for primer extension analysis. TATA, TATA box; Inr, initiator element.
AID
Methods A 0471
/
6714$$$161
193
quires a linear DNA template; for such studies, the G-less cassette assay is ideal because closed circular DNAs can be used as templates for transcription. Second, the runoff transcription assay is often not well suited for investigating transcription in crude enzyme systems, because high levels of nonspecific background transcription initiated at ends, nicks, or cryptic promoters in some DNA templates can obscure the correctly initiated transcript; in these instances, the G-less cassette assay is preferred because background transcription can be reduced by including only ATP, CTP, and UTP as substrates for elongation by RNA polymerase II and by treating reaction products with ribonuclease T1 , which cleaves RNA after G-residues. Assays for Investigating Specific Steps in PromoterDependent Transcription by RNA Polymerase II Although the general assays described above are useful for quantitating promoter-dependent transcription by RNA polymerase II, they are not sufficient for dissecting its mechanism. Promoter-dependent transcription by RNA polymerase II is a complex, multistep process requiring (i) assembly of polymerase and its initiation factors into a functional preinitiation complex, (ii) ATP-dependent unwinding of promoter DNA surrounding the transcriptional start site to form an ‘‘open complex,’’ (iii) transcription initiation (i.e., synthesis of the first phosphodiester bond of nascent transcripts), (iv) promoter escape (i.e., conversion of the unstable initiation complex into a stable elongation complex), and (v) elongation of full-length transcripts (1, 2). Below we describe a variety of assays that have been used successfully to elucidate many aspects of the mechanism of promoter-dependent transcription by RNA polymerase II and the roles of individual transcription factors in this process.
Sarkosyl and Heparin Challenge Assays In studies investigating the mechanism of transcription initiation by RNA polymerase II, it is often important to carry out transcription reactions under conditions in which (i) initiation is limited to a single round per promoter and (ii) events occurring during initiation and synthesis of the first few phosphodiester bonds of nascent transcripts can be distinguished from events occurring during subsequent steps in transcription. This is readily accomplished using Sarkosyl (N-lauroylsarcosine) or heparin challenge
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
194
assays. Elongation of 4- to 10-nucleotide-long transcripts by RNA polymerase II is resistant to inhibition by 0.2–0.5% Sarkosyl (7–9) and by 10–100 mg/ ml heparin (10, 11), whereas assembly of the preintiation complex and synthesis of the first 2–3 phosphodiester bonds of nascent transcripts are not. Thus, if Sarkosyl or heparin is added to transcription reactions shortly after addition of ribonucleoside triphosphates, full-length runoff transcripts will be synthesized only from those promoters at which initiation has already occurred. A convenient way to limit initiation to a single round per preinitiation complex is to initiate transcription with only the subset of ribonucleoside triphosphates needed for synthesis of the first 4–10 phosphodiester bonds of transcripts; following addition of Sarkosyl or heparin, full-length transcripts are synthesized by addition of the remaining ribonucleoside triphosphates.
The Template Competition Assay This assay is a modification of the template competition assay originally described by Chamberlin and co-workers (12). The template competition assay is well suited for investigating the requirements for interaction of RNA polymerase II and its transcription factors with promoters, and it has proven quite valuable in biochemical studies dissecting the mechanism of assembly of the preinitiation complex (11, 13, 14). Although the template competition assay is more difficult to perform than some other assays for identification and analysis of RNA polymerase II preinitiation intermediates, it offers several advantages. Unlike the gel-shift assay, for example, which subjects putative preinitiation intermediates to the nonoptimal transcription reaction conditions present in native polyacrylamide gels, the template competition assay is carried out entirely under optimal transcription reaction conditions. In addition, unlike the gel-shift assay, which does not permit easy assay of the transcriptional activities of putative preinitiation intermediates identified as bands in native polyacrylamide gels, the template competition assay permits identification of ‘‘functional’’ preinitiation intermediates by direct measurement of their transcriptional activities. Template challenge assays are performed as diagrammed in Fig. 2. Two different plasmids, each containing the same promoter, are used as DNA templates. The plasmids are linearized by restriction enzyme digestion, so that they direct synthesis of
AID
Methods A 0471
/
6714$$$161
runoff transcripts of different lengths. During an initial preincubation, templates I and II are incubated in separate reaction mixtures with various combinations of RNA polymerase II and transcription factors; at this stage, it is important that the transcription factor of interest is present at a subsaturating concentration. The two reaction mixtures are combined to begin a second preincubation and, where necessary, the remaining transcription proteins required for assembly of the complete preinitiation complex are added to reaction mixtures. Following the second preincubation, which must be long enough to allow binding reactions to come to equilibrium, transcription is carried out using the Sarkosyl or heparin challenge assay to limit initiations to a single round per preinitiation complex; if initations are not limited to a single round, misleading results can be obtained because transcription proteins released from the first template following initiation may be able to assemble into preinitiation complexes and promote initiation on the second template. The relative molar amounts of full-length runoff transcripts synthesized from each template are determined and expressed as the molar ratio (I/II) of transcription from templates I and II. If the transcription factor of interest is capable of interacting stably and stoichiometrically with preinitiation intermediates on template I in the presence of the set of transcription proteins provided during the first preincubation, the molar ratio of transcripts synthesized on the two templates will be greater than that of a control reaction in which the two templates are mixed at the beginning of the first preincubation. On the other hand, if the transcription factor interacts catalytically, or not at all, with preinitiation intermediates on template I during the first preincubation, it should distribute equally on both templates during the second preincubation, and the molar ratio of transcripts synthesized from the two templates should be the same as that of the control reaction.
Assays That Bypass Requirements for Subsets of Initiation Factors and ATP-Dependent Open Complex Formation Negatively supercoiled and ‘‘bubble’’ templates have proven useful for investigating the roles of individual transcription factors in promoter-dependent transcription initiation by RNA polymerase II. Negatively supercoiled templates bypass the requirement for TFIIH and ATP in open complex formation,
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
most likely because the stored free energy of supercoiling facilitates unwinding of promoter DNA around the transcriptional start site. The utility of negatively supercoiled DNA templates first became apparent with the observation that initiation by RNA polymerase II at a negatively supercoiled human IgH promoter requires only TFIIB and TBP or TFIID and is independent of ATP, whereas initiation by polymerase at the same relaxed IgH promoter requires the full complement of general initiation factors and depends on ATP (15, 16). Subsequent investigation revealed that TFIIH and ATP are dispensable for initiation at promoters on all negatively supercoiled templates tested; however, the requirement for other initiation factors varies depending on the promoter sequence (16–18). For example, while transcription initiation by RNA polymerase II at the negatively supercoiled IgH promoter requires only TFIIB and TFIID or TBP, initiation at the negatively supercoiled adenovirus E4 promoter requires TFIIB, TFIID or TBP, and TFIIF, and initiation at the supercoiled AdML promoter requires TFIIB, TFIID or TBP, and TFIIF, but is strongly stimulated by TFIIE. Thus, by choosing supercoiled templates containing the appropriate promoters, it is possible to devise assays that can be used for studying the
195
mechanisms of action of different subsets of initiation factors. Because open complex formation on negatively supercoiled templates is driven by the energy of supercoiling, it is not possible to control precisely the position of the region of unwound DNA. In addition, negatively supercoiled templates have the significant disadvantage that they must remain supercoiled in the presence of RNA polymerase II and transcription factors and therefore must be used in transcription systems devoid of nucleases and topoisomerase I. Bubble templates, which contain premelted promoters with short stretches of noncomplementary bases surrounding the transcriptional start site, bypass the requirement for TFIIH and ATP in initiation and provide an alternative to negatively supercoiled templates. Because bubble templates are prepared synthetically, by hybridization of either synthetic oligonucleotides (19) or larger singlestranded DNA fragments (20), the position of the premelted region of DNA can be precisely controlled. Furthermore, because bubble templates do not need to be supercoiled, they are much less sensitive to inactivation by nucleases and therefore can be used in relatively crude transcription systems. Detailed
FIG. 2. The template competition assay. The bottom panel is a mock autoradiogram of a denaturing polyacrylamide gel analyzing the transcripts synthesized in the template competition assay outlined in the top panel.
AID
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
196
procedures for preparing supercoiled and premelted DNA templates can be found in Refs. (15, 19–21).
Abortive Initiation Assay The abortive initiation assay has been widely used in investigations of the requirements for synthesis of the first phosphodiester bond of nascent transcripts by both prokaryotic and eukaryotic RNA polymerases (18, 22–24). The abortive initiation assay exploits the ability of RNA polymerase to utilize dinucleotides to prime synthesis of transcripts at its promoters. If only a dinucleotide and the next ribonucleoside triphosphate encoded by the DNA template are provided as substrates for transcription, RNA polymerase II will reiteratively synthesize trinucleotide transcripts that are rapidly released from the polymerase active site. Dinucleotideprimed transcription initiation by RNA polymerase II can occur over an approximately nine basepair region surrounding transcriptional start site (25). Thus, at the AdML promoter, for example, the dinucleotides CpA and CpU support trinucleotide synthesis in the presence of [a-32P]CTP; CpC, ApC, and UpC support trinucleotide synthesis in the presence of [a-32P]UTP; and UpC supports trinucleotide synthesis in the presence of [a-32P]ATP (Fig. 3). Dinucleotide-primed trinucleotide transcripts are easily separated from unincorporated [a-32P]NTPs by electrophoresis through 28% acrylamide/3% bisacrylamide denaturing gels (23). Assays for Investigating the Mechanism of Elongation by RNA Polymerase II A variety of assays that exploit the intrinsic ability of RNA polymerase II to initiate transcription in the
absence of accessory factors at single-stranded regions of DNA or at nicks and ends of double-stranded DNA have been developed for monitoring polymerase activity during purification, for probing aspects of polymerase catalytic activity, and for investigating the mechanisms of action of elongation factors. Transcripts are internally labeled using [a-32P]NTPs and can be conveniently analyzed by denaturing polyacrylamide gel electrophoresis, followed by autoradiography or PhosphorImager analysis, or by counting radioactivity of samples directly after removal of unincorporated [a-32P]NTPs from labeled RNA products by trichloroacetic acid precipitation or DE-81 adsorption (5). The simplest assays for promoter-independent transcription by RNA polymerase II utilize sheared, single-stranded calf thymus or salmon sperm DNA as template (26). In addition, we describe below a series of specialized DNA templates that have been developed for assay of promoter-independent transcription by RNA polymerase II (Fig. 4).
Tailed, Dumbbell, and Bubble Templates DNA duplexes which are extended at one 3*-end are excellent templates for a number of RNA polymerases including RNA polymerase II (Fig. 4). Extended ‘‘tailed’’ duplexes can be synthesized enzymatically using terminal transferase and dCTP (the favored base) (27). To restrict transcription to initiation from only a single end of the DNA, one end is usually removed by digestion with a restriction enzyme that yields a short DNA duplex containing one tailed 3*-terminus. If small RNA transcripts are
FIG. 3. Requirements for dinucleotide-primed abortive initiation at the AdML promoter. A*, U*, and C* refer to [a-32P]ATP, [a-32P]UTP, and [a-32P]/CTP, respectively.
AID
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
to be analyzed, this small transcriptionally active DNA fragment can be removed by standard DNA purification techniques. Tailed templates can also be formed by ligating an oligonucleotide onto a linear DNA duplex containing a 5*-overhanging ‘‘sticky’’ end (28). RNA polymerase II binds at the tail–duplex junction, where it initiates an RNA chain complementary to a few bases of the single strand adjacent to the duplex (29). The site of transcription initiation can be fixed more precisely by initiating with a dinucleotide primer complementary to specific template bases on the single-stranded tail (30). An advantage of tailed templates is that they are used very efficiently by RNA polymerase II such that most, if not all, of the DNA duplexes are occupied by transcriptionally engaged polymerase. This is in contrast to the relatively low template occupancy in most promoter-dependent reactions reconstituted with general transcription factors. An unusual feature of tailed templates, however, is the propensity for the nontemplate strand to be displaced during RNA synthesis such that the transcript remains base-paired with the template strand and a long RNA:DNA hybrid is produced. This aspect of transcription on tailed templates has recently been described in detail (26). In addition, because transcription initiates at the end of the template, full-length transcripts are nearly the same length as the template DNA. This can lead to confusion regarding the identities of reaction products since RNA polymerase II has the ability to label the nontemplate strand of DNA by adding a radioactive ribonucleotide to the 3*-end (31). Interestingly, this activity of RNA polymerase II is strongly stimulated by the transcription factor elongin (SIII) (32). The ‘‘dumbbell’’ template, on which RNA polymer-
ase II can extend a ribonucleic acid primer in a template-dependent manner (33), is the basis of another useful assay (Fig. 4). The dumbbell template is a chemically synthesized oligonucleotide that can base-pair extensively across its length, except for a short single-stranded gap; the few 3*-terminal bases are ribonucleotides added to a deoxyribonucleotide chain. The dumbbell structure is thought to mimic a transcription bubble with RNA bases hydrogenbonded to a DNA template strand. RNA polymerase II covalently attaches ribonucleotides to this single chain, which can be extended by copying the DNA template strand until its 5*-end is reached. The size of the oligonucleotide that can be obtained through chemical synthesis will necessarily limit the length of transcripts extended; thus, dumbbell templates cannot be used to study elongation of lengthy transcripts. Analysis of Escherichia coli RNA polymerase has been facilitated by the reconstitution of a synthetic transcription bubble using double-stranded DNA containing an internal segment that is mismatched and cannot base-pair (34). In a more complete reconstruction of a transcription bubble, an RNA chain annealed to the template strand of the bubble can be used as a primer and extended by E. coli RNA polymerase (35). Analysis of elongation by RNA polymerase II would benefit from this approach, but such experiments have not yet been reported. Earlier reports suggested that certain DNA sequences on supercoiled plasmids, which presumably yield transiently melted regions, allow initiation by purified RNA polymerase II albeit inefficiently (36–39). As described above, synthetic bubble-containing templates have also been useful for studies of promoterdependent transcription by RNA polymerase II.
FIG. 4. Specialized DNA templates for investigating elongation by RNA polymerase II.
AID
Methods A 0471
/
6714$$$161
197
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
198
Isolation and Analysis of RNA Polymerase II Elongation Complexes Utility The isolation of elongation complexes has proven to be extremely useful in examining individual steps in the transcription elongation process. A number of procedures are now commonly used to separate complexes from proteins and small solutes. This makes possible substantial control over reaction conditions, including the removal and provision of specific protein factors and specific subsets of the four ribonucleoside triphosphate substrates and manipulation of the DNA and RNA in an elongation complex. These advances in elongation complex isolation have been instrumental in the identification and detailed analysis of the nascent RNA cleavage reaction of RNA polymerase II. This is an elongation complexassociated activity of the enzyme that is activated by elongation factor SII and is involved in transcription through a variety of obstacles to elongation. By exploiting cloned regions of DNA containing either G-less (or U-less) stretches of template or naturally occurring arrest sites, it has become routine to generate large amounts of specific, template-engaged elongation complexes that can be isolated and used as starting material for studying the elongation process. These preparations provide highly purified populations of a specific elongation complex, which, when provided with an appropriate set of ribonucleoside triphosphates, resumes synchronous elongation. By manipulating reaction conditions, it is possible to incorporate radioactive nucleotides into specific positions in the transcript. The first few residues of the transcript can be labeled to high specific activity with a radioactive nucleotide; one can ensure that the label is restricted to the 5*-end of the transcript by incubating reaction mixtures for a very short time with low concentrations of highly radioactive NTP substrates or by omitting one or more of the four NTPs on an appropriately engineered template. If subsequent elongation is conducted in the absence of additional labeled substrate, transcripts become labeled to the same specific activity regardless of their length. For some applications the labeled nucleotide(s) needs to be incorporated at or near the transcript 3*-end. There are a number of ways that this can be done, including stepwise ‘‘walking’’ of RNA polymerase on a template by providing and
AID
Methods A 0471
/
6714$$$161
removing one or more of the NTPs. We have found it useful to exploit the SII-activated, nascent RNA cleavage activity of RNA polymerase II to attain a unique 3*-end labeled RNA (40–42). Complexes bearing unlabeled RNA chains are assembled at a naturally occurring arrest site. They are isolated using an anti-RNA antibody (see below) and washed free of NTPs. In the presence of elongation factor SII and Mg2/, a specific oligonucleotide is removed from the 3*-end of the RNA chain. The remaining 5*-end of the RNA is held in an active elongation complex. With the knowledge of the sequence at which cleavage takes place, a single labeled nucleotide of choice is added with or without other required nucleotides to position the label at a unique internal, or 3*-terminal, position. This strategy is also effective in placing photoactive nucleotide derivatives into strategic locations within an RNA chain (42). Velocity Sedimentation and Gel Filtration Early work using velocity sedimentation as an analytical tool enabled the identification of initiated transcription complexes (43). The limited resolution and long duration of the ultracentrifugation procedure, coupled with the difficulty of obtaining sufficiently large amounts of material for detailed structural and mechanistic studies, has restricted the wide use of this separation technique. Both high ionic strength and the detergent Sarkosyl will inhibit interaction of elongation factors such as TFIIF, elongin (SIII), and SII with RNA polymerase II (8, 44–47), as well as interfere with other protein–protein and protein–nucleic acid interactions. Consequently, gel filtration in the presence of high ionic strength or Sarkosyl separates elongation complexes from the majority of proteins in a nuclear extract, including most of the general initiation and elongation factors (45, 47, 48); if a gel of appropriate porosity is chosen, elongation complexes can even be separated from large molecules such as free RNA polymerase II (49). As is the case for velocity sedimentation, drawbacks to this approach include sample dilution and the length of time needed for highresolution separations. Nevertheless, a number of laboratories have very effectively used gel filtration to assemble and isolate purified populations of elongation complexes to study elongation and, in particular, the nascent RNA cleavage reaction and elongation on templates assembled into chromatin (47, 48, 50–57).
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
Affinity Isolation of Elongation Complexes Perhaps the most popular and effective recent innovation has been the immobilization of elongation complexes through an affinity ligand (Fig. 5). Important advances in our understanding of the structure and function of elongation complexes have resulted from purifying the specific complexes via their RNA (58), DNA (59, 60), or RNA polymerase (61) moieties. The first two approaches are discussed below. The latter approach has thus far been carried out only with E. coli RNA polymerase due to the relative ease with which a holoenzyme containing a hexahistidine-tagged subunit can be generated in bacteria (62, 63). In principle, however, this method could be applied to appropriately modified RNA polymerase II. DNA-mediated immobilization. DNA polymerases can use biotinylated derivatives of dUTP as a substrate for incorporation into DNA in vitro (64). Biotinylated DNA binds tightly to avidin or streptavidin immobilized on a solid support such as agarose.
199
The first use of biotin–streptavidin immobilized DNA to dissect the transcription reaction into multiple steps was reported by Arias and Dynan (65). The ability to rapidly change the buffer composition enabled these workers to define substrate requirements for various stages in transcription initiation (66). Biotinylated DNA templates were initially generated by filling in the ends of restricted DNA with biotinylated dUTP; biotinylated templates can also be prepared by PCR with biotinylated oligonucleotides as primers. The use of biotinylated DNA templates has been extended to the study of mechanisms of transcription elongation by RNA polymerase II (59, 60, 67). It is preferable in these studies to immobilize only the upstream end of the transcribed DNA duplex. Thus, only the upstream primer needs to be biotyinylated. The biotinylated base should also be incorporated far enough upstream such that enough spacing DNA remains upstream of the promoter to allow transcription factor binding. As well, streptavidin coupled to paramagnetic particles (Dynabeads
FIG. 5. Methods for immobilizing RNA polymerase II elongation complexes. Diagram of immobilized elongation complexes are not to scale. (Top) Elongation complexes immunosorbed to solid-phase-bound protein A (shaded semicircle) via their RNA using an antiRNA monoclonal antibody (‘‘Y’’). (Bottom) Elongation complexes adsorbed to solid-phase-bound avidin (shaded semicircle) via a biotinylated terminal nucleotide on one strand of the DNA duplex. Transcription can be initiated (/1) from an authentic promoter in the presence of general transcription factors.
AID
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
200
M280; Dynal Inc., Great Neck, NY) can be used as the solid phase, and immobilized material can be washed exhaustively by repeated rounds of magnetic concentration followed by resuspension (59). The use of this system was instrumental in the identification and purification of a Drosophila elongation factor known as P-TEFb (68). This approach was also used to demonstrate the SII-activated, nascent RNA cleavage reaction carried out by RNA polymerase II elongation complexes (60). An advantage of DNA-mediated immobilization of elongation complexes over RNA-mediated immobilization is that transcript release can be readily assayed by treating the desired elongation complexes with protein fractions, removing them by centrifugal or magnetic force, and assaying for labeled transcripts remaining in the supernatant. Xie and Price successfully employed this approach to identify and purify a novel Drosophila protein that catalyzes ATP-dependent transcript release by RNA polymerase II (69). RNA-mediated immobilization. Although bacteriophage and other RNA polymerases can incorporate biotinylated nucleotides into RNA for affinity selection of the transcript, RNA polymerase II does not use these base analogs to an appreciable extent (64; and D. Reines, unpublished results). Instead, a monoclonal antibody designated D44 (70), which recognizes RNA in a sequence-independent manner, has been used for extensive analysis of the structure and function of RNA polymerase II elongation complexes. Its benefits include (i) a nearly infinite supply of reagent, (ii) no need to derivatize components of the elongation complex, (iii) a binding interaction that is complete after a few minutes on ice, (iv) separation of elongation complexes by virtue of the fact that they are synthetically active, and (v) concentration of transcriptionally active components with removal of unoccupied templates and inactive RNA polymerase II. Template walking is easily accomplished via the rapid exchange of reaction solutes by brief centrifugation and resuspension in a buffer of choice. The D44 antibody is an IgG3 produced by a hybridoma resulting from the fusion of a myeloma with splenocytes from an unimmunized NZB/NZW F1 mouse (70). This strain of mice displays features of autoimmune disease including circulating anti-nucleic acid antibodies. D44 was initially identified by its immunoreactivity with E. coli rRNA and the syn-
AID
Methods A 0471
/
6714$$$161
thetic copolymer polyribo(G,C). Interestingly, D44 did not react well with poly(A,G), poly(A,C), poly(C,U), poly(G,U), poly(A,U) or any of the homopolymers tested in enzyme-linked immunosorbent assays. The triple-base copolymers poly(GCU) and poly(GCA) are also strongly preferred over poly(GAU) and poly(ACU) (70). Although this analysis suggests that the guanine and cytosine content of RNA is an important part of the epitope, the specific antigenic determinants recognized by D44 IgG are not understood. RNA chain length is another variable that can influence the avidity of the interaction with D44: larger RNAs generally bind the antibody more efficiently. This limitation can be overcome by increasing the concentration of IgG in the reaction, although caution is recommended since at very high concentrations D44 will interact with DNA (71, 72). For most purposes DNA cross-reactivity is not a problem. Experimentally we have found that a variety of RNAs react with the antibody. Nevertheless, pilot experiments are recommended to ensure that a particular RNA serves as a target for D44 IgG. As an IgG, RNA-bound D44 can be absorbed to formalin-fixed Staphylococcus aureus cells (Immunoprecipitin; BRL-Life Technologies, Inc.) or protein A – Sepharose beads. We have found the former less expensive and easier to handle. A suspension of washed S. aureus is added to elongation complexes with D44 IgG and incubated on ice for 10 min. Solid-phase-adsorbed elongation complexes remain active for transcript extension and can be treated with nucleases for footprinting experiments (73). Experiments to analyze the nascent RNA cleavage activity stimulated by elongation factor SII have taken advantage of the fact that the small oligonucleotides liberated by hydrolysis are released into the soluble fraction and can be readily removed from protein, DNA, and elongation complex-associated RNA by microcentrifugation for 5 min at 15,000g (40). Elongation complexes can be released from the solid phase under standard reaction termination conditions, which include sodium dodecyl sulfate, and in a form suitable for analysis of proteins or nucleic acids by denaturing gel electrophoresis. Native elongation complexes are eluted from the solid phase by providing an excess of rRNA competitor for S. aureusadsorbed D44 to bind (72) or by hydrolyzing the RNA that tethers the protein and DNA to the S. aureus surface (42). D44-mediated immunoprecipitation of RNA-containing complexes can easily be
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
adapted for detection of other RNA-binding proteins (74).
201
12. Kadesch, T. R., Rosenberg, S., and Chamberlin, M. J. (1982) J. Mol. Biol. 155, 1–29. 13. Conaway, R. C., Garrett, K. P., Hanley, J. P., and Conaway, J. W. (1991) Proc. Natl. Acad. Sci. USA 88, 6205–6209.
Assumptions in the Use of Isolated Elongation Complexes Are partially assembled transcription complexes or those that have been stopped on the template experimentally and subjected to purification regimens truly representative of those complexes in vivo? Both transcription initiation and elongation are dynamic processes, and it has been documented that structural and functional states of RNA polymerase II can change over time. Although these experiments have great utility in elucidating the steps involved in transcription, the investigator should bear in mind the possibility that experimentally frozen snapshots of transcription complexes give insight into structural or functional states that can be populated, but they may not necessarily represent an intermediate found in vivo. These issues are discussed in more detail by Krummel and Chamberlin (75).
14. Aso, T., Conaway, J. W., and Conaway, R. C. (1994) J. Biol. Chem. 269, 26575–26583. 15. Parvin, J. D., and Sharp, P. A. (1993) Cell 73, 533–540. 16. Timmers, H. Th. M. (1994) EMBO J. 13, 391–399. 17. Parvin, J. D., Shykind, B. M., Meyers, R. E., Kim, J., and Sharp, P. A. (1994) J. Biol. Chem. 269, 18414–18421. 18. Holstege, F., Tantin, D., Carey, M., van der Vliet, P. C., and Timmers, H. Th. M. (1995) EMBO J. 14, 810–819. 19. Pan, G., and Greenblatt, J. (1994) J. Biol. Chem. 269, 30101– 30104. 20. Tantin, D., and Carey, M. (1994) J. Biol. Chem. 269, 17397– 17400. 21. Holstege, F. C. P., van der Vliet, P. C., and Timmers, H. Th. M. (1996) EMBO J. 15, 1666–1677. 22. Gralla, J. D. (1996) in Methods in Enzymology (Adhya, S., Ed.), Vol. 273 pp. 99–109, Academic Press, San Diego. 23. Luse, D. S., and Jacob, G. A. (1987) J. Biol. Chem. 262, 14990–14997. 24. Dvir, A., Garrett, K. P., Chalut, C., Egly, J. M., Conaway, J. W., and Conaway, R. C. (1996) J. Biol. Chem. 271, 7245– 7248. 25. Samuels, M., Fire, A., and Sharp, P. A. (1984) J. Biol. Chem. 259, 2517–2525.
REFERENCES
26. Edwards, A. M., and Kane, C. M. (1996) in Methods in Enzymology (Adhya, S., Ed.), Vol. 274, pp. 419–436, Academic Press, San Diego.
1. Roeder, R. G. (1996) Trends. Biochem. Sci. 21, 327–335. 2. Reines, D., Conaway, J. W., and Conaway, R. C. (1996) Trends. Biochem. Sci. 21, 351–355. 3. Weil, P. A., Luse, D. S., Segall, J., and Roeder, R. G. (1979) Cell 18, 469–484. 4. Sawadogo, M., and Roeder, R. G. (1985) Proc. Natl. Acad. Sci. USA 82, 4394–4398. 5. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
27. Kadesch, T. R., and Chamberlin, M. J. (1982) J. Biol. Chem. 257, 5286–5295. 28. Lorch, Y., LaPointe, J. W., and Kornberg, R. D. (1987) Cell 49, 203–210. 29. Dedrick, R. L., and Chamberlin, M. J. (1985) Biochemistry 24, 2245–2253. 30. Mougey, E. B., and Dennis, D. (1988) Anal. Biochem. 171, 256–265. 31. Lewis, M. K., and Burgess, R. R. (1980) J. Biol. Chem. 255, 4928–4936.
6. Triezenberg, S. J. (1995) in Current Protocols in Molecular Biology, Vol. 1 (Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., Struhl, K., Albright, L. M., Coen, D. M., Varki, A., Eds.) pp. 4.8.1–4.8.5, Wiley, New York.
33. Jeon, C., and Agarwal, K. (1996) Proc. Natl. Acad. Sci. USA 93, 13677–13682.
7. Hawley, D. K., and Roeder, R. G. (1985) J. Biol. Chem. 260, 8163–8172.
34. Tripatara, A., and deHaseth, P. L. (1993) J. Mol. Biol. 233, 349–358.
8. Hawley, D. K., and Roeder, R. G. (1987) J. Biol. Chem. 262, 3452–3461.
35. Daube, S. S., and von Hippel, P. H. (1992) Science 258, 1320– 1324.
9. Conaway, R. C., and Conaway, J. W. (1988) J. Biol. Chem. 263, 2962–2968.
36. Dynan, W. S., and Burgess, R. R. (1981) J. Biol. Chem. 256, 5866–5873.
10. Zheng, X. M., Moncollin, V., Egly, J. M., and Chambon, P. (1987) Cell 50, 361–368.
37. Lescure, B., Bennetzen, J., and Sentenac, A. (1981) J. Biol. Chem. 256, 11018–11024.
11. Conaway, R. C., and Conaway, J. W. (1990) J. Biol. Chem. 265, 7559–7563.
38. Hirose, S., Tsuda, M., and Suzuki, Y. (1985) J. Biol. Chem. 260, 10557–10562.
AID
Methods A 0471
/
6714$$$161
32. Takagi, Y., Conaway, J. W., and Conaway, R. C. (1995) J. Biol. Chem. 270, 24300–24305.
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
202
39. Sekimizu, K., Horikoshi, M., and Natori, S. (1984) J. Biochem. (Tokyo) 96, 1859–1865. 40. Gu, W., and Reines, D. (1995) J. Biol. Chem. 270, 11238– 11244.
58. 59.
41. Gu, W., Wind, M., and Reines, D. (1996) Proc. Natl. Acad. Sci. USA 93, 6935–6940.
60.
42. Powell, W., and Reines, D. (1996) J. Biol. Chem. 271, 22301– 22304.
61.
43. Horikoshi, M., Sekimizu, K., and Natori, S. (1984) J. Biol. Chem. 259, 608–611.
62.
44. Price, D. H., Sluder, A. E., and Greenleaf, A. L. (1989) Mol. Cell. Biol. 9, 1465–1475.
63.
45. Bengal, E., Goldring, A., and Aloni, Y. (1989) J. Biol. Chem. 264, 18926–18932.
64.
46. Kephart, D. D., Marshall, N. F., and Price, D. H. (1992) Mol. Cell. Biol. 12, 2067–2077.
65.
47. Izban, M. G., and Luse, D. S. (1991) Genes Dev. 5, 683–696. 48. Bengal, E., Flores, O., Krauskopf, A., Reinberg, D., and Aloni, Y. (1991) Mol. Cell. Biol. 11, 1195–1206. 49. Kang, M. E., and Dahmus, M. E. (1993) J. Biol. Chem. 268, 25033–25040. 50. Izban, M. G., and Luse, D. S. (1992) J. Biol. Chem. 267, 13647–13655. 51. Izban, M. G., and Luse, D. S. (1992) Genes Dev. 6, 1342–1356.
66. 67. 68. 69.
52. Izban, M. G., and Luse, D. S. (1993) J. Biol. Chem. 268, 12874–12885.
70.
53. Izban, M. G., and Luse, D. S. (1993) J. Biol. Chem. 268, 12864–12873.
71.
54. Chen, Y., Chafin, D., Price, D. H., and Greenleaf, A. L. (1996) J. Biol. Chem. 271, 5993–5999.
72. 73.
55. Guo, H., and Price, D. H. (1993) J. Biol. Chem. 268, 18762– 18770.
74.
56. Christie, K. R., Awrey, D. E., Edwards, A. M., and Kane, C. M. (1994) J. Biol. Chem. 269, 936–943.
75.
57. Wu, J., Awrey, D. E., Edwards, A. M., Archambault, J., and
AID
Methods A 0471
/
6714$$$161
Friesen, J. D. (1996) Proc. Natl. Acad. Sci. USA 93, 11552– 11557. Reines, D. (1991) J. Biol. Chem. 266, 10510–10517. Marshall, N. F., and Price, D. H. (1992) Mol. Cell. Biol. 12, 2078–2090. Wang, D. G., and Hawley, D. K. (1993) Proc. Natl. Acad. Sci. USA 90, 843–847. Borukhov, S., Sagitov, V., and Goldfarb, A. (1993) Cell 72, 459–466. Kashlev, M., Martin, E., Polyakov, A., Severinov, K., Nikiforov, V., and Goldfarb, A. (1993) Gene 130, 9–14. Wang, D., Meier, T. I., Chan, C. L., Feng, G., Lee, D. N., and Landick, R. (1995) Cell 81, 341–350. Langer, B. R., Waldrop, A. A., and Ward, D. C. (1981) Proc. Natl. Acad. Sci. USA 78, 6633–6637. Arias, J. A., and Dynan, W. S. (1989) J. Biol. Chem. 264, 3223–3229. Arias, J. A., Peterson, S. R., and Dynan, W. S. (1991) J. Biol. Chem. 266, 8055–8061. Wiest, D. K., Wang, D., and Hawley, D. K. (1992) J. Biol. Chem. 267, 7733–7744. Marshall, N. F., Peng, J., Xie, Z., and Price, D. H. (1996) J. Biol. Chem. 271, 27176–27183. Xie, Z., and Price, D. H. (1996) J. Biol. Chem. 271, 11043– 11046. Eilat, D., Hochberg, M., Fischel, R., and Laskov, R. (1982) Proc. Natl. Acad. Sci. USA 79, 3818. Eilat, D., Hochberg, M., Pumphrey, J., and Rudikoff, S. (1984) J. Immunol. 133, 489–494. Reines, D. (1991) Anal. Biochem. 196, 367–372. Gu, W., Powell, W., Mote, J., and Reines, D. (1993) J. Biol. Chem. 268, 25604–25616. Ashley, C. T., Wilkinson, K. D., Reines, D., and Warren, S. T. (1993) Science 262, 563–566. Krummel, B., and Chamberlin, M. J. (1992) J. Mol. Biol. 225, 221–237.
06-19-97 15:14:08
metaas
AP: Methods A
Assays for Investigating Transcription by RNA Polymerase II in Vitro Daniel Reines,* Arik Dvir,† Joan Weliky Conaway,†,‡ and Ronald C. Conaway†,1 *Department of Biochemistry, Emory University School of Medicine, Atlanta, Georgia 30322; †Program in Molecular and Cell Biology, Oklahoma Medical Research Foundation, 825 N.E. 13th Street, Oklahoma City, Oklahoma 73104; and ‡Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, Oklahoma 73190
With the availability of the general initiation factors (TFIIB, TFIID, TFIIE, TFIIF, and TFIIH), it is now possible to investigate aspects of the mechanism of eukaryotic messenger RNA synthesis in purified, reconstituted RNA polymerase II transcription systems. Rapid progress in these investigations has been spurred by use of a growing number of assays that are proving valuable not only for dissecting the molecular mechanisms of transcription initiation and elongation by RNA polymerase II, but also for identifying and purifying novel transcription factors that regulate polymerase activity. Here we describe a variety of these assays and discuss their utility in the analysis of transcription by RNA polymerase II. q 1997 Academic Press
Messenger RNA synthesis is a major site for the regulation of gene expression. Eukaryotic messenger RNA synthesis is a complex biochemical process catalyzed by multisubunit RNA polymerase II and governed by the concerted action of a diverse collection of transcription factors (TF) that control polymerase activity at both the initiation and the elongation stages of transcription (1, 2). Over the past 10 years, substantial progress defining the molecular mecha1 To whom correspondence should be addressed. Fax: (405) 2711580.
nisms underlying the initiation and elongation stages of eukaryotic messenger RNA synthesis has been achieved. This progress is due in large part to two major advances: (i) the development of purified, reconstituted RNA polymerase II transcription systems amenable to detailed mechanistic analysis and (ii) the creation of novel assays for dissecting the mechanism of transcription in these systems. With the resolution and purification of the general initiation factors (TFIIB, TFIID, TFIIE, TFIIF, and TFIIH), basal transcription by RNA polymerase II can now be reconstituted with purified proteins from a large number of structurally diverse promoters. This ‘‘minimal’’ transcription system is proving valuable not only as a tool for investigating the mechanism of transcription by RNA polymerase II, but also as an assay system for identifying and purifying novel transcription factors that regulate polymerase activity. The development of well-defined transcription systems has been accompanied by a proliferation of powerful assays for investigating transcription. A number of these assays exploit specialized DNA templates. In this article, we describe the most commonly used variations of these assays and discuss their applications. Because basic procedures for purifying RNA polymerase II and the general initiation factors and for reconstituting optimized basal transcription have recently been thoroughly reviewed in Methods in Enzymology (Vols. 273 and 274), we will not discuss them here.
192
AID
1046-2023/97 $25.00 Copyright q 1997 by Academic Press All rights of reproduction in any form reserved.
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
DESCRIPTION OF THE METHOD General Assays for Measuring Promoter-Dependent Transcription by RNA Polymerase II RNA polymerase II will initiate transcription accurately in vitro from a large number of structurally diverse promoters. A variety of assays for detecting and quantitating initiation by RNA polymerase II from promoters are currently in use. These include the runoff transcription (3), G-less cassette (4), S1 nuclease, RNase protection, and primer extension assays (5, 6) (Fig. 1). Of these assays, the S1 nuclease, RNase protection, and primer extension assays are superior for confirming that RNA polymerase II is initiating transcription in vitro at the correct transcriptional start sites, but they are laborious and time-consuming to perform. The runoff transcription and G-less cassette assays are simpler to perform and are therefore preferred for most measurements of promoter-dependent transcription by RNA polymerase II. Although the runoff transcription assay is the method of choice for most purposes, it has several disadvantages that preclude its use in select circumstances. First, the runoff transcription assay cannot be used to investigate the role of DNA topology in transcription by RNA polymerase II, because it re-
FIG. 1. Commonly used assays of promoter-dependent transcription by RNA polymerase II. The asterisk in the bottom panel designates a radioactive label at the 5*-end of an oligonucleotide used for primer extension analysis. TATA, TATA box; Inr, initiator element.
AID
Methods A 0471
/
6714$$$161
193
quires a linear DNA template; for such studies, the G-less cassette assay is ideal because closed circular DNAs can be used as templates for transcription. Second, the runoff transcription assay is often not well suited for investigating transcription in crude enzyme systems, because high levels of nonspecific background transcription initiated at ends, nicks, or cryptic promoters in some DNA templates can obscure the correctly initiated transcript; in these instances, the G-less cassette assay is preferred because background transcription can be reduced by including only ATP, CTP, and UTP as substrates for elongation by RNA polymerase II and by treating reaction products with ribonuclease T1 , which cleaves RNA after G-residues. Assays for Investigating Specific Steps in PromoterDependent Transcription by RNA Polymerase II Although the general assays described above are useful for quantitating promoter-dependent transcription by RNA polymerase II, they are not sufficient for dissecting its mechanism. Promoter-dependent transcription by RNA polymerase II is a complex, multistep process requiring (i) assembly of polymerase and its initiation factors into a functional preinitiation complex, (ii) ATP-dependent unwinding of promoter DNA surrounding the transcriptional start site to form an ‘‘open complex,’’ (iii) transcription initiation (i.e., synthesis of the first phosphodiester bond of nascent transcripts), (iv) promoter escape (i.e., conversion of the unstable initiation complex into a stable elongation complex), and (v) elongation of full-length transcripts (1, 2). Below we describe a variety of assays that have been used successfully to elucidate many aspects of the mechanism of promoter-dependent transcription by RNA polymerase II and the roles of individual transcription factors in this process.
Sarkosyl and Heparin Challenge Assays In studies investigating the mechanism of transcription initiation by RNA polymerase II, it is often important to carry out transcription reactions under conditions in which (i) initiation is limited to a single round per promoter and (ii) events occurring during initiation and synthesis of the first few phosphodiester bonds of nascent transcripts can be distinguished from events occurring during subsequent steps in transcription. This is readily accomplished using Sarkosyl (N-lauroylsarcosine) or heparin challenge
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
194
assays. Elongation of 4- to 10-nucleotide-long transcripts by RNA polymerase II is resistant to inhibition by 0.2–0.5% Sarkosyl (7–9) and by 10–100 mg/ ml heparin (10, 11), whereas assembly of the preintiation complex and synthesis of the first 2–3 phosphodiester bonds of nascent transcripts are not. Thus, if Sarkosyl or heparin is added to transcription reactions shortly after addition of ribonucleoside triphosphates, full-length runoff transcripts will be synthesized only from those promoters at which initiation has already occurred. A convenient way to limit initiation to a single round per preinitiation complex is to initiate transcription with only the subset of ribonucleoside triphosphates needed for synthesis of the first 4–10 phosphodiester bonds of transcripts; following addition of Sarkosyl or heparin, full-length transcripts are synthesized by addition of the remaining ribonucleoside triphosphates.
The Template Competition Assay This assay is a modification of the template competition assay originally described by Chamberlin and co-workers (12). The template competition assay is well suited for investigating the requirements for interaction of RNA polymerase II and its transcription factors with promoters, and it has proven quite valuable in biochemical studies dissecting the mechanism of assembly of the preinitiation complex (11, 13, 14). Although the template competition assay is more difficult to perform than some other assays for identification and analysis of RNA polymerase II preinitiation intermediates, it offers several advantages. Unlike the gel-shift assay, for example, which subjects putative preinitiation intermediates to the nonoptimal transcription reaction conditions present in native polyacrylamide gels, the template competition assay is carried out entirely under optimal transcription reaction conditions. In addition, unlike the gel-shift assay, which does not permit easy assay of the transcriptional activities of putative preinitiation intermediates identified as bands in native polyacrylamide gels, the template competition assay permits identification of ‘‘functional’’ preinitiation intermediates by direct measurement of their transcriptional activities. Template challenge assays are performed as diagrammed in Fig. 2. Two different plasmids, each containing the same promoter, are used as DNA templates. The plasmids are linearized by restriction enzyme digestion, so that they direct synthesis of
AID
Methods A 0471
/
6714$$$161
runoff transcripts of different lengths. During an initial preincubation, templates I and II are incubated in separate reaction mixtures with various combinations of RNA polymerase II and transcription factors; at this stage, it is important that the transcription factor of interest is present at a subsaturating concentration. The two reaction mixtures are combined to begin a second preincubation and, where necessary, the remaining transcription proteins required for assembly of the complete preinitiation complex are added to reaction mixtures. Following the second preincubation, which must be long enough to allow binding reactions to come to equilibrium, transcription is carried out using the Sarkosyl or heparin challenge assay to limit initiations to a single round per preinitiation complex; if initations are not limited to a single round, misleading results can be obtained because transcription proteins released from the first template following initiation may be able to assemble into preinitiation complexes and promote initiation on the second template. The relative molar amounts of full-length runoff transcripts synthesized from each template are determined and expressed as the molar ratio (I/II) of transcription from templates I and II. If the transcription factor of interest is capable of interacting stably and stoichiometrically with preinitiation intermediates on template I in the presence of the set of transcription proteins provided during the first preincubation, the molar ratio of transcripts synthesized on the two templates will be greater than that of a control reaction in which the two templates are mixed at the beginning of the first preincubation. On the other hand, if the transcription factor interacts catalytically, or not at all, with preinitiation intermediates on template I during the first preincubation, it should distribute equally on both templates during the second preincubation, and the molar ratio of transcripts synthesized from the two templates should be the same as that of the control reaction.
Assays That Bypass Requirements for Subsets of Initiation Factors and ATP-Dependent Open Complex Formation Negatively supercoiled and ‘‘bubble’’ templates have proven useful for investigating the roles of individual transcription factors in promoter-dependent transcription initiation by RNA polymerase II. Negatively supercoiled templates bypass the requirement for TFIIH and ATP in open complex formation,
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
most likely because the stored free energy of supercoiling facilitates unwinding of promoter DNA around the transcriptional start site. The utility of negatively supercoiled DNA templates first became apparent with the observation that initiation by RNA polymerase II at a negatively supercoiled human IgH promoter requires only TFIIB and TBP or TFIID and is independent of ATP, whereas initiation by polymerase at the same relaxed IgH promoter requires the full complement of general initiation factors and depends on ATP (15, 16). Subsequent investigation revealed that TFIIH and ATP are dispensable for initiation at promoters on all negatively supercoiled templates tested; however, the requirement for other initiation factors varies depending on the promoter sequence (16–18). For example, while transcription initiation by RNA polymerase II at the negatively supercoiled IgH promoter requires only TFIIB and TFIID or TBP, initiation at the negatively supercoiled adenovirus E4 promoter requires TFIIB, TFIID or TBP, and TFIIF, and initiation at the supercoiled AdML promoter requires TFIIB, TFIID or TBP, and TFIIF, but is strongly stimulated by TFIIE. Thus, by choosing supercoiled templates containing the appropriate promoters, it is possible to devise assays that can be used for studying the
195
mechanisms of action of different subsets of initiation factors. Because open complex formation on negatively supercoiled templates is driven by the energy of supercoiling, it is not possible to control precisely the position of the region of unwound DNA. In addition, negatively supercoiled templates have the significant disadvantage that they must remain supercoiled in the presence of RNA polymerase II and transcription factors and therefore must be used in transcription systems devoid of nucleases and topoisomerase I. Bubble templates, which contain premelted promoters with short stretches of noncomplementary bases surrounding the transcriptional start site, bypass the requirement for TFIIH and ATP in initiation and provide an alternative to negatively supercoiled templates. Because bubble templates are prepared synthetically, by hybridization of either synthetic oligonucleotides (19) or larger singlestranded DNA fragments (20), the position of the premelted region of DNA can be precisely controlled. Furthermore, because bubble templates do not need to be supercoiled, they are much less sensitive to inactivation by nucleases and therefore can be used in relatively crude transcription systems. Detailed
FIG. 2. The template competition assay. The bottom panel is a mock autoradiogram of a denaturing polyacrylamide gel analyzing the transcripts synthesized in the template competition assay outlined in the top panel.
AID
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
196
procedures for preparing supercoiled and premelted DNA templates can be found in Refs. (15, 19–21).
Abortive Initiation Assay The abortive initiation assay has been widely used in investigations of the requirements for synthesis of the first phosphodiester bond of nascent transcripts by both prokaryotic and eukaryotic RNA polymerases (18, 22–24). The abortive initiation assay exploits the ability of RNA polymerase to utilize dinucleotides to prime synthesis of transcripts at its promoters. If only a dinucleotide and the next ribonucleoside triphosphate encoded by the DNA template are provided as substrates for transcription, RNA polymerase II will reiteratively synthesize trinucleotide transcripts that are rapidly released from the polymerase active site. Dinucleotideprimed transcription initiation by RNA polymerase II can occur over an approximately nine basepair region surrounding transcriptional start site (25). Thus, at the AdML promoter, for example, the dinucleotides CpA and CpU support trinucleotide synthesis in the presence of [a-32P]CTP; CpC, ApC, and UpC support trinucleotide synthesis in the presence of [a-32P]UTP; and UpC supports trinucleotide synthesis in the presence of [a-32P]ATP (Fig. 3). Dinucleotide-primed trinucleotide transcripts are easily separated from unincorporated [a-32P]NTPs by electrophoresis through 28% acrylamide/3% bisacrylamide denaturing gels (23). Assays for Investigating the Mechanism of Elongation by RNA Polymerase II A variety of assays that exploit the intrinsic ability of RNA polymerase II to initiate transcription in the
absence of accessory factors at single-stranded regions of DNA or at nicks and ends of double-stranded DNA have been developed for monitoring polymerase activity during purification, for probing aspects of polymerase catalytic activity, and for investigating the mechanisms of action of elongation factors. Transcripts are internally labeled using [a-32P]NTPs and can be conveniently analyzed by denaturing polyacrylamide gel electrophoresis, followed by autoradiography or PhosphorImager analysis, or by counting radioactivity of samples directly after removal of unincorporated [a-32P]NTPs from labeled RNA products by trichloroacetic acid precipitation or DE-81 adsorption (5). The simplest assays for promoter-independent transcription by RNA polymerase II utilize sheared, single-stranded calf thymus or salmon sperm DNA as template (26). In addition, we describe below a series of specialized DNA templates that have been developed for assay of promoter-independent transcription by RNA polymerase II (Fig. 4).
Tailed, Dumbbell, and Bubble Templates DNA duplexes which are extended at one 3*-end are excellent templates for a number of RNA polymerases including RNA polymerase II (Fig. 4). Extended ‘‘tailed’’ duplexes can be synthesized enzymatically using terminal transferase and dCTP (the favored base) (27). To restrict transcription to initiation from only a single end of the DNA, one end is usually removed by digestion with a restriction enzyme that yields a short DNA duplex containing one tailed 3*-terminus. If small RNA transcripts are
FIG. 3. Requirements for dinucleotide-primed abortive initiation at the AdML promoter. A*, U*, and C* refer to [a-32P]ATP, [a-32P]UTP, and [a-32P]/CTP, respectively.
AID
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
to be analyzed, this small transcriptionally active DNA fragment can be removed by standard DNA purification techniques. Tailed templates can also be formed by ligating an oligonucleotide onto a linear DNA duplex containing a 5*-overhanging ‘‘sticky’’ end (28). RNA polymerase II binds at the tail–duplex junction, where it initiates an RNA chain complementary to a few bases of the single strand adjacent to the duplex (29). The site of transcription initiation can be fixed more precisely by initiating with a dinucleotide primer complementary to specific template bases on the single-stranded tail (30). An advantage of tailed templates is that they are used very efficiently by RNA polymerase II such that most, if not all, of the DNA duplexes are occupied by transcriptionally engaged polymerase. This is in contrast to the relatively low template occupancy in most promoter-dependent reactions reconstituted with general transcription factors. An unusual feature of tailed templates, however, is the propensity for the nontemplate strand to be displaced during RNA synthesis such that the transcript remains base-paired with the template strand and a long RNA:DNA hybrid is produced. This aspect of transcription on tailed templates has recently been described in detail (26). In addition, because transcription initiates at the end of the template, full-length transcripts are nearly the same length as the template DNA. This can lead to confusion regarding the identities of reaction products since RNA polymerase II has the ability to label the nontemplate strand of DNA by adding a radioactive ribonucleotide to the 3*-end (31). Interestingly, this activity of RNA polymerase II is strongly stimulated by the transcription factor elongin (SIII) (32). The ‘‘dumbbell’’ template, on which RNA polymer-
ase II can extend a ribonucleic acid primer in a template-dependent manner (33), is the basis of another useful assay (Fig. 4). The dumbbell template is a chemically synthesized oligonucleotide that can base-pair extensively across its length, except for a short single-stranded gap; the few 3*-terminal bases are ribonucleotides added to a deoxyribonucleotide chain. The dumbbell structure is thought to mimic a transcription bubble with RNA bases hydrogenbonded to a DNA template strand. RNA polymerase II covalently attaches ribonucleotides to this single chain, which can be extended by copying the DNA template strand until its 5*-end is reached. The size of the oligonucleotide that can be obtained through chemical synthesis will necessarily limit the length of transcripts extended; thus, dumbbell templates cannot be used to study elongation of lengthy transcripts. Analysis of Escherichia coli RNA polymerase has been facilitated by the reconstitution of a synthetic transcription bubble using double-stranded DNA containing an internal segment that is mismatched and cannot base-pair (34). In a more complete reconstruction of a transcription bubble, an RNA chain annealed to the template strand of the bubble can be used as a primer and extended by E. coli RNA polymerase (35). Analysis of elongation by RNA polymerase II would benefit from this approach, but such experiments have not yet been reported. Earlier reports suggested that certain DNA sequences on supercoiled plasmids, which presumably yield transiently melted regions, allow initiation by purified RNA polymerase II albeit inefficiently (36–39). As described above, synthetic bubble-containing templates have also been useful for studies of promoterdependent transcription by RNA polymerase II.
FIG. 4. Specialized DNA templates for investigating elongation by RNA polymerase II.
AID
Methods A 0471
/
6714$$$161
197
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
198
Isolation and Analysis of RNA Polymerase II Elongation Complexes Utility The isolation of elongation complexes has proven to be extremely useful in examining individual steps in the transcription elongation process. A number of procedures are now commonly used to separate complexes from proteins and small solutes. This makes possible substantial control over reaction conditions, including the removal and provision of specific protein factors and specific subsets of the four ribonucleoside triphosphate substrates and manipulation of the DNA and RNA in an elongation complex. These advances in elongation complex isolation have been instrumental in the identification and detailed analysis of the nascent RNA cleavage reaction of RNA polymerase II. This is an elongation complexassociated activity of the enzyme that is activated by elongation factor SII and is involved in transcription through a variety of obstacles to elongation. By exploiting cloned regions of DNA containing either G-less (or U-less) stretches of template or naturally occurring arrest sites, it has become routine to generate large amounts of specific, template-engaged elongation complexes that can be isolated and used as starting material for studying the elongation process. These preparations provide highly purified populations of a specific elongation complex, which, when provided with an appropriate set of ribonucleoside triphosphates, resumes synchronous elongation. By manipulating reaction conditions, it is possible to incorporate radioactive nucleotides into specific positions in the transcript. The first few residues of the transcript can be labeled to high specific activity with a radioactive nucleotide; one can ensure that the label is restricted to the 5*-end of the transcript by incubating reaction mixtures for a very short time with low concentrations of highly radioactive NTP substrates or by omitting one or more of the four NTPs on an appropriately engineered template. If subsequent elongation is conducted in the absence of additional labeled substrate, transcripts become labeled to the same specific activity regardless of their length. For some applications the labeled nucleotide(s) needs to be incorporated at or near the transcript 3*-end. There are a number of ways that this can be done, including stepwise ‘‘walking’’ of RNA polymerase on a template by providing and
AID
Methods A 0471
/
6714$$$161
removing one or more of the NTPs. We have found it useful to exploit the SII-activated, nascent RNA cleavage activity of RNA polymerase II to attain a unique 3*-end labeled RNA (40–42). Complexes bearing unlabeled RNA chains are assembled at a naturally occurring arrest site. They are isolated using an anti-RNA antibody (see below) and washed free of NTPs. In the presence of elongation factor SII and Mg2/, a specific oligonucleotide is removed from the 3*-end of the RNA chain. The remaining 5*-end of the RNA is held in an active elongation complex. With the knowledge of the sequence at which cleavage takes place, a single labeled nucleotide of choice is added with or without other required nucleotides to position the label at a unique internal, or 3*-terminal, position. This strategy is also effective in placing photoactive nucleotide derivatives into strategic locations within an RNA chain (42). Velocity Sedimentation and Gel Filtration Early work using velocity sedimentation as an analytical tool enabled the identification of initiated transcription complexes (43). The limited resolution and long duration of the ultracentrifugation procedure, coupled with the difficulty of obtaining sufficiently large amounts of material for detailed structural and mechanistic studies, has restricted the wide use of this separation technique. Both high ionic strength and the detergent Sarkosyl will inhibit interaction of elongation factors such as TFIIF, elongin (SIII), and SII with RNA polymerase II (8, 44–47), as well as interfere with other protein–protein and protein–nucleic acid interactions. Consequently, gel filtration in the presence of high ionic strength or Sarkosyl separates elongation complexes from the majority of proteins in a nuclear extract, including most of the general initiation and elongation factors (45, 47, 48); if a gel of appropriate porosity is chosen, elongation complexes can even be separated from large molecules such as free RNA polymerase II (49). As is the case for velocity sedimentation, drawbacks to this approach include sample dilution and the length of time needed for highresolution separations. Nevertheless, a number of laboratories have very effectively used gel filtration to assemble and isolate purified populations of elongation complexes to study elongation and, in particular, the nascent RNA cleavage reaction and elongation on templates assembled into chromatin (47, 48, 50–57).
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
Affinity Isolation of Elongation Complexes Perhaps the most popular and effective recent innovation has been the immobilization of elongation complexes through an affinity ligand (Fig. 5). Important advances in our understanding of the structure and function of elongation complexes have resulted from purifying the specific complexes via their RNA (58), DNA (59, 60), or RNA polymerase (61) moieties. The first two approaches are discussed below. The latter approach has thus far been carried out only with E. coli RNA polymerase due to the relative ease with which a holoenzyme containing a hexahistidine-tagged subunit can be generated in bacteria (62, 63). In principle, however, this method could be applied to appropriately modified RNA polymerase II. DNA-mediated immobilization. DNA polymerases can use biotinylated derivatives of dUTP as a substrate for incorporation into DNA in vitro (64). Biotinylated DNA binds tightly to avidin or streptavidin immobilized on a solid support such as agarose.
199
The first use of biotin–streptavidin immobilized DNA to dissect the transcription reaction into multiple steps was reported by Arias and Dynan (65). The ability to rapidly change the buffer composition enabled these workers to define substrate requirements for various stages in transcription initiation (66). Biotinylated DNA templates were initially generated by filling in the ends of restricted DNA with biotinylated dUTP; biotinylated templates can also be prepared by PCR with biotinylated oligonucleotides as primers. The use of biotinylated DNA templates has been extended to the study of mechanisms of transcription elongation by RNA polymerase II (59, 60, 67). It is preferable in these studies to immobilize only the upstream end of the transcribed DNA duplex. Thus, only the upstream primer needs to be biotyinylated. The biotinylated base should also be incorporated far enough upstream such that enough spacing DNA remains upstream of the promoter to allow transcription factor binding. As well, streptavidin coupled to paramagnetic particles (Dynabeads
FIG. 5. Methods for immobilizing RNA polymerase II elongation complexes. Diagram of immobilized elongation complexes are not to scale. (Top) Elongation complexes immunosorbed to solid-phase-bound protein A (shaded semicircle) via their RNA using an antiRNA monoclonal antibody (‘‘Y’’). (Bottom) Elongation complexes adsorbed to solid-phase-bound avidin (shaded semicircle) via a biotinylated terminal nucleotide on one strand of the DNA duplex. Transcription can be initiated (/1) from an authentic promoter in the presence of general transcription factors.
AID
Methods A 0471
/
6714$$$161
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
200
M280; Dynal Inc., Great Neck, NY) can be used as the solid phase, and immobilized material can be washed exhaustively by repeated rounds of magnetic concentration followed by resuspension (59). The use of this system was instrumental in the identification and purification of a Drosophila elongation factor known as P-TEFb (68). This approach was also used to demonstrate the SII-activated, nascent RNA cleavage reaction carried out by RNA polymerase II elongation complexes (60). An advantage of DNA-mediated immobilization of elongation complexes over RNA-mediated immobilization is that transcript release can be readily assayed by treating the desired elongation complexes with protein fractions, removing them by centrifugal or magnetic force, and assaying for labeled transcripts remaining in the supernatant. Xie and Price successfully employed this approach to identify and purify a novel Drosophila protein that catalyzes ATP-dependent transcript release by RNA polymerase II (69). RNA-mediated immobilization. Although bacteriophage and other RNA polymerases can incorporate biotinylated nucleotides into RNA for affinity selection of the transcript, RNA polymerase II does not use these base analogs to an appreciable extent (64; and D. Reines, unpublished results). Instead, a monoclonal antibody designated D44 (70), which recognizes RNA in a sequence-independent manner, has been used for extensive analysis of the structure and function of RNA polymerase II elongation complexes. Its benefits include (i) a nearly infinite supply of reagent, (ii) no need to derivatize components of the elongation complex, (iii) a binding interaction that is complete after a few minutes on ice, (iv) separation of elongation complexes by virtue of the fact that they are synthetically active, and (v) concentration of transcriptionally active components with removal of unoccupied templates and inactive RNA polymerase II. Template walking is easily accomplished via the rapid exchange of reaction solutes by brief centrifugation and resuspension in a buffer of choice. The D44 antibody is an IgG3 produced by a hybridoma resulting from the fusion of a myeloma with splenocytes from an unimmunized NZB/NZW F1 mouse (70). This strain of mice displays features of autoimmune disease including circulating anti-nucleic acid antibodies. D44 was initially identified by its immunoreactivity with E. coli rRNA and the syn-
AID
Methods A 0471
/
6714$$$161
thetic copolymer polyribo(G,C). Interestingly, D44 did not react well with poly(A,G), poly(A,C), poly(C,U), poly(G,U), poly(A,U) or any of the homopolymers tested in enzyme-linked immunosorbent assays. The triple-base copolymers poly(GCU) and poly(GCA) are also strongly preferred over poly(GAU) and poly(ACU) (70). Although this analysis suggests that the guanine and cytosine content of RNA is an important part of the epitope, the specific antigenic determinants recognized by D44 IgG are not understood. RNA chain length is another variable that can influence the avidity of the interaction with D44: larger RNAs generally bind the antibody more efficiently. This limitation can be overcome by increasing the concentration of IgG in the reaction, although caution is recommended since at very high concentrations D44 will interact with DNA (71, 72). For most purposes DNA cross-reactivity is not a problem. Experimentally we have found that a variety of RNAs react with the antibody. Nevertheless, pilot experiments are recommended to ensure that a particular RNA serves as a target for D44 IgG. As an IgG, RNA-bound D44 can be absorbed to formalin-fixed Staphylococcus aureus cells (Immunoprecipitin; BRL-Life Technologies, Inc.) or protein A – Sepharose beads. We have found the former less expensive and easier to handle. A suspension of washed S. aureus is added to elongation complexes with D44 IgG and incubated on ice for 10 min. Solid-phase-adsorbed elongation complexes remain active for transcript extension and can be treated with nucleases for footprinting experiments (73). Experiments to analyze the nascent RNA cleavage activity stimulated by elongation factor SII have taken advantage of the fact that the small oligonucleotides liberated by hydrolysis are released into the soluble fraction and can be readily removed from protein, DNA, and elongation complex-associated RNA by microcentrifugation for 5 min at 15,000g (40). Elongation complexes can be released from the solid phase under standard reaction termination conditions, which include sodium dodecyl sulfate, and in a form suitable for analysis of proteins or nucleic acids by denaturing gel electrophoresis. Native elongation complexes are eluted from the solid phase by providing an excess of rRNA competitor for S. aureusadsorbed D44 to bind (72) or by hydrolyzing the RNA that tethers the protein and DNA to the S. aureus surface (42). D44-mediated immunoprecipitation of RNA-containing complexes can easily be
06-19-97 15:14:08
metaas
AP: Methods A
ASSAYS FOR TRANSCRIPTION BY RNA POLYMERASE II
adapted for detection of other RNA-binding proteins (74).
201
12. Kadesch, T. R., Rosenberg, S., and Chamberlin, M. J. (1982) J. Mol. Biol. 155, 1–29. 13. Conaway, R. C., Garrett, K. P., Hanley, J. P., and Conaway, J. W. (1991) Proc. Natl. Acad. Sci. USA 88, 6205–6209.
Assumptions in the Use of Isolated Elongation Complexes Are partially assembled transcription complexes or those that have been stopped on the template experimentally and subjected to purification regimens truly representative of those complexes in vivo? Both transcription initiation and elongation are dynamic processes, and it has been documented that structural and functional states of RNA polymerase II can change over time. Although these experiments have great utility in elucidating the steps involved in transcription, the investigator should bear in mind the possibility that experimentally frozen snapshots of transcription complexes give insight into structural or functional states that can be populated, but they may not necessarily represent an intermediate found in vivo. These issues are discussed in more detail by Krummel and Chamberlin (75).
14. Aso, T., Conaway, J. W., and Conaway, R. C. (1994) J. Biol. Chem. 269, 26575–26583. 15. Parvin, J. D., and Sharp, P. A. (1993) Cell 73, 533–540. 16. Timmers, H. Th. M. (1994) EMBO J. 13, 391–399. 17. Parvin, J. D., Shykind, B. M., Meyers, R. E., Kim, J., and Sharp, P. A. (1994) J. Biol. Chem. 269, 18414–18421. 18. Holstege, F., Tantin, D., Carey, M., van der Vliet, P. C., and Timmers, H. Th. M. (1995) EMBO J. 14, 810–819. 19. Pan, G., and Greenblatt, J. (1994) J. Biol. Chem. 269, 30101– 30104. 20. Tantin, D., and Carey, M. (1994) J. Biol. Chem. 269, 17397– 17400. 21. Holstege, F. C. P., van der Vliet, P. C., and Timmers, H. Th. M. (1996) EMBO J. 15, 1666–1677. 22. Gralla, J. D. (1996) in Methods in Enzymology (Adhya, S., Ed.), Vol. 273 pp. 99–109, Academic Press, San Diego. 23. Luse, D. S., and Jacob, G. A. (1987) J. Biol. Chem. 262, 14990–14997. 24. Dvir, A., Garrett, K. P., Chalut, C., Egly, J. M., Conaway, J. W., and Conaway, R. C. (1996) J. Biol. Chem. 271, 7245– 7248. 25. Samuels, M., Fire, A., and Sharp, P. A. (1984) J. Biol. Chem. 259, 2517–2525.
REFERENCES
26. Edwards, A. M., and Kane, C. M. (1996) in Methods in Enzymology (Adhya, S., Ed.), Vol. 274, pp. 419–436, Academic Press, San Diego.
1. Roeder, R. G. (1996) Trends. Biochem. Sci. 21, 327–335. 2. Reines, D., Conaway, J. W., and Conaway, R. C. (1996) Trends. Biochem. Sci. 21, 351–355. 3. Weil, P. A., Luse, D. S., Segall, J., and Roeder, R. G. (1979) Cell 18, 469–484. 4. Sawadogo, M., and Roeder, R. G. (1985) Proc. Natl. Acad. Sci. USA 82, 4394–4398. 5. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
27. Kadesch, T. R., and Chamberlin, M. J. (1982) J. Biol. Chem. 257, 5286–5295. 28. Lorch, Y., LaPointe, J. W., and Kornberg, R. D. (1987) Cell 49, 203–210. 29. Dedrick, R. L., and Chamberlin, M. J. (1985) Biochemistry 24, 2245–2253. 30. Mougey, E. B., and Dennis, D. (1988) Anal. Biochem. 171, 256–265. 31. Lewis, M. K., and Burgess, R. R. (1980) J. Biol. Chem. 255, 4928–4936.
6. Triezenberg, S. J. (1995) in Current Protocols in Molecular Biology, Vol. 1 (Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., Struhl, K., Albright, L. M., Coen, D. M., Varki, A., Eds.) pp. 4.8.1–4.8.5, Wiley, New York.
33. Jeon, C., and Agarwal, K. (1996) Proc. Natl. Acad. Sci. USA 93, 13677–13682.
7. Hawley, D. K., and Roeder, R. G. (1985) J. Biol. Chem. 260, 8163–8172.
34. Tripatara, A., and deHaseth, P. L. (1993) J. Mol. Biol. 233, 349–358.
8. Hawley, D. K., and Roeder, R. G. (1987) J. Biol. Chem. 262, 3452–3461.
35. Daube, S. S., and von Hippel, P. H. (1992) Science 258, 1320– 1324.
9. Conaway, R. C., and Conaway, J. W. (1988) J. Biol. Chem. 263, 2962–2968.
36. Dynan, W. S., and Burgess, R. R. (1981) J. Biol. Chem. 256, 5866–5873.
10. Zheng, X. M., Moncollin, V., Egly, J. M., and Chambon, P. (1987) Cell 50, 361–368.
37. Lescure, B., Bennetzen, J., and Sentenac, A. (1981) J. Biol. Chem. 256, 11018–11024.
11. Conaway, R. C., and Conaway, J. W. (1990) J. Biol. Chem. 265, 7559–7563.
38. Hirose, S., Tsuda, M., and Suzuki, Y. (1985) J. Biol. Chem. 260, 10557–10562.
AID
Methods A 0471
/
6714$$$161
32. Takagi, Y., Conaway, J. W., and Conaway, R. C. (1995) J. Biol. Chem. 270, 24300–24305.
06-19-97 15:14:08
metaas
AP: Methods A
REINES ET AL.
202
39. Sekimizu, K., Horikoshi, M., and Natori, S. (1984) J. Biochem. (Tokyo) 96, 1859–1865. 40. Gu, W., and Reines, D. (1995) J. Biol. Chem. 270, 11238– 11244.
58. 59.
41. Gu, W., Wind, M., and Reines, D. (1996) Proc. Natl. Acad. Sci. USA 93, 6935–6940.
60.
42. Powell, W., and Reines, D. (1996) J. Biol. Chem. 271, 22301– 22304.
61.
43. Horikoshi, M., Sekimizu, K., and Natori, S. (1984) J. Biol. Chem. 259, 608–611.
62.
44. Price, D. H., Sluder, A. E., and Greenleaf, A. L. (1989) Mol. Cell. Biol. 9, 1465–1475.
63.
45. Bengal, E., Goldring, A., and Aloni, Y. (1989) J. Biol. Chem. 264, 18926–18932.
64.
46. Kephart, D. D., Marshall, N. F., and Price, D. H. (1992) Mol. Cell. Biol. 12, 2067–2077.
65.
47. Izban, M. G., and Luse, D. S. (1991) Genes Dev. 5, 683–696. 48. Bengal, E., Flores, O., Krauskopf, A., Reinberg, D., and Aloni, Y. (1991) Mol. Cell. Biol. 11, 1195–1206. 49. Kang, M. E., and Dahmus, M. E. (1993) J. Biol. Chem. 268, 25033–25040. 50. Izban, M. G., and Luse, D. S. (1992) J. Biol. Chem. 267, 13647–13655. 51. Izban, M. G., and Luse, D. S. (1992) Genes Dev. 6, 1342–1356.
66. 67. 68. 69.
52. Izban, M. G., and Luse, D. S. (1993) J. Biol. Chem. 268, 12874–12885.
70.
53. Izban, M. G., and Luse, D. S. (1993) J. Biol. Chem. 268, 12864–12873.
71.
54. Chen, Y., Chafin, D., Price, D. H., and Greenleaf, A. L. (1996) J. Biol. Chem. 271, 5993–5999.
72. 73.
55. Guo, H., and Price, D. H. (1993) J. Biol. Chem. 268, 18762– 18770.
74.
56. Christie, K. R., Awrey, D. E., Edwards, A. M., and Kane, C. M. (1994) J. Biol. Chem. 269, 936–943.
75.
57. Wu, J., Awrey, D. E., Edwards, A. M., Archambault, J., and
AID
Methods A 0471
/
6714$$$161
Friesen, J. D. (1996) Proc. Natl. Acad. Sci. USA 93, 11552– 11557. Reines, D. (1991) J. Biol. Chem. 266, 10510–10517. Marshall, N. F., and Price, D. H. (1992) Mol. Cell. Biol. 12, 2078–2090. Wang, D. G., and Hawley, D. K. (1993) Proc. Natl. Acad. Sci. USA 90, 843–847. Borukhov, S., Sagitov, V., and Goldfarb, A. (1993) Cell 72, 459–466. Kashlev, M., Martin, E., Polyakov, A., Severinov, K., Nikiforov, V., and Goldfarb, A. (1993) Gene 130, 9–14. Wang, D., Meier, T. I., Chan, C. L., Feng, G., Lee, D. N., and Landick, R. (1995) Cell 81, 341–350. Langer, B. R., Waldrop, A. A., and Ward, D. C. (1981) Proc. Natl. Acad. Sci. USA 78, 6633–6637. Arias, J. A., and Dynan, W. S. (1989) J. Biol. Chem. 264, 3223–3229. Arias, J. A., Peterson, S. R., and Dynan, W. S. (1991) J. Biol. Chem. 266, 8055–8061. Wiest, D. K., Wang, D., and Hawley, D. K. (1992) J. Biol. Chem. 267, 7733–7744. Marshall, N. F., Peng, J., Xie, Z., and Price, D. H. (1996) J. Biol. Chem. 271, 27176–27183. Xie, Z., and Price, D. H. (1996) J. Biol. Chem. 271, 11043– 11046. Eilat, D., Hochberg, M., Fischel, R., and Laskov, R. (1982) Proc. Natl. Acad. Sci. USA 79, 3818. Eilat, D., Hochberg, M., Pumphrey, J., and Rudikoff, S. (1984) J. Immunol. 133, 489–494. Reines, D. (1991) Anal. Biochem. 196, 367–372. Gu, W., Powell, W., Mote, J., and Reines, D. (1993) J. Biol. Chem. 268, 25604–25616. Ashley, C. T., Wilkinson, K. D., Reines, D., and Warren, S. T. (1993) Science 262, 563–566. Krummel, B., and Chamberlin, M. J. (1992) J. Mol. Biol. 225, 221–237.
06-19-97 15:14:08
metaas
AP: Methods A